Strong viremia control in vaccinated macaques does not prevent gradual Th17 cell loss from central memory

Abstract

It has been proposed that microbial translocation might play a role in chronic immune activation during HIV/SIV infection. Key roles in fighting bacterial and fungal infections have been attributed to Th17 and Tc17 cells. Th17 cells can be infected with HIV/SIV, however whether effective vaccination leads to their maintenance following viral challenge has not been addressed. Here we retrospectively investigated if a vaccine regimen that potently reduced viremia post-challenge preserved Th17 and Tc17 cells, thus adding benefit in the absence of sterilizing protection. Rhesus macaques were previously vaccinated with replication-competent Adenovirus recombinants expressing HIVtat and HIVenv followed by Tat and gp140 protein boosting. Upon SHIV_89.6P challenge, the vaccinees exhibited a significant 4 log reduction in chronic viremia compared to sham vaccinated controls which rapidly progressed to AIDS (Demberg et al. J. Virol. 81:3414, 2007). Plasma and cryopreserved PBMC samples were examined pre-challenge and during acute and chronic infection. Control macaques exhibited a rapid loss of CD4⁺ cells, including Th17 cells. Tc17 cells tended to decline over the course of infection although significance was not reached. Immune activation, assessed by Ki-67 expression, was associated with elevated chronic viremia of the controls. Significantly increased plasma IFN-γ levels were also observed. No increase in plasma LPS levels were observed suggesting a lack of microbial translocation. In contrast, vaccinated macaques had no evidence of immune activation within the chronic phase and preserved both CD4⁺ T-cells and Tc17 cells in PBMC. Nevertheless, they exhibited a gradual, significant loss of Th17 cells which concomitantly displayed significantly higher CCR6 expression over time. The gradual Th17 cell decline may reflect mucosal homing to inflammatory sites and/or slow depletion due to ongoing low levels of SHIV replication. Our results suggest that potent viremia reduction during chronic SHIV infection will delay but not prevent the loss of Th17 cells.

Introduction

The main route of HIV transmission worldwide is via heterosexual contact across mucosal epithelia. The virus first establishes a small localized infection in the submucosa before rapidly spreading systemically (Haase, 2005; Miller et al., 2005). Despite the rapid systemic dissemination, the majority of infected cells reside in the gut (Ling et al., 2010; Mattapallil et al., 2005). One hallmark of HIV infection of people and SIV infection of macaques is early rapid depletion of CD4⁺ T-cells in the gut mucosa, which also occurs in natural non-human primate hosts of SIV, although the natural hosts generally do not progress to simian AIDS (Brenchley and Douek, 2008a; Brenchley et al., 2004; Gordon et al., 2007; Mattapallil et al., 2005). In addition to CD4⁺ T cell loss, chronic immune activation of CD8⁺ T-cells is associated with disease progression, and in fact can be used to distinguish elite controllers from HIV progressors (Bello et al., 2009; Loke et al., 2010; Vollbrecht et al., 2010). HIV infection also leads to mucosal epithelial barrier damage, in part driven by the viral envelope protein, gp120 (Nazli et al., 2010). This damage to the epithelium can lead to enhanced microbial translocation, which has been associated with disease progression and may contribute to chronic immune activation (Brenchley et al., 2006; Klatt et al., 2010; Marchetti et al., 2008; Troseid et al., 2010).

IL-17 producing cells are of current interest due to their association with control of fungal and bacterial infections. These cells can be divided into three major phenotypes: Th17 cells (CD4⁺), Tc17 cells (CD8⁺) and IL-17⁺ γδ T-cells. The most commonly studied and best characterized are Th17 cells. They play a distinct role in the clearance of bacterial and fungal infections and have been shown to be involved in autoimmune disease in several animal models and in humans (Crome, Wang, and Levings, 2010; Damsker, Hansen, and Caspi, 2010; Hashimoto et al., 2010; Kamada et al., 2010; Wu et al., 2010; Zhao et al., 2009). Tc17 cells on the other hand seem to play a role similar to Th1 cells in anti-tumor immunity (Kuang et al., 2010) and also function in the clearance and control of viral infection (Hamada et al., 2009). IL-17⁺ γδ T-cells (Vδ1, Vδ2 cells) have been associated with responses to Candida albicans (Vδ1 cells) and to mycobacterium (Vδ2 cells) in HIV infected patients (Fenoglio et al., 2009) and with clearance of Listeria monocytogenes infection in mice (Xu et al., 2010). The role of γδ T-cells in protective immunity is further reviewed by Cua and Tato (Cua and Tato, 2010).

The origin of Th17 cells has not been definitively established; however the current consensus is that murine Th17 cells are related to Tregs, both originating from the same precursors, whereas human Th17 cells are more closely related to Th1 cells (Annunziato et al., 2008; de Jong, Suddason, and Lord, 2010; Romagnani et al., 2009; Torchinsky and Blander, 2010). Presumably the origin of non human primate Th17 cells will resemble that of humans, but to date this is not known. In addition to IL-17, Th17 and Tc17 cells produce a vast array of other cytokines including TNF-α, IL-1, IL-2, IL-10, IL-21, IL-22 and IFN-γ (Klatt and Brenchley, 2010; Kuang et al., 2010; Ndhlovu et al., 2008; Torchinsky and Blander, 2010). A few reports have shown that retroviral infection, including HIV (Fenoglio et al., 2009; Maek et al., 2007) and HTLV-1 (Dodon et al., 2004), leads to IL-17⁺ cell expansion. However, most studies have found Th17 cells to be lost or declining during SIV and HIV infections, either by homing to the mucosa or due to their susceptibility to infection (Brenchley et al., 2008; Hunt, 2010; Ndhlovu et al., 2008; Prendergast et al., 2010). The majority of these studies have compared IL-17⁺ cells in SIV-infected natural hosts and susceptible macaque species, or in healthy volunteers, HIV-infected non-progressors (elite controllers) and disease progressors. A recently published paper by Nigam et al. (2011) shows the distribution of Th17 and Tc17 populations in different tissues in healthy and SIV infected unvaccinated macaques. However, the question of whether vaccines that succeed in significantly reducing viral loads following challenge can prevent the loss of Th17 and Tc17 cells has not been addressed.

