Module 6. Protein Concentration Determination
1.
Introduction
When conducting biochemical research, it is often necessary to accurately determine the concentration of a protein
solution. Several methods are available, each having particular advantages and disadvantages. The method of choice in
this module is the commonly used Bradford Assay. This is a great opportunity to learn this practical and useful
biochemical laboratory skill.
In Module 3, you designed an experiment to set up a standard curve for BSA using the Bradford assay. This curve showed
the relationship between Absorbance at 595 nm and BSA concentration. As Beer’s law suggests, you found that solutions
with higher protein concentrations absorbed more light.
In Module 4, you purified the DHFR protein and you will determine the concentration of protein in the samples from
Module 4. Doing the Bradford assay for these samples alone would tell you very little unless you have a standard for
comparison. For example, if a sample with an unknown protein concentration gives an absorbance value of 0.4 in the
Bradford assay, what is the protein concentration of the unknown? If you have a standard solution with a known protein
concentration that also gives an absorbance of 0.4, then you can reasonably assume that the unknown sample has the same
concentration as the standard since they give the same absorbance result in the Bradford assay.
Suppose that you have a number of samples, and their concentrations vary (just like in Module 3). It would be useful to
have a number of standards that span the full range of likely concentrations of your protein samples. That is, in fact, a
standard curve! You will prepare a Bradford Assay for a series of standards of known concentration of BSA, ranging from
low to high concentration (think back to Module 3) as well as for your protein samples from Module 4. You will then
measure the Abs595 on all standards and your samples from Module 4. You will then plot absorbance versus
concentration for each BSA standard to create the standard curve. Using this standard curve, you can read (interpolate) the
concentration of your samples given its absorbance reading in the assay.
2.
Purpose of the lab
In Module 3, you designed an experiment to construct a standard curve for the Bradford assay. In Module 4, you purified
the protein DHFR.
Your assignment is to design and implement a protocol to establish the standard curve for BSA and determine the
concentration of protein in the samples from Module 4.
You will again make solutions with different known BSA concentrations, add each to a colorimetric reagent, and measure
the resulting absorbance values. You will draw a graph that shows absorbance on the y-axis and protein concentration on
the x-axis. This is called a standard curve, a kind of a ruler that tells you what absorbance values to expect for specific
protein concentrations. You will use this standard curve to determine the concentration of a protein sample; the instructor
has the protein samples stored. This will be done by mixing the protein sample saved from Module 4 with the colorimetric
reagent in exactly the same way as was done for the standards. The resulting absorbance will be compared to the standard
curve. You will be able to find the concentration on the standard curve that gives you the same absorbance as your
unknown. From this information, you can determine the concentration of your samples from Module 4.
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3.
Agenda for the Day
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4.
You must have worked through Math Moment problems before coming to class. Upon entry, each student shows
their work to instructor to receive points.
Presentation on Bradford Assay.
In class, groups review Math Moment Problems to prepare for experimental design.
Experimental design: Groups discuss and agree on experimental protocol. Each student writes down a step-bystep protocol for the experiment.
Conduct the experiment in groups. Each student individually records data in their personal laboratory notebook.
Clean up
Background
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Useful information can be found Chapter 3A under section” Buffer dilutions” and 3B under section “The
Bradford Assay” in the textbook (Boyer); review Module 3 as necessary.
BioRad Protein Assay Protocol has a sample protocol – model your protocol on the “standard protocol for
microtiter plates” that starts on page 6: http://www.bio-rad.com/webroot/web/pdf/lsr/literature/LIT33.pdf
Coomassie Blue has three different colors depending on its solvent acidity or double bond location.
Red (cationic)
Acidic environment
A(max)=470 nm
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Green (neutral)
Neutral environment
Blue (anionic)
Basic environment
A(max)=595 nm
When a protein binds to Coomassie molecule, a protein-dye complex will form. In this complex, the dye is
stabilized in the unprotonated (anionic) form.
The more protein molecules bind to the reagent, the deeper the color and the higher the absorbance will be.
