SDS-PAGE - Trouble in running SDS-PAGE (Jun/02/2002 )
I have never Degassed my acrylamide I keep my APS and TEMED for months , I run my gels at a fixed voltage of 300V (20 mins) and I don't get streaks
Streaks are caused either by too much protein, by buffer conditions and by loading whole cells also by not making the gel up properly - some people just don't have the knack!
the first can be quickly and simply eliminated
The second is the most likely (common victims = arrogant postdocs ) things that go wrong:
1) Used 10x running buffer instead of 1X - very common or made the running buffer wrong
2)Had too much NaCl in you sample or you have very acidic or basic buffer - 1)EtOH ppt or TCA ppt (search google for protocol as I would recommend this regardless) 2) buffer exchange before running a gel
3) Sample buffer: not enough SDS (if you have a membrane protein then this is plausable)
B Meraptoethanol has gone off - Just put some more in (can't go wrong)
4) Made the gel solutions (Tris pH8.8 / pH 6.5 wrong) also common
If you are loading whole cells boil them for 15 mins in sample buffer with extra BMercaptoethanol (2ul in 50ul sample )
If you don't have the knack ask someone more experianced for Hands on assistance - this goes double for postdocs
APS and TEMED catalyse the polymerisation - if you leave them out then you are in for a long wait for your gel to set
I would recommend using the Maniatis protocol not the AMERSHAM.
Also I would strongly recommend using pre-dissolved APS
Can someone help us?
Sounds like your annode running buffer is mixing with the cathode running buffer during the run, make sure the levels of the two are sufficently different that one does not overflow into the other when someone knocks the aparatus / walks through the room
Other possibilities include not removing the isopropanol used during the setting of the resolver properly before pouring the stacker, and not using a large enough stacker (should have min 0.5 cm between bottom of wells and start of resolver)
Unless you have dodgy equipment you should be able to run the gels at up to 300V with no ill effect.
after u prepare ur buffers adjust the pH correctly and filter all the buffers before u use . Also check if ur acrylamide is fresh or not . run ur gel as soon as u load the samples. all the best
I'm running SDS-PAGE according to Amersham's manual. But I am finding it hard to set the gels. Most protocols tell you to use 0.05% APS. But it's hardly working. I used 0.01 g of APS and 10 ul (instead of 5ul) of TEMED for 10 ml of resolving gel and it worked. But as for the stacking gel, it didn't work. It has lower percentage of acrylamide just 4% so any tips on setting the gel without any anxieties.
I would like to know whether it is better using APS powder and accordingly adjusting the volume or using freshly prepared APS solution. Moreover the manual says that while degassing the solution (omitted APS and TEMED) the flask should be swirled all the time for 15 minutes. Is it true or we can leave it still? And then further on it says that after adding APS and TEMED, it should be swirled gently without producing bubbles? Is that possible? I even get bubbles while degassing. So how am I going to prevent that? Any tips.
Cowboy method for making SDS PAGE gels...... 0.1g of APS powder in water and vortex. Add 100ul to 10ml of separating gel (Final concentration 0.1% APS). Add everything needed for separating gel, except TEMED. Vortex to mix and let stand briefly. Gel with not polymerise without TEMED so no worries!! Add 25ul of TEMED to 10ml of separating gel and vortex again. Pour immediately and overlay with 100% EtOH (removes any bubbles). Separating gels polymerises within minutes. Pour off EtOH.
Stacking gel procedure is the same as above (50ul APS to 5ml stacking gel and vortex all together). Add 12.5ul TEMED to 5ml stacking gel, vortex, pour immediately and plce in well comb.
Again it sets hard in minutes. I have done it this way for years and no bothers as of yet!!
Streaks are caused by the reoxidation of cysteine residues in protein samples during electrophoresis. There is also documented research on the presence of "artifacts" at 66 kDa when mercaptoethanol is used as the denaturing agent in the loading buffer. I too had this problem. When denaturing protein samples use DTT, a much stronger reducing agent, and boil for 10 mins. This completely denatures proteins and the DTT prevents reoxidation.
I was running SDS electrophoresis today and after coomassie blue staining I had a clear gel (no bands and no marker). What can be the problem? The protein concentration is in order, the pH of the bufers I checked before running the gel and I used a new coomassie solutions. I was running the same samples as before. The protein was stored for 1 month in -20°C and for approximately three times thawed and freezed. The gel ran faster than usualy. I'm new in it and I don't know what happened:-(Thanks for your advices.
U have to aliquate your protein sample in many tube /sample if u want to keep it in -20°C .
and when u get it to run gel- 1 tube per time , don't storage it again.
Doan Lan, try using an alcohol as your overlay for the gel rather than water. It usually gives a cleaner edge to the gel and a "normalized" concentration of the acrylamide. Also if you are using 6x loading dye switch to 2X and try again. I usually get fuzzy bands if I use the 6x dye as opposed to the 2x.
i have been using SDS-PAGE for quite sometime and it always work very well, however i don't understand what is the function of stacking gel and the mechanisms of separating proteins by SDS-PAGE. can anyone tell me?