Name: isolation of cd4+ t-cells and analysis of circulating t-follicular helper (ctfh) cell subsets from peripheral blood using 6-color flow cytometry
Uploaded: 2019-01-07T14:00:00.000+00:00
Duration: 7 min 39 s
Description: Watch this Scientific Journal Video about Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper cTfh Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry at JoVE.com

The work was supported by a grant from Leukaemia UK to ETB and MJA.

Blood samples were obtained from normal subjects (NS) (n = 12) as well as patients with marginal zone lymphoma (MZL) (n = 7) and other types of B-cell non-Hodgkin&#39;s lymphoma (BNHL) (6 FL patients, 2 lymphoplasmacytic lymphoma patients and 1 low-grade B-cell non-Hodgkin&#39;s lymphoma not otherwise specified patient). Patients were recruited from the hematology clinics at Leicester Royal Infirmary after having given informed, written consent, with ethical approval in place for all studies. Ethical approval was obtained by Leicestershire, Northamptonshire and Rutland Research Ethics Committee 1, reference 06/Q2501/122 for patient samples and the Health Research Authority (HRA) NRES Committee East Midlands-Derby, reference 14/EM/1176 for normal subjects.
1. Isolation of CD4+ T-cells from Whole Peripheral Blood
<ol>
	<li>Take fresh peripheral blood (2 to 15 mL) into K2EDTA tubes by standard venipuncture and process as soon as possible to maintain maximum viability. 
	NOTE: Use standard laboratory personal protective equipment (coat, gloves, glasses) and carry out the work in a class II Biosafety Cabinet.</li>
	<li>Bring the blood and all necessary reagents (CD4+ enrichment cocktail, 2% fetal bovine serum/phosphate buffered saline (FBS/PBS), density gradient media, and freezing medium if cryopreserving cells) to room temperature.</li>
	<li>Mix the blood with a commercially available cocktail of antibodies for the enrichment of human CD4+ T Cells. Use 50 &#181;L of the reagent for every 1 mL of blood. Use a 50 mL conical polypropylene tube for 5 to 15 mL of blood. 
	&#8203;NOTE: If using 2 to 4 mL of blood, perform this step in a 14 mL conical polypropylene tube.</li>
	<li>Incubate the mixture at room temperature for 20 min.</li>
	<li>Dilute the mixture with an equal volume of 2% FBS/PBS</li>
	<li>Layer this diluted mixture on top of the density gradient media slowly to avoid disturbing the interface and proceed to centrifugation immediately to avoid diffusion of the blood into the density gradient medium.
	<ol>
		<li>For 2 to 3 mL of blood, use 3 mL of density gradient medium in a 14 mL conical polypropylene tube, but for 4 mL of blood, use 4 mL of density gradient medium in a 14 mL conical polypropylene tube and for 5 to 15 mL blood, use 15 mL of density gradient medium in a 50 mL conical polypropylene tube.</li>
	</ol>
	</li>
	<li>Centrifuge the layered mixture for 20 min at 1,200 x g with the brake off at 20 &#176;C. 
	NOTE: A temperature below ambient may result in the contamination with red blood cells and granulocytes. A lower speed will cause the CD4+ isolation process to fail. Leaving the break engaged will decrease cell yield. Use a bench top centrifuge with rotors that can be closed to avoid aerosols.</li>
	<li>Remove the enriched CD4+ cell layer from the interface using a Pasteur pipette. 
	NOTE: The enriched cell layer will resemble a standard &#34;buffy coat&#34; of white cloudy cells, but as only CD4+ cells are present, this will naturally be smaller and more difficult to see.</li>
	<li>Add 2% FBS/PBS to the CD4+ cells up to a total volume of 10 mL and then wash twice with 2% FBS/PBS by centrifuging for 10 min at 400 x g during each wash.</li>
	<li>Resuspend the pellet in 2% FBS/PBS and count the cells stained with a vital dye (trypan blue) to determine the cell numbers and viability.</li>
	<li>Optional step: resuspend 5 x 105 to 1 x 106&#160;cells in freezing medium (10% dimethyl sulfoxide in FBS) in 1 mL aliquots. 
	NOTE: Levels of some surface markers may be altered by cryo-preservation and thawing. It is, therefore, very important that all samples are processed consistently. In order to minimize the differences in analytical variables, the samples were cryo-preserved and batched for analysis in this study.</li>
</ol>
2. Flow Cytometry
<ol>
	<li>Thaw the cryopreserved CD4+ cells quickly in a water bath at 37 &#176;C and wash once in pre-warmed medium (RPMI 1640 + L-glutamine supplemented with 10% FBS and 1x penicillin-streptomycin).</li>
	<li>Add the cells to 9 mL of medium, centrifuge for 5 min at 400 x g and remove the supernatant.</li>
	<li>Resuspend the cells in 1% bovine serum albumin (BSA) in PBS (50 &#181;L) so that each condition to be tested uses between 5 x 105 and 1 x 106 cells.</li>
	<li>Mix the cells with Staining Buffer (50 &#181;L). 
	NOTE: Brilliant Stain Buffer prevents various staining artefacts that may interfere with data analysis when two or more BV or BUV stains are used simultaneously owing to inherent chemical properties of these dyes. In single or unstained conditions, the 50 &#181;L of brilliant stain buffer can be substituted for 50 &#181;L of 1% BSA/PBS.</li>
