Purification of Thymocyte and T Cell Subsets

Abstract

Many analytical or cell culture procedures require homogeneous starting cell populations that cannot be obtained directly from organ dissection. Here, we describe two enrichment procedures to achieve this goal and discuss their respective advantages in specific experimental contexts. Notes in this chapter include some tips on how to determine the appropriate level of purity (see Note 1).

1. Introduction

This chapter describes two procedures for preparing homogeneous thymocyte and T cell suspensions from organs that contain a heterogeneous mixture of cell types. Both techniques rely on the availability of appropriate antibodies against surface molecules differentially expressed on desired and unwanted cells.

The first procedure uses antibody-coated magnetic beads to retain (“positive selection”) or discard (“negative selection”) cells expressing specific surface molecules. The positive selection approach is of broad applicability, as one only needs a single cell-specific surface antigen, but has the significant disadvantage of leaving the purified cells coated with beads. This has the potential to interfere with cell activation or differentiation in culture (antibodies that bind the T cell receptor, for example, are detrimental for certain procedures as they can activate T cells). In addition, the presence of cell aggregates tends to limit cell purity, as one desired cell in the aggregate will positively select the whole aggregate. The negative selection approach is more complex, because it requires antibodies against all other cell types present in the starting population. However it yields cells “untouched” (i.e., without attached antibodies or beads), which is of particular importance if the cells are to be cultured or tested functionally in subsequent steps.

The second procedure uses flow cytometric sorting of cells stained with appropriate fluorescent antibodies. As with the bead purification procedure, antibodies can be used in flow cytometric sorting to positively or negatively mark cells, and in the latter case leaves cells untouched. In contrast, positively marked cells carry fluorescent antibodies that have to be taken into account when subsequent staining steps are considered, and, as mentioned above, can contribute to activation or differentiation. The choice of the sorter is generally dictated by locally available equipment. Except for experienced users, professional operators generally perform sorts; prior consultation with operators is highly recommended to discuss staining and sorting strategies, including gate definitions.

Several experimental considerations guide the choice of one procedure over the other. The first regards biological consequences on purified cells. Especially with negative selection, bead purification induces only minimal cell change or damage. In contrast, hydrodynamic stress, electrical charging, and repeated centrifugations are unavoidable side effects of flow cytometric sorting and can result in reduced survival and functionality. Time is also an important consideration. Bead purification is generally faster, particularly when large cell numbers are desired, as processing time is not proportional to input size. In contrast, the time required for sorting is directly proportional to input size, potentially affecting cell viability for large inputs. Specific situations (e.g., purification of extremely rare subsets) will require pre-purification steps. Cell yields are often higher with bead purification than with flow cytometric sorting. Last but not least, cost is a major consideration, and in many cases tips the scale in favor of bead purification when it can reach purities consistent with the study objectives. This is particularly true in applications requiring high cell numbers and repeated measurements.

However, flow cytometric sorting has decisive advantages. It offers unequaled flexibility for defining purification criteria. It is irreplaceable when gradient expression levels (e.g., CD44 ^lo vs. CD44^hi) or co-expression of two marks (e.g., CD4 ⁺ CD8 ⁺ from CD4⁻ CD8⁺ or CD4⁺CD8⁻ thymocytes) defines desired populations. Despite major progress in bead purification approaches, cell purity is a second key asset of flow sorting. Although purities greater than 95 % can be achieved by bead purification with appropriate antibodies, they are routinely much lower when the frequency of the target population falls below 10 % of the starting cell suspension. In contrast, flow cytometric sorting can provide highly purified cells (>99 %) even when they are at a very low frequency in the starting population.

Regardless of the procedure chosen, it is important to carefully delineate the experimental plan well in advance. Estimate the size of the starting population (and when applicable the number of experimental animals) by calculating frequency in the target population and expected yield. When working with small cell numbers or precious samples, it is highly recommended to perform pilot studies with similar cells that are more easily replaceable. Slower flow cytometric sorting speeds may be required for infrequent target populations to increase purity and yield.

