Optimizing whole blood stimulation conditions for intracellular cytokine staining: A solution for resource-limited laboratories and field studies
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Optimizing whole blood stimulation conditions for intracellular cytokine staining: A solution for resource-limited laboratories and field studies
Gnodja Tégmaba1,2*, Kadanga Motomdéwa Monique1, Bara Fagdéba David1, Simfele Hombamane Christelle1, Kolou Maléwé3, Mbengue Babacar2
1Unité de Recherche en Immunologie et Immunomodulation (UR2IM) /Laboratoire de Microbiologie et de Contrôle de Qualité des Denrées Alimentaires (LAMICODA)/Ecole Supérieure des Techniques Biologiques et Alimentaires (ESTBA), Université de Lomé, Lomé-Togo
2Service d’Immunologie, Faculté de Médecine, de Pharmacie et d’Odontologie (FMPOS), Université Cheikh Anta Diop (UCAD), Dakar-Sénégal
3Faculté des Sciences de la Santé (FSS), Université de Lomé, Lomé-Togo
Abstract
Introduction: Intracellular cytokine staining (ICS) combined with flow cytometry is a preferred method for evaluating and characterizing cell-mediated immune responses. While peripheral blood mononuclear cells (PBMCs) are commonly used in immunological studies, their isolation poses challenges in remote or resource-limited settings. This study aimed to establish optimal conditions for whole blood stimulation to assess cytokine-producing T cells, specifically interferon gamma (IFN-γ), interleukin-17A (IL-17A), and interleukin-10 (IL-10).
Methods: Eight healthy male donors, aged 28–35 years, participated in this study. Of these, 3 donors were included to assess CD4+Tbet+IFNγ+, CD4+GATA-3+IL-4+, CD4+RORgt+IL-17A+, and CD4+FOXP-3+IL-10+cells using whole blood samples and PBMCs from the same donors and fives were recruited to investigate only the effects of varying whole blood samples volumes and different container types assessing only CD4+Tbet+IFNγ+. Cells were identified through flow cytometry after stimulation of PBMCs and whole blood samples using phorbol 12-myristate 13-acetate (PMA) and ionomycin.
Results: This study has two main limitations including the very small sample size and the cell culture plate which did not allow stimulation at 200 µL of whole blood. However, the results remain debatable with the use of Falcon tubes. The percentage of CD4⁺Tbet⁺IFN-γ⁺ cells did not differ significantly when a cell culture plate was compared to a Falcon®-type tube using whole blood sample. However, cytokine response was significantly higher when 100 µL of whole blood was used in cell culture plates compared to 200 µL in Falcon®-type tubes.
Conclusion: This study suggests that 100 µL of whole blood is a practical alternative to PBMCs, with 96-well plates serving as the optimal container for T-cell stimulation in resource-limited settings.
Keywords
Whole blood sample, Intracellular cytokine staining, Flow cytometry, T cells
Introduction
T cells play an essential role in the specific immune response. These lymphocytes, derived from the thymus, can be further categorized into subpopulations based on their phenotypic markers and the cytokines they express, which reflect their functional roles (1). Numerous diagnostic methods are designed to study T cells as well as the pathologies associated to their dysfunctions. Among the various methods available for analyzing T-lymphocyte functions, intracellular cytokine staining (ICS) by flow cytometry is considered as the gold standard (2). This approach not only detects cytokine expression at the single-cell level but also enables the simultaneous detection, quantification, and phenotypic characterization of T cells (3). ICS can be performed on Peripheral Blood Mononuclear Cells (PBMCs) (4) or directly on Whole Blood (WB) (5).
PBMCs are typically isolated from whole blood using Ficoll density gradient centrifugation allowing for stimulation, staining and immediate analysis or cryopreservation. However, the isolation process is labor-intensive, time-consuming, and expensive, involving lengthy centrifugation steps that can be challenging in settings with unreliable electricity. These constraints make PBMC isolation less practical in low-income countries. Additionally, PBMC isolation often requires large blood volumes, which may not be ethically acceptable or culturally appropriate in certain populations, such as pediatric patients or pregnant women (6-9).
