SDS Polyacrylamide Gel Electrophoresis (SDS-PAGE) Analysis of Purified. Fluorescent Protein. Table of Contents Fall 2012

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1 SDS Polyacrylamide Gel Electrophoresis (SDS-PAGE) Analysis of Purified Fluorescent Protein Table of Contents SDS Polyacrylamide Gel Electrophoresis (SDS-PAGE) Analysis of Purified Fluorescent Protein Student Version Introduction... 1 Laboratory Exercise... 2 Important Laboratory Practices... 2 Day 1: Prepare the SDS-PAGE samples... 3 Day 2: Run SDS-PAGE Gel... 4 Day 3: Dry the gel... 7 Day 4: Complete gel drying... 8 Worksheet: SDS-PAGE... 9 Acknowledgements... 11

2 SDS Polyacrylamide Gel Electrophoresis (SDS-PAGE) Analysis of Purified Fluorescent Protein Introduction Student Version Bacteria can be genetically engineered to produce a protein of interest. This protein may be used as a clinical drug or a research tool. In either case, it is important to verify the purity of the protein. A simple solution is to visualize the purified sample on an electrophoresis gel. The method commonly used to separate proteins is called SDS-PAGE (or sodium dodecyl sulfate polyacrylamide gel electrophoresis). In this procedure, an electrical field moves proteins through a gel matrix. SDS-PAGE, like horizontal agarose gel electrophoresis, separates the molecule of interest (protein in this case) by size. However, analyzing proteins is a bit more complicated than analyzing DNA. Unlike DNA, proteins can be positively charged as well as negatively charged. Furthermore, protein secondary and tertiary structures must also be overcome if proteins are to be accurately separated based on size. The detergent SDS plays a major role in addressing these issues along with the sulfur chemical, dithiothreitol. Proteins are globular in secondary and tertiary structure due to disulfide bonds, hydrophobic interactions, and hydrophilic interactions with their aqueous environment. Therefore, something must be done to break the secondary and tertiary structure of the proteins in the sample for accurate analysis of peptide size to occur. Sodium dodecyl sulfate (SDS) is a detergent possessing both a hydrophobic end (the dodecyl group) and a hydrophilic end (the sulfate group). The tertiary structure of most proteins often relies upon hydrophobic interactions at the core of the protein. The hydrophobic end of SDS breaks these interactions through interactions with the hydrophobic side chains of the amino acids. Similarly, a sulfate group can disrupt hydrogen bonding in secondary protein structure. Disulfide bonds holding tertiary or quaternary structure together can be broken by using a reducing agent, such as beta-mercaptoethanol (BME) or dithiothreitol (DTT). Finally, heating the protein sample also aids the denaturation and unfolding process allowing chemicals like SDS and DTT to interact with the protein. In addition to denaturing the protein, SDS also serves an additional purpose. Because each protein is coated with SDS molecules and the sulfate group has a negative charge, SDS also serves to give each protein molecule a net negative charge. This means that when an electrical field is applied to the gel in buffer, each protein molecule will move toward the positive electrode. This allows the acrylamide to separate the proteins based on size. You can think of polyacrylamide as a synthetic version of agarose. As the SDS-coated protein molecules move through the gel matrix, smaller molecular weight proteins are able to navigate through the pores in the matrix more quickly than larger ones. Thus, the proteins in the sample are separated by size and relative molecular weight. Once proteins have been separated by size, they must be visualized for analysis. As in agarose gel electrophoresis, two dyes are used during the visualization process. Loading dye again serves a dual role to enable you to visualize the progress of the protein movement in the gel matrix and increase the density of the sample to weigh down in the wells of the gel. The dye, bromophenol blue, provides the colored dye front during electrophoresis and glycerol gives the sample heavier density. Once the gel run is complete, a separate dye will bind to the protein in the polyacrylamide gel for the proteins to be visible during analysis. In this lab, a blue dye, called Coomassie Brilliant Blue (CBB), serves the same role as ethidium bromide in DNA gels. When the polyacrylamide gel is soaked with Coomassie dye, the protein bands become visible as the blue dye molecules bind to the peptides. Acetic acid in the staining solution fixes the proteins and dye in place. Because CBB is nonpolar, methanol is used as the solvent in the fix/stain solution. Excess dye is removed during the destaining step with a wash solution consisting of the fixative solution minus the dye through a diffusion process. In this protocol, your gel will be preserved following the staining and destaining steps. The gel drying solution has glycerol to prevent the gel from cracking. Compressing the gel between pieces of cellophane preserves the gel in a clear sheet. Once the cellophane and acrylamide dry and harden, it is possible to handle it easily and save the results for recording keeping. 1

