Tris-Tricine protocols

[KLAUS LEHNERT]

Hi netter's

does anyone out there have a good (and tried) protocol for
Tris-Tricine Polyacrylamide gels?
I have tried the protocols in the "original" publication
(Schaeger+Jagow 1987), I'm using high-quality reagents, but at
the very best I'm getting very fuzzy bands, and even when I run
the gel (in a BioRad Mini-ProteanII) for twice as long (i.e.
duoble the time the bluemarker needs to run out), my smallest
marker band (Aprotinin, 6.5kDa) is still in the upper half of
the gel.

Any suggestions?

KLAUS

[Volker Nölle]

We use a modified and easy protocol of Tricine-SDS-gels with good results:

16.5 % gel:

4 ml 49.5 % Acryl/Bis (100:3)
4 ml 3 M Tris 0.3 % SDS pH 8.45
1.6 g Glycerol
2.4 ml MQ

4 % stacking gel:

0.5 ml 49.5 % Acryl/Bis (100:3)
1.55 ml Tris 0.3 % SDS pH 8.45
4.2 ml MQ

Anode buffer:
0.2 M Tris pH 8.9

Kathode buffer:
0.1 M Tris pH 8.25
0.1 M Tricine
0.1 % SDS

Run overnight at 12-16 mA in a gel as big as possible (about 20 cm).

Dye with colloidal Coomassie (do not use Destain before !):

0,8 g Coomassie G-250 in 400 ml MQ=20
add 400 ml 1 M H2SO4
stir for 3 h, filtrate (folden filter)
add 88 ml 10 M KOH and 124 ml 100 % TCA

dye for some hours or overnight at room temperature
then 1 or 2 hours in water
sensitivity: 0,2 =B5g protein

good luck!

Volker Noelle
Institute of Biochemistry
University of Cologne
Germany

email: a223...@smail.rrz.uni-koeln.de

[Dr. Frank O. Fackelmayer]

In article <5g7n7v$3d8$1...@scream.auckland.ac.nz>,
mmm_kl...@mednov1.auckland.ac.nz (KLAUS LEHNERT) wrote:

Hi Klaus,
I also had some problems with the original protocol of Schaegger. I´ve
changed some things and now it works like a charm.

1. Always run 12%gels (not 16.5% or with two layers of resolving gel)
2. Don´t use any glycerol in the gel (replace its volume by water)
3. Use a minigel (some 6cm high), not a long gel !!!!!
(From what I´ve learned while optimizing the procedure, this is the most
important step. Although your bands are quite close to each other, the
separation is great, at least down to 3kD. Using longer gels does not only
produce fuzzy bands, but also takes an awful lot of time.)
4. Run your minigel at low current (30V/approx. 15mAmps) until samples are
in the stacking gel, than change to higher current (say 30mAmps) and run
your gel until the coomassie blue just runs off the gel. In my case, this
takes some 60min, but of course it is dependent on your gel dimensions.
Try varying your electric parameters to have the gel finished in one hour.
5. Stain your gel with fixating Coomassie Blue (Steck et al. 1980, Anal.
Biochem. 107:21-24 is great for that).

If the problems remain, change your acrylamide solution.

Hope this helps,
Frank

[Shiwani Arora]

KLAUS LEHNERT wrote:
>
> Hi netter's
>
> does anyone out there have a good (and tried) protocol for
> Tris-Tricine Polyacrylamide gels?
> I have tried the protocols in the "original" publication
> (Schaeger+Jagow 1987), I'm using high-quality reagents, but at
> the very best I'm getting very fuzzy bands, and even when I run
> the gel (in a BioRad Mini-ProteanII) for twice as long (i.e.
> duoble the time the bluemarker needs to run out), my smallest
> marker band (Aprotinin, 6.5kDa) is still in the upper half of
> the gel.
>
> Any suggestions?
>
> KLAUS

I have routinely used tricine gels for analyzing my small 14 kDa
protein. I use the following recipes:

Separating Gel: (15%T, 6%C)
BioRad's minigels, 0.5 mm spacers, two gels.

12 ml total volume

1.9 ml distilled water
4 ml gel buffer
6 ml 30% polyacrylamide, Bis stock
60 ul 20%SDS
5 ul TEMED
50 ul 10% APS (Ammonium persulfate)

Stacking Gel:

3%T, 3%C
2 gels, BioRad's miniprotean, 0.5 mm spacers

3.18 ml distilled water
1.24 ml gel buffer
0.5 ml 30% polyacrylamide, Bis stock
25 ul 20%SDS
5 ul TEMED
50 ul 10% APS
5 ml total volume.

Gel Buffer:
3M Tris, pH 8.45
0.3% SDS
(72.6 g Tris, 3ml of 20%SDS, in 200 ml final volume)

Fill up the top buffer tank with 1X Cathode buffer
10 X Cathode Buffer:
1 M Tris, pH 8.25 (Do not adjust pH)
1 M Tricine
1% SDS
(60.5 g Tris, 90 g Tricine, 25 ml of 20% SDS stock, final volume 500 ml)
Dilute 20 ml of 10 X Cathode Buffer to 200 ml with water to make 1 X
Cathde Buffer.

Fill up the lower tank with 1 X Anode buffer.
10 X Anode buffer:
2.0 M Tris, pH 8.9
(121 g Tris in 500 ml final volume)
Dilute 40 ml of 10 X Anode Buffer to 400 ml final volume with water to
get 1 X Anode Buffer.

Keep your buffer stocks in fridge. Do not reuse buffers.Dilute your
cathode and Anode buffers just before use.
Run your gels at 50 mA constant current. The bromophenol blue dye should
reach the bottom of the gels in about 3 hours.

Try this protocol. Let me know if you run into any problems. Good luck.