MOLECULAR WEIGHT DETERMINATION & BY ELECTROPHORESIS and MALDI-TOF MS Introduction: Electrophoresis is a commonly used technique in bioanalytical chemistry. In gel electrophoresis, molecules migrate through a gel medium according to their charge (applied by the electrodes). In the case of proteins, charge is proportional to mass and therefore molecular weight resolution is achieved. Mass separation is also achieved by the composition of the gel. These basic principles apply to all types of electrophoresis but there are small differences depending on the molecule of interest. Electrophoresis is an attractive techni que due to low cost and simplicity. It is a powerful separation technique, however only gives a crude estimate of molecular weight and identity. It is often used in in combination with more precise analytical methods. Since proteins are so big in size many need to be digested into smaller pieces to fit the mass range of instruments. Enzymatic digestion can be performed on specific bands in the gel after electrophoresis or in solution of a mixture of proteins. The benefit of in gel digestion is the separation of different species, but the extraction is a fairly long and laborious process. In solution digestion is quicker and less work; but provides no separation prior to digestion resulting in a much more complex sample for mass spectrometry. Due to time constraints, you will perform in solution digestion on a solution of a single protein which was also used for the Electrophoresis. MALDI Matrix Assisted laser desorption Ionization is a form of mass spectrometry. Samples are crystalized with a matrix, spotted on a target and analyzed in a time of flight mass spectrometer. The ionization in this case is performed by a laser. There are many technical considerations in maldi such as type of matrix; sample preparation, interfering agents and competitive ionization. It will help you develop your discussion if you familiarize with these aspects a little prior to the experiment or while writing your report. The first week you will perform the digestion and the electrophoresis. The second week will be sample cleanup, preparation and MALDI. Prelab: 1) What different types of biomolecules are analysed by gel electrophoresis? 2) What is reverse pipetting? Why is it a benefit to use this pipetting technique for these applications? 3) Briefly describe the basic components of a generic MALDI-TOF mass spectrometer and their function. 4) You will need to be aware of what you are looking for in the MALDI samples; which requires knowledge of the theoretical tryptic peptide masses, including modifications. Also the intact masses of B.S.A., myoglobin and lysozyme and IgG. There are many databases/tools to help with this: a. Go to the website http://web.expasy.org/peptide_mass/ b. Type in PO2769 in the box. 1
Notes: c. Select: [M+H] +, average, and choose iodacetamide in the drop down box for cysteines treated with. Select tryspin as the enzyme and allow for 1 missed cleavages it should give peptides with 0 missed as well as 1 missed. d. Click perform cleavages. Once the data appears, scroll down to the bottom and export the data in raw text format; then copy to excel. You don t need to hand this in, bring it to the MALDI portion of the lab for you to use. e. Notice: You are given both modified and unmodified masses. You should look for both! - Your station is located across from the GC-MS (between it and the ICP). Everything you need should be located in the drawers below or on the bench. - Refrigerated samples and reagents are in the fridge in the room between MCAL and McKenna Lab if already not on the bench. Please replace them to the fridge once finished. - The digestion procedure is not complicated, but requires organization while doing the electrophoresis so plan ahead! You should get it started first, but make sure to keep track of the times! Protocol: Week 1: SDS-PAGE and In-Solution Digestion Part A: Electrophoresis 1. Turn on the water bath and ensure adequate water. Bring the water to a boil. 2. Prepare 1x Tris-Glycine Running buffer from the 10x stock. You will need 800 mls in total. 3. The sample buffer and reducing agent are supplied as 4x and 20x respectively, and stored at room temp. They need to be at 1x in the sample, see table below. 4. Use the multicolor 500 ul centrifuge tubes to prepare. ADD THE B2ME LAST AND IN THE HOOD ITS VERY VOLATILE AND STINKS! 2
