Immunogen-Specific Strengths and Limitations of the Activation-Induced Marker Assay for Assessing Murine Antigen-Specific CD4⁺ T Cell Responses

The identification and quantification of Ag-specific CD4⁺ T cells is important for studying immune responses to vaccination, infection, and autoimmunity. Intracellular cytokine staining (ICS) and ELISPOT assays are gold-standard techniques for detecting Ag-specific CD4⁺ T cells and are based on cytokine production following restimulation with specific Ag (1), whereas MHC/peptide tetramer reagents provide direct labeling of Ag-specific cells by binding to TCRs for detection by flow cytometry (2). Typically, cytokines that are produced in abundance that can be detected by ICS via flow cytometry are IL-2, TNF-α, and IFN-γ (3). Moreover, ICS and ELISPOT assays only identify cells that produce cytokines above certain detectable thresholds. In 2016, the Crotty laboratory (4, 5) introduced the activation-induced marker (AIM) assay that identifies Ag-specific CD4⁺ T cells based on the upregulation of TCR stimulation-induced surface markers. This cytokine-independent method measures the specific upregulation of cell-surface activation markers following stimulation with specific Ag in culture (often ∼18 h but varies by study) compared with cells cultured in the absence of Ag (unstimulated). Commonly used AIMs include: OX40, CD25, PD-L1, CD40L, and CD69 (4–6). The AIM assay is particularly useful in identifying Ag-specific T follicular helper (Tfh) cells, a CD4⁺ T cell subset that is important for providing help to B cells, that have tightly regulated cytokine expression that makes them difficult to detect by cytokine-dependent methods, such as ICS (4–7). In addition, the AIM assay is useful in that whole protein Ag can be used for in vitro culture stimulation and identification of Ag-specific cells without needing to know specific MHC-restricted epitopes that other methods, such as MHC/peptide tetramer reagents, require (4–6). Since its introduction, the AIM assay has been used in multiple human and nonhuman primate studies to evaluate CD4⁺ and CD8⁺ T cell responses to vaccination and infection (4–6, 8–16).

Recently, several laboratories have begun to use the AIM assay in murine studies to evaluate Ag-specific T cell responses. One study used the AIM assay to detect hemagglutinin-specific Tfh cells based on upregulated CD25, OX40, and CD154 expression following PR8 influenza infection or hemagglutinin immunization in C57BL/6 mice (17). Another study used the AIM assay to clone HIV envelope–specific Tfh cells following envelope trimer immunization to generate TCR-transgenic mice (11). Our laboratory recently used the AIM assay to validate the presence of Ag-specific CD4⁺ T cells induced by adjuvanted plague subunit vaccination in mice (18). Despite being proven as a valuable tool to identify Ag-specific T cells, the AIM assay has not been validated for its strengths and limitations in various murine models. In this study, we used known TCR-transgenic mouse models to evaluate the efficacy of the AIM assay for identifying Ag-specific CD4⁺ T cells in response to various types of immune responses, including immunization, infection, and autoimmunity. Using adoptive transfer of congenically marked TCR-transgenic donor cells, we quantified the frequency of donor Ag-specific TCR-transgenic CD4⁺ T cells that upregulated expression of OX40 and CD25 (AIM⁺ cells) with the AIM assay. Our experimental designs allowed us to determine whether the AIM assay efficacy varied for effector versus memory cells induced by protein immunization and for acute versus chronic infection. Our findings reveal important considerations for using the AIM assay for various applications in mouse models of immunization, infectious disease, and autoimmunity.

Naive congenically marked (CD45.1) SMARTA TCR-transgenic CD4⁺ T cell splenocytes specific to the lymphocytic choriomeningitis virus (LCMV) glycoprotein (GP)_61-80 epitope (19) were i.v. transferred into naive C57BL/6 (CD45.2) mice (The Jackson Laboratory, Bar Harbor, ME). For adoptive transfers for LCMV Armstrong infections and GP immunizations, 2 × 10⁴ SMARTA CD45.1⁺CD4⁺ T cells were transferred into recipient mice. For adoptive transfers for LCMV clone 13 infections, 2 × 10³ SMARTA CD45.1⁺CD4⁺ T cells were transferred into recipient mice. Naive congenically marked (CD45.2) OT-II TCR-transgenic CD4⁺ T cell splenocytes specific to the chicken OVA_323-339 epitope (20) were i.v. transferred into naive congenically marked (CD45.1) C57BL/6 background mice. For adoptive transfers for OVA immunizations, 2 × 10⁴ or 1 × 10⁵ OT-II CD45.2⁺CD4⁺ T cells were transferred into recipient mice. For polyclonal and affinity two-dimensional micropipette (2D-MP) experiments, Foxp3 EGFP reporter mice (21) were obtained from The Jackson Laboratory (strain number 006772). NOD.BDC2.5 TCR-transgenic mice (22) were acquired from The Jackson Laboratory (strain number: 004460). Animal experiments were conducted under approved University of Utah Institutional Animal Care and Use Committee protocols.