In contrast, whole blood offers a more practical alternative. It retains all host serum factors and cellular components, providing a more physiological environment for stimulation by maintaining cell-to-cell communication (10). Whole blood procedures require minimal material, fewer preprocessing steps, and only small blood volumes, which is advantageous in resource-limited settings. However, a key challenge with whole blood lies in determining the number of cells available for stimulation, as this may vary between individuals and depends on the blood volume used. Although, the use of whole blood sample does not solve the electricity problem in low-income countries, it does solve the budgetary constraints, ethical and societal problems related to sample collection.
Recent studies have highlighted the potential of whole blood for analyzing cellular functionality using ICS and flow cytometry. Nevertheless, critical aspects such as the optimal blood volume and type of cell culture vessel for studying cytokine-producing T cells remain poorly understood.
This study aims to optimize whole blood stimulation conditions for intracellular cytokine staining, focusing on cytokine-producing T cells. We investigated the expression of key cytokines, including interferon gamma (IFN-γ),
interleukin-4 (IL-4), IL-17A, IL-10 in CD4+ T lymphocytes. Furthermore, we assessed the use of 15 ml Falcon-type tubes and 96-well U-bottom plates to establish optimal stimulation conditions using varying volumes of whole blood samples.
Methods
Study population
This descriptive, cross-sectional study was conducted in two stages with a total of eight (08) male healthy donors aged 28 – 35 years recruited at the blood collection and distribution point of Centre Hospitalier Universitaire Sylvanus Olympio (CHU SO). Firstly, three healthy donors were included to assess cytokines producing T cells using whole blood samples and PBMCs from the same donors. Secondly, fives donors were recruited to investigate only the effects of varying whole blood volumes and different container types. Venous blood samples were collected from each participant in ethylene diamine tetra acetic acid (EDTA) tubes and immediately transported to the Unité de Recherche en Immunologie et Immuno-modulation (UR2IM) /Laboratoire de Microbiologie et de Contrôle Qualité des Denrées Alimentaires (LAMICODA)/ Université de Lomé, Togo for cellular analyses. All protocols were started within the next two hours after blood collection on the same day.
Immunophenotyping of T cell subpopulations
Peripheral Blood Mononuclear Cells (PBMCs) isolation and stimulation: PBMCs from fresh whole blood samples were isolated using the Ficoll Paque (PAN-Biotech) density gradient centrifugation method on ice as described by Simfele et al. (11). The cells were suspended in culture medium (Roswell Park Memorial Institute 1640 supplemented with gentamicin 50 μg/mL, penicillin streptomycin 100 μg/mL, L-glutamine 2 mM/mL (RPMI+++), and fetal bovine serum (FBS) at 10% (PAN-Biotech) at the concentration of 2×105 cells/µL and then the cells were counted and their viability assessed by Trypan blue exclusion method. A One hundred microliters (100 µL) of PBMCs suspension per well was stimulated in duplicate as described by Tchopba et al. (12) using 96- wells U-bottom plates (Greiner Bio One, Frickenhausen, Germany) with 50 µL of 1X cell stimulation cocktail (phorbol 12-myristate 13-acetate (PMA), Ionomycin, Brefeldin A and Monensin; eBioscience), and incubated at 37°C in presence of 5% CO2 for 6 hours. Thereafter, cells were harvested and wash before staining.
Whole blood stimulation: Fresh whole blood was cultured in RPMI complete medium (RPMI+++ and fetal bovine serum (FBS) at 10%), in 96- wells U-bottom plates (Greiner Bio One, Frickenhausen, Germany) and 15 ml polypropylene conical bottom (Greiner Bio One, Frickenhausen, Germany) under sterile conditions. Different volumes of whole blood were tested in 15 mL Falcon® tubes to determine the optimal stimulation volume. Equal volumes of EDTA-treated whole blood were mixed with RPMI-1640 medium and 1X cell stimulation cocktail in duplicate as follow: for 100 µL blood samples we add 100 µL of 1X cell stimulation cocktail with RPMI-1640 medium and for 200 µL blood samples, 200 µL of 1X cell stimulation cocktail with RPMI-1640 medium were added. All tubes incubations were conducted as described above. After stimulation, red blood cells were lysed from stimulated blood samples using RBC Lysis Buffer (Roche® Diagnostics GmbH, Mannheim, Germany) according to the manufacturer’s instructions. After that, cells were stained under the same conditions with PBMCs.
