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I know that the bradford assay is a very standard way of measuring protein concentration after e.g. a purification. However, in the lab that I work in now they normally only use nano drop at the 280nm wavelength.

How trustworthy is this concentration? Say that I e.g. get 2.5 mg/mL, could I really trust this?

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That depends strongly on your protein and how exact you need this concentration. Both tests (Bradford and the measurement at 280nm) only do an approximation.

The measurement at 280nm relies on the interaction of aromatic aminoacids with the UV light. If you have a protein with few or no aromatic amino acids, your measurement will be wrong.

The same is true for the Bradford assay. Here you form a complex between Coomassie Brilliant Blue and unpolar and cationic side chains of the amino acids. This test is more precise, since it includes more possible side chains, but is still dependend on the composition of the protein.

More precise is the BCA assay, since it utilizes the reduction of copper ions by the peptide bond of the protein. What you could do is run a BCA assay (or at least a Bradford assay) from your sample and measure the same sample on the Nanodrop at 280nm and compare the results.

You can find an overview over the methods here and here.

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  • $\begingroup$ I will aks the same as i did on Joe Boyle's answer, based on his common on extinction coefficient: when using the extinction coefficient for the protein in Nanodrop, how does it then compare to other methods? Does it then get more reliable $\endgroup$
    – CuriousTree
    Jan 16, 2017 at 15:56
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    $\begingroup$ @CuriousTree knowing the exact epsilon value for a particular protein helps, but you're still stuck with the basic limitation of UV-based measurements - it entirely depends on the number of aromatic amino acids. The thing to keep in mind for any kind of quantitation assay is that you need to have a relevant standard. Not every protein is the same as, or even similar to BSA. $\endgroup$
    – MattDMo
    Jan 16, 2017 at 22:52
  • $\begingroup$ @CuriousTree When you know your protein very well, it might help, but in the moment you are analyzing protein mixtures (or something complex like cell lysates), this will not help. $\endgroup$
    – Chris ♦
    Jan 18, 2017 at 11:16
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The nanodrop should have an option that allows you to input an "extinction coefficient". This is a measurement of how much one mole of your protein will absorb at 280nm. The nanodrop will use this value to give you an accurate concentration reading.

To find the extinction coefficient for your protein, put its whole sequence into this server http://web.expasy.org/protparam/.

A few tips:

  • Use the exact buffer your protein is in as a blank.

  • Repeat your nanodrop reading a few times using a new pipette tip each time to ensure reproducibility

  • The purer your protein the more accurate your concentration reading will be! If there is a major contaminant in there, you will not get an accurate concentration reading.
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  • $\begingroup$ thanks! Do you think this is comparable to Bradford then? because its specific for the protein? $\endgroup$
    – CuriousTree
    Jan 16, 2017 at 15:52
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    $\begingroup$ Neither technique is perfect but they can both give good estimates. Bradford depends on your protein interacting with the Bradford reagent in a similar way to the known standard, unfortunately I don't have much experience using it. If you need a precise concentration and have the time/resources then quantitative amino acid analysis would be my suggestion: link $\endgroup$
    – Joe Boyle
    Jan 16, 2017 at 17:55
  • $\begingroup$ @JoeBoyle if my peptide has no/few aromatic residues and I still want to use Nanodrop, will the use of Expasy Protparam extinction coefficient input for Nanodrop become accurate enough? $\endgroup$
    – littleworth
    May 17, 2022 at 9:21
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    $\begingroup$ @scamander If your protein has no aromatics you won't be able to use 280nm absorbance but if it has a low number you can still use the extinction coefficient. As usual, the quality of the reading will depend on how pure your protein is. $\endgroup$
    – Joe Boyle
    May 18, 2022 at 10:07
  • $\begingroup$ @JoeBoyle Thanks so much for your answer. Mine is a peptide (comes in powder). How do you measure protein purity? $\endgroup$
    – littleworth
    May 19, 2022 at 0:15
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If you are dealing with a pure protein where there is nothing else present that will absorb at 280nm and if the E(1%, 280) of the pure protein is known or may be calculated from the amino acid sequence, then A-280 measurement is a very accurate, fast, nondestructive method of determining protein concentration.

It is certainly much more reliable than any method that uses a standard curve, where the color produced may depend on the standard protein used.

It also has the additional advantage that the protein may be fully recovered after the measurement is made, often a consideration when dealing with precious samples.

It must be emphasized that the protein needs to be pure for this method to work. When dealing with impure protein samples, the 'rule of thumb' is that a 1mg/ml solution will have an A-280 of 1, or E(0.1%, 280) = 1. But of course other substances such as DNA or toluene (in a yeast extract, maybe) may absorb at 280nm and play havoc with your readings.

A-280 measurements, being nondestructive, are usually considered adequate for monitoring eluates from chromatographic columns.

If the amino acid sequence is known and you wish to use the Perkins (1986) method to calculate the E(1%, 280) it is probably worth bearing in mind that this method takes no account of a bound prosthetic group (such as haem or tightly bound NAD), which could also significantly contribute E(1%, 280) value.

Some good refs

Perkins, S. J. (1986). Protein volumes and hydration effects. The calculation of partial specific volumes, neutron scattering matchpoints and 280-nm absorption coefficients for proteins and glycoproteins from amino acid sequences. Eur. J. Biochem. 157, 169 - 180. [Pubmed] [pdf]

Gill, S. C. & von Hippel, P. H. (1989). Calculation of protein extinction coefficients from amino acid sequence data. Anal. Biochem. 182, 319 - 326. [published erratum appears in Anal. Biochem. (1990) 189, 283].

Sober, E. K. & Sober, H. A. (1970). Molar extinction coefficients and E (1%, 280) values for proteins at selected wavelengths of the ultraviolet and visible region. In Handbook of Biochemistry. Selected Data for Molecular Biology, 2nd edn. Sober, H. A., Ed. pp C-71 - C-98. The Chemical Rubber Company, Cleveland, Ohio.

Edit

I can give an example of the Perkins method, if anyone is interested.

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  • $\begingroup$ Best answer for purified proteins. With 280nm absorbance you can calculate a reasonably correct extinction coefficient from your sequence and then make a quick measurement on your nanodrop (using the UV module, not the protein module). This is more accurate than Bradford where you have a sequence-dependent error that you cannot correct for. Strictly speaking it is only valid for the unfolded state. You can measure in 6M Guanidinium chloride to be sure. In my experience, this is usually not necessary though and the value is pretty close for folded proteins, too. $\endgroup$
    – Raik
    Dec 27, 2018 at 14:17
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The choice between colorimetric and direct quantification at A280 depends on both the protein to be quantified and the buffer being used. Generally A280 works well when you have a purified protein solution and the protein is well characterised. Entering the correct extinction coefficient will improve the accuracy of the concentration calculation. If using A280 it is also worth checking that your buffer does not absorb highly in the 280nm range. You can do this by blanking with water and running your buffer as a sample. If your buffer absorbs highly then you may be better off selecting another method.

For cell lysates and uncharacterised proteins you are better off with a colorimetric assay such as Bradford, BCA or Pierce 660. These can be run in microvolume but take care in the preparation, especially with assays such as Bradford that may have a lot of particulates.

Also, as you are measuring against a standard curve for colorimetric assays, the more similar your standards are to your sample, the better the results.

Andy

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