The 18 Indian rhesus macaques (RMs; Macaca mulatta) whose peripheral blood mononuclear cells (PBMCs) were analyzed in this study served as unvaccinated controls for our previous vaccination studies (Table 1) (30–33). All animals were housed at the Wisconsin National Primate Research Center (WNPRC) and cared for in compliance with the guidelines of the Weatherall report (34) and the National Research Council’s Guide for the Use and Care of Laboratory Animals (35). All procedures were performed in accordance with protocols approved by the University of Wisconsin Graduate School Animal Care and Use Committee. Detailed descriptions of caretaking procedures (i.e., housing, feeding, environmental enrichment, and medical care) and efforts to minimize animal suffering (i.e., anesthesia and euthanasia) can be found in the “Research Animals and Ethics Statement” sections in the Materials and Methods of our previously published studies (30–33). MHC class I genotyping was conducted by the WNPRC.

As described in our previous studies, RMs were subjected to repeated intrarectal challenges with SIVmac239, once every 2 weeks (30–33). Depending on the study, each challenge dose contained either 4.8 × 10⁵ (31–33) or 6.52 × 10⁶ viral RNA (vRNA) copies (30). Plasma viral loads were quantified at 7 and 10 days after each challenge, and animals were no longer challenged if a detectable viral load measurement (>15 vRNA copies/mL plasma) was obtained. In one of our studies, animals were subjected to escalating dose challenges if they remained uninfected after the first eight challenges (30). For all other studies, the same dose was used for all challenges. Viral loads were determined by extracting total RNA from EDTA-anticoagulated plasma and using this RNA as template for a two-step reverse-transcription PCR with Gag-specific primers and a Gag-specific fluorescent probe, as previously described (30–33, 36). Viral loads for each animal are shown in Figure 1.

We conducted a retrospective analysis of RhCMV- and SIVmac239-specific CTL populations in our historical cohort of unvaccinated RMs. Animal inclusion criteria were as follows: (1) no history of vaccination against SIVmac239, (2) a MHC class I haplotype including the Mamu-A^∗01 allele, the Mamu-A^∗02 allele, or both, and (3) the presence of cryopreserved PBMC samples at an acute SIVmac239 infection timepoint, a chronic SIVmac239 infection timepoint, or both. Acute infection timepoints were defined as those 35–70 days (5–10 weeks) after the infecting challenge. Chronic infection timepoints were defined as those >70 days (>10 weeks) after the infecting challenge. PBMC timepoints for each animal are listed in Table 2. Despite the absence of any defined Mamu-A^∗01-restricted, RhCMV-derived CTL epitopes, we chose to include Mamu-A^∗01⁺ animals in our study, since Mamu-A^∗01-restricted CTLs specific for SIVmac239-derived peptides (i.e., Tat SL8 and Gag CM9) are some of the best-characterized SIVmac239-specific CTL populations (7, 13, 26, 27).

PBMCs were isolated from EDTA-anticoagulated blood by Ficoll-Paque Plus (GE Healthcare) density centrifugation and washed once in R10 medium [RPMI 1640 with GlutaMAX (Gibco), 10% heat-inactivated FBS (Gibco), antibacterial/antimycotic (Lonza)]. In some cases, red blood cells were lysed by resuspending PBMCs in ACK lysing buffer (Lonza or Gibco) and incubating for 5 min at room temperature. PBMCs were washed again in R10 medium (if ACK-treated), quantified, then pelleted by centrifugation and resuspended in cryopreservation solution (90% FBS, 10% DMSO; or 67.5% RPMI 1640, 22.5% FBS, 10% DMSO) at 4°C. PBMCs were cooled to −80°C, then transferred to liquid nitrogen freezers for long-term storage. For the immunophenotyping assays in this study (described below), cryopreserved PBMCs were thawed in a 37°C water bath, washed twice in R10 medium, and re-quantified. Approximately 1.5 × 10⁶ PBMCs were used per test.