Initially a 95–99 % yield may seem ideal; however it is useful to know the nature of contaminants. After flow sorting, the operator should “rerun” a small portion of the purified sample, although in some cases additional stains may be needed. For bead-purified cells, purity checks are often done by staining an aliquot of the purified cells and analyzing them by flow cytometry. Antibodies used for purification, whether positive or negative, can potentially impede binding of those used for a purity check. Thus, depending on each specific situation, purities should be verified with antibodies that do not cross-block those used in the purification step.

In this chapter, we provide detailed procedures to prepare purified T cell subsets (from thymus or peripheral organs) using either depletion (“negative selection”) or flow cytometry sorting. These procedures can easily be adapted to specific requirements and reagents. Protocols for bead purification by positive selection depend heavily on the specific reagents being used and will therefore not be described in detail here. As with every antibody-mediated experimental procedure, pilot analyses titrating the amount of antibodies (and related reagents such as beads) are highly recommended to optimize yield and purity prior to proceeding with the actual experiment.

2. Materials

All necessary efforts should be made to keep organism contaminants out. If the end procedure is sterile, all reagents in purifying T cells should also be sterile (and the flow cytometry operator should be notified in advance that the cells must remain sterile during the sort). Medium should be made just before the experiment. If the sterility of the starting reagents is in question, sterile filter (≤45 nm pore size) the final buffer or media before use.

1. Heat-inactivated fetal bovine serum (FBS) (see Note 2).

2. Isolation buffer: 500 mL PBS pH 7.4 (no Ca²⁺, no Mg²⁺), 0.1 % FBS (or bovine serum albumin), 2 mM EDTA pH 8.0.

3. Complete medium: RPMI 1640 medium, 10 % FBS, 100 U/mL penicillin, 0.1 mg/mL streptomycin, 0.292 mg/mL l -glutamine, 20 mM HEPES (see Note 3).

4. Red cell lysis buffer: 8290 mg/L ammonium chloride, 1 g/L potassium bicarbonate, 37 mg/L EDTA pH 8.0.

5. Nylon tissue filters (pore size 100 μm, cut to 2 or 4 cm ²).

6. 1 or 3 mL syringe pestles.

7. 60 mm petri dishes.

8. 5, 15, and 50 mL polypropylene conical tubes.

9. 5 mL polystyrene round-bottom tubes 12 × 75 mm style (flow-sorting procedure).

10. 50 μm filters (flow-sorting procedure).

11. Appropriate fluorochrome-conjugated antibodies.

12. Magnetic beads compatible with appropriate purified antibodies.

13. Rocker.

14. Two curved forceps.

15. Scissors appropriate for dissection.

16. 37 °C incubator with 5 % CO₂.

17. Vertical magnet holding 15 or 50 mL conical tube (DynaMag 15, 50, or equivalent).

3. Methods

3.1. Obtaining Single-Cell Suspensions

Minimizing total purification time is critical in this procedure. Often a trial run with cells is advisable, not only to practice the procedure, but also to get an idea of the efficiency for the specific application. Depending on the desired final purity and the fragility of the cell type to be isolated, two to five times as many target cells should be in the starting population.

1. Prepare the workspace before euthanizing the animal(s). For each organ, set up one 60 mm petri dish on ice with 3 mL complete medium and a 2 cm² piece of nylon filter on the bottom of the dish. Prepare one syringe pestle (1 or 3 mL syringe) (excluding thymus), plus one 4 cm² 100 μm filter per organ, and keep them in a clean container. For the thymus, no syringe pestle is needed.

2. Remove thymus, spleen, and/or lymph nodes (LN)s from animal(s) (see Note 4 on LN selection). Before adding the thymus to the petri dish with medium, roll it on gauze or clean tissue paper to remove any blood, and carefully dissect it from surrounding connective or lymphoid tissue (which tends to stick to the gauze or paper). Do not pull tissue from the thymus and make sure not to damage the thymus or allow it to dry. When removing lymph nodes, carefully remove all fat.

