Hello. So I've been trying to oxidize antibodies gathered from a modified CHO cell line. So I've been able to confirm that the antibody does bind to the target antigen with western blot. So I know that they function as they should. But upon oxidation and conjugation with hydrazide-biotin (sigma aldrich) the antibodies won't bind to streptavidin-HRP. So any subsequent western blot would come up completely blank. My oxidation protocol so far has been to try different molar concentrations of sodium periodate (5, 10, 20mM conc.) and to vary the oxidation time at 10 minutes and 30 minutes (reaction at room temp, on wrist action shakers). The oxidation is carried out under slightly acidic conditions of 5.6 pH using sodium acetate (50mM) (buffer exchange using columns and PBS). Hydrazide-biotin in final solution is 5mM (2 hours, room temp, on wrist action shakers followed by desalting using Zeba columns). So far I've been having zero luck. Any thoughts? Please let me know. ANY help is appreciated. Thank you!!
10 min even at 5 mM is a long time and you are most likely oxidizing other residues. Try 3 min at 2.5 mM. Are you adding the periodate using a concentrated solution? Are you quenching the reaction? I prefer running the reaction at pH 7.0 using phosphate buffer.
Hello,
Thank you for your suggestion! I am using 100mM NaIO4 as stock solution, and diluting it to make 50mM. I did consider quenching the solution using polyethylene glycol at 0.250mL/mL of oxidized antibody solution. I have tried at pH of 5.6 using NaOAC and around 7.2-7.4 using PBS. Is quenching absolutely necessary? A published paper in our lab didn't specify quenching so we forewent in favor of just desalting and buffer exchange using millipore micron 10 filters and zeba columns.
Quenching is absolutely necessary given that the periodate is in excess and the rapidness of the reaction.
Thank you! I'll be sure to incorporate the step in my next experiment.
Thank you again.
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