Cells staining and gating strategy: To assess cytokine expression in CD4⁺T cells, cell-surface staining was performed using APC-conjugated anti-CD4 (clone A161A1) antibodies (1:50) and incubated at 4°C for 30 minutes. Intracellular staining followed, using Fix-Perm (eBioscience) reagent for fixation and permeabilization and Fc block (human TruStain FcX®, BioLegend, San diego, CA, USA) to prevent non-specific binding. Four staining panels were used: 1. Anti-human T-bet-PE (clone 4B10) and Anti-human IFN-γ-FITC (clone 4S.B3); 2. Anti-human GATA3-PE (clone 16E10A23) and Anti-human IL-4-FITC (clone MP4-25D2); 3. Anti-human RORγt-PE (clone AFKJS-9) and Anti-human IL-17A-FITC (clone BL168); 4. Anti-human FoxP3-FITC (clone 206D) and Anti-human IL-10-PE (clone JES3-9D7). Samples were washed, resuspended in fix-perm buffer, and analyzed using a flow cytometer (Cytoflex, Beckman Coulter, Brea, California, USA). Data were acquired using CytExpert 2.1 software (Cytoflex, Beckman Coulter, Brea, California, USA), and gating strategies were used to identify CD4⁺ T-cell subpopulations from PBMCs (Fig. 1) and whole blood samples (Fig. 2). Unstained and single-stained cells served as negative and positive controls, respectively. All antibodies were obtained from BioLegend (BioLegend®CA, USA).
Fig. 1: Gating strategy for assessing Cytokine producing T-cells from PBMCs. Lymphocytes gate (total T cells) were set based on cells granularity and size defined by Forward scatter (FSC) and side scatter (SSC) (1A) then from them, we identified CD4+T-cells (1B) which were then separated into different subpopulation CD4+Tbet+IFNγ+ (1C), CD4+GATA-3+IL-4+(1D), CD4+RORγt+IL-17A+ (1E), and CD4+FOXP-3+IL-10+ (1F).
Fig. 2: Gating strategy for assessing Cytokine producing T-cells from whole blood. Lymphocytes gate (total T cells) were set based on cells granularity and size defined by Forward scatter (FSC) and side scatter (2A) then from them, we identified CD4+T-cells (2B) which were then separated into different subpopulation CD4+Tbet+IFNγ+ (2C), CD4+GATA-3+IL-4+ (2D), CD4+RORγt+IL-17A+ (2E), and CD4+FOXP-3+IL-10+ (2F).
Statistical analysis
Data were compiled in Excel and analyzed using GraphPad Prism version 5.02 (GraphPad Software, La Jolla, CA, USA). Differences between groups were assessed using non-parametric Mann-Whitney U tests with statistical significance set at p < 0.05.
Ethics statement
Ethical approval for this study was obtained from the bioethics committee for health research of Togo (“Comité de Bioéthique pour la Recherche en Santé”, CBRS) under approval number 039/2024/CBRS. Written informed consent was obtained from all participants prior to sample collection.
Results
Comparison of cytokines-producing T cells between whole blood and PBMCs
We compared the frequencies of cytokine-producing T cells, including CD4+Tbet+IFNγ +(Th1), CD4+GATA3+IL-4+ (Th2), CD4+RORγt+IL-17A+ (Th17) and CD4+FoxP3+IL-10+ (Treg), between whole blood and PBMCs as shown in Fig 2 and summarized in Table 1. Our findings revealed that Th1 cells were significantly more abundant in PBMCs compared to whole blood (Fig. 3A). In contrast, no significant differences were observed for Th2 (Fig. 3B), Th17 (Fig. 3C), or Treg cells (Fig. 3D), Although, all these T cells subsets were higher in PBMCs than in whole blood. Additionally, the standard deviation (SD) was notably larger for PBMCs than for whole blood, indicating greater variability in cell frequencies between individuals when PBMCs were used.