3. For the thymus, take two curved forceps and gently tease apart the organ.

4. For the LNs and spleen, use the rubber end of the syringe pestle to dissociate the cells from the organ tissue, and ensure that you have dissociated any observable clumps as well. It is helpful to use the lid of the dish to store 5 mL additional medium for step 5. At this point, wash the used end of the syringe pestle in this medium, by swirling it in the medium; then discard the pestle.

5. Tilt the dish containing the cells at a 30° angle, draw up the medium with the free cells, and wash this medium back over the dish. Gently pipet up the medium from the dish again and run it through the 4 cm² 100 μm filter into a 15 mL conical tube. Before passing the medium through the filter, use the pipet tip to push the filter half way into the top of the tube, and then pull the pipet tip a few mm back from the filter. Pulling the pipet back prevents the medium from directly hitting the filter and splashing out of the tube. Leave the filter balanced in the top of the conical tube. Next, use the additional 5 mL medium to wash the dish and pass that through the same filter into the conical tube. The dish and filter can now be discarded. Pellet the cells at 150 × g, for 5 min at 4 °C, and decant the supernatant.

6. For spleen cell preparation, lyse red blood cells by resuspending the cell pellet in 2 mL red cell lysing buffer for 2 min on ice. Then immediately, dilute out the red cell lysing buffer by adding 8 mL complete medium. Pellet the cells at 150 × g, for 5 min at 4 °C, and decant the supernatant.

7. Resuspend the organs at 10–20 × 10⁶ cells/mL in complete medium (see Note 5 on expected yields).

8. Count the cells using a dye that distinguishes dead cells (for example: trypan blue). Calculate the cell concentration and total number. Set aside an aliquot (10⁶ cells is ideal) from each starting cell suspension for flow cytometric analysis.

9. In unmanipulated mice, spleen and LN cells can be combined to maximize the size of the starting population. For T cells, when large cell numbers are not needed, LN may be a preferable starting point because of the greater frequency of T cells in LN than in spleen.

3.2. Magnetic Bead Purification of T Cell Subsets by Depletion

Several commercial kits are available to purify target populations (e.g., CD4 or CD8 T cells), most of them using proprietary reagents and materials (magnets). The following procedure uses Dynabeads (LifeTech) to negatively select the desired population. Provided the beads carry the appropriate secondary reagent, Dynabeads can also be used with user-provided antibodies (see Note 6). Volumes and bead numbers are given for 10⁷ starting cells. They should be scaled up proportionally with higher cell numbers. Up to 5 × 10⁷ cells can be processed in a 15 mL conical tube; use several tubes or scale up to 50 mL tubes if processing more cells.

1. Pellet cells to be purified by spinning at 150 × g, for 5 min, at 4 °C.

2. Resuspend cells in 100 μL isolation buffer. If any cell clumping is observed, refilter before proceeding.

3. Add 20 μL FBS and 20 μL kit antibody.

4. Incubate for 20 min at 2–8 °C (wash beads during this incubation and keep on ice until needed; see steps 5 and 6).

5. Wash the beads (to eliminate free antibody that might have been released during storage): first resuspend beads in the vial by vortexing for 30 s. Pipet 200 μL beads into a new 15 mL tube and resuspend in equal volume or at least 1 mL isolation buffer.

6. Put the tube without the lid on magnet for 1 min, remove buffer by pipetting, and discard. Remove tube from magnet and resuspend beads in the same volume of isolation buffer as the original bead volume.

7. After antibody incubation, wash cells by adding 2 mL isolation buffer. Pellet the cells at 150 × g, for 5 min, at 4 °C and decant the supernatant.