	<li>Add the antibodies to cells (as per the &#34;volume used&#34; column in Table 1) and incubate on ice in the dark for 30 min.</li>
	<li>Wash the cells twice in 1% BSA/PBS by centrifuging for 3 min at 600 x g during each wash.</li>
	<li>Resuspend in 1% BSA/PBS (400 &#181;L) and transfer to a flow cytometry acquisition tube.</li>
	<li>Vortex the cells gently before acquiring data on a flow cytometer:</li>
</ol>
3. Flow Cytometry Controls and Set-up
<ol>
	<li>Using an unstained sample, set photodiode voltages so that lymphocytes can be separated from obvious debris and dead cells (events with high side scatter (SSC) area and low forward scatter (FSC) area)16. Using single stained samples, set photomultiplier (PMT) voltages so that positive fluorescence can be discerned from background fluorescence while making sure that all events fall within the detectable scale. 
	NOTE: An improvement would be to plot CV against PMT voltage for each PMT using dimly fluorescent beads to find the minimum voltage for optimum resolution (The &#34;Peak 2&#34; method19).</li>
	<li>Account for spectral overlap by generating a compensation matrix using single stains with capture beads. Add commercially available compensation beads (60 &#181;L), and negative control beads (60 &#181;L) to 1% BSA/PBS (100 &#181;L), and vortex. 
	NOTE: The capture beads used must match the host species and IgG isotype of the antibody being used. The compensation matrix should be re-calculated if the lot number of any tandem dyes changes, as they can display considerable variability in their emission spectra between lots.</li>
	<li>Add a single antibody (20 &#181;L) and vortex.</li>
	<li>Incubate at room temperature in the dark for 30 min.</li>
	<li>Centrifuge at 200 x g for 10 min and discard the supernatant.</li>
	<li>Resuspend in 1%BSA/PBS (500 &#181;L) and vortex.</li>
	<li>Acquire the sample with a flow cytometer using the designated compensation matrix generator in the acquisition software according to the manufacturer&#39;s instructions.</li>
	<li>Use FMO controls to guide the placement of gates for positive marker fluorescence to account for dye spill over, which is the main source of background fluorescence in experiments using &#8805;4 colours16. Follow the general cell-surface staining protocol as described in Step 3, but for each condition, omit one of the fluorophores from the staining step (where CD45RA is omitted, also omit the live/dead stain). Acquire 100,000 lymphocytes for each condition.</li>
	<li>Set the gate positivity at &#8804;0.5% of cells. See Figure 2 for an illustrated example of FMO controls.</li>
</ol>
4. Data Analysis
<ol>
	<li>Employ the flow cytometer&#39;s acquisition software to set a threshold of 5,000 units on the FSC parameter to exclude very small debris.</li>
	<li>Use a stopping gate to acquire 10,000 cTfh cells (CD4+ CD45RA- CXCR5+). 
	NOTE: The individual user can collect more than 10,000 cells. This number was a compromise between collecting enough cells to provide meaningful results and the time taken for collection.</li>
	<li>Using an FSC-Area / SSC-Area dot plot, draw an ellipse or polygon gate to select the population of lymphocytes whilst excluding debris and overtly dead cells (events with a high SSC-Area and low FSC-Area).</li>
	<li>Using an FSC-Area / FSC-Width dot plot, draw a polygon gate to select single cells while excluding doublets (doublets have an increased area but similar width to single cells).</li>
	<li>Using an FSC-Area / CD45RA and viability marker dot plot, draw a rectangular gate to select live, CD45RA- cells (cells with a low fluorescence for this marker).</li>
	<li>Using an FSC-Area / CD4 dot plot, draw a rectangular gate to select CD4+ cells (cells with a high fluorescence for this marker).</li>
	<li>Using a CD4 / CXCR5 dot plot, draw a rectangular gate to select CXCR5+ (i.e., cTfh) cells (cells with a high fluorescence for this marker).</li>
	<li>Using a CXCR3 / CCR6 dot plot, place a quad gate to subdivide cTfh cells into cTfh1,2 and 17 cells (cells that are high and low for each marker). 
	NOTE: Cells with the phenotype CD4+ CXCR5+ CXCR3+ CCR6+ are poorly characterized. We refer to these cells as cTfh1/1718 because conventional helper T-cells (CD4+ CXCR5-) that are transitioning between Th17 and Th1 show expression of CXCR3 and CCR6 and have been described as cTfh1/1719.</li>
	<li>Using a PD-1 histogram, use the range tool to subdivide cTfh cells (or if preferred, the individual cTfh1,2,17 or 1/17 subsets) into PD-1-/+ or hi populations. 
	NOTE: The distinction between PD-1+ and PD-1hi is not well defined in the literature. The threshold was set as the same PD-1 intensity required to detect bona-fide Tfh in human tonsil lymphocyte suspensions using flow cytometry with the same antibody panel. Alternatively, a marker for ICOS can be added to the antibody panel, as only PD1hi cells are ICOS+.</li>
</ol>

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The authors have nothing to disclose.

