To detect the CAR-positive rate of CD19 CAR-T cells, should Protein L or an anti-idiotypic antibody be used?

If used for clinical sample QC, do not rely on a single dye. For release testing, we use flow cytometry with anti-scFv/idiotype antibodies to detect CAR positivity, and simultaneously use qPCR or ddPCR to measure VCN, with the two methods corroborating each other. Flow cytometry antibodies require methodological validation: specificity using non-transduced T cells and CD19-negative tumor cells; precision with at least 3 replicates from the same donor; upper/lower limits of quantification. For antibody suppliers, choose those that can provide GMP or RUO lots, lot-to-lot COA, and no sodium azide interference. For clinical QC, it is not recommended to directly add preservative-containing conventional flow cytometry antibodies to the system; centrifugation and washing are preferable. Protein L can serve as a rapid indicator for process development, but for clinical release testing, anti-idiotype or CD19 recombinant protein is recommended, as it is more specific for CD19-CAR.

We ultimately used recombinant anti-FMC63 idiotype antibodies, available from both ACRO and Sino, and customizable from Genscript. When selecting, don't just look at the CAS number; check whether the recognized epitope falls within the scFv framework region or CDR. Some antibodies are sensitive to CAR conformation, and the signal may change after fixation/permeabilization. We recommend doing three things: titrate with CAR+ and CAR- cell lines; compare live-cell staining and fixed-permeabilized staining; repeat 3 times with the same lot to check CV. For lot-to-lot stability, we tested 3 lots and the MFI variation was within about 10%, but this varies by vendor, so it's best to ask the supplier for CoA and stability data. Lead time is generally 1-2 weeks; custom anti-idiotype antibodies may take 4-6 weeks. The price is higher than Protein L. Based on 100 test pricing, anti-scFv is usually 2-5 times that of Protein L, and the custom multiplier is even higher.

Protein L itself binds to immunoglobulin kappa light chains and generally stains FMC63-derived scFv, but common causes of high background include: dead cell adhesion, nonspecific Fc receptor binding, failure to perform Fc Block during staining, and excessive antibody concentration. You can first try CD16/CD32 blocking, viability dye to exclude dead cells, and titrate down the Protein L-FITC concentration, adding 2% FBS or BSA to the wash buffer. In our previous experience, the main cause of high background was the addition of serum to the culture system or carrier proteins in IL-2, which led to weak-positive smearing. For clinical sample QC, Protein L background is more heterogeneous in lymphoma patient samples, so it is recommended to at least switch to anti-scFv for validation. If the background cannot be reduced, do not force its use.

Besides anti-idiotypic antibodies, biotinylated CD19-Fc or CD19-His recombinant proteins can be used to detect CAR binding capacity, with a principle closer to functionality. Note that soluble CD19 may cause internalization or competition, so staining should be performed at 4°C for a short time. We have performed dual staining with CD19-Fc and anti-scFv, and the correlation between the two is good, with a Pearson r of approximately 0.95 or above. In terms of lead time, CD19 recombinant proteins are mostly in stock and are cheaper than anti-idiotypic antibodies. When clinical sample background is high, co-culture of CAR+ target cells to measure cytokines/killing can be used as a functional supplement. It is recommended to first perform small-scale testing: select a CAR+ cell line or a known positive donor, perform antibody titration and blocking experiments, and confirm that the signal can be competed away by unlabeled anti-scFv.