Samples: Sample Concentration (mg/ml) Volume Sample Required (ul) Volume Sample Buffer (ul) Volume B2mE* (ul) Volume Mill-Q (ul) Final Volume (ul) IgG 2 25 25 5 45 100 B.S.A. 2 7 25 5 63 100 Myoglobin 2 20 25 5 50 100 Lysozyme 2 7 25 5 63 100 Protein Mix 2ug/mL 25 25 5 45 100 Protein Shake 5 5 25 5 65 100 *this is beta-2-mercaptoethanol. 5. The prestained standards (Precision Plus protein standard) do not need to be boiled. You will only need 10 ul and can be loaded directly on to the gel. Load 2 lanes if you choose. 6. Give the samples a quick vortex and tap on the counter to collect liquid at the bottom. Boil the samples for approximately 5 minutes using the floating rack in a beaker on the hot plate. Assembling the Gel Apparatus: 7. Remove the gels from the pouch. Remove the tape at the bottom. Slide the comb out by GENTLY pulling up in one smooth motion. Be careful. If you damage the stacking gel in this step, those lanes become useless. Rinse the wells with running buffer using a Pasteur pipette about 4x. 8. Set the gel holder to the open position on a clean, flat surface 3
9. Place the cassettes with the wells of the gel facing inward If using only one gel, the buffer dam will replace the second cassette. Make sure the assembly remains balanced and does not tip over. The cassettes will rest at a 30 degree angle. (B) 10. Gently push the cassettes together, making sure they rest firly and squarely against the green gasket at the bottom of the assembly. Align the short plates and check that they align just below the notch at the top of the green gasket. (C) 11. Maintain gentle pressure on each gel cassette. Slide the green arms of the clamping frame one at time over the gels, locking them in place, The wing clamps lift the gel cassettes into the notch, 4
forming a tight seal. Check to ensure everything is square and sitting flat, then pour a small amount of running buffer into the inner chamber to ensure it is not leaking. 11. Place the assembly inside the module. Then fill the outer chamber with the rest of the running buffer, approximately to the line indicating 1 gel. Loading Samples and Running the Gel: 12. Load 40 ul into each well (except standards only 10) using reverse pipetting technique and the gel-loading tips provided. Make sure you arrange your samples so that you will be able to tell what s what should it get flipped in the process. 13. Place the lid onto the gel box. It will snap into place if done correctly. 14. Connect the electrodes to the power supply and switch it on. Set the voltage to 150, then press run. Look for a steady stream of air bubbles. They should be obvious, if not there is no conductivity. Observe the dye front to make sure it begins to migrate. It should be relatively 5
straight across all the lanes. If not, consult the instructor within 5 minutes of starting the run. Removing the cassette and staining the gel: 15. Once the dye front reaches the bottom, press Stop. Turn the power supply off and disconnect the electrodes. Remove the lid from the apparatus and remove the get cassette from the assembly. 16. To open the cassette, align the arrow on the opening lever with the arrow on the cassette. Apply downward pressure and the cassette will snap apart. 17. Pull the two plates apart from the top, and gently remove the gel and place in staining reservoir. BE CAREFUL IT IS REALLY EASY TO RIP THE GEL. There is a lip at the bottom which is thicker and can be used to lift off if necessary. The gel can be dislodged around the edges by carefully running a pipette tip around the edges between the gel and the cassette. 18. Rinse the gel 5x with Millipore water. Pour the Imperial protein stain, just enough to cover the gel. Shake for a few minutes. You might see bands after about 5-10 minutes, but it might take longer. Leave stain on overnight. 19. Cover your gel with saran and label with your lab# and names. The lab instructor will decant stain the next morning and replace with Millipore water periodically throughout the next day, until the background is resolved. A picture can be taken during the following lab period when you are doing the next part of the lab. Part B: Digestion Solutions: 1) 100 mm NH4HC03 (F.W. 79.056): Weigh 0.0395 g. Bring volume to 5 mls with DDH20 4) 100 mm D.T.T. Dilute the stock given to you accordingly with NH4HC03 5) 200 mm IAA - Dilute the stock given to you accordingly with NH4HC03 5) Trypsin Should be added to the samples at a ratio of 1:50. Protocol: 1) Prepare 100 mm D.T.T. using the 1M stock provided to you. There is 20 ul of 1M in the tube provided. 2) Aliquot 50 ug (notice quantity not volume!!) of the B.S.A. (2mg/mL) Add D.T.T at a final concentration (in the samples) of 10mM (1/10) 3) Dilute the B.S.A aliquot to a concentration of 1 mg/ml using NH4HC03. 4) Incubate for 45 min @37 degrees Celsius in the water bath. 6
