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Western Protocol (see bottom for native gels)

Day 1 (~3hrs w/o block, 4hrs w/)

Transfer and blocking

1. Bring sample in PCR tubes 2. Put blue Novex “Tricine SDS sample buffer 2X” mixed with 5% Beta mercaptoethanol (BME; often found in 455C hood or under printer hood). Novex buffer found on shelf above western bench or Dahabada’s shelf. a. Dilute to 1X ex: ~30 μl: 20 μl sample, 10 μl Novex/BME b. To make 5% BME in novex sample buffer: 475ul novex+25ulBME 3. Fill heat block in hood with water and boil tubes for 5 min between 90-100 deg C. a. If heat too long, all liquid will evap but can maybe use this step to concentrate a sample that might not fill in the wells. 4. Grab Invitrogen 10-20% tricine gel (1mm) from red box in left fridge drawer. 5. Remove white strip from bottom of gel 6. Make solution of 50ml of Novex running buffer (from 10x stock) and 450ml water in 500ml graduated cylinder, cover with parafilm, and shake to mix. a. Measure carefully! Important b. Can reuse once 7. Take out comb from gel. Fill wells with 1000ul pipette running buffer to wash them out. Dump running buffer in sink. Repeat 2 more times for 3 times total. 8. Fill wells one more time with running buffer. 9. Set up tank: 4 pieces (tank, gel place holder, conducting wires, sealer) a. Wells of gel go towards inside of tank 10. Once tank is set up, lock it and pour running buffer in to fill middle part of tank (make sure it’s not leaking to the outer parts). Load wells with sample, then fill outer parts. Pour the rest of the running buffer on the outer parts of tank. 11. Load gel with samples and at least 1 standard/ prestained marker ladder (8 μl of see blue plus for 10-20% tricine). a. Use special, thin pipette tips. b. Don’t use all 10 lanes until experienced because outer lanes might tear or smile. Leave at least one of the outer lanes open if you can. c. Make a note/diagram of which sample was loaded in each lane d. If doing 2 gels, load them asymmetrically so you know which is which 12. Plug in wires and push DC start. Use 100V for 10min and 130 for rest of the time. Make sure you see bubbles/conduction. Will take between 1 and 1.5 hrs. 13. In the meantime, wash out the fiber pads with miliQ water over and over to make sure no left over SDS in them. Wash the next tank with soap and miliQ water. Make sure soap is completely washed away. 14. Put out 2 pyrex pans and pour western transfer buffer/running buffer in one of them. a. Transfer buffer recipe 10x: b. 14.54g tris base (12mM) c. 72.07g glycine (96mM) d. to make 1x: need to add MeOH (20% of final volume) and ph 1X to 8.3 15. For 1 gel: Put 1 nitrocellulose membranes and 2 filter papers (all come together in a sandwich) in transfer buffer, and 2 fiber pads. Only use flat/smooth forceps when handling nitrocellulose membranes. Double if have 2 gels. a. Invitrogen LC2001 .45um pore size, 20/pack 16. Stop gel when sample buffer (pink line) has moved into gel foot (very bottom notch). Turn power off, take lid off tank, take gel out. 17. Use Japanese scraper knife to break shell of gel. Leave bigger piece of shell on table. Cut off lanes (stacking) and pink line (foot) and throw away. 18. Dip knife in the running buffer in the tank and place under gel and flip gel into transfer buffer tray. If have 2 gels, can clip