Our vaccine approach is based on priming with replication-competent Adenovirus vectors and boosting with envelope protein. This strategy has elicited strong protection in both SIV and SHIV challenge models in rhesus macaques (Demberg et al., 2007; Malkevitch et al., 2006; Patterson et al., 2008; Patterson et al., 2011; Patterson et al., 2004) and in the HIV challenge model in chimpanzees (Lubeck et al., 1997). Recently, priming of rhesus macaques with adenovirus 5 host range mutant (Ad5hr) recombinants encoding HIVenv and HIVtat followed by boosting with Tat and Env protein led to strong protection against a SHIV_89.6P challenge (Demberg et al., 2007). While peak viremia in the vaccine and control groups was similar, the vaccinated animals controlled infection rapidly thereafter, leading to a significant 4-log decrease in chronic phase viremia compared to sham-vaccinated controls. CD4⁺ cells were preserved in contrast to control animals that lost CD4⁺ T-cells almost completely within two weeks. Here, using cryopreserved PBMC and plasma samples from this study and from six additional control animals similarly sham-vaccinated and challenged (Patterson et al., 2008), we asked whether the vaccinated macaques preserved their IL-17-secreting cells, and avoided chronic immune activation in contrast to the controls. Such an outcome would add benefit to partially protective vaccines by potentially reducing viral-induced mucosal damage, preventing the occurrence of opportunistic infections, slowing disease progression, and extending the time period before HAART therapy has to be initiated.

Material and methods

Peripheral blood and plasma specimens

Cryopreserved ficoll-paque plus (GE Healthcare, Piscataway, NJ, USA) gradient derived PBMC from rhesus macaques of Indian origin were used in this study. The PBMC were from two groups of animals from a previously published vaccine study (Demberg et al., 2007). One group of 8 macaques was vaccinated with replication competent Ad5hr-HIV_IIIBtat and Ad5hr-HIV_89.6pgp140ΔCFI followed by boosting with gp140ΔCFI and Tat proteins adjuvanted with MPL-SE and alum, respectively. The other group of 3 controls was supplemented with 6 similarly treated control animals from another study (Patterson et al., 2008). All controls received empty Ad5hr vector and were boosted with adjuvant only or with PBS (3 with MPL-SE plus alum, 3 with MPL-SE only, and 3 with PBS). Vaccinated and 3 control animals were challenged intravenously with SHIV_89.6P at week 50 (12 weeks following the last boost; 30 MID₅₀). The supplemental controls were challenged with the same SHIV_89.6P stock at week 44 (8 weeks following the last adjuvant boost; 90 MID₅₀). Cells were evaluated by flow cytometry and real time PCR at three different time points: pre-challenge/time of challenge (wk48 or wk44), 4 weeks post challenge (wk54/wk48) and during the chronic phase (24 weeks post challenge). Matched plasma samples, derived from EDTA-treated blood, were obtained at similar time points.

PBMC stimulation

PBMC from all animals were thawed, washed with R10 (RPMI1640 supplemented with penicillin/streptomycin, L-glutamine and 10% FBS, all Life Technologies, Carlsbad, CA, USA) counted and plated into two wells of a 24-well plate (Nunc, Thermo-Scientific, Waltham, MA, USA). One well was stimulated for 4 hours in 1 ml of R10 containing 50 ng/ml of PMA (Sigma-Aldrich, St Louis, MO, USA) and 250 ng/ml of Ionomycin (Sigma-Aldrich) to evaluate IL-17-producing cells. One hour into the stimulation, 1 µl of Golgi Stop (BD Bioscience, San Jose, CA, USA) was added. The unstimulated well was used to evaluate CD8⁺ and CD4⁺ T cell activation measured by Ki-67 and background cytokine secretion by flow cytometry.

Limulus Amoebocyte Lysate (LAL) assay

Plasma LPS levels were evaluated using the Hycult chromogenic endpoint LPS detection assay (Limulus amoebocyte lysate assay, Hycult Biotech, Uden, The Netherlands) according to the manufacturer’s instructions. Plasma samples were diluted 1:30 in DPBS (Lonza, Basel, Switzerland) and heat inactivated for 5 min at 75°C prior to assay. EU levels obtained were converted to pg/ml with 1 EU = 100 pg/ml.

16s rDNA quantitative real time PCR

Bacterial DNA was purified from 500 µl of non-inactivated plasma from macaques in the original study (Demberg et al., 2007) by a modified alkaline lysis protocol. Reagents from a GeneJet plasmid purification kit (Fermentas, Glen Burnie, MD, USA) were used under aseptic conditions to avoid bacterial contamination. Plasma samples were spun at 4000×g for 10 min to pellet any free or complexed (opsonized by complement and/or antibody) bacteria. The supernatant was aspirated, the pellet was resuspended in kit buffer using pulse vortexing, and the suspension was lysed using the SDS containing kit lysis buffer. The mixture was vigorously pulse vortexed for 10s to free genomic DNA. Neutralizing buffer was added and the lysate was again pulse vortexed. The mixture was applied onto Nucleospin Tissue XS DNA spin columns (Macherey-Nagel, Bethlehem, PA, USA) and centrifuged according to the manufacturer’s instructions. Pellets were washed with kit buffer, and samples were dissolved in 30 µl TrisHCL-buffer (pH=8) without EDTA, brought up to 70 µl with PCR grade water (Fermentas), and used directly in triplicate in the PCR assay. In parallel, free bacterial DNA was isolated from 200 µl of plasma using the Qiagen Blood Mini Kit (Qiagen, Valencia, CA, USA) following the manufacturer’s instructions. No carrier DNA was added.

For PCR detection of bacteria in plasma samples, we used universal primers for 16s RNA: p201 and p1370 (Table 1) obtained following a two-step HPLC purification from Eurofins MWG Operon (Huntsville, AL, USA). We used Invitrogen SYBR greenER premix reagents (Life Technologies) and pre-treated the mastermix minus primer with PCR grade DNAse I (Life Technologies) as described (Tseng et al., 2003). All samples were run in triplicate. E. coli DNA standards (0.25, 0.5, 1 and 2 pg) were included on each plate. DNAse I pre-treatment enhanced the sensitivity of the assay but did not eliminate all background. A consistent lower detection limit of 0.125 pg per reaction, or approximately 160 copies of E.coli 16s rDNA was achieved based on calculations reported elsewhere (Farrelly et al., 1995).