8. Resuspend the cells in 800 μL isolation buffer and add pre-washed beads.

9. Incubate for 15 min at 25 °C with tilt and rotation on a rocker at a speed sufficient to keep the beads in suspension.

10. Add 1 mL isolation buffer, and gently pipet up and down five times with a large-bore pipet (5 mL or similar).

11. Put tube without lid on magnet for 2 min. Beads and bead-attached cells will be pulled to the tube wall on the magnet side.

12. Being careful not to dislodge the beads, transfer supernatant containing non-adherent cells to a new tube. For applications that require special media or buffers, wash cells with that medium or buffer as soon as possible after transferring. Discard beads at this point.

13. Count purified cells to determine recovery and take an aliquot (if possible, at least 10⁵ cells) to quantify purity by flow cytometric analysis (see Note 7).

14. Purity from magnetic bead isolation depends on the application (routinely >85 % for CD4 or CD8 T cell purification from LN).

3.3. Magnetic Bead Enhancement Pre-flow Cytometric Sorting

A similar procedure used in Subheading 3.2 can be used to increase the frequency of rare target cells prior to flow cytometric sorting. This is especially useful when starting from high cell numbers (>10⁸) for which sorting times quickly become prohibitive. In this case, because the objective is to remove most of the nontarget cells rather than achieving high purity, significant savings can be achieved by lowering bead numbers and antibody concentrations. While pilot experiments are needed to titer down these reagents, a good starting point is to use 1/8 of the recommended amounts of beads and antibodies while keeping cell concentration intact and all incubations at 4 °C. This “light” procedure routinely achieves >40 % cell purity with very little loss of desired cells.

1. Pellet cells to be run through kit by spinning at 150 × g, for 5 min, at 4 °C.

2. Resuspend cells in 100 μL isolation buffer per 10⁷ cells. If any cell clumping is observed, refilter before proceeding.

3. Add 20 μL FBS cells and 2.5 μL kit antibody.

4. Incubate for 20 min at 2–8 °C (wash beads during this incubation and keep on ice until needed; see steps 5 and 6).

5. To wash the beads: first resuspend beads in the vial by vortexing for 30 s. Pipet 25 μL beads into a new 15 mL tube and resuspend in equal volume or at least 1 mL isolation buffer.

6. Put the tube without the lid on magnet (DynaMag 15, 50, or equivalent) for 1 min, remove buffer by pipetting, and discard. Remove tube from magnet and resuspend beads in the same volume of isolation buffer as the original bead volume.

7. After antibody incubation, wash cells by adding 2 mL isolation buffer. Pellet the cells at 150 × g, for 5 min, at 4 °C and decant the supernatant.

8. Resuspend cells in 975 μL isolation buffer per 10⁷ cells and add pre-washed beads.

9. Incubate for 15 min at 4 °C with tilt and rotation on a rocker at a speed sufficient to keep the beads in suspension. (The 4 °C incubation reduces efficiency, but better preserves cell viability in advance of flow cytometric sorting.)

10. Add 1 mL isolation buffer per 10⁷ cells, and gently pipet up and down five times with a large-bore pipet (5 mL or similar). 11. Put tube without lid on magnet for 2 min.

11. Transfer supernatant containing enhanced homogeneity to a new tube, and wash with complete medium as soon as possible after transferring.

12. Count cells to determine recovery and take an aliquot of 10⁵ cells to quantify purity by flow cytometric analysis (see Notes 7 and 8).

13. Purity will be >40 % with very little loss of desired cells.

3.4. F low Cytometric Sorting

Before sorting cells, consider doing a pre-depletion with beads (see Subheading 3.3), especially when preparing rare cells (<5 % of the input). Although this increases the pre-sort preparation time, it often reduces the total cell manipulation time before downstream procedures. When the target frequency is under 1 %, the pre-depletion strategy should seriously be considered as the sorter error rates can substantially decrease purity in this situation (also see Subheading 3.4.2). The procedure outlined below assumes basic understanding of flow cytometry procedures and does not address operation of the cell sorter itself, which is generally left to a professional operator.

