Lab 4: protein SDS/PAGE and Western blot, analyze conventional PCR (via agarose gel) & qPCR data

Please print these lab materials before each lab.

Lab Objectives
  • Run extracted total protein from previous labs on SDS-PAGE Gel/Western Blot
  • Run conventional PCR samples on an agarose gel
  • Download qPCR data and discuss analysis

Making an agarose gel (This has been done ahead of time)

Supplies and Equipment:

  • Micropipettes (1-1000 μl)
  • Sterile filter pipette tips (1-1000 μl)
  • Tip waste jar
  • 1L flask
  • agarose
  • 1X TAE
  • Ethidium bromide
  • Microwave
  • Gel rigs
  • Kimwipes
  • Lab coat
  • Safety glasses
  • gloves

  1. Weigh 2g of agarose and mix with 150mL 1x TAE in a 1L flask
  2. Microwave solution for ~ 3 minutes. Keep an eye on the solution so that it does not boil over. You want the solution to be clear - no precipitate and no bubbles.
  3. Cool solution (you should be able to touch the flask for a few seconds), then add 12uL ethidium bromide(EtBr). WARNING: EtBr is a carcinogen be sure to wear gloves and appropriately dispose tip waste.
  4. Mix thoroughly by swirling, then pour into gel tray.
  5. Add gel combs. Using a clean pipet tip, pop any bubbles that could get in the way of your PCR product.
  6. After gel is set, wrap in plastic wrap (label with your initials and date) and place gel in the fridge if not using immediately.

Agarose Gel Electrophoresis
Last week you performed a conventional PCR in class using your reverse transcribed cDNA samples as template and primers your designed. This week we will be checking if amplification was successful using electrophoresis.

Procedure Background
  • Nucleic acid molecules are separated by applying an electric field to move the negatively charged molecules through an agarose matrix. Shorter molecules move faster and migrate farther than longer ones because shorter molecules migrate more easily through the pores of the gel. This phenomenon is called sieving.
  • The most common dye used to make DNA or RNA bands visible for agarose gel electrophoresis is ethidium bromide usually abbreviated as EtBr. It fluoresces under UV light when intercalated into DNA (or RNA). By running DNA through an EtBr-treated gel and visualizing it with UV light, any band containing more than ~20 ng DNA becomes distinctly visible. EtBr is a known mutagen, however, so safer alternatives are available.
  • A DNA ladder is a solution of DNA molecules of different lengths used in agarose gel electrophoresis. It is applied to an agarose gel as a reference to estimate the size of unknown DNA molecules
  • If amplification was successful you should see one clear band between 150-400bp (in length depending on your gene) and the negative controls will have no band. If there is a band in the negative control than there might be contamination in your reagents and you can not be sure that the intended gene was actually amplified.
  • Often contamination requires carful rePCR in case the contamination occurred during reaction setup. However, if any reagents are contaminated troubleshooting may be required to obtain a clean PCR product.
  • PCR bands outside of the intended size range could indicate unspecific amplification and will require either optimization of the reaction cocktail and thermal cycling parameters or redesigning the primers.

  1. Place gel in gel box and fill with 1x TAE buffer (to fully cover wells)
  2. Remove combs from wells
  3. Load 7uL 100bp ladder in far left lane
  4. Load 20uL of your PCR sample into the gel (retain the remaining vol at -20ºC)
  5. Run gel at ~ 100V for ~ 1hr
  6. Visualize the gel on the UV transilluminator

Protein Extraction and Analysis Part 2

SDS - Polyacrylamide Gel Electorophoresis (SDS-PAGE)

Supplies and Reagents
  • micropipettes (1-1000 μL)
  • sterile filter pipette tips (1-1000 μL)
  • sterile gel loading tips
  • 1.5 mL screw cap tubes
  • microcentrifuge tube rack
  • lab coats
  • safety glasses
  • gloves
  • lab pen
  • timers
  • heating block with water bath
  • tube "floatie" (8 tube capacity)
  • glass container for boiling water that can accommodate "floatie"
  • protein gel box (SR provided)
  • SDS/PAGE gels
  • gel loading tips
  • trays for staining gels
  • power supply
  • platform rocker/shaker
  • plastic wrap
  • 2X SDS reducing sample buffer
  • protein ladder marker
  • gel running buffer
  • light box
  • digital camera