Table 1.

Real-time PCR primers.

Target	Forward primer 5’ to 3’	Reverse Primer 5’ to 3’	Amplicon size (bp)	Accession number	Reference
IFN-γ	GCAACAAAAAGAAACGGGATGAC	CTGACTCCTTTTTCGCTTCC	148	NM_001032905	In house
MxA	AGGAGTTGCCCTTCCCAGA	TCGTTCACAAGTTTCTTCAGTTTCA	78	AF135187.1	Dr. Fenizia
TNF-α	AGCCCATGTTGTAGCAAACC	GCTGGTTATCTGTCAGCTCCA	104	DQ902483	In house
CCL-3 (MIP-1α)	CCTCCTGCTGCTTCAGCTAC	CTCCTTACTGGGGTCAGCAC	146	AF457195	In house
CCL-4 (MIP-1β)	CTTCCTCGCAACTTTGTGGT	GCTTGCTTCTTTTGGTTTGG	88	NM_002984	In house
CCL-5 (RANTES)	AGTGGCAAGTGCTCCAACC	CGAACCCATTTCTTCTCTGG	86	DQ913730	(Hofmann-Lehmann et al., 2002)
CXCL8 (IL-8)	GAGTGGACCACACTGTGCCA	AAACTTCTCCACAACCCTCTGC	108	NM_001032965	(Hardstedt et al., 2005)
IL-10	AGAACCACGACCCAGACATC	GGCCTTGCTCTTGTTTTCAC	119	DQ890063	In house
18s	GCCCGAAGCGTTTACTTTGA	TCCATTATTCCTAGCTGCGGTATC	81	NR_003286	(Medeiros et al., 2003)
p201	GAGGAAGGIGIGGAIGACGT				Tseng et al., 2003
p1370		AGICCCGIGAACGTATTCAC			Tseng et al., 2003
CD14	GGTTCCTGCTCAGCTACTGG	TGGTGCCGGTTATCTCTAGG	95	XP_001087125.1	In house

Accession numbers were obtained from http://www.ncbi.nlm.nih.gov/nuccore.

Cytokine/chemokine and CD14 real time PCR

RNA was isolated from week 48 (pre), 56 and 70 cryopreserved PBMC using the Qiagen RNeasy Plus mini kit according to the manufacturer’s directions, followed by the immediate generation of cDNA using the Qiagen QuantiTect kit with an extension time of 1 hour at 42°C. Primers (Table 1) except for CD14 were designed to be exon spanning using the UCSC Genome Browser (http://genome.ucsc.edu/). Gene expression levels were normalized against 18s RNA as reference gene. For the calculation of expression levels we used the efficiency corrected (Pfaffl) equation (Pfaffl, 2001) except for CD14 and the interferon-α/β induced gene MxA (myxovirus resistance A gene). Samples were run in triplicate in a 25 µl reaction using Invitrogen SYBR greenER premix with ROX (Life Technologies) on an Applied Biosytems ABI7000 cycler (Life Technologies) under the following conditions: 2 min 50°C, 10 min 95°C and 40 cycles of 30s 95°C, 15s 59°C and 30s 72.5°C followed by standard melting curve analysis.

Multiplex bead assays (Luminex xMAP)

Previously frozen undiluted plasma samples from the original study (Demberg et al., 2007) were evaluated for the expression of cytokines and chemokines at four different time points: before challenge (wk48), at peak viremia (wk52), close to viral set point (wk61) and during the chronic phase (wk70). The non-human primate specific Invitrogen 5-plex chemokine kit (IL-8, MCP-1, MIP-1α, MIP-1β and RANTES) and the 5-plex cytokine kit (IFN-γ, IL-2, IL-4, IL-10 and TNF-α) (both Life Technologies) were used according to the manufacturer’s instructions. Duplicate samples were read on a Luminex 100 machine (Luminex Corporation, Austin, TX, USA).

Flow Cytometry

PBMC were cultured as stated above and transferred into 5 ml flow tubes, washed once with DPBS and incubated with the viability dye Aqua (Life Technologies) to discriminate between live and dead cells (done on 6 animals at all time points). After washing, cells were further stained for 25 min at RT while protected from light with a cocktail of the following surface antibodies: CD3, CD4, CD8, CD20, CD27, CD28, CD95 and CD196 (CCR6) (Table 2). Cells were washed in DPBS and fixed with 150 µl of BD Cytofix/Cytoperm solution (BD Bioscience) for 15 min at RT in the dark. After fixation, cells were washed in BD Perm/Wash solution and stained intracellularly for 25 min at RT in the dark with a cocktail of the following antibodies: IFN-γ, IL-17, Ki-67 or Isotype control (Table 2). Cells were washed again with BD Perm/Wash solution and resuspended in 150 µl of DPBS with 2% Formaldehyde (Tousimis, Rockville, MD, USA). A minimum of 250,000 events in the lymphocytic gate were acquired on the same day on a custom 4 laser LSR II cytometer (BD Biosciences) using DIVA 6.2 software. Only samples with at least 100 events in the central and effector memory compartments were included in the analyses. Results were analyzed in FlowJo version 9.0.2 for Macintosh (Tree Star, Inc., Ashland, OR, USA) and graphs were generated in GraphPad Prism 5.0.3 for PC (GraphPad Software, Inc., La Jolla, CA, USA).

Table 2.

