Introduction
Herpes simplex virus 2 (HSV2) infects 30% of North American adults and 50–80% of adults in developing countries [1–3]. There is substantial epidemiological evidence that HSV2 infection enhances both HIV susceptibility [4–6] and sexual transmission [7–10]. HSV2 infection may be associated with a higher blood HIV viremia [11], and HSV2 suppression decreased not only the amount and frequency of genital HIV shedding, but also the blood HIV viral load [4,11].
This epidemiological ‘negative synergy’ [12] between HIV and HSV2 may be an important factor driving HIV transmission in sub-Saharan Africa, since approximately 50% of the general population in the region are infected with HSV2 [1], and mathematical modeling predicts that at this prevalence, more than half of new HIV infections may be directly attributed to HSV2 [5,13]. Negative synergy between HIV and HSV2 may also have particular relevance for HIV ‘core groups’, such as female commercial sex workers (FSWs), who are at high risk for both infections [14]. HSV2 infection has been associated with an almost six-fold increase in HIV incidence in FSWs [6], and over 80% of new HIV infections in men in some African cities may be acquired via transactional sex with HIV-infected FSWs [15].
Despite these established epidemiological associations, the biologic basis of the negative synergy between HIV and HSV2 is unclear [3]. A physical breach in the mucosal barrier due to genital micro-ulceration and/or mucosal infiltration of HIV target cells during HSV2 reactivation are possible mechanisms for increased HIV susceptibility [7]. In HIV–HSV2 co-infected individuals, HSV2 infection has been associated with increased levels and frequency of genital HIV shedding [1,16], which is likely to be the basis for enhanced HIV sexual transmission. Underlying these increases in HIV shedding may be enhancement of HIV-long terminal repeat (LTR) transcription and HIV replication by HSV2 proteins [17–20], and co-infection of the same human CD4+ cell by HIV and HSV2 [21], resulting in increased HIV replication.
The exact pathogenesis of HIV sexual transmission remains undefined. The CCR5 co-receptor appears necessary for sexual acquisition of HIV [22], and R5-tropic strains of HIV attach to CD4 and then bind CCR5 prior to entering and infecting cells [23]. HIV may also bind the DC-SIGN lectin on mucosal dendritic cells (DCs), circumvent endosomal antigen processing and exploit DC migration to infect CD4+ T cells in trans at remote sites [24–27].
To elucidate the potential mucosal immune basis for negative HIV–HSV2 synergy, a detailed ex vivo analysis of the female genital tract (FGT) mucosal immune milieu was performed in a well-defined cohort of Kenyan FSWs. We characterized the associations of HIV and HSV2 with changes in the FGT immune milieu by measuring genital tract immune cell populations (CD4+/CCR5+/CD69+ T cells and DC-SIGN+ or TLR9+ mucosal DC), cytokine/chemokine protein levels and immune gene expression profiles. The findings suggest a mucosal vicious circle in which HSV2 infection increases HIV susceptibility, HIV infection impairs HSV2 mucosal immune control, and local HSV2 reactivation enhances both HSV2 and HIV transmission. These findings demonstrate that HSV2 infection may alter the FGT immune milieu to increase HIV acquisition by the susceptible, and to amplify HIV transmission from the co-infected, and have direct implications for the design of novel HIV prevention strategies.
Methods
Participant enrolment and sampling
Women were enrolled through a dedicated FSW clinic in the Pumwani district of Nairobi, Kenya, from November 2004 to January 2005. Informed consent was obtained from all study participants, and the study protocol was approved by the Research Ethics Boards at the University of Toronto, the University of Manitoba and Kenyatta National Hospital (Nairobi, Kenya). Genital tract samples were collected in the following order: a cervico-vaginal lavage (CVL) of the ectocervix with 2 ml of sterile phosphate-buffered saline (PBS), re-aspirated from the posterior fornix region; a vaginal swab for Trichomonas vaginalis culture and Gram stain; a cotton-tipped swab from the cervical os placed into viral transport medium (Hank's Balanced salt solution, 20 mmol/l HEPES, 0.5% gelatin, 10 mg/ml gentamicin, 4 mg/ml amphotericin); an endocervical swab for Neisseria gonorrhea culture; a scraping of the external cervical os using a plastic cell scraper (Benzi Jinshuo Applicator Co., Liaoning, China) in 10 ml PBS, which was combined with a cervical lavage using 1 ml PBS to collect loosened cells; and a cervical cytobrush (Histobrush, Spectrum Lab, Dallas, Texas, USA), which was rotated through 360° in the cervical os, and placed into 2 ml of RNA storage buffer (RNALater; Ambion, Austin, Texas, USA). Cervical samples were not collected from women who were actively menstruating. All samples were placed on ice and transported to the laboratory within 2 h. One tube (5 ml) of blood was drawn into ethylenediamine tetraacetic acid, and three tubes (24 ml) into heparin. Blood was transported to the laboratory at room temperature. All immunologic and viral shedding assays were performed by laboratory staff blinded to HSV2 and HIV-1 infection status.
