CHRISTIAN LAB WESTERN BLOT PROTOCOL

CHRISTIAN LAB WESTERN BLOT PROTOCOL There is actually 2 parts to a western blot: A. SDS-PAGE: Separates protein by size. Smaller proteins migrate faster through the gel than larger proteins. Size separation is most accurate in the middle of the gel. Higher gel percentages will separate smaller proteins and lower gel percentages will separate larger proteins (see Appendix A). B. Immunoblot: The proteins that have been separated by SDS-PAGE have been transferred to a nitrocellulose membrane, thus maintaining their relative positions. We can now determine the size and relative amount of our specific protein using an antibody raised against that protein. To visualize the interaction, a secondary antibody is used that is coupled to horse raddish peroxidase (HRP). In presence of a chemiluminescent substrate, HRP generates a light signal which can be detected with X-ray film or using a specialized camera. Day 1: prepare samples and/or pour gels (2h) Day 2: (pour gels) run gels, transfer to nitrocellulose, blocking & add primary Ab (4-5h) Day 3: develop western blot (2h) A. SDS-PAGE: 1. PREPARE SAMPLES MATERIALS (see Appendix B for recipes) Lysis Buffer: the one you choose depends on what type of protein you are looking at and how strong you need the lysis buffer to be Examples of possible choices: 1% Triton-100 LB, RIPA LB Protease and Phosphatase Inhibitors 1. phenylmethylsulfonyl fluoride (PMSF): protease inhibitor,10mg/ml in isopropanol, use at 1/100 2. Aprotinin (protease inhibitor): stock 1000X 3. Leupeptin (Leu): stock at 1mg/ml, use at 1/100 4. Sodium OrthoVanadate (Van): tyrosine phosphatase inhibitor, stock at 100 mm, use at 1/100 5X Sample buffer (SB) Working solutions Lysis buffer + inhibitors (LB+I) make up enough lysis buffer containing protease and phosphatase inhibitors (see above) for all your samples + 10% 1X Sample buffer dilute 5X sample buffer in water to make 1X sample buffer S.L. Christian September 2008 edited: Feb 2011 page. 1

PROCEDURE 1. Lyse cells in appropriate LB+I 2. Determine protein concentration using BCA assay (Appendix C) 3. Make up samples to contain 1X SB and equal protein concentrations up to the same total volume (see below for example) Concentration Volume cell Volume 3X BPB Total (µg/µl) lysate (µl) H 2 0 (µl) (µl) X=7 µg 14- X µl 7 µl 21 µl Example 0.585 11.96 2.04 7 21 µl 1. incubate at 100 C (ie. boil) for 5 min. 2. Let samples cool for a minute, then do quick spin in centrifuge to get all liquid to bottom of tube. 3. Make up standards to same volume as samples (eg. 15 µl of sample buffer + 4 µl of prestained standards) 2. POUR GELS Note: Acrylamide is toxic, wear gloves at all times! MATERIALS SDS-PAGE Separating Gel Recipes Makes 7 ml= 2 gels 6% 8% 10% 12% 15% dh 2 O 993 µl x 4 862 µl x 4 3 ml 800 µl x 3 825 µl x 2 1.5 M Tris 937 µl x 2 938 µl x 2 938 µl x 2 975 µl x 2 975 µl x 2 ph 8.8 30% 750 µl x 2 675 µl x 3 825 µl x 3 1000 µl x 3 937.5 µl x 4 Acrylamide (29:1) 10% SDS 75 75 75 75 75 10% APS 75 75 75 75 75 TEMED 6 4.5 3 3 3 SDS-PAGE Stacking Gel Recipe 4.5% ; 3 ml per 2 gels dh 2 O 860 µl x 2 0.5 M Tris ph 6.8 760 µl 30% Acrylamide 500 µl 10% SDS 30 µl 10% APS 30 µl TEMED 3 µl S.L. Christian September 2008 edited: Feb 2011 page. 2

