Mouse Naïve CD4⁺ T Cell Isolation and In vitro Differentiation into T Cell Subsets

All experimental procedures are performed using protocols approved by the office of Environmental Health and Safety at the Rosalind Franklin University of Medicine and Science. C57BL/6 mice (purchased from NCI) used for this protocol were housed under specific pathogen-free conditions, and all animal experiments were performed using protocols approved by the Institutional Animal Care and Use Committee (IACUC) at the Rosalind Franklin University of Medicine and Science.

1. Preparation of Instruments, Supplies, and Reagents

1. Coat 48-well or 24-well plates with anti-CD3 and anti-CD28 (Table 1) in sterile PBS, cover and wrap in parafilm to prevent evaporation and contamination, and then incubate O/N at 4 ^oC the day before T cell isolation. Alternatively, coat wells 1 h at 37 ^oC. Perform Th0, Th1, Th2, and iTreg conditions by coating both anti-CD3 and anti-CD28 at 1μg/ml in 0.5-1 ml total volume (48-well plate). For Th17, we typically coat anti-CD3 at 2 μg/ml and anti-CD28 at 1 μg/ml.
   NOTE: We have found that a lower anti-CD3 concentration is optimal for Th1, Th2, and iTreg generation while a higher anti-CD3 concentration is optimal for Th17 generation. Thus, anti-CD3 and anti-CD28 concentrations should be titrated and optimized for individual laboratories (Table 2).
2. Sterilize a set of dissection tools (scissors and forceps) by autoclave or disinfectant solution. A tube or small beaker containing 70% ethanol will be needed for quick sterilization of tools between dissections. Finally, a spray bottle containing 70% ethanol will be needed for sterilizing the surface of the mouse before dissection.
3. Prepare a dissection surface by wrapping a flat piece of Styrofoam or corkboard in aluminum foil. Dissecting pins or needles are used to fix the animal in place.
4. Prepare dishes or plates for the collection of tissues. If pooling tissues, separate 60mm dishes containing 3ml of sterile PBS + 1% FBS (PBS+) for spleen and lymph node tissues is sufficient. If keeping tissues from individual mice separate, prepare a 24-well plate containing 1ml of PBS+ in the desired number of wells.
5. Prepare complete RPMI media for cell culture and pre-warm before use (Table 1).
   NOTE: We routinely use RPMI but other media such as IMDM and DMEM may be equally effective or superior depending on the lab and the experiment.
6. Cut 2×2 cm squares of nylon mesh with a 120 μm pore size and autoclave to sterilize (Table 1).
7. Optional: if using the high-speed magnetic cell sorting system such as autoMACS for CD4⁺ cell enrichment, pre-warm running and rinsing buffers in a 37 ^oC water bath to prevent damage to the machine.
8. NOTE: Enrichment is recommended to increase the sorting yield and decrease sorting time.
9. All preparation steps, except for dissection, should be performed in a sterile tissue culture hood.

2. Isolation of Lymph Nodes and Spleen from Mice

1. Euthanize the required number of animals using an institutionally-approved technique. Use C57BL/6 female mice, age 5-10 wks, due to their high percentage of naïve T cells and lower body fat in comparison to males or to older mice.
   NOTE: Cells derived from male mice will perform similarly in vitro and the protocol steps remain the same. Female mice, 5-10 wks old, will typically yield 3-6 x 10⁶ naïve CD4⁺ T cells per mouse. However, using different strains of mice may affect yield and performance. For example, cells from C57BL/6 mice are better for generating Th1 cells while cells from Balb.c mice are better for generating Th2 cells.
2. Spread the limbs of the animal and pin them in place as shown in Figure 1. Cut the skin longitudinally from the anus to the chin while being careful not to puncture deeper tissue.
3. Spread open the skin with forceps and pin the skin in place to allow easy access to lymph nodes as shown in Figure 1.
4. Harvest the accessible lymph nodes from the locations shown in Figure 1. Grabbing the lymph node with a pair of forceps while separating tissue with another pair of forceps will allow for easy removal. Place the tissue in the collection dish.
5. Harvest the spleen from the location shown in Figure 1. In this case, careful cutting into the peritoneum is needed to gain access to the spleen. Place the tissue in a collection dish.
6. Repeat the procedure if more tissues will be harvested. Otherwise proceed to the next steps. At this point, perform all remaining steps on ice until the plating of the T cells after sorting.

