Subdominant CD8⁺ T-Cell Responses Are Involved in Durable Control of AIDS Virus Replication^▿

Animals and CD8⁺ cell depletion.

Indian rhesus macaques (Macaca mulatta) from the Wisconsin National Primate Research Center colony were typed for MHC class I alleles Mamu-A*01, -A*02, -A*08, -A*11, -B*01, -B*03, -B*04, -B*17, and -B*29 by sequence-specific PCR (23; W. M. Rehrauer et al., unpublished results). Animals were cared for according to the regulations in the Guide for the Care and Use of Laboratory Animals of the National Research Council (41), as approved by the University of Wisconsin Institutional Animal Care and Use Committee. Macaques were infected with SIVmac239 as part of previous studies (3, 37; unpublished data). None of the animals in the present study received SIV-specific vaccine formulations prior to SIV challenge, except for animal 95096, which received a lipopeptide construct containing the Gag CM9 epitope. Elite controller animals maintained chronic-phase viremia at or below 500 vRNA copy eq/ml of plasma (55). We transiently depleted ECs of peripheral CD8⁺ cells by administering monoclonal antibody cM-T807 (produced by Centocor and provided via the National Institutes of Health Nonhuman Primate Reagent Resource) intravenously in a single dose at 50 mg/kg of body weight.

CD8⁺ T-cell and NK cell quantification.

The kinetics of the depletion of the animals of CD8⁺ cell populations and the reemergence of these populations were monitored by staining Ficoll density gradient-purified peripheral blood mononuclear cells (PBMC) with fluorescently labeled antibodies specific for CD8 (clone DK25, labeled with phycoerythrin [PE] or allophycocyanin, from DakoCytomation, Carpinteria, CA), CD16 (clone 3G8, labeled with Pacific blue, from BD-Pharmingen, San Jose, CA), and CD3 (clone SP34-2, labeled with Alexa 700 or fluorescein isothiocyanate [FITC], from BD-Pharmingen). Briefly, 500,000 PBMC were incubated in 100 μl of complete tissue culture medium in the presence of the antibodies for 30 min at room temperature. Fluorochrome-stained samples were washed twice, fixed, and run on a BD-LSR-II flow cytometer (Becton Dickinson, San Jose, CA) using FACSDiva software. Data were analyzed by using FlowJo 6.1 software (Treestar, Ashland, OR). Absolute counts were calculated by multiplying the frequency of CD8⁺ T cells or CD3⁺, CD8⁺, CD16⁺ NK cells within the lymphocyte gate with the lymphocyte counts per microliter of blood obtained from matching complete blood count results.

Tetramer staining.

Epitope-specific CD8⁺-T-cell populations were enumerated by staining PBMC with Mamu-A*01, -A*02, and -B*17 tetrameric complexes loaded with synthetic epitope peptides largely as described previously (45). Briefly, 0.5 to 2 million PBMC were suspended in 100 μl of complete culture medium and incubated for 1 h at 37°C with tetramers labeled with PE. Next, antibodies recognizing CD3 (FITC-labeled SP34-2) and CD8 (clone SK1 labeled with peridin chlorophyll protein; BD Biosciences, San Jose, CA) were added, and samples were incubated for an additional 40 min at room temperature. Cells were then washed twice with fluorescence-activated cell sorter (FACS) buffer (phosphate-buffered saline containing 2% fetal bovine serum) and fixed in 1% paraformaldehyde. Data were acquired on a FACSCalibur flow cytometer (Becton Dickinson) and analyzed using FlowJo software (Treestar).

ICS.

Intracellular cytokine-staining (ICS) assays were performed as described in detail previously (54). Briefly, 500,000 PBMC were incubated for 1.5 h at 37°C in 100 μl of complete medium with anti-CD28 (clone L293; BD-Pharmingen), anti-CD49d (clone 9F10; BD-Pharmingen), and synthetic peptides (pools of 10- to 15-mers or single peptides representing minimal optimal CD8⁺ T-cell epitopes) based on the SIVmac239 protein sequence. Then 10 μg of brefeldin A per ml was added to prevent protein transport from the Golgi apparatus, and the cells were incubated a further 5 h at 37°C. Cells were then washed and stained for surface expression of CD4 (clone SK3 labeled with allophycocyanin; BD Biosciences) and CD8 (clone SK1 labeled with peridin chlorophyll protein; BD Biosciences) and fixed overnight in 1% paraformaldehyde at 4°C. The following day, cells were permeabilized in FACS buffer containing 0.1% saponin and stained for expression of the cytokines gamma interferon (IFN-γ; clone 4S.B3 labeled with FITC [BD-Pharmingen]) and interleukin-2 (IL-2; clone MQ1-17H12 labeled with PE [BD-Pharmingen]). Antibodies against these cytokines were titrated for optimal staining. After intracellular staining, PBMC were fixed in 1% paraformaldehyde for 2 h at 4°C. Events were then collected on a FACSCalibur flow cytometer and analyzed with FlowJo.