PROCEDURE Assembling plates and Pouring Gels 1. Acrylamide and 10% APS at 4 C, everything else at RT. 2. Wearing gloves, wash front gel plate (has notch) and back gel plate (no notch but has rounded corners) with dilute 7X and scrubber. Rinse extensively with distilled water and dry. Lay plates down on paper towel. 3. Rinse and dry 3 spacers. Make sure they are all the same size. 4. Wash the plates with 100% EtOH and wipe with kimwipes. 5. Take the back plate and wrap a yellow (0.75 mm) gel wrap around it. The breaks in the gel wrap should correspond to where the corners are. 6. Lay the plate down on the table and place spacers against the left and right edges. The rounded part of the spacer should fit into the rounded corner. Make sure the spacer touches the gel wrap but it is important that it the spacer is not under the gel wrap (this will cause leaks). 7. Carefully lay the top plate (clean side down) onto the bottom plate. 8. Pick up the two plates and square them up. Use the extra spacer to push one of the spacers back into place, if necessary. Put two clamps over the spacer (be sure to use the clamps that close all the way; the ones with wider openings are for assembling the plates into the gel box). 9. Push the other spacer back into the correct position and place two clamps over it. 10. Put two clamps on the bottom side and stand up the assembled gel plates. 11. Use the level to make sure the top of the plates are level. 12. Make a mark 2.5-3 cm below the notch in the front plate. Pouring Separating Gel 1. Recipes for gels with different % acrylamide are listed in MATERIALS section. Add the reagents in the order indicated. 2. Pipet the separating gel mix between the plates up to the mark. Touching the pipet to the back plate will ensure smooth pouring and avoid bubbles. 2. Using a P-1000, gently overlay the acrylamide mix with 0.5-1 ml of isopropanol. 3. Let gel polymerize for about an hour. 4. You will need the APS again for the stacking gel, so put it in the fridge (good for ~1 week). Pouring Stacking Gel 1. Get the APS and acrylamide from fridge. 2. Find the appropriate comb to make the wells for the gel and rinse it off. Make sure it will fit between the spacers on your gel. 3. Pour off the isopropanol from the top of the gel. (blot dry with Whatman paper) 4. Rinse the top of the gel with distilled water from squirt bottle and pour this off. 5. Tilt gel onto one corner and let it any remaining fluid drain to one side while you make the stacking gel mixture. 6. Add first three reagents of stacking gel mixture in a small glass flask. 7. Use a piece of 3MM paper to remove the last bit of liquid from the top of the gel. 8. Add the APS and the TEMED to the stacking gel mix and pipet the stacking gel mix between the gel plates to within a few mm of the top of the notch. 9. Insert the spacer into the stacking gel mix, trying not to get any bubbles beneath the teeth. It is ok if extra acrylamide spills over the edge. Do this on a paper towel. Work quickly because the stacking gel will polymerize within a few minutes. S.L. Christian September 2008 edited: Feb 2011 page. 3

10. If necessary, pipet a bit of stacking gel mix around the sides of the spacers to bring the level back up to the top of the notch. 3. LOAD SAMPLES, RUN GEL, TRANSFER TO NITROCELLUOSE MATERIALS 10X Running Buffer 1X Running Buffer 60.6 g Tris 200 ml 10X Running Buffer 288 g Glycine 1800 ml dh 2 O 20 g SDS Adjust to 2 L with dh 2 O Transfer Buffer (~1L/transfer box) 2.9 g Glycine 5.8 g Tris base add water to 800 ml, stir until dissolved 200 ml Methanol PROCEDURE 1. Put gel into apparatus for running gels 2. Pour 1X running buffer into upper and lower chambers 3. Carefully remove comb, running buffer will flow into wells and rinse out unpolymerized acrylamide 4. If running only one gel, clamp spacer plate on the other side. 5. Load the samples using gel loading tips. Place the tip close to the bottom of the well. The sample is denser than the running buffer and will sink to the bottom of the well. Be careful not to expel a big air bubble at the end and force sample out of the well. 6. load equal volume of 1X sample buffer in all empty lanes 7. When all samples are loaded, place the cover on the gel box matching color of electrodes. 8. Turn on the power supply. Gel Running Conditions Run gel at 100V until dye front is near the bottom (1 2h). (Optional: For overnight, run at 20-30 V. In the morning, turn up the power to 100 V for a while to refocus the bands.) 1. Turn off the power supply. 2. Place the gel box in a pan or sink. 3. Undo the clamps and remove the gel plates. 4. Using a spacer, pry apart the plates. The gel should stick to one plate or the other. 5. If you are going to do a transfer, you should have already set up the transfer cassette. 6. Turn the plate upside down over the transfer cassette, prod the gel with the spacer and let it fall into the buffer. Mini-gel transfer to nitrocellulose 1. While gel is running: a) Make transfer buffer. b) Get pan for assembling sandwich and rinse with distilled water. S.L. Christian September 2008 edited: Feb 2011 page. 4

c) Get transfer box, cassettes, pads and rinse with distilled water. d) Get filter forceps and rinse. e) Get 4 pieces of 3MM filter paper per gel (6 x 9 cm) f) Get 1 nitrocellulose membrane per gel (6 x 8 cm). Equilibrate in transfer buffer for 30 minutes before transfer. g) Fill pan with transfer buffer. h) Place cassette in pan with black side down. i) Soak pad in transfer buffer and place on cassette. 2. Assemble sandwich-wear gloves! a) Soak 2 pieces 3MM paper and place on pad. b) Place gel on 3MM paper. Gel should be face down so that blot comes out with same orientation as loaded. Smooth out with fingers. c) Label upper right hand side of nitrocellulose with pencil to maintain orientation. Use forceps to place nitrocellulose on gel. Run pipette over it to get rid of bubbles. d) Place 2 pieces of 3MM on nitrocellulose. Smooth out with pipette. e) Soak pad and place on top. f) Close cassette and put in transfer box with black side of cassette facing the black side of the transfer box insert. g) Put white plastic insert with ice into transfer box. h) Fill transfer box with transfer buffer. 3. Run at 70 volts for 75 min. Starting current = 0.19A for two gels. OR Run at 100 volts for 1h with stirring (optional: To transfer overnight, run at 20-30 V. Omit ice insert) B. IMMUNOBLOT MATERIALS 10X TBS (Tris buffered saline) 175.3 g NaCl 48.5 g Tris base Adjust volume to 2L ph to 7.5 (add approx. 29ml conc. HCl) TBST (0.05) 2L 1L 10X TBS 200 ml 100 ml water 1800 ml 900 ml 10% Tween-20 10 ml 5 ml PROCEDURE 1. Remove nitrocellulose membrane from cassette (discard gel and filter papers) 2. Block with 20 ml 5% milk/tbst or 10 ml 5% BSA/TBS, as appropriate for antibody used, for 1 h at RT with rocking 3. Add primary antibody ON at 4 C with rocking 4. Wash 15-30 min, 3 changes, with TBST S.L. Christian September 2008 edited: Feb 2011 page. 5

