Module 5. Determination of Protein Purity by SDS-PAGE
1.
Introduction
You are given a tube of clear solution that you know contains protein molecules. Your
supervisor asks you to find out how many different proteins are in the tube and what their sizes
are. For example, the answer might be that there are 2 different proteins, one is 20 kDa and the
other one is 57 kDa. What experiment can you do to answer this question?
The method you might use is SDS-PAGE. SDS-PAGE is a method that separates protein by
size in a polymer matrix. The proteins travel from top to bottom; the smaller proteins migrate
faster, landing further down the gel than the larger proteins. The reason the larger molecules do
not migrate as far down the gel is that they are slowed down more by the
thick polymer matrix (see figure on right).
In the gel you will use, there are 10 wells, so you can run 10 samples at
the same time through the matrix. You always use a molecular weight
marker (ladder) that shows where different sizes of proteins land on a
gel (See figure on left for the ladder we will use). You can then
compare your samples to the ladder to find out what size proteins are
present in your samples (See figure on bottom right).
We will dilute all protein samples to a predetermined concentration and
volume before mixing with the denaturing sample buffer. To
completely denature the samples we heat them in a steaming water bath
for 10 minutes. It is important to load the right amount of protein into a well to obtain results where
the band on your gel is not smeared but can be clearly seen.
Purpose of the lab
In Module 4, you purified the protein Dihydrofolate reductase from B. stearothermophilus (BsDHFR) from a mixture of
proteins. In this module, you will assess how well the column chromatography step you performed worked and how pure
the resulting purified protein is by doing SDS-PAGE. The protocol for this part is described in the module. You will also
explore the relationship between the amount of protein loaded on the gel and the appearance of the resulting band in an
SDS-PAGE gel and add this part to the protocol.
In this module your assignment is to design and conduct an experiment to look at the relationship between amount of
protein loaded in a well and the appearance of the resulting band in an SDS-PAGE gel.
Note: A general protocol for SDS-PAGE is provided but you must modify it to accomplish your goal.
2.
Agenda for the Day
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3.
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 SDS-PAGE.
In class, groups review Math Moment Problems.
In class, groups review Experimental Protocols and come up with a consistent protocol.
Conduct the experiment in groups. Follow the provided protocol for 5 lanes and your own protocol for the other 5
lanes. You will need to run all wells at the same time so prepare all samples and load all samples before running
the gel. Each student individually records data in their personal laboratory notebook.
Clean up
Background
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Read about SDS-PAGE in Ch 6 in your book (pages 174-176 and 188-190).
Watch the following video:
http://www.piercenet.com/browse.cfm?fldID=21518847-2D72-475F-A5B9-B236EC5B641E#SDS-PAGE-video
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See manufacturer’s manual instructions with useful information at
http://www.piercenet.com/instructions/2161472.pdf
The catalog number for the molecular weight marker we will use is Fisher
BP3601-1.
Please,
read
about
it
on
line
(http://www.fishersci.com/ecomm/servlet/fsproductdetail_10652_1448604
__-1_0). There is a picture of these markers at the front of this module.
It is important not to load too much or too little protein onto the SDSPAGE gel (made of polyacrylamide). Overloading (too much protein)
results in precipitation and aggregation of proteins, producing streaks and
smears. Underloading (too little protein) is disappointing because only the
most abundant proteins (or nothing) will show up on the gel. It is
important to adjust the concentrations of sample so that an appropriate
amount of protein can be loaded onto a gel. Usually 10 µg per lane of
purified protein gives a reasonable result but you may be able to see as little as 50 ng. A typical gel well holds 30
µl easily, and perhaps 40 µl if the well dividers are in good shape. The concentrations of your MIX, F, W and
DHFR samples are unknown at this time but are likely between 0.05 mg/mL and 2 mg/mL.
Math Moment
1. You have a protein stock that has the concentration 46 µM. The molecular weight of the protein is 18 kDa. What is the
concentration of the protein in mg/mL?
2. Let’s say you want to load 10 µg of protein into a well. For the protein in the question above, how many µL of protein
sample should be loaded into one well?