Quantification of viral RNA in plasma.

vRNA was detected in EDTA-anticoagulated plasma by quantitative reverse transcription-PCR by using a modification of a previously published protocol (9). Viral RNA was isolated from plasma as described previously (9). Viral RNA was reverse transcribed and quantified using a one-step quantitative reverse transcription-PCR kit (Invitrogen, Carlsbad, CA) with the LightCycler 1.2 (Roche, Indianapolis, IN). The final reaction mixtures (20-μl total volume) contained 0.2 mM (each) deoxynucleoside triphosphates, 3 mM MgSO₄, 0.015% bovine serum albumin, 150 ng of random hexamer primers (Promega, Madison, WI), 0.8 μl of SuperScript III reverse transcriptase and Platinum Taq DNA polymerase in a single enzyme mix, 600 nM (each) amplification primers (forward [SIV1552], 5′-GTCTGCGTCATCTGGTGCATTC-3′, and reverse [SIV1635], 5′-CACTAGCTGTCTCTGCACTATGTGTTTTG-3′), and 100 nM probe (5′-6-carboxyfluorescein-CTTCCTCAGTGTGTTTCACTTTCTCTTCTGCG-3′). The reverse transcription reaction was performed at 37°C for 15 min and then at 50°C for 30 min. An activation temperature of 95°C for 2 min was followed by 50 amplification cycles of 95°C for 2 min and 62°C for 1 min with ramp times set to 3°C/s. Synthetic 10-fold serial dilutions of an SIV gag in vitro transcript served as an internal standard curve for each run. Copy numbers for samples were determined by interpolation onto the standard by curve using the LightCycler software version 4.0.

Ex vivo suppression assay.

PBMC were isolated by centrifugation over a Ficoll density gradient. To generate target cells, a portion of the PBMC were depleted of CD8⁺ cells by using anti-CD8 nonhuman primate microbeads on an AutoMACS bead separation unit (Miltenyi, Auburn, CA) according to the manufacturer's protocol. Depletions were >99% effective. We stimulated the CD8⁻ fraction with 5 μg/ml concanavalin A (Sigma) for 18 to 24 h, after which cells were washed and incubated in R15-50 (RPMI medium-15% fetal calf serum-1% l-glutamine-50 U/ml interleukin-2 [National Institutes of Health AIDS Reference Reagent Program, Germantown, MD]) for an additional day. Target cells were then incubated with clonal SIVmac239 at a multiplicity of infection of 0.00005 for 4 h, washed twice, and added to coculture assay mixtures. To generate effector cells, PBMC derived from the same blood draw were cultured in R15-50 for 2 days. These effectors were labeled with anti-CD8 (clone SK1 labeled with FITC; BD Biosciences) and anti-CD3 (clone SP34-2 labeled with PE-Cy7; BD Biosciences) antibodies for live-cell sorting using a MoFlo cell sorter (DakoCytomation, Fort Collins, CO). Effector cells were either mock sorted (antibody labeled and run through the cell sorter), depleted of NK cells (CD3⁻, CD8⁺), depleted of CTL (CD3⁺, CD8⁺), or depleted of all NK and CTL to >99% purity. After sorting, the viability and quantity of effector cells were confirmed by trypan dye exclusion. For the suppression assay, 150,000 target cells were mixed with autologous effectors at an effector/target cell ratio of ∼3:1. The absolute number of effector cells was adjusted to ensure that the number of non-CD8⁺ effector cells remained constant for each condition, and each population was tested in duplicate for each experiment. Cells were cultured in 2 ml of R15-50 in 24-well plates. Culture medium (0.5 ml) was removed on days 3, 5, and 7 for quantitation of viral RNA and replaced with fresh medium. On day 7, cells were removed from culture and stained for surface expression of CD3, CD4, and CD8 and intracellular Gag p27 (clone 55-2F12; National Institutes of Health AIDS Research and Reference Reagent Program) by using Fix and Perm (CALTAG, Burlingame, CA). Events were collected on a FACSCalibur flow cytometer and analyzed with FlowJo.