This protocol represents a simple and efficient way to analyze peripheral blood cTfh cells, enabling the detection of all relevant subsets identified in the literature thus far. Blood samples can be easily and efficiently obtained as part of standard out-patient clinics and serial samples can be collected in parallel with clinical data. In turn, this enables prospective studies evaluating cTfh subsets as biomarkers for disease progression or response to treatment. These studies would be particularly warranted in disease ...

T-follicular helper cells (Tfh) are a CD4+ T-cell subset that was initially characterized in lymphoid tissues1. These cells express PD-1 and CXCR5 surface receptors, secrete IL-21 and IL-4 and show nuclear expression of the transcription factor, BCL-62,3. As their name suggests, they are found in germinal centers and are essential for high affinity antibody production1.
Dysregulated Tfh responses have been implicated in disease pathogenesis, most notably autoimmune disease, where they promote the expansion of autoreactive B cells4. They also play a role in the tumor microenvironment of both solid5,6 and lymphoid cancers7. Conversely, genetic defects of surface proteins essential for Tfh function such as inducible T-cell costimulator (ICOS) result in human immunodeficiency syndromes8. CD4+ CXCR5+ cells in human peripheral blood are termed circulating T-follicular helper cells (cTfh) and are believed to be the memory compartment of Tfh cells in tissues9. The purpose of the method described here is the analysis of cTfh subsets following CD4+ cell isolation from peripheral blood samples.
Several cTfh subsets have been defined and the efficiency with which they provide B-cell help differs from one subset to another9,10,11,12. The relative proportions of these subsets are altered in a number of diseases, most prominently autoimmune disease in which there is almost always a relative increase in the more functional PD-1+/hi and/or cTfh2 or cTfh17 subsets in comparison to the less functional PD-1- and/or cTfh1 subsets12. The extent of these changes frequently associate with clinical parameters including disease activity and autoantibody titers, indicating a potential role of cTfh subset distribution as a prognostic biomarker in disease, which may reflect the activity of Tfh in lymphoid tissues9,12,13,14. Additionally, taking blood samples from participants is quick, safe and acceptable, and so allows serial monitoring for the analysis of disease progression or response to therapy.
The use of isolated CD4+ T-cells over traditional peripheral blood mononuclear cell (PBMNC) suspensions enables higher throughput flow cytometry experiments by reducing the time required to acquire a substantial number of cTfh cells for analysis. This is particularly helpful when sorting cells from rare cTfh subsets using flow activated cell sorting (FACS). To aid the efficiency, these suspensions can be cryopreserved to enable &#34;batching&#34; of samples to be used in the flow cytometry experiment. On testing, the CD4+ purity was not reduced by cryopreservation.
While different laboratories used different markers to categorise cTfh cells in the early stages of their discovery, the method presented here makes use of a unified scheme of two groups of cell-surface markers as proposed by Schmidt et al.12,15 to enable the simultaneous identification of cTfh and their nine recognized subsets in a single flow cytometry experiment.
As only cell surface markers are used, the cells do not require fixation or permeabilization, and thus can remain alive for downstream functional studies. This could be facilitated by cell sorting using FACS with the same antibody panel. This panel could be expanded to include other markers, allowing for the restrictions of the flow cytometer being used.
The analysis of multi-color flow cytometry experiments can be challenging due to the inherently subjective nature of gating on 2-dimensional dot plots, especially when cell populations do not have a clear bi-modal distribution in marker fluorescence, as is the case for cTfh cells and their subsets. For this reason, it is imperative to set up effective controls to reduce the artefacts to enable better resolution of populations and to set gating strategies confidently. As such, antibody panel design and the set-up of basic controls for a flow cytometry experiment, i.e., using compensation and FMO controls are outlined in Step 3.4.2 and 3.4.3,respectively.
All cTfh cells are defined as CD4+ CXCR5+ CD45RA-. The level of expression of the characteristic Tfh activation marker PD-1 can then be determined to identify the subsets of PD-1-, PD-1+ or PD-1hi cTfh cells. Then, using a combination of the chemokine receptors CXCR3 and CCR6, which are differentially expressed by traditional Th1,2 or 17 cells, cTfh can be characterized as cTfh1,2 or 17-like by a profile of CXCR3+ CCR6-, CXCR3- CCR6-, and CXCR3- CCR6+, respectively.
The antibody panel used by our laboratory is displayed in Table 1. The user may have to adapt their fluorophore selection to account for the laser and light filter configuration available on their local flow cytometer.
The following considerations influence the choice of fluorophores. Use bright fluorophores where possible. In particular, use the brightest available fluorophores on the dimmest (less highly expressed) markers. Dimmer markers include PD-1, CXCR3 and CCR6, and to a lesser extent, CXCR5. We specifically made use of the newer BB, BV and BUV fluorophores which provide excellent brightness and thus enable easier resolution of distinct populations of the cells.
Spread the fluorophore selection across the emission spectra as much as possible to minimize spectral overlap and thus the level of compensation required. A free, online tool that can be used to assist designing a flow cytometry panel can be found here: http://www.bdbioscienes.com/us/s/spectrumviewer. To save space on the emission spectrum, we employed a &#34;dump channel&#34; by using a viability dye with an emission wavelength that overlaps with that of APC-H7 (conjugated to CD45RA) to enable the detection (and exclusion) of both dead and/or CD45RA+ using a single detector.
Here, a protocol is presented for the isolation of peripheral blood CD4+ T-cells and their subsequent analysis by flow cytometry to determine the proportions of the different and recently described circulating subsets.