3. We must not put the protein in the well by itself but rather first mix it with denaturing buffer. It is important to denature
the protein with this buffer and by heating before running the sample on an SDS-PAGE because this way the 3D shape does
not influence the rate at which the protein migrates and the gel results are due to size only. You are using a 4X sample
buffer. This means that you must dilute it 4-fold to reach your final sample concentration (V2/V1 = 4). You plan to add 10
µL of sample buffer to your sample. What volume of protein solution should you add the sample buffer to in order to have
the final concentration of the sample buffer be 1X?
4. You are using a 4X sample buffer. You will mix 10 µL of sample buffer with 30 µL of protein sample. Your protein
sample has the concentration 0.828 mg/mL. You want there to be 10 µg of protein in the sample after you mix protein with
sample buffer. What dilution of your protein do you need to make in order to be able to use 30 µL of protein and achieve
10 µg of protein in your sample?
5. You do not know the concentration of your protein but you suspect it to be in the range 0.2 mg/mL to 2 mg/mL. You
know you ideally want to have around 10 µg of protein in a well but this is an approximation. If you assume your protein
concentration is 0.5 mg/mL, and you will use 30 µL of protein sample with 10 µL of loading dye, what dilution of your
protein solution do you need to make first? Describe how you will make the dilution.
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6. Now assume the concentration is 0.2 mg/mL. What is the highest amount of protein that you can have in a volume of
30 µL (provide your answer in the units of µg)? Do you expect this to show up on a gel? (Hint: look again at the point in
the Background section of this module)
7. You have a protein of an unknown concentration but you know that the range is 0.2 mg/mL to 2 mg/mL. You know that
typically amounts around 10 µg give a good result on an SDS-PAGE gel. You have 9 wells (1 ladder and 9 others). Using
5 wells, how would you design an experiment to look at the effect of amount of protein loaded on the band appearance on
the gel? Remember that you are mixing 10 µL of sample buffer with 30 µL of protein sample. You will need to make 5
different dilutions of the original protein solution. How will you do that?
8. Find a picture of the ladder/marker we are using for the experiment. Draw a figure of a gel with 10 lanes, with the
ladder/marker (the one we are using) in the first lane. Label the lanes on the marker lane. In lane 2, draw what you expect
a mixture of proteins to look like (let’s say there are 1000 different proteins with molecular weights between 2 kD and 130
kD). Think about a smear rather than individual bands. In lane 3 draw what a monomeric 18 kD protein would look like.
In lane 4, draw what it would look like if the protein in well 3 dimerized. In lane 5, draw what it would look like if there
were both monomers and dimers of the 18 kD protein present. Remember DTT and 2-mercaptoethanol reduce disulfide
bonds and make monomers out of dimers etc.
5.
Supplies Provided
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Your purified protein and other fractions from last week: MIX (Original mixture), F (flow-through), W (wash) and
DHFR (purified protein).
Eppendorf tubes
40 mM Hepes buffer, pH 6.8
Ready made gel
Thermo Scientific Tris-HEPES-SDS Running Buffer (Product No. 28398)
Transfer pipettes
Lane Marker (Fisher EZ-RUN marker, catalog number BP3601-1)
4X sample buffer
Staining solution (InstantBlue from Expedeon)
SDS Page equipment (Biorad models)
Micropipettors and tips
Waterbath at 70 °C
Staining trays
Shaker
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6.
Protocol
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Label four 1.5 mL eppendorf tubes O (Original mixture), F (flow-through), W (wash) and P (purified protein).
For each of your protein samples (MIX, F, W, and DHFR), add 10 µL 4X Sample Buffer to 30 µL of protein
sample in the appropriately labeled eppendorf tube.
Heat Eppendorf tubes for 3-5 minutes in a water bath at ~70°C.
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7.
Common Mistakes and Some Advice
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Caution: Inserting pipette tip too far into the cassette when loading may cause the cassette to separate! Hold the
tip gently above the well and let the solution enter into the well drop wise and slowly.
It is helpful to have the running buffer be cold.
