Lab 3 Exercises

Lab 3 Exercises

Completion and Submission of Assignment: 25 points

Use the lab 3 material provided in Blackboard and any internet search engine to answer the following questions (please type). Each question is worth 2 points. This is a two part lab. The online portion is the following week.   

1) What is the purpose of using SDS in this experiment?

2) Why is the reducing agent mercaptoethanol used in this experiment?

3) Why are pre-cast gels commonly used now in research labs and industry? 

4) Why would it be useful to determine the molecular weight of a protein? 

5)  Why is it important to not release the plunger before all the sample has been ejected?

6) After incubated with SDS, which direction do proteins move in the gel?

7) In SDS-PAGE, do higher MW or lower MW proteins move further in the gel?   

8) What is the purpose of bromophenol blue? 

9) How is the molecular weight of the protein determined?

10) Why is the tape removed at the bottom of the gel? 

5 point question

11) Watch the following video from Biorad. Write a short 6-10 step by step procedure based on the video for running a gel using the Biorad system. 

Experiment 3: Protein Molecular Weight Determination via SDS-PAGE Electrophoresis and Biorad Quantity One 1D Image Analysis  

Safety Considerations: Major hazards: electrical/shock hazard and use of acrylamide. Students must wear gloves and safety glasses during this experiment. All waste must be disposed of according to the instructor’s/GA’s directions. Students will be required to clean up their area upon completion of the experiment.

Introduction:

Proteins vary significantly in their properties including molecular weight, amino acid composition, pI, shape and solubility. Using known amino acid compositions/protein sequences (e.g. from bioinformatics lab 2), the exact molecular weight of a polypeptide can be calculated; however, even if one knows this information (from gene or peptide sequence data), molecular weight determination of known proteins must still be undertaken to confirm identity a particular protein during purification, etc. Molecular weight determination of a protein is therefore an essential technique used in the biochemistry laboratory. Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) is the most common method for the rapid determination of protein molecular weight. More sophisticated techniques such as mass spectrometry allow for highly accurate molecular weight determination of peptides and proteins; however, these techniques require costly instrumentation/sample preparation and are thus less flexible for the day to day analysis of proteins in the laboratory.  

Polyacrylamide gels are formed by mixing the monomer, acrylamide, the cross- linking agent, methylenebisacrylamide, and a free radical generator, ammonium persulfate, in aqueous buffer. Free radical polymerization of the acrylamide occurs and at various points, the acrylamide polymers are bridged to each other. The pore size in polyacrylamide gels is controlled by the gel concentration and the degree of polymer cross-linking. Higher percentage gels are more suitable for the separation of smaller proteins. Polyacrylamide gels can also be prepared to have a gradient of gel concentrations. Typically the top of the gel (under the sample wells) has a concentration of 5%, increasing linearly to 20% at the bottom. Gradient gels can be useful in separating protein mixtures that cover a large range of molecular weights. Gels of homogeneous concentration (e.g. 12%) are better suited for achieving wider separations of proteins that occupy narrow ranges of molecular weights. It should be noted that acrylamide is a neurotoxin and can be absorbed through the skin. However, in the polymerized form it is nontoxic. In this lab, we will be using pre-cast gels to reduce the hazards associated with the presence of unpolymerized acrylamide. Moreover, these “state of the art,” high resolution gels are commonly used in both industry and research labs throughout the world.

Sodium dodecyl sulfate (SDS) is a detergent which consists of a hydrocarbon chain bonded to a highly negatively charged sulfate group. SDS binds strongly to most proteins and causes them to unfold to a random, rod-like chains. No covalent bonds are broken in this process. Therefore, the amino acid composition and sequence remains the same. Since the protein’s specific, three-dimensional shape is abolished, the protein no longer possesses biological activity. Proteins that have lost their specific folding patterns and biological activity but have their intact polypeptide chains are called denatured. Proteins can contain covalent linkages known as disulfide bonds. These bonds are formed between two cysteine amino acid residues that can be located in the same or different polypeptide chains. High concentrations of reducing agents, such as beta-mercaptoethanol, can break disulfide bonds. This allows SDS to completely dissociate and denature the protein.

In most cases, SDS binds to proteins in
a constant ratio of 1.4 grams of SDS
 per gram of protein. The amount of negative charge of the SDS is much more than the negative and positive charges of the amino acid residues. The large quantity of bound SDS efficiently masks or obliterates the intrinsic changes of the proteins of interest. Consequently, SDS denatured proteins are net negative in charge and since the binding of the detergent is proportional to the mass of the protein, the charge to mass ratio is constant. When loading the gel, all protein samples will contain buffer, SDS, beta-mercaptoethanol as the reducing agent to break disulfide bonds, glycerol to create density greater than that of 
the electrode buffer and the negatively charged tracking dye bromophenol blue. The tracking dye will migrate ahead of the smallest proteins in these samples towards the positive bottom electrode. During SDS electrophoresis, the proteins migrate through the gel towards the positive electrode at a rate that is inversely proportional to their molecular weight. In other words, the smaller the denatured protein, the faster it migrates. The molecular weight of an unknown protein is obtained by the comparison of its position after electrophoresis to the positions of standard SDS denatured proteins. After proteins are visualized by staining and destaining the gel, their migration distance is measured. The log10  of the molecular weights of the standard proteins are plotted versus their migration distance. The molecular weight of unknown proteins are then easily calculated from the standard curve using either Excel or the Biorad Quantity One software.