3. Tissue Processing and CD4⁺ Enrichment

1. Process the lymph nodes and spleen in different dishes to increase the yield as lymph node cell preparations do not require erythrocyte lysis, which may result in minor leukocyte death. Therefore, the following steps describe a method for processing splenic and lymph node populations separately followed by combining prior to labeling for sorting.
2. Remove the tissue (spleen or pooled lymph nodes) from the collection dish and place in a new 60 mm dish containing 3 ml of PBS+. Place a square of sterile nylon mesh over the tissue using sterilized forceps.
3. Take the thumb side of a syringe plunger (3-10 ml) and gently smash the tissue against the mesh. Almost all of the tissue should go into suspension. Alternatively, place the tissue between two autoclaved frosted slides and grinded into suspension.
4. Pipette the suspension up and down a few times to break up the remaining soluble clumps. Place a new square piece of nylon mesh over the opening of a 15 ml conical tube and filter the suspension through to remove the remaining debris.
5. Repeat steps 2-4 for additional lymph node and splenic tissue.
6. Once the suspension has been filtered into a 15ml tube fill the remainder of the tube with PBS+ and invert a few times. Centrifuge the cells at 475 x g for 5 min at 4 ^oC.
7. For the splenic cells, aspirate the supernatant following centrifugation and resuspend the cell pellet in 1 ml of ice-cold 1X ACK solution per spleen for 1 min to lyse erythrocytes. Then add 10 ml of PBS+ on top of the ACK, invert the tube, and centrifuge 475 x g for 5 min at 4 ^oC.
8. For lymph node cells, aspirate the supernatant and resuspend in 2ml of PBS+. If desired these can be combined with the splenic cells after their ACK lysis and washing step.
9. Aspirate the supernatant after centrifugation of the splenocytes and resuspend in another 10 ml of PBS+. At this point the lymph node suspension can be added on top of the spleen suspension if tissue pooling is required for sorting. Centrifuge the tube again at 475 x g for 5 min at 4 ^oC.
10. The cell pellet can now be prepared for CD4 enrichment. If this step will not be performed, proceed to the sorting preparation steps.
11. For CD4 enrichment using the high-speed magnetic cell sorting system, resuspend the cellular pellet with CD4 beads. Typically 15 μl of beads diluted in 85 μl of PBS+ is used to label the tissues from 1 mouse. Ramp up the volume of bead mixture as needed, resuspend the pellet, and incubate for 15-30 min at 4 ^oC.
12. Wash the cells with 10 ml of PBS+, pass the suspension through nylon into a new 15 ml tube to remove debris, and centrifuge again at 475 x g for 5 min at 4 ^oC.
13. Resuspend the pellet in 100 μl of PBS+ per mouse and proceed to positive selection on the high-speed magnetic cell sorting system. If using a manual separation system, follow the manufacturer’s instructions for enrichment. Alternatively, resuspend the samples in FACS buffer: PBS with 1mM EDTA (pH = 8) and 1% BSA, sterile-filtered.
14. Remove the positive fraction and wash again with 10ml of PBS+. The enriched CD4⁺ fraction is now ready to be labeled for sorting.

4. Sorting Naïve CD4⁺ T cells

1. Label the cell pellet with PBS+ containing fluorescent antibodies directed against CD62L, CD44, CD25, and CD4. If using the antibodies listed in Table 1, dilutions for each are provided. For staining, a 100 μl volume of the antibody cocktail is used per tissues from one mouse.
2. Resuspend the pellet in antibody cocktail and incubate for 15-30 min at 4 ^oC in the dark.
3. Wash the cells with 10ml of PBS+ and centrifuge 475 x g for 5 min at 4 ^oC.
4. Pass the mixture over another nylon filter or cell strainer cap to remove any leftover debris. Pipette the labeled cells into a tube that is compatible for the cell sorter. Keep cells on ice and protected from the light until the sort.
5. Prepare collection tubes by pipetting 1-2ml of complete RPMI media or FBS into the bottom of tubes compatible for collection on a cell sorter.
6. Sort the naïve CD4⁺ T cells as a CD4⁺CD25^–CD62L⁺CD44^– population (Figure 2). Wash the collected cells with complete RPMI media and centrifuge as described above.
7. Resuspend the pellet in complete RPMI, count the naïve cells, and adjust the concentration to 1 x 10⁶ cells/ml.