<table><tbody><tr><td>RosetteSep Human CD4+ T Cell Enrichment Cocktail</td><td>STEMCELL TECHNOLOGIES</td><td>15062</td><td></td></tr><tr><td>Ficoll-Paque PLUS</td><td>GE Healthcare Life Sciences</td><td>17144003</td><td></td></tr><tr><td>BUV395 Mouse Anti-Human CD183&amp;nbsp;Clone 1C6/CXCR3</td><td>BD Horizon</td><td>565223</td><td></td></tr><tr><td>BV711 Mouse Anti-Human CD196 (CCR6)&amp;nbsp;Clone 11A9</td><td>BD Horizon</td><td>563923</td><td></td></tr><tr><td>BV421 Rat Anti-Human CXCR5 (CD185)&amp;nbsp;Clone&amp;nbsp;RF8B2</td><td>BD Horizon</td><td>562747</td><td></td></tr><tr><td>BB515 Mouse Anti-Human CD4&amp;nbsp;Clone &amp;nbsp;RPA-T4</td><td>BD Horizon</td><td>564419</td><td></td></tr><tr><td>PE Mouse Anti-Human CD279&amp;nbsp;Clone MIH4</td><td>BD Pharmingen</td><td>557946</td><td></td></tr><tr><td>APC-H7 Mouse Anti-Human CD45RA Clone&amp;nbsp; HI100</td><td>BD Pharmingen</td><td>560674</td><td></td></tr><tr><td>LIVE/DEAD Fixable Far Red Dead Cell Stain Kit, for 633 or 635 nm excitation</td><td>ThermoFisher Scientific</td><td>L34973</td><td></td></tr><tr><td>Brilliant Stain Buffer</td><td>BD Horizon</td><td>563794</td><td></td></tr><tr><td>Anti-Mouse Ig, &amp;kappa;/Negative Control Compensation Particles Set</td><td>BD CompBead</td><td>552843</td><td></td></tr><tr><td>FACS Aria II Flow Cytometer</td><td>BD Biosciences</td><td>644832</td><td></td></tr><tr><td>FACSDiva 6.1.3</td><td>BD Biosciences</td><td>643629</td><td>Flow Cytometer Acquisition Software</td></tr><tr><td>FlowJo 10.2</td><td>Treestar Inc.</td><td></td><td>Flow Cytometry Data Analysis Software</td></tr><tr><td>Anti-CXCR3 antibody</td><td>BD Horizon&amp;nbsp;</td><td>565223</td><td></td></tr><tr><td>Anti-CCR6 antibody</td><td>BD Horizon&amp;nbsp;</td><td>563923</td><td></td></tr><tr><td>Anti-CXCR5 antibody</td><td>BD Horizon&amp;nbsp;</td><td>562747</td><td></td></tr><tr><td>Anti-CD4 antibody</td><td>BD Horizon&amp;nbsp;</td><td>564419</td><td></td></tr><tr><td>Anti-PD-1 antibody</td><td>BD Pharmingen&amp;nbsp;</td><td>557946</td><td></td></tr><tr><td>Anti-CD45RA antibody</td><td>BD Pharmingen&amp;nbsp;</td><td>560674</td><td></td></tr><tr><td>Viability Marker</td><td>ThermoFisher Scientific&amp;nbsp;</td><td>L34973</td><td></td></tr></tbody></table>

isolation of cd4+ t-cells and analysis of circulating t-follicular helper (ctfh) cell subsets from peripheral blood using 6-color flow cytometry