To reiterate, the protein unknowns in this lab will be denatured with the anionic detergent sodium dodecyl sulfate (SDS) at ~ 100°C. Under the experimental conditions, the proteins will have a mobility in the gel that is inversely proportional to the logarithm of their molecular weights. Pre-stained proteins of known molecular weights will be separated by electrophoresis in parallel with the unknown samples. The standards will be used to estimate the molecular weights of the unknown protein samples by either graphical analysis using either Excel or Biorad Quantity One software. Since the standard proteins are prestained, the individual bands will be visible during electrophoresis and can be observed while they migrate through the gel. Each group will be assigned an unknown protein (unknown number will be given) of interest and the molecular weight of that particular protein will be determined.

Procedure (warning: this experiment involves a potential electrical hazard and can only be done under the direct instructor’s supervision):

1. Bring a beaker of water to a boil in a microwave (your GA will assist with this step). Remove from heat.
2. Dilute 33 mL your unknown protein with 67 mL of sample buffer containing SDS, mercaptoethanol, glycerol and tracking dye in a microcentrifuge tube.
3. Using a flotation device/rack to hold the tube, place the capped tube in the hot water bath for 5 minutes.
4. Remove the tubes from the flotation device and let cool for 1-2 minutes. 
5. Prepare the SDS-PAGE running buffer according to your instructor’s/GA’s instructions. Typically, 500 mL of running buffer is prepared by diluting a pre-bought 10X concentrate (50 mL diluted to 500 mL, 10 fold dilution). 
6. Precast polyacrylamide gels will vary slightly in design depending on the manufacturer. Procedures for their use will be similar (your instructor will demonstrate the exact procedure for gels used in this lab). Open the pouch containing the gel cassette with scissors. Remove the cassette and place it on the bench top with the front facing up. 
Note: The front plate is smaller (shorter) than the back plate.
7. Some cassettes will have tape at the bottom of the front plate. Remove all of the tape to expose the bottom of the gel to allow electrical contact.
8. Remove the comb by placing your thumbs on the ridges and pushing (pressing) upwards, carefully and slowly.
9. Wash the wells with running buffer X3.
10. Different electrophoresis units will vary by manufacturer. Your instructor will provide directions on how to properly set-up the gel system including safety guidelines for each type of unit. 
11. In general, place the gel cassette in the electrophoresis unit in the proper orientation. The protein samples will not separate in gels that are not oriented correctly or the unit may leak. Follow the directions accompanying the specific apparatus as outlined by your instructor.  
12. Add the diluted running buffer into the upper and lower buffer chambers. The sample wells and the back plate of the gel cassette should be submerged under buffer.
13. Load standards and samples (10 mL) into the assigned wells for each group. Note: samples and standards for each lab group will be assigned depending on the numbers of students and types of samples to be analyzed each term. Your instructor will set-up a practice gel system in order to practice pipetting samples before loading samples in the actual experimental gel system.
14. To pipette samples, place a fresh fine tip on the micropipette. Next place the lower portion of the fine pipet tip between the two glass plates, below the surface of the electrode buffer, directly over a sample well. The tip should be at an angle pointed towards the well. The tip should be partially against the back plate of the gel cassette but the tip opening should be over the sample well.
Do not try to jam the pipet tip in between the plates of the gel cassette. Eject all the sample by steadily pressing down on the plunger of the automatic pipet. Do not release the plunger before all the sample is ejected (this step is very important!). Premature release of the plunger will cause buffer to mix with sample in the micropipette tip. Release the pipet plunger after the sample has been delivered and the pipet tip is out of the buffer.
15. Place the cover on the gel unit and attach the power leads according the manufacturer’s and instructor’s instructions. Warning: this step can only be carried out under the direct supervision of the instructor.
16. Set the proper power supply voltage and length of run according to the manufacturer’s and instructor’s directions. Warning: this step can only be carried out under the direct supervision of the instructor.
17. Once the run is complete, remove the gel from the electrophoresis unit. Warning: this step can only be carried out under the direct supervision of the instructor.
18. Remove the gel from the cassette based on the manufacturer’s and instructor’s instructions.
19. Remove the residual SDS from the gel by placing the gel in a Tupperware dish containing ~ 200 mL of distilled water. Gently shake the gel for 10 minutes using an orbital shaker. Repeat this step for a total of 3X making sure to replace the distilled water after each wash.
20. Remove the last volume of distilled water and add 20-50 mL of stain to the gel (the gel should be completely covered with stain).   
21. Stain the gel for 1 hour under gentle agitation (orbital shaker). 
22. Remove the stain from the gel and replace with ~200 mL distilled water. Gently shake the gel for 2-5 hours using an orbital shaker. Repeat this step X3 replacing with fresh water after each step. 
23. Determine the MW of each unknown protein by comparison to the protein standards. This procedure will be accomplished using the Biorad Quantity One software. Students will also be required to determine the MW of each protein by manually measuring the band/protein standards and using Excel to plot a graph of Log MW versus migration distance. Your instructor/GA will assist with both of these exercises.  

References:

Lab adapted from Edvotek Biotechnology Education Company.