While encapsulating ADMSCs in sodium alginate, providing a 3D environment for the cells, I come across different viability rates. Trypan blue shows a lower viability rate, as low as 50%. On the other hand, I get a viability of 90% using a dual fluorescent dye; using acridine orange (AO) and propidium iodide (PI). I'm really puzzled. Why do I get different rates with the very same scaffold?
Is there any other way through which I could confirm the accurate viability?
Hi, Hanie, the fluorescent dyes will give you much accurate results. I think your problem with Trypan bule is methogological. Here many such points have been discussed, just look here: https://www.researchgate.net/post/Trypan_blue_destroy_cells_during_counting_Why
Good luck!
Over staining is often a factor for fluctuation between samples. Detection sensitivity is a 2nd factor. We usually need calibration by using one and the same sample
If the Trypan Blue concentration is too high, it can also stain viable cells a light blue, which can skew your viability counts. Try different Trypan Blue dilutions until you get one that gives more consistent results.
Trypan Blue can kill cells by itself so depending on the concentration you use and the amount of time you put the cells it may give you wrong results. Try other viability assays such as Alamar Blue and MTT (but always normalize your results to total DNA) Since they test metabollic activity and this may be influenced by amount of cells you have.
50% viability in sodium alginate seems not quite an accurate result to me. I have assessed the viability and proliferation potential of hADSCs embedded in alginate hydrogels by MTT and by flow cytometry after LIVE&DEAD fluorescence staining (calcein AM and PI): http://www.biomedcentral.com/1472-6750/12/35
The unexpected viability rate decrease was a result of excess sodium present in the solution. I purified my alginate and made a solution using DMEM. Alas! That was the trick and my viability stayed constant after that!
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