Currently, I am trying to do a Bradford Assay on my MS2 capsid protein that is fused with APOBEC1 and I am having very low absorbances for this protein. My Bradford Standards and MS2-APOBEC1 absorbances are shown below. For the SDS PAGE (attached as a PNG) of MS2-APOBEC1(57 kDa biggest band on the last lane), I loaded 16 μL of protein. The buffer that my protein is in is 25mM Tris-HCL pH 7.3, 250mM NaCl, and 250mM Imidazole. Based on my SDS PAGE it shows that I have an abundance of protein, but when I did a Bradford assay I got a lower absorbance than the 2 μg sample of BSA. I have done a Bradford assay using this same buffer on a different protein and I get good results. My main question is what is directly affecting the absorbance of MS2-APOBEC1? What other techniques would be helpful to determine the protein concentration of MS2-APOBEC1?
BSA
2μg = 0.0297 Absorbance
5μg = 0.0878 Absorbance
10μg = 0.2551 Absorbance
15μg = 0.3815 Absorbance
20μg = 0.5015 Absorbance
MS2-Apobec1
20 μL = 0.0073 Absorbance
30 μL = 0.0084 Absorbance
Hi there,
Instead of Bradford why don't you use UV spectra? If you know the sequence of your protein and considering your sample is pure, Absorbance at 280nm is the easier and the more accurate way to get protein concentration
Yes I agree, A280, and use the theoretical extinction coefficient determined from the sequence.
You seem to have a circular argument in deducing that your buffer is okay in the Bradford assay. It MAY be okay, but how do you know the reading was correct for the other protein?
Your readings for your protein are so low (almost zero), it is hard not to think that an error with the sample could be a possible explanation too; or your controls are too high to get accurate readings after subtraction of blanks. What OD readings are you seeing for your controls?
Dear Saul
Does your protein contains many S-S bonds?
Because in my experience, as for example happen for the igG (see attached picture) proteins with strong structural stability may be not completely unfolded when mixed with the Bradford reagent and therefore only a fraction of their residues is stained and detected from the reagent.
AS Dominique Liger suggest you can try UV quantification. If you do not have nanodrop or similar, you can use plastic UV transparent cuvettes (eg.
https://www.sigmaaldrich.com/IT/en/substance/branduvcuvettemicro1234598765?context=product) which are much cheaper than quarz cuvettes, and work also with low sample volumes.
In the unlucky case that you protein sequence do not contain W or Y and you can not determine it concentration by UV, you can try to estimate it performing a densitometric analisys from SDS-page using BSA as standard and Image J to determine band intensity.
You can find some information about it at:
https://proteocool.blogspot.com/2021/09/tips-of-week-13-densitometric-protein.html
good luck
ciao
Manuele
Hey,
Why don't you try lambert beer's law formula to check concentration? We use it for our estimation. Bradford assay actually is a very speedy reaction as compared to biuret Or lowry methods so that might be a reason that there is a problem occuring. Maybe the protein can be assayed using biuret Or lowry Or folin ? You can try anyone and check or else directly putting in formula for A=e.c.l
A= absorbance, C= concentration of desired protein l= path length e= molar extinction coefficient
Hi, just read that arginine residues are a must in protein sequence for Bradford assay because without those aminoacids the dye won't bind to protein and react. You can check whether your protein has arginine residues by sequencing, if there are none Bradford won't work.
Best of luck!
Aparajita
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