Protein Assay
Coomassie blue method of Bradford, using a commercial reagent from Bio-Rad
Very often we need to determine the concentration or amount of protein in a particular solution.
We may be purifying a protein and need to follow the amount, or trying to determine how much
of a sample to load into a gel or put in to an enzyme assay.
One commercial version (made by BioRad) uses the Commassie Brilliant Blue dye-binding assay
developed by Bradford. It works like this:
The absorbance maximum for an acidic solution of the dye Coomassie Brilliant Blue G250 shifts from 465 to 595 nm when it binds to proteins (primarily to basic and aromatic
amino acid residues). When we have a set concentration of this dye in solution, we can
measure the amount of protein by reading the absorbance at 595 – the more protein, the
more molecules of dye will be absorbing at this wavelength.
To carry out the assay, one simply mixes the protein with the dye solution -- it is a simple
one-step assay. A standard curve with known concentrations of protein (usually, but now
always, BSA) establishes the relationship between the amount of protein added and the
absorbance at 595 nm. By subjecting measured volumes of the unknowns to the same
assay conditions (amount of dye and volume of assay) and determining the A595, one can
determine the concentration of protein in the original sample (amount of protein/vol)
There are four protocols recommended by the manufacturer. They differ primarily in the volume
in which the assay is carried out. The one we used most of the time is the Micro assay
The Micro Assay
Set up tubes:
protein + H2O
0.8 mL
Add dye concentrate
0.2 mL
MIX WELL, wait 5’, read A595
This really is all there is to the assay, but some issues arise that you must consider, which will be
discussed further below:
1. You must have a standard curve every time you do the assay, using a protein solution of
known concentration.
2. Your samples must be tested in such a way as to give absorbances in the readable range of the
assay. (once you have bound all the dye in solution, you could keep adding protein forever
and you would get no increase in absorbance.
3. Interfering substances may need to be considered
Standard curve in the Micro Assay:
To carry out the BioRad assay:
1) Obtain some BSA (bovine serum albumin) standard that is 0.5 mg/mL (any concentration is
OK, this is just easy)
2) Figure out how many uL of the standard are needed to get the ug indicated below. Then
figure out how much water needed to bring the volume to 0.8 mL
3) Make triplicate samples of the protein + H2O.
4) Make an arbitrary sample for each unknown –such as 10 uL + 790 uL of water
Amt of
protein (ug)
BLANK
1.0
2.0
4.0
6.0
8.0
10.0
100.0 *
Sample 1
Sample 2
*
Vol Standard
Vol of H2O
#ul of
0.5 mg/mL stock
(to 0.8 mL)
0
0.8
10 uL
10 uL
0.79
0.79
A595
mean
SD
0.000
Normally you will not do this one – just do it the first time so you can see what it looks like
5) Add 0.2 ml of the Conc Reagent to each tube (standard and unknowns) and MIX WELL
(You can do this in glass tubes and vortex. Or, you can do the reaction in the plastic cuvettes,
and mix by putting parafilm over the top and inverting. Do mix, and mix immediately. Do
not add all the reagent, and let them sit. The reagent is dense, and will not combine equally
with the protein in your assay.)
6) Now, Before you go to the spectrophotometer, look at the unknowns. Is the color similar to
the color in the tubes containing less that 10 ug? If it is too blue, make another sample using
much less (you can use a smaller volume, or dilute and use the same volume) If it is too
light, make another using more.
7) Play around making protein samples until you get three that appear to be in the 1-10 ug range
(they do not have the be the same dilution & volume!)
8) Now, read all the samples, and record the A595s.
9) Plot the data with A595 on the y axis and the amount (ug) of standard in the assay on the x
axis. Draw the best line you can (it is not ideal, actually to use the computer for this, because
the behavior of this assay is NOT LINEAR (even though it looks like it)
10) Read the unknown A595 values, and determine the amount of protein in the sample.
11) Determine the concentration by taking the amount / volume used. (If you made a dilution in
order to get the sample in range, say a 1/10 dilution, you will have to multiply that back,
because your sample concentration is 10X more concentrated than what you determined from
the diluted sample.
12) You can use the mean, and figure the protein concentration from that, but you need to look at
your triplicate A595 values and see that they are close before you do that. If you need to know
how much the assay really varied in terms of protein concentration, you will need to
determine the concentration from each A595 that you read, and then figure the mean (and
standard deviation) of that.