Diagnostic procedures
HSV2 infection status was determined by IgG enzyme-linked immunosorbaent assay (ELISA) (Kalon Biological, Guildford, UK) [6]. HIV-1 serology used a synthetic peptide enzyme immunoassay (Detect HIV; Biochem ImmunoSystems Inc., Montreal, Canada), positive tests were confirmed using a recombinant antigen enzyme immunoassay (Recombigen HIV-1/2 EIA; Cambridge Biotech Corporation, Galway, Ireland), and CD4 T-cell counts were assayed for all participants (FACScan; Becton Dickinson Immunocytometry Systems, San Diego, California, USA). Cytomegalovirus (CMV) infection status was determined using the AxSYM CMV IgG assay (Abbott Laboratories, Abbott Park, Illinois). Trichomonas vaginalis culture was performed by In Pouch TV (Biomed Diagnostics, San Jose, California, USA), a Gram stain was performed, and blood was tested for syphilis serology (rapid plasma reagin; RPR). Bacterial vaginosis was defined as a Nugent score of 7–10 [28]. If a genital ulcer was present on physical examination, swabs were taken for Haemophilus ducreyii culture. No diagnostics were performed for Chlamydia trachomatis. All diagnosed infections were treated according to Kenyan national guidelines, including the provision of antiretrovirals for HIV-1-infected women.
Viral shedding in the genital tract
An endocervical swab was collected into 2 ml of viral transport medium (VTM; see above). Cervical and blood plasma HIV-1 RNA was measured using the Versant bDNA kit (limit of detection 50 copies/ml; Bayer Diagnostics, Emeryville, California, USA). HSV2 DNA was quantified with the RealArt HSV1/2 LC PCR kit (lower limit of detection 10 copies/ml; Artus Biotech, Hamburg, Germany). CMV shedding has been found to be associated with increased detection of HIV shedding in both male and female genital tracts [29–31]; hence cervical CMV shedding was also measured in this study. CMV DNA was quantified in 250 μl of CVL secretions with the RealArt CMV LC polymerase chain reaction (PCR) kit (lower limit of detection 1 copy/ml; Artus Biotech). All shedding assays were performed by laboratory personnel blinded to participant clinical status.
Assays of immune proteins
Levels (pg/ml) of 15 cytokines and chemokines were measured in cervico-vaginal lavage by Cytokine Bead Array (BD Biosciences, San Diego, California, USA) [32]; these included interleukin (IL)-2, IL-4, IL-5, IL-10, tumor necrosis factor (TNF)-α, interferon (IFN)-γ, IL-1β, IL-8, IL-6, IL-12p70 regulated on activation normal T-cell expressed and secreted (RANTES), Monokine induced by IFN-γ (MIG/CXCL9), macrophage chemotactic protein (MCP1/CCL2), and interferon inducible protein-10 (IP-10/CXCL10). ELISA was used to measure CVL IFN-α (Multisubtype IFNα ELISA; PBL Biomedical Laboratories, Piscataway, New Jersey, USA).