Viral genome sequencing.

Complete SIV genomes were sequenced as described in a previous study (43). Eighteen primer pairs were used in a single-step reverse transcription-PCR to produce overlapping amplicons, which were sequenced on a 3730 DNA analyzer (Applied Biosystems, Foster City, CA). Sequences were assembled using CodonCode aligner (CodonCode, Deadham, MA). DNA sequences were conceptually translated by using the MacVector 8 trial version.

Depletion of CD8⁺ cells resulted in massive viral replication.

We identified ECs from among animals infected in previous challenge experiments. ECs rapidly controlled acute infection with SIVmac239 and maintained low or undetectable viremia (≤500 vRNA copy eq/ml of plasma) during chronic infection. Six ECs were used in the present study (Table 1); four of these expressed Mamu-B*17, and two did not. Data on the epitopic breadths of cellular immune responses to SIVmac239 during acute infection were available for four of these animals (data summarized in Table 2); we have reported on the acute-phase responses of animals 95096 and AJ11 previously (37, 44). Animals AJ11 and 01064 made particularly broad responses, with PBMC from each animal recognizing more than 20 separate 15-mer pools (Table 2). Animals 95096 and 95071 were tested by ICS for IFN-γ (data not shown). These experiments revealed that CD8⁺ lymphocyte responses in animal 95096 were dominated by those directed to Tat pool A, which contained the Mamu-A*01-restricted Tat SL8 epitope, and responses in animal 95071 were dominated by those directed to Vpr pool B, which did not contain a known minimal optimal epitope. Acute-phase cellular immune responses in AJ11 and 01064 were assessed by using an IFN-γ enzyme-linked immunospot assay. AJ11's acute-phase repertoire was dominated by a population responding to Nef pool C, which did not contain a previously described minimal optimal epitope. 01064 had essentially codominant populations recognizing Nef pool D, which contained the Mamu-A*02-restricted epitope Nef YY9-159, and Vif pool E, which contained no known epitopes. Although they were gathered from different experiments, the available data indicate that ECs targeted multiple SIV antigens during the acute phase of infection, before viremia was controlled (Table 2).

To transiently deplete animals of CD8⁺ lymphocytes, ECs were given recombinant antibody cM-T807, which recognizes CD8α, intravenously at 50 mg/kg on day 0. Circulating CD8⁺ T cells were almost completely removed from peripheral blood for approximately 3 weeks (Fig. 1a). CD8α is also present on NK cells (CD3⁻, CD8⁺, CD16⁺), of which animals were depleted with the same kinetics as CD8⁺ T cells (Fig. 1b). While there was some variation among animals, the numbers of both CD8⁺ T cells and NK cells appeared to return quickly to predepletion levels.

During the period of CD8⁺ cell depletion, plasma viremia increased in each animal by a factor of 2 to 4 logs. Most strikingly, animal 95071 had undetectable viremia on the day of treatment but over 8 million copy eq/ml 14 days later (Fig. 1c).

Viremia was controlled when CD8⁺ cells repopulated the periphery.

Peripheral CD8⁺ cells reappeared coincident with a reassertion of control over viral replication in all six ECs. Previous studies with normal-progressor macaques had shown that transient depletion of CD8⁺ lymphocytes resulted in increased viral burdens, but they had not evaluated total SIV-specific responses by the rebounding CD8⁺ population (18, 27, 28, 36, 50, 51). We reasoned that analyzing the antigen specificity of returning CD8⁺ T cells could identify T-cell correlates of control in these animals. One of the hallmarks of ECs is the ability of their antigen-specific CD8⁺ T cells to proliferate (38), so we hypothesized that populations of SIV-specific CD8⁺ T cells would undergo a generalized expansion upon return to the periphery. At 21 days after depletion, when CD8⁺ cells first returned, we used Mamu-A*01, -A*02, and -B*17 tetramers loaded with commonly recognized epitopes to measure the antigen-specific CD8⁺-T-cell response.

