Laboratory 6 Week 1: Cell Fractionation
Introduction:
Eukaryotic cells are complex and contain many kinds of membrane organelles.
For instance, they contain nuclei, mitochondria, vacuoles etc. Two methods exist to
study the organelles in more detail. The first is by using a variety of techniques to
visualize the nuclei while still inside the cells by microscopy (i.e. cell staining and
immunofluorescence). The second method involves suspending the cells in solution, and
breaking them open (lysing the cells). Then the various organelles are then separated
from each other by centrifugation, which then allows them to be used for further study. It
is this second method that we will use in this laboratory.
A procedure called cell fractionation is used to break open the cells and separate
the various organelles. To perform cell fractionation, we first will suspend our cells in
solution, and then we break open the cells, or lyse them. This will release the organelles
inside into solution. Next, we can separate the organelles by centrifuging our solution.
By using centrifugation, we can easily separate the various organelles, since the various
organelles are of different mass, and density (for instance, nuclei are significantly heavier
than mitochondria etc.). During centrifugation, different organelles will pellet at the
bottom at specific speeds based on the mass and densities of the organelles. For instance,
the heavier (larger) the organelles, the less velocity is needed to pellet the organelle. The
lighter (smaller) the organelle, centrifugation must occur at a greater velocity to pellet the
organelle.
Therefore, if we want to separate nuclei from mitochondria, we will centrifuge at
low speed. At low speed, the nuclei will pellet at the bottom, while the mitochondria will
stay suspended in the solution. The left over solution after centrifugation is called the
supernatant, and will contain lighter organelles (i.e. mitochondria and dissolved proteins).
If we then want to separate the mitochondria from the rest of the supernatant, we can
centrifuge at the appropriate speed that would pellet the mitochondria, and then remove
the resulting supernatant.
If we first centrifuged or cell suspension at the speed appropriate to pellet
mitochondria, we would bring the mitochondria to the bottom of the tube. However, we
would pellet everything that is heavier than the mitochondria (i.e. nuclei etc.). Therefore,
in order to get nuclei separated from mitochondria, we must centrifuge at the lower speed
first to obtain the nuclei, and centrifuge the supernatant at the higher speed to collect the
mitochondria. This type of separation protocol is called differential centrifugation. In
this procedure, it is possible to separate the organelles to purification because objects a
similar size to our desired organelles will also pellet at the same speeds. However, these
objects are significantly less concentrated in our suspension. Therefore, our pellets will
are enriched for the organelle we are attempting to isolate.
Each pellet that is isolated can then be resuspended by adding solution, and can
be called a fraction (for instance, the resuspended nuclear pellet is considered the nuclear
fraction). Additionally, a sample of the original suspension is considered a fraction, and
is called the crude fraction, as it contains all the organelles and soluble proteins. Lastly,
a sample of the final supernatant is also considered a fraction and is called the soluble
fraction, and contains everything that was not pelleted by centrifugation.
Using a Centrifuge
There are various types of centrifuges one can use. Various kinds of centrifuges are
available: clinical centrifuges seldom run faster than 5,000 rpm but are able to readily
sediment cells, nuclei or chloroplasts. High-speed centrifuges go as fast as 25,000 rpm
and can sediment smaller organelles such as mitochondria. Ultracentrifuges can produce
speeds of 60,000 - 75,000 rpm and can sediment membranous organelles such as
microsomes and golgi components. The high speed and ultracentrifuges are always
refrigerated and often operated in a vacuum to reduce the heat generated when a rotor is
rapidly spinning in air.
To centrifuge your sample, one must first get the appropriate rotor and place it on the pin
in the centrifuge. The rotor you choose to use must fit the tubes in which you are placing
your cell suspensions. The pin is rotated by the centrifuge motor, thus spinning your
rotor. The relative centrifugal force your cell suspension is subjected to is dependent on
the speed you are spinning your rotor, as well as the radius of the rotor. You can think of
the rotor radius as the distance your centrifuge tube is from the axis of rotation (the pin).
In general, your centrifuge tubes are not placed in the rotor in a straight up and down
fashion, but are placed at an angle, with the bottom of the centrifuge tube being further
from the axis of rotation than the top. The force that that the organelles are being
subjected to in order to pellet them is specified by the formula R.C.F. = 1.119x 10-5
(rpm)2r, where RCF is relative centrifugal force, rpm is the revolutions per minute of the
rotor and r is the distance (in cm) of the particle from the axis of rotation. Given that the
rotor radius is larger at the bottom of the rotor than the radius at the top of the rotor, the
RCF will be different along the length of the tube. Can you determine whether the top or
bottom of your centrifuge tube will be subjected to greater force? The radius used for our
purposes is generally called r(average) and is usually the mean of the maximum and
minimum possible radii. Note, many centrifuge rotors come with conversion charts
between RCF (which is noted by the units in multiples of gravity (x g)) and rpm.