For viral infections, mice were either infected with 2 × 10⁵ PFU of LCMV Armstrong via i.p. injection or 2 × 10⁶ PFU of LCMV clone 13 via i.v. injection. For protein immunizations, mice were immunized via i.m. injection (quadriceps) with 2–10 μg rLCMV GP or 10–50 μg of OVA with the addition of AddaVax (InvivoGen) adjuvant at a 1:1 ratio. rLCMV GP-expressing 293A cells were kindly provided by Dr. C. Davis (Emory University), and rGP was purified as described previously (23).

Single-cell suspensions of spleens and pooled inguinal and lumbar draining lymph nodes (dLN) were prepared using 70-μm cell strainers. RBCs were lysed from splenocyte cell suspensions by incubation in Ammonium-Chloride-Potassium Lysing Buffer (Life Technologies). Cell suspensions were resuspended in RPMI 1640 media supplemented with 5% FBS. CD4⁺ T cells were purified from spleens of LCMV Armstrong– and clone 13–infected mice and enriched before culture using a magnetic CD4⁺ T cell–negative selection kit (STEMCELL Technologies). T cell–depleted APCs were enriched by staining splenocyte suspensions with PE-conjugated anti-CD4 and anti-CD8 Abs followed by magnetic separation using a magnetic column and an Anti-PE MicroBeads Kit (Miltenyi Biotec). For experiments with NOD.BDC2.5 mice, inguinal (iLN) and pancreatic LN (pLN) and pancreatic islets were isolated from 14-wk-old prediabetic male mice. LN single-cell suspensions were prepared by disrupting the tissue using frosted glass slides. To isolate islet cells, pancreata were perfused with a solution of collagenase 4 (600 units/ml; catalog number LS004188; Worthington Biochemical) in 1× HBSS with 5% FBS and digested for 30 min at 37°C. Individual islets were picked by hand and further digested with Cell Dissociation Buffer (catalog number 13151-041; Life Technologies) at 37°C for 15 min. Blood was collected via submandibular venipuncture.

A total of 1 × 10⁶ cells from isolated tissues was plated in individual wells of 96-well or 48-well (islets) flat-bottom plates (Greiner Bio-One) in RPMI 1640 supplemented with 10% FBS, 1% l-glutamine, 1% penicillin/streptomycin, and 50 μM 2-Mercaptoethanol (RF10 media). For restimulation with protein Ags for the AIM assay, rLCMV GP or OVA protein at a concentration of 5–50 μg/ml was used. For peptide stimulation, RF10 media was supplemented with 2–10 μg/ml of either GP_61-80 or OVA_323-339 peptides. For plate-bound tetramer restimulation, plates were coated with 1:300 2.5HIP tetramer [I-A(g7) LQTLALWSRMD] in 1× PBS for a minimum of 30 min at 37°C and gently washed twice prior to use. Control wells included RF10 media without protein or peptide. Cells were incubated for 18 h at 37°C with 5% CO₂ before transfer to round-bottom plates for FACS staining. The percent AIM⁺ cells was determined by taking the percent OX40⁺CD25⁺ cells in the sample cultured in the presence of protein/peptide Ag and subtracting the percent OX40⁺CD25⁺ cells in the sample cultured in the absence of exogenous Ag.

CXCR5 surface staining was performed using a three-step protocol (24) using purified rat anti-mouse CXCR5 primary Ab (2G8; BD Biosciences) in 1× PBS supplemented with 2% FBS plus 3.4% BSA (Sigma-Aldrich) plus 2% mouse serum (CXCR5 staining buffer) (Sigma-Aldrich), a secondary Biotin-SP conjugated Affini-pure F(Ab’)₂ Goat Anti-Rat IgG (Jackson ImmunoResearch Laboratories), and then with a fluorochrome-conjugated streptavidin (allophycocyanin or PE-Cy7; Invitrogen) in 1× PBS supplemented with 2% FBS (FACS buffer). Cells were surface stained in FACS buffer for 15–30 min on ice with fluorochrome-conjugated Abs: CD4 (RM4-5), B220 (RA3-6B2), PD-1 (RMP1-30, 29F.1A12), OX40 (OX-86), CD25 (PC61), PD-L1 (10F.9G2), CD44 (IM7), CD154 (MR1), CD45.1 (A20), CD45.2 (104), LIVE/DEAD Fixable Near-IR Dead Cell Stain (from BD Biosciences, eBioscience, BioLegend, Vector Laboratories, and Invitrogen). Transcription factor staining was performed by fixing, permeabilizing, and staining using the Foxp3 Permeabilization/Fixation kit and protocol (eBioscience) with fluorochrome-conjugated Abs: Bcl6 (K112-91) and Foxp3 (FJK-16s) (BD Biosciences and eBioscience, respectively).

ICS was performed following 5-h stimulation with GP_61-80 or OVA_323-339 peptides and brefeldin A (GolgiPlug; BD Biosciences) in RPMI medium supplemented with 5% FBS at 37°C with 5% CO₂. Cells were then stained for surface Ags, followed by permeabilization, fixing, and staining using the Cytofix/Cytoperm kit and protocol (BD Biosciences) with fluorochrome-conjugated Abs: IFN-γ (XMG1.2), TNF-α (MP6-XT22), and IL-2 (JES6-5H4). Cell sorting was performed using an FACSAria II (BD Biosciences). Flow cytometry data were analyzed using FlowJo v10 software.