3 Laboratory Exercise The protocol outlined below describes the procedure for running your purified protein sample on an SDS-PAGE gel. First you will heat the sample with a reducing agent in loading buffer to denature the proteins. Then you will load the sample on a polyacrylamide gel, which will separate the proteins based on size. After the gel run is complete, the gel is stained with Coomassie to visualize the proteins. After destaining, the gel can be dried and saved in your lab notebook. Objectives - student should be able to: 1. Successfully denature their FP purification lab samples and run it on an SDS-PAGE gel. 2. Understand why the various components are used to prepare and run samples. 3. Stain, destain, and dry their gels. a. Add reagents to the bottom of the reaction tube, not to its side. b. Add each additional reagent directly into previously-added reagent. c. Do not pipette up and down, as this introduces error. Rack or vortex tubes to mix. d. Make sure contents are all settled into the bottom of the tube and not on the side or cap of tube. A quick spin may be needed to bring contents down. Important Laboratory Practices a. Pipet slowly to prevent contaminating the pipette barrel. b. Change pipette tips between each delivery. c. Change the tip even if it is the same reagent being delivered between tubes. Change tip every time the pipette is used! Keep reagents on ice when needed Check the box next to each step as you complete it. 2

4 Day 1: Prepare the SDS-PAGE samples 1. Obtain 4 clean microfuge tubes and label as follows: 1) GFP lysate 2) Wash 3) Purified GFP 4) Blank Also label each tube with your group number. 2. Using a P-20, add 5 µl of 5X Loading Dye containing DTT to each tube. 1" 2" 3" 4" 3. Using a P-20, add 20 µl of the GFP lysate to tube #1 you labeled in step 1. Using clean tips in between, repeat for the wash and purified GFP tubes. Add 20 µl of distilled water into the tube labeled blank. Note: All tubes should now contain 25 µl of your sample and loading dye mixture. 4. Place a tube cap lock onto each microfuge tube and heat the samples for 3 min at 85 C in a heat block. Note: This heat step denatures your protein. 1" 2" 3" 4" 5. Leave the samples at room temperature for a few minutes till cool enough to handle. If you will not be loading the SDS-PAGE gel today, store the protein samples you have prepared at 4 C. 3

5 Day 2: Run SDS-PAGE Gel 1. [Skip this step if you are proceeded nonstop from Day 1.] Remove your samples from the fridge and heat them at 37 C for 2 minutes. 2. Leave the samples at room temperature for a few minutes till cool enough to handle. 3. Obtain a vertical gel box and share a 12-well Tris-Glycine gel. Note: Two groups will share a gel and a gel box. Gloves should be worn at all times when working with acrylamide gels. 4. Open the package and rinse the gel briefly with water. Remove the tape covering the bottom part of the gel. 5. Place the gel cassette onto the gel rig such that the opening to the wells are facing inward and the label of the gel is legible from the outside. Note: Verify that the cassette is seated properly and facing the correct direction. Also, be careful not to over-tighten the clamps (stop tightening once you feel resistance). That may crack the gel cassette. 6. Carefully pour 1X Running Buffer into the top buffer chamber. Once the chamber is above the opening of the wells, wait a minute to verify that the chamber is not leaking through to the bottom. Tighten the clamps a little bit more if you observe leaking. Once you have verified that the chamber is not leaking, remove the comb and save it for later. Note: Be sure to save the comb. It will be used to remove the bottom of the gel in a later step. 7. Fill the bottom chamber of the gel rig to the designated fill line. If necessary, add additional buffer.!! 4

6 8. Using a P-20, load the gel as follows: Lane 1: 15 µl Blank sample Lane 2: 5 µl of the protein ladder Lane 3: 15 µl Lysate sample Lane 4: 15 µl Wash sample Lane 5: 15 µl Purified GFP sample Lane 6: Skip Lanes 7-12: repeat lane assignments for Team 2 Note: Try to depress the micropipette button as slowly as possible to avoid splashing any sample into the next well. Keep an eye on your sample during loading. Remove pipet tip slowly to prevent disturbing the sample. 9. Once all of the samples have been loaded, carefully place the cover onto the gel box. Connect the electrode leads to the power supply. Set the power supply to 200V and allow the gel to run for 40 min or until the dye front has reached near the bottom. Once the dye front reaches the bottom, turn off the power supply and remove the gel box cover. Teacher s Note: The gel run can be as short as 30 minutes. Shorter runs merely decrease the distance between bands but do not alter any conclusions that would otherwise be made. 10. Carefully transfer the gel box to a basin to catch the running buffer. Pour the used 1X Running Buffer from the basin and gel box back into a buffer bottle using a funnel. Pour slowly enough to avoid pouring the gel pieces into the bottle! 11. Loosen the clamps and remove the gel cassette. Rinse it briefly with distilled water. 12. Carefully use a spatula to break the seals between the pieces of the plastic cassette covering the gel. To avoid ripping the gel, try to minimize the degree to which the gel is opened until you have completely broken all the seals. Carefully remove the top half of the cassette and throw it in the trash. Note: The gel may stick to either side of the cassette. Simply remove the half to which it has stuck least. 5