Comments, notes and advice about this assay:
Calculating concentration vs reading it from a graph
Although it is true the spec reads concentration, in this assay we are not reading protein
concentration directly, but rather the effect that the protein in the assay has on the dye. It is
helpful to think of the protein added to the assay in terms of the amount (the # of ug) added to the
assay (1 mL volume and 1/5 dilution of dye). That way you can easily determine the
concentration by dividing amount (ug) you found divided by volume (uL) that you put into the
assay. (Correct for dilution in the volume or after the concentration has been calculated)
People often plot protein assays as absorbance vs. protein concentration. The difficulty with this
is that it is not always clear what they mean, and it is not always easy to figure out how to
calculate back to your sample. It depends on how you put the assay together. It’s valid, but you
have to be very careful to interpret it correctly, and I have seen people spend hours and hours
agonizing over this -- it is not worth it. For this class, please do it as described here. (I do not
want to spend hours trying to validate your results, or arguing with you about what your method
means.)
Linearity (or lack of same)
This assay looks linear, but is actually not linear. If you look at a small enough part of the curve
(such as the narrow 1- 10 ug window) it sort of is, and people like linear functions because they
can draw neat lines with rulers, or get computers to plot and read the graph for them. That’s fine
as long as you don’t let the computer do things you wouldn’t do from the actual data. Watch for
inflections, and be aware that a calculated slope/intercept will not be valid if it is not in the
“readable range” of the data.
Note that the blank is zero protein, but does contain the dye reagent. The CBB does absorb
some at 595 without protein around, and you do not want to see this (it just raises the
background, by an absorbance of about 0.045).
Tubes, cuvettes and mixing
Mixing of the samples is critical. The concentrated reagent is very dense. If it is not mixed well
with the protein in the solution, the assay results will be horrible.
The glass 12x75 tubes are very convenient for setting up the reaction because they are easy to
handle, easy to vortex and it is easy to use your own personal spectrophotometer (a.k.a. your eye)
to determine where the useful range of your curve is. You will need to transfer to a cuvette to
read. You may read in glass or in plastic disposable cuvettes. (don’t use quartz, because it is not
good for them, and they are expensive). You can do the reaction in the plastic cuvettes, which
saves materials and time, but make sure you get sufficient mixing.
You can not used data that is uninterpretable
Although the assay is not strictly linear, it is nearly so, and the data needs to be within this range
to be usable. If the ODs are out of the “linear range” when you read them, you may still be able
to get a protein determination, as long as there is some degree of dependence of A595 on protein
(and if you use the real values from the graph, not a calculated slope and intercept) . However, if
your absorbance is too high, and the curve too flat, you need to re-do the assay using less of the
sample (less volume, or a dilution of the sample). You cannot just dilute the assay tube.
If you have a lot of protein, you may bind all of the available CBB -- this is the upper end of your assay.
Once the CBB is all bound, any additional protein sits around. You cannot dilute the whole reaction, because
that will just dilute the completely bound dye. SO … if a sample contains too much protein, dilute it.
An alternative, of course, is to use a tiny little amount. This is OK too, just keep in mind that the
micropipettors lose both precision and accuracy in the very low range. It is usually better to
make a dilution with larger volumes (20 -100 uL) than to try to use 1 uL or less (unless you have
very small amounts of sample available).
Interfering substances
Samples may contain substances which interfere with the Bradford assay (detergents, high salt
there is a whole list of things). Some raise or lower the absorbance of the blank (they interact
with the dye itself) and some affect the binding of the protein to the dye. Interfering substances
(and the concentrations that are a problem) can be looked up, or you can determine empirically
whether and how much they interfere. If a critical set of samples has an interfering substance in
the buffer, you can use a small enough volume that the concentration is below the level that
interferes (as long as your protein concentration is high enough). On the other hand, you may
need to find a way to get rid of the interfering substance or find another assay.
Safety and Disposal
The concentrated reagent contains phosphoric acid, and is very acidic. Be careful with it. It
would be appropriate to wear goggles or safety glasses when setting up the reaction, since you
will be vortexing these solutions.
Also, in order not to contaminate the stock at all, take some out – enough to do your assay (and
them some), and when you are done, discard it (do not return it to the stock bottle).
For the CR and for the samples, discard into the tub of water near the sink. We will adjust the
pH of the solution in the tub before discarding down the sink.