Procedure Background
  • SDS-PAGE is the process of separating proteins from one another on the basis of molecular weight. A mixture of proteins is subjected to an electric field and pulled through a polyacrylamide matrix towards the cathode. However, proteins must be treated prior to separating them in this manner. This is because the charge on any given protein is dependent upon the amino acid sequence as well as the pH of the solution. Thus, a crude protein extract from cells or tissue contains a hetergenous mixture of proteins with varying charges. Without treating them in some manner, the proteins will migrate independent of their molecular weight. Additionally, proteins have various tertiary or quarternary structure that can influence the rate at which they migrate in an electric field. In order to address these issues, protein samples are prepared in a specific fashion to linearize the proteins and impart the same charge to all proteins in the sample to ensure that they become homogenous and their migration rate during SDS-PAGE is solely due to molecular weight.
  • Protein samples are combined with a reducing sample buffer. This reducing sample buffer contains sodium dodecyl sulfate (SDS), B-mercaptoethanol, glycerol, Bromophenol blue and a buffer. SDS imparts an overall negative charge to all the proteins in a sample. This ensures that all of the proteins will migrate in the same direction (towards the cathode) when placed in an electric field. B-mercaptoethanol is a reducing agent. It accepts electrons from disulfide bonds formed between two cysteine residues. It serves as one step to help break any tertiary or quartenary structure of proteins in the sample. Glycerol simply serves as a sinking agent for your sample. It has a greater density and will allow your sample to sink into the buffer contained in the wells of the gel. Bromophenol blue is a negatively charged dye that allows one to visually track the migration of your samples through the polyacrylamide gel. Bromophenol blue migrates at the same rate of proteins ~5-7kDa. Finally, the buffer is present to maintain the appropriate pH for your sample. Once samples have the appropriate amount of reducing sample buffer, they are boiled. Boiling causes the proteins to fully denature, eliminating any tertiary or quartenary structure and leads to linear chains of amino acids.
  • Samples are then run through polyacrylamide gels. Traditionally, a single polyacrylamide gel is actually comprised of two gels with different percentages of polyacrylamide, pH and buffer. The top portion of the gel, relative to the bottom portion of the gel, has a lower percentage of acrylamide, a lower pH and a lower concentration of buffer. This gel is referred to as the stacking gel. The bottom gel has higher mounts of all three components listed above and is called the running gel.
  • The low percentage of polyacrylamide in stacking gels allows all the proteins in the sample, regardless of molecular weight, to quickly and easiy migrate through the gel. When the samples begin to enter the lower gel containing a higher percentage of polyacrylamide (running gel), the proteins are now all "stacked" upon one another. This allows all of the proteins in a sample to enter the running gel at essentially the same time. Additionally, the differences in pH and buffer content between the stacking and running gels leads to a local increase in voltage around the sample, which helps drive the sample from solution in the well into the polyacrylamide matrix of the stacking gel.
  • After SDS-PAGE is complete (when the dye front has reached the bottom of the gel), the gel is either set up for Western blotting or is stained to reveal the proteins. There are a number of stains that can be used, depending on the sensitivity needed to visualize proteins of interest, but we will use Coomassie Brilliant Blue. This is a non-selective stain, meaning it binds all proteins regardless of their amino acid makeup. Additionally, it is cheap and rather sensitive. The dye will initially turn the entire gel blue and even a short exposure (5 minutes) often results in over staining. Thus, it is necessary to destain the gel to wash the dye out of the areas of the gel where no protein is present. After destaining, the proteins should appear as blue/purple bands and the rest of the gel should remain relatively clear.

Also see Manufacturers Protocol / Manual: Precise™ Protein Gels

  1. Begin boiling water on hot plate.
  2. In a fresh, 1.5mL SCREW CAP tube add 15uL of your protein stock and 15uL of 2X Reducing Sample Buffer. Return your protein stock to the box in the -20C freezer labeled protein samples.
  3. Mix sample by flicking. Briefly centrifuge (10s) to pool liquid in bottom of tube.
  4. Boil sample for 5 mins.
  5. While sample is boiling, observe assembly of gel box and gels. Rinse gel wells thoroughly as demonstrated.
  6. When sample is finished boiling, immediately centrifuge for 1min. to pool liquid.
  7. Slowly load your entire sample into the appropriate well using a gel loading tip.
  8. Put lid on gel box and plug electrodes into appropriate receptacles on the power supply.
  9. Turn power supply on and set voltage to 150V. Run for 45mins. CHECK YOUR AGAROSE GEL RESULTS. MAKE SURE EVERYTHING IS SET UP FOR WESTERN BLOT.
  10. Turn off power supply and disconnect gel box from power supply.
  11. Remove lid from gel box.
  12. Disengage the tension wedge.
  13. Remove gel from gel box.
  14. Carefully crack open cassette to expose gel.
  15. Trim wells at top of gel.
  16. Notch a designated corner of the gel to help you remember the correct orientation of the gel (i.e. which is the top/bottom of the gel, which is the right/left side(s) of the gel)
  17. Proceed to Western Blotting protocol.

