Western Blots

1. Run an SDS gel as usual. (It's nice to run a lane of prestained markers, which will transfer on to the blot and control for the transfer).

2. Transfer: make 4 liters of transfer buffer. Cut 3 pieces Whatman and one piece of nitrocellulose to the size of the gel. Pry the gel plates apart, cut the stacker off with a razor blade, then pick up the gel by pressing a dry Whatman sheet on top and lifting up carefully. Put the gel down, paper side down on a glass plate. Wet the gel and paper by squirting water underneath the paper and on top of the gel. Look carefully for bubbles between the gel and paper, and with a gloved finger carefully smooth these off the edge (make sure the gel and your finger are very wet or you will tear the gel doing this). Wet the nitrocellulose in transfer buffer and lay it on top of the gel, again smoothing out bubbles. Wet the remaining two Whatmanns and lay on top of the sandwich. As a final bubble removal, use a 5 ml pipette like a rolling pin on the sandwich.

To transfer in a Biorad transfer cell: Fill the box ~3/4 with transfer buffer. Wet the two brillo pad-like stuffers with transfer buffer. Place one wet stuffer on the black half of the plastic holder, then pick up the glass plate carrying your sandwich, place it next to the holder, and slide the sandwich off the plate onto the brillo pad without bending the sandwich. Place the second wet brillo on top, close the holder, and place it in the transfer cell with the black side of the holder facing the black electrode (protein moves towards the red electrode). Fill up the cell the rest of the way with transfer buffer. Place a small stirbar in the cell. It's best to cool the cell during transfer, although not absolutely necessary, so place a cooling coil hooked up to running tap water in the cell. (At high amps without cooling, long transfers can melt the apparatus). Place the lid on the cell, making sure the black and red leads on the lid match the red and black sides of the cell. Place the apparatus on a stir plate and set stirring, and turn on power. Transfer is at ~.33 amps (40-80 volts depending on the type of apparatus) for at least one hour. Overnight is okay, although you may want to turn down the power.

Some comments on transfer: some proteins behave anomolously in transfer. Large proteins or basic proteins might require longer transfer. Some proteins can go all the way through the nitrocellulose if you transfer too long. The electrical contacts on the Biorad apparatus are notoriously bad, so make sure current is running after you turn on the volts. You can clean the contacts with a wet Q-tip. The main problem encountered by first time blotters is air bubbles in the sandwich, which give big splotches of no transfer on the blot. These go away with careful technique and experience. Semi-dry transfer apparati have given uneven results for people I know.

3. Ponceau staining the blot. After transfer, take the sandwich apart and remove the nitrocellulose blot. Ponceau is reversable protein dye; this staining allows you to check the transfer and see any bubbles that might have been in the sandwich, and to check the loading of the gel. In a pipette tip box lid on a rotator, stain the blot in ponceau for 20 min. The ponceau solution is reusable, so pour it back in the bottle after use. Destain by rinsing the blot in distilled water, until the protein bands show up well. The marker lanes will fade, so make them permanent by marking over the bands with a colored pencil (Colerase brand is good). To get a permanent record of the gel, Xerox the ponceau stained blot.

4. Blocking. To remove the ponceau and block the blot, soak in blotto for >30 sec. A note: some epitopes or antibodies are sensitive to milk. An alternative to blotto is 2% BSA in TBS/Tween; people block with this for 30min to 1 hr.

5. Primary incubation. Make a solution of 1℃ antibody in blotto (or BSA/TBS/Tween). Need only ~5 mls or less for a modest size blot (14X8 cm). Antibody concentration should be optimized; a general guidline is ~1:2000 dilution for a red-hot polyclonal, or 1:30 dilution of tissue culture supernatant for a good monoclonal. Do the incubation in a sealameal bag for 1 hr at room temp on a rocker. Can go overnight in the fridge. Remember: it's critical to do a control with no 1° antibody in order to determine if your bands are due to cross reactivity with the secondary antibody.

6. Wash 3X3min in TBS/Tween in a pipette tip box lid. Block again 30 sec in blotto.

7. 2° incubation. In a sealameal bag incubate blot in 2° antibody. For alkaline phosphatase, use a 1:3000 dilution of Biorad goat anti-mouse or anti-rabbit alkaline phosphatase conjugated antibody in blotto. Incubate on rocker 1 hr at 25℃.

8. Wash 3X3min in TBS/Tween.

9. Alkaline phosphatase development: Add developing solution and incubate on rotator in box lid until bands come up. Reagents are light sensitive, so it's best to cover with aluminum foil while developing. It goes faster at 37°. Can develop up to several hours, although it's mostly done in an hour. The background fades when the blot dries, so don't be afraid to let the background go up. Rinse in water and let dry between Whatmann sheets. Bands fade in the light, so keep it in your notebook.

10. Chemiluminescence detection: this method is much more (10X?) sensitive than alkaline phosphatase. Use a 1:3000 dilution of Biorad goat anti-mouse or goat anti-rabbit horseradish peroxidase conjugated secondary antibody. After washing the blot, pour off all the wash buffer and add a 1:1 mixture of Amersham "ECL" detection reagents, just enough liquid to cover the blot. Mix for 1 minute, then working quickly, dab the blot against Whatmann paper to remove excess liquid while leaving the blot moist, wrap the blot in saran wrap, and expose serially to film in a cassette for 30 seconds, 1.5 min, and 5 minutes. I find that regular S&S nitrocellulose gives a terrible background on 5 min. exposures with this detection method, but that S&S "supported nitrocellulose" is much better.

Solutions:

Transfer buffer Ponceau stain (reuse)

57.6 g glycine 0.2% ponceau S in 3% TCA

12.0 g tris

800 ml MeOH

H20 to 4 liters

TBS/Tween Blotto

20 mM Tris pH 7.4 5% nonfat dry milk in TBS

500 mM NaCl (keeps if you add 5mM sodium azide)

0.05% Tween-20

Alk. Phos. developer

47.5 ml 0.15M Tris pH 9.6

200 µl 1M MgCl2

500 µl 5 mg/ml BCIP in dimethyl formamide (BCIP= 5-bromo-4- chloro-3-indoyl phosphate, toluidine salt), US Biochemical). Store this stock at -20° wrapped in foil; it's light sensitive.

2.5 ml 2 mg/ml NBT (p-nitro blue tetrazolium chloride). Store at -20°.

Alternative Development methods

Alkaline phosphatase is a good, sensitive, general purpose detection method. 125I detection is good for quantitation, but isn't any more sensitive, and results can be slow due to long exposure times. The HRP conjugated secondary followed by DAB detection method offers no advantages; it's not very sensitive. Chemiluminescence can be substantially more sensitive than alk. phos. There are a couple of chemistries available for this, and it is probably the method of choice where extreme sensitivity is required. As is the case with alk. phos., the detection isn't linear, so it's not good for quantitation. Remember, if you are using HRP conjugated secondaries to do chemiluminescence, HRP is inhibited by azide, so you don't want to use blotto with azide in it.

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