5. Add appropriate secondary antibody (HRP conjugate) 1hr at RT with shaking (1:5000 in TBST) 6. Wash 15-30 min, 3 changes, with TBST 7. Prepare Chemiluminescent Substrate, mix 2 ml Reagent A plus 2 ml Reagent B 8. Incubate membrane 5 min at RT 9. blot corners of membrane on filter paper 10. wrap in SaranWrap, tape in autorad cassette 11. expose to camera and take picture Strip & Reprobe (optional if you want to reprobe the same membrane with a different Ab) 1. Rinse membrane with TBS 2. wash 3 x 10 min with TBS ph 2.0 (100 ml 10XTBS, 900 ml dh 2 O, 1.86ml conc HCl) 3. rinse with TBS 4. block 1h with appropriate blocking buffer 5. Add primary antibody ON at 4 C with rocking 6. follow step 5-12 above S.L. Christian September 2008 edited: Feb 2011 page. 6

APPENDIX A From the BioRad Catalogue S.L. Christian September 2008 edited: Feb 2011 page. 7

APPENDIX B 1% Triton X-100 lysis buffer- Store at 4 C Reagent Final concentration For 500 ml, use Water up to 500 ml 1 M TrisHCl, ph 8 20 mm 10 ml 10% Tx-100 1% 50 ml glycerol 10% 100 ml of 50% (w/v) in water EDTA 2 mm 2 ml of 0.5 M, ph 8.0 NaCl 137 mm 13.7 ml of 5M RIPA-Tris based Final Stock Volume 50 mm Tris-HCl (ph 7.6) 1M 25 ml 5 ml 0.02% sodium azide 10% 1 ml 200 µl 0.5% sodium deoxycholate 10% 25 ml 5ml 0.1% SDS 10% 5 ml 1 ml 1% NP-40 10% 50 ml 10 ml 150 mm NaCl 5 M 15 ml 3 ml H20 up to 500 ml up to 100 ml RIPA- Phosphate based Final Stock Volume 1X PBS 10X 50 ml 10 ml 0.5% sodium deoxycholate 2.5g 0.5g 0.1% SDS 10% 50 ml 10 ml 1% NP-40 10% 50 ml 10 ml H20 up to 500 ml up to 100 ml 5X SDS-PAGE Sample Buffer with 1X Glycerol (20 ml) 312.5 mm Tris base 0.757 g or 6.25 1 M Tris, ph 6.8 10% Glycerol 2 g 11.5% SDS 2.3 g 500 mm DTT 1.54 g (Omit for Non-Reducing Gels) ph to 6.8 Adjust volume to 20 ml 0.1% Bromophenol Blue 20 mg (0.020 g) (Add last after phing) Make 1 ml aliquots and store at 20 C S.L. Christian September 2008 edited: Feb 2011 page. 8

APPENDIX C BCA protein assay From Pierce, includes 2 mg/ml BSA standard 1. Set up 1.5 ml µfuge tubes with standards or samples in final volume of 50 ul. Standards should receive same amount of lysis buffer as sample tubes. Example: Tube # µl Water µl Lysis buffer µl 2 mg/ml BSA in µg BSA in µg/µl BSA water assay 1 40 10 0 0 0 2 37.5 10 2.5 5 0.5 3 35 10 5 10 1 4 32.5 10 7.5 15 1.5 5 30 10 10 20 2 6 27.5 10 12.5 25 2.5 Samples 40 10 µl sample 2. All standards should get same volume of identical lysis buffer used for samples. If assaying a different sample volume, adjust amount of lysis buffer added to standards and increase or decrease volume of water accordingly. 3. Mix Pierce BCA reagent: 50 parts soln A + 1 part soln B. 4. Add 1 ml to each sample. Vortex. 5. Standard protocol: 30 minutes in 37 o water bath. Enhanced protocol: 30 min in 60 C water bath 6. Vortex, cool 5 minutes. 7. Read A562 vs tube #1 as blank. Read all samples within 10 minutes. Use quartz cuvettes. For standards, just empty cuvette between each as you go in ascending order. Then wash out with water and do samples. 8. To calculate protein concentrations of samples, plot ug/ul BSA std (Y axis) vs OD562 (X axis). Do curve fit and get linear equation. Y = µg protein. Take ODs of samples (X) and plug into equation. S.L. Christian September 2008 edited: Feb 2011 page. 9