7 13. Obtain a gel tray and fill it halfway with water. Place the cassette into the water in the tray with the gel facing down. Use your finger to carefully remove the gel from the cassette into the water. Throw the remaining plastic cassette into the trash. Gently shake the tray back and forth a few times to rinse the gel. Note: Water tension will help pull the gel off the cassette. To avoid ripping the gel, only use your finger initially to detach the gel from the plastic. The water tension should do the rest. 14. Carefully pour the water into a sink. Note: Lightly keep one or more fingers on the gel to prevent it from leaving the tray. Be careful not to use too much force on the gel or you will rip it. 15. Carefully pour enough Coomassie stain into the tray to barely cover the gel. Allow the gel to stain for 10 min. Note: Avoid splashing! Coomassie dye will stain clothing and skin, so take care when using it. 16. Once staining is complete, place a funnel in the mouth of the Commassie stain bottle and carefully pour the staining solution back into the bottle. Note: Be sure to lightly hold the gel in place with a finger while you pour or you accidentally may pour your gel into the bottle. However, take care to use as little force as possible when holding the gel or you may rip it. 18. Carefully pour enough Destain Solution into your tray to cover your gel. Place 4 crumpled KimWipes into each corner of the tray to absorb the dye more quickly. Note: Avoid placing the KimWipes on your gel. It might stick. Close the cover on your tray and allow the gel to destain overnight. 6

8 Day 3: Dry the gel (Optional) 1. Squeeze the excess liquid from the KimWipes into the tray and dispose the KimWipes in trash. Pour off the Destain into a chemical waste container. Note: Be sure to lightly hold the gel in place with a finger while you pour or you may accidentally pour your gel into the bottle. However, take care to use as little force as possible when holding the gel or you may rip it. 2. Carefully pour enough water into the tray to cover the gel. Let gel rinse then carefully pour off the water. Cut two 11x11 cm pieces of cellophane. 3. Carefully pour enough Gel Drying solution into the tray to cover the gel. Note the time or start a timer. The gel should not remain in the gel drying solution for longer than 5 minutes. Soak the first cellophane sheet in the gel drying solution with the gel for 1 minute. 4. Place one of the plastic plates on your benchtop. Once the cellophane sheet has soaked for a minute, carefully place it on the plastic plate. Use your fingers to carefully swipe away any bubbles produced under or on top of the sheet. Note: To avoid bubbles, carefully lay the sheet onto the plate slowly from one end to the other. Pick up the sheet and lay it down again if significant bubbles are produced. 5. Carefully remove the gel from the gel drying solution and place it on the cellophane sheet on the plastic plate. Use your fingers to wipe away any bubbles produced under or on top of the gel. Note: To avoid bubbles, carefully lay the gel onto the cellophane slowly from one end to the other. Pick up cornes of the gel and lay it down again if significant bubbles are produced. 7

9 6. Soak the second cellophane sheet in the gel drying solution for 1 minute. After the second cellophane sheet has soaked for a minute, carefully lay it on top of the gel. With your finger, swipe away any bubbles produced under the cellophane sheet. Note: To avoid bubbles, carefully lay the cellophane onto the gel slowly from one end to the other. Pick up the cellophane and lay it down again if significant bubbles are produced. 7. Cover the gel with the second plastic plate. Use the binder clips to seal the plastic plate, gel, and cellophane sandwich. Allow the gel to dry overnight. 8. Place a funnel into the mouth of the bottle of Gel Drying solution and carefully pour the excess gel drying solution back into the bottle. Day 4: Complete gel drying (Optional) Remove the binder clips and one of the plastic plates. The gel and cellophane should have hardened into a single plastic sheet. If desired, you may trim the excess cellophane from the edges of the gel. Take a picture of the gel for each group member. Tape the gel in your lab notebook. 8

10 Name Date Period Worksheet: Polyacrylamide Gel Electrophoresis 1. Briefly describe how you made the purified protein sample you analyzed in this lab. 2. Briefly explain one reason a scientist might wish to perform SDS-PAGE. 3. Briefly explain two roles that SDS plays during the procedure. 4. Briefly explain what role acrylamide plays in the electrophoresis process. 9

11 5. Professor Farnsworth s graduate student forgot to add DTT to the loading dye before running her gel. What (if any) changes do you expect to see on her gel? Be sure to explain the reasoning for your answer. 6. Briefly explain the difference between the roles that Bromophenol Blue and Coomassie Brilliant Blue dyes play in SDS-PAGE. 7. Why did you include the crude lysate in your experiment? Why did you include the wash buffer in your experiment? 8. Professor Gonzales would like to make 500 ml of 1M Tris-HCl, ph 6.8 to use in the SDS-PAGE procedure. Briefly explain what he should do to make this solution. MW of Tris base = g. 10

12 BABEC Educational SDS-PAGE Kits For Research Use Only. Not for use in diagnostic procedures. BABEC thanks LifeTechnologies for their generous support of gels for SDS-PAGE lab. BABEC thanks New England BioLabs (NEB) for their generous support of reagents for SDS-PAGE lab. 11

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