PBMCs were stained with the appropriate BV421-conjugated pMHCI tetramers (NIH Tetramer Core Facility at Emory University, Atlanta, GA, United States) for 30 min at room temperature. PBMCs were then stained for an additional 30 min with a cocktail containing the following fluorophore-conjugated monoclonal antibodies (mAbs) and amine-reactive viability dye: anti-CCR7 FITC (clone 150503, BD Biosciences), anti-CD8a BV711 (clone RPA-T8, BioLegend), anti-CD28 BV785 (clone CD28.2, BioLegend), anti-CD14 APC (clone M5E2, BioLegend), anti-CD20 APC (clone 2H7, BioLegend), anti-CD159a APC (clone Z199, Beckman Coulter), anti-PD-1 PE-Cy7 (clone EH12.2H7, BioLegend), and Far Red live/dead dye (LIVE/DEAD fixable Far Red dead cell stain kit, Invitrogen). Cells were washed, then fixed and permeabilized using Cytofix/Cytoperm (BD Biosciences) and Perm/Wash Buffer (BD Biosciences), respectively. Cells were stained intracellularly with anti-CD3 PerCP-Cy5.5 (clone SP34-2, BD Biosciences), anti-granzyme B Alexa Fluor 700 (clone GB11, BD Biosciences), and anti-CTLA-4 PE-CF594 (clone BNI3, BD Biosciences). Following a final wash in Perm/Wash Buffer, samples were acquired on a special-order BD LSR II flow cytometer. Data was analyzed using FlowJo software (version 10.4); gating strategy is shown in Supplementary Figure 1. To assess the extent of SIVmac239-induced disease progression in our RMs, we calculated approximate CD4:CD8 T cell ratios using the aforementioned pMHCI tetramer staining data from resting PBMCs. CD4:CD8 T cell ratios were calculated using the following formula: (100 – % of CD8⁺ T cells among live CD3⁺ CD14^– CD20^– CD159a^– lymphocytes)/(% of CD8⁺ T cells among live CD3⁺ CD14^– CD20^– CD159a^– lymphocytes). This data is depicted in Supplementary Figure 2.

Peripheral blood mononuclear cells in R10 medium (1.5 × 10⁶ per tube) were incubated with anti-CD28 and anti-CD49d costimulatory mAbs (BD Biosciences, clones L293 and 9F10, respectively), anti-CD107a PE (clone H4A3, BioLegend), and minimal optimal peptide (final concentration of 2 μM) for 1 h at 37°C. Following addition of brefeldin A (BioLegend) and GolgiStop (BD Biosciences), samples were incubated for an additional 5 h at 37°C. Cells were washed once, then stained with the following cocktail of fluorophore-conjugated mAbs and amine-reactive viability dye: anti-CD8a BV785 (clone RPA-T8, BioLegend), anti-CD14 APC (clone M5E2, BioLegend), anti-CD20 APC (clone 2H7, BioLegend), anti-CD159a APC (clone Z199, Beckman Coulter), anti-PD-1 PE-Cy7 (clone EH12.2H7, BioLegend), and Far Red live/dead dye (LIVE/DEAD fixable Far Red dead cell stain kit, Invitrogen). Following a wash step, cells were fixed and permeabilized using Cytofix/Cytoperm (BD Biosciences) and Perm/Wash Buffer (BD Biosciences), respectively. Cells were then stained intracellularly with anti-CD3 PerCP-Cy5.5 (clone SP34-2, BD Biosciences), anti-TNF-α APC-Cy7 (clone MAb11, BioLegend), anti-IFN-γ BV421 (clone B27, BioLegend), and anti-CTLA-4 PE-CF594 (clone BNI3, BD Biosciences). Cells were washed once more in Perm/Wash Buffer, and samples were acquired on a special-order BD LSR II flow cytometer. Data was analyzed with FlowJo software (version 10.4); gating strategy is shown in Supplementary Figure 3. To evaluate T cell functionality, we created the following gates: CD107a alone (degranulation), a Boolean or gate to quantify overall response frequencies (CD107a or IFN-γ or TNF-α), and a Boolean and gate to assess polyfunctionality (CD107a and IFN-γ and TNF-α). For each of the above gates, we subtracted the corresponding frequency for the unstimulated negative control (Microsoft Excel). We also analyzed exhaustion marker (PD-1 and CTLA-4) expression in the Boolean or gate subpopulation. For degranulation, polyfunctionality, and exhaustion marker analyses, the Boolean or gate frequency for a stimulated sample had to be least two times greater than the or gate frequency for the unstimulated negative control to be included in our comparisons of multiple animals.