The laboratory
In the cell fractionation laboratory, we will use differential centrifugation to create
fractions enriched for specific organelles. We will first suspend whole plant cells in a
mild salt solution (mannitol grinding medium) and lyse them using a mortar and pestle.
A sample of the resulting solution will be the crude fraction. By doing differential
centrifugation, we will obtain nuclear, mitochondrial and soluble fractions as well. The
soluble fraction will contain objects such as soluble proteins, ribosomes and nucleic
acids.
Materials
1. Cauliflower or Spinach
2. Razor blades (7)
3. Balances
4. Weighing Dishes (located next to balances in both 284 and 286)
5. Mortar and Pestles (In Cold Room – Chilled)
6. Grinding Sand (Next To Balance in 286)
7. 50 mL Graduated Cylinders (24)
8. Paring knives (2)
9. Ice buckets (6)
10. Oak Ridge Tubes (12) – Chilled in Cold Room
11. Cheesecloth
12. Scissors (3 pairs) – near cheesecloth
13. 250 mL beakers (12)
14. Microfuge Tubes
15. Small glass stirring rods
16. Microscope slides
17. Microscopes
18. Coverslips
19. Methyl Green Pyronin
20. Spectrophotometers
21. 5 mL Microcuvettes
22. Ziplock Bags
23. 13 X 100 test tubes (40)
24. Distilled water (4 x 100 mL bottles)
25. Sharpies (8)
26. Bin for dirty dishes
27. Bin for dirty cuvettes
28. Protein solution (0.1 mg/ml)
Make 2 bottles/100 mL
For 200 mls, take a 400 mL beaker
Fill Beaker with 100 mL deionized water
Weigh out 1 g of BSA and mix into the water
Bring volume up to 200 mL and aliquot into 100 mL bottles
29. Mannitol Grinding Medium
Use a 1 L flask and add 850 mL Deionized water
Add 54.66g D-Mannitol
Add 0.82g KH2PO4
Add 2.42g K2HPO4
pH to 7.2 (This step is extremely important-BE ACCURATE!)
Bring volume up to 1L with deionized water and aliquot in 500 mL
bottles (2)
Store Media in the Cold Room
30. Mannitol Assay Medium
Use a 600 ml beaker and add 400 mL Deionized water
Add 27.33 g D-Mannitol
Add 0.41 g KH2PO4
Add 1.21 g K2HPO4
Add 0.38 g KCl
Add 0.51 g MgCl2 X 6 H2O
pH to 7.2 (This step is extremely important – BE ACCURATE!)
Bring volume up to 500 mL with deionized water and aliquot into 2
250 mL Bottles
Store media in the Cold Room
31. 0.4 M Perchloric Acid
Take 34.5 mL concentrated Perchloric Acid
Bring volume up to 1 L with deionized water
32. Protein Dye solution
Obtain a 1 L beaker
Place 50 mL of 95% ethanol in the beaker
Add 600 mg of Brilliant Blue G
Add 750 mL Perchloric Acid and Mix
Bring volume up to 1L with deionized water
Experimental Protocol
Part A: Fractionation of Cauliflower or Spinach Cells (1 head of cauliflower or
bag or spinach per lab section)
1. Using a single-edge razor blade, remove a total of 20 g of the outer 2-3 mm of the
cauliflower surface. [Alternate: Whole class works together to remove a total of
about 220 g of the outer 2-3 mm of the cauliflower surface. See pictures under
the “Results” hyperlink.]
2. Place the cauliflower tissue in a chilled mortar with 40 mL of ice-cold mannitol
grinding medium and 5 g of cold purified sand. Grind the tissue with a chilled
pestle for 4 minutes (on ice!). [Alternate: Whole class puts the entire 220 g of
cauliflower tissue into the kitchen blender. 300 mL of the ice-cold mannitol
grinding medium is poured on top. The blender is “pulsed” several times to break
up the largest chunks of cauliflower tissue. The cauliflower is then homogenized
at “liquefy” speed for 1 minute. See pictures under the “Results” hyperlink.]