When loading the samples into wells, take your time. If you rush,
you may introduce bubbles or the sample may float to an adjacent
well.
When heating samples, close the caps tightly to prevent “popping”.
We are using a Bio-Rad Mini-PROTEAN Cell apparatus (see
picture on right). Make sure that the rubber seal is in the correct
orientation. If not, remove the gasket from the inner frame (Figure
on the right), turn it around so the flat side is facing outwards and
re-insert into the inner frame.
Advice for experimental design
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9.
Set up your gel running apparatus (instructor will help you) and place your gel into the gel holder with the short
plate pointing toward the inner tank.
Add sufficient volume of Tris-HEPES-SDS running buffer into the inner tank of the gel running apparatus to cover
the sample wells by 5-7 mm.
Add Tris-HEPES-SDS running buffer to the outer tank to ensure proper cooling. The buffer in the outer tank
should be at least 2/3 up the side of the container.
Using a transfer pipette, rinse the sample wells thoroughly with Tris-HEPES-SDS running buffer to remove air
bubbles and to displace any storage buffer.
Add 10 µL of Fisher EZ-RUN marker into lane #1 on the gel. Caution: Inserting the pipette tip too far into the
cassette may cause the cassette to separate.
Carefully apply your protein samples (~30 - 40 μL) to your wells #2(MIX), #3(F), #4(W), and #5 (DHFR).
Connect the gel rig leads to the power supply and turn the power supply on. Electrophorese at 100 V for about 45
minutes. Do not touch the gel box after you connect it to the power supply. Make sure bubbles are going up
from the electrode when you start the run.
Turn the power supply OFF. Disconnect the gel rig leads from the gel box. Remove gel from the gel tank.
Insert a coin in one of the slots on the side and twist to open the cassette. Pull the top plate away from the bottom
plate, exposing the gel. Loosen the gel on the bottom with water and place in a staining tray.
Carefully cover the gel with InstantBlue gel staining solution (instructor will do this). You should see bands
appearing on the gel in about 15 – 30 minutes. Pour off the gel stain (save it – instructor will help) and store the
gel in distilled water. Take a picture of the final stained gel with your cell phone.
The protocol above describes how to run the ladder, MIX, F, W, and DHFR samples on the gel. This part will use
5 of the 10 lanes in the gel.
You must, in addition, design a protocol to examine the effect of amount of protein loaded and appearance of band
using the remaining 5 wells.
You may choose to focus on one or two of the fractions, for example MIX or DHFR, or try two different amounts
of each sample. This is your choice.
You must have designed the protocol in detail including how much of each solution (protein sample, buffer to
dilute sample, sample buffer) to add when preparing the sample that will be loaded into the wells. Remember, you
can load at most 40 μL into each well. You must mix 1 part 4x sample buffer with 3 parts protein solution.
Vocabulary
SDS-PAGE, DTT, 2-mercaptoethanol, ladder/marker, staining, lane
10.
Safety
You must wear safety glasses when conducting the experiment. You must never eat or drink in the laboratory. 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. Caution: Do not touch the power source/gel running apparatus while the power is on.
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11.
Clean up
Save the running buffer in the original bottle. Do not throw anything in the biohazard waste. Return pipettors in the correct
boxes; the last person puts the boxes in the cabinet in the back of the lab. Clear and dry the gel boxes and return them to the
cabinet. Dry your ice buckets and place them back in the cabinet. Place all other items where you got them. Make sure
they are clean. Leave your bench ready for the next class to start working. Remember to turn the waterbath off if you are
the last user.
12.
Homework
The data from this experiment will become part of a full laboratory report that covers modules 4-6 (one per group). This
report will consist of an abstract, an introduction, a methods section, and a result and discussion section (results and
discussion are combined into one section). Please, see the “General Advice on Report” document and follow the
requirements (use it as a check list) that are listed there when preparing your report. Prepare a professionally formatted,
concise report your group feels proud of, each group member must contribute sections and each group member must edit
the entire document. This report is due one week after completion of Module 6 at the beginning of the lab session.
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