To our surprise, the reassertion of control of viral replication was associated not with generalized proliferation of all previously detected SIV-specific CD8⁺ T-cell populations but rather with differential increases in the frequencies of particular CD8⁺ T-cell specificities. The populations of only a few specificities expanded in most animals, and previously subdominant populations tended to show a much greater degree of increase than immunodominant ones. Remarkably, we could detect only a single CD8⁺ T-cell response in the Mamu-B*17-positive macaque AJ11. This response was directed against the Nef MW9 epitope (5.67% of CD3⁺, CD8⁺ T cells) (Fig. 2a), recognized by a subdominant population of CD8⁺ T cells during acute infection (37). The frequency of Nef MW9-specific CD8⁺ T cells increased 22-fold after depletion.

Indeed, tetramer assays revealed strong increases in the frequencies of previously subdominant populations in each of the ECs. Prior to depletion, the immunodominant response in animal 95096 was directed against the well-described Mamu-A*01-restricted Gag CM9 epitope. The frequency of Gag CM9-specific T cells changed very little after depletion (1.2-fold increase) (Fig. 2b). However, the previously subdominant population of CD8⁺ T cells recognizing the Mamu-B*17-restricted Env FW9 and Nef IW9 epitopes expanded ∼10-fold following depletion in this animal. Similarly, a strong immunodominant response to the Mamu-B*17-restricted epitope Nef IW9 occurred in animal 95071 before depletion (9.1% of CD3⁺, CD8⁺ T cells) (Fig. 2c). As CD8⁺ T cells rebounded following depletion, the Nef IW9-specific population was still dominant (10.6% of CD3⁺, CD8⁺ T cells; 1.2-fold increase), but there was an eightfold expansion of cells specific for a different Mamu-B*17-restricted epitope, Vif HW8 (1.0 to 8.2%) (Fig. 2c). The Mamu-B*17-positive animal 98016 made strong responses against the Vif HW8 and Nef IW9 epitopes after depletion (41- and 2,436-fold increases, respectively), although these responses were barely detectable prior to depletion (Fig. 2d). Animal 01064 expressed Mamu-A*02, and not Mamu-B*17, but its SIV-specific CD8⁺ T-cell response showed a similar pattern. Before depletion, it made a dominant response to the Mamu-A*02-restricted Gag GY9 epitope. This response expanded fourfold after depletion, while the previously subdominant population of CD8⁺ T cells recognizing the Nef YY9 epitope expanded 15-fold, becoming the newly dominant SIV-specific population (Fig. 2e).

Control of viral replication was correlated with previously subdominant populations of epitope-specific CD8⁺ T cells and broad CD4⁺ T-cell responses against Gag.

Our analysis at 21 days after depletion was limited to tetramer staining due to constraints on blood draw volume with rhesus macaques. Therefore, we obtained maximal blood draw volumes the following week to measure the cellular immune responses to all nine viral gene products. At 28 days postdepletion, we performed ICS for IFN-γ and IL-2 by using previously identified minimal optimal epitope peptides (3, 31, 40, 54) as well as pools of 15-mer peptides spanning the entire SIV proteome. This approach allowed us to detect responses to both known and previously undescribed epitopes recognized by CD4⁺ and CD8⁺ T cells.

For animals AJ11 and 95096, ICS revealed that only a few SIV-specific CD8⁺ T-cell responses were present (Fig. 3a and b). In accordance with the tetramer result, ICS assays detected only a single CD8⁺ T-cell response in AJ11, directed against the Nef MW9 epitope recognized by a previously subdominant T-cell population. ICS also detected three CD8⁺ T-cell responses to previously described epitopes in 95096, in accordance with the results of tetramer staining. However, the Mamu-B*17-restriced epitope Nef IW9 failed to elicit IFN-γ or IL-2 responses in 95096, although tetramer stains 1 week earlier detected a population of T cells that bound this peptide (0.99% of CD3⁺, CD8⁺ T cells) (Fig. 2b). 95096 also made CD8⁺ T-cell responses to three peptide pools, one each from Gag, Vif, and Nef, which could not be accounted for by the responses to known minimal optimal epitopes. These responses are therefore likely directed against previously undescribed epitopes (Fig. 3b). ICS additionally revealed a robust antiviral CD4⁺ cell response in both 95096 and AJ11, directed primarily against Gag (Fig. 4a and b).