The relative 2D affinity (2D-MP) of polyclonal IAb:GP_66-77-specific cells from C57BL/6-FOXP3 EGFP mice (The Jackson Laboratory) was measured using the previously described 2D-MP assay (25–27). Seven days following LCMV infection and AIM assay culture, splenic CD4⁺ T cells were negatively enriched by magnetic separation (130-104-454; Miltenyi Biotec) and stained and sorted to obtain FOXP3⁻(GFP⁻)CD25⁻OX40⁻ AIM-negative CD4⁺ T cells and FOXP3⁻(GFP⁻)CD25⁺OX40⁺ AIM-positive CD4⁺ T cells. Human RBCs were coated with IAb:GP_66-77 monomer or negative control H2Db:NP_366-374 monomer obtained from the National Institutes of Health Tetramer Core. Quantification of peptide–MHC (pMHC) density with anti-IA/IE Ab (M5/114/15/2; eBioscience) and TCR surface density with anti-mouse TCR-β PE Ab (H57-597; BD Biosciences) were determined using Quantibrite PE quantification beads (BD Biosciences). Quantification of binding events and TCR/pMHC affinity calculations was calculated as previously described (25–27). Each cell was tested to the monomer of interest and then to the control monomer to determine Ag specificity of the cell.

All experiments were analyzed using Prism 9 (GraphPad). Statistically significant p values <0.05 are indicated and determined using a two-tailed unpaired Student t test, paired Student t test, or Mann–Whitney U test.

To test the efficacy of the AIM assay to identify effector Ag-specific CD4⁺ T cells after adjuvanted protein immunization in mice, we adoptively transferred congenically marked naive SMARTA (CD45.1⁺) or OT-II (CD45.2⁺) TCR-transgenic CD4⁺ T cells into recipient C57BL/6 mice. SMARTA (CD45.1⁺) TCR-transgenic CD4⁺ T cells are specific for the LCMV GP_61-80 epitope (19), whereas OT-II (CD45.2⁺) TCR-transgenic CD4⁺ T cells are specific for the OVA_323-339 epitope (20). Recipient mice of SMARTA CD4⁺ T cells were then immunized i.m. with rLCMV GP and recipients of OT-II CD4⁺ T cells were immunized i.m. with chicken OVA in AddaVax adjuvant. Donor SMARTA (CD4⁺CD45.1⁺) or OT-II (CD4⁺CD45.2⁺) cells from pooled inguinal and lumbar dLN were then analyzed 7 d postimmunization. As OX40 (CD134) and CD25 (IL-2Rα) are the primary surface receptors used in both human and murine AIM assay studies (6, 17) to denote activated Ag-specific CD4⁺ T cells, we first evaluated the expression of these receptors on donor SMARTA and OT-II cells after 18-h stimulation in culture with rGP or OVA proteins or GP_61-80 or OVA_323-339 peptides (Fig. 1A). As murine regulatory T (Treg) cells highly express both OX40 and CD25 (28), Foxp3⁺ cells were excluded from our analyses (Fig. 1A). We first assessed whether stimulation with whole recombinant proteins or TCR-transgenic cell-specific peptides improved AIM assay identification of Ag-specific cells. Following culturing with Ags, the AIM assay marked ∼60–80% of SMARTA TCR-transgenic CD4⁺ T cells as OX40⁺CD25⁺AIM⁺ cells. Meanwhile, only ∼2–7% of SMARTA cells that were cultured without Ag expressed OX40 and CD25 (Fig. 1B, 1C). We tested two concentrations of adoptively transferred donor OT-II cells: a lower (2 × 10⁴ CD4⁺ T cells) concentration for direct numerical comparison with the transferred number of donor SMARTA CD4⁺ T cells and a 5-fold higher concentration (1 × 10⁵ CD4⁺ T cells). Similar to the observation in the SMARTA system, both chicken OVA protein and OVA_323-339 peptide stimulation induced ∼60–80% of OT-II TCR-transgenic CD4⁺ T cells to have upregulated expression of OX40 and CD25 (Fig. 1B, 1C). In addition, when the protein and peptide stimulation conditions were compared between each group, there was no difference in the frequencies of OX40⁺CD25⁺ SMARTA or OT-II CD4⁺ T cells (Fig. 1B, 1C). Together, these data show that the AIM assay effectively identifies the majority of protein immunization-induced Ag-specific CD4⁺ T cells.