Antibodies used in Flow Cytometry

Antigen and color	Clone	Supplier
KI-67 FITC	B56	BD Biosciences
IL-17 PE	eBio64CAP17	eBioscience
CD28 Pe-Cy5	CD28.2	BD Biosciences
CD196 (CCR6) PerCP-Cy5.5	11A9	BD Biosciences
CD95 Pe-Cy7	DX2	eBioscience
CD3 Pacific Blue	SP34-2	BD Biosciences
CD4 Qdot605	L200	Invitrogen (Custom conjugate)
CD8 eFlour650	RPA-T8	eBioscience
IFN-γ APC	B27	BD Biosciences
CD27 Alexa700	0323	eBioscience
CD20 APC-Cy7	2H7	Biolegend

Statistical analysis

Differences in viral loads over time between control and vaccinated macaques were tested by the Wilcoxon rank sum test. Changes in IL-17⁺ and CCR6⁺ expressing cells over time and differences between the groups in the expression of Ki-67 were analyzed on square root or cube root-transformed values respectively, using repeated measures ANOVA corrected for multiple comparisons. Differences in viral loads at single time points and in LPS levels were evaluated by the exact Wilcoxon rank sum test. PCR data were assessed by ANOVA corrected for multiple comparisons on log-transformed data.

Results

Strong control of viremia in vaccinees does not prevent gradual loss of Th17 cells

We investigated the preservation of IL-17-producing cells among CD4⁺ and CD8⁺ memory T cells in peripheral blood in vaccinated and control animals. As previously published, following intravenous SHIV_89.6P challenge the vaccinated macaques exhibited a significant 4-log decrease in chronic viremia compared to controls and maintained peripheral blood CD4⁺ T-cells (Demberg et al., 2007). The addition of supplemental controls (Patterson et al., 2008) did not change this relationship appreciably. Macaques in the combined control group were unable to suppress viral replication and showed a significantly higher viral load of about 3.5 logs compared to the vaccinated animals (p<0.0001, Fig. 1A). In addition, CD4⁺ T-cells were rapidly depleted (Fig. 1B) after a short initial proliferative burst during the early phase of infection as described elsewhere (Kaur et al., 2000). For analysis of IL-17-producing cells, Th17 and Tc17 cells were gated as shown in Fig. 2. Th17 cells have a memory phenotype (Brenchley et al., 2008; Kader et al., 2009b). In our hands, at the pre-challenge time point the majority of Th17 cells in all the macaques displayed a significantly higher central memory (CM) phenotype (CD28⁺CD95⁺) than effector memory (EM) phenotype (CD28-CD95⁺) (data not shown). This observation is in accord with the distribution of memory populations among CD4 positive cells in healthy animals, where CM cells account for approximately 60% and EM cells only 9% of the CD4 population. We therefore define Th17 cells as CD3⁺CD4⁺ cells with a CM phenotype that are IL-17 positive. Pre-challenge, the total population of Th17 cells in all macaques ranged from 6% to 39%, but by 4 weeks post-challenge the cells were present at very low frequency in the control animals due to the rapid overall loss of CD4⁺ T-cells (Fig. 1B). Further, a loss of CD4 CM cells in the control macaques post-challenge, with a concomitant rise in CD4 EM cells, is illustrated in Table 3. In contrast, the CD4 CM population remained stable in the vaccinated macaques. By 24 weeks post-challenge, only 3 of 9 control macaques had sufficient cells in the CD4 CM population to allow analysis of Th17 cells. Therefore, post-challenge dynamics of Th17 cells were investigated in the vaccinated group only. Here despite control of viremia to very low levels in the chronic phase of infection (Fig. 1A) and maintenance of close to pre-challenge CD4 counts (after initial rapid expansion and contraction of CD4 cells, Fig. 1B), we observed a significant decline in the population of Th17 cells in comparison to pre-infection levels (pre vs chronic and wk 4 vs chronic: p= 0.027 for both; Fig 3A). In addition we monitored the expression of CCR6 (a homing marker for inflamed tissue) over time on these cells. At the pre-challenge time point, approximately 33% of IL-17⁺ cells in the vaccinated macaques expressed the CCR6 receptor on the cell surface. Over the course of infection, the remaining IL-17⁺ cells significantly upregulated the expression of the CCR6 receptor. This was seen by wk4 post challenge and was maintained in the chronic infection phase (pre vs wk4 and vs chronic: p = 0.0003 for both; Fig. 3B).

Fig. 1. Viral load and CD4 count after SHIV_89.6P challenge.

(A) Geometric mean viral loads post challenge. A 4-log significant difference in median chronic viremia between control and vaccinated macaques over weeks 6 to 24 was previously reported [37]. With 6 additional controls, the 3.5 log difference remained significant (p<0.0001) (B) Mean CD4⁺ T-cell counts in blood over the course of the study. The dashed line indicates 100 cells/µl. The solid line marks the restoration of CD4 T cells in vaccinated macaques to pre-challenge levels. Post challenge, CD4⁺ T-cell counts declined to <10 cells/µl in controls. Error bar = standard error of the mean (sem).

Fig. 2. Flow Cytometry gating strategy.

On the left hand side, singlets were further gated on the lymphocytic population followed by a live/dead gate and selection of CD3⁺ cells. CD3⁺ cells were gated on CD4 or CD8 with further gating for central memory (CM), effector memory (EM), and total memory (TM) using CD28 and CD95. As an example, CD4 CM cells and CD8 TM cells were gated as shown for IL-17 secretion.

Table 3.

CD4 central and effector memory cells in vaccinated and control macaques over time post-challenge.

	% Positive Cells ± sem
Vaccinated Macaques	Pre-challenge	Wk 4 post-challenge	Wk 24 post-challenge
CD4CM	53.8 ± 3.5	50.2 ± 5.9	56.8 ± 3.0
CD4EM	9.2 ± 3.9	20.8 ± 8.5	12.7 ± 3.4
Control Macaques
CD4CM	65.2 ± 4.1	37.3 ± 6.9	25.7 ± 5.7
CD4EM	8.5 ± 3.4	55.5 ± 8.7	69.0 ± 7.9

Fig. 3. Loss of Th17 cells and up-regulation of CCR6 on Th17 cells during the course of infection in the vaccinated macaques.

(A) Successive decline in the total population of Th17 cells (pre vs wk 4 and pre vs chronic, p = 0.027 for both) in vaccinated, protected macaques. (B) Up-regulation of CCR6 during the course of SHIV_89.6P infection. Significantly higher expression was seen in the total Th17 population (pre vs wk4 and pre vs chronic, p = 0.0003 for both). Plotted are mean values with sem.

