I would suggest to dilute your sample and use the BCA assay instead of Bradford, which is less compatible with SDS and urea. Alway use your buffer as control (blank) to subtract the "noise" deriving from the buffer.

Next time, for your protein extraction, I would also suggest you to prepare an alternative version of your buffer (50 mM Tris-HCl, pH 6.8, 2% SDS, 8 M urea, 10% glycerol, 2% mercaptoethanol) without the bromophenol blue, since the blue is used to visualize sample loading on SDS-PAGE gel, but it is not necessary in protein extraction procedure. Probably the absence of the blue color could help you in determining protein concentration with less "color" interference.

Good luck for your experiment

When you said failed, in what way ? No UV max at 280nm ? Bromophenol blue contains aromatic chromophores absorbed at 280nm too. Thus will masked your protein absorption if its diluted.

5% SDS and 3M urea are compatible with BCA assay. dilute your sample to 3M urea or less and also make mercaptoethanol less than 25mM by dilution. then determine the concentration and multiply by diluting factor to get original concentration...

Other way is to use commercial kits available (there is one from BxxRad at least) that are compatible with detergents, which will be your major problem in that sample. But remember to prepare standards in the same buffer conditions.

Manyak's solution will be likely also work, so give a try as BCA assay is so common that every lab has the reactants.

It can be done in two ways:

1) Precipitate a volume (say 100ul) of ur protein and resuspend it in 0.1N NaOH, and then proceed for any spectrophotometric estimation method. Calculate the concentration of ur actual protein sample accordingly.

2) Dilute ur protein atleast 20-50 times and go for for any spectrophotometric estimation method, (for blank use 20-50 times diluted buffer)

I usually go with method 2 , where i dilute my proteins and estimate using Bradford method.

All d best

How do you fail using 280 nm?

My suggestion would have been using absorbance at 280 nm measuring spectra and perhaps correcting for buffer contributions.

I know the extinction coefficient will be slightly affected by the urea (Pace et al 1995 Protein science). The SDS might be scattering light and perhaps the BPB absorbs at 280. Is some other component absorbing/scattering at 280 nm?

You should exchange the buffer that contains your protein(s). The simplest way to do this: run a gel filtration column. The pros: you get your proteins in the buffer you desire free of salt. The cons: your proteins will be diluted and you should concentrate them.

An alternative: use centriprep or centricon filters, You will exchange the buffer and you will concentrate the protein at the same time. The pros: it is faster and relatively easy. The cons: you should check that your proteins do not run dry.

Hope it heps.

Hi,

you can use one of the desalting columns (like illustra NAP 25 (5) from GE helthcare, or similar one, it takes just 20-30 min and you will dilute your sample ~2.5-3 times), in that way you can do buffer exchange to the buffer which will not interfere with protein determination.

Thanks everyone for the suggestions. Anyway, I accidentally found Nanodrop 1000 in my lab. Does anyone measure the protein concentration with the similar sample buffer as I stated in my question? Thanks.

Dear Kokwai

Its good that you found the solution. I would suggest not go with Nanodrop as i have experienced false estimation using this.

I wud strongly suggest u to use a known standard concentration in same buffer and reconfirm the standard concentration using Nanodrop, if comes OK then proceed.

All d best

the best way to mesure a protein concentration in such buffer is to use : NI™ Protein Assay A Non-Interfering™ Protein Assay.

Good luck !

Are the BCA or RcDc Protein Assay (Bio-Rad) only the best for preparing a known standard?

My proteins are not purified one. I just want to measure a crude protein concentration before loading it into gel.

The reason I said I failed the A280 using cuvett because the values I had got were the same even I tried using different dilutions (diluted it using the same buffer).

I blank it using the SDS-8M urea buffer (without crude protein) and I just wanted to do a simple confirmation test which I used back the blank sample to measure OD. I was supposed to get zero but I was not, there showed some value.

Dear Kokwai, you should take the nanodrop values with care. At least in my case I found that is quite reliable for assessing the range of concentration of my samples (5 or 10 mg/mL) but not to get an exact value of the actual concentration. The "feature" can be more problematic in your case, as you have Urea and SDS in your buffer.

RcDc assay is good as it is still linear when SDS and/or Urea is present, you should be adviced that your standard (BSA, usually) MUST be prepared in the same buffer, and your blank MUST also be the neat buffer, not water.

I guess that you measured the standard again after blanking and the ABS value was not zero... what was the value? If it was very low then you can take the resultant concentration with care, but if you objective is no more than a loading check... it should be in principle good enough.

Good luck!

Go for Bradford Assay. Its easy and cost effective and at the same moment best method as per my experience.

I would suggest to dilute your sample and use the BCA assay instead of Bradford, which is less compatible with SDS and urea. Alway use your buffer as control (blank) to subtract the "noise" deriving from the buffer.

Next time, for your protein extraction, I would also suggest you to prepare an alternative version of your buffer (50 mM Tris-HCl, pH 6.8, 2% SDS, 8 M urea, 10% glycerol, 2% mercaptoethanol) without the bromophenol blue, since the blue is used to visualize sample loading on SDS-PAGE gel, but it is not necessary in protein extraction procedure. Probably the absence of the blue color could help you in determining protein concentration with less "color" interference.

Good luck for your experiment

yup off course no need to use Bromophenol blue in your extraction buffer. Bradford works well and is cheaper than BCA.

i think there is no need to check protein prior SDS-PAGE running. Do your work in same conditions (amount spores and volume of sample buffer added) and run SDS-PAGE along. Different concentration of BSA was also run in gel seperately for standard. After Staining, calculate protein with Densitometer.

When you suggest some protein assays please think about the method before just just suggesting according to the price. And do not hesitate to read about the assays first. Bradford assay will not work as in the mentioned buffer you have a 2% SDS concentration which is indeed interfering with the assay!

Dilution in water would surely solve your problem.

We use a buffer similar to yours and measure our protein content using the Advanced Protein Assay from Cytoskeleton. To my knowledge it's the only assay suitable for these buffer contents (especially high urea and sds concentration). It's a very simple procedure, just dilute the concentrated buffer 5-times, then add your sample, wait ten minutes and read out at 495nm, it's both suitable to cuvettes and to 96-well plates. Good luck!

Hey Rubina,

I talked about two buffers in my post:

The first one meant the buffer containing your sample (proteins of interest): we use 7 M urea, 2 M thiourea, 40 mM Tris, 4% CHAPS, protease and phosphatase inhibitors.

The second buffer mentioned in my post meant the 5x-concentrate that is part of the Protein Assay we use.

Sorry, if you got mixed up by this...

Greetings, Julia

This Non-interfering protein assay kit works well for estimating protein concentration in nasty buffers:

http://www.gbiosciences.com/PDF/Protocol/NI_Protein_Assay.pdf

It worked well for me with urea-extracted mouse brain lysates (7M urea, 2M thiourea, 4% CHAPS, 30 mM Tris, pH 8.5).