Cervical immune cell populations
Cells from the cervical scraping were filtered through a 100 μm filter (Fisher Scientific, Pittsburgh, Pennsylvania, USA), washed and divided into two equal aliquots for staining with a panel of DC or T-cell markers. Cells in the T-cell aliquot were stained with CD69-FITC, CCR5-PE, CD4-PerCP and CD3-APC (BD Pharmingen, San Jose, California, USA); cells in the DC aliquot were stained with CD1a-FITC, TLR9-PE (Imgenex San Diego, California, USA), DC-SIGN-APC (eBioscience, San Diego, California, USA) and CD4-PerCP (BD Pharmingen). Appropriate isotype control antibodies were used in parallel. Samples were processed using FACSCalibur (Becton-Dickinson Immunocytometry Systems). Mononuclear and larger granular cell populations were identified by distinct forward and side scatter properties [33]. Flow cytometry was performed by research personnel blinded to participant HSV2 infection and shedding status. Cell numbers enumerated by flow cytometry (in the T-cell or DC aliquot) were multiplied by two to determine ‘cells per cytobrush’ and then log10 transformed for analysis and presentation.
Cervical immune gene expression
RNA was extracted from one cervical cytobrush by a modified guanidine isothiocyanate method [34], using Trizol LS (Sigma, St. Louis, Missouri, USA). RNA was quantified spectrophotometerically, for integrity and contaminating DNA removed with DNAse (Ambion). RNA was reverse-transcribed to complementary DNA (cDNA) with random-nanomer primers and Omniscript reverse transcriptase (Qiagen, Valencia, California, USA). Quantitative PCR (QPCR) assay was performed (ABI PRISM 7900 Sequence Detection System; Perkin Elmer Applied Biosystems, Foster City, California, USA) in 384-well plate format with the SYBR green method using Platinum Taq polymerase. Assays using exon sequence PCR primers (Primer Express software; Perkin Elmer Applied Biosystems) were performed in triplicate, and repeated twice with independent cDNA preps. Quantitative PCR values (cycle threshold; CT), were obtained during exponential amplification, and verified using dissociation curves. Genomic DNA template (human placenta gDNA) dilution standard curves (5 log range) were conducted with each primer pair to verify a linear template–CT relationship (r2 for all primer pairs > 0.9). We assayed several candidate housekeeping genes [β-actin, B2M, GAPDH, HPRT, RPL32 and TATA binding protein (TBP)]. Only TBP, GAPDH and B2M expression levels did not fluctuate with HIV or HSV2 infection status, with TBP showing the least variability. Target gene expression levels for each primer set were normalized to TBP expression, and reported as an average ratio.
Statistical analysis
All analyses were performed with SPSS version 11.0 software (Chicago, Illinois, USA). Associations of continuous variables with HIV–HSV2 infection status or shedding were determined using a Mann–Whitney U non-parametric test. Comparisons between discrete variables were performed using the chi-squared test with calculations of likelihood ratios (LR). Bivariate correlations were calculated using the Pearson two-tailed test.
Results
Epidemiological associations between HIV and HSV2
The study population consisted of a cross-sectional sample of 102 Kenyan FSWs from a well-characterized Nairobi cohort [35]. Eleven women were taking antiretroviral therapy, and were excluded from analysis: of the remainder, 55 were HIV uninfected, and 36 were HIV infected, antiretroviral therapy-naive (Table 1). HIV–HSV2 epidemiological synergy was apparent. HIV-infected sex workers were more likely to be infected by HSV2 (36/36, 100% versus 42/55, 76%; LR = 14.5; P < 0.001; Fig. 1), and also to be shedding HSV2 DNA in the cervix (9/36, 25% versus 0/42, 0%; LR = 15.3; P < 0.001). Cervical HSV2 shedding was only observed in HIV-infected FSWs, and clinically apparent genital ulceration was present in just 11% (1/9) FSWs shedding HSV2. HSV2 shedding was not associated with alterations in blood CD4 cell count (shedders, 442 cells/μl versus non-shedders, 557 cells/μl; P = 0.2) or blood HIV viral load (22 101 versus 20 434 copies/ml or 3.73 versus 3.99 log10 copies/ml; P = 0.8). Cervical HIV shedding was detected in 10 of 36 FSWs (28%), and tended to be more common in those shedding HSV2 (4/9, 44% versus 6/27, 22%; P = 0.2).
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Table 1: Demographic characteristics of study participants, by HIV serostatus.
Fig. 1: Study profile and infection status of female sex worker participants. All female sex workers (FSWs) were screened and enrolled during the biannual clinic resurvey, November 2004–January 2005. HIV indicates HIV type 1; HSV2, herpes simplex virus type 2. Statistical comparisons used the Mann–Whitney U non-parametric test or chi-squared test with calculation of likelihood ratios.