We also wanted to compare the AIM assay to ICS and tetramer staining for its ability to detect immunization-induced polyclonal Ag-specific CD4⁺ T cells. We immunized C57BL/6 mice with rGP in AddaVax adjuvant. Lymphocytes from dLN were analyzed 7 d postimmunization. For AIM assay analyses, cells were either stimulated with rLCMV GP or unstimulated in RF10 media for 18 h. For cytokine staining, cells were cultured with LCMV GP_61-80 peptide for 5 h followed by ICS. For tetramer staining, lymphocytes were stained with IA(b) LCMV GP_66-77 MHC II tetramer for 2 h. Our results indicate that IAb:GP_66-77 tetramer staining identified a significantly higher frequency of polyclonal GP-specific CD4⁺ T cells compared with the AIM assay, whereas the frequency of cytokine-producing (ICS⁺) CD4⁺ T cells was relatively similar to the AIM⁺ frequency (Fig. 1D, 1E). We also looked at the polyclonal response to OVA immunization and found that the AIM assay identified fewer OVA-specific CD4⁺ T cells compared with the ICS assay (Fig. 1F).

To better validate the sensitivity of the AIM assay in identifying Ag-specific CD4⁺ T cells longitudinally at the effector and memory cell stages of the response, we adoptively transferred naive SMARTA (CD45.1⁺) CD4⁺ T cells into recipient C57BL/6 mice. Recipient mice were then either immunized by i.m. injection with rLCMV GP or infected i.p. with 2 × 10⁵ PFU of LCMV Armstrong, an acute viral strain of LCMV in which infection is cleared by day 8 postinfection. For analyses of Ag-specific CD4⁺ T cells, spleens from infected mice and pooled inguinal and lumbar dLN from immunized mice were collected at 7, 14, and 60 d postimmunization/infection to represent early effector, late effector, and memory time points, respectively. In addition, we harvested naive SMARTA splenocytes from naive SMARTA TCR-transgenic mice. To control for batch effects from Ab staining and flow cytometric analyses, adoptive transfers, immunizations, and infections of recipient mice were staggered so spleens and dLN for all four time points (days 0, 7, 14, and 60) were collected on the same day. Lymphocytes from spleen and dLN were cultured either with GP or without Ag stimulation. In addition, ex vivo cell preparations from dLN and spleen were stored packed on ice and at 4°C overnight (instead of plated for cell culture) and stained at the same time as Ag-stimulated and unstimulated cells following their 18 h of culture.

As predicted, naive SMARTA cells had minimal upregulation of OX40 and CD25 expression when stimulated overnight without Ag or ice-packed overnight. When cultured with GP, the AIM assay marked 7% of naive SMARTA cells (Fig. 2A). Seven days postinfection with LCMV Armstrong, surprisingly, the AIM assay identified only ∼17% of total SMARTA CD4⁺ T cells, as measured by upregulated expression of OX40 and CD25, and was the lowest frequency of AIM⁺ (OX40⁺CD25⁺) cells observed among all three time points (Fig. 2A, 2B). In addition, day 7 effector SMARTA cells had an OX40⁺CD25⁺ background staining of ∼2.5%, which was higher than ex vivo control cells from the same mouse (Fig. 2A). Interestingly, the frequency of viral infection–induced AIM⁺ SMARTA cells significantly increased to ∼35–40% at day 14 postinfection and increased further to ∼50% of memory SMARTA cells 60 d postinfection (Fig. 2A, 2B). In contrast to viral infection, following adjuvanted GP protein immunization, the AIM assay identified similar frequencies (means between 40 and 60%) of SMARTA cells at effector (days 7 and 14) and memory (day 60) time points (Fig. 2A, 2B). Interestingly, whereas rGP protein restimulation used for the AIM assay of cultured splenocytes or PBMCs induced only a small portion (∼18–45%) of viral infection–induced day 7 effector SMARTA cells to upregulate AIM markers, restimulation with GP_61-80 peptide resulted in a significant increase in the efficacy of the AIM assay, resulting in ∼70–90% of SMARTA cells identified as AIM⁺ (Fig. 2C).

To address whether APCs from day 7 LCMV-infected mice were defective for inducing AIM marker expression on day 7 effector SMARTA cells, we purified CD4⁺ T cells from spleens of day 7 acutely infected mice and cultured them for 18 h with APCs from either naive or acutely infected (day 7) mice (with the addition of either GP protein or GP_61-80 peptide). Our results show that effector SMARTA cells upregulated similar expression of OX40 and CD25 to become AIM⁺ when cultured with GP protein and APCs from either naive or day 7 infected mice, indicating that reduced AIM⁺ frequency of day 7 effector SMARTA cells following acute LCMV infection is not due to defects in the APCs from acutely infected mice (Fig. 2D). Together, these data show that the AIM assay using protein stimulation typically identified the majority of Ag-specific immunization-induced effector and memory CD4⁺ T cells and viral infection–induced memory CD4⁺ T cells, whereas viral infection–induced day 7 effector cells are somewhat refractory to protein stimulation, resulting in poor AIM assay efficacy. Importantly, this latter defect can be rescued with the use of epitope-specific peptide restimulation, resulting in the majority of effector SMARTA cells being marked as AIM⁺.

