Western blotting

Western blotting

This protocol is designed for empty Invitrogen Novex gel cassettes and Novex blotting unit

Pouring and running the gel

Stand the empty gel cassette on a paper towel somewhere it can’t easily fall over (for example, leaning against the side of a plastic box)

Using the ratios given in Sambrook and Russell appendix 8, mix water, Tris, acrylamide and SDS to make 10ml resolving gel per gel cassette. *

*for low cell number, when working the aging time-course, use a 15% acrylamide (30% solution, 37.5:1, Biorad) gel

To a 10ml aliquot of resolving gel, add the necessary amount of 10% APS¹, invert to mix, then add TEMED¹ and mix by inversion again

Pour ~8ml into the gel cassette – it should reach the second plastic ridge

Layer 500ml water on top of the gel – add this down the side of the cassette slowly

Once the gel has set (about 30min), draw off the water layer with a tissue 

To 2.5ml of 5% stacking gel, add 25µl 10% APS, invert to mix, then 2.5µl TEMED, invert to mix again

Pipette the stacking gel into the top of the gel cassette (being careful to avoid bubbles), and carefully insert the comb (some acrylamide will dribble out, catch this with paper towel)

Once set (10min), remove the comb and wash the gel and comb under running water

Denature protein samples in loading dye for 3 min at 95° or 10 min at 65° (if using high urea loading dye), and defrost BioRad ladder

Meanwhile, mount the gel in the tank and pour in 1x Tris-glycine running buffer (~700ml). Make sure the buffer isn’t leaking from upper to lower chamber

Cool the protein samples to room temperature², spin briefly and load into wells using a Biorad extended tip. Load 5µl ladder.

Run at 75V until samples reach the resolving gel then increase to 175V

As the ladder is prestained, it is relatively easy to judge how far to run the gel
 

Blotting*

Make 500ml transfer buffer and soak the blotting pads in ~400ml

Cut 1 sheet of PVDF³ membrane per gel to 8x7cm and wet for 15s in methanol, then soak for 5min in transfer buffer. Cut two sheets of Whatman 3MM per gel to same size, wet these with transfer buffer just before use

Open the gel cassette, cut off the wells and the foot of the gel, rinse briefly with deionised water to remove residual SDS

Put 1 sheet of wetted 3MM paper on the gel, then use the gel knife to remove from cassette onto parafilm so the gel is face up.

Lay the membrane carefully on the gel, followed by the other sheet of wetted 3MM paper. Roll over a pipette to remove bubbles

Repeat the above steps for a second gel if being used.

Put two pre-soaked blotting pads in the bottom of the electrode, then the first gel sandwich, with the MEMBRANE SIDE UP. 

If transferring two gels, add another soaked pad and the second membrane sandwich also membrane side up.

Add enough soaked blotting pads to be ~5mm above the top of the electrode.

Hold lower electrode above tank, apply upper electrode, squeeze together and push into tank. Seal in place with holder – the electrode assembly should be leak proof.

Pour in enough buffer to just cover the pads, then fill the outer chamber with distilled water.

Put lid on and transfer at 25V for 1-2hrs (current should be ~100mA)

Remove membrane, rinse 3x with water (don’t skip this step)

(Not for fluorescent detection) Soak membrane in Ponceau S solution for ~1 minute

Meanwhile, wash the pads, squeeze out and leave to dry

(Not for fluorescent detection) Pour Ponceau S back in bottle and wash membrane with water until the water no longer turns pink. 

If desired, store membrane at 4° in TBS for a few days before probing

Soak the gel overnight in Coomassie stain to visualise residual protein – use designated staining tray (will be blue!). Next day dispose of stain in the sink and wash gel repeatedly with water to destain. For fluorescent western, be very careful that no trace of Coomassie is transferred to the membrane (use different gloves!)

Probing the blot for fluorescent detection

Notes:

The membrane must not be exposed to Tween until AFTER the blocking step

Do not touch the membrane in the vicinity of the bands

Use designated plasticware for the antibody incubations
Some antibodies, eg from CellSignal have optimised conditions for each antibody – follow them, but keep the volumes as in this protocol.

Do all tray incubations on a shaker and tube incubations on rollers

Two antibodies can be utilised simultaneously with this protocol, provided they are from different species. Use appropriate secondaries to detect in the 700nm (CST 5366) and 800nm (CST5257) regions since the Li-cor doesn’t detect the ones normally used for IF.

Don’t stain the membrane with Ponceau

Be careful with Coomassie as the slightest trace on the membrane shows up in the fluorescent detection

Block membrane with 5% milk in TBS at room temperature for one hour or overnight at 4°

Replace blocking solution with 5% milk in TBST containing primary antibody(s) I normally do this in a 50ml Falcon tube on rollers, in which case 3ml of solution is required. Incubate for 1 hour at room temperature or overnight at 4°. NB: Some antibodies, particularly from NEB, should be used in 5% BSA in TBST. If doing this, wash the membrane 3x 5min with TBST before applying the primary in 3ml in a Falcon tube as above

Rinse the membrane with TBST then wash three times with TBST for five minutes.

