Even if you have already used a Spectrawave visible
light spectrophotometer, reviewing the video should
help you with today's work, as it provides valuable information
on the protein assay itseslf.
Research in the life sciences has focused heavily on
molecular biology – the study of DNA and gene regulation – ever
since molecular biologists learned to clone eukaryotic
genes back in the early 1980s. That is because DNA carries
the genetic code for the structure and function of every
living organism on earth. What information do we find
in the genetic code? We find the code for amino acid
sequences of polypeptides, the primary components of
proteins! Proteins are an essential component of every
structural component of the human body, including all
cells and all organelles within our cells. Often they
are the major component. Furthermore, all enzymes are
proteins. Enzymes are responsible for all of the metabolic
and synthetic reactions that keep us alive.
Obviously, to understand the structures and functions
of enzymes, organelles, cells, tissues, organs, and organ
systems we must study the structures and functions of
individual proteins. A starting point to such a study
is to know how much protein we have. The most common
approach to determining how much protein we have in a
sample is to conduct a colorimetric assay to determine
protein concentration. In a colorimetric assay a protein
sample is mixed with a reagent that changes color in
the presence of protein, with color intensity proportional
to the concentration of protein in the sample.
The biuret protein assay was published as a method to
determine protein concentration in the 1940s, although
the reaction itself was studied as long ago as the early
19th century. The biuret assay was commonly employed
well into the 1980s and is still in use because it is
so convenient and inexpensive to prepare and easy to
use. When we assay a protein sample we lose some of it
because the colorimetric reagent destroys the protein
in the process. Because the biuret assay consumes a lot
of protein many laboratories use methods that employ
a much more sensitive color reagent such as the Bradford
assay. We use the biuret assay for this laboratory study
because you can prepare your own reagent from inorganic
reagents (i.e., from "scratch"), giving you
an opportunity to practice your solution-making skills.
In addition to reviewing and printing off this protocol,
please examine the on-line resource on mixtures and solutions
and the video clip on using the WPA S1200 Spectrawave
spectrophotometer. You will also have to finish part
A of the protocol (see below).
Today you will practice your solution making skills
by preparing a complex solution (biuret reagent). We
will also have you learn to calibrate a spectrophotometer
and we'll have you test your solution with calibration
standards containing bovine serum albumin (BSA).
- Prepare biuret color reagent
- Add color reagent to a reference tube and two calibration
standards containing protein
- Calibrate a spectrophotometer
- Read and record absorbance values
***You must wear eye protection
throughout this laboratory session***
A) Prepare biuret color reagent
Parts 1-3 are to be completed before coming to lab.
This information is part of your protocol, to be recorded
in your notebook as you carry out the work.
- Read the formula for biuret reagent carefully and
completely. Biuret reagent consists of 0.9% (W/V) sodium
potassium tartrate, 0.3% copper sulfate x 5 H2O, and
0.5% potassium iodide, all dissolved in order in 400
ml 0.2 M NaOH (f.w. 40.0), then brought to a final
volume of one liter. This is one of several ways in
which one might describe a solution formula.
- Scale the formula down to make a final volume of
just 200 ml of reagent. Scaling down the formula means
to adjust all of the amounts and volumes to make a
smaller volume of solution. For example, to make 200
ml biuret reagent you will need just 1.8 gm of sodium
potassium tartrate (scaled down from 9 gm per liter).
The amount 1.8 gm in 200 ml is in the same proportion
as 0.9 gm in 100 ml, which is a 0.9% solution.
- Write down each step that you will conduct to prepare
your reagent, starting with the beginning volume of
water and the mass of the first reagent that you will
dissolve. Write each step clearly and omit nothing,
including each time you q.s. to volume. NOTE: All chemicals
including the NaOH are available as solid reagents.
You will have to prepare everything fron scratch.
Parts 4-7 of part A is to be completed in lab.
- To use a top loading electronic balance place a piece
of weigh paper on the pan. Tare the balance by [pressing
the lever]. To weigh out dry chemical first make sure
that your spatula is clean. If not, then rinse and
dry it before reaching into the chemical jar. Carefully
place dry chemical onto the paper until you have the
desired mass. If you add too much then don't return
the excess to the jar. Drop it in a waste basket or
flush it down the sink. We only return an excess to
the jar if the stuff is far too expensive to waste.
