Optimizing Assays to Find Rare Antigen-Specific T cells in Cryopreserved PBMCs

Background

Immunomonitoring of T cell immune responses spans many therapeutic areas, including infectious and autoimmune diseases, and it is especially important for vaccine research. Immunomonitoring can be a difficult task because methods and protocols vary so widely.

Several functional assays are commonly used to count antigen-specific CD8⁺ T cells, and the protocols for these assays vary a great deal between labs. This makes it hard to interpret data from multi-center clinical trials or compare results between laboratories.

To address these issues in immunomonitoring of clinical trials, the Association for Immunotherapy of Cancer (CIMT) formed a CIMT monitoring panel to standardize protocols for measuring T cell antigen immune responses.

Study Design

Thirteen centers from 6 European countries took part in this study. Each center received the same samples and was asked to count antigen-specific T cells and assess their function, using tetramer staining plus a functional assay of their choice. Common techniques included ELISPOT assays, HLA-multimer staining, and intracellular cytokine staining (ICS).

Pre-tested samples of peripheral blood mononuclear cells (PBMC), synthetic peptides, and PE-conjugated HLA-tetramers were sent to each center. PBMCs were isolated from HLA-typed healthy volunteers by Ficoll density gradient separation. Each sample was tested for T cell reactivity against CMV and influenza. All centers received an HLA-A negative control, as well as HLA-A positive samples combining CMV- and influenza-reactive PBMCs.

The study had 2 phases. In Phase I, all centers ran the assays using their own standard protocols. In Phase II, each center received optimized protocols based on the Phase I findings.

Phase I: Protocol Variability

For the tetramer-staining assay, labs could choose to stain samples with antibodies (Ab) for CD8⁺ alone, CD3⁺CD8⁺, or CD4⁺CD8⁺, using their preferred Ab clone, fluorescent dye, and Ab concentration.

For the functional assays, synthetic peptides were provided, and each group could choose the INF-γ ELISPOT assay, FACS-based intracellular INF-γ staining, or both, at an antigen concentration of their choice (ranging from 1-10 µg/ml). To reduce variability in FACS analysis, sample plots, gate settings, and quadrants were provided to all centers.

For tetramer-staining, centers reported the number of viable cells post-thawing, the cytometer model, and the number of lymphocytes and/or CD8⁺ cells analyzed. Results were reported as the percent of tetramer-positive cells among CD8⁺, CD3⁺CD8⁺, or CD4⁺ lymphocytes, depending on the antibody cocktail used. For the functional assays, each center reported the ELISPOT plate type, reagents and conditions, and number of cells tested.

Tetramer Staining Results

Tetramer results from Phase I showed that the number of CD8⁺ cells analyzed significantly affected the sensitivity of tetramer staining. When fewer than 30,000 CD8⁺ T cells were counted, only 70% of antigen-specific responses were detected. When more than 30,000 CD8⁺ cells were counted, detection rose to 89%. When antigen-specific T cells were present at high frequencies, the number of counted cells made no difference.

Interestingly, Ab clone, Ab concentration, and cytometer type did not cause any significant differences. So the main factor driving detection of antigen-specific T cells by tetramer staining is the number of CD8⁺ cells used. Based on this, Phase II recommended using at least 1×10⁶ PBMCs for this assay.

IFN-γ ELISPOT Results

Most groups chose the INF-γ ELISPOT as their functional assay, and results varied widely between centers.

Some centers included a resting phase of 2-20 hours after thawing the cells, which detected 73% of positive reactivity (spot-forming cells per seeded PBMC). Skipping the resting phase detected only 30% of positive cells.

Replicate reproducibility within a center was also affected by the number of replicates used: duplicates often failed the Student’s t-test, while triplicates were enough to reach statistical significance.

Adding allogeneic APCs to bind and present the synthetic peptides had a negative effect on detection (28% of responses detected vs. 58% without them). Looking at the number of cells seeded per well, wells with more than 4×10⁵ PBMC detected 71% of positive samples, compared to only 43% for wells with fewer cells. Again, when antigen-specific T cells were present at high frequencies, cell counts did not affect detection rates.

Based on these findings, Phase II set minimum requirements for the INF-γ ELISPOT protocol: (1) run triplicates for each test antigen, (2) avoid allogeneic APCs, (3) include a resting phase, and (4) use more than 4×10⁵ PBMCs per well.

Study Limitations

Notably, lab experience with these assays had no effect on performance — experienced labs did no better than labs that had just adopted the techniques. This underscores the value of standardized protocols for immunomonitoring assays.

The study did not address specific detection limits for the ELISPOT assays, variability between ELISPOT plate readers, or serum source effects on background and specificity. It also did not report whether live/dead cell stains were used in the tetramer assays, or how combining these stains might have affected assay sensitivity.

Conclusion

This study identified several factors that should generally be applied when running tetramer staining and INF-γ ELISPOT assays with cryopreserved PBMC samples. These protocol adjustments matter most when measuring antigen-specific T cell populations present at low frequencies.

Reference:

The CIMT-monitoring panel: a two-step approach to harmonize the enumeration of antigen-specific CD8+ T lymphocytes by structural and functional assays. Britten CM, Gouttefangeas C, Welters MJ, Pawelec G, Koch S, Ottensmeier C, Mander A, Walter S, Paschen A, Müller-Berghaus J, Haas I, Mackensen A, Køllgaard T, thor Straten P, Schmitt M, Giannopoulos K, Maier R, Veelken H, Bertinetti C, Konur A, Huber C, Stevanović S, Wölfel T, van der Burg SH. Cancer Immunol Immunother. 2008 Mar;57(3):289-302. Epub 2007 Aug 25.

Sanguine supplies research-grade human PBMCs for studies like this.