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Day 5: SDS-PAGE and Electrophoretic Transfer

Assignments Due

Preparation

Overview of Experiment

In this session a denaturing 10% polyacrylamide gel (SDS-PAGE) is run to separate the proteins based on size. We use the discontinuous SDS-PAGE system developed by Laemmli, UK (Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227 (5259): 680-685, 1970); a resolving or separating (LOWER) gel and a stacking (UPPER) gel are cast with different pore sizes, pH, and ionic strength. Additionally, different mobile ions are used in the gel and running buffers. This buffer discontinuity concentrates large sample volumes in the stacking gel and allows high resolution in the separating gel.

One half of the gel is silver stained to visualize the protein bands in order to evaluate the progress of the purification procedures -- Silver stain is NONSPECIFIC and gives a qualitative picture of the efficacy of the purification scheme. The other half of the gel is electrophoretically transferred onto a PVDF membrane that becomes our western blot -- in a later class we will develop the membrane with an antibody generated against murine adenosine deaminase to identify SPECIFICALLY which band corresponds to adenosine deaminase and to estimate molecular mass (m).

Procedures

Gel Contents

SDS-PAGE Sample Preparation

  1. Using the concentration of protein in each sample (in mg/ml or µg/µl), determine the volume to load into each well of the gel:

    For the crude sample you need to load 7.5-10 µg of protein per well. Prepare the sample in 12 µl; dilute if necessary with the Tris/glycine/SDS running buffer. The crude samples with multiple proteins must have more protein loaded since the total protein will be distributed in many bands.

    For the size exclusion and the Q fractions, you need to load 0.5 to 2 µg per well. Prepare the sample in 12 µl; dilute if necessary with the Tris/ glycine/SDS running buffer.
    *if your fractions are very dilute, use the maximum sample volume (12 µl).

  2. Add 5X-SDS gel-loading buffer to each sample to give 1X final conc. and MIX WELL. This solution contains a dye (bromophenol blue) to monitor the electrophoretic front and glycerol to encourage the sample to settle to the bottom of the well during loading. β-mercaptoethanol is also included in the tracking dye to enhance denaturation by disrupting disulfide bonds between cysteines.

    1X SDS gel-loading buffer:
    50 mM Tris-Cl (pH 6.8)
    2% SDS
    2% β-mercaptoethanol
    0.1% bromophenol blue
    10% glycerol

  3. Heat the samples in a boiling water bath for 4 minutes.
  4. Pulse spin the samples to collect the liquid into the bottom of the vials.
  5. Put samples on ice until ready to load the gel.
  6. Mix sample by pipetting before loading the gel.

Plate Assembly

Precast polyacrylamide mini gels from Bio-Rad [record the specific details about the type of gel you use in your lab notebook] are assembled as described below. These instructions are adapted from the BioRad Ready Gel Cell Instruction Manual.

  1. Get ONE gel per team.
  2. Take pre-cast gel out of its pouch and remove the comb from the top of the gel.
  3. Carefully pull the tape off the bottom of the gel (the tape must be removed so the bottom of the gel is exposed to running buffer).
  4. Rotate the cams of the clamping frame outward; remove the electrode assembly.
  5. Place the pre-cast gel into the slot at the bottom of the electrode assembly.
    NOTE: make sure that the SHORT glass plate faces INWARD toward the notches on the green U-shaped gaskets.
    **since you will only be using ONE gel, you will need to use the mini cell buffer dam**
  6. Press gel and buffer dam up against the gaskets.
  7. Transfer the electrode assembly and gels into the clamping frame.
  8. Gently press down on the electrode assembly while closing the two cam levers of the clamping frame.
    NOTE: Gently pressing the top of the electrode assembly forces the top of the short plate on each gel against the rubber gasket; if you do not push down, the upper buffer chamber may leak.
  9. Carefully lower the assembled unit into the tank.
  10. Pour approximately 125 ml of Tris/glycine/SDS running buffer (25 mM Tris, pH 8.3, 192 mM glycine, 0.1% (w/v) SDS) into the center of the inner core until the liquid level is slightly above the top of the inner glass plate. Observe the bottom of the inner core for leaks. If no leaks are found, pour approximately 200 ml of buffer into the tank. Check for bubbles at the point of contact between the gel and the buffer. Remove bubbles if necessary using a disposable transfer pipet.
  11. The wells are outlined in "black": use regular small tips for the pipettors and slowly load your samples into the wells of the gel.

    Half of the gel will be stained for protein and the other half will be transferred for western analysis. Each half of the gel should contain standards: the prestained standards (Precision Plus Protein Kaleidoscope Standards, BioRad) are ready to load (DO NOT BOIL); load 5 µl.