We performed further experiments to try to improve the efficacy of the AIM assay for detecting virus infection–induced effector cells by either titrating the duration of the cell culture/restimulation period or the Ag concentration used. Culture and restimulation with GP protein for 6 h resulted in a very low frequency (≤12%) of SMARTA cells that were AIM⁺, whereas 12, 18, and 24 h resulted in a significantly higher percent of AIM⁺ cells, with the 12-h culture period resulting in modestly higher percent of AIM⁺ cells compared with 18- and 24-h culture (Supplemental Fig. 1A). When we titrated the Ag concentration during restimulation, whereas ∼70% of day 7 effector SMARTA cells were marked as AIM⁺ when stimulating with GP_61-80 peptide, only ∼10–43% of SMARTA cells were OX40⁺CD25⁺AIM⁺ when stimulated with 5, 10, 20, or 40 μg/ml of GP protein, with a slight trend toward an increased percentage when using 20 and 40 μg/ml (Supplemental Fig. 1B). In addition, we tested whether increasing the frequency of effector SMARTA cells would alter the AIM assay efficacy. To do this, we adoptively transferred varying numbers (200, 2,000, or 20,000) of naive SMARTA cells prior to LCMV infection. Seven days post–LCMV Armstrong infection, we observed that despite the large difference in the frequency of CD4⁺ T cells that were effector SMARTA cells in the AIM assay culture (Supplemental Fig. 1C, left), the percent AIM⁺ of SMARTA cells was similarly low in all three experimental groups (Supplemental Fig. 1C, right).

We analyzed our data to assess whether using either OX40 or CD25 as a single marker to identify AIM⁺ cells, instead of combined positivity for both OX40 and CD25, could mark a higher frequency of SMARTA cells. We assessed the frequency of marker-positive effector and memory SMARTA cells (OX40⁺, CD25⁺, or OX40⁺CD25⁺) in cultured cells following no stimulation or GP protein stimulation (Supplemental Fig. 2). Interestingly, following culture with no stimulation of day 7 effector cells, the frequency of OX40⁺ and CD25⁺ single-positive cells was significantly higher than the frequency of OX40⁺CD25⁺ (double-positive), suggesting that the combined use of both markers aids in reducing the “background” of marker-positive cells when cultured in the absence of Ag (Supplemental Fig. 2A). In contrast, the background of AIM-positive cells (whether using single or double markers) appeared to be lower at memory time points (Supplemental Fig. 2B, 2D) compared to the day 7 effector time points (Supplemental Fig. 2A, 2C). In general, once the background of marker-positive cells was subtracted out, the calculated percent AIM⁺ of SMARTA cells was significantly higher for OX40⁺ cells compared with OX40⁺CD25⁺ cells for both effector and memory cells induced by LCMV infection (Supplemental Fig. 2A, 2B), but not for cells induced by protein immunization (Supplemental Fig. 2C, 2D). These findings suggest that both single- and double-marker strategies for identifying murine Ag-specific CD4⁺ T cells may have differing strengths and disadvantages.

The AIM assay was designed as an alternative method to ICS protocols to quantify Ag-specific CD4⁺ T cells that were poor cytokine secretors, including CXCR5-expressing Tfh cells (4). To test whether the AIM assay culture conditions alter the frequency and expression of Ag-specific CXCR5⁺Bcl6⁺ germinal center (GC) Tfh or CXCR5^negative non-Tfh populations, we analyzed day 7 SMARTA CD4⁺ T cells from spleens of LCMV Armstrong–infected recipient mice or from dLN of GP⁺ AddaVax-immunized mice. Culturing day 7 SMARTA CD4⁺ T cells with or without LCMV GP for 18 h at 37°C with 5% CO₂ significantly decreased the frequency of infection-induced GC Tfh cells compared with ex vivo control cells by ∼2-fold (Fig. 3A, 3B). Although there was no significant difference in the frequency of immunization-induced GC Tfh cells, culturing day 7 SMARTA CD4⁺ T cells did significantly result in reduced expression of the transcription factor Bcl6 in CXCR5⁺ Tfh cells by ∼2-fold, regardless of stimulation with GP (Fig. 3A, 3C). However, the same reduction in Bcl6 expression was not observed in infection-induced CXCR5⁺ Tfh cells, although there was a significant increase in both cultured conditions (Fig. 3C). As the AIM assay culturing conditions significantly altered either the frequency of Ag-specific GC Tfh cells or the expression of Bcl6 in CXCR5⁺ Tfh cells, these data suggest that the AIM assay may underreport the quantity of murine Ag-specific GC Tfh cells.

To determine whether the AIM assay is biased in identifying non-Tfh compared with Tfh cell populations, we analyzed day 7 SMARTA CD4⁺CXCR5^negative non-Tfh cells and SMARTA CD4⁺CXCR5⁺ Tfh cells postinfection with LCMV Armstrong or postimmunization with rGP in AddaVax adjuvant. In LCMV infection–induced SMARTA CD4⁺ T cells, the AIM assay using rGP to restimulate cells only identified ∼10% of CXCR5⁺ Tfh cells as compared with ∼30% of non-Tfh cells, indicating a significant bias toward non-Tfh cells (Fig. 3D, 3E). In contrast to culturing with GP protein, GP_61-80 peptide restimulation resulted in efficient AIM marker upregulation by both CXCR5⁺ and CXCR5^negative effector SMARTA cell subsets that were induced by viral infection (Supplemental Fig. 3A, 3B). In immunization-induced SMARTA CD4⁺ T cells, the AIM assay following culture with rGP did not result in a bias for identifying CXCR5⁺ versus CXCR5^negative cells (Fig. 3D, 3E). Together, these data show that when using recombinant protein stimulation, the AIM assay is suboptimal for identifying infection-induced Ag-specific CXCR5⁺ Tfh cells compared with non-Tfh cells and that AIM assay culture conditions may alter expression of key transcription factors, including Bcl6.

