Flow Cytometric Clinical Immunomonitoring Using Peptide-MHC Class II Tetramers: Optimization of Methods and Protocol Development

Abstract

With the advent of novel strategies to induce tolerance in autoimmune and autoimmune-like conditions, clinical trials of antigen-specific tolerizing immunotherapy have become a reality. Besides safety, it will be essential to gather mechanistic data on responding CD4⁺ T cells to assess the effects of various immunomodulatory approaches in early-phase trials. Peptide-MHC class II (pMHCII) multimers are an ideal tool for monitoring antigen-specific CD4⁺ T cell responses in unmanipulated cells directly ex vivo. Various protocols have been published but there are reagent and assay limitations across laboratories that could hinder their global application to immune monitoring. In this methodological analysis, we compare protocols and test available reagents to identify sources of variability and to determine the limitations of the tetramer binding assay. We describe a robust pMHCII flow cytometry-based assay to quantify and phenotype antigen-specific CD4⁺ T cells directly ex vivo from frozen peripheral blood mononuclear cell samples, which we suggest should be tested across various laboratories to standardize immune-monitoring results.

Keywords: antigen-specific T cells; flow cytometry; protocol standardization; tetramerization; tetramers.

Figure 1

Shorter staining protocol resulted in higher numbers of tetramer-positive CD4⁺ T cells. Three staining protocols were compared as indicated. (A) The number of HLA-DRB1*0101 HA_306–318-positive CD4⁺ T cells per million CD4⁺ T cells is depicted for the three protocols in peripheral blood mononuclear cells of one representative individual. (B) Flow cytometry plots for two patients stained with HLA-DRB1*0101-collagen II_259–273, using either the short or the long protocol. The negative control FMO plots are shown. (C) The number of HLA-DRB1*0101-collagen II_259–273⁺ CD4⁺ T cells per million CD4⁺ T cells using the short versus the long staining protocol in Donors A and B, as depicted in panel (B).

Figure 2

The streptavidin–biotin ratio is important for optimal tetramer formation. Research tetramers were compared with tetramers purchased from MBL and ProImmune (PI). (A) Flow cytometry plots of a representative donor depicting FMO-PE and staining with HLA-DRB1*0401-collagen II_259–273 tetramers using the short staining protocol (representative of three HLA-DRB1*04:01⁺ and three HLA-DRB1*01:01⁺ donors with rheumatoid arthritis). (B) The number of tetramer-positive cells per million CD4⁺ T cells identified using HLA-DRB1*0401-collagen II_259–273 and HLA-DRB1*0101-collagen II_259–273 tetramers from two sources (n = 3 replicates per donor), staining at 4°C. (C) Flow cytometry dot plots depicting the number of tetramer-positive cells using tetramers generated from monomers as depicted, from two sources using the same formula to calculate the streptavidin–biotin ratio and each stained at a concentration of 4.2 µg/ml (representative of three individual donors). In panels (A,B), staining with research and MBL tetramers was carried out at 4 and 37°C and with PI tetramers at 37°C. In panel (C), all tetramers were stained at 4°C.

Figure 3

Gating strategy for tetramer analysis. Live cells were gated using forward and side scatter. Subsequently, single cells were gated using forward scatter height and area followed by side scatter height and area (upper panels). Live CD3⁺ cells were gated using Lineage/LIVE/DEAD marker negative and CD3-positive cells. Next, CD4⁺ T cells were gated by gating on the CD3⁺ CD4⁺ double-positive cells. The FMO sample was used to determine the tetramer-positive gate. The gate was set on the CD4-postive cells that are negative in the tetramer channel (middle panels). This gate was used to identify the CD4⁺ tetramer⁺ cells in the rest of the samples (bottom panels). Representative of 40 experiments.