3.4.1. Flow Cytometric Sorting: General Procedure

1. Plan ahead. Consult the flow cytometry facility and inquire about specifics of available equipment and procedures. Discuss the choice of fluorochromes and evaluate the time needed for cell purification and to verify purities. It is also important to choose a sorting nozzle of appropriate diameter, as a general rule at least five times as large as the cells in the starting population. For lymphocytes, a 70 μm nozzle provides adequate speed without damaging the cells. However, a 100 μm nozzle may be better suited when sorting larger cells or when using the double-sort procedure as described in Subheading 3.4.2. For most applications, cells should be kept cold, and it is important to verify that the sorter has a chillable (4 °C) sample holder.

2. Resuspend cells to be stained for flow sorting at 2 × 10⁷ cells/mL complete medium (use 15 mL tube for <6.0 × 10⁷ cells, 50 mL for >6.0 × 10⁷ cells). Before staining for sort, a titration of the antibodies is advisable; 0.25 μg antibody per 10 ⁷ cells is a good starting point. If using the protocol in Subheading 3.3 prior to sorting, ensure that the antibodies used for flow cytometric sorting were not blocked by the antibodies in the bead kit (see Notes 7–9).

3. Do not forget to prepare cell samples for flow cytometer setup if needed. Typically set aside 0.5 × 10⁶ cells per compensation tube. These will be the single-color tubes used to set up the flow cytometric sorter or analyzer for purity checks.

4. Add appropriate antibodies, mix well, and incubate at 4 °C for 45 min in the dark.

5. Wash with complete medium using four times staining volume. Pellet the cells at 150 × g, for 5 min, at 4 °C and decant the supernatant.

6. Resuspend cells to be flow sorted at 2 × 10⁷ cells/mL complete medium in 5 mL round-bottom polystyrene tubes with caps.

7. Filter the starting population immediately before sorting.

8. Define sorting gates in collaboration with the operator (see Note 10).

9. Use polypropylene collection tubes that are large enough to hold the volume collected. For example two to three million cells will fit in a 5 mL polypropylene round-bottom tube; however more than that will need a polypropylene 15 mL conical tube (see Note 11 on plastics in flow cytometric sorting). Add a “cushion” of 50 % FBS and 50 % complete medium to the collection tubes, a total of 1 mL for 5 mL tubes and 2 mL for 15 mL tubes.

10. Upon finishing each sorted sample, immediately fill the collection tube containing the purified cells with complete medium. If the sorted cells have filled the tube, transfer its contents as soon as possible to a larger tube and then fill that tube with complete medium. Diluting the sheath fluid with complete medium improves cell viability.

11. Take a small sample of purified cells and run in on the sorter to verify purity. Depending on the target population, it may be necessary to stain with other markers (e.g., for intracellular proteins, see Notes 7–9 on staining post-sort).

12. Spin cells at 65 × g, for 30 min, at 4 °C. Resuspend in an appropriate volume and count cells (see Note 12).

13. Cells are ready for downstream applications.

3.4.2. Secondary Protocol for Sorting Populations ≥0.01 %

This procedure uses two consecutive sorts to isolate good purity (>95 %) populations from very rare cells (less than 0.01 % of the starting population) if bead isolation procedures are unavailable or inadequate for pre-sort enrichment (see Subheading 3.3).

1. Perform the primary sort following the procedure defined in Subheading 3.4.1 through step 10 with the following changes:

   1. Only use a 100 μm nozzle.

   2. In step 8, use a wide gate to select the entire desired population and maximize yields. Unwanted cells gated in error will be removed in the second sort.

2. After step 10, pellet the cells at 150 × g, for 7 min, at 4 °C and decant the supernatant.

3. Resuspend the cells in a small volume, ≥300 μL complete medium.

4. Perform the secondary sort, using a tight gate that is slightly inside the edges of the desired population. This will increase purity to acceptable levels (>95 %).