HSV2 infection associated with increases in HIV target cells in the genital mucosa
The potential association of HSV2 infection status in HIV-negative FSWs with changes in genital CD4/CCR5+ T-cell or CD1a/DC-SIGN+ iDC populations was assessed (Fig. 2a; representative FACS plot of cervical cell populations). There were no differences in the absolute number of cervical CD4+ T cells, but there was a 0.3 log10 increase (three-fold higher absolute cell numbers) of CD4/CCR5+ T cells in the cervix of HSV2-infected FSWs (P < 0.05; Fig. 2b). Cervical CD4+ T cells were also more activated, with a greater absolute number and proportion (46% in HSV2+ versus 16% in HSV2) expressing CD69 (P < 0.001 for both; Fig. 2b). Strikingly, there was an entire log10 increase (ten-fold higher absolute cell numbers) in DC-SIGN+ iDCs (P < 0.001; Fig. 2c), and an increased proportion of cervical iDCs expressed DC-SIGN (11.9% in HSV2+ versus 4.6% in HSV2–, P < 0.05; Fig. 2c). Further characterization demonstrated that over 90% of cervical CD1a+ iDCs co-expressed CD11c, suggesting that these cells were of myeloid origin (data not shown).
Fig. 2: Association of chronic herpes simplex virus type 2 (HSV2) infection with changes in cervical immune cell populations in HIV-uninfected sex workers. (a) A representative forward (FSC) versus side (SSC) scatter contour plot of a cervical cytobrush specimen is shown with granular (larger cells) and lymphocyte (smaller mononuclear cells) gates drawn. Dendritic cell populations found in the granular gate are shown as CD1a (x-axis) versus TLR9 or DC-SIGN. In the lymphocyte gate, CD3 versus CD4 T-cell populations are shown, as well as CD3+ CD4+ T cells versus CCR5 or CD69. (b) The number (log10 transformed ‘cervical cells/cytobrush’ or percentage of cell subset) and characteristics of immune cell populations in the cervix of female sex workers (FSWs) are shown for women uninfected by HSV2 (N = 13; white bars) and infected by HSV2 (N = 42; shaded bars). CD4+ T cells and subsets are shown: total CD4+ T cell numbers (left); the total number of CD4+ T cells expressing CCR5 (centre left); the total number of CD4+ T cells expressing the activation marker CD69 (centre right); and the percentage of CD4+ T cells expressing CD69 (right). (c) immature dendritic cells (iDCs) and subsets are shown: total iDC numbers (left); the total number of iDCs expressing DC-SIGN (centre); and the percentage of iDCs expressing DC-SIGN (right). The data represent mean ± standard deviation. * P < 0.05 and ** P < 0.001.
Overall, HSV2 infection was associated with increases in genital HIV target cell populations, even in the absence of genital ulceration or HSV2 reactivation. We did not find any significant association between HSV2 infection and cervical cytokine or chemokine levels in HIV-uninfected FSWs. Since HSV2 shedding was not detected in HIV-negative FSWs, changes in the genital immune milieu related to HSV2 reactivation could not be examined.
HIV infection associated with iDC depletion in the genital mucosa
In comparison with HIV-negative women, there was a profound depletion of iDC populations in the genital mucosa of HIV–HSV2 co-infected FSWs. Cervical CD1a+ iDCs were reduced (P < 0.05; Fig. 3a), particularly iDC subsets expressing either DC-SIGN (P < 0.05) or Toll-like receptor 9 (TLR9; P = 0.05), an innate immune receptor that binds CpG motifs on HSV2 DNA to trigger signaling cascades that culminate in type 1 interferon and inflammatory cytokine release, and a potentially important mediator of HSV2 immune control [36,37]. This iDC depletion was independent of systemic immune status (blood CD4+ T-cell counts or CD4/CD8 ratio; data not shown). HIV-infected FSWs had increases in cervical CD3+ T cells (P < 0.05) and CD8+ T cells (P = 0.001; Fig. 3a), without differences in activation levels. Cervical CD4+ T-cell numbers did not vary with HIV status (P = 0.7), but the CD4/CD8+ T-cell ratio was decreased in both the blood (P < 0.001) and cervix (P < 0.001) of HIV-infected FSWs, and to a similar degree at both sites (ratio 0.5 in blood versus 0.6 in the cervix; P = 0.5). There were no significant differences in CCR5+CD4+ cervical cell populations (absolute numbers, log10 transformed or % CCR5 expression by CD4 cells) between HIV-uninfected and infected women, although there was a trend (P = 0.07) for a decrease in the proportion of CCR5 expressing CD4+ T cells in HIV-infected FSWs. Although there were no significant differences in cervical cytokines/chemokines levels between HIV-infected and uninfected women, there were some trends to increased cytokines/chemokines in HIV-infected FSWs (MCP, IP-10, IL-10, IL-5, IL-2; P < 0.1 for all). HIV infection status was not associated with overall differences in the expression of TLRs 1–10 or FoxP3.