If doing multiple membranes with different antibodies, wash them separately, but they can have the secondary antibody applied together if convenient.

Apply the secondary antibody (or antibodies) in TBST with 5% milk for 1hr at room temperature. Our NEB secondaries are used at 1:15,000. Cover the plastic box with aluminium foil during this step.

Wash membrane three times with TBST for five minutes. Wash once for 5 minutes with TBS (no TWEEN) to remove residual TWEEN. Cover the plastic box with aluminium foil during these washes.

Scan the membrane on the new Licor CLX. You can scan it wet (pass the roller over it) or dry (dry is more sensitive)-it has to be fully dried by letting it sit in the dark on 3MM paper at RT for a while or by placing it in a 37° incubator for 20min on 3MM paper covered in foil.

High sensitivity protocol*

As fluorescent protocol, but:

Block with 10ml Odyssey Blocking Buffer (Licor)

Use primary antibody in 3ml Odyssey Blocking Buffer (Licor) + 0.1% Tween 20

Use PBS-T for washes (PBS+0.1% Tween-20)

Use secondary antibody 1:15,000 in 10ml Odyssey Blocking Buffer (Licor) + 0.1% Tween 20

*use High sensitivity protocol when working with low cell number. 

Probing the blot for chemiluminescent detection

Notes

Some antibodies, eg from CellSignal have optimised conditions for each antibody – follow them, but keep the volumes as in this protocol.

Do all tray incubations on a shaker and tube incubations on rollers

Block membrane with 5% milk in TBST at room temperature for one hour or overnight at 4°

Replace blocking solution with more 5% milk in TBST containing primary antibody. I normally do this in a 50ml Falcon tube on rollers, in which case 3ml of solution is required. Incubate for 1 hour at room temperature.

Rinse the membrane with TBST + 5% milk then wash four times with TBST + 5% milk for five minutes.

If doing multiple membranes with different antibodies, wash them separately, but they can have the secondary antibody applied together if convenient.

Apply the secondary antibody in TBST with 5% milk for 1hr at room temperature

Wash membrane four times with TBST for five minutes. Wash once for 5 minutes with TBS (no TWEEN) to remove residual TWEEN.

Pat the membrane dry, and apply 3ml SuperSignal West Pico to the membrane

Leave for 5 min then dry on tissue paper and wrap in Saran

Image using a gel doc system or film

Stripping the membrane

450ml β- Me

5ml SDS (20%)

3.125ml 1M Tris (pH 6.8)

41.425 ml H₂O

50ml falcon tube

Soak the membrane at 50^oC for 30 mins, then wash 3 x in 1x TBST. Block the membrane as usual. 

Staining the membrane

Membranes can be stained after blotting, but can no longer be used for fluorescent detection:

Amido black:  Stain 15-30min with amido black solution, destain 5-10 min repeatedly with amido black destain

Ponceau:         Stain for a few minutes with Ponceau solution then rinse repeatedly with water. 

Solutions

2x Protein loading dye:    

100mM Tris-Cl pH6.8

4% SDS

0.2% Bromophenol Blue

20% glycerol

200mM DTT (add fresh before use)

Urea Sample buffer:   10ml :

70mM Tris pH6.8      700µl of 1M

8M urea         4.8g

5% SDS       2.5ml 20% (Biorad stuff)

1mM EDTA        20µl of 0.5M

water to 9ml, warm gently to dissolve

100mM DTT              Add from 1M just before use

0.01% Orange G

10x Tris-glycine running buffer: 

30.2g Tris base

188g glycine

10g SDS

dH₂O to 1L

10x Transfer buffer: 

7.28g Tris base

36g glycine

Water to 500ml

Dilute to 1x in 20% methanol final just before use

NB: Currently 10x TBS is made by media service, so just dilute and add TWEEN.

10x TBS: For 500ml:

15g Tris HCl

40g NaCl

1g KCl

Add dH₂O to 400ml

Add 10ml conc. HCl

Check pH, adjust if needed to pH7.4

Add dH­₂O to 500ml

TBST: 

1x TBS

0.1% Tween-20

Coomassie stain: 

Mix 200ml ethanol with ~700ml water in 1L bottle

Add 18.8ml phosphoric acid (85%) 

Add 80g ammonium sulphate

Let this dissolve with occasional mixing (takes a few minutes)

Meanwhile, dissolve 0.8g coomassie Brilliant Blue G250 in 50ml water in a 50ml Falcon tube. Ensure this is fully dissolved (shake well – can you see residual powder when you invert the tube?)

Add the coomassie solution to the bottle and add water to 1L

Ponceau S:   

0.1% Ponceau S

5% acetic acid

Amido black:

0.1% napthol blue-black

10% methanol

2% acetic acid

Amido black destain: 

50% methanol

7% acetic acid

[1] Amounts of APS and TEMED vary with gel percentage, see Sambrook and Russell Appendix 8 

[2] Samples with high concentrations of urea should not be chilled on ice as they tend to freeze.

[3] We use Millipore PVDF membrane certified for fluorescence for fluorescent westerns

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