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Weird BSA standard curve for Bradford assay - (Jun/12/2009 )

hai! I was trying to quantify protein of whole brain tissue homogenate. the buffer used in the sample is RIPA buffer. So, when i did the serial dilution for BSA standard, i used RIPA as diluent. But i found that the standard curve look so abnormal n weird. As i know that, detergent can affect the Bradford assay. But then, what can i use to dilute my BSA to draw my standard curve? and wat can i can do with the sampe which is already mix with the RIPA buffer? is there any other idea to help out this? because i only have bradford reagant.
Anyone knows bout this? can someone help? thanks!

-kathyliaw93-

you can try diluting your samples and ripa diluted bsa with water and performing a micro-bradford determination.

-mdfenko-

I routinely use RIPA to lyse cells for various applications. I agree with mdfenko. Perform a microassay and just dilute your samples/standards in water (or PBS), but add a volume of RIPA equal to the sample volume to each of your standards.

For example...
I use a total of 1 ml for the Microassay. (800 Ál water or PBS + 200 Ál Bio-Rad Protein Assay Reagent)

If performing a Bradford on 5 Ál of sample then add 5 Ál of RIPA to the Blank or Standard + ? Ál volume Standard + remaining volume up to 800 Ál with Water. Then add Protein Reagent, mix, incubate, and read.

I hope this isn't too confusing!

-Roo-

Hmm,what are the importance and the uses of BSA Standard curves?
Any links to view?

-IANIRON-

I found this thread by searching and am glad to be reminded that detergent throws off the Bradford assay.

My standard curve looked fine, as it was BSA diluted in TE. I never tried to make a standard curve in any detergent-containing buffer.

However, based on this standard curve, I was told by the Bradford assay that all my samples were within 2-fold of each other. But then the bands on the gel (GAPDH) looked like there could be more like a 20-fold difference between the highest and lowest concentration.

The Bradford assay did rank the samples in almost the right order from highest to lowest concentration -- but the relative quantities were all off. I could also detect this by looking at the concentrations the Bradford told me -- and then doing the 2-fold or 1.5-fold dilutions which should bring them all to the same concentration -- only to find that it hardly had any effect on the relative differences.

Is this the sort of thing expected from samples lysed/diluted in RIPA buffer, which is only about 1.2% detergents? (No loading buffer/sample buffer had been added)

-MDavies-

MDavies on Nov 9 2009, 03:08 AM said:

I found this thread by searching and am glad to be reminded that detergent throws off the Bradford assay.

My standard curve looked fine, as it was BSA diluted in TE. I never tried to make a standard curve in any detergent-containing buffer.

However, based on this standard curve, I was told by the Bradford assay that all my samples were within 2-fold of each other. But then the bands on the gel (GAPDH) looked like there could be more like a 20-fold difference between the highest and lowest concentration.

The Bradford assay did rank the samples in almost the right order from highest to lowest concentration -- but the relative quantities were all off. I could also detect this by looking at the concentrations the Bradford told me -- and then doing the 2-fold or 1.5-fold dilutions which should bring them all to the same concentration -- only to find that it hardly had any effect on the relative differences.

Is this the sort of thing expected from samples lysed/diluted in RIPA buffer, which is only about 1.2% detergents? (No loading buffer/sample buffer had been added)


you should always either add buffer to your standards or, at least, run buffer blanks to determine the offset caused by the buffer.

-mdfenko-

Roo on Tue Jul 7 19:39:47 2009 said:


I routinely use RIPA to lyse cells for various applications. I agree with mdfenko. Perform a microassay and just dilute your samples/standards in water (or PBS), but add a volume of RIPA equal to the sample volume to each of your standards.

For example...
I use a total of 1 ml for the Microassay. (800 Ál water or PBS + 200 Ál Bio-Rad Protein Assay Reagent)

If performing a Bradford on 5 Ál of sample then add 5 Ál of RIPA to the Blank or Standard + ? Ál volume Standard + remaining volume up to 800 Ál with Water. Then add Protein Reagent, mix, incubate, and read.

I hope this isn't too confusing!


Hi all,

I am new to the forum and I had a question regarding the Bradford assay. I use a recipe for lysis buffer to lyse insect cells and I want to measure the protein concentration by bradford microassay. As such, I set up my bradford similarly to what Roo mentioned but using a final volume of 500ul

i.e. For the standard curve, BSA @ 1mg/ml(1ul,2ul,3ul......) + lysis buffer (5ul) + water (394ul, 393ul, 392ul.....) + Bradford reagent(100ul) . For my sample I add 5ul(already in lysis buffer) in 395ul water + 200ul Bradford reagent

When I read my sample concentration off the standard curve, what is the concentration units - is it ug/ml or ug/ul?
Also, what dilution factor do I need to multiply by to get the actual sample concentrations?
If using a 96 well plate, is the linear range the same as if using cuvettes?


Forgive the simplistic questions but I am getting confused trying to figure this out!

Thanx

-asp1979-

are you really adding 100ul reagent to the standards and 200ul to the sample or is it just a typo?

assuming it is a typo, you can evaluate the standard curve as mass (rather than concentration), hence, you have 1ug, 2ug, 3ug,...

you can determine concentration afterward (Xug/5ul).

-mdfenko-

mdfenko on Fri Feb 4 20:25:34 2011 said:


are you really adding 100ul reagent to the standards and 200ul to the sample or is it just a typo?

assuming it is a typo, you can evaluate the standard curve as mass (rather than concentration), hence, you have 1ug, 2ug, 3ug,...

you can determine concentration afterward (Xug/5ul).


Thanks. Yes sorry that ws a typo. it should be 100ul.

-asp1979-

what i have been doing and what seems to make sense is to dilute your lysis buffer to say 1:50 and use that as blank and diluent. Then dilute your samples to 1:50 as well with water and results should be comparable.

i do this even with a detergent compatible assay.

-azrael201-