Fig. 3: Associations of HIV serostatus and cervical HIV-1 RNA shedding. (a) HIV infection was associated with depletion in cervical CD1a+ immature dendritic cells (iDCs), TLR9+ iDCs and DC-SIGN+ iDCs, and increases in CD3+ and CD8+ T cells. The data represent mean ± standard deviation for N = 55 HIV uninfected and N = 33 HIV-infected female sex workers (FSWs). (b) HIV shedding in N = 10 FSWs was associated with increases in both chemokines [interferon inducible protein-10 (IP-10), macrophage chemotactic protein (MCP), monokine induced by IFN-γ (MIG), regulated on activation normal T-cell expressed and secreted (RANTES)] and pro-inflammatory cytokines [interleukin (IL)-1β, IL-8, IL-6 and interferon (IFN)-γ]. (c) In HIV shedders, the amount of cervical HIV RNA correlated with the level of cervical herpes simplex virus (HSV) DNA (r 2 = 0.9, P < 0.001). * P < 0.05 and ** P < 0.001.
Immune associations of genital tract HIV shedding
Cervical HIV shedding was detected in 10 of 36 FSWs (28%), and was strongly associated with immune activation in the genital tract. Levels of chemokines (IP-10, MCP, RANTES; all P < 0.001 and MIG; P < 0.05) and inflammatory cytokines (IL-1β, IL-8, IL-6 and IFN-γ; all P < 0.05; Fig. 3b) were elevated in the cervico-vaginal secretions of the 10 HIV shedders. In this subgroup, absolute HIV levels were inversely correlated with TLR9+ iDC numbers (r2 = −0.7; P < 0.05) and with mRNA expression of TLR8 (r2 = −0.9; P = 0.001) and TLR9 (r2 = −0.7; P < 0.05). Cervical HIV load correlated with both the number and proportion of activated cervical CD4+ T cells expressing CD69 (r2 = 0.7; P < 0.05; and r2 = 0.7; P < 0.05, respectively). Neither blood CD4 cell counts, blood HIV viral load, nor cervical immune cell populations varied significantly with HIV RNA shedding status.
Immune associations of genital tract HSV2 shedding
Cervical HSV2 shedding was only detected in HIV-infected sex workers, and the level of cervical HIV RNA was strongly correlated with HSV2 DNA levels (r2 = 0.9; P < 0.001: Fig. 3c). Activated cervical CD4+ CD69+ T cells were increased in the genital mucosa of FSWs shedding HSV2 (202 versus 68 cells/cytobrush; P < 0.05, Fig. 4a), as were cervical levels of chemokines (P < 0.05 for all; Fig. 4b), but not pro-inflammatory cytokines. HSV2 levels correlated with the number and proportion of CD4 T cells expressing CCR5 (r2 = 0.7; P < 0.05; and r2 = 0.7; P = 0.02; respectively) (Fig. 4c), and with levels of the chemokines IP-10, MCP, MIG and RANTES (all r2 > 0.6; all P ≤ 0.05). Cervical iDC numbers were unchanged in FSWs shedding HSV2 (13 588 versus 10 696 cells/cytobrush; P = 0.2), although fewer iDCs expressed TLR9 (3.1 versus 11.6%; P = 0.001). HSV2 DNA levels correlated inversely with the number of DC-SIGN+ iDCs (r2 = −0.8; P < 0.05) and TLR9+ iDCs (r2 = −0.8; P < 0.05) (Fig. 3d, e). This suggests that TLR9+ and DC-SIGN+ iDCs may mediate local immune control of HSV2 reactivation at a genital level.