5. Setting Up the In vitro Differentiation

1. Aspirate the wells of the anti-CD3 and anti-CD28 coated plates and wash each well with 1ml of sterile PBS (no FBS). For 48-well plates, plate 0.5 x 10⁶ cells/well in 0.5 ml of RPMI. For 24-well plates, culture 1 x 10⁶ cells/well in 1.0 ml of RPMI.
2. If testing the influence of a factor such as a drug add the substance to the appropriate wells either before, during, or after the differentiation process depending on the experiment. Next, supplement the culture media with cytokines and blocking antibodies accordingly. Th0: 30 U/ml hIL-2. Th1: 15 ng/ml rmIL-12, 30 U/ml hIL-2, and 5,000 ng/ml 11B11.Th2: 10 ng/ml rmIL-4, 30 U/ml hIL-2, 2,000 ng/ml soluble 37.51, and 5,000 ng/ml XMG1.2. iTreg: 15 ng/ml hTGFβ, 30 U/ml hIL-2, 5,000 ng/ml XMG1.2, and 5,000 ng/ml 11B11. Th17: 20 ng/ml rmIL-6, 3 ng/ml hTGFβ, 5,000 ng/ml XMG1.2, and 5,000 ng/ml 11B11.
   NOTE: The conditions presented above were found to be optimal for maximizing cytokine production as measured at the end of the differentiation experiment (Representative Results section). However, each condition should be optimized for individual laboratories (Table 2). Furthermore, the addition of soluble anti-CD28 in addition to the plate-bound anti-CD28 has been found to enhance Th2 cytokine production in our laboratory, but has no effect on other T cell subsets:
3. Incubate the plate at 37 ^oC with 5% CO₂ for 4-5 d prior to analysis of gene expression and cytokine production. Alternatively, remove the cells from the TCR stimuli after 2 d, adjusted to 1 x 10⁶ cells/ml with fresh media, and plate in new wells (non-coated) to enhance proliferation and final yield. In this case, fresh polarizing cytokines and antibodies are not needed but 10 U/ml of IL-2 should be added to new media for Th0, Th1, Th2, and iTreg cultures.

6. Analysis of Differentiation

1. For cytokine analysis by flow cytometry, restimulate each well with 10ng/ml PMA and 1,000 ng/ml ionomycin or 1 μg/ml anti-CD3 for 4-6 hr in the presence of a golgi inhibitor such as brefeldin A (Table 1). Perform intracellular cytokine staining depending on the experiment (Figure 3).
   NOTE: Stain non-lineage-specific cytokines or transcription factors, such as IL-4 (not shown) and IFNγ (Figure 3) for Th17 cultures, for each condition as negative controls and as an indicator of differentiation efficiency. Furthermore, stain Foxp3 for Th17 cultures as the reciprocal development of Tregs may occur with the addition of TGFβ.
2. For protein quantification, collect the supernatants from each well and assay by cytokine-specific ELISA (Figure 3). If there is concern that the factor being tested has influenced proliferation compared to control samples, the cells can be washed, counted, normalized, and then restimulated with 1 μg/ml of anti-CD3 for 24 h before collecting supernatants for ELISA assays. If assaying IL-4 from Th2 conditions, washing and restimulating is necessary to remove IL-4 that may be leftover from the initial differentiation culture.
3. For gene expression analysis by real time PCR, harvest cells after the incubation period, wash, normalize, and re-stimulate for 2-6 h with 1 μg/ml of anti-CD3 prior to harvesting mRNA (Figure 3).