The resulting absorbance is measured at 595 nm. In this lab, we will use the closest available wavelength 564 nm.
You will need to prepare a standard curve by plotting average blank-corrected 564 nm measurements for each
BSA standard vs. its concentration or amount.
You will use the standard curve to determine the protein concentration of your protein samples from Module 4.
All assays have limits. Amounts of substance below some minimum will be undetectable. Beyond some
maximum amount or concentration an assay becomes saturated, that is, increases in amount or concentration do
not affect absorbance. We generally try to work within the linear range of an assay, that is, where absorbance is
directly proportional to concentration.
Often a sample is so concentrated that when you assay the prescribed volume of sample the result is off scale –
the assay reagent is saturated. To fix this problem you need to dilute the sample. You may make a dilution or
multiple dilutions of your protein solution, make sure to record how you did the dilution, and take the dilutions
you used into account when calculating the final concentration. Read the concentration from the standard curve
and then multiply the result by the dilution factor you used to get the actual concentration of the sample.
Make multiple dilutions of your unknown. If the absorbencies of your protein sample do not fall into the linear
range of your standard curve, make different dilutions of your samples and repeat the experiment. You will need
to repeat the standard curve as well if you repeat the sample analysis. Both must be done simultaneously.
When samples are so concentrated that you cannot pipet a small enough amount accurately, you may have to
conduct serial dilutions.
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5.
Math Moment
1. There are two ways to prepare dilutions: parallel and serial dilutions. You need to prepare a standard curve for the
linear range 125-1000 µg/ml. Use the parallel dilution method (C1V1=C2V2) to set up the following dilutions 0, 0.05, 0.1,
0.2, 0.3, 0.4 and 0.5 mg/mL to cover the desired range. Volume of each dilution needed is 50 uL. Fill in the blanks in the
table below with your answers.
Concentration of standard
protein
2.0 mg/mL
C1
2.0
2.0
2.0
2.0
2.0
2.0
2.0
D1
D2
D3
D4
D5
D6
Volume of
standard solution
needed
V1 (µL)
Diluted concentration
to be prepared
C2 (mg/mL)
Volume of
dilution needed
V2 (µL)
Volume of
solvent needed
V=50-V1 (µL)
0
0
0.05
0.1
0.2
0.3
0.4
0.5
50
50
50
50
50
50
50
50
2. What is your smallest volume that you can comfortably measure with your P20 pipettor? Can you measure all of
needed volumes (see table, especially the first row). If your answer is no, what can you do to solve the problem? Could
you perhaps use a different final volume (V2)?
3. Another way to make dilutions called serial dilutions. In this
method, you make one dilution, then use part of the first dilute
solution to make an even more dilute solution, and so on. This is very
practical method when you need to prepare a solution diluted by a
large factor. In the example shown in the figure below, the dilution
factor is 10. Do you understand why? The reason is that V2/V1 = 100
µL/10 µL = 10. In the figure, the researcher transferred 90 µL of
solvent (buffer) to each of the 5 wells. Then he took 10 µL of the
standard stock protein solution and added it to the first well and
mixed. Then he took 10 µL of solution in the first well and add it to
the second well and mixed. Then he took 10 µL of solution in the second well and added it to the third well and mixed
and so on. Based on this information and the figure, fill out the empty cells in the table below. Assume that the standard
stock protein solution had the concentration of 10 mg/mL. Use the units listed in the column headings. Note, the data for
the first well is filled out as an example.
Well #
Volume of protein solution
added to well (V1) (µL)
Volume of buffer added
(µL) to well
Total volume in well
after dilution is made
(V2) (µL)
Final protein concentration in well (C2)
(µg/mL)
1
2
3
4
5
10
90
100
1000
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4. Let’s say that you are trying to cover the range described in question 1: 125-1000 µg/ml using the serial dilution
approach. Does the serial dilution in question 3 cover this range? If not, what change could you make to your serial
dilution approach to cover the linear range 125-1000 µg/ml. Hint: Think about changing the dilution factor, which means
changing V1 and V2. Make a decision on what your V1 and V2 are going be and fill out the table below with your new
values. Assume again that the standard stock protein solution has the concentration of 10 mg/mL. Note: The range of
concentration you cover does not have to fit exactly into the desired range, you should just make sure that it is covered as
well as you can. For example, the lowest standard solution in your assay does not have to be exactly 125 µg/ml.