Fig. 4: Associations of herpes simplex virus (HSV) shedding in HIV-infected female sex workers (FSWs). (a) Women shedding HSV type 2 (HSV2) had increased genital CD69+ CD4+ T cells, and decreased number and proportion of TLR9+ genital immature dendritic cells (iDCs). (b) HSV2 shedding was associated with increases in chemokines [interferon inducible protein-10 (IP-10), macrophage chemotactic protein (MCP), regulated on activation normal T-cell expressed and secreted (RANTES) and monokine induced by interferon (IFN)-γ (MIG)] (all P < 0.005), but not with differences in pro-inflammatory cytokine levels. (c) The level of HSV2 DNA was positively correlated with the proportion of cervical CD4+ T cells expressing CCR5 and (d) negatively correlated with the number of cervical iDC expressing DC-SIGN (r 2 = −0.8; P < 0.05), (e) and TLR9 (r 2 = −0.8, P < 0.05). The data represent mean ± standard deviation. * P < 0.05 and ** P < 0.001. IL, interleukin.
Immune associations of other genital tract infections
Other genital co-infections may also alter HIV susceptibility [38]. Although CMV is also a Herpes virus and CMV shedding was more common in HIV-infected FSWs (22 versus 2%; LR = 10.6; P < 0.05), no associations were seen between CMV shedding and immune cell populations, cytokines or gene expression. BV was detected in 42 and 27% of HIV-uninfected and infected FSWs, respectively, but no significant associations were seen between genital immune milieu and BV or Nugent score in either group. Infection by T. vaginalis, N. gonorrhoeae and T. pallidum were uncommon (Table 1) and were not associated with genital immune changes.
Discussion
This study confirms the substantial epidemiological synergy reported between HIV and HSV2, and is the first to demonstrate that this may be underpinned by a profound negative ‘mucosal synergy’ between these viruses in the female genital tract. In HIV-negative at-risk women, HSV2 infection was associated with increases in genital mucosal target cell populations that would be expected to increase susceptibility to HIV infection. These mucosal immune changes were evident even in the absence of genital ulceration or HSV2 reactivation, implying that HSV2 induces a persistent state of increased mucosal HIV susceptibility. The ten-fold increase in cervical iDCs expressing the DC-SIGN lectin, and the three-fold increase in cervical CD4+ T cells expressing the HIV co-receptor CCR5, provides a putative biological explanation for the observation that HSV2 seropositivity is a strong independent risk factor for HIV acquisition [6–9] even in the absence of symptomatic ulceration.
Both DC-SIGN and CD1a are important in mucosal immune responses. Upon antigen encounter, immature dendritic cells mount innate immune responses and undergo a coordinated series of dynamic cellular events that lead to maturation, antigen peptide loading onto MHC class II molecules, migration to lymph nodes and antigen presentation to T cells to induce adaptive immune responses. DC-SIGN binding of various viral and microbial pathogens triggers DC activation and/or maturation, as well as signaling via Toll-like receptors [25,39]. In addition, CD1a molecules can bind and efficiently present antigen to T cells in a maturation-independent manner [40]. Intravaginal inoculation of mice with HSV2 led to rapid recruitment of submucosal DCs to the infected epithelium, followed by local stimulation of IFNγ production from HSV-specific CD4+ T cells [41] and submucosal DCs were the primary cells responsible for priming and mounting protective T-cell helper 1 responses during HSV2 infection [42]. HSV2 infection of rhesus macaques impaired iDC maturation and promoted the release of chemokines RANTES and MIP1α, offering another possible explanation for the association of increased mucosal iDCs with HSV2 infection, and for increased genital chemokine levels during HSV2 reactivation. Together with these studies, our in vivo demonstration that HSV2 infection was associated with increased cervical DC-SIGN+ CD1a+ iDCs suggests that these cells may be important in mediating local immune control of HSV2.
