Here we describe a robust method of determining immune cell identity and purity through epigenetic signatures detected using quantitative PCR (qPCR). DNA demethylation at a specific locus serves as a unique identifier for a particular cell type and allows for identification of CD8+, regulatory, or Th17 T cells.
Immune cell subtype population frequencies can have a large effect on the efficacy of T cell therapies. Current methods, like flow cytometry, have specific sample requirements, high sample input, are low throughput, and are difficult to standardize, all of which are detrimental to characterization of cell therapy products during their development and manufacturing.
The assays described herein accurately identify and quantify immune cell types in a heterogeneous mixture of cells using isolated genomic DNA (gDNA). DNA methylation patterns are revealed through bisulfite conversion, a process in which unmethylated cytosines are converted to uracils. Unmethylated DNA regions are detected through qPCR amplification using primers targeting converted areas. One unique locus per assay is measured and serves as an accurate identifier for a specific cell type. The assays are robust and identify CD8+, regulatory, and Th17 T cells in a high throughput manner. These optimized assays can potentially be used for in-process and product release testing for cell therapy process.
The use of T cells in cellular immunotherapy has increased significantly over the last decade, especially with the advent of chimeric antigen receptor (CAR) T cell technology1. While T cell-based therapies have been met with great clinical success, they are heterogeneous and cell characteristics and identities can vary significantly between donors2. The heterogeneity of T cell populations can have major impacts on therapeutic efficacy and complicate conclusions from clinical trials. For this reason, it is crucial to understand T cell heterogeneity during product manufacturing and formulation to determine the optimal formulations of T cell immunotherapies.
In a broad sense, T cells can be divided into two groups: CD4+ helper T cells, which secrete immunomodulatory cytokines, and CD8+ cytotoxic T cells, which directly lyse cells presenting the cognate antigen on major histocompatibility complex (MHC) I molecules. CD4+ helper T cells can differentiate into a myriad of specific subsets that produce a unique set of cytokines. Some suggest that a defined ratio of CD4+ to CD8+ T cells in the final cell product maximize in vivo efficacy and persistence but could complicate cell manufacturing3. Regulatory T cells (Tregs), a subset of CD4+ T cells, are immunosuppressive and diminish the activation of immune cells. Regulatory T cells have been implicated in mediating tumor tolerance and, if present during CAR T cell manufacturing, can inhibit the antitumor immune response of CAR T cells4,5,6. Other CD4+ subsets such as T helper 17 (Th17) T cells promote tumor clearance and can be leveraged to increase the efficacy of T cell therapies. Thus, increasing Th17 CD4+ T cells is of special interest in solid tumor settings where increased production of interleukin 17 (IL-17) promotes antitumor immune response5,7,8,9. Understanding the importance of Th17 cells in tumor responses has prompted more investigation into strategies to generate these cells in vitro7,9. Therefore, methods for detecting these T cell subsets are crucial for optimizing T cell therapies and should be robust, easy, scalable, and reproducible.
Flow cytometry is the most common method to determine the identity of an immune cell. However, flow cytometry requires live, intact cells that must be analyzed on the same day they are harvested. For intracellular cytokine staining (ICS), protein secretion inhibition is required to retain cytokines within the T cells. However, different inhibitor compounds have differential effects on the secretion of specific cytokines, forcing users to create specialized cocktails for the detection of the cytokine of interest10,11. Additionally, the fidelity of flow cytometry relies on antibody clones that bind specifically and potently to their target. The use of different antibody clones can cause varying results and lead to imprecise conclusions, making the method difficult to standardize12. Further, analysis of data collected via flow cytometry, and thus conclusions drawn from the data, can vary greatly between users based on how gates are set13,14. For these reasons, flow cytometry is not ideal for precise quality control during cell therapy development.
The assessment of genomic methylation at specific gene loci is an alternative method of determining cell identity. The rationale for using DNA methylation to identify specific cell types has been described in multiple articles and relies on specific methylation patterns present in a given cell population15,16,17. In target cells, certain sites contain unmethylated nucleotides, while in non-target cells these sites are methylated. This pattern can be detected through bisulfite conversion, a process that exclusively converts unmethylated cytosine nucleotides (C) to uracil (U). Primers and probes can be designed against the converted sites, then amplified and detected through qPCR16.
