COMPARING PROTEIN PROFILES
SAMPLE PREPARATION
--------------------------------------------------------------------------------------------------------------------1. Label the top of a 1.7 mL tube containing 500 l of Camiolo Protein Extraction Buffer, so that you
know which sample is in which tube.
2. Place approximate 0.1 grams of your tissue sample (about the size of half of an eraser head) into
the Extraction Buffer. Make sure the tissue sample is completely submerged.
3. Homogenize your tissue sample briefly; agitate by inverting several times for 1 minute, then
centrifuge on high for one minute.
4. Transfer approximately 100l of sample liquid to a newly labeled 1.7 ml tube and add another
100l of Camiolo buffer. Make sure to not transfer any solid material.
Your proteins are now ready for concentration determination using the Bradford assay.
DETERMINING PROTEIN CONCENTRATION
Prepare Samples: You will be determining the protein concentration of your samples in groups of 4-8
students using protein standards of known concentrations
1. Label 1 cuvette identifying YOUR specific protein sample. For the protein standards, label 8
cuvettes with the known concentrations of the protein standards (see the photo below).
2. Add 1 ml of Bradford reagent into each appropriately labeled cuvette.
3. Pipette 20 l of the your protein sample, or of the known protein standard, into each appropriately
labeled cuvette, vortex briefly or invert several times.
4. Incubate at room temp at least 5 minutes, not longer that 1-hour, and measure absorbance at
wavelength of 595 nm.
Collecting Data
5. After incubation, clean off the clear non-ribbed side of each cuvette using a Kimwipe.
6. Place the 0.00 mg/ml cuvette in the Spectrophotometer. Line up the clear sides with the arrow.
Push the Zero button and remove cuvette when you see 0.00. You should only have to Zero one
time. Preforming this step “blanks” the instrument, making all the subsequent measurements
relative to this “blank.”
7. With the other cuvettes, wipe off the sides using a Kimwipe and place them in the
Spectrophotometer. Line up the clear sides with the arrow. Push the Test button and record the
absorbance values.
DETERMINING PROTEIN CONCENTRATION OF SAMPLES
Standards:
Protein concentration
(mg/ml)
0.0 (blank)
0.125
0.25
0.50
0.75
1.00
1.50
2.00
Absorption
0
Use Excel to find out the equation for the linear best fit line and write it below (these instructions are
for Excel in Windows 7, yours may differ):
1. Open an Excel spread sheet containing a column for protein standard concentrations, and an
additional column with their corresponding absorbances (as shown above).
2. Highlight the values, click on insert, and choose the scatter plot with no line function.
3. Go to chart layouts, choose trendline. In order to see the equation, go into trendline options and
click “display equation on chart”.
4. Determine protein concentrations of your insect samples using displayed equation.
Tissue Samples:
Sample name
Absorption
Protein concentration
(mg/ml)
PROTEIN STANDARDS AND PROTEIN SAMPLE ANALYSIS
If the absorbance value of your sample falls beyond the maximum absorbance value of the protein
standard of the highest concentration, it will be necessary to add less of your protein sample for the
protein assay.
For protein samples that are greater than >0.5 A from the protein standard of highest concentration,
dilute your protein sample by ¼ for the Bradford protein concentration analysis, by adding 5l of your
protein sample and 15l of DI water into your cuvette with 1ml of Bradford reagent. Then measure
the Absorbance, determine the concentration, and multiple the concentration by 4 in order to calculate
the concentration of protein in your sample.
For protein samples with values between 0.2 and 0.5 A higher than the Absorbance of the highest
concentration protein standard, dilute your protein sample by ½ by adding 10l of your protein sample
and 10l of DI water into your cuvette with 1ml of Bradford reagent. Then measure the Absorbance,
determine the concentration, and multiple the concentration by 2 in order to calculate the concentration
of protein in your sample.
Only the protein samples with absorbance values that fall beyond the absorbance of the protein
standard of highest concentration need to be diluted. In other words, if your protein sample absorbance
falls within the range of the protein standard absorbance values, it will not be necessary to dilute your
sample.
ANALYZING DATA AND CALCULATING DILUTIONS FOR SDS-PAGE
You will want to load an equal amount of protein for your protein gel. This is most easily achieved if
you start with all of your samples at the same concentration. Your concentrations should be > 1
mg/ml, so you will need to dilute your samples to be a standard 1 mg/ml of total protein. Below is an
example of how to calculate your dilution. The dilution will be made with Laemmli Sample Buffer,
which has the detergent SDS. SDS is necessary to denature and charge the proteins so that they can be
separated by SDS-Polyacrylamide gel electrophoresis (SDS-PAGE)
Calculation for sample dilution to 1 mg/ml
For each sample, calculate how much of the sample to add to Laemmli Sample Buffer to make 0.1 ml
(= 100 l) volume. Example: if the measured concentration of your sample was 3 mg/ml, use the
following formula to make a final volume of 0.1 ml of 1 mg/ml protein:
(final concentration) X (final volume) = (sample concentration ) X (sample volume)
(final concentration) X (final volume)
--------------------------------------------- = sample volume
Sample concentration
(1 mg/ml) (0.1 ml) = (3 mg/ml) (x)
(1 mg/ml) (0.1 ml)
----------------------- = (x)
(3 mg/ml)
x = 0.0333 ml = 33.3 l
Volume of Laemmli Sample Buffer needed = (100 l – sample volume)
Volume of Laemmli Sample Buffer needed = (100l – 33.3l) = 66.7l Laemmli Sample Buffer for a
final concentration of 1 mg/ml.