Tc17 cells showed a trend towards depletion from PBMC over time in control animals, but were maintained in vaccinees controlling viremia

During SIV infection the balance between Th17 and Tc17 is altered in peripheral blood (Kader et al., 2009a; Nigam et al., 2011). Here we investigated the dynamics of the Tc17 cell population longitudinally in the vaccinated and control macaques. In contrast to Th17 cells which are mainly CM T-cells, Tc17 cells were more evenly divided between CM and EM phenotypes (data not shown). Therefore we report here data on total CD8 memory cells. Percentages of CCR6 expressing Tc17 cells were similar within control and vaccinated groups pre-challenge and at 4 weeks post-challenge. However, with disease progression Tc17 cells expressing CCR6 increased significantly compared to pre and wk4 levels (p = 0.0001 for both) but only among control animals (Fig. 4A). In comparison to the control animals, vaccinated macaques expressed slightly but not significantly higher levels of CCR6⁺ Tc17 cells at the pre and wk4 post-challenge time points. CCR6⁺ Tc17 cells in vaccinated animals were lower at the chronic phase time point when compared to controls, and overall did not increase over the course of infection (Fig. 4A). In comparison to the CCR6⁺ Th17 cells in the vaccinated group which increased over the course of infection (Fig. 3B), the CCR6⁺ Tc17 cells appeared to be more stable.

Fig. 4. CCR6 expression on Tc17 cells and population dynamics over the course of infection.

(A) Mean levels of CCR6 expression on Tc17 cells from the control animals (black bars) and vaccinated animals (open bars). Significantly increased expression on total memory cells (pre vs chronic and wk4 vs chronic, p = 0.0001 for both) was observed for control animals. (B) Tc17 memory population dynamics over the course of infection in control animals (black bars) and vaccinated animals (open bars). Error bar = sem.

The Tc17 population (total memory) varied from roughly 1% to 17% among all animals at the pre-challenge time point. Interestingly, the total Tc17 population tended to decline over time post-challenge in the control animals (Fig. 4B), but this effect did not reach significance. A similar decline in the Tc17 population was recently reported in unvaccinated, SIV infected macaques and was also observed in rectal tissue, supporting the trend observed here (Nigam et al., 2011). A similar trend was not seen in the vaccinated animals (Fig. 4B).

Despite rapid depletion of CD4⁺ cells in control macaques, enhanced LPS plasma levels in the chronic phase were not detected

As CD4 cells and thus also Th17 cells were depleted in control macaques following viral challenge, we assumed this loss might be associated with enhanced levels of lipopolysaccharides in plasma due to enhanced microbial translocation. This assumption was supported by the even more profound depletion of Th17 and Tc17 cells known to occur from the gut mucosa following pathogenic SIV infection (Nigam et al., 2011). We detected LPS in all plasma samples prior to challenge (Fig. 5). By chance, the control animals had significantly higher mean levels of LPS in plasma both prior to challenge (654 pg/ml) and at the chronic phase time point (622 pg/ml) compared to the vaccinees (345 pg/ml and 419 pg/ml, respectively; p = 0.015 and 0.036, respectively) (Fig. 5). However, post-challenge an expected increase in plasma LPS levels for the control animals was not seen, despite the rapid decline in total CD4 (and therefore Th17) cells and uncontrolled viremia (Fig. 1A, B). LPS levels remained similar for both groups in the chronic phase (wk24 post-challenge) compared to pre-challenge values in spite of the significant differences in challenge outcome (Fig. 5).

Fig. 5. Lipopolysaccharide (LPS) levels in plasma, measured by LAL assay.

Shown are LPS levels for paired samples before and after challenge (wk24, chronic phase).

To validate the lack of elevated LPS levels, we further tested the original study animals (Demberg et al., 2007) for free and pelleted bacterial DNA in plasma at the same time points. The PCR data confirmed our previous observations using the LAL assay; all samples evaluated for 16s rDNA by real time PCR were negative (data not shown).

To further validate the findings from the LAL and 16s rDNA PCR assays, we tested the same animals for CD14 mRNA by real-time PCR using total mRNA from PBMCs at wk48, wk56 and wk70. Increased levels of CD14 mRNA might suggest enhanced activation of monocytes and macrophages due to increased levels of bacteria or bacterial endotoxins (LPS) in the blood resulting from microbial translocation and/or mucosal barrier damage. The results were in concert with the LAL and 16s PCR data. CD14 mRNA levels were not different at wk56 post-challenge compared to pre-challenge values, either in the control group (0.8 fold change) or in the vaccinated group (non-significant 1.4 fold change). At wk70 CD14 mRNA levels further increased in the vaccinated group, however, the moderate 2.2 fold change was still not significantly different compared to pre-challenge (wk48) levels (data not shown). Similarly, the control group showed no elevation in CD14 mRNA levels at wk70 compared to pre-challenge values (1.2 fold change).

Chronic immune activation was associated with increased viremia in the absence of microbial translocation

Chronic immune activation in HIV infected individuals has been associated with bacterial DNA plasma levels (Jiang et al., 2009). In spite of the lack of increase in LPS levels with time of infection, we nevertheless looked at chronic immune activation by Ki-67 and IFN-γ staining in all the control and vaccinated macaques pre-challenge, 4 weeks post-challenge and at wk24 post-challenge during the chronic phase. Overall IFN-γ levels were low (data not shown). We detected significantly elevated immune activation in the chronic phase of infection in control macaques, as shown by increased percentages of Ki-67⁺ CD8⁺ CM (p = 0.0043) and EM cells (p= 0.0006) at wk24 post challenge and CD4⁺ CM cells at wk4 and wk24 post-challenge (p = 0.014 and p<0.0001), compared to values in vaccinated animals in which activation increased by wk4 post challenge and dropped during the chronic phase back to almost pre challenge levels (Fig. 6A–C). In contrast, CD4⁺ EM cells were not elevated in control macaques at any time point (Fig. 6D). The increase in Ki-67⁺ activated cells was in concert with the increased viremia both at wk 4 and wk 24 post-challenge in the control macaques compared to the vaccinated macaques (p <0.0001 for both; Fig. 6E). Overall, the control animals exhibited a sustained high level of viremia throughout the chronic phase of infection (Fig. 1A).