To validate the sensitivity of the AIM assay for the identification of virus-specific cells in polyclonal CD4⁺ T cells, Foxp3 reporter mice (which carry a GFP reporter gene that is used to identify GFP⁺ Treg cells) (21) were infected i.p. with 2 × 10⁵ PFU of LCMV Armstrong. For analyses of Ag-specific CD4⁺ T cells, spleens from infected mice were collected at 7 d postinfection. In this study, we wanted to compare the AIM assay to ICS and tetramer staining for its ability to detect infection-induced Ag specific CD4⁺ T cells in a polyclonal response. Although the AIM assay identified ∼1–4% of splenic CD4⁺ T cells as GP-specific following GP–protein stimulation, IAb:GP_66-77 tetramer staining and ICS assay identify between 5 and 11% of CD4⁺ cells from the spleens of the same mice (Fig. 4A–D). Together, these data suggest that the AIM assay is less sensitive for the detection of murine virus-specific effector CD4⁺ T cells compared with other Ag-specific T cell assays.

To determine whether OX40⁺CD25⁺AIM⁺ cells are Ag-specific and to measure their TCR affinity for MHC/peptide Ag, we sorted AIM⁺ (OX40⁺CD25⁺) and AIM⁻ (OX40⁻CD25⁻) CD44^hiFoxp3-negative (gated as GFP⁻ to exclude Treg cells) day 7 splenic effector cells from LCMV-infected Foxp3 GFP reporter mice (Fig. 4E). Sorted cells were then tested for binding frequency and affinity using 2D-MP assay. Approximately 90% of AIM⁺ sorted cells were binders to IAb:GP_66-77 Ag (Fig. 4F). In contrast, ∼45% of AIM-negative sorted cells also exhibited binding to IAb:GP_66-77 Ag (Fig. 4F). Furthermore, AIM⁺ T cells, although having a large percentage of high-affinity cells in comparison with AIM⁻ T cells, also include some low-affinity GP_66-77-specific T cells (Fig. 4G). Each cell was tested for binding to IAb:GP_66-77 monomer and an irrelevant H2Db:NP_366-374 monomer to demonstrate Ag specificity (Fig. 4H). Interestingly, the low-affinity AIM⁺ population of cells (Fig. 4G) would have been missed by tetramer staining (26), yet are able to respond to Ag to upregulate AIM markers. Although inclusive of some low-affinity T cells, the AIM⁺ population had a higher geometric mean affinity than AIM⁻ cells. Furthermore, the range of affinity for both AIM⁺ and AIM⁻ cells spans ∼10,000-fold (Fig. 4G). We have previously shown that 8 d post–LCMV Armstrong infection, ∼50% of sorted CD4⁺CD62L⁻ T cells are specific for GP_66-77 by 2D-MP, with an affinity spanning >1000-fold (including both high- and low-affinity cells), whereas in contrast sorted IAb:GP_66-77 tetramer-positive cells yields only high-affinity cells by 2D-MP (26). In this study, we show that the AIM assay, although skewing toward the identification of high-affinity virus-specific effector CD4⁺ T cells, marks both high- and low-affinity cells and therefore encompasses an expanded profile of Ag-reactive T cells.

Chronic LCMV infection impairs the immune response by causing T cells to become functionally exhausted under conditions of persistent Ag stimulation (29–31). We hypothesize that the T cell dysfunction that occurs during chronic infection will include impairment of cells to upregulate AIM assay markers following Ag restimulation. To compare the sensitivity of the AIM assay in identifying Ag-specific cells in chronic versus acute viral infection, we adoptively transferred naive SMARTA (CD45.1⁺) cells into recipient C57BL/6 mice and then infected either i.p. with 2 × 10⁵ PFU of LCMV Armstrong (acute strain) or i.v. with 2 × 10⁶ PFU of LCMV clone 13 (chronic strain). Splenocytes from 7 and 30 d postinfection were either stimulated with rGP protein or GP_61-80 peptide. As predicted, the AIM assay marked a significantly lower percent of SMARTA cells as AIM⁺ at both days 7 and 30 in LCMV clone 13–infected mice compared with mice infected with the acute LCMV Armstrong strain (Fig. 5A, 5B).