The assay described herein measures the frequency of immune cell subtypes within heterogeneous populations by quantifying the number of specific unmethylated loci compared to a housekeeping gene. Copies of the unmethylated regulatory elements for the CD8 B gene are measured in the CD8 assay, FoxP3 in Treg, and IL-17 in Th17 in this assay. These loci were identified by isolating the specific cell population of interest (e.g., CD8+ T cells) and performing bisulfite sequencing to identify loci that are unmethylated in only that cell type15. Primers and probes are developed against the uniquely unmethylated loci and samples are analyzed via qPCR. A standard curve of known copy numbers is created from dilutions of a high copy number standard plasmid, allowing for the conversion of a Ct value to a transcript copy number.
To evaluate assay performance, control samples are required, including a reference gDNA and a calibrator plasmid. The reference genomic material is a pooled blood sample from multiple donors with known immune cell frequencies and is used for a quality control (QC) assay performance check. The calibrator sample is a synthesized plasmid that contains the CD8, FoxP3, and GAPDH gene sequences in an equimolar ratio. It is used as a measure of the bisulfite conversion efficiency, because bisulfite conversions within different genomic regions will vary in efficiency. The ratio of the target to GAPDH within the calibrator is 1, but due to differences of the bisulfite conversion efficiency, the ratio can vary. This calibration factor is applied to the CD8 and Treg assays. FoxP3, the gene used in the Treg assay, is located on the X chromosome. Because this assay measures only unmethylated copies of FoxP3, and only one copy of FoxP3 is unmethylated in Tregs, this assay is sex agnostic and can be used with both male and female samples18. The Th17 assay does not utilize a calibrator sample or GAPDH as a housekeeping gene. Instead, another primer set targeting the methylated loci present in non-Th17 cells is included and the total number of cells is reflected by the sum of the methylated and unmethylated copies of IL-17. The data obtained from the qPCR is entered into a preset analysis template that performs quality control checks on the data and calculates the percentage of the target cell within the starting population. This provides automated and unbiased data analysis, thereby removing user subjectivity and improving standardization capabilities.
This assay uses gDNA, which can be isolated by a leukapheresis procedure using cultured or fixed cells, or frozen cell pellets. The low sample requirement, flexibility in sample starting material, high accuracy, and standardized analysis address the limitations associated with flow cytometry and are ideal for quality control purposes during cell therapy process development.
All human samples were obtained from a commercial source following their guidelines for human research ethics.
1. Genomic DNA isolation
NOTE: Genomic DNA isolation is performed using a commercially available kit (see Table of Materials) from any hematopoietic cells, such as peripheral blood mononuclear cells (PBMCs), purified T cells, or any other hematopoietic cell type. In this experiment, peripheral blood mononuclear cells (PBMCs) and T cells from three different human donors were used.
2. Sample preparation
3. Bisulfite conversion
NOTE: Ensure that the incubation times during the bisulfite conversion follow the recommended times. Over-incubation or under-incubation will affect the results of the assay.
4. CD8 and Treg assays
Initial plasmid Copy Number/3 µL | Volume | Diluent DNA [1] | Final Copy Number per 3 µL | Label |
31250 | 1000 µL | - | 31250 | STD#1 |
31250 | 200 µL | 800 µL | 6250 | STD#2 |
6250 | 200 µL | 800 µL | 1250 | STD#3 |
1250 | 200 µL | 800 µL | 250 | STD#4 |
250 | 200 µL | 800 µL | 50 | STD#5 |
1250 | 30 µL | 1200 µL | 30 | STD#6 [2] |
[1] 10 ng/µL Lambda DNA in TE (10 mM Tris, 1 mM EDTA, pH 8.0) | ||||
[2] Use STD#3 to prepare STD#6 |
Table 1: Preparation of standard dilutions. A 4-log dilution standards spanning 31250-30 copies were prepared according to the table using TE/lambda DNA as the diluent. The same standard dilutions were used for both CD8 and GAPDH. The standards were stored at -20 °C.