3. Filter the suspension through four layers of cheesecloth into a beaker (also wring
out the juice); measure and note the volume, then save a small measured amount
(~ 2-3 mL) in two microfuge tubes for assays and microscopy. Label the “crude”
fraction. Transfer the rest to a chilled 40 mL centrifuge tube. Keep the centrifuge
tubes on ice until all groups are ready to centrifuge.
4. Centrifuge the filtrate at 600 x g (4200 rpm, JA20 rotor; 4000 rpm, SS34 rotor)
for 10 min at 4° C. Make sure that the centrifuge tubes are balanced; the tubes
opposite each other should have the same total volume. Place a 50 mL graduated
cylinder on ice. The pellet from this centrifugation is the nuclear pellet.
5. Decant the supernatant from the centrifugation into the chilled graduated cylinder
(save the nuclear pellet on ice). Measure the supernatant volume and pour the
supernatant into a clean 40 mL chilled centrifuge tube. Spin at 10,000 x g (17,000
rpm, JA20 rotor; 16,000 rpm SS34 rotor) for 20 minutes at 0° - 4° C. Again, be
sure that the centrifuge tubes are balanced. Meanwhile, resuspend the nuclear
pellet in 5 mL mannitol assay medium. Put another 50-mL graduated cylinder on
ice.
6. After the 20 minute centrifugation, transfer the supernatant to the graduated
cylinder; note the volume and transfer to a clean tube (Label “soluble” fraction).
Resuspend the mitochondrial pellet in 8.0 mL of ice- cold Mannitol Assay
Medium. (Use a glass rod or a Pasteur pipette; make sure all the clumps are
completely dispersed). Label “Mitochondria”. Store all tubes on ice until the end
of the period.
7. You now have 4 samples: crude, nuclear, soluble, mitochondrial.
8. Prepare wet mounts of each and examine by microscopy. Capture a
representative image at 400X of each preparation for your notebook. Record
observations of the kinds of particles present and their relative frequencies.
9. Prepare slides of all four preparations with methyl green pyronin:





Place a small drop of each fraction on a clean slide
Add a drop of stain
Cover with a coverslip
Observe at 400X with bright field: nuclei should stain green, cytoplasm red or
pink, and mitochondria can be seen as small dots.
Capture an image of all four preparations for your Results Section
10. Measure the protein concentration in each aliquot as described below and record the
concentrations in your notebook. You will need the protein concentrations later to
report the specific activity of your preparations.
11. After observing each fraction microscopically and measuring the protein
concentration of each fraction, you will need to freeze the fractions for use next week.
Put about 1 mL of the crude, nuclear, soluble, and mitochondrial fractions into
labeled 1.5 mL microfuge tubes. Put ALL of the remaining mitochondrial suspension
(about 7.0 mL) into a labeled sterile 10 mL plastic tube with a screw cap. This larger
amount of the mitochondrial fraction will be needed for activity assays next week.
Write the names of all group members as well as the day and time of your lab on the
outside of a small Ziploc bag. Put all your labeled samples in the small Ziploc bag.
Place all of the small Ziploc bags into a large Ziploc bag labeled with the day and
time of the lab. Store the large Ziploc bag in the cell biology freezer.
Part B: Determination of Protein Concentration of Your Fractions
Standard Curve for Protein Concentration Determination
Proteins are necessary for cells to function properly. Without them, cells would
not be able to have the proper structure, and would not be able to carry out important
processes for life. Additionally, the different organelles have different amounts and
different types of proteins. Therefore, our different fractions will contain different
amounts of protein.
In order to determine how much protein is in each fraction, we must create a
standard curve by using protein solutions of known concentrations and determining their
absorbance using the spectrophotometer. This standard curve will then be used to
determine the protein concentrations in your fractions. A protein that is frequently used
for standard curves is Bovine Serum Albumin (BSA). BSA readily dissolves in water to
form a colorless solution. Dissolved BSA reacts with specific dye known as Coomassie
Blue G-250. Your resulting solutions will turn various shades of blue depending on the
concentration of protein that is in your sample. If your sample has a high protein
concentration, your solution will turn deep blue. If your solution has a low protein
concentration, it will turn a brownish blue. If your solution has no protein in it, then it
will be brown.
In this lab you will: A) mix known concentrations of BSA dissolved in buffer
with Coomassie Blue G-250, B) measure the absorbance of the resulting blue colored
solution and C) plot the measured absorbance of the protein-dye conjugate versus the
known concentration of BSA in that sample. The resulting graph will be the protein
standard curve that you will then use to determine the amount of protein in your
samples of unknown protein concentration.