Fig. 6. Chronic immune activation among CD8 and CD4 memory populations in control and vaccinated macaques.

(A) Mean Ki-67 expression among CD8⁺ CM and (B) EM cells pre- and post-challenge (wk4 and wk24). Ki-67⁺ cells were elevated in controls compared to vaccinated macaques during chronic infection (wk24) for both CD8⁺ CM (p = 0.0043) and EM (p = 0.0006) cells. (C) Mean Ki-67 expression among CD4⁺ CM and (D) EM cells pre- and post-challenge (wk4 and wk24). Ki-67⁺ cells were elevated in controls compared to vaccinated macaques at wk4 and wk24 for CD4⁺ CM cells (p = 0.014 and p<0.0001, respectively) (C), whereas no differences were observed for EM cells (D). (E) Geometric mean viral loads illustrating increased viral burdens in control compared to vaccinated macaques post-challenge at wk4 and wk24 (p<0.0001 for both).

Elevated plasma IFN-γ and elevated MxA mRNA are indicative of chronic inflammation in control macaques

In addition to cellular markers of activation, we tried to identify a cytokine/chemokine activation signature in plasma at the protein level in the animals from the original vaccine study (3 controls, 8 vaccinated) (Demberg et al., 2007), examining plasma from both groups at pre- and post-challenge time points (pre, wk2, wk11, wk20) using the Luminex bead array. Surprisingly, the cytokines IL-2, IL-4, IL-10 and TNF-α were below the lower limit of detection in both groups. However, we observed a transient non-significant increase in the levels of five additional cytokines/chemokines (Rantes, MIP-1α, MIP-1β, MCP-1, and IL-8) during acute infection (wk2). Only IFN-γ was significantly elevated in control macaques at wk11 and wk20 compared to the vaccinated animals (p = 0.036 and 0.0036, respectively; Fig. 7A). This increase in plasma IFN-γ was marginally but not significantly correlated with the corresponding viral loads (p = 0.052; data not shown).

Fig. 7. Plasma IFN-γ levels post-challenge normalized to pre-challenge values and real time PCR analysis of MxA and IL-8 mRNA.

(A) Plasma levels of IFN-γ as indicated over the course of infection. The mean of duplicate assays are plotted. The dotted line indicates the time of challenge. IFN-γ levels in control macaques compared to the vaccinated group were elevated at wk11 and wk20 post-challenge (p = 0.036 and 0.0036, respectively). (B) Mean fold increases of MxA and IL-8 mRNA at wk6 and wk20 post-challenge compared to pre values for the vaccinated and control groups are shown. Significant elevations in MxA mRNA are indicated as follows: A: p = 0.047; B: p = 0.0026; C: p = 0.0021. No significant differences between levels in control and vaccinated macaques were observed. Significant elevations in IL-8 mRNA are indicated as follows: D: p<0.0001; E: p = 0.047. IL-8 mRNA dropped significantly in the vaccinated and control groups between wk6 and wk20 (p = 0.0005 and p = 0.048, respectively). Error bar = sem.

To further explore cytokine/chemokine expression, we conducted real-time PCR experiments using cDNA derived from total mRNA isolated from PBMC at pre, wk6 and wk20 post-challenge timepoints, assaying MxA, TNF-α, IFN-γ, MIP-1α, MIP1β, Rantes, IL-8 and IL-10. The PCR results differed somewhat from the protein expression data. Compared to pre-challenge levels, only mRNA of MxA, one of several IFN-α/β induced genes, was significantly upregulated post-challenge in controls at wk6 and wk20 (p = 0.0026, p = 0.0021, respectively) and in the vaccinated group at wk20 only (p = 0.047) (Fig. 7B). In addition to MxA, IL-8 expression post-challenge was transiently elevated in both control and vaccinated groups compared to pre-challenge values at wk6 (p<0.0001 and p = 0.047 for vaccinated and controls, respectively), but declined by wk20 (Fig. 7B). The decreases in IL-8 expression between wk6 and wk20 in the vaccinated and control groups were statistically significant (p = 0.0005 and p = 0.048, respectively; Fig. 7B).

Discussion

In this study we addressed the hypothesis that vaccinated macaques, partially protected against a SHIV_89.6P challenge and showing strong control of chronic viremia, although not exhibiting sterilizing immunity, would exhibit enhanced preservation of Th17 and Tc17 cells and decreased microbial translocation compared to unprotected control animals. Such an outcome would suggest that partially protective vaccines might have a potential benefit by reducing immune activation, potentially preventing opportunistic bacterial infections and slowing disease progression. In addition to monocytes/macrophages, Th17 cells play a role in control of bacterial infections (Crome, Wang, and Levings, 2010; Cua and Tato, 2010; Damsker, Hansen, and Caspi, 2010). Here, for the first time to our knowledge, they were investigated longitudinally in a preclinical HIV vaccine study in non-human primates that achieved a highly protective outcome. We observed a very rapid decline in peripheral blood CD4 cells in unvaccinated control animals by 4 weeks post-challenge (Fig. 1B). In vaccinated macaques which exhibited strong control of viremia and maintained CD4 counts over time (Fig. 1A,B) we nevertheless observed a significant loss of Th17 cells by 24 weeks post-challenge (Fig. 3A). Th17 cells are targets for HIV/SIV replication (El Hed et al., 2010; Kader et al., 2009b; Klatt and Brenchley, 2010) and can be depleted during infection. Th17 cells express CXCR4 under inflammatory conditions and express CCR5 constitutively (Lim et al., 2008; Miller Sanders, Cruse, and Lewis, 2010). Both co-receptors are used by the dual-tropic SHIV_89.6P challenge virus, perhaps contributing to the loss of the Th17 population even in conjunction with low levels of viremia.