Aberrant T-follicular helper (Tfh) cell activity is detectable in autoimmune conditions and their presence is associated with clinical outcomes when the lymph node microenvironment in B-cell non-Hodgkin&#39;s lymphoma is analyzed. Subsets of circulating T-follicular helper cells (cTfh), the circulating memory compartment of Tfh cells in the blood, are also perturbed in disease and therefore represent potential novel predictive biomarkers. Peripheral blood-based testing is advantageous because it is relatively non-invasive and allows simple serial monitoring.This article describes a method for isolating CD4+ T-cells from human blood, and further analysis by flow-cytometry to enumerate cTfh cells and the proportions of their various subsets (cTfhPD-1-/+/hi, cTfh1,2,17 and cTfh1/17). The level of these subsets was then compared between normal subjects and patients with lymphoma. We found that the method was robust enough to obtain reliable results from routinely collected patient material. The technique we describe for the analysis can be easily adapted to cell sorting and downstream applications such as RT-PCR.

High CD4+ purity was achieved using the CD4+ isolation protocol, which was reliable across all blood samples tested by us (mean: 96.6%, SD: 2.38, n=31) (Figure 3).
Identification of cTfh (CD4+ CXCR5+ cells) in a representative normal subject is presented (Figure 4A). The proportion of total cTfh cells within C...

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Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper cTfh Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

Here, a protocol for the isolation and characterization of CD4+ T-cell subsets from human peripheral blood is described. Purified CD4+ T-cells are analyzed by flow cytometry to determine proportions of T-follicular helper cell subsets.

Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper (cTfh) Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

Aberrant T-follicular helper (Tfh) cell activity is detectable in autoimmune conditions and their presence is associated with clinical outcomes when the ...

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12.1K Views.  University of Leicester. Here, a protocol for the isolation and characterization of CD4+ T-cell subsets from human peripheral blood is described. Purified CD4+ T-cells are analyzed by flow cytometry to determine proportions of T-follicular helper cell subsets.

Video: Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper cTfh Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

Isolating Human Peripheral Blood Mononuclear Cells and CD4+ T cells from S&#233;zary Syndrome Patients for Transcriptomic Profiling

Cutaneous T-cell lymphomas (CTCL) are derived from the transformation and uncontrolled proliferation of mature skin-homing T cells, and mycosis fungoides (MF) and S&#233;zary syndrome (SS) represent the most common subtypes. Despite a number of studies on characterizing gene expression, genetic alterations, and epigenetic abnormalities of CTCL, the molecular pathogenesis of MF/SS remains unclear. MF refers to the more common CTCL with a skin-predominance, and is usually limited to skin, whereas&#160;SS is an aggressive leukemic variant of CTCL with widespread skin involvement and is characterized by neoplastic distribution mainly involving blood, skin, and lymph node. Focusing on clinical practice, the identification of gene expression biomarkers has&#160;enormous potential to improve diagnosis and treatment of MF/SS. Indeed, recent transcriptomic studies have identified potential diagnostic biomarkers from differences in gene expression between normal and malignant T cells, which may improve our understanding of SS biology, and reveal potential therapeutic targets. This manuscript describes a detailed reproducible protocol for the isolation of peripheral blood mononuclear cells from fresh whole blood from patients diagnosed with SS, selection of CD4+ memory T cells (CD4+CD45RO+ T cells), chemical stimulation, and preparation of RNA suitable for transcriptomic profiling to discover novel prognostic molecular markers to gain additional insight in disease etiology. The stimulation using chemical agonist to activate nuclear regulation provides more specific assessment for pathways important in the dynamic transcription regulation and gene expression and eliminates confounding defects that may arise from upstream signaling defects arising from TCR antigen loss at the cell membrane. The data obtained from comparison of transcriptome of unstimulated to stimulated SS T cells unmasks functional regulatory gene expression defects not evident from analysis of quiescent unstimulated cells. Furthermore, the method outlined from this approach can be adapted for studying T cell gene expression defects in other T cell immune diseases.

We present a simple protocol for the isolation of peripheral blood mononuclear cells from whole blood obtained from patients diagnosed with S&#233;zary Syndrome, followed by selection of CD4+ T cells, their stimulation with phorbol12-myristate13-acetate and A23187 ionophore, and preparation of RNA for transcriptomic profiling.

Cutaneous T-cell lymphomas (CTCL) are derived from the transformation and uncontrolled proliferation of mature skin-homing T cells, and mycosis fungoides ...