However, DC-SIGN has also been demonstrated to permit the propagation of productive HIV infection in CD4+ T cells at low viral titers [27], such as are present in the genital mucosa soon after sexual acquisition of HIV [43]. Therefore, HIV may exploit the local increases in DC-SIGN+ iDCs mediating HSV2 immune control in the genital mucosa, to enhance host susceptibility to HIV infection after sexual exposure.
HIV infection was associated with an overall depletion of cervical iDCs, as well as depletion of iDC subpopulations expressing DC-SIGN+ and/or TLR9+, regardless of HIV disease stage. HIV nef has been shown to reduce surface expression of CD1a on iDCs through redistribution into late endosomal/lysosomal compartments [44]. Therefore, we cannot determine whether HIV infection was associated with a true depletion of iDCs, or a downregulation of CD1a surface expression by iDCs. Regardless, the reduced numbers of cervical iDCs expressing CD1a was strongly associated with local HSV2 reactivation in HIV-infected FSWs, again implying that these iDCs mediate local HSV2 immune control, and that their depletion in the context of HIV infection may further enhance the sexual transmission of both HSV2 and HIV.
The association of HSV2 and HIV shedding levels in FSWs co-infected by both HIV and HSV2 suggests that HSV2 infection may increase HIV transmission from this core group to their male clients. During reactivation, HSV2-encoded proteins may directly increase transcription from HIV-LTR [17,19–21]. HSV2 shedding was also associated with increased chemokine levels and the amount of HSV2 viral DNA was positively correlated with CCR5+ CD4+ cells. CCR5 receptor engagement by HIV envelope glycoproteins activates signalling that culminates in the release of various chemokines that generate a chemokine gradient that recruits T cells to sites of inflammation [45–47]. The increase in CCR5+ cells and chemokines during episodes of HSV2 shedding may recruit activated CD4+ T cells to the genital tract, and therefore enhance local replication of HIV [43] and consequently increase HIV transmission during sexual intercourse.
Overall, we observed strong immune synergy between HSV2 and HIV specifically in the mucosal genital compartment that may have direct implications for HIV prevention strategies. Our ex vivo approach allowed direct characterization of the female genital immune milieu, and elucidation of the immediate HSV2-associated mucosal changes that may underlie increased sexual HIV transmission. The resulting static ‘snap shot’ of the FGT immune milieu does not permit study of the extremely dynamic processes of immune cell migration to and from the genital mucosa, but does allow precise definition of the cell populations present in the mucosa at the time of sexual exposure to HIV, which is likely to be a critical determinant of transmission.
In sub-Saharan Africa, over half of new HIV infections occur in women, with young women from 15–24 years of age being at particularly high risk [48]. Given the expected lack of a protective HIV vaccine in the near future, these statistics highlight the urgent need for an accessible alternative to prevent HIV infection [49]. Our finding of increased DC-SIGN+ iDCs and CCR5+CD4+ T cells in the cervix of HSV2-infected women implies that microbicides targeting these molecules may hold promise, and such compounds are in early clinical testing [50,51]. Since DC-SIGN+ dendritic cells in the rectal mucosa also bind HIV avidly, an effective microbicide targeting this molecule might be important for both men and women practicing anal intercourse [52]. Our results strongly suggest that HSV2 suppression, both in HIV-infected and uninfected individuals, holds promise as an HIV prevention strategy, and supports the rationale for large-scale clinical trials that are currently testing this hypothesis [4].
Acknowledgements
We thank Dr. Mario Ostrowski for helpful discussions and critical reading of the manuscript, Jane Kamene and the Pumwani clinic nurses for study recruitment and providing treatment, Ann Miangi, Nyakio Chinga and the laboratory staff at the University of Nairobi Microbiology Annex for specimen processing and performing diagnostic assays. Above all, we thank the women of the Pumwani ML cohort for their continued participation and support of our studies.
Sponsorship and role of the funding source: This work was supported by the Canadian Institutes of Health Research (CIHR; R.K., grant; K.F., grant; F.A.P., grant), the Canada Research Chair Programme (R.K., salary support); the National Institutes of Health (NIH; F.A.P., grant); the Canadian Network for Vaccines and Immunotherapeutics (CANVAC; R.K., grant; A.R., salary support) and the Ontario HIV Treatment Network (OHTN; A.R., fellowship; R.K., grant). The funding source had no role in study design, data collection, analysis or interpretation, or in report writing or submission.
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