How should the dosage of anti-human CD3 monoclonal antibody for flow cytometric detection of human T cell surface be titrated?

For a small lab, I suggest buying 50 tests instead of 100 tests, unless you run more than 10 samples per week. The 100-test format looks cheaper per unit, but repeated freeze-thaw cycles after the antibody is opened will cause the titer to drop quickly, ultimately leading to more waste. When aliquoting, work quickly under light-protected conditions and dispense into PCR tubes or low-protein-binding cryogenic vials. Aliquot each tube according to one week's usage, for example, 5-10 μL per tube, add sealing film, and store at -20°C. Do not store at -80°C, as some fluorescent antibodies will aggregate or precipitate. Take one tube and place it at 4°C protected from light for use, and try to use it up within one week. If only 2-3 tests are used per experiment, aliquot into even smaller tubes to avoid bringing the entire vial back to temperature. Mix gently before aliquoting; do not vortex fluorescent antibodies, to avoid bubbles and protein denaturation. Also record the lot number and aliquoting date.

Do not titrate solely based on the volume in the instructions. My approach is based on the final staining system: after red blood cell lysis, resuspend the white blood cells to approximately 1×10^6/100 μL, and perform a gradient of the antibody from 0.125, 0.25, 0.5, 1, 2, and 4 μL/test, with the total staining volume per tube fixed at 100 μL, at 4°C protected from light for 30 minutes. During analysis, gate the lymphocytes and calculate the staining index, i.e., the MFI of the positive population minus the MFI of the negative population divided by twice the SD of the negative population, and select the lowest concentration at which the staining index reaches a plateau. Common causes of positive population shift include antibody excess, excessively high voltage, incomplete red blood cell lysis, or failure to perform Fc blocking. The 5 μL/test in the instructions is only a suggested starting point; different red blood cell lysis batches and cell numbers will cause changes, so it is more stable to base it on cell number, but the volume must be fixed.

For whole blood red blood cell lysis samples, background control is more critical than antibody amount. I would first use fresh whole blood and the same lysis buffer, add Fc receptor blocking reagent for 10-20 minutes, then stain with CD3 FITC. For gating, first use CD45 and side scatter to gate lymphocytes, then gate CD3+ to avoid interference from granulocytes and debris. During titration, also set up isotype control and FMO tubes; the FMO tube does not include CD3 FITC, while all other reagents are the same, and it is used to determine the positive gate boundary. If the positive population shifts overall, first confirm whether the voltage and compensation are fixed, and do not use the isotype control as the sole basis for adjusting voltage. It is recommended to re-titrate each new batch of antibody or new lysis buffer, and at least perform one concentration curve, rather than directly continuing to use the amount from an old batch.

If I had to decide whether to use volume or cell number, I would combine both: use cell number as the basis, but fix the staining volume at 100 μL. This is because antibody binding is mainly affected by cell number, antigen density, and free antibody concentration; after red blood cell lysis, the white blood cell count varies greatly among different samples, so using volume alone can easily lead to insufficient antibody when there are many cells. For titration experiments, it is best to use fresh blood from the same donor or fixed whole blood quality control material, prepare multiple aliquots at one time, with each tube containing approximately 1×10^6 cells. After gradient staining, observe the MFI of the positive population, the background of the negative population, and the separation distance, and choose the lowest concentration that can completely separate the populations without increasing background. A shift in the positive population may also come from antibody aggregates, so centrifugation or filtration can be performed before use, especially after fluorescent antibodies have been repeatedly frozen and thawed. Therefore, it is more reliable to first perform a concentration gradient and then fix the amount used.