I need help with making precoated ELISA plates, especially preserving. I never did this before, but it will save me a lot of time. What I found with google search is: incubate with capture antibodies O/N, then next morning, decant, block for an hour, decant again, then add Sucrose (2%-5%), then preserve with a desiccant (CaCl2), not mentioned the amount needed, Any insight?
Also, how long will they be good to use, before they get expired? I'm coating them with 19C7 antibodies to detect Troponin I.
Thanks
When preparing pre-coated ELISA plates for my work I would coat with my antibody for 2 hours at 37oC or overnight at 4oC in carbonate/bicarbonate coating buffer at pH 9.6. Following aspiration of the coating solution I wash with TBST. Following this I would either block for 1 hour at RT leave un-blocked depending on your ELISA protocol. I then use a 3% Trehalose solution to stabilize the plates, coating with a volume 10 ul more than the a.b coat for 1 hour at RT. After this stabilization step the Trehalose is tipped off, plate patted dry and then dried in an incubator at approx 30oC overnight. I would then store my plates either in ziploc bags or vacuum seal bags with a silica desiccant at 4oC.
Use Polystyrene strips.
carbonate/bicarbonate buffer (pH9.6) for coating. After blocking and then drying, use desiccant pack (usually comes with electronics) in a Ziploc bag to store the strips.
See more details in the link below
http://www.abcam.com/index.html?pageconfig=resource&rid=11422
When preparing pre-coated ELISA plates for my work I would coat with my antibody for 2 hours at 37oC or overnight at 4oC in carbonate/bicarbonate coating buffer at pH 9.6. Following aspiration of the coating solution I wash with TBST. Following this I would either block for 1 hour at RT leave un-blocked depending on your ELISA protocol. I then use a 3% Trehalose solution to stabilize the plates, coating with a volume 10 ul more than the a.b coat for 1 hour at RT. After this stabilization step the Trehalose is tipped off, plate patted dry and then dried in an incubator at approx 30oC overnight. I would then store my plates either in ziploc bags or vacuum seal bags with a silica desiccant at 4oC.
Just like Goodwin G. Jinesh suggested, I used to do the coating in the carbonate buffer, but at some point I tested the coating in PBS (phosphate buffered saline) and, since I observed no difference in reference signal values, used the PBS ever since
Hi,
I agree with previous comments. In my case, I use in particular a 50 mM carbonate/bicarbonate buffer pH=9.6 (in which you can add 0.10 g/L thimerosal as preservative) to coat my capture antibodies on polystyrene microtiter plates (NUNC Maxisorp) with an overnight incubation at 4°C. Of note, it is possible to keep the plate longer in this condition (with coating buffer at 4°C) if you can't perform the next steps right after the overnight incubation but for short time period only i.e. a few days (less than one week). I never tried the procedure you mentioned.
Hope it helps!
Forget all this blocking voodoo. Coating at pH 9.6 in 0.1 mol/L carbonate buffer with 2 – 5 µg/mL overnight is mostly sufficient. The solid phase is overloaded anyhow (binding capacitiy is at 100 ng/cm²). Store the coated plates frozen until use. Blocking (if there are some free binding sites available) happened during the first washing with PBS Tween 20 by the detergent. Blocking is only necessary for commercial assays where storage of the test kit at room temperature for longer periods is required. If you will do really a blocking, use a protein, which is proven free of your antigen. Do not be surprized by checking BSA.
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