Well #
Volume of protein solution
added to well (V1) (µL)
Volume of buffer added
(µL) to well
Total volume in well
after dilution is made
(V2) (uL)
Final protein concentration in well (C2)
(µg/mL)
1
2
3
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5. A researcher will set up an assay to measure substance X. When X is mixed with assay reagent a complex is formed
that absorbs light at wavelength 400 nm (some other colorimetric reagent, not Coomassie). The researcher will use a
spectrophotometer requires that she puts 2 ml volume in each cuvette. To get the right proportion of assay reagent to
sample, we make our sample volume 0.5 ml and add 1.5 ml color reagent to each tube. The stock solution of X is 5
mg/mL. The researcher made the following standard solutions from the stock, see table below.
Table 1. Example of how to plan a standard curve. The concentration of protein in the stock solution was 5 mg/ml. This
example is for illustration purposes only.
amount of substance X (mg)
0 (reference)
0.01
0.02
0.05
0.1
0.2
0.5
1
1.5
2
volume of stock solution (µl)
0
2
4
10
20
40
100
200
300
400
volume of buffer (µl)
500
498*
496*
490
480
460
400
300
200
100
*It is common to use pipettors that give us volumes that are accurate to no more than 2 significant figures.
She mixed 500 µL of each standard with 1.5 mL of the colorimetric reagent and obtained an absorbance at 400 nm for
each. The data is shown below. The relationship is not perfectly linear, rather it shows a typical extinction pattern where
the higher concentrations are not fully in the linear range.
She then prepared three assay tubes for the unknown sample, containing 500 µl, 50 µl, and 5 µl sample, respectively. She
added enough buffer to make the volume 500 µl (just like the standards) and then added 1.5 mL of colorimetric reagent.
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Suppose the solutions gave absorbance readings of 0.86, 0.12, and 0.01,
respectively. The last absorbance is off scale, of course. The intercept
should be zero, but we cannot count on very low absorbances giving us
sufficiently accurate readings. Use the readings from the two tubes
(0.86 and 0.12) to calculate the concentration of the original unknown
solution. Remember that the X-axis of your standard curve is in units
of mg. Are the results identical? Which result should you use or
should you take an average? Note: Using the one absorbance reading
that falls closest to the middle of the linear range gives the most
accurate results. In the example above the center is an absorbance of
0.5, corresponding to 1 mg of substance. At the upper end of the range
the color reagent approaches saturation, so that not only do you have
less resolution among absorbance readings, but the reagent is less
sensitive to differences in protein concentration.
6.
Supplies Provided
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7.
Ice buckets and ice
BSA protein standard at a concentration of 2.0 mg/mL - KEEP IT ON ICE
Bradford Reagent (already diluted to 1X)
Eppendorf tubes
Micropipettors and tips (multi-channel pipettors)
96-well microtiter plates
Plate shaker
UV -VIS microtiter plate spectrophotometer
Experimental Design
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Take a look at the range of concentrations you used and resulting absorbencies in Module 3 to generate your
standard curve. If your data points formed an initial increase followed by a plateau, you should make the highest
concentration in your standard curve lower so that all data points fall in the linear range.
The instructor will give you a standard solution of BSA (protein) that has the concentration 2.0 mg/mL.
In addition to establishing the standard curve, this time you will add the unknown protein samples in wells on the
same plate as the standards. Unknown proteins need to have exactly same amount of liquids in the wells as
standards. In other words, you will add 10 μL of each solution into a well containing 200 µL of Coomassie
Reagent as described in the Biorad protocol. All measurements need to have as constant factors as possible, same
temperature, same incubation time, same light exposure etc.
