I have problem quantifying my protein. Actually, I am working with Magnetic nanoparticles (Dynabeads) to bind my RpsA protein with a Binding buffer (ph 8), with differents conditions of temperature and incubation time.
At the moment of quantification of supernatants I use Nanodrop and Bradford, however this results are incongruent between them. For example, In Nanodrop, the best temperature is 4ºC (10min) (all lectures) but Bradford results 37ºC (10min), both decrease at 30 minutes. I don`t know which is the true result.
My protein is RpsA (62KDa), I bound 15ug (bradford) to 15ul of dynabeads
I appreciate your help.
Personally, I would go with the Bradford data and use a standard curve of either BSA or gamma-globulin (note, they give different slopes). I would avoid incubations at higher temperatures due to the possibility of proteolysis. You could always verify by SDS-PAGE/Coomassie staining.
I'm going to use your advices.
Thank you so much for your help,
Personally, I am quite conflicted about that. Apart from dialyzing against pure water and drying all these techniques are in my opinion approximations. Of course with most proteins one can encounter in a lab this not an option or something one would like to do.
For a Bradford one has to keep in mind, that the result is always obtained against the behavior of BSA and a dye, eg. Coomassie. The behavior of your protein might be different leading to higher or lower value. The same is true for the NanoDrop or any photometrical measurements, because primary sequence, folding, solutions properties play a role and can lead to "incorrect" values.
In your context it is "just" a qualitative comparison of the same protein under different conditions. I would quantify with both techniques, if the number of sample allow that, and put equal amounts on the same SDS-PAGE/Coomassie gel. Based on the result of staining intensity, you might be able to judge, in which sample the discrepancies occur. This way you can find optimal conditions, but I would be careful to take the protein amounts as granted, especially if the protein amount influences further experiments.
Dear Felix Ernst, I really appreciate your advice, In my opinion you're right with the fact of using SDS-PAGE in order to identify the origin of discrepance.
Thank you so much!
Indeed, if you're not interested in the absolute amount of protein but rather the relative amount as you compare one binding condition to the next then wouldn't just one of the assays be sufficient? In other words, if you graphed the results for a given amount of nanoparticles for a given amount of protein at one temperature but varying time on the x-axis, then couldn't the y-axis be the % remaining protein of the initial amount? For example, if the readout from the Nanodrop gives x amount of starting protein then the result you want is what percent of that amount is remaining after the binding treatment and irregardless of what the true starting amount was. If you want to check for a discrepancy between the two methods then you might need an orthogonal method such as RP-HPLC at 215 nm because SDS-PAGE, if stained by Coomassie, is re-affirming what the Bradford assay found using Coomassie; essentially doing the Bradford assay again but in a solid matrix rather than liquid. If you want the absolute amount then you might have to do total amino acid analysis. Here's an example: http://www.alphalyse.com/quantitive_aaa.html
A Bradford assay is definitely more sensitive for protein quantification than the Nanodrop.
Both methods are different and both results can be wrong. It takes time to determine closest quantities by a particular method for a real concentration. But once this will be done for a specific protein at specific conditions you always can use it for further measurements.
UV method is used in Nanodrops. Most common UV method is a measuring of a sample at 280 and 260 nm, a result can be recalculated by Warburg formula C (UV)=1.55 * (A280) - 0.78 * (A 260). The final result can vary basing on three factors: a contamination by nucleotides, an extinction coefficient and a measured quantities should be in a linear part of an absorption curve. To avoid variations two more wavelength can be measured 230 and 320, and results can be used for corrections. I believe Nanodrops uses second option. But an extinction coefficient should be determinated any way. I'm not sure about temperature's dependence 4 and RT but some shift can be.
Bradford's method is based on affinity of Coomassie blue G-250 particles to specific composition of AA at acidic conditions at RT. This combination has max absorption at 595 nm. Definitely a result is depended on AA sequences, dye dilution, acidity of reagent, a measured quantities should be in linear part of a curve and a curve should be calibrated. Most important here are two points: to find proper protein for a calibration and detergents should not exist in buffer. For example BSA calibration can give up to 1.5 more concentrated sample than a real one. Tweens and Tritons bind coomassie with conversion to blue color too, I didn't check another detergents, but it's easy to check for each particular one.
BCA is convenient in a case detergents are used in buffer.
I like Dzmitry answer, because it really gets into the different technical aspects. i recommend to everyone, who thinks "Bradford is the best" to actually try enzyme kinetics and their analysis, where precise unbiased quantification is important. Folding changes and different flexibilities can influence the intercalation of Coomassie into proteins, so that if comparing homologous variants of enzymes one can easily get into trouble if you rely on Bradford. I don't say NanoDrop is without flaws, but if buffer conditions are the same, in my experience it is more reliable than Bradford.
The A280 method is based on the absorption of Trp, Tyr and Cys residues and hence the actual abundance of these residues in a protein. Molar absorptivities are (I think!) 5500, 1490 and 125, respectively, so you can determine a molar absorption coefficient for your protein if you know its aa composition. I believe this is described in Pace, CN et al, 1195, protein science 4, p2411 (but I haven't actually checked that). Coomassie staining has similar caveats (only reaction with certain aa residues), but I don't know any formula that can be used to obtain absolute amounts of protein for such staining.
All you need to know about different methods of protein quantification
https://www.labome.com/method/Protein-Quantitation.html
Protein concentration measurement based on A280 and Bradford(using Coomasie reagent) are both accurate although at low concentration A205 measurement will yield better result. But the downside is A205 will have interference from other UV absorbing peaks, but this technical difficulties can be overcome by using nanodrop method at 205nm. For low concentration the best reagent is BCA as the assay method is based on copper chelation agent. However, BCA assay involves an incubation period of 30 minutes at 37 degrees.
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