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SDS-PAGE gel staining

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1. Silver Staining of SDS-PAGE Gels

Reference

[bibliplug last_name=’Blum’ keywords=’gel staining’]

Solutions required

These volumes are sufficient for one big SDS-PAGE gel from the Bio-Rad system. Halve the volumes for mini-gels.

Currently, the lab only has 10% formaldehyde and not 37%. Therefore, you will require 3.7 times more formaldehyde than what is recommended below.

  • Fix solution [50% (v/v) methanol, 12% (v/v) acetic acid, 0.05% (v/v) formaldehyde]. Methanol (100 ml), acetic acid (24 ml) and formaldehyde [100 μl of a 37% (v/v) solution] were diluted to 200 ml with dd.H2O.
  • Wash solution I [50% (v/v) ethanol]. Ethanol (100 ml) was diluted to 200 ml with dd.H2O.
  • Pretreatment solution [0.02% (w/v) sodium thiosulfate]. Sodium thiosulfate (Na2S2O3.5H2O) (40 mg) was dissolved in dd.H2O.
  • Impregnating solution [0.2% (w/v) silver nitrate, 0.075% (v/v) formaldehyde]. Silver nitrate (AgNO3) (400 mg) was dissolved in dd.H2O (199.85 ml) and formaldehyde [150 μl of a 37% (v/v) solution] was added just before use.
  • Develop solution [6% (w/v) sodium carbonate, 0.0004% (w/v) sodium thiosulfate, 0.05% (v/v) formaldehyde].   Sodium carbonate (Na2CO3) (12 g) and sodium thiosulfate (Na2S2O3.5H2O) (4 ml of pre-treatment solution) were mixed together in 199 ml dd.H2O. Formaldehyde [100 μl of a 37% (v/v) solution] was added just before use.
  • Stop solution [50% (v/v) methanol, 12% (v/v) acetic acid]. Methanol (50 ml) and acetic acid (12 ml) were mixed together and made up to 100 ml with dd.H2O.
  • Wash solution 2 [50% (v/v) methanol]. Methanol (50 ml) was diluted to 100 ml with dd.H2O.

Equipment required

  • A Tupperware dish, cleaned with detergent and washed with distilled water and 70% ethanol.

Procedure

  • Disassemble the electrophoresis gel plates from the SDS-PAGE apparatus and rinse briefly with distilled water.
  • Separate the glass plates with a plastic spacer if using the big Bio-Rad system or very carefully with a spatula if using the other mini-gel system.
  • If the gel appears to be sticking to one of the plates, do not try to pull it loose. Rather place the plates and gel into the Tupperware and add distilled water until the plates are submerged. The plates will separate easily and the gel will pull away.
  • Discard the water and follow the steps below
1. Fix >1 h [or overnight (o/n)]
2. Wash 1 [50% ethanol] 3 x 20 min
3. Pretreatment 1 min
4. Wash 2 [50% methanol] 3 x 20 s
5. Impregnation 20 min
6. Water rinse 2 x 20 s
7. Develop Until bands appear; keep an eye!
8. Water wash 2 x 2 min
9. Stop At least 10 min

NB: The washing procedure is always accompanied by shaking.

2. Coomassie Blue Staining of SDS-PAGE Gels

This method of protein staining has a reported sensitivity limit of around 1 μg/band whereas silver staining extends to below 200 ng/band. However, Coomassie staining is suitable for gels prepared for purification tables, and if the target protein is available in large amounts. However, always use a silver stain gel to assess protein purity if one wishes to use the protein as an antigen for antibody production.

There are methods on the web where staining and destaining times are reduced by microwaving. Although this does speed up the process, the hot methanol fumes are not pleasant and people will start complaining. Also, one should consider methanol flammable.

You will notice that the destaining solutions below do not contain acetic acid. Acetic acid is not required for destaining — it has historically been included in these solutions.

Solutions required

  • Coomassie stain (0.5% [m/v] in 50% [v/v] methanol). Coomassie Blue R-250 (0.5 g) and methanol (250 ml) are added to a 500 ml volumetric flask and made up to volume with distilled water. The solution is stirred until the stain is completely dissolved (30 min – 1 hour at RT).
  • Destain solution I (50% [v/v] methanol in distilled water).
  • Destain solution II (5% [v/v] methanol in distilled water).

Procedure

  • Disassemble the electrophoresis gel plates from the SDS-PAGE apparatus and rinse briefly with distilled water.
  • Separate the glass plates with a plastic spacer [if using the big Bio-Rad system] or very carefully with a spatula if using the other mini-gel system.
  • If the gel appears to be sticking to one of the plates, do not try to pull it loose. Rather place the plates and gel into the Tupperware and add distilled water until the plates are submerged. The plates will separate easily and the gel will pull away.
  • Drain off the water, and add excess Coomassie stain and keep on a shaker for 4 h.
  • After 4 h, decant the stain back into the bottle [it can be reused 3-5 times] and pour excess destain I onto the gel and shake for 4 h.
  • Pour off destain I and pour in excess destain II. Shake until completely de-stained and fully rehydrated. The gel can be stored in destain II until photographed or dried.

3. Imidazole-SDS-Zinc Staining

Coomassie Blue staining (CBB), SYPRO Ruby and imidazole–SDS–zinc staining are simpler to perform than silver staining, which is comparatively tedious. CBB and SYPRO Ruby staining, however, require overnight staining for maximal signal strength, while silver staining takes less than 2 h, and imidazole–SDS–zinc staining may be completed in less than 20 min for most 1D mini–gels. Imidazole–SDS–zinc staining is more sensitive than CBB and silver staining, and manifests the less protein staining variability. Imidazole–SDS–zinc staining, unlike SYPRO Ruby and CBB, is rapid and, because no fixative is used, is also completely reversible, making it compatible not only with downstream analysis by mass spectrometry and Edman sequencing, but also with immunoelution/western blotting techniques.

Reference

[bibliplug last_name=’Gillespie’ category=’Resources’]

Equipment

  • Glass petri dish

Solutions

  • Pretreater Imidazole–SDS solution [200 mM imidazole, 0.1% (m/v) SDS, 10–15 min]
  • Distilled water
  • Developer: 200 mM zinc sulfate

Method

  1. Pretreat by soaking the gels in imidazole–SDS solution for 10-15 minutes.
    [200 mM imidazole, 0.1% (m/v) SDS]
  2. Rinse briefly in distilled water for 30 seconds.
  3. Develope in 200 mM zinc sulfate until the gel background turns intensely white with transparent protein bands (15–60 s).
  4. The extent of development is best monitored during manual agitation of the gel over a dark surface and is stopped by discarding the developer and rapidly rinsing with running distilled water for 10–15 s
  5. As development of background continues for a few seconds after the developer is discarded, the reaction is best stopped just as the bands of interest became visible.
  6. Photography of gels is best achieved using epi–white light, on a dark background. No filter is required but bands in the low nanogram range (< 5 ng) are best photographed using a 610 nm longpass filter.