Reagents | Amount |
Lambda DNA (50 ng/µL in TE, pH8.0) | 1 µL |
TaqMan assay | 0.5 µL |
Water, Nuclease-free | 0.5 µL |
Master Mix | 5 µL |
TOTAL | 7 µL |
Table 2: Preparation of qPCR cocktail. Prepare the qPCR master mix in two separate tubes, one each for CD8 and GAPDH, excluding the template DNA according to the table.
Step | Time | Temp | Cycles |
Pre-incubation | 35 min | 95°C | 1X |
Amplification | 15 sec | 95°C | 50X |
1 min | 61°C | ||
Cooldown | 5 sec | 42°C | 1X |
Table 3: qPCR run parameters for the CD8 and Treg assay. The qPCR was run according to the parameters specified in the table.
5. Th17 assay
Reagent | Amount |
Water, Nuclease-free | 9.5 µL |
Hot Start PCR Master Mix | 12.5 µL |
Th17 PCR Primer | 1 µL |
TOTAL | 23 µL |
Table 4: Pre-amplify the DNA for the Th17 assay. The pre-amplification reaction master mix was prepared according to the table.
Step | Time | Temp | Cycles |
Pre-incubation | 35 min | 95°C | 1X |
Denaturation | 1 min | 95°C | 12X |
Annealing | 45 sec | 55°C | |
Elongation | 30 sec | 72°C | |
Final elongation | 10 min | 72°C | 1X |
Hold | Indefinite | 4°C |
Table 5: Preamplify Th17 DNA. Th17 DNA was pre-amplified for 12 cycles before the actual qPCR.
All three methylation assays begin with an input of gDNA and result in a percentage of CD8+ T cells, Treg, or Th17 cells within the entire cell population. The data generated from the qPCR following bisulfite conversion was analyzed using the provided analysis template. This template used the standard curves obtained from the standard samples to estimate the copy number of the targets and total cell counts in the test samples. The percent cell type was then calculated using the formulas integrated in the analysis templates. Figure 1 shows representative data from the CD8+ assay, obtained from analyzing both peripheral blood mononuclear cells (PBMCs) and T cells from three different donors using both flow cytometry and the described methylation assay. The trends between the two methods across all donors in both cell types were similar. Figure 2 shows representative data from the Treg assay. Three purified Treg donors were analyzed using both methylation-based qPCR and flow cytometry and the results were plotted on the graph shown. Results for both methods were relatively similar. However, in all cases flow cytometry yielded higher values. Figure 3 shows the comparison of Th17 cells detected via flow cytometry and methylation. In this graph, there are stimulated and unstimulated conditions for the experimental methods. Cells require stimulation with PMA and ionomycin and treatment with a protein transport inhibitor to increase the amount of IL-17A present within each cell so that it can be detected via flow cytometry20. Methylation status can be detected regardless of the stimulation status. Flow cytometry yielded lower levels of Th17 cells when compared to methylation. The sensitivity and specificity of the assay was demonstrated by the limit of blank (LOB), limit of detection (LOD), and limit of quantitation (LOQ) values displayed in Table 6.
Figure 1: Flow and methylation comparison of CD8+ percentages for both PBMCs and T cells for three different donors. PBMCs and T cells from three different donors were assayed for the percentage of CD8+ T cells in the entire population using either flow cytometry or methylation. Please click here to view a larger version of this figure.
Figure 2: Flow and methylation comparison of Treg percentages for purified Treg for three different donors. Purified Treg cells were assayed for the percentage of Treg cells in the entire population using either flow cytometry or methylation. Please click here to view a larger version of this figure.
Figure 3: Flow and methylation comparison of Th17 percentages for stimulated and unstimulated experimental groups. Stimulated and unstimulated T cells were analyzed using both flow cytometry and methylation. Please click here to view a larger version of this figure.