corner of one to distinguish them. 19. Assemble the “sandwich” as follows being careful to not get air bubbles between layers and to NOT touch the membrane with gloves (only forecpes): a. Place cassette with the black side down in a baking pan type tray. If the cassette is laid flat, it will break the hinges so this way, it can be opened only 90deg. b. Fiber pad c. sheet of filter paper d. gel: bottom (where colored bands were) towards hinges e. nitrocellulose membrane f. sheet of filter paper g. push (don’t roll) out air bubbles side to side and up and down with empty 15ml conical tube. h. Fiber pad (everything should be as wet as possible) 20. Carefully close cassette without moving anything, and close lock all the way. 21. Repeat for second cassette but tank may be run with only 1 cassette. 22. Place the insert/cassette holder in the tank so that the insert slides in more towards the back, leaving a larger space in the front (can check by placing lid on). 23. Drop in small stir bar. 24. Place cassette(s) with black side of each facing the black half of the insert. 25. Pour whatever transfer buffer was used to equilibrate gels and papers into the tank. 26. Take the tank, a small stir bar, and the transfer buffer to the cold room (418B) and set up on magnetic stir plate. On your way there, stop at the end of the hall and fill the white ice holder for the transfer tank with ice. 27. Set stirring to as high as possible, making sure that the bar has clearance in the tank. Fill completely with transfer buffer. 28. Put lid on tank and wires into box on shelf. It should say pause, so push “run” and you want it at constant .4amps and 200V but voltage may fluctuate. a. 200V for 50min b. 250V for 45min c. 80V for 1hr 29. After 50min, press pause and open cassette and use tweezers to pick up layers to check if transfer is complete. Transfer is complete when all color has gone from gel to nitrocellulose membrane. If transfer is NOT complete, can NOT re-run. 30. Disassemble cassette and discard everything but the membrane (transfer buffer with MeOH needs to be wasted or be kept for future use). Leave cassette, fiber pads, insert and tank to soak overnight in hot water with a splash of 5% SDS. Be sure to rinse very well the following day. 31. Get a tip lid for each gel. Put 1x PBS (tablets sigma P4417-100TAB) in it and do two quick shake and washes. Use tweezers to transfer the membrane. Microwave membrane in the tip lid (petri dish might melt) in ~50ml PBS for ~45sec at 50% power on each side (press: time, 45, power, 5), allowing membrane to sit in hot PBS for 4 min between and after flipping. Don’t put lid on box in microwave. Then dump the PBS. No need to let it sit. 32. Make blocking solution: 2.5g powdered milk (in fridge, use block from ECL kit) into 50ml of TBST (5% milk). Shake vigorously and place on shaker to ensure components are fully dissolved. Microwave to make sure Fully dissolved otherwise might get specks on membrane upon development. a. I used 2% block: ECL block 2g/100ml PBST or TBST 33. Block for 1hr at room temp or overnight at 4degC in 20ml milk solution. If blocking overnight, keep extra milk in cold room too. Place the bright side of the membrane up (maybe down, heard both) and mark with pencil. 34. Check with Spencer lab to see if can use their detector machine tomorrow. End of Day 1