WesternBreeze Chromogenic Western Blot Immunodetection

Supplies and Reagents
  • Nanopure water
  • gel staining tray
  • Blocking Solution
  • rotary shaker
  • Primary Antibody Solution
  • Antibody Wash
  • Secondary Antibody Solution
  • Chromogenic Substrate
  • timers
  • lab coats
  • safety goggles
  • gloves
  • SDS-PAGE gel
  • Tris-Glycine transfer buffer
  • filter paper
  • nitrocellulose membrane
  • semi-dry transfer station

After the SDS-PAGE has separated the proteins on your gel according to size and conformation, the Western Blot is used to probe the separated proteins with protein-specific antibodies. We will be using Hsp70 in this lab. The proteins are inaccessible to the antibody when they are in the gel, so we use an electric charge to transfer them to the nitrocellulose membrane. Once on the membrane, the proteins are accessible to binding by the antibody.
The blocking solution prevents non-specific binding of the antibody. Blocking solutions are made of dilute protein and will actually bind to places on the membrane where there is not already protein. This step helps to eliminate false positive results.
Primary antibodies are generated when a host (in this case a mouse) is exposed to the protein of interest. The secondary antibody (anti-mouse) will bind directly to the species-specific portion of the primary antibody. The secondary antibody is linked to a reporter molecule.
This kit contains some chemicals that are irritants and one (nitro blue tetrozolium) that is a possible carcinogen. Make sure you take the appropriate safety precautions when following the protocol.
Do not touch the surface of the membrane, even with gloves on. This could adversely impact your results.
All washing, blocking and incubating steps are performed on a rotary shaker at 1 revolution per second.
Add solution to the trays slowly to avoid formation of bubbles under the membrane. Decant from the same corner of the dish to ensure removal of all previous liquids.
(Adapted from the WesternBreeze protocol and Wikipedia)

Western Blot Protocol
This is a lengthy protocol with many incubation steps. There will be one gel for the entire class. As a class, you should assign yourselves steps so that everyone can participate and so that we don't waste time. When it is not your turn to attend to the gel, you can do protein and RNA extractions for your project.
  1. Soak the filter paper, membrane and gel in Tris-Glycine Transfer Buffer for 15 minutes.
  2. Assemble the blotting sandwich in the semi-dry blotting appartus:
    1. Anode (+++)
    2. filter paper
    3. membrane
    4. gel
    5. filter paper
    6. cathode (---)

  1. Transfer the blot for 30 minutes at 20V
  2. Remove the gel from the sandwich and rinse off adhering pieces of gel with transfer buffer.
  3. Wash membrane 2 times, for 5 minutes each, with 20 mL of pure water.
  4. Put the membrane in the plastic box and add 10 mL of Blocking Solution. Cover and incubate overnight on a rotary shaker set at 1 revolution/second.
  5. Your TA will do the rest of the steps. After class tomorrow you can come and see your results.
  6. Decant liquid.
  7. Rinse the membrane with 20 mL of water for 5 minutes, then decant. Repeat.
  8. Incubate the membrane in 10 mL of Primary Antibody Solution. Decant the solution.
  9. Rinse the membrane with 20 mL of Antibody Wash for 5 minutes, then decant. Repeat 3 times.
  10. Incubate the membrane in 10 mL of Secondary Antibody Solution for 30 minutes. Decant.
  11. Wash the membrane for 5 minutes with 20 mL of Antibody wash, then decant. Repeat 3 times.
  12. Rinse the membrane with 20 mL of pure water for 2 minutes, then decant. Repeat twice.
  13. Incubate the membrane in 5 mL of Chromogenic Substrate until a purple band appears. This will occur between 1-60 minutes after adding the Chromogenic Substrate.
  14. Dry the membrane on a clean piece of filter paper to the open air.