Assay | LoB | LoD | LoQ |
CD8 | 0 | 19 | 52 |
Gapdh | 0 | 7 | 21 |
FoxP3 | 0 | 3 | 12 |
Th17 TpG | 0 | 35 | 73 |
Th17 CpG | 0 | 38 | 73 |
Table 6: LoB, LoD, LoQ of copy numbers for methylation-based assays. Sensitivity and specificity of the assay is detailed by the LoB, LoD, and LoQ values obtained by MIQE guidelines21,22. In general, as few as 40 copies can be accurately detected by these assays.
With the advent of new immunotherapeutics, there is a need for standardized methods of detecting immune cell identity and purity. Assessment methods for in-process and release testing that are robust, validated, and scalable remain to be established and pose a major challenge to the commercialization of cell-based therapies. While flow cytometry is currently the most common method for immune cell phenotyping, high sample quality and quantity requirements make it difficult for regular use. Further, implementing flow cytometry in a good manufacturing practice (GMP) environment is limited by operator-dependent gating strategies and the requirement of reference standards for each marker used13,14. Although automated gating has been shown to improve assay robustness, it is not yet a well-established quality control strategy. While gene expression profiling may also be used to characterize cell therapy product identity and purity, the data are semiquantitative and labor intensive. Additionally, RNA and microRNA are relatively less stable than DNA and can contribute to the lack of robust and reproducible results. Thus, utilizing the epigenetic DNA methylation status of a specific locus provides a stable, easy to perform, robust, and scalable method of identifying and quantifying a cell type of interest.
The detection of methylation patterns to phenotype cells relies on three critical steps: First, the assay requires the use of gDNA, which is isolated according to the described methods. High quality DNA is required and if not used, bisulfite conversion may be affected23,24. If low quality DNA is obtained, further purification is recommended to ensure the assay reliability. Second, successful bisulfite conversion of the unmethylated cytosine to uracil is required. The most critical step in bisulfite conversion is DNA denaturation23,25. High temperature denaturation using ammonium bisulfite, as opposed to sodium bisulfite, increases the conversion efficiency and consistency and is the recommended process for these assays25,26. Users must ensure that the reaction occurs at 80 °C and that sample mixing is done frequently. For these reasons, a digital thermomixer is recommended (see Table of Materials). Third, proper pipetting technique must be followed during the qPCR preparation. Three technical replicates are required during the qPCR reaction to adhere to the minimum information for publication of quantitative real-time PCR experiments (MIQE) guidelines27. The inclusion of more technical replicates is acceptable but not required. Improper pipetting technique and/or not including technical replicates will lead to unreliable qPCR results.
If the critical steps are followed and desired results are still not obtained, multiple controls within the assay can be leveraged to pinpoint the issue. Proper bisulfite conversion is the most crucial step in this assay. Improper bisulfite conversion would be highlighted by a failed calibration factor and/or reference values calculated by the analysis template. If the bisulfite conversion was performed incorrectly (e.g., if the reaction was carried out at RT), there would be no amplification during qPCR. Also, no amplification would be seen if an error was made during the qPCR reaction preparation (e.g., if the qPCR master mix was not added). These two issues can be separated by investigating the standard samples. Amplification in the standard samples, but not the reference, calibrator, and experimental samples, would indicate that the bisulfite conversion was not performed correctly. No amplification in any of the samples would indicate that the qPCR was not performed correctly.
While this assay addresses the stringent sample requirement and analysis variability associated with other phenotyping methods, specifically flow cytometry, there are limitations that must be noted. This assay targets a single locus that is used as a unique identifier for the target cell, which prohibits the multiplexed analysis that is commonly performed with flow cytometry. This makes identifying complex cell types like Th17 difficult. However, the use of methylation patterns at a single locus has been shown to be an accurate phenotypic marker of multiple cells and has been used in multiple clinical trials15,16. This is true especially in Tregs where assessing FOXP3 methylation signatures is an accurate and unambiguous method for detecting true Tregs from transiently FOXP3 expressing cells28. The loss of multiplexed analysis is compensated by the accuracy of the loci interrogated. Future iterations of the assay could include multiple qPCR dyes and quenchers to allow for the detection of multiple loci in one qPCR reaction.
