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F A Q
 

    basic information       
1  What is the organizing concept of FBM?

Run Gels FAST!  

Tris was a 30-year mistake in the history of DNA electrophoresis. Our research indicates that the simple substitution of a Tris-free solution in DNA electrophoresis allows you to use much higher voltages to separate DNA fragments 4- to 15-times faster in your standard gel rigs - without giving up anything. Run the most common DNA sizes, from 200 to 800 bp, at 4 to 6 times faster than usual. Even smaller? -- run them even faster.

You do not need to run routine gels over lunch break or overnight. You can stop running gels the way your mentor did.
 


2  Do I just dilute the 20X FBM media with water to form a 1X solution - then make my gels and run them in the same 1X solution?

YES. LB®, SB™, and LA® replace TBE and TAE. Usually there is no need to change any protocols. Just cross out TAE and TBE, and write in LB, SB, or LA. Our newest product, LB, is similar to SB but conducts even less current - for even less gel heating. Check out our How to Use LB and our How to Use SB page. LB is recommended for most PCR analysis, for oligos, and for fragments under 2500 bp. LA is recommended for standard plasmid digests or whenever the DNA fragments are larger than 2000-3000 bp. See our How to Use LA page.
 


3  Can I run SB and LB gels at standard conditions?
YES. But why wait over an hour to get a result when you can get crisp bands within 15 minutes in denaturing and in nondenaturing gels? Crank the voltage to speed DNA migration.
 

4  Do I need to buy any new electrophoretic equipment with this new conductive medium?
NO. LB, SB, and LA are superb one-for-one replacements for your current buffers (TBE and TAE). But you might be limited by your older power source if it does not supply the higher voltage to take full advantage of low-conductance gels.
 

5  Why am I not using LB or SB already?

They were only recently discovered - LB in 2004 and SB in 2003 - by FBM researchers.

U.S. Pat. 7,163,610 was awarded to FBM in 2007, and U.S. Pat. 7,811,437 in 2010.


6  How do I decide between LB and SB?

LB is the general choice for analysis of smaller DNA fragments, such as PCR products. Both LB and SB provide excellent separation of these smaller DNA fragments in standard agarose and polyacrylamide gels. LB generates less heat than SB, and so LB is favored for the highest voltage runs and for visualizing sub-100 bp fragments, including oligos. In most situations, SB and LB are nearly interchangeable.

SB generates more heat than LB due to sodium having a smaller shell of hydration and thus being more electrokinetically mobile. At moderate voltages, a user would not notice a difference, but at the highest tolerable voltages, the SB gel would start to melt when an LB gel would not.

For general use, 1X LB or SB can be used with standard agarose gels. For resolving DNA fragments over 2000 bp, gels made using 0.3-0.5% of a strengthened agarose (such as FMC's Seakem® Gold) and 1X LB are recommended. For very small (less than 100 bp) double-strand DNA, 0.2 to 0.5X LB works well in standard agarose at 2 to 2.5%.

An investigator using LB for the first time should initially try a voltage setting four times higher than they would use for TAE or TBE. If using SB, the initial voltage setting should be set at three times higher than for TAE or TBE. Once experience is gained, voltage settings can generally be adjusted somewhat higher.
 


7  What is best for standard plasmid digests or for separating large DNA fragments?
Large DNA can also be run fast in a gel. If you use LB or SB, a commercially available high-strength agarose in a 0.3 to 0.5% gel is best to separate fragments over 2000 bp.  If you prefer to use a standard agarose gel at 0.7 or 0.8%, use LA to optimally resolve large DNA fragments. LA is borate-free and tris-free and is a replacement for TAE for standard plasmid restriction digests. Borate, which is present in LB and SB, causes reversible crosslinks with DNA, reducing the separation of larger DNA fragments unless low % gels are made of high-strength agarose. See our tutorial on large DNA
 

   

    compatibilities           


8  Are FBM media compatible with my other DNA procedures (gel extractions, sub-cloning, etc.)?
YES. We tested common commercial kits to extract DNA and to clone fragments isolated from gels made with FBM conductive media. Your DNA might even be happier in a cooler medium such as LB, SB, or LA.
 

9  Do FBM media work in polyacrylamide gels?

Most customers use 2% or 3% agarose gels instead of polyacrylamide because SB and LB resolve small fragments of DNA very well. This is nice since acrylamide pre-cast gels have a more limited shelf life than agarose gels and because agarose gels are easier to make.

There is a feature of polyacrylamide gels about which investigators have long known to be careful. Ammonium persulfate (APS) is used to polymerize the acrylamide in order to create the polyacrylamide gel. APS is a salt, a high ionic load. Different investigators use differing amounts of APS to polymerize their gels, depending upon both the speed of polymerization desired and the age of the APS solution (it loses effectiveness even after one week, and so some investigators compensate by using more in their gels).  To handle the ionic load of APS, for decades many investigators have preferred to "pre-run" their polyacrylamide gels so that the APS migrates out of the gel. They then load their samples and run the gel normally. If care is not taken to handle the APS ionic load, curved migration paths and poor resolution can result. The high salt content of APS is one reason to favor agarose over polyacrylamide. The agaroses used for electrophoresis are of high-purity with negligible salt content.
 


10  Are FBM media compatible with colored versions of PCR polymerases