Sodium borate buffer gel problems - (May/05/2010 )
I recently switched to using sodium borate buffer after reading Brody and Kern's papers, as well as several of the forum posts here suggesting sodium borate buffer was vastly superior to TAE (which I was previously using).
After imaging my gel, the bands were very distinct (3kbp and below), but above 3kbp the ladder was not well separated (my gel was .8%, tried to run it at 300V, the machine only let me run up to 200V), after 10 minutes or so it migrated probably 1/3 down the gel, and both the buffer and gel were very hot (the end of the gel was very slightly melted). I ran it at 120V in hopes it would further separate the larger bands, but after maybe 10-15 minutes (not exactly sure as I was also finishing a mini prep), there was nothing on the gel. I tried staining it afterwards with EtBr (I had cast the gel with EtBr beforehand and put .5 micrograms/mL as the final concentration into the running buffer) but still nothing. A colleague suggested the bands had run off the gel, but it seems odd that after 10-15 minutes at 120V it would have run out of the gel.
I'm also trying to figure out if I'm using the sodium borate buffer correctly. I ordered sodium tetraborate decahydrate from Sigma Aldrich as per the request of my company's media core facility, who then made a .15M solution with pH 8.4. For the gel I diluted it down to 15mM solution and used that for the gel and running buffer (I realize now that I should have diluted down to 10mM).
Would incorrectly made sodium borate buffer have caused the problems I had? The running buffer being so hot makes me think the buffer I used wasn't made correctly.
I am ordering boric acid this time from Sigma Aldrich and will use the recipe I found on this site (47g boric acid + 8g NaOH, fill up to 900 mL water, dissolve and then fill up to 1L of H2O). Sorry for the long post. Any thoughts would be most appreciated.
Too much ionic strength will cause the gel to run "hot", though I wouldn't have thought running 1.5x buffer would have made that much difference. The hotter a gel gets, the faster it will run as the pore size should increase, perhaps this was your problem?
Sodium borate is not appropriate for separating bands over 3 kbp. It is good only for smaller bands which are common in routine PCR.
I recently tried to make a 2.5% gel with sodium borate buffer and while heating it, it turned to a slight yellow. Can anyone please tell me if this is normal or is something wrong with my buffer? My buffer recipe was: 45g of sodium borate + 8g of NaOH for a 10X stock (1 litre, adjusted ph to 8.5), and used 1x to make the gel.
bob1 on May 5 2010, 08:07 PM said:
I'm not sure, but I will report back on how it goes with a solution I will make myself and at 1X this time instead of 1.5X.
vladooo on May 6 2010, 03:19 AM said:
My bands are 3kbp or less, it was just the ladder that was pretty scrunched up and I was hoping that there was a way (running it at a lower voltage) to make the ladder separate a little more for a picture perfect gel. From what you wrote it seems like I am stuck with the ladder not separating at its bands higher than 3kbp.
Thanks for the responses. I'll report back when I get the new materials in and try it again.
I made the buffer myself this time, using one of the recipes listed on this site actually (8g NaOH, 47g boric acid, fill up to 900mL H2O, mix, fill up to 1000mL, and adjust to pH ~8.2 to make 20X solution). The 1X buffer after running at 300V was cool to the touch, and just for the heck of it, I ran RNA samples using this 1X SB buffer. Lo and behold it worked beautifully (I post-stained with SYBR green II stain), well separated bands that were bright and easy to see, and I only ran it for 20 minutes (1% native agarose gel).
I also tested it on DNA samples this morning, but for some reason the ladder appeared very faint (I used 6 uL of a .05 ug/uL 1kb ladder) compared to the DNA (used ~1ug). Same thing with the RNA I ran on the same gel - the 1ug of RNA appeared very bright compared to the 3ug of RNA ladder I loaded. I've used these amounts of ladder before, but I'm not sure if the buffer would cause this or the new stain I'm trying (SYBR green II stain - used 4ul for 20mL of 1X SB buffer post-staining, and placed on shaker for 20 min). Any ideas? I'll run it again using EtBr but I'm not sure why the ladder would be so much lighter than the samples since I've used this amount of ladder before.
Thanks for the responses guys.