Fig. 3: Percentage of T cells cytokines-expression in whole blood samples and PBMCs (n=3) stimulated with PMA, Ionomycin, Brefeldin A and Monensin. *, p < 0.05; ns, non-significant. The Mann-Whitney U test was used to compare differences among groups. A: Th1 (CD4⁺Tbet⁺IFNγ⁺); B: Th2 (CD4⁺GATA3⁺IL-4⁺); C: Th17 (CD4⁺RORγt⁺IL-17A⁺); D: Treg (CD4⁺FoxP3⁺IL-10⁺). Scatter plots show the mean and standard error of the mean (SEM). Mann–Whitney U test was used for statistical comparisons. Each dot represents an individual data point, and the horizontal black lines represent the mean
Table I: Variability in cytokine-producing T cells between whole blood and PBMCs, expressed as mean. Standard deviation (SD) reflects the variability around the mean
| Cell type | Whole Blood (n=3) | PBMCs (n=3) | p-value |
| Mean (SD) | Mean (SD) | ||
| CD4+Tbet+IFNγ+ | 9.85 (4.67) | 48.37 (21.96) | 0.0500 |
| CD4+GATA3+IL-4+ | 2.87 (1.98) | 41.66 (37.45) | 0.1000 |
| CD4+RORγt+IL-17A+ | 5.49 (1.03) | 17.88 (12.02) | 0.1000 |
| CD4+FoxP3+IL-10+ | 3.71 (1.78) | 10.11 (9.70) | 0.2000 |
Effect of Falcon® tube used for stimulation on T cells response
Since the significant difference was observed with the CD4⁺Tbet⁺IFNγ⁺ cells and due to budgetary constraints, this subtype was chosen to examine whether the type of plastic container used for stimulation influences cytokine-producing T cell expression. As shown in Fig. 4E, no significant difference was observed in the percentage of CD4⁺Tbet⁺IFNγ⁺ cells when comparing stimulations performed in cell culture plates versus Falcon® tubes with 100 µL of whole blood.
Impact of volume of whole blood samples stimulated
The effect of increasing the volume of whole blood used for stimulation was also assessed. While an increase to 200 µL resulted in a lower response, the difference was not statistically significant when using Falcon® tubes (Fig. 4F). Interestingly, when 100 µL of whole blood was used in a cell culture plate, the response was significantly higher compared to 200 µL in a Falcon® tube.
Fig. 4: Impact of plastic container type on the percentage of CD4+Tbet+IFNγ + (n=5). (E) Comparison of cell culture plates with 100 µL of whole blood (P 100) and Falcon® tubes with 100 µL of whole blood (FT 100). (F) Comparison of plastic containers (Falcon® tubes vs. plates) and blood volumes (100 µL vs. 200 µL). *p < 0.05; ns, not significant; Falcon® tubes with 100 µL of whole blood (FT 200). Each dot represents an individual data point, and the horizontal black lines represent the mean. Statistical analysis was performed using the Mann–Whitney U test.
Discussion
Whole blood stimulation assays are widely utilized in clinical research, for applications such as disease characterization, evaluating host immunity, diagnosis testing, treatment monitoring, and drug development (13). Compared to PBMCs, whole blood is more readily accessible in clinical settings, making it a potentially convenient alternative for studying immune responses. This study aimed to optimize the conditions under which whole blood could serve as an alternative to PBMCs for assessing T cell responses using intracellular cytokine staining (ICS) by flow cytometry. Specifically, we sought to determine the optimal volume of whole blood and the most suitable cell culture containers for stimulation. Our study included firstly three healthy male donors, with both whole blood and PBMCs stimulated under identical conditions for comparison building on previous studies of cell-mediated responses using intracellular cytokine staining (ICS) in PBMCs and whole blood (2).