Comparisons of CTL populations were performed using Welch’s t-test (unpaired two-sample t-test assuming unequal variances; Microsoft Excel). Associations between CTL phenotypic characteristics and setpoint SIVmac239 viral loads were evaluated by Pearson correlation analysis (GraphPad Prism). p-values < 0.05 were considered statistically significant and are reported in tables, figures, or figure legends.

As a preliminary investigation into the suitability of RhCMV-specific CTLs for retargeting-based immunotherapeutic strategies, we conducted a retrospective analysis of 18 unvaccinated, SIVmac239-infected RMs to analyze and compare the phenotypic characteristics and effector functions of RhCMV- and SIVmac239-specific CTLs during SIVmac239 infection (Table 1). Five of the 18 RMs controlled SIVmac239 replication to below 10,000 vRNA copies/mL plasma during chronic infection (Figure 1). Three of the five controllers (r10055, r00008, and r10018) expressed one of the elite control-associated RM MHC I alleles, Mamu-B^∗08 or Mamu-B^∗17 (37, 38). However, one of three Mamu-B^∗08⁺ RMs (r11063) did not control SIVmac239 replication, with setpoint viral loads well above 10⁵ vRNA copies/mL plasma. The extent of SIVmac239-induced immunosuppression was monitored by flow cytometry-based computation of the CD4:CD8 T cell ratio in resting PBMCs of each animal. Average CD4:CD8 T cell ratios decreased from 1.64 pre-SIVmac239 infection to 1.14 in acute infection, and to 0.81 in chronic infection (Supplementary Figure 2).

To determine whether RhCMV-specific CTLs undergo changes in frequency, phenotype, or effector function repertoire during SIVmac239 infection, we studied RhCMV-specific CTL populations in the PBMCs of eleven Mamu-A^∗02⁺ RMs prior to SIVmac239 infection and during chronic SIVmac239 infection (defined as >10 weeks after the infecting SIVmac239 challenge; Table 2). We focused on two recently identified CTL populations specific for the RhCMV-derived peptides IE-1 VY9 (25) and IE-2 AN10 (unpublished data). We identified these antigen-specific CTL populations by two methods: pMHCI tetramer staining of resting PBMCs, and CD107a degranulation assay with ICS (CD107a/ICS) in which PBMCs were stimulated with the minimal optimal peptides IE-1 VY9 or IE-2 AN10. At least one of the two RhCMV-specific CTL populations was present in ten of eleven RMs tested (Figure 2). Nine RMs had detectable IE-1 VY9-specific populations, while six had detectable IE-2 AN10 populations. Both CTL populations were present in five of the eleven animals. The tetramer staining and CD107a/ICS analyses yielded relatively consistent results, suggesting that most circulating RhCMV-specific CTLs are responsive to stimulation with their cognate peptide. Frequencies of RhCMV-specific CTLs did not differ significantly between pre-SIVmac239 infection and chronic SIVmac239 infection timepoints (all p-values > 0.05, Welch’s t-test). On average, RhCMV IE-1 VY9-specific CTLs comprised 4–7% of circulating CD8⁺ T cells, while RhCMV IE-2 AN10-specific CTLs accounted for 1–2% of circulating CD8⁺ T cells. Remarkably, tetramer frequencies and CD107a/ICS response frequencies for RhCMV IE-1 VY9 exceeded 10% of circulating CD8⁺ T cells in two animals, and RhCMV IE-1 VY9-specific CTLs accounted for 34% of circulating CD8⁺ T cells in one animal (r00008).