Day 2 (~5hrs in a hurry)

Incubation and detection

35. Rinse membrane briefly 2X with TBST; no need to wash 36. Dilute antibody (6E10) in 20ml blocking solution (1 : 10,000) by putting 1ul of antibody per 10ml of milk. Incubate 1hr at room temp or overnight in 4degC. Primary antibodies are in freezer in orange tube tray. a. If using A11 polyclonal antibody: dilute 1:1000 or 10ul in 10ml. b. Can reuse primary twice with 1wk rest between uses c. asyn antibodies: mab211 1:333, pabFL140 1:333 with 1:5000 secondary d. actin 1:2000 with 1:5000 secondary e. 10ml of solution will cover membrane if using a lot of antibody 37. sign up for the spencer lab alpha imager on google calendar 38. after primary rinse membrane briefly 3X with TBST 39. Wash 3 times for 7min each in TBST, with 2 quick rinse and dumps between each wash 40. Wash 1 time for 7 min in TBS 41. Dilute secondary Ab in blocking solution 1:10,000 or 1ul in 10ml and incubate for 1hr at room temp. Use anti-mouse for 6E10 in fridge in cardboard box. a. If using A11: dilute 1:10,000 or 1ul in 10ml and use anti rabbit. b. HRP 1:10,000 in PBST or TBST not block 42. Take out orange box Amersham ECL advance western blotting detection kit to reach room temp. 43. After secondary wash 3 times for 7min each in TBST. With 2x quick washes in between. 44. Wash 1 time for 7 min in TBS. Then empty TBS. 45. Mix detection solution: 1ml of A with 1ml of B. 46. Ask the spencer lab to log you in to the alpha imager. 47. Coat bright side of membrane 1ml at a time with a 1000ul pipette, using the whole 2ml’s. Develop right away. a. With 2 gels, develop 1st gel before adding detection solution to 2nd gel. 48. Don’t touch computer with gloves, Open program on desktop called “FlourChem” and push acquire button at top. 49. You should now see the 3 colored buttons (focus, expose, and acquire) at top. Put shelf in and put world map thing on shelf and focus on lines with knob on top of machine. a. Close aperture meaning put it on 1.8 50. Put membrane on shelf centered with largest bands of ladder towards back a. Need to keep ECL solution on membrane 51. Close door of machine and follow directions for chemiluminescence #1 taped to shelf. Make sure transillumination and reflective buttons are not clicked. a. Can check in “Display” box on right • Chemi Display • Turbo • Auto expose 52. click “super speed”, click Preview. You should see yellow line then blue then green go across screen. Y → B → G (green means it is ready, pink means it can’t see it). Once the green shows up, click Ultra and click Acquire. a. Turbo/super speed checked = lower resolution. b. Auto expose often picks times that are longer than needed. Can turn off and pick your own time. Don’t expose for more than 4min, all ECL reagent will get used up and won’t get a good picture 53. To take a picture of see blue, leave at chemifilter/#1, turn on reflective white, open aperture to 8. Turn off chemi display, leave on ultra and hit acquire. a. If see larger than 30min expose time, unclick auto expose and change exposure time to 10min. change speed to medium high and click acquire image. If nothing after 10min, then you have nothing on your membrane that we can see. b. If you get a lot of dark bands, can cover that part of the gel with a paper towel and re-read to get the lighter bands to be more exposed. c. 5min at normal speed/ultra resolution is different than 5min at med/high d. Hit focus to cancel preview then can hit preview again 54. To save: file, save, users, Bitan lab, make own folder a. Must save once with “save as” or “save all” ant this saves as black background, white bands b. Then go back and save modified grayscale and this will save any changes you’ve made to the contrast or if you reversed the image. 55. To get large blow up of membrane: edit, edit activation, crop membrane, edit, crop 56. Can also print on tiny printer next to computer by pushing print button in top right. 57. If want bands to be black and background to be white, go to “contrast adjustment” box and hit reverse.

Stripping buffer

Light stripping: Low pH: 9.38g Glycine (250mM), 5g SDS (final 1%), 500ml water, ph 2.0 Harsher stripping: SDS/BME/heat (for 100ml): 0.76g tris base (final 50mM or seen recipes with 62.5mM), 2g SDS (final 2%), 700ul B-merccaptoethanol (final 100mM)→water to 100ml. ph to 6.8 1. Heat stripping buffer to 50c in water bath 2. Incubate blot with stripping buffer for 30min to 2hrs with gentle shaking 3. Rinse blot several times with TBS 4. Blot is now ready for reblocking.

Native gels

Native gels are just like SDS gels BUT: 1. Use a Native sample buffer and do NOT denature with heat 2. Use 10-20% tris-glycine mini gels from Invitrogen (I’ve seen people use the tricine gel with the tris/glycine running buffer and it seemed fine). 3. Use tris/glycine (no SDS) running buffer: it will run much more slowly

10X Tris/Glycine Running Buffer, [concentration of 1X is 25 mM Tris, 192 mM glycine, pH 8.3] 10X: 29 g Tris Base + 144 g Glycine → water to 1 L, ph to 8.3 keep at room temp, good for 1 yr.

Native Sample Buffer, [62.5 mM Tris-HCl, pH 6.8, 40% glycerol, 0.01% w/v bromophenol blue]

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