    Load 15 µl of each of your samples (remember to load the SAME volume for ALL).
    Be sure to keep a record of the well position of each sample.
  12. In the EMPTY wells, load 15 µl of 1X-SDS gel-loading buffer in Tris/glycine/SDS running buffer; this step ensures even migration of the dye front as the gel runs.
  13. Place the lid on the unit and connect the electrodes so that the anode is at the bottom of the gel. (The color scheme can be followed, red to red, etc.)
    PLEASE NOTE: Banana plug fittings are not to be turned or twisted. Only push on and pull off by grasping the plug (or the entire lid for the boxes) without turning.
  14. Running conditions: Tris-HCl SDS gels are run at 200 V constant voltage. A typical run will take approximately 30 minutes.


    NOTE: While the gel is running, pay attention to the current display; if the current drops substantially or if the gels are running very slow and the dye front looks distorted, stop the run and add buffer to the inner chamber. The buffer level must be ABOVE the inner plate.

    Warning: Be certain the power supply reads "OFF" before handling the electrode connections. Electrical shock hazards are always present when using laboratory power supplies. Do not contact buffer reservoirs or wiring while the power supply is activated.

  15. When the run is complete, disassemble the gel. Using a razor blade (Bio-Rad key), carefully cut off the stacking gel from the gel; carefully cut the gel in half.



BIO-RAD Precision Plus Protein Kaleidoscope Standards

Color

Molecular Mass (m), kD

Blue

250

Purple

150

Blue

100

Pink

75

Blue

50

Green

37

Pink

25

Blue

20

Blue

15

Yellow

10

NOTE: These standards must be recorded in your lab notebook.


Silver Stain of SDS-PAGE Gels (can detect as little as 0.5 ng per band)

Adapted from:
Morrissey, J. H. (1981). Silver stain for proteins in polyacrylamide gels: a modified procedure with enhanced uniform sensitivity. Anal. Biochem.117.2: 307-310.

Silver stain is up to 100X more sensitive for detection of protein than the Coomassie stain and will be used to stain the SDS-PAGE gels. Take care not to touch the gel with your fingers as the stain will develop the deposits left there.

Solutions

40% methanol, 10% acetic acid
*5µg/ml dithiothreitol (DTT)
*0.1% AgNO3
*3% Na2CO3 w/ formaldehyde
Milli-Q H2O (RO H2O is NOT clean enough for the silver stain procedure)
[*these solutions were prepared in Milli-Q H2O]

Procedure

  1. Place the SDS-PAGE gel in a container (plastic storage boxes work well; these are in the lab closet) and cover with 40% methanol, 10% acetic acid solution. Shake on rocker for 10 minutes.
  2. Wash the SDS-PAGE gel with excess Milli-Q H2O three times for 5-10 minutes each time, on the rocker.
  3. Pour off the water after last wash and add 5µg/ml DTT solution to cover gel. Shake for 10 minutes and pour off DTT solution.
  4. Add 0.1% AgNO3 solution and shake for 10 minutes.

    CAUTION: WEAR GLOVES! The AgNO3 solution is toxic and should not touch your skin.
  5. Rinse with Milli-Q H2O.
  6. Add 3% NaCO3 w/ formaldehyde and watch gel for desired intensity. If the developer soultion in the tray turns yellow or brown within one minute, pour off and apply fresh developer. When desired intensity of gel bands is reached, rinse briefly with Milli-Q H2O and photograph the results (each team should bring a DIGITAL CAMERA to lab--iPhone/iPad will NOT work): place gel on light box; please clean light box with a Kimwipe after disposing of gel.

Note: The incubation times above are acceptable for gels up to 1.5 mm thick. Extend times by 5 minutes each for thicker gels.


Western Blot: Electrophoretic Transfer of Proteins from SDS-PAGE

*REFERENCE: Towbin, H., Staehelin, T., and Gordon, J. (1979). Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: Procedure and some applications. Proc. Natl. Acad. Sci. USA 76 (9): 4350-4354.

{Look at the Bio-Rad Mini Trans-Blot Cell Instruction Manual for detailed directions and illustrations.}

  1. Obtain a polyvinylidene difluoride (PVDF) membrane (Immun-Blot® PVDF Membranes, Bio-Rad) with the same dimensions as the gel and mark it in a corner with a ball point pen to identify the orientation of the membrane with the gel. Do not handle the gel or membrane with your bare hands--wear gloves! Do not allow the membrane to bend or fold.
  2. Obtain 700 ml per buffer tank of transfer buffer (10 mM CAPS, pH 11.0, 10 % methanol).
  3. Wet the membrane in methanol for a couple seconds and immediately place in transfer buffer; be sure not to let the membrane dry out after it is wetted with methanol.
    NOTE: the PVDF membranes are hydrophobic and do not wet well in aqueous solutions; if you SKIP the Methanol step, the transfer will be poor. If the membrane dries out, re-wet with methanol before putting in aqueous solutions.
  4. Completely saturate the pre-cut filter paper and fiber pads of the transfer unit by soaking them in transfer buffer. Avoid trapping air bubbles in the fiber pads or filter paper during the wetting process.