Animal 95071 had an exceptionally strong and broad CD8⁺ T-cell response at 4 weeks postdepletion, with CD8⁺ T cells recognizing at least 30 distinct epitopes. This response was dominated by CD8⁺ T cells recognizing Vif HW8 (10.9% of CD8⁺, IFN-γ⁺ T cells). Other responses were weak in comparison (Fig. 3c). Although this animal's response to Nef IW9 (10.6% of CD3⁺, CD8⁺ T cells) (Fig. 2c) appeared to be dominant in tetramer assays, CD8⁺, IFN-γ⁺ T-cell responses to Nef IW9 were far more modest (0.55% of lymphocytes expressing CD8 and IFN-γ), similar to our observations with animal 95096. 95071 also made extremely strong CD4⁺ T-cell responses to SIV, even during the CD8⁺ cell depletion phase when viral replication was high (data not shown). These responses expanded upon recovery of CD8⁺ cells and were dominated by responses to Rev pool A, with a substantial response to Gag pool B (Fig. 4c).

Animal 98016 also had broad CD8⁺ and CD4⁺ T-cell responses following depletion (19 epitopes were recognized by CD8⁺ and CD4⁺ T-cell subsets), although they were smaller in magnitude than those in 95071. There was a strong response to Vif HW8 (2.3%) (Fig. 3d), although Nef IW9-specific CD8⁺ T cells had shown higher frequencies in tetramer assays. The strongest CD8⁺ T-cell response in this animal was directed against Nef pool D (3.3% of lymphocytes expressing CD8 and IFN-γ), while the response by the highest-frequency CD4⁺ T cells recognized Gag pool I (0.18% of lymphocytes expressing CD4 and IFN-γ) (Fig. 4d).

Animal 00078, which did not express Mamu-A*01, -A*02, or -B*17, made at least 13 distinct CD8⁺ T-cell responses, dominated by IFN-γ⁺ cells responding to pools of peptides from Vif (Vif pool E, 0.72%) (Fig. 3e) and Nef (Nef pool D, 0.26%). 00078 made at least 20 CD4⁺ T-cell responses, with high-frequency responses to pools of peptides from Gag (Gag pool B, 0.22%; Gag pool G, 0.2%) (Fig. 4e) and Nef (Nef pool F, 0.14%). The majority of these responding CD4⁺ cells were positive for both IFN-γ and IL-2 (data not shown). Similarly, animal 01064, which expresses Mamu-A*02 but not Mamu-B*17, made at least 18 distinct CD8⁺-T-cell responses, the largest targeting the Nef YY9 epitope (1% of lymphocytes expressing CD8 and IFN-γ) (Fig. 3f). Other strong responses were directed against Rev peptide pools B and C (total of 0.8% of lymphocytes expressing CD8 and IFN-γ) and Vif pool E (0.34% of lymphocytes expressing CD8 and IFN-γ). Interestingly, although we detected a strong response to the previously dominant Gag GY9 epitope in tetramer assays, this epitope never stimulated an IFN-γ response in 01064 following depletion (Fig. 3f and data not shown). CD4⁺ cells from 01064 recognized nine different peptide pools after depletion, with the strongest response directed against Gag pool B (0.13% of lymphocytes expressing CD4 and IFN-γ) (Fig. 4f).

EC macaques showed different patterns of epitope recognition after CD8 depletion treatment. The epitopic breadths of responses in animals 95071 and 98016 increased over those seen at baseline, while the numbers of antigens targeted by other animals were similar pre- and postdepletion. Despite these differences, there were nonetheless clear similarities in the total cellular immune responses as viremia declined. We observed differential increases in the frequencies of SIV-specific CD8⁺ T-cell populations, with previously subdominant responses increasing to a much greater degree than dominant ones. Furthermore, expanding CD8⁺ T-cell populations in each EC preferentially targeted epitopes or pools from Nef and Vif. ECs also each had strong and broad CD4⁺ T-cell responses. Strikingly, while the dominant CD8⁺ T-cell responses were different in each EC, three of six ECs shared a dominant CD4⁺ T-cell response to Gag pool B, while the other three ECs made strong but subdominant responses to Gag pool B peptides (Fig. 4).