Cancer Research

Cell-based Flow Cytometry Assay to Measure Cytotoxic Activity

Cytolytic activity of CD8+ T cells is rarely evaluated. We describe here a new cell-based assay to measure the capacity of antigen-specific CD8+ T cells to kill CD4+ T cells loaded with their cognate peptide. Target CD4+ T cells are divided into two populations, labeled with two different concentrations of CFSE. One population is pulsed with the peptide of interest (CFSE-low) while the other remains un-pulsed (CFSE-high). Pulsed and un-pulsed CD4+ T cells are mixed at an equal ratio and incubated with an increasing number of purified CD8+ T cells. The specific killing of autologous target CD4+ T cells is analyzed by flow cytometry after coculture with CD8+ T cells containing the antigen-specific effector CD8+ T cells detected by peptide/MHCI tetramer staining. The specific lysis of target CD4+ T cells measured at different effector versus target ratios, allows for the calculation of lytic units, LU30/106 cells. This simple and straightforward assay allows for the accurate measurement of the intrinsic capacity of CD8+ T cells to kill target CD4+ T cells.

This protocol describes a sensitive, cell-based cytotoxicity assay. By enumerating the decrease in frequency of live target CD4+ T cells in the presence of an increasing number of effector CD8+ T cells, this assay allows for the direct assessment of cytolytic activity of antigen-specific CD8+ T cells.

Cytolytic activity of CD8+ T cells is rarely evaluated. We describe here a new cell-based assay to measure the capacity of antigen-specific CD8+ T cells ...

Immunology and Infection

Retroviral Transduction of Helper T Cells as a Genetic Approach to Study Mechanisms Controlling their Differentiation and Function

Helper T cell development and function must be tightly regulated to induce an appropriate immune response that eliminates specific pathogens yet prevents autoimmunity. Many approaches involving different model organisms have been utilized to understand the mechanisms controlling helper T cell development and function. However, studies using mouse models have proven to be highly informative due to the availability of genetic, cellular, and biochemical systems. One genetic approach in mice used by many labs involves retroviral transduction of primary helper T cells. This is a powerful approach due to its relative ease, making it accessible to almost any laboratory with basic skills in molecular biology and immunology. Therefore, multiple genes in wild type or mutant forms can readily be tested for function in helper T cells to understand their importance and mechanisms of action. We have optimized this approach and describe here the protocols for production of high titer retroviruses, isolation of primary murine helper T cells, and their transduction by retroviruses and differentiation toward the different helper subsets. Finally, the use of this approach is described in uncovering mechanisms utilized by microRNAs (miRNAs) to regulate pathways controlling helper T cell development and function.

Many experimental systems have been utilized to understand the mechanisms regulating T cell development and function in an immune response. Here a genetic approach using retroviral transduction is described, which is economic, time efficient, and most importantly, highly informative in identifying regulatory pathways.

Helper T cell development and function must be tightly regulated to induce an appropriate immune response that eliminates specific pathogens yet prevents ...

Analysis of Human T Cell Activity in an Allogeneic Co-Culture Setting of Pre-Treated Tumor Cells

Cytotoxic T cells play a key role in the elimination of tumor cells and are, therefore, intensively studied in cancer immunology. The frequency and activity of cytotoxic T cells within tumors and their tumor microenvironment (TME) are now well-established prognostic and predictive biomarkers for numerous tumor types. However, it is well-known that various tumor treatment modalities, including radiotherapy, chemotherapy, immunotherapy, and targeted therapy, modulate not only the immunogenicity of the tumor but also the immune system itself. Consequently, the interaction between tumor cells and T cells requires more intensive study in different therapeutic contexts to fully understand the complex role of T cells during tumor therapy. To address this need, a protocol was developed to analyze the activity and proliferative capacity of human cytotoxic (CD8+) T cells in co-culture with pre-treated tumor cells. Specifically, CD8+ T cells from healthy donors are stained with the non-toxic proliferation marker carboxyfluorescein diacetate succinimidyl ester (CFSE) and stimulated using CD3/CD28-coated plates. Subsequently, T cells are co-cultured with pre-treated tumor cells. As a readout, T cell proliferation is quantified by measuring CFSE signal distribution and assessing the expression of surface activation markers via flow cytometry. This can be further complemented by quantifying cytokine release using enzyme-linked immunosorbent assay (ELISA). This method facilitates the evaluation of treatment-induced changes in the interaction between tumor cells and T cells, providing a foundation for more detailed analyses of tumor treatment modalities and their immunogenicity in a human ex vivo setting. Additionally, it contributes to the reduction of pre-clinical in vivo analyses.

The present protocol describes an experimental workflow that allows for the ex vivo analysis of human T cell stimulation in an allogenic co-culture system with pre-treated tumor cells.

Cytotoxic T cells play a key role in the elimination of tumor cells and are, therefore, intensively studied in cancer immunology. The frequency and ...