Prepare multiple dilutions of your unknown protein sample to optimize the chances that one of the dilutions gives
you a protein absorbance that falls approximately in the middle of your standard curve. If your standard protein
sample absorbance does not fall approximately in the middle of the linear range of your standard curve, you will
need to make new dilutions and run BOTH the unknown samples and the standard samples and run the entire
assay again.
It helps to have a reasonable estimate of ranges of concentrations of sample that one can expect. Even with such
an estimate it is good to prepare samples with a range of dilutions, in case a sample is so concentrated that its
absorbance readings are out of range. For us a reasonable estimate might be 0.05 mg/mL to 0.5 mg/mL.
You must prepare each experiment (well) in triplicate (or you can choose more replicates). Remember to account
for the volume needed when preparing your protocol.
The recommended wavelength for Bradford Assay is 595 nm. The plate reader in the lab does not read that
specific area but very close, 564 nm. Please, in your lab notebook and procedure, write down this wavelength.
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8.
Common Mistakes and Some Advice
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9.
Remember good pipetting technique. Use the correct pipettor (P20, P200, or P1000) for the volume you
measuring. Please, handle pipettors gently.
When adding the reagent to the wells, do not empty the pipettor to the second stop (that makes lots of bubbles).
As you are expelling liquid, touch the tip on the sidewall of the well.
Once you have the absorbance values, take a picture of the computer screen and then type the values in a data
sheet in excel. Plot your data in the lab class, bring an electronic device or laptop that allows you to do that. This
allows you to check that your standard curve is linear.
Remember that the standard curve can either show absorbance versus amount of substance or absorbance versus
of concentration of sample.
A point of confusion is the distinction between BSA concentrations of the standard solutions (the dilutions you
make in eppendorf tubes) and the BSA concentrations in the wells of the microtiter plate. You will make a set of
solutions (your standards) in Eppendorf tubes. You will then take 10 uL of each solution and mix with 200 uL of
Coomassie reagent in a well in a microtiter plate. After you mix 10 uL of BSA solution with the Coomassie
reagent, the BSA concentration in the well is lower (by 21-fold) compared to the BSA standard in the eppendorf
tube. When preparing your standard curve, if you express your x-axis as a concentration, be clear about whether it
is the BSA concentration in the well in the microtiter plate (after combining protein with Coomassie Reagent) or
the BSA concentration of the standard solution you prepared in your Eppendorf tube. You can also display your
x-axis as amounts.
You must make sure that your unknown absorbance value is close to the middle of the linear range of your
standard curve. If not, the assay must be repeated.
If you have to repeat the assay, you must repeat BOTH standards and unknowns. You must, of course, change the
dilution of the unknown sample so that the absorbance falls in the correct range.
Vocabulary
BSA, standard, incubate, aliquot, reagent (dye), colorimetric spectroscopy, 96-well plate, microtiter plate, Bradford
reagent, Coomassie reagent, unknown, dilution factor, linear range.
10.
Safety
You must wear safety glasses when conducting the experiment. You must never eat or drink in the laboratory. You will
need to wait in line to use the plate reader. Please, be patient when waiting for your turn to use the plate shaker and plate
reader. Be gentle with the plate reader and plate shaker, these are delicate instruments. Any observed violations of these
rules will result in lower final grade and/or removal from the lab. These safety items are solely the responsibility of the
student.
11.
Clean up
For clean-up, return the remaining original solution to the instructor. Discard dilutions in the sink and eppendorf tubes in
the regular trash. Do not throw anything in the biohazard waster. Wash 96-well plates and leave them at the sink to dry.
Mark the well that you used with a marker. Return pipettors in the correct boxes, the last person puts the boxes in the
cabinet in the back of the lab. Dry your ice buckets and place them back in the cabinet. Place all other items where you
got them from. Make sure they are clean. Leave your bench ready for the next class to start working.
12.
Homework
See Module 5.
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