I'm looking to run a Western Blot to identify expression of collagen type I from whole cell lysates of Saos-2 osteoblast cell lines.

I've listed reagents at the bottom of the post.

Experimental conditions are as follows:

Seed cells in 6 well plate, once confluent mineralize for 0, 3, 6, 9 and 12 days.

Scrape well contents and pellet.

Lyse pellet with RIPA buffer.

Quantify protein using Biorad DC assay.

1. Mix LSB with protein sample so that LSB is 1X and there is 28ug of protein to load

2. Load samples into gel and run gel at 80V for 10mins to allow protein to pass through stacker

3. Once on resolving gel run at 120V until dye front is right at bottom of gel/runs off of gel

4. Set up apparatus for transferring on to nitrocellulose membrane

5. Pre-soak membrane, sponges and filter paper for 5-10mins in transfer buffer

6. Make up “sandwich” with paper, gel, membrane and foam sponges use falcon tube or 5ml pipette to roll out any air bubbles

7. Run at 200V, 400mA for 1hr 30mins, with ice block in tank with a magnetic stirrer

8. Ponceau stain to check transfer – wash off with H20

9. Wash blot with PBS-T 0.1% for 5 mins (x2)

10. Block for 30mins with 10% Milk powder in PBS-T 0.1%

11. Make up primary Ab in 5% Milk powder in PBS-T 0.1% (1:1000 for Col 1 abcam Ab) – leave overnight in fridge

12. Wash Primary off with PBS-T 0.1% for 15mins

13. Add secondary Ab in 5% milk powder in PBS-T 0.1% (1:4000 for Licor Anti rabbit 800CW) – leave for 50mins-1hr

14. Wash for 15 mins (x2) in PBS-T 0.1%

15. Read on Licor

I'm often seeing a band at 250kDa which is about double the expected band of 130kDa which I suspect may be a dimer.

The antibody is meant for native conditions but there are a variety of posts on the abcam website where people have produced the band using denaturing conditions - I've tried full denaturing conditions resulting in loss of signal/many bands which I imagine are degradation products of the full Col 1 structure?

Abcam themselves said that it is possible to run it with reducing but non-denaturing conditions - again this resulted in a 250kDa band.

When I run a "true" native gel, I've stripped SDS/sodium deoxycholate out of everything (gels, running buffer, transfer buffer, LSB and lysis buffer) but this results in the ladder running strangely, and the samples not migrating as they should, they get rather bunched up further up the gel as now charge will play a role in protein migration.

Any tips/advice would be appreciated, I've seen some posts on Reserachgate before but none that are the same as the exact scenario I'm encountering and none of them have a conclusive answer to this.

Additionally, if anyone has a native-PAGE protocol they would recommend please let me know.

Thank you for your time,

David.

Reagents:

Antibody:

Abcam34710

Stacker Gel:

Water – 2.8ml

Tris-Hcl 1M pH 6.8 – 0.5ml

30% Acrylamide protogel – 0.67ml

10% SDS – 40ul

25% Ammonium persulfate – 20ul

TEMED – 10ul

Resolving Gel:

Water

Tris-Hcl pH 8.8

30% Acrylamide protogel

10% SDS

25% Ammonium persulfate

TEMED

Running buffer (10X – dilute to 1X in DI Water):

30.3g Tris

144g Glycine

10g SDS

Transfer buffer (10X – dilute to 1X with 200ml Methanol, 100ml 10X Transfer buffer and 700ml DI Water)

144g Glycine

30G Tris

LSB:

6% SDS

22.5% glycerol

93.8mM Tris-Hcl ph 6.8

bromophenol blue

+15% fresh B-mercaptoethanol added fresh on day of use