To determine whether the phenotypic characteristics or effector functionalities of RhCMV-specific CD8⁺ CTLs become altered during SIVmac239 infection, we examined a number of characteristics integral to the functionality and efficacy of a CTL. These features included memory phenotype, exhaustion marker expression, direct markers of cytolytic effector function (granzyme B expression and degranulation upon stimulation), and expression of cytokines upon stimulation. CTLs specific for the RhCMV-derived peptides IE-1 VY9 and IE-2 AN10 were defined by tetramer staining and CD107a/ICS assays, as described above. Memory phenotype, granzyme B expression, and PD-1 expression were evaluated in tetramer⁺ CTLs in resting, unstimulated PBMCs. Both CTL populations were predominantly of the terminally differentiated effector memory (T_EM2) phenotype, defined as CD28^– CCR7^– CTLs (39), and the frequencies of T_EM2 CTLs did not change during SIVmac239 infection (Figure 3A). High frequencies of CTLs specific for both RhCMV epitopes expressed granzyme B, and granzyme B⁺ frequencies remained at similarly high levels during chronic SIVmac239 infection (Figure 3B). Interestingly, the exhaustion marker PD-1 was expressed at relatively high levels by both RhCMV-specific CTL populations before and during SIVmac239 infection, although the frequencies of PD-1⁺ CTLs and PD-1 median fluorescence intensities (MFIs) did not differ significantly between these timepoints (Figures 4A,B). We evaluated CTL effector functions and expression of the exhaustion marker CTLA-4 via CD107a/ICS assays in which PBMCs were stimulated with the RhCMV-derived minimal optimal peptides. In contrast to PD-1, very low frequencies of RhCMV-specific CTLs expressed CTLA-4, even during chronic SIVmac239 infection (Figure 4C). CTLA-4 MFIs for RhCMV-specific CTLs were similarly low (Figure 4D). At both pre- and chronic SIVmac239 infection timepoints, the majority of CTLs responding to either of the RhCMV peptides were polyfunctional, which was defined as simultaneous expression of all three effector function markers (CD107a⁺ IFN-γ⁺ TNF-α⁺) (Figure 3C). In summary, we observed no significant differences in the phenotypic characteristics or effector function profiles of RhCMV IE-1 VY9- or RhCMV IE-2 AN10-specific CTLs between pre- and chronic SIVmac239 infection timepoints.

HIV/SIV-specific CTLs are often characterized as “dysfunctional” or “exhausted” during chronic infection (10–13). To further evaluate our hypothesis that RhCMV-specific CTLs are refractory to the generalized immune dysfunction that occurs during SIVmac239 infection, we directly compared our two RhCMV-specific CTL populations with four well-characterized, immunodominant SIVmac239-specific CTL populations during chronic SIVmac239 infection. These included Mamu-A^∗01-restricted populations specific for Tat SL8 and Gag CM9 (7, 26, 27), as well as Mamu-A^∗02-restricted populations specific for Nef YY9 and Gag GY9 (28, 29). We analyzed the PBMCs of seven Mamu-A^∗01⁺ RMs, ten Mamu-A^∗02⁺ RMs, and one Mamu-A^∗01⁺ Mamu-A^∗02⁺ RM by tetramer staining and CD107a/ICS assays at acute and chronic SIVmac239 infection timepoints (Tables 1, 2). Interestingly, we observed no statistically significant differences in the frequencies of CTLs specific for any of the four SIVmac239-derived epitopes between acute and chronic infection (Supplementary Figure 4). Notably, the frequencies of Tat SL8-specific CTLs in chronic infection were two- to four-fold lower than in acute infection, but these differences were not statistically significant (p = 0.2204 for tetramer frequencies, p = 0.3392 for CD107a/ICS response frequencies). Furthermore, we observed no statistically significant differences in the frequencies of T_EM2, granzyme B⁺, polyfunctional, PD-1⁺, or CTLA-4⁺ SIVmac239-specific CTLs between acute and chronic infection timepoints (Supplementary Figures 5, 6). The average granzyme B⁺ frequencies of Tat SL8-specific CTLs did decrease twofold from acute to chronic infection, although this reduction was not statistically significant (p = 0.2486).