- To dissolve your chemical stir the water or partially
completed solution using the stir bar with stir plate
and add the chemical directly from the weigh paper.
Scrape it in if necessary. If some material sticks
to the sides of your flask then use a pasteur pipet
to wash it down into the solution.
***To minimize the splash/spill hazard please keep
the stopper in the flask during stirring and storage***
- To q.s. to volume first make sure that
all of the dry reagent is dissolved. Pour the solution
into a cylinder of appropriate volume and then top
off with deionized water. Return the solution to the
flask and stir briefly to mix.
- Your completed reagent should be clear blue. Once
it is mixed then turn off the stir plate.
B) Prepare a reference and two calibration standards
To check the quality of your Biuret reagent we will
have you prepare a reference tube with which to calibrate
your spectrophotometer and two calibration standards
containing different amounts of bovine serum albumin.
- Obtain three 13 x 100 mm culture tubes. Mark the
top of one tube "R" for "reference," a
second tube "2 mg BSA," and a third tube "10
- Pour deionized water from the stock bottle into a
small beaker for use as a working aliquot.
- Pipet 1 ml deionized water into the reference tube.
- Decide what volume of 20 mg/ml BSA standard contains
2 mg of BSA and pipet that volume into the 2 mg tube.
Likewise, prepare your 10 mg tube.
- Add deionized water to each tube containing BSA to
normalize all volumes to 1.0 ml.
- Use your serological pipet with pipet aid to add
5 ml of your biuret reagent to each of the three tubes.
A TA or instructor can show you how to use the pipet
aid to draw up and deliver solution. Let the tubes
stand for 10 min while you prepare your spectrophotometer.
C) Prepare your spectrophotometer and test your reagent
Your spectrophotometers should require very little warm-up
- Press the on/off button once to turn on the instrument.
It will go through a set of self-diagnostic checks.
When the Self Diagnostics screen pops up with three
items checked, press F2 (OK).
- We will use the instrument to take single absorbance
measurements. Selecting that mode requires several
steps. First, press F2 again (Make a measurement).
- Now press F1 (Single/Multi λ/Ratio) and F1
again to select single wavelength (lambda symbol).
- Press F1 a third time to select absorbance. If you
keep pressing F1 it will alternate between % T (transmittance)
and absorbance. We want absorbance values.
- Now press F2 (Set λ) and use the up/down arrowheads
to set the desired wavelength (540 nm). When you have
the desired wavelength press F2 again (accept λ)
- Now carefully insert the reference tube, using both
hands. Press R to calibrate the instrument. After a
short delay the absorbance (Abs) will read 0.000.
- Insert your first (2 mg) sample and press T. After
a short delay the absorbance will be displayed. Record
it and then repeat for the second(10 mg) sample.
- The exact absorbance that you measure for a standard
is influenced by the accuracy with which the standard
was prepared, your pipetting accuracy, and by the quality
of your biuret solution. Our assay gives a typical
absorbance between 0.05 and 0.1 for a 2 mg sample and
between 0.3 and 0.5 for a 10 mg sample.
- Decide whether or not each absorbance is within a
reasonable range and check with an instructor if your
results deviate from the expected. After that, all
that remains is to clean up.
D) Clean up your glassware and work area
- Carefully empty the culture tubes with used biuret
reagent into the nearest sink and dispose of them in
the glass trash receptacle. Rinse any spilled reagent
from your peg rack and return it to the front bench
- Rinse the cylinders and flask with deionized water
from a tap and leave them to air dry. Use soap and
water and if necessary a bottle brush to remove reagent
stuck to the glassware.
- Clean off the surface of your stir plate.
- Clean off your spatula and rinse it with deionized
- Brush off any spilled reagent from the balance and
from the bench surface, disposing of it in the nearest
- Rinse the serological pipet by drawing up and expeling
deionized water a few times. Leave it and the pipet
aid on the bench – they are reusable.