NOTE: You will be provided with a protein solution containing BSA at a concentration
of 100 µg/mL (0.1 mg/mL). This is known protein concentration. You will do serial
dilutions of your known protein concentration to make other proteins solutions of known
concentration.
1. Make sure that the spectrophotometer has been turned on and is fully warmed up.
2. Label two sets of 13 x 100 glass tubes #1-6a and # 1-6b. (You will do each of the
following steps in duplicate).
3. Your BSA working solution is 100 µg/mL. You must now calculate what volume of
the BSA working solution you must add to the tubes such that the two sets of six
tubes will contain 0, 10, 20, 40, 70, or 100 µg protein respectively. If in doubt, check
with the instructor. Write your volumes in the table below.
4. To each tube add the appropriate amount of distilled water to bring the final volume
up to 1 mL. All tubes must have the same final volume. Write your volumes in the
table below.
5. Add 1 mL of Coomassie Blue G-250 Dye Solution to each tube. Mix by inverting
(use parafilm over the top of the test tube), swirling or gently vortexing with a vortex
mixer. (Remember, frothing denatures proteins; denatured proteins precipitate, and
precipitated proteins are not accurately measured. Only soluble protein is measured).
6. Tube #1 contains no protein so it is considered the "blank”. Pour the contents of tube
#1 into a cuvette . Place this cuvette in the cuvette holder labeled “B”.
7. Allow the color to develop for 10 minutes. While the color is developing, zero the
instrument using the cuvette in the “B” position.
8. Sequentially place the contents of the other tubes into cuvettes and place them
sequentially into the cuvette holders 1, 2, 3, 4 and 5 on the rotating turret. Close the
lid.
9. Ten (10) minutes after adding the protein dye solution, measure and record (see the
Table below) the absorbance of the cuvettes in positions 1, 2, 3, 4 and 5 at 620 nm.
[Note: the color will continue to develop and get darker with time. The standard
curve will only be useful when compared to samples of unknown protein
concentration that have been similarly exposed to the protein dye solution for 10
minutes.]
10. To create a standard curve, plot the average absorbance at 620 nm (A620) of each
pair of samples on the Y-axis versus the µg protein (BSA)/mL on the X-axis. Look at
the graph and find the linear region. You will use the linear range to determine the
protein concentration in your unknown sample.
Tube
Number
g
BSA
to be
added
1a
1b
2a
2b
3a
3b
4a
4b
5a
5b
6a
6b
0
0
10
10
20
20
40
40
70
70
100
100
Volume
BSA
solution
to be
added
Volume
of water
to be
added
Concentration Measured
of BSA in
Absorbance at
ug/mL
620 nm
Average
Absorbance
at 620 nm
Determination of Protein Concentration in your fractions:
Preparation of samples: use one set of each fraction for the protein assay. Dilute your fractions
with water indicated in the table below.
1. For a 1/20 dilution, pipette 50 µL of the protein containing cauliflower suspension into a
clean cuvette; add 950 µL distilled water and invert to mix. For a 1/100 dilution, add 10
µL of the protein containing cauliflower suspension + 990 µL distilled water. Starting
with your lowest dilution for each fraction, then continue with the following instructions:
2. Add 1 mL of Protein Dye Solution to the 1 mL of diluted protein containing cauliflower
suspensions. Add 1 mL of Protein Dye Solution to 1 mL of distilled water to serve as
your blank.
3. Determine the absorbance of the first dilution for each cauliflower fraction. See a photo
of the results of this procedure at:
http://199.17.130.29/berg/307s04/Labs/Images/cellfraction.jpg
4. If the absorbance is out of the linear range of your standard curve, then the absorbance
reading is invalid. You must repeat the measurement using the greater dilution of your
cauliflower fraction.
5. If the absorbance reading is in the linear range of the curve, and above the lowest
detectable amount, record this value and use it to determine the protein concentration of
that fraction.
6. Using the BSA standard curve, determine the protein concentration of each fraction in
µg/mL
7. When finished, rinse all tubes thoroughly in warm running tap water and then
submerse them in pan of water for later washing. Cuvettes should be carefully
rinsed with warm running tap water and the inside surfaces of the cuvettes should
be gently rubbed with wet cotton swabs to remove any adherent Coomassie stain.
The cuvettes should be rinsed one final time in RO water and turned upside down
to dry on a piece of absorbent material or paper towel.