We hypothesized that cell-intrinsic T cell dysfunction induced by chronic LCMV clone 13 infection decreased the sensitivity of virus-specific CD4⁺ T cells to the AIM assay. To test this, we purified CD4⁺ T cells from spleens of day 30 acute and chronically infected mice and cultured them for 18 h for the AIM assay by mixing them with enriched APCs from either naive or chronically infected (day 30) mice. SMARTA cells from chronic infection had lower induced AIM marker expression compared with SMARTA cells from acute infection, whether they were stimulated with GP protein (Fig. 5C) or GP_61-80 peptide (Fig. 5D). Interestingly, when cultured with naive APCs, SMARTA cells from acute infection have significantly higher frequencies of AIM⁺ cells than when cultured with chronic APCs, suggesting impaired Ag presentation in APCs from chronically infected mice (Fig. 5C). In line with this idea, the use of GP_61-80 peptide resulted in significantly improved AIM⁺ SMARTA cells from acute infection when cultured with APCs from chronic viral infection (Fig. 5E); however, the use of peptide Ag did not improve AIM marker responsiveness in SMARTA cells from chronic infection (Fig. 5E). Together, these findings indicate that virus-specific CD4⁺ T cells from chronically infected mice are dysfunctional in their responsiveness to upregulate AIM markers following restimulation. In addition, APCs from chronically infected mice are suboptimal in their capacity to present Ags in the AIM assay; however, this defect can be overcome by using exogenous peptide as the stimulating Ag.

To evaluate the AIM assay in an autoimmunity setting, we tested the AIM assay on lymphocytes from iLN and pLN and islets from BDC2.5 TCR-transgenic mice. Lymphocytes were stimulated with plate-bound 2.5HIP tetramer or remained unstimulated during 18-h culture. Compared to ex vivo cells, unstimulated AIM-cultured cells exhibited some upregulation of CD25, whereas stimulation with plate-bound 2.5HIP tetramer resulted in upregulation of both OX40 and CD25 (Fig. 6A). Subtracting nonspecific OX40/CD25 staining, AIM⁺ cells accounted for ∼25% of CD44^hi cells from pancreatic islets, whereas iLN and pLN exhibited significantly lower frequencies (Fig. 6B). We evaluated whether other AIM markers could provide improved identification of CD44^hi BDC2.5 cells from islets and found that Ag-specific AIM upregulation of CD154/CD40L marked ∼60% of these cells (Fig. 6C, 6D). These results indicate that the AIM assay may be used to evaluate autoreactive T cells and that different AIM marker combinations may improve their detection.

Since its introduction in 2016 (4, 5) the AIM assay has been a crucial tool for immunologists to identify Ag-specific T cells for vaccine studies in humans and nonhuman primates (5, 8), including publications evaluating Ag-specific T cell responses to SARS-CoV-2 infection and immunization (10, 14, 16, 32). The AIM assay was developed based on the idea that CD4⁺ Tfh cells were particularly “stingy” producers of cytokines, making conventional methods such as ICS or ELISPOT unsuitable to detect them (4, 5). Recently, several murine studies have used the AIM assay for specific purposes (11, 17); however, the utility of this assay for evaluation of murine Ag-specific CD4⁺ T cells responding to vaccination, infection, and autoimmunity has not been studied. In this work, we use known TCR-transgenic mouse models, including LCMV-specific SMARTA, OVA-specific OT-II, and chromogranin-specific BDC2.5 TCR-transgenic CD4⁺ T cells, to verify the efficacy and identify strengths and weaknesses of the AIM assay under various types of T cell activation and differentiation in vivo. Our findings suggest that the AIM assay is capable of identifying the majority of immunogen-specific CD4⁺ T cells following protein immunization throughout different phases of the immune response, whereas viral infection–induced CD4⁺ T cells respond poorly to the AIM assay, particularly among effector CD4⁺ T cells and especially under conditions of chronic infection/Ag.

We observed that the AIM assay is relatively effective for identifying Ag-specific CD4⁺ T cells following adjuvanted protein immunization, marking the majority of SMARTA and OT-II cells following induced immune responses with their respective Ags. Across both SMARTA and OT-II TCR-transgenic models, the results were very similar, with the AIM assay marking the majority of TCR-transgenic cells following 18-h culture to restimulate with Ag. A previous study by Crotty and colleagues (11) demonstrated that ∼72% of day 8 effector SMARTA cells from mice immunized with KLH-GP61 in alhydrogel induced expression to become CD69⁺CD40L⁺AIM⁺ cells following 9 h of restimulation with GP_61-80 peptide. Their results are similar to ours, in which SMARTA TCR-transgenic cells induced by GP protein immunization induced ∼80% of SMARTA cells to be OX40⁺CD25⁺, following 18-h restimulation with either whole GP protein or GP_61-80 peptide. Our results demonstrate that for both OVA-specific OT-II cells and GP-specific SMARTA cells induced by protein immunization, the AIM assay was similarly effective for identifying Ag-specific cells whether whole protein or specific peptide was used for the AIM assay culture. This point is important when considering that in preclinical mouse models of immunization, specific immunodominant epitopes may not be well defined, and being able to quantify specific CD4 T cell responses against a whole protein Ag is still very beneficial.