- Straighten up the bench surface, leaving it as you
Planning the Protein Assay
The biuret assay can provide a quantitative estimate of the concentration
of protein so that we might analyze experimental results or optimize
an experiment. Recall that biuret reagent changes color with intensity
proportional to the concentration of protein in a sample (within limits).
To estimate the protein concentration in a sample for which the concentration
is not known we need to use standards for comparison.
samples containing known amounts of protein. When we mix color reagent
with the standards we obtain a range of color intensities to which to
compare the unknowns.
One could estimate protein concentration of unknowns by comparing each
unknown with the set of standards, but that method has obvious drawbacks.
It relies on our judgment and of course there is what to do when
the color change of an unknown falls between the color changes of two
standards. Last time we introduced you to a device called a spectrophotometer,
converts color change to a quantity called an absorbance value.
By measuring absorbance values corresponding to a set of protein standards
we can plot a standard curve of absorbance versus amount of protein.
Absorbance and amount of protein are continuous variables, so we should
add a trend line that relates absorbance to amount over the entire usable
range of the assay.
We can estimate the amount of protein in an unknown from its absorbance
by reading the corresponding amount from the standard curve. Concentration
of the unknown is simply the estimated amount divided by the volume of
sample that was added to the tube.
You will need to plan your standard curve ahead of time.
Today you will start by conducting a protein assay. We will have you
prepare the standard curve in your notebook, in class, and use it to
estimate protein concentrations for two unknowns. We will then have you
use the information to accomplish the kinds of objectives that are a
part of many laboratory protocols.
- Prepare a set of protein standards, add color reagent, determine
- Prepare unknowns for assay, add color reagent, and measure absorbances
- Plot a standard curve in your notebook and add a trend line
- Estimate concentrations for your unknowns
- Calculate how to dilute your unknowns using two different approaches
- Estimate fraction yields for your two unknowns
***You must wear eye protection throughout this laboratory session***
A) Prepare protein standards
- Referring to the table in step 3 below, calculate the volume of 20
mg/ml protein standard to go into each assay tube.
- We want to normalize the volumes to 1 ml per tube before adding color
reagent. Otherwise, variable volumes will affect color intensity and
distort the results. Determine the volume of water to add to each tube
so that the starting volume of every tube is 1 ml. Bring these numbers
to lab with you
- Under the heading "Protein standards," set up a table in
your notebook as in the example below. Fill in the two volume columns
with the values that you calculated prior to lab. Leave the absorbance
column blank for now.
||Amount protein (mg)
||Volume 20 mg/ml BSA (ml)
||Volume water (ml)
||Absorbance @ 540 nm
- Obtain 11 13/100 mm culture tubes and place them in a peg rack, open
- Label one tube "R" using a black Sharpie marker. Label
near the top of the tube, so that the label does not obstruct the light
passing through the color reagent later. Place the tube in a peg rack.
Label the remaining 10 tubes 1 through 10.
- Use a variable volume pipettor to deliver 20 mg/ml BSA and water
into your tubes, according to the information listed in your table.
You will need two different pipettors, one to deliver volumes under
200 µl and one to deliver
volumes from 200 µl to 1 ml (1000 µl). It is very easy
to forget which tube you are on. Experienced researchers check off
each item in the notebook line by line and/or set each finished tube
back one row in the rack to avoid confusion.
B) Add color reagent, incubate, read and record the absorbance values
- When you are ready, use a serological pipet to add 5 ml color reagent
to your reference tube and to each of your standards.
- While your tubes are incubating at the bench (10 minutes),
turn on and calibrate your spectrophotometer as you learned last time.
The reference tube does not require the 10 minute incubation..
the order in which you added color reagent, read and record the absorbance
(not transmittance) corresponding to each of your standards. For best
results the time interval between adding color reagent and reading
absorbance should be close to the same for each tube.
- Keep your reference
tube to re-check your calibration when you read your unknowns.
C) Prepare unknowns and read absorbance values
You will need to prepare unknowns for comparison with the standards. Of course you should record all of the information
in your notebook as you go along.