Water (for blank)
Crude homogenate
Crude homogenate
Nuclear fraction
Nuclear fraction
Mitochondrial
fraction
Mitochondrial
fraction
Supernatant
Fraction
Supernatant
fraction
Dilution
Abs 620 nm
NA
1/20
1/100
1/20
1/100
1/20
0.00
1/100
1/20
1/100
[Protein] from
standard curve
0
[Protein] of the
original sample
0
Laboratory #6 Second Week-SDS Polyacrylamide Gel
Electrophoresis
Learning Objectives:
1. Learn to do SDS-Polyacrylamide gel electorphoresis, a common technique in Cell
Biology
2. Learn how to use stoichiometry to determine how much sample to run on your gel
3. Learn how determine how many proteins are in your individual samples, and how
to determine their size in molecular weight
4. Learn how to determine protein content differences between organelles
Experimental Objectives:
1.
2.
3.
4.
to separate the proteins in your cell fractions by electrophoresis
to stain the proteins for easy visualization
to determine the molecular mass of some of the unknown proteins
to determine whether the various fractions have unique proteins
Gel electrophoresis is one of the most commonly used techniques in both Molecular
and Cellular Biology. In fact, many of you have probably run a gel already. The purpose
of doing gel electrophoresis is to separate charged molecule by running them through a
matrix (a gel) when subjected to electrical current. The properties of each molecule will
determine how far it will run through a gel. Molecules are separated on the gel on the
basis of charge, size, and shape, with small molecules migrating further (towards the
bottom of the gel) and larger molecules migrating slower (finishing towards the top of the
gel).
The most common molecules that Biologists run on gels are nucleic acids (DNA and
RNA) and proteins. DNA and RNA are usually run on agarose gels, whereas proteins are
most commonly run on SDS-Polyacrylamide gels. However, sometimes DNA and RNA
are also run on SDS-Polyacrylamide gels.
Nucleic acids have a negative charge, which helps us in getting them to run on a gel.
In order to get the nucleic acids to migrate through the gel, we need to load them orient
the electrical field in the correct manner. Each gel has a set of wells at the top, which is
where we load our samples. In order to then get them to migrate through the gel, we need
to then subject the samples to an electrical field that is properly oriented. To orient the
field correctly, we place the negative electrode (cathode) at the top of the gel, and the
positive electrode (anode) at the bottom of the gel. When the field is active, the
negatively charged nucleic acids will migrate towards the positive electrode, and into the
gel, properly separating by size.
Proteins however do not have a negative charge, so how do we get them to migrate
through the gel? The simple answer is that we use the detergent SDS (Sodium Dodecyl
Sulfate), by placing it both in the gel, the running buffer and the sample buffer. The SDS
is a negatively charged molecule and acts as a denaturant, removing both secondary and
tertiary structure from the proteins. The SDS molecules that bind the protein also confer
a large net negative charge on it, approximately proportional to the molecular weight of
the protein. Therefore, when we subject the proteins to our electrical field, the now
negatively charged molecules will migrate through the gel matrix toward the anode, with
smaller proteins migrating faster, and larger proteins migrating faster. Actually, they will
migrate toward the anode at a velocity that is roughly proportional to the log of the
molecular weight of each polypeptide. . Since the denatured proteins migrate through
the gel on the basis of size, it is possible to estimate molecular weight of individual
polypeptides. Standard proteins of known size are electrophoresed on the same gel along
with sample proteins
In this lab, we will take the fractions we made from our cell fractionation and run
them on an SDS-Polyacrylamide gel. Alongside our samples will run a ladder containing
proteins of known molecular weight, which we use to prepare a standard curve. We will
then use the standard curve to estimate the molecular weight of various proteins in our
fractions.
Materials:
Mini-Protean electrophoresis equipment (Bio-Rad)
Mini-Protean Ready Gels (12%) (Bio-Rad)
Cell fractions from previous experiment
microfuge tubes
4x Sample Buffer
5x Running Buffer (dilute to 1x before use)
mg/mL
SDS-PAGE prestained standards
Destain I (50% methanol, 5% acetic acid, freshly made)
Destain II (7% acetic acid, 5% methanol, freshly made)
Coomasie Blue stain
Power supply
Micropipettors, tips,
Boiling water bath
Unknown protein @ 1
Procedure:
NOTE: The Mini-Protean III is a miniature vertical slab gel unit intended for
rapid electrophoresis of protein samples of small volume. Because its gels are small,
proteins may be separated in less than 45 minutes. The Mini-Protean III can run two
polyacrylamide gel sandwiches simultaneously, if needed. The polyacrylamide gels used
in this lab are precast gels (4% acrylamide in the stacking gel and 12% acrylamide in the
running gel). The upper, or stacking gel (4% acrylamide) slightly restricts protein
migration and serves to concentrate the protein samples. The running gel (12%) serves to
separate individual polypeptides into discrete bands. For a general idea of how this looks
see supplemental Figure 2.