In addition to vaccination studies, the AIM assay has been used to study human Ag-specific T cells responding to various types of infectious diseases, including group A Streptococcus (12), SARS-CoV-2 (10, 14, 16), hepatitis B virus, CMV (6), and HIV (6, 15, 33). Given the extensive differences that occur in various types of bacterial, viral, and parasitic infections, including Ag amount, Ag persistence, inflammatory cytokines, disease pathologies, differences in T cell lineages, etc., it is possible that T cells respond differently to AIM marker upregulation depending upon the functional state of these T cells in these various types of disease settings. Although the exact frequency of human Ag-specific T cells induced by infections that are effectively marked by the AIM assay is difficult to know, using murine TCR-transgenic systems combined with acute and chronic infection mouse models allows for a better understanding of whether viral infection and persistence alter the efficacy of this assay. Our results indicate that following acute infection, Ag-specific memory (day 60) cells respond relatively well to induce AIM markers, whereas day 7 effector cells have a relatively poor capacity for OX40/CD25 upregulation following AIM culture in the presence of whole-protein Ag restimulation. Interestingly, this defect was rescued by using GP_61-80 peptide for restimulation instead of whole protein during the AIM culture. Besides our use of SMARTA TCR-transgenic cells, we also analyzed polyclonal responses to acute LCMV using other standard assays, such as ICS and tetramer staining. Compared to both of these assays, the AIM assay identified significantly lower frequencies of CD4⁺ T cells in the spleen that were Ag-specific. 2D-MP assay of AIM⁺ and AIM⁻ sorted cells confirmed that AIM⁺ cells include sampling of the entire range of affinity for Ag, including both high-affinity and some low-affinity T cells. In addition, we observed that the 18-h AIM culture conditions that were used resulted in reduced frequencies of SMARTA cells that were CXCR5⁺Bcl6^hi GC Tfh, which was made evident when we directly compared AIM cultured cells with ice-packed ex vivo splenocytes from the same samples. In contrast, effector cells induced by protein immunization did not result in reduced detection of Tfh cells within the AIM⁺ cells, although there was an observed reduction in Bcl6 mean fluorescence intensity. Together, these findings suggest that the AIM assay is not as effective for identifying murine virus-specific effector CD4⁺ T cells induced by acute infection.

Our study identified that virus-specific CD4⁺ T cells from chronic viral infection possess cell-intrinsic defects in their ability to upregulate OX40/CD25 markers following AIM culture, whether they were restimulated with whole GP protein or GP_61-80 peptide. In addition, APCs from chronically infected mice exhibited Ag-presentation defects when presenting Ag derived from whole GP protein. Together, these findings suggest that the AIM assay may dramatically undercount the true number of Ag-specific cells and does not identify the chronically stimulated or dysfunctional cells from chronically infected mice. Although our experiments do not address the sensitivity/efficacy of the AIM assay under all conditions in human studies, our results can still be used for cautionary purposes when evaluating Ag-specific T cells that may have become dysfunctional through chronic Ag stimulation or tolerogenic mechanisms.

Our study focused on validating the sensitivity of the AIM assay for identifying murine Ag-specific CD4⁺ T cells using known TCR-transgenic systems for immunization, infection, and autoimmunity. Based on our findings using various murine experimental model systems, for experimental designs and interpretations that use the AIM assay, careful consideration should be taken for selecting AIM markers and experimental conditions that are suited to effectively identify Ag-specific CD4⁺ T cells. We emphasize that our study only evaluated the AIM assay in mouse studies and does not specifically address whether these same strengths and/or shortcomings apply to human and nonhuman primates. Furthermore, although we evaluated the AIM assay using several mouse TCR-transgenic models, additional differences in AIM assay sensitivity/efficacy may occur due to intrinsic TCR affinity differences that exist within polyclonal Ag-specific CD4⁺ T cell populations within the immune response. Given these considerations and limitations of our study and acknowledging that many exceptions may exist, our results generally indicate that the AIM assay has varying sensitivity during different immune responses, with the highest efficacy for detecting Ag-specific T cells in adjuvanted protein immunization, reduced efficacy in acute infection, and very poor sensitivity to detect Ag-specific CD4⁺ T cells responding to chronic infection. Especially for immunization studies, the AIM assay is incredibly useful for the detection of cells that have tightly regulated/limited cytokine production such as Tfh and Th nonpolarized cells (34–36). Thus, the AIM assay can be a particularly useful tool for basic and preclinical murine vaccine studies to measure the relative frequency of vaccine Ag-specific cells, particularly for immunogens in which no TCR-transgenic or MHC tetramer reagents are available. However, the AIM assay for detecting murine CD4⁺ T cell responses has some limitations that should be considered, particularly for immunological or pathological conditions in which T cell dysfunction may occur, that should be considered when designing experiments, interpreting data, or making conclusions.

The authors have no financial conflicts of interest.

Flow cytometry data collection for this publication was supported by the University of Utah Flow Cytometry Core Facility. We thank Dr. Carl Davis at Emory University for providing the 293A-sGP cell line and GP purification protocol. We also thank the National Institutes of Health Tetramer Core Facility (Emory University) for providing I-A^b GP66-77 and I-A^g7 2.5HIP tetramers.