5) During incubation, prepare dilute the 500 mm stock solution to 50mM and obtain some ice from the 4 th floor in a beaker 6) Remove tube from 37 degrees and quickly place on ice to cool to room temperature. 7) Immediately add IAA at final concentration of 20mM in the sample. 8) Incubate 30 min @ r.t. in the dark (cover tube with foil). 9) Add 0.8 ul DTT to neutralize I.A.A and prevent trypsin alkylation. Incubate 20 min @ r.t. 10) Remove the trypsin from the freezer (one of the 0.5 ml tubes) 11) Add trypsin at a ratio of 1 trypsin:50 protein and incubate overnight @ 37 degrees. Return the trypsin to the freezer immediately. Commented [JB1]: M The technician will look after putting your samples away in the fridge the next day. Make sure they are clearly labelled with your group #. They will be stored in the freezer until the following week. WEEK 2: MALDI-TOF MS. You will perform the sample clean up and prepare samples for MALDI in the MCAL laboratory. The spotting on the target and analysis will be performed upstairs in room 534 with the assistance of the technician or TA. Part A: C18 Ziptip Peptide Clean-up Protocol for LC/MS Introduction: Many buffers and compounds are not Mass Spectrometry compatible, such as those found in biological fluids or those used in the tryptic digestion process. (ie Urea, guanidine, DTT, etc.) Each of these tips contains a C18 sorbent to effectively bind the peptides, wash contaminants and finally elute peptides in mass spectrometry compatible solvents. Important Notes: - Sample must be free of organic solvents. - Avoid introducing air into the membrane, this will dry it and effect binding - Have everything prepared and ready before starting - DO NOT WASTE TIPS THEY COST $3 EACH! - The ziptips are for the digests only not needed for the protein samples Solutions: 7
0.1% FA 5% Acetonitrile, 0.1% FA 50% Acetonitrile, 0.1% F.A. Wet and Equilibrate Tip 1. Prepare all the solutions. 1 ml will be plenty. Use the 1.5 ml Eppendorf tubes. 2. Transfer 10 ul of your uncleaned digest sample into a 1.5 ul Eppendorf tube. 3. Wet tip: Aspirate and Dispense tip (discard on Kimwipe) with 10 L of 50% Acetonitrile. Repeat 2x. 4. Equilibrate: Aspirate and Dispense with 10 L of 0.1% F.A. Discard. Repeat 2x. 5. Bind: Aspirate the 10 L of sample into the tip. Depress the plunger, but do not go all the way to the stop. Re-asiprate the sample, (release the plunger) and repeat this cycle for a total of 10x. 6. Wash: Wash the tip by aspirating 10 L of 5% Acetonitrile/0.1% F.A. Discard wash. Repeat 2x. 7. Elute: Aspirate 10 L 50% Acetonitrile/0.1% F.A. Dispense into a microcentrifuge tube. Repeat 2x with a fresh 10 L Acetonitrile. Part B: Maldi TOF MS Solutions: 1. Matrix Solvent A: 30% Acetonitrile in 0.1% TFA (also called TA-30). 2. Matrix Solvent B: Ethanol 3. Matrix A: 20 mg/ml CHCA (alpha-hyrdoxy cinammic acid) in Matrix Solvent A. 4. Matrix B: 20 mg/ml DHB in Matrix Solvent A. 5. Matrix C: 20 mg/ml SPA in Matrix Solvent A 6. Matrix D: 20 mg/ml SPA in straight Ethanol. Samples: 1. B.S.A peptide digest both Matrix A and Matrix B. You will also be provided with a control digest; previously prepared in the same way you have done so. This is just in case the digestion you performed did not work properly. 2. Mixed Proteins (4 components) Matrix C (SPA only) in water only. 3. B.S.A. intact protein Matrix C (SPA only) 4. Lysozyme intact protein in Matrix C(SPA only) X 2 : one in water and one in NH4HC03 5. You will analyze all of the intact proteins (excluding the mixture) using Matrix D, but these are not prepared in tubes. Matrix D is not mixed with the sample; it will be spotted directly on the target. 8
Protocol: The matrices are already prepared for you. After ziptipping your peptides, you will just need to prepare the samples (mix with matrix) and spot on the target. 1. Mix together 5 ul of Matrix with 5 ul of each sample in 500 ul Eppendorf tubes NOTE: Matrix A and B for the peptide digest : prepare a separate sample with each. Matrix C is for the intact proteins. 2. Spot on the target: For the peptides; 1 ul of the mixed matrix and sample on the spot. For intact proteins, 0.3 ul of Matrix D first. Then 1 ul of mixed sample and Matrix C on top. 3. Dry the target It should only take a few minutes to dry. Write down some observations regarding the different types of crystals you see. 4. Analyze on the MALDI! Either the TA or the Emy will be helping you with this. Post Lab Questions: 1. What other types of Mass Spectrometry are frequently used in proteomic analysis? 2. What was the physical difference in the appearance of the matrix crystals? How does this affect the analysis? 9
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