We then compared the phenotypic and functional characteristics of the four SIVmac239-specific and two RhCMV-specific CTL populations at the chronic SIVmac239 infection timepoint. Both RhCMV-specific CTL populations had significantly higher frequencies of T_EM2 CTLs than all SIVmac239-specific CTL populations, except for Gag CM9-specific CTLs (Figure 5). Nevertheless, T_EM2 frequencies for Gag CM9-specific CTLs (mean = 74.8%) were lower than those of RhCMV IE-1 VY9- (mean = 97.1%) and RhCMV IE-2 AN10- (mean = 94.4%) specific CTLs. Both RhCMV-specific CTL populations contained higher frequencies of granzyme B⁺ CTLs than any of the four SIVmac239-specific CTL populations (Figure 6). The differences in granzyme B⁺ frequencies between RhCMV IE-1 VY9- and SIVmac239-specific CTLs were all statistically significant, while the granzyme B⁺ frequencies of RhCMV IE-2 AN10-specific CTLs were significantly higher than those of CTLs specific for Tat SL8 and Nef YY9, but not Gag CM9 and Gag GY9. Most strikingly, frequencies of polyfunctional CTLs were significantly higher in both RhCMV-specific CTL populations than any of the four SIVmac239-specific CTL populations (Figure 7). However, we observed no significant differences in the frequencies of degranulating (CD107a⁺) CTLs among responding RhCMV- and SIVmac239-specific CTLs (Supplementary Figure 7).

To determine whether RhCMV-specific CTLs exhibit features of exhaustion during chronic SIVmac239 infection, we compared the frequencies of CTLs expressing the exhaustion markers PD-1 and CTLA-4 in RhCMV- and SIVmac239-specific CTL populations. On average, both RhCMV-specific CTL populations had lower PD-1⁺ frequencies than two of the SIVmac239-specific CTL populations (Gag CM9 and Nef YY9), and PD-1⁺ frequencies comparable to the other two SIVmac239-specific CTL populations (Tat SL8 and Gag GY9) (Figure 8A). Comparisons of PD-1 MFIs for the RhCMV- and SIVmac239-specific CTL populations yielded similar results, indicating that both RhCMV-specific CTL populations had significantly lower PD-1 MFIs than SIVmac239 Gag CM9- and Nef YY9-specific CTLs (Figure 8B). We found that lower frequencies of RhCMV-specific CTLs expressed CTLA-4 upon stimulation than any of the four SIVmac239-specific CTL populations studied (Figure 9A). Gag CM9-specific CTLs had significantly higher CTLA-4⁺ frequencies than both RhCMV-specific CTL populations, while Nef YY9-specific CTLs had significantly higher CTLA-4⁺ frequencies than RhCMV IE-2 AN10-specific CTLs, but not RhCMV IE-1 VY9-specific CTLs. CTLA-4 MFIs showed a similar trend, with both RhCMV-specific CTL populations exhibiting significantly lower CTLA-4 MFIs than SIVmac239 Gag CM9- and Nef YY9-specific CTLs (Figure 9B).

Five of the 18 RMs in this study controlled chronic-phase SIVmac239 viremia to less than 10,000 vRNA copies/mL plasma. Due to the association between SIV-mediated immunosuppression and increased RhCMV replication, we surmised that associations between chronic-phase SIVmac239 viremia and the phenotypic characteristics of RhCMV-specific CTLs may exist. Additionally, we sought to evaluate potential relationships between phenotypic characteristics of SIVmac239-specific CTLs and viral loads. Pearson correlation analyses revealed multiple statistically significant associations between CTL phenotype and chronic-phase SIVmac239 viremia (Table 3). Setpoint SIVmac239 viral loads were positively correlated with RhCMV IE-1 VY9-specific CTL polyfunctionality and RhCMV IE-2 AN10-specific CTL granzyme B expression. CTLA-4 expression was positively correlated with setpoint viral load for three of the four SIVmac239-specific CTL populations we analyzed. Surprisingly, PD-1 expression by Gag CM9-specific CTLs was negatively correlated with setpoint viral load. When we analyzed all four SIVmac239-specific CTL populations together, we detected positive correlations between T_EM2 and granzyme B⁺ frequencies, and setpoint viral load.