Determination of Regulatory T Cell Subsets in Murine Thymus, Pancreatic Draining Lymph Node and Spleen Using Flow Cytometry

Our immune system consists of a number and variety of immune cells including regulatory T cells (Treg) cells. Treg cells can be divided into two subsets, thymic derived Treg (tTreg) cells and peripherally induced Treg (pTreg) cells. They are present in different organs of our body and can be distinguished by specific markers, such as Helios and Neuropilin 1. It has been reported that tTreg cells are functionally more suppressive than pTreg cells. Therefore, it is important to determine the proportion of both tTreg and pTreg cells when investigating heterogeneous cell populations. Herein, we collected thymic glands, pancreatic draining lymph nodes and spleens from normoglycemic non-obese diabetic mice to distinguish tTreg cells from pTreg cells using flow cytometry. We manually prepared single cell suspensions from these organs. Fluorochrome conjugated surface CD4, CD8, CD25, and Neuropilin 1 antibodies were used to stain the cells. They were kept in the fridge overnight. On the next day, the cells were stained with fluorochrome conjugated intracellular Foxp3 and Helios antibodies. These markers were used to characterize the two subsets of Treg cells. This protocol demonstrates a simple but practical way to prepare single cells from murine thymus, pancreatic draining lymph node and spleen and use them for subsequent flow cytometric analysis.

Herein, we present a protocol to prepare single cells from murine thymus, pancreatic draining lymph node and spleen to further study these cells using flow cytometry. In addition, this protocol was used for determining the subsets of regulatory T cells using flow cytometry.

Our immune system consists of a number and variety of immune cells including regulatory T cells (Treg) cells. Treg cells can be divided into two subsets, ...

Discrimination of Seven Immune Cell Subsets by Two-fluorochrome Flow Cytometry

Immune cell characterization heavily relies on multicolor flow cytometry to identify subpopulations based on differential expression of surface markers. Setup of a classic multicolor panel requires high-end instruments, custom labeled antibodies, and careful study design to minimize spectral overlap. We developed a multiparametric analysis to identify major human immune populations (CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, NK cells and monocytes) in peripheral blood by combining seven lineage markers using only two fluorochromes. Our strategy is based on the observation that lineage markers are constantly expressed in a unique combination by each cell population. Combining this information with a careful titration of the antibodies allows investigators to record five additional markers, expanding the optical limit of most flow cytometers. Head-to-head comparison demonstrated that the vast majority of immune cell populations in peripheral blood can be characterized with comparable accuracy between our method and the classic &#34;one fluorochrome-one marker approach&#34;, although the latter is still more precise for identifying populations such as NKT cells and &#947;&#948; T cells. Combining seven markers using two fluorochromes allows for the analysis of complex immune cell populations and clinical samples on affordable 6-10 fluorochrome flow cytometers, and even on 2-3 fluorochrome field instruments in areas with limited resources. High-end instruments can also benefit from this approach by using extra fluorochromes to accomplish deeper flow cytometry analysis. This approach is also very well suited for screening several cell populations in the case of clinical samples with limited number of cells.

Here, we present a flow cytometric protocol to identify CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, NK cells and monocytes in human peripheral blood by using only two fluorochromes instead of seven. With this approach, five additional markers can be recorded on most flow cytometers.

Immune cell characterization heavily relies on multicolor flow cytometry to identify subpopulations based on differential expression of surface markers. ...

Flotation-Based T Cell Isolation, Activation, and Expansion from Human Peripheral Blood Mononuclear Cell Samples Using Microbubbles

The process of isolating T cells from peripheral blood mononuclear&#160;cells (PBMCs) to establish ex vivo cultures is crucial for research, clinical testing, and cell-based therapies. In this study, a simple, novel protocol to isolate, activate, and expand T cells from PBMCs ex vivo is presented. This study utilizes functionalized buoyancy-activated cell sorting (BACS) technology to isolate and activate T cells. Briefly, the protocol involves the positive selection of CD3+ cells from leukopak-derived PBMCs, followed by a 48 h co-stimulation with pre-conjugated anti-CD28-bound streptavidin microbubbles (SAMBs) prior to transduction in 24-well plates. Functionalized microbubbles offer a unique opportunity to buoyantly activate cells, leading to proliferative phenotypes that allow for expansion with minimal exhaustion. This technique offers reduced exhaustion because the co-stimulatory microbubbles remain buoyant and return to the top of the culture medium, thus reducing the amount of time that the expanding cells are in contact with the co-stimulatory factors. The results indicate that the isolated and cultured T cells receive enough stimulation to activate and proliferate but not to an extent that leads to overactivation, which then leads to exhaustion, as demonstrated by the presence of excessive PD-1.

The objective of this study is to demonstrate the feasibility of flotation-based separation to isolate, activate, and expand primary human T cells.

The process of isolating T cells from peripheral blood mononuclear&#160;cells (PBMCs) to establish ex vivo cultures is crucial for research, clinical ...