This assay can be used as an alternative method for flow cytometry in determining cell phenotype and identity. It should be noted that this assay does not yield the exact values that flow cytometry does (Figures 1-3). This is due in part to the variability of data analysis associated with flow cytometry. Depending on the gating strategy, results from flow cytometry can vary significantly. This is especially the case when using complex staining protocols to look at intracellular targets, such as IL-17, where the coefficient of variation (CV) can be as high as 15%14. Between multiple users, the assay presented consistently had a CV of <15%, supporting the robustness of the assay and improved standardization capabilities. The largest differences between epigenetic and flow cytometry-based phenotyping is seen when cell stimulation is required (Figure 3). The detection of Th17 cells via flow cytometry was accomplished using a common protocol for T cell stimulation and intracellular cytokine staining29,30,31. The differences seen in Th17 phenotyping between epigenetic measurement and flow cytometry could be due to the kinetics of IL-17 production. While methylation signatures are steady, protein production takes time and must be present in sufficient quantities to be detected by fluorescent antibodies20,32. The 6 h incubation with protein transport inhibitor and cell stimulation cocktail may need to be extended to see the level of Th17 cells detected via epigenetic-based phenotyping methods. More studies are needed to determine the precise reason why the values do not match and determine the most accurate phenotyping method.
In this report, we detail how to identify and quantify immune cell types in a heterogeneous mixture of cells in a simple and robust manner. The assays are designed and optimized for potential use for in-process and release testing of cell-based therapeutics. The described assays meet the requirement for highly qualified raw materials to be used in cell therapy applications. By addressing the shortcomings of flow cytometry and other molecular methods such as stability, sample requirements, prior stimulation, cellular permeabilization for intracellular staining, and subjectivity of data analysis, the described assays are in line with the goals of commercialization of cell-based therapies.
The project was funded by Thermo Fisher Scientific Intramural grant.
Name | Company | Catalog Number | Comments |
2.0 ml Eppendorf Safe-Lock Tubes | Eppendorf | 22363344 | |
Analysis template for CD8 assay | Click Here | ||
Analysis template for Th17 assay | Click Here | ||
Analysis template for Treg assay | Click Here | ||
Attune NxT Acoustic Focusing Cytometer, blue/red/violet6/yellow | Thermo Fisher Scientific | A29004 | |
CTS PureQuant CD8+ T-Cell Assay | Thermo Fisher Scientific | A43674 | |
CTS PureQuant Th17 Assay | Thermo Fisher Scientific | A43676 | |
CTS PureQuant Treg Assay | Thermo Fisher Scientific | A43675 | |
Dynabeads SILANE Genomic DNA Kit | Thermo Fisher Scientific | 37012D | Used for genomic DNA isolation and purification after bisulfite conversion. Contains elution buffer to dilute samples. |
DynaMag-2 Magnet | Thermo Fisher Scientific | 12321D | Used as a magnetic rack during DNA isolation and purification |
eBioscience Essential Human T cell Phenotyping Kit | Thermo Fisher Scientific | A42923 | |
eBioscience Essential Human Th1/Th17 Phenotyping Kit | Thermo Fisher Scientific | A42927 | |
eBioscience Essential Human Treg Phenotyping Kit | Thermo Fisher Scientific | A42925 | |
Eppendorf SmartBlock 2 mL | Eppendorf | 5362000035 | |
Eppendorf ThermoMixer C | Eppendorf | 5382000023 | |
HulaMixer Sample Mixer | Thermo Fisher Scientific | 15920D | Recommended sample mixer |
NanoDrop | Thermo Fisher Scientific | ND 1000 | |
Nonstick, RNase-free Microfuge Tubes, 1.5 mL | Applied Biosystems | AM12450 | |
QuantStudio 12S Flex | Applied Biosystems | 4470661 | |
TE, pH 8.0, RNase-free | Thermo Fisher Scientific | AM9849 | |
Vortex Mixer | Thermo Fisher Scientific | H2KT17113 |
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