To visualize the proteins after electrophoresis, the gel must be stained with
Coomassie Blue for 30 minutes, and then destained. To handle and to preserve the gel,
the gel can be enclosed in a sealable plastic bag with a small amount of Destain II. The
gel can be photographed with a digital camera to provide all students with a digital image
of their gel.
Each gel will hold up to 10 samples. Each group will use one gel and will arrange
their samples asymmetrically so that they can accurately identify lanes after the gel has
been stained. Place a drawing of a gel in your notebook identifying the order in which
you will be running the samples.
Each Mini-Protein III electrophoresis apparatus will accommodate two gels. Two
groups will use one Mini-Protein III apparatus and one power supply. Two groups will
electrophorese their gels on one apparatus.
1. Find the small fractions that you froze at the end of the cell fractionation
experiment.
2. By now you should have determined the protein concentration of each cell
fraction. Now, determine what volume of each sample (crude, nuclear, soluble,
mitochondrial) will contain 50 µg of protein. For example, if you calculated that
your protein yield was 5 mg protein /mL (5 µg/µL), you would need 10 µL of
that fraction to give you a 50 µg concentration. Place the appropriate volume
(based on protein concentration) of each sample into a labeled microfuge tube.
3. Add 1/4 volume of 4x Sample Buffer to each sample. For example, if the volume
of your protein sample is 10 µL, add 2.5 µL of sample buffer. Also add 2.5 µL
sample buffer per 10 µL of your protein standards.
4. Place the microfuge tubes containing your samples and sample buffer in a boiling
water bath and boil the samples for 2 minutes. Remove the microfuge tubes and
place tubes on ice.
5. Insert the precast gel to the gel apparatus as demonstrated by the instructor.
6. Add 1x Running Buffer to the buffer chambers of the electrophoresis apparatus as
demonstrated by the instructor. [Note: About 500 mL Running Buffer must be
prepared for each group of students.]
7. It is usually best to avoid the outmost lanes. Load the boiled protein samples into
the bottom of a well as demonstrated by your instructor, in the order shown in
Table 1.
Lane 1
Lane 2
Lane 3
Empty
SDS-PAGE molecular weight standards
mitochondrial fraction
Lane 4
Lane 5
Lane 6
Lane 7
Lane 8
Lane 9
Lane 10
soluble fraction
nuclear fraction
crude fraction
SDS-PAGE molecular weight standards
your choice
your choice
Empty
1. Connect the leads to a power supply (red to red and black to black), and
electrophorese the samples until the bromophenol blue dye front has traveled to
the very bottom of the gel (200 V for ~ 45 minutes).
2. After electrophoresis, carefully remove the fragile gel from between the glass
plates, and submerge the gel in Coomassie Blue stain. Shake gently on the shaker
for at least 30 minutes.
3. Remove the gel from the stain solution and place in Destain I for 15 minutes to
1hr. Remove and put in Destain II 1 - 4 hours until the background is clear. (If
you leave it too long in the destain, even the proteins will become destained).
4. Put your destained gel on a piece of saran wrap or in Ziploc bag and photograph it
with a digital camera. Include a centimeter ruler in your photograph so that you
can easily quantify your measurements. You should put a print of this image into
your notebook.
5. Use an image modifying software (eg: Photoshop) to make a grayscale TIF file
out of your image and save it as sdsgel.TIF. Save the file somewhere on your
computer where you can easily find it when needed.
6. Use Scion Image (NIH Image) to measure the migration distances on of the
various proteins on your gel. You will need to analyze the grayscale sdsgel.TIF
file. You will need to set the scale in millimeters using the ruler that you included
in your photograph. Once the scale is set, start your measurements at the
beginning of the resolving gel and end them at the forward edge of the band under
consideration. Remember to measure the migration distance of the bromophenol
blue for each lane so that you can calculate the Rf for each band. Record all of
your data in spreadsheet for easy data manipulation and graphing. You may print
a copy of your spreadsheet and put it into your notebook.