One hundred microliters (100 µL) of whole blood and PBMCs were used to compare the cytokine-producing T cell subpopulations. For surface marker labeling, We chose to perform staining after red blood cell (RBC) lysis (14), as this method seemed more appropriate for our experimental setup. Although some studies advocate staining before lysis (15,16). The results highlighted inter-individual variability in responses, which was more pronounced when PBMCs were used compared to whole blood. According to Moris et al., (2) who reported highly consistent CD4+ T cell responses between whole blood and PBMCs in their clinical trials, these differences observed in our study could stem from variations in cell concentration (unknown cells number or concentration, smaller volume of whole blood used) or differences in stimulation conditions. As whole blood stimulation is closest to in vivo setting compared to peripheral blood mononuclear cells (13). Moreover, the non-significant differences observed in CD4⁺GATA3⁺IL-4⁺; CD4⁺RORγt⁺IL-17A⁺; CD4⁺FoxP3⁺IL-10⁺ cells despite being higher in PBMcs could be related to our sample size which was too small due to lack of resources.
To explore the hypothesis that whole blood volume stimulated influences in vitro responses, fives donors were included to investigate the effects of varying whole blood volumes and different container types (Falcon tubes vs. 96-well U bottom plates) on the stimulation results focusing on Th1 (CD4+Tbet+IFNγ+) cells, as this population showed statistically significant differences between whole blood and PBMC assays.
Our findings revealed no significant difference in Th1 cell percentages when comparing stimulation in cell culture plates to Falcon tubes, suggesting that Falcon tubes can serve as viable alternatives for cell stimulation (3). However, increasing the volume of whole blood from 100 µL to 200 µL led to a decrease in response, although this difference was not statistically significant. Since the 96-well U bottom plates could not hold the total volume of whole blood sample including RPMI complete medium and stimulant (PMA) with 200 µL whole blood, we first compared the plate versus Falcon tube with 100 µL blood volume. The results showed no significant difference, so we assumed to compare the plate with different blood volumes using the Falcon tube.
Conversely, stimulation with 100 µL of whole blood in 96-well U bottom plates resulted in significantly higher Th1 responses compared to 200 µL in Falcon tubes. These results indicate that using 100 µL of whole blood in cell culture plates provides the optimal condition for characterizing T cell responses through ICS.
Measurements of vaccine immunogenicity in preclinical development and clinical trials routinely include quantification of antigen-specific Th1 and Th17 cells by ICS, and ELISA-based quantification of cytokines-release (17). Thus, this conclusive evaluation took into account the cellular aspect (Th1, Th2, Th17 and regulatory T cells) and the ICS to be applicable. In addition, it will constitute a basis for its applicability for the measurement of immunological markers with prognostic and diagnostic values in clinical biology. However, our study has some limitations. The small sample size may limit the generalizability of our results, and further studies should include larger cohorts with diverse population in age, sex and even their immune status and additional cell types to validate our observations. Additionally, the use of PMA and Ionomycin as activating stimuli may have affected CD4 expression levels (18), potentially leading to artificially lower proportions of cytokine-producing CD4+ T cells. Alternative stimulation protocols or adjustments to the current method could address this limitation in subsequent experiments.
Conclusion
This study suggests that 100 µL of whole blood assays can serve as a practical and reliable alternative to PBMC-based assays, for assessing T cell responses, with 100 µL of whole blood in 96-well cell culture plates being the optimal conditions for T-cell stimulation in resource-limited settings. These findings provide a foundation for further research to refine whole blood-based assays, making them more accessible and practical for a range of clinical and research applications.
Conflict of interest
The authors declare that they have no financial or personal relationship(s) which may have inappropriately influenced them in writing this article.
Authors contribution
Gnodja Tégmaba: Conceived and designed the study, enrolment, performed the experiments, formal analysis, writing original draft, participated in writing-review and editing.
Kadanga Motomdéwa Monique: Performed the experiments, formal analysis, participated in writing-review and editing.
Bara Fagdéba David: Performed the experiments, formal analysis, participated in writing-review and editing.
Simfele Hombamane Christelle: Performed the experiments, formal analysis, participated in writing-review and editing.
Kolou Maléwé: Conceived and designed the study, participated in writing-review and editing, contributed reagents/materials, supervision and validation.
Mbengue Babacar: Conceived and designed the study, participated in writing-review and editing, contributed reagents/materials, supervision and validation.
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