5. Follow steps 10–13 in Subheading 3.4.1.

4. Notes

1. It is important to define objectives for target cell purity. High cell purity (99 % or more) is often necessary when purified cells will undergo multiple rounds of cell division (e.g., activating T cells). In this case, preferential expansion of contaminating cells has the potential of significantly altering experimental outcomes. Although high cell purity is always desirable, realistic objectives should be defined based on the properties of expected contaminants and on the dynamic range of assays to be run on purified cells.

2. FBS should be heat inactivated by incubation at 56 °C for 50 min.

3. HEPES can be used to buffer media outside of a CO ₂ incubator.

4. Because LNs from different body areas are to be exposed to different antigenic environments, care should be exercised in deciding whether to pool populations obtained from distinct anatomic sites. Typically, in unmanipulated laboratory mice housed in specific pathogen-free facilities, the major site of microbial exposure is the gut. Consequently, the frequency of activated cells is greater in mesenteric LNs that drain intestinal tissues, and these should generally be processed separately. In contrast, peripheral LNs (including popliteal, inguinal, axillary, and cervical) have few germinal-center and activated T cells, and it is legitimate to pool these populations.

5. Typical cell yields from organs are as follows for a 6–8-week-old female mouse: thymus, 150–250 × 10⁶; spleen 50–80 × 10⁶; and LNs 30–40 × 10⁶.

6. Although commercial bead kits typically include specific antibody mixes designed for purification of homogeneous cell populations, it is possible to use beads with user-prepared antibodies. In such cases, it is paramount to verify that the bead-bound secondary reagent (e.g., streptavidin or anti-IgG) binds the primary antibodies used for purification. In addition, pilot experiments should titer each antibody for its ability to remove undesired cells.

7. When verifying purity of “negative” bead purification, it is essential to avoid staining with the same antibodies as those used for purification (or antibodies that cross-block each other). Otherwise, unwanted cells may escape detection by staining if the epitope is masked by the purifying antibody. Commercial kits often use near-saturating levels of anti-CD8α antibody to purify CD4 ⁺ T cells and of anti-CD4 to purify CD8 ⁺ T cells. In such situations, cell purity can be verified using anti-CD8β, a-non cross-blocking anti-CD4, anti-TCRβ (H57), anti-MHC class II, an Fc blocking antibody, and a live/dead discriminating dye (DAPI for example, see Note 8). Three of the four commonly used anti-CD4 monoclonal antibodies (GK1.5, H129, and RM4–5) cross-block, whereas the fourth one (RM4–4) does not.

8. Live/dead dyes like DAPI remain in sorted cell suspensions and could be detrimental. Instead of these membrane-permeable dyes, use LiveDead fixable dyes that only bind dead cells and are washed away with the excess antibody. Also consider sorting live cells based on FSC and SSC and then testing viability post-sort on a small sample used only for purity check.

9. The fluorochromes (especially PE) used for flow sorting can remain associated with sorted cells for an extended time, unless they are diluted by proliferation. If additional stains are needed, make sure to leave open fluorochromes for these stains.

10. Gating strategies obviously depend on each application. In general, it is advisable to tightly gate the target population. This slightly reduces the yield but minimizes contaminations. It is also recommended, when possible, to exclude cells with unwanted markers by negative gating. Several markers staining with the same fluorochrome can be combined to exclude multiple cell types. This also contributes to the exclusion of cell aggregates (which should be primarily excluded by light scatter-based doublet discrimination).

11. Polystyrene collection tubes have the potential to build up static electricity, which can interfere with sorting, and should be avoided for cell collection.

12. Flow sorters often misreport cell counts, and there is unavoidable cell loss during the centrifugation of sorted cells. If possible, manually recount sorted cells after centrifugation with a live/dead discriminating dye.