Jump to content

  • Log in with Facebook Log in with Twitter Log in with Windows Live Log In with Google      Sign In   
  • Create Account

Submit your paper to J Biol Methods today!
Photo
- - - - -

Weird BSA standard curve for Bradford assay


  • Please log in to reply
9 replies to this topic

#1 kathyliaw93

kathyliaw93

    member

  • Members
  • Pip
  • 2 posts
0
Neutral

Posted 12 June 2009 - 10:28 AM

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!

#2 mdfenko

mdfenko

    an elder

  • Active Members
  • PipPipPipPipPipPipPipPipPipPip
  • 2,805 posts
133
Excellent

Posted 15 June 2009 - 07:12 AM

you can try diluting your samples and ripa diluted bsa with water and performing a micro-bradford determination.
talent does what it can
genius does what it must
i do what i get paid to do

#3 Roo

Roo

    Enthusiast

  • Active Members
  • PipPipPipPipPip
  • 31 posts
0
Neutral

Posted 07 July 2009 - 11:39 AM

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!

#4 IANIRON

IANIRON

    member

  • Members
  • Pip
  • 1 posts
0
Neutral

Posted 20 July 2009 - 08:45 PM

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

#5 MDavies

MDavies

    member

  • Active Members
  • Pip
  • 8 posts
0
Neutral

Posted 09 November 2009 - 12:08 AM

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)

#6 mdfenko

mdfenko

    an elder

  • Active Members
  • PipPipPipPipPipPipPipPipPipPip
  • 2,805 posts
133
Excellent

Posted 09 November 2009 - 09:05 AM

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.
talent does what it can
genius does what it must
i do what i get paid to do

#7 asp1979

asp1979

    member

  • Members
  • Pip
  • 3 posts
0
Neutral

Posted 04 February 2011 - 11:07 AM

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

#8 mdfenko

mdfenko

    an elder

  • Active Members
  • PipPipPipPipPipPipPipPipPipPip
  • 2,805 posts
133
Excellent

Posted 04 February 2011 - 12:25 PM

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).

Edited by mdfenko, 04 February 2011 - 12:26 PM.

talent does what it can
genius does what it must
i do what i get paid to do

#9 asp1979

asp1979

    member

  • Members
  • Pip
  • 3 posts
0
Neutral

Posted 05 February 2011 - 01:38 AM

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.

#10 azrael201

azrael201

    member

  • Active Members
  • Pip
  • 27 posts
0
Neutral

Posted 09 February 2011 - 03:43 PM

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.




Home - About - Terms of Service - Privacy - Contact Us

©1999-2013 Protocol Online, All rights reserved.