Assembly of the gel holder cassette
The following procedures should be performed while wearing gloves to prevent contamination of the membrane.

  1. Open the gel holder cassette by sliding and lifting the latch. Note that one panel of the cassette is clear and the other panel is tinted smoky gray. The clear panel is the anode (+) side and the gray panel is the cathode (-) side. For the electrode module, the cathode is the smoky gray electrode panel located in the center of the buffer tank. Always insert the gel cassette so that the gray plastic faces the gray plastic of the cathode electrode. The anode and cathode are identified by the black (cathode) and red (anode) disks on top of the electrode module.
  2. Place the opened gel holder in a shallow container (plastic storage box) so that the gray panel is flat on the bottom of the container. The clear panel should rest at an angle against the wall of the vessel. Pour some transfer buffer into the container (just enough to cover the panel).
  3. Place the fiber pad on the gray panel of the cassette panel and be sure all air is removed. When assembling the fiber pads, filter paper, gel, and membrane, center all components in the holder. If any of the material extends past the edge of the cassette, it will catch on the guide rails, causing a distortion of both the blot-gel contact and the transfer pattern.
  4. Place a piece of the saturated filter paper on top of the fiber pad. Be sure all air is removed from the filter paper and the layer below it. Place the gel on top of the paper. Align the gel in the center of the cassette. Transfer will be incomplete for any portion of the gel that has bubbles on the surface or lies outside the pattern of circles of the gel holder cassette. Make sure that no air bubbles are trapped between the gel and the filter paper. Bubbles can be removed by rolling a plastic test tube over the filter paper and gel.
  5. Place the prewetted PVDF membrane on top of the gel. This is best done by holding the membrane at opposite ends so that the center portion will contact the gel first. Then, gradually lower the ends. Next, roll a plastic test tube over the top of the membrane to exclude all air bubbles from the area between the gel and membrane.
  6. Complete the sandwich by placing another piece of saturated filter paper on top of the membrane and placing a saturated fiber pad on top of the filter paper.
  7. Close the cassette. Hold it firmly so the sandwich will not move and secure the latch. At this point, movement of the sandwich might disrupt the gel-membrane contact, which would cause incomplete transfers or swirling transfer patterns.
  8. Place the gel holder in the buffer tank so that the gray panel of the holder is facing the gray cathode electrode panel.
  9. Prepare a "dummy" cassette with two fiber pads (takes up volume so less transfer buffer is needed).
  10. Put the cooling tray in the back of the chamber. (The cooling tray containing "freezer pack" can be found in the chest freezer in B06.)
  11. Add the transfer buffer from the plastic container to the tank.
  12. If necessary, add more transfer buffer to cover the gels. (Do NOT fill the tank all the way to the top.)
  13. Place a stir bar in the buffer tank and use a stir motor to keep temperatures low during the transfer.
  14. Turn on the magnetic stirrer, and put the lid in place. Be sure that the electrode wires on the lid are attached to the appropriate pins of the electrode module. Plug the unit into the power supply. Normal transfer polarity is cathode to anode, i.e. red wire to red outlet and black wire to black outlet on the power supply.
  15. Turn on the power supply to initiate transfer. Set the voltage to 50V and transfer for 1 hour.

    Warning: Be certain the power supply reads "OFF" before handling the electrode connections. Electrical shock hazards are always present when using laboratory power supplies. Do not contact buffer reservoirs or wiring while the power supply is activated.
  16. After transfer is complete, turn off the power supply and disconnect the electrodes. Disassemble the cassette and mark the side of the membrane that was in contact with the gel if it is not already marked.
  17. Make pen marks on the membrane beside the colored standards and record their location as a sketch in the notebook.
  18. Place the membrane inside a clean paper towel and store at room temperature in your drawer.

PLEASE NOTE: The exposed wires in the electrophoresis equipment are platinum and are very expensive to replace. Please, only rinse these components with RO water, do not use a brush, and set to air dry. Be careful handling the devices to avoid breaking the wires. For drying, stack the units on the desk top so that no component is inside the other.

Do not throw away the pre-cut fiber pads--they are $5 each!

Copyright, Acknowledgements, and Intended Use
Created by B. Beason (bbeason@rice.edu), Rice University, 15 June 1999
Updated 29 May 2013