Cell Cycle Analysis: Assessing CD4 and CD8 T Cell Proliferation After Stimulation Using CFSE Staining and Flow Cytometry

Source: Perchet Thibaut1,2,3, Meunier Sylvain1,2,3, Sophie Novault4, Rachel Golub1,2,3 
1 Unit for Lymphpoiesis, Department of Immunology, Pasteur Institute, Paris, France 
2 INSERM U1223, Paris, France 
3 Universit&#233; Paris Diderot, Sorbonne Paris Cit&#233;, Cellule Pasteur, Paris, France 
4 Flow Cytometry Platfrom, Cytometry and Biomarkers UtechS, Center for Translational Science, Pasteur Institute, Paris, France
The cell cycle is a universal process of life. During the cell cycle, a cell undergoes several modifications to divide into two daughter cells. This mechanism occurs throughout an organism&#39;s life in response to its needs. Cell divisions and embryonic development produce a full organism from a single-celled zygote. During adulthood, the cell cycle is central to many critical biological processes, such as tissue repairs.
Mechanisms of cell division are tightly controlled events where the cell undergoes stepwise modifications before final division. Cells that are not yet in the cycle are described as being in the Gap 0 (G0) phase. During this stage the cell is considered quiescent. When the cell starts to cycle, four distinct phases are recognized: Gap 1 (G1), Synthesis (S), Gap 2 (G2) and Mitosis (M). G1 phase is a checkpoint for resources needed by the cell for DNA synthesis. Then, S phase occurs, and DNA replication starts, followed by the G2 interphase, another checkpoint that controls all elements necessary to for the cell to divide. Finally, the cell enters mitosis and divides into two daughter cells.
Cell division is a highly informative parameter in many different biological systems. In the field of immunology, analysis of leukocyte proliferation can indicate the mechanism of the immune response. Other domains of investigation also rely on cell cycle analysis. For instance, analysis of the cell cycle during tumor development has improved our understanding of cancer.
Many fluorescent dyes are now available for tracking cell proliferation. These dyes differ in their chemical and spectral properties. Two different classes of dyes exist: protein dyes permanently combine with protein by forming a covalent bond, and membrane dyes stably intercalate within cell membranes via strong hydrophobic associations. In vitro and in vivo studies of immune cell proliferation by flow cytometry are among the most common applications of both classes of cell tracking dyes (1, 2).
CFSE (Carboxyfluorescein succinimidyl ester) is a fluorescent dye that marks dividing cells. Initially, all the cells receive the same amount of dye; dividing cells evenly split the dye they received between their two daughter cells. Consequently, the cell cycle can be followed by the progressive decrease of dye intensity in the cells. CFSE staining is followed by conventional multi-parametric flow cytometry, a high-throughput, fluorescence-based technology that allows phenotypic and functional characterization of cells based on their degree of CFSE staining (3).
In the following experiment, we assess the proliferation of CD4+ and CD8+ T cells in-vitro, following CD3 stimulation, using CFSE staining and flow cytometry.

Cell Cycle Analysis: Assessing CD4 and CD8 T Cell Proliferation

Source: Perchet Thibaut1,2,3, Meunier Sylvain1,2,3, Sophie Novault4, Rachel Golub1,2,3
1 Unit for Lymphpoiesis, Department of Immunology, Pasteur ...

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An In Vitro Technique to Study T Cell Interaction with Supported Planar Lipid Bilayers

Source: Steblyanko, M., et al. <a href="https://app.jove.com/v/58143">Assessment of the Synaptic Interface of Primary Human T Cells from Peripheral Blood and Lymphoid Tissue.</a> J. Vis. Exp. (2018).
This video demonstrates an in vitro technique to study immune synapse formation between primary human T cells and supported lipid bilayers (SLB). T cell surface proteins interact with respective ligands immobilized on SLB to form an immune synapse, followed by the movement of lytic granules towards the synapse, which is visualized using fluorescence microscopy.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Isolation of CD4+ T Cells for an ...

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Cellular Immunodiagnostics

Selective Isolation of CD4+ V&delta;1+ &gamma;&delta; T Cells from Human Peripheral Blood Mononuclear Cells

Source: Welker, C., et al. Isolation and Ex Vivo <a href="https://app.jove.com/v/53482">Culture of V&#948;1+CD4+&#947;&#948; T Cells, an Extrathymic &#945;&#946;T-cell Progenitor.</a> J. Vis. Exp. (2015).
This video demonstrates a protocol for the magnetic labeling-based isolation of a low-concentration population of CD4+V&#948;1+&#947;&#948; T cells from human peripheral blood mononuclear cells (PBMCs). Using a combination of FITC-conjugated anti-V&#948;1 antibodies, magnetic anti-FITC microbeads, and anti-CD4 antibody-coated superparamagnetic beads, the CD4+V&#948;1+ T-cells can be successfully purified from the PBMC population.

Source: Welker, C., et al. Isolation and Ex Vivo Culture of V&#948;1+CD4+&#947;&#948; T Cells, an Extrathymic &#945;&#946;T-cell Progenitor. J. Vis. Exp. ...

Isolation of CD4⁺ T-cells and Analysis of Circulating T-follicular Helper (cTfh) Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

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Isolating Human Peripheral Blood Mononuclear Cells and CD4+ T cells from Sézary Syndrome Patients for Transcriptomic Profiling