MONOCLONAL ANTIBODY STAINING PROCEDURE

I. SAMPLE – one or more of the following preparations

A. Suspension of single cells from tissue (e.g., lymph node, spleen, bone marrow, placental cells)
B. Tissue culture cells
C. Ficoll – hypaque separated mononuclear cells

II. REAGENTS

A. Antibodies

1. Primary antibodies: usually purchased or your own monoclonals

a. if these are conjugated with a fluorochrome (e.g., FITC or PE) use the DIRECT STAINING PROCEDURE b. if the primary antibody is not conjugated to a fluorochrome, use the INDIRECT STAINING PROCEDURE

2. Secondary antibodies: fluoresceinated polyclonal antibodies

B. BUFFER: Phosphate-buffered saline (PBS Ca ²⁺ and Mg²⁺ free) + 2 % newborn calf serum (or 0.2% BSA) + 0.1% sodium azide.

C. Formaldehyde preservative: 2% solution in protein-free PBS

D. PBS: protein and azide free

Buffer and formaldehyde solution can be provided by the FLOW CYTOMETRY CORE LAB if they are not routinely used by the investigator.
III. TUBES

A. Use appropriate 12 x 75 mm polystyrene/polypropylene tubes. Please note that Falcon snap-cap tubes are the only ones that fit on the Becton Dickinson flow cytometer for certain. If you want to stain in another tube type, please transfer the cells finally to those specified here.
B. Mark all tubes with easily readable numbers which correspond to your protocol sheet (see below).
C. Use 1% formaldehyde solution for re-suspension of all potentially biohazardous specimens and cap them.

IV. PROTOCOL

A. Obtain protocol sheets from the FLOW CYTOMETRY CORE LAB (sample attached – please Xerox)
B. Fill out a sheet each time you do an experiment. Include all of the information requested.

SINGLE COLOR PROTOCOL SHEET (Excel required)

DUAL COLOR PROTOCOL SHEET (Excel required)

DIRECT STAINING PROCEDURE

1. Prepare your cells as a suspension of single cells in a manner appropriate for the specimen you wish to examine. Make sure the cells are viable. As the final step, wash at least once with 1 ml of cold BUFFER. Resuspend the cells at 10<7 cells/ml (thus 50 microliters = 5 x 10<5 cells) in cold BUFFER. 2. Meanwhile add 50 microliters of BUFFER and then the appropriate amount of monoclonal antibody to the bottom of tubes. Note: For multi-color staining, add all your fluorochrome-conjugated antibodies at the same time. 3. Add 50 microliters of the cell suspension to the bottom of the tubes. 4. Vortex briefly and incubate 30 minutes at 4^° C in the dark. 5. Wash twice with 1 ml of buffer; centrifuge at 250g for 5 minutes. 6. Resuspend samples in 1 ml of buffer and hold at 4^°C in the refrigerator (or on ice) prior to analysis.

INDIRECT STAINING PROCEDURE

1-5. Process cell samples as above using a working dilution of unlabelled monoclonal antibody.

6. Resuspend cell pellet in 100 microliters of working dilution of the fluoresceinated second antibody.

7. Vortex briefly and incubate for 20 minutes in the refrigerator. 8. Wash twice with ~ 1 ml of buffer; centrifuge at 250g for 5 minutes. 9. Resuspend samples in 1 ml of buffer and hold at 4^°C in refrigerator (or on ice) protected from light prior to analysis.

NOTE: If your samples are Ficoll-Hypaque separated whole blood cells you might still have residual red blood cells in your preparation. These red cells interfere with flow cytometric analysis of lymphocytes and have to be removed by NH<4Cl lysis. See attached recipe and procedure.

PRESERVING PROCEDURE
If cells are not going to be read on the flow cytometer the same day, or they are considered potentially biohazardous, do not re-suspend them after the staining procedure. Stop at Step 5 of DIRECT STAINING or at Step 8 of INDIRECT STAINING, and continue as below.

1. Add to the pellet 0.5 ml of cold protein-free PBS, and vortex; then add 0.5 ml of cold formaldehyde solution (see attached recipes). 2. Vortex again and incubate in the refrigerator. Make sure to keep cells in the dark as exposure to light may cause loss of fluorescence.

AMMONIUM CHLORIDE LYSING SOLUTION – 10X

Reagents	Amount:
Ammonium Chloride, ACS	82.9 g
Potassium Bicarbonate, USP	10.0 g
Ethylenediamine tetraacetic acid (EDTA) disodium salt	0.37 g
Water, glass distilled	qsad 1.0 liter

Adjust pH to 7.2 and keep in tightly closed container at 4^°C. To prepare a 1X working solution (to be used at room temperature), dilute 10X 1:10 with glass distilled water. Keep tightly closed and discard at the end of the day.

LYSING PROCEDURE FOR LYSIS OF RESIDUAL RED BLOOD CELLS

IN FICOLL-HYPAQUE SEPARATED MONONUCLEAR CELLS

1. After the staining procedure, take off as much of the washing buffer as possible without disturbing the pellet.
2. Vortex briefly and add 1 ml of room temperature 1X NH<4Cl lysing solution to your cells, vortex again and expose your cells to it for 2-3 minutes at room temperature. Note: do not exceed this time. 3. Centrifuge for 5 minutes at 200g. Remove the lysing buffer completely and resuspend in BUFFER or 1% formaldehyde solution for analysis.

PREPARATION OF 2% FORMALDEHYDE STOCK SOLUTION (2 METHODS)

METHOD 1:

Formaldehyde preservative – 2% formaldehyde solution in protein-free phosphate-buffered saline (PBS).

Prepare as follows:

Add 2 g paraformaldehyde powder (e.g., Sigma, St. Louis, MO) to 100 ml of 1 X PBS. Heat to 70^°C (do not exceed this temperature) in a fume hood until the paraformaldehyde goes into solution (note that this happens quickly as soon as the suspension reaches 70^°C). Allow the solution to cool to room temperature. Adjust to pH 7.4 using 0.1 M NaOH or 0.1 M HCl, if needed. Filter and store at 4^°C protected from light.

METHOD 2:

Formaldehyde preservative – 2% formaldehyde solution in protein-free PBS.

Prepare as follows:

10% formaldehyde*	20 ml
10 x PBS	10 ml
Distilled water	70 ml

* 10% formaldehyde solution (e.g., Polysciences, Warrington, PA, Ultrapure, Cat.#04018), depolymerized paraformaldehyde, EM grade, methanol-free solution.