In contrast to the loss of Th17 cells seen here in SHIV infected macaques; Maek et al. (2007) reported that HIV positive patients show an increase in both Th17 and CD3⁺CD4⁻IL-17⁺ populations compared to healthy volunteers. Further, Ndhlovu et al. (2008) reported that suppression or control of viral replication below 50 copies/ml preserves IL-17 producing cells in HIV infected children, whereas children with higher viral loads showed a significant decline in these cells. By 24 weeks post-challenge, in 4 animals with viral loads of <50 copies/ml and a fifth with <100 copies/ml, we observed a decline in Th17 cells compared to pre-challenge levels (data not shown), contradicting these findings. Further evidence contradicting the preservation of Th17 cells during infection was reported by Kader et al. (2009a) and Nigam et al. (2011) who showed that HAART treatment of SIV infected rhesus monkeys cannot restore the Th17 population to pre-infection levels.

Loss of Th17 cells during HAART might be due either to ongoing low level infection despite control of plasma viremia, or to homing of the cells to inflamed tissues. Together with the decline of Th17 cells over time in the protected macaques we observed an increase in CCR6 receptor expression (Fig. 3B), indicating potential homing to inflammatory sites (Potzl et al., 2008; Yamazaki et al., 2008). CCR6, a hallmark of human Th17 cells, may serve as an activation marker on rhesus macaque Th17 cells, and expression levels may change with homing of the cells to target tissues. Unfortunately due to limitations of PBMC samples and the flow cytometry staining panel, we were unable to investigate expression of the gut mucosal homing marker α4β7 which is expressed on a large proportion of Th17 cells (Kader et al., 2009b; Nigam et al., 2011). In fact cells that express CCR6 also express the gut homing markers α4β7 or αEβ7 to some extent (Nigam et al., 2011). It would be of interest to investigate whether these gut homing markers also increase in parallel with the higher expression of CCR6. Homing in the presence of low-level, persistent infection could explain in part the loss of these cells from the circulation over time as previously proposed (Klatt and Brenchley, 2010). In fact, Ciccone et al. (2011) recently reported higher levels of Th17 cells in the colon compared to the peripheral blood of HIV-infected long-term non-progressors and those on long-term ART therapy.

Here we also report for the first time investigation of Tc17 cells overtime in a highly effective preclinical vaccine study. Only a few reports have appeared on Tc17 cells with regard to SIV/HIV infection (Kader et al., 2009a; Maek et al., 2007; Ndhlovu et al., 2008; Nigam et al., 2011). Interestingly, the upregulation of CCR6 expression on Tc17 cells seems to be similar to that of Th17 cells. The downward trend of these cells over time in the unprotected control group but not in the vaccinated group might be due to enhanced migration of this population to inflamed tissue sites, potentially following the “footsteps” of Th17 cells (Fig. 4B). The significantly higher expression in control macaques of CCR6 at wk74 compared to the pre and wk4 time points (Fig. 4A) supports this assumption. Currently the role of Tc17 cells in vivo is unclear. In cancer patients Tc17 cells cluster around the edges of tumors, secrete high amounts of inflammatory cytokines, and express CCR6 and an effector phenotype, but exhibit limited cytotoxic capacity (Kuang et al., 2010; Nigam et al., 2011). These findings are in accord with our results showing that the cells have in part an effector phenotype and express CCR6, indicative of greater activation. Overall the function and contribution of Tc17 cells in SIV and HIV infection remain elusive and warrant further investigation.

Plasma LPS levels are routinely used as an indication of microbial translocation. Here, no increase in LPS levels was observed post-challenge during the chronic phase in either control or vaccinated macaques using a sensitive LAL assay. Further, bacterial DNA was not detected, and CD14 mRNA levels were not increased post-challenge. Overall microbial translocation might be slow in onset and require a sustained period of chronic infection rather than the rapid CD4 depletion and disease course seen in the SHIV model. This suggestion is supported by the recent report of Redd et al. (2011) which proposed that increased microbial translocation and elevated LPS levels are a consequence of advanced HIV disease and AIDS rather than a cause. Numerous discrepancies in the literature regarding microbial translocation and HIV-1 disease progression have been reported (Redd et al., 2011). Similar discrepancies have occurred in the SIV literature as well. LPS levels were not predictive of SIV disease progression in rhesus macaques (Leinert et al., 2010) whereas high levels of LPS prior to SIV infection were associated with faster disease progression in pigtailed macaques (Klatt et al., 2010). In some aspects, the course of SHIV_89.6P infection seen here in control animals resembles SIV infection of natural hosts, where high viremia occurs in the absence of microbial translocation (Brenchley and Douek, 2008b; Pandrea et al., 2007). However, in contrast to our results, rhesus monkeys challenged with non-pathogenic SIV_agm display a transient drop in CD4 counts but no preferential drop in mucosal Th17 cells occurs (Paiardini, 2010). Further, although increased apoptosis and immune activation are seen at mucosal sites in the rhesus/SIV_agm model, only a transient, non-significant increase in LPS is seen before levels drop below pre-challenge values with control of viremia (Pandrea et al., 2007).

Several mechanisms might contribute to control of LPS during SHIV infection. During early stages of HIV infection, endotoxin core antibodies (EndoCab) can help control LPS blood levels (Brenchley et al., 2006) but may also interfere with the detection of LPS by the LAL assay (Bennett-Guerrero et al., 2001). The expansion of EndoCab antibody (IgG) responses has been shown in live attenuated Shigella vaccine trials (Simon et al., 2009) and in cystic fibrosis patients after infection with Pseudomonas cepacia (Nelson et al., 1993). Enhanced EndoCab antibodies have been reported in plasma of an HIV-infected African cohort and might have contributed to “protection” from increased LPS levels (Redd et al., 2009). Moreover, it was recently shown that plasma LPS levels correlated inversely with serum anti-LPS antibodies in untreated HIV patients, suggesting a possible role of the antibody in decreasing LPS levels (Lim et al., 2011). We did not measure EndoCab antibodies due to limited cross-reactivity of available reagents (only IgM reactive) with rhesus macaque proteins (Leinert et al., 2010). The role and influence of anti-LPS antibodies during HIV/SIV infection should be investigated in future studies.