- Obtain two 15 ml plastic conical centrifuge tubes containing samples of unknown concentration. One should be
labeled A, B, or C and the other should be labeled D, E, or F.
- Label two tubes with the letter code for one of your unknowns, followed by 1 or 2 (e.g., A-1 and A-2). Label
two more tubes for your second unknown.
- In your notebook set up a table for your unknowns as you did for the standards, but with four columns. Label columns
1-4 "tube number," "volume of unknown," "volume of water," and "absorbance @ 540
- As you prepare your unknowns, fill out columns 1-3. For each unknown, tube 1 should contain 0.1 ml (100µl)
of sample. Plan to put 1.0 ml of sample into each tube 2.
We prepare two tubes for each unknown so that if one tube contains too much or too little protein to be measured,
the other tube should give us a usable reading.
- Pipet the tabled volume of unknown into each respective tube, followed by the tabled volume of water.
- Add color reagent, incubate, check your spectrophotometer calibration, and read your absorbances, recording them in column 4 of your unknowns table.
- Check that at least one absorbance reading for each of your unknowns falls within the range of absorbance values
of your standards. If your data are "good," then you have all of the information you need with which
to complete the rest of the work before you leave for the day.
D) Clean up
Please clean all equipment and supplies and straighten
up your bench area.
Prepare your standard curve and estimate protein concentrations
Your protein standard curve will serve as a tool for estimating protein
concentrations in lab. Therefore you want it to be fairly large – it
should take up most of the width of the page and be well-proportioned
as suggested in the graphing tutorial that you completed earlier in the
course. Choose informative labels for the axes as you learned previously.
Plot your standard curve data and then include a best fit trend line.
The relationship may be linear or somewhat curvilinear. Use your best
judgment to fit your trend line, and do remember that it is not good practice to extrapolate, either toward or away from the origin.
From the absorbances for your unknowns estimate each protein concentration.
Remember that concentration of the unknown is the amount of protein divided
by the volume of sample used, not total volume in the assay tube. By
convention we nearly always report protein concentrations as milligrams/milliliter
(mg/ml). For each unknown also remember to use the single absorbance
value that falls within the most linear part of the standard curve.
Plan to make dilutions
Show all work. First, we will have you dilute a specific starting volume to a desired final concentration of protein. This is the
kind of dilution that you would perform in order to make a working solution. Second, you will determine how to prepare
each of your samples to a desired final volume and concentration.
- Your first problem is to determine how to dilute 150 µl of
each of your two unknowns to a final concentration of 1 mg/ml. You
you determined c1 using your standard curve, and your desired
final concentration of 1 mg/ml is c2.
In your notebook record the three known variables for diluting each
of your unknowns. Calculate v2 , showing all calculations in your notebook.
Write down both v2 and the volume to add to v1.
- Your second problem is to determine how to dilute the each unknown
to obtain a final volume of 150 µl at a final
concentration of 1.5 mg/ml. Again record the three known variables
and determine the unknown variable for each unknown. Show all calculations.
Estimate fraction yields
A common approach to learning how something works is to take it apart. We apply that principle to living tissue
when we conduct what we call a tissue fractionation. We usually start by homogenizing the tissue, then we separate the homogenate into components, often employing a method called differential centrifugation. Centrifugation yields a solid component (the pellet) that we resuspend in a volume of liquid. It also yields a liquid component, the supernatant, that we process further. When we conduct a fractionation we want to be able to report
how much of each component we have, usually in terms of the amount of protein recovered.
Hypothetical fraction volumes
The following table lists fractions and volumes obtained from a hypothetical fractionation of whole liver tissue.
For the purpose of this exercise, assume that each of your two unknowns was a sample from the fraction with corresponding label (e.g., unknown A was a sample from fraction A, etc.).
Fraction volume (ml)
whole liver homogenate
nuclear fraction (500 xg pellet)
mitochondria fraction (10,000 xg pellet
microsome fraction (50,000 xg pellet
first ammonium sulfate cut
For each of the two unknowns that you selected, use the volume in the table to determine the total protein in
the fraction. Enter the results in your notebook, showing how you did the calculation.