Both NK and CD8⁺ T cells from ECs suppressed SIV replication in a novel ex vivo assay.

Since the cM-T807 antibody depleted animals of peripheral NK cells as well as CD8⁺ T cells, it is possible that the rebound in viremia we observed was due simply to the removal of NK cells, with CD8⁺ T-cell populations expanding in response to the antigens but not actively controlling viral replication. Following the animals' return to set-point viremia, we sought to determine the relative contributions of NK and CD8⁺ T cells in suppressing viral replication by using a novel ex vivo assay. We cocultured animals' PBMC with activated, autologous target cells superinfected with clonal SIVmac239. These unfractionated PBMC suppressed viral replication (Fig. 5a), so we reasoned that the depletion of effector PBMC of NK and/or CD8⁺ T cells should inhibit suppression, analogous to in vivo CD8 depletion. Indeed, as we saw in vivo, the removal of all CD8⁺ cells from the effector PBMC resulted in levels of viral replication equal to that seen when no effectors were added to the cultures (Fig. 5d). Removing either NK or CD8⁺ T-cell populations from the cultures also led to increased levels of viral replication. The relative contribution of each subset was variable among the animals tested (Fig. 5b and c), though consistent for each animal across multiple experiments at different times postdepletion. These data indicate that both CD8⁺ T cells and NK cells from these animals have antiviral functions and likely contribute to maintaining elite control of SIV.

Viral sequencing revealed variations in multiple epitopes recognized by CD8⁺ T cells.

We sequenced the entire genome of the replicating plasma virus from each animal at the peak of viremia to investigate the potential relationship between escape mutations and the expansion of populations of CD8⁺ T cells specific for a given epitope. We were particularly interested in epitopes targeted by high-frequency Mamu-B*17-restricted CD8⁺ T cells, due to the association between Mamu-B*17 expression and the control of viremia (55). Responses to the Nef IW9 epitope are typically immunodominant in acute infection in Mamu-B*17-positive macaques, and this epitope showed variations at position 1 in all four Mamu-B*17-positive ECs. Three of these Mamu-B*17-positive ECs had CD8⁺ T cells which bound Mamu-B*17 tetramers containing IW9 peptide (Fig. 2), while the frequency of CD8⁺ T cells secreting IFN-γ in response to this peptide was far lower (Fig. 3). It is therefore possible that the variant sequence is able to drive the expansion of populations of CD8⁺ T cells that bind the tetramer but do not produce cytokines in response to the IW9 peptide. Animal 95096's virus harbored a glutamine-for-histidine substitution at position 2 in the Env FW9 epitope, a substitution not seen in Mamu-B*17-positive progressors (Fig. 6 and data not shown). Nonetheless, the population of Env FW9-specific CD8⁺ T cells expanded in 95096 after depletion, though the percentage of cells secreting IFN-γ (0.2%) (Fig. 3b) was lower than that detected by the FW9 tetramer (0.94%) (Fig. 2b). By contrast, virus from animal 95071 showed mixed-base heterogeneity in this epitope at position 8, a mutation also observed in virus from Mamu-B*17-positive progressors (Fig. 6 and data not shown). This may be a more effective escape mutation, since both tetramer and ICS assays detected Env FW9-specific CD8⁺ T cells only at very low frequencies in 95071.

Mutations within epitopes recognized by CD8⁺ T cells tended to correlate with the diminution or absence of responses (Fig. 6). Animals 98016 and 01064 had replacements within the Mamu-A*02-restricted epitope Gag GY9 usually recognized by immunodominant responses. These animals had expanding Gag GY9 tetramer-positive populations (Fig. 2d and e), but we detected no IFN-γ responses to the Gag GY9 peptide (Fig. 3d and e). Virus from animals 95096 and AJ11 had variations in the Mamu-B*17-restricted Env LF11, and we detected no responses to this epitope (Fig. 6). Animal 95096's virus had variations in the Mamu-B*17-restricted Pol MF8 and Mamu-A*01-restricted Tat SL8, to which no responses were detected by tetramers or ICS (Fig. 6). However, escape mutations could not always explain the lack of certain responses. No variation was detected in Vif HW8 in the four Mamu-B*17-positive ECs. Animals 95071 and 98016 made extremely strong responses to this epitope after depletion, but no response was detected in 95096 or AJ11.