While elevated LPS plasma levels were not observed in the SHIV_89.6P model, we did observe chronic immune activation in CD8⁺ CM and EM, and CD4⁺ CM cells, evidenced by Ki-67⁺ cells, in association with high viremia in the control macaques, confirming previous reports that activation is correlated with viral load and disease progression (Leligdowicz et al., 2010; Redd et al., 2010; Tuaillon et al., 2009). In contrast to our findings (Fig. 6D), others have reported strong Ki-67 expression in the CD4⁺ EM compartment following CD8⁺ T cell depletion in the SHIV_89.6P model (Mueller et al., 2009), however, this might be attributed to proliferation of CD4⁺ T-cells following the depletion. In the SIV model, strong Ki-67 expression has been seen on CD4⁺ CM cells (Okoye et al., 2007), similar to our findings, but also on CD4⁺ EM cells. The latter observation follows distinction of “transitional” EM and EM cells as measured by CCR7 and CCR5. Without this further division of memory subpopulations, almost all the reported Ki-67⁺ cells would have been phenotyped as CM cells, in agreement with our findings (Fig. 6C). Ki-67 has been correlated with expression of HLA-DR and CD38 in HIV infected individuals (Chattopadhyay and Roederer, 2010). Moreover, as markers of chronic immune activation, CD38 and HLA-DR have been recently reported to correlate well in monkeys (Manrique et al., 2011). The inclusion of these markers in future studies might provide a more detailed picture regarding chronic immune activation.

To examine immune activation in greater depth, we sought to define an activation profile based on cytokines/chemokines in plasma. In general strong IFN-γ responses are associated with control of viral infection. A recent report showed that higher IFN-γ levels during acute HIV infection were associated with lower viral loads 12 months later at set point (Roberts et al., 2010). Here, plasma IFN-γ was slightly (but not significantly) elevated in the vaccinated group of macaques compared to controls during acute infection (Fig. 7A), most likely reflecting recall of adaptive immunity, whereas IFN-γ levels rose rapidly thereafter in controls in response to continuing high viremia.

Our investigations of gene transcription by PCR revealed significant drops in IL-8 mRNA in the vaccinated and control groups at wk20 post-challenge (Fig. 7B). Overall, however, total IL-8 mRNA remained slightly higher in the vaccinated group (Fig. 7B). This might indicate that Th17 or Tc17 cells migrated to mucosal sites. IL-8 is an IL-17-regulated gene (Raffatellu et al., 2008), therefore higher IL-8 mRNA levels in vaccinated animals during the chronic phase might be explicable by the slower decline in Th17 cells and stable Tc17 population. In the control macaques the lower IL-8 mRNA levels in PBMC might be due to diminished gene activation attributed to loss or migration of Th17 cells and the potentially limited compensatory capacity of Tc17 cells which tended to decline or migrate into tissues (strong CCR6 upregulation) (Fig. 4B).

The type I interferons, IFN-α/β, are mainly produced by plasmacytoid dendritic cells (pDC) through activation by TLR9 and TLR7 (Fitzgerald-Bocarsly and Jacobs, 2010; Swiecki and Colonna, 2010). During in vitro HIV/SIV infection, pDCs secrete high amounts of IFN-α (Harris et al., 2010; Swiecki and Colonna, 2010) which activates the MxA gene (Haller, Staeheli, and Kochs, 2009). MxA is a well defined surrogate marker for the IFN-α family which exhibits high diversity in humans (Nyman et al., 1998) and rhesus macaques (Easlick et al., 2010). In early phase of chronic SIV infection the down regulation of an initial robust IFN-α response distinguishes nonpathogenic infection of the natural host from pathogenic infection of rhesus macaques (Harris et al., 2010). Here, using activation of the MxA gene as a surrogate marker IFN-α, we asked whether high levels of IFN-α are similarly associated with disease progression and low levels with control of viremia. MxA levels were elevated rapidly in control macaques with significantly enhanced expression levels seen at wk6 and wk20 post-challenge (Fig. 7B), whereas MxA expression in the vaccinated group was not significantly elevated until wk20. Stronger induction was seen in the control group, as expected for a pathogenic infection (Harris et al., 2010). However, the increased MxA expression in the vaccinated group by week 20 suggests eventual disease progression.

The lack of correlation observed between plasma cytokine protein levels and mRNA levels can be attributed to the different samples used for analysis. The cytokine/chemokine levels in plasma represent a pool to which many cell types from multiple tissues contributed. In contrast mRNA levels measured by real-time PCR were obtained on purified PBMC, thus representing only a fraction of potential cell contributors.

The dual-tropic SHIV_89.6P is able to infect and deplete not only memory CD4 cells but also naïve cells in the rhesus macaque model (Zhang et al., 2000), leading to more rapid depletion of CD4⁺ T-cells compared to the SIV model. Thus the SHIV infected animals in general progress faster to AIDS. In this challenge model we show that a partially protective vaccine was unable to prevent the loss of Th17 over time, despite preservation of CD4 cells potent control of viremia. However no loss of Tc17 cells over the course of infection was seen in vaccinated animals, as has been reported for unvaccinated, SIV-infected animals (Nigam et al., 2011), similar to the trend observed here for the control group. Higher levels of immune activation as measured by Ki-67 on CD4 and CD8 cells, plasma IFN-γ and MxA mRNA levels were observed in the control group in accordance with uncontrolled viremia in these animals. Interestingly elevated plasma LPS levels were not observed in either group during SHIV_89.6P infection calling for further investigation of microbial translocation in the mucosa and the role of Th17 and Tc17 cells. Overall, despite the control of viremia to the threshold of detection in the strongly protected vaccinated macaques, we detected a loss of Th17 cells from the PBMC. In control animals, in addition to the severe depletion of CD4 cells from the blood, a slow loss of Tc17 cells was seen over time potentially indicating that Tc17 and Th17 cells home to the gut mucosal tissue, where virus persists and where damage is most profound. Due to the classic role of Th17 cells in bacterial infection and the emerging role of Tc17 cells, it is likely that both cell types home to the inflamed tissue sites and engage to control bacterial, fungal and viral infections. Unfortunately no fresh tissue samples were available from these animals for cellular analyses nor were frozen tissues for immunohistochemical staining for Th17 and Tc17 cells and bacteria in the GI tract. Questions to address in the future include design of strategies to better preserve these cells, characterization of their role in the gut mucosa, and methods to harness their functions in vaccine approaches to prevent HIV induced epithelial damage and microbial translocation and to fight disease progression.