I am using Nanodrop to measure protein concentration in lysate since it is fast and only requires small amount of sample. When I use water as blank, the result showed two high peak at 230 and 280 nm, and the absorption in 280 nm is 27 mg/ml. However, when I used the lysis buffer as blank, the result only showed 1 wide peak at 260 nm and the protein concentration now is only 12 mg/ml. My lysis buffer contains Tris-HCl, DTT, EDTA, protein inhibitor, triton x-100 and 8M Urea and lyse the cell by sonication. I do not know why using water as blank then my samples showed high peak at 280 nm which supposes to be protein but not in case of lysis buffer? I am new with protein and have no experience at all.
Buffer additives, such as DTT, Triton x-100, have high absorbanc in the range 260-280nm. It often interfers protein quantification. I think it is not a good approach to measure the concentration of protein from lysates. Nucleic acids, contaminant proteins and buffer additives affect the accuracy of your measurement significantly. Moreover, the concentration obtained from cell lysate is the total protein concentration.
It is better to run SDS-PAGE for the soluble fraction of the lysate, then the concentration of your protein can be estimated from band corresponds to your target protein.
Protein concentration in the lysate can also be measured using one of several fairly easy and sensitive colorimetric assays, including Bradford, BCA, Lowry, and others. Some of the components in the lysis buffer will cause interference with some of the assays, so check the product literature for tolerances to lysis buffer components. The lysis buffer should be used as the blank, not water. Because the lysate is quite concentrated, a very small portion of the lysate can be diluted and used to measure the protein concentration. These assays require only a few µg of protein.
I agree with Alemu and Adam.
I will add that you will likely find that your quantification is unlikely to be linear using this approach. You could test this by either diluting an extract into the extraction buffer, or spiking an extract with BSA in extraction buffer. The presence of DTT and non-protein contaminants may hide the signal of the actual protein in your sample.
I've had issues using UV280 to reliably quantify even purified individual proteins in DTT containing buffers.
You'll probably be better off using a quantification kit, as Adam suggested. Thermo and Biorad have a good table for identifying a kit that may be compatible with your extraction buffer (https://tools.thermofisher.com/content/sfs/brochures/TR0068-Protein-assay-compatibility.pdf) (http://www.bio-rad.com/webroot/web/pdf/lsr/literature/Bulletin_6852.pdf).
Your inclusion of Triton and DTT might complicate matters, depending on their concentration. Very few quantification kits are compatible with both at high concentrations. As long as the Triton at/below 1% you might get away with Pierce 660.
Brian is right. The best option would have been Pierce 660 unless the Triton concentration is below 1%. If indeed the concentration is around 12 mg/ml, you may dilute your sample to lower the Triton concentration. Another option for complex buffers is using tryptophan fluorescence (Wiśniewski 2015, Anal Chem, Article Fast and Sensitive Total Protein and Peptide Assays for Prot...
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Dear Hoa, Nanodrop certainly is not an ideal choice to measure protein concentrations, because the absorbance spectrum of various amino acids differs and is based upon the UV behavior in a solution with different types of molecules along with your proteins. So, it is a havoc.
You must know that protein readings around 280 nm are mainly based on the net content of aromatic amino acids tyrosine (Y) and tryptophan (W) within a protein, whereas, if your protein is rich in phenylalanine (F), which is also an aromatic amino acid, but it absorbs best at 260 nm than the 280 nm. So it is a matter of protein composition as well. If you are using water as blank here, (which is wrong, as you need to blank with your lysis buffer) and this water is giving you clear peaks for various spectra, so it is heavily contaminated, just throw it.
I prefer for you to do this quantification business through fluorometric means, mainly through the Qubit device, as it is well manageable and convenient as compared to BCA or Bradford assays and is quite sensitive too.
https://www.thermofisher.com/order/catalog/product/Q33211
After blank with lysis buffer, protein concentration will drop definitely as compared to blank your nanodrop with the water, as your device is adjusted to zero in accordance with the heaviness of lysis buffer now.
Also, it is important to know that proteins absorbs at 205 nm with great sensitivity than the 280 nm, so conventional spectrophotometers may be more useful to quantify proteins, though the process is laboursome, but this approach is more useful if protein concentration is below 100 ug / ml. This 205 nm is a safer wavelength, as most of the contaminants absorb from 230 - 240 nm range.
You can replace the DTT from your lysis buffer with 2-MEA, as if the DTT not dissolved well or stored well, it is not very effective and may interfere with UV-based observations like the nanodrop. 2-MEA is more stable and uniform.
Degradation products result from the enzymatic treatments and lysis procedures, absorb UV light immensely so absorbance increases dramatically, which is a fact that Nanodrop won't cope with this and you have false high values then in terms of absorbance.
Also you should remove 8 M Urea from your lysis buffer, as Urea-treated proteins are never measured through nanodrop. For Urea case, you need to quantify proteins through BCA, Qubit etc. As, 8M Urea is a very heavy denaturant and unfolds or linearizes the proteins, so protein is no more in its parent conformation and this may alter absorbance spectra through nanodrop.
I hope these all facts about protein measurement will certainly help you for your further future endeavors with proteins.
Sincerely....
Dialyze your protein into a different buffer to remove the urea and detergent or use a different assay for protein measurement. And you should always blank against the buffer that your protein is in.
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