COMPARING PROTEIN PROFILES FROM DIFFERENT TISSUES
Sample Preparation
1. Label 1.7 mL tubes with 500 l of Protein Extraction Buffer (Camiolo Buffer—0.075M KOAc, 0.3M
NaCl, 0.01M EDTA, 0.1M L-arginine, 25% Triton X-100), so that you know which sample is in
which tube.
2. Place approximate 0.07 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 for 1 minute; agitate by inverting several times for 5 minutes, then
centrifuge on high for one minute.
4. Transfer approximately ~400 l of sample liquid to a newly labeled 1.7 ml tube. 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 and lid identifying YOUR specific protein sample. For the protein standards, label 8
cuvettes and lids 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
Absorption
(mg/ml)
0.0 (blank)
0
0.125
0.25
0.50
0.75
1.00
1.50
2.00
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: Raw data
Sample name
Tissue Samples: Processed data
Sample name
Absorption
Absorption
Protein concentration
(mg/ml)
Protein Standard 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
Volume of
sample needed
Volume of
Laemmli needed
5. Dilute your samples in a clean labeled 0.65 ml tub. Place the 0.65 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 Polyacrylamide Gel Electrophoresis (SDS-PAGE)
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
dam. If you are running one gel in the
cassette leave the dam in place. If you
running two gels in the cassette,
the dam. (DO NOT THROW THE
DAM AWAY!)
buffer
gel
are
remove
4. Put on gloves and get a package that holds your gel.
The gel incorporates a 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 of 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
(10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 glycine/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 200 volts and start the run. If you
protein standards are prestained, then watch the protein standard to know when to stop the gel. The
smallest molecule of protein standard should about 1.5 centimeters from the bottom of the gel (about
20-30 minutes). 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 runs off the gel.
9. To stop the running of the gel, turn off the power supply and remove the gel from the gel cassette.
Open the gel casing and carefully (gel is thin and delicate) separate the two halves of the casing. 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. At this point you can either stain the gel of transfer it to nitrocellulose for Western analysis. Below is
a description of Commassie staining of gel.
11. 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!).
12. Stain the gel, shaking gently for 30 minutes. After 30 minutes, recycle the stain by pouring it back into
the bottle.
13. Replace the stain with destain (which will smell like vinegar) – completely covering the gel. Destain
for about 30 minutes. Pour the blue destain into the destain waste container and add fresh destain into
the tray. Cover the tray and destain overnight.
14. 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? Do similar tissues show more relatedness to each other (based on the # of similar
sized proteins)? Why are the protein profiles different for these tissues if the DNA in the organism is the
same in all of the cells? What kind of information can you get from a stain gel compared to a Western
analysis?