Sample name
Final protein
concentration
1 mg/ml
1 mg/ml
1 mg/ml
1 mg/ml
1 mg/ml
1 mg/ml
1 mg/ml
1 mg/ml
Volume of
sample needed
Volume of
Laemmli needed
Dilute your samples in a clean labeled 1.7 ml tube. Place the 1.7 ml tubes into 95°C heat block
for 5 minutes. This will completely denature the extracted proteins.
Proteins are now extracted and denatured and ready for gel electrophoresis. Proteins can be stored in 20°C until ready to run gel, however, they will need to be denatured again prior to running on gel.
SDS-PAGE ELECTROPHORESIS
Vertical electrophoresis for proteins with acrylamide gel is a bit trickier, more complex and more
hazardous than horizontal electrophoresis done with DNA and agarose gels.
1. If samples are frozen, take them out of the freezer and thaw. Place liquid protein samples into a
95°C heat block for five minutes to denature the proteins so they can easily move through the gel.
2. Inside the gel box is the gel cassette. Take the
gel cassette out of the gel box
3. On one side of the gel cassette is a buffer dam.
If you are running one gel in the gel cassette
leave the dam in place. If you are running two
gels in the cassette, remove the dam. (DO NOT
THROW THE DAM AWAY!)
4. Put on gloves and get a package that holds your gel. The gel incorporates
the stacking gel that has been dyed red. In addition, there is no comb to
remove as the top of the teeth are fixed in place. This not only prevents
the teeth from being accidentally torn off during comb removal, but that
all the wells are uniform, with the teeth remaining vertical. This prevents
sample cross contamination/inability to load desired volume. The teeth
also protrude above the lower front plate of the gel cassette preventing
sample flow over into adjacent wells.
5. Open the sides of the gel cassette. Place the gel in the gel cassette so you see the numbers written
correctly from the outside of the gel cassette (This is critical so ask if you are unsure). Place the
second gel in the same way or make sure the dam is placed correctly in the gel cassette. Close the
sides of the gel cassette
6. Place the gel cassette into the gel box. Carefully remove the comb (The gel is very fragile), and
load the gel with protein samples (15l if your concentration is at 1 mg/ml)). Don’t forget to
record which sample is in which lane. Don’t forget to load a protein standard to monitor
movement of proteins (5l). If your sample protein concentration is < 1 mg/ml, then you will need
to calculate how much sample you need to add to be equivalent to 15l of 1 mg/ml protein. For
example, if you had 0.5 mg/ml concentration you would need to add:
(15l)(1mg/ml) = (x)( 0.5mg/ml)
x=(15l)(1mg/ml)/0.5mg/ml = 30l
However most of you should have >1mg/ml protein concentration and will not need to do this step.
7. Here is another critical step. Fill the inside of the gel cassette to the top with Tris tricine/SDS
buffer. Outside the gel cassette, fill the gel box up to appropriate line (2 gel or 4 gel).
8. Plug the gel box into the power supply. Set the power supply to 100 volts and start the run. If you
protein standards are prestained, then watch the protein standard to know when to stop the gel. Let
the gels run approximately 15-20 min and turn the power supply up to 130 volts. Turn up the
power to 150 volts in another 10 min. Let gel run until smallest molecule of protein standard is
about 1.5 centimeters from the bottom of the gel. If the standards are not prestained, then monitor
the progression of the gel with the dye front and stop the gel when the dye front migrates about 1
cm from the bottom of the gel. Total run time will be 40-50 min.
9. To stop the running of the gel, turn off the power supply and remove the gel from the gel cassette.
Use a flathead tool to crack open the gel casing and carefully (gel is thin and delicate-don’t poke
tool) separate the two halves of the casing. Be careful not to jab the gel, do not put the tool in
between the casing more than The gel will stay on one side or the other of the casing and usually
the bottom will curl up, that is a good place to gently pull gel up off the plate and place into the
staining tray. BE CAREFUL WITH GEL, IT IS VERY EASY TO TEAR!
10. Pour enough Coomassie blue stain to cover the gel. This is a powerful stain, be careful not to
splash it on self or clothes, as it will stain any and all proteins it touches (including yours!).
11. Stain the gel, shaking gently for 30 minutes. After 30 minutes, recycle the stain by pouring it back
into the bottle.
12. Replace the stain with destain (which will smell like vinegar) – completely covering the gel.
Destain for about 60 minutes. Pour the blue destain into the destain waste container and add fresh
destain into the tray. Cover the tray and destain overnight.
13. The next day, place the gel on a light box for examination.
FINAL ANALYSIS
Remember – the distance the protein band has traveled is related to the size of the protein. Also, how
dark the band is tells the concentration of the protein– the darker the band, the more of that protein
there is.
What similarities and differences did you see between the protein profiles of the different samples. Is
this what you expected? How do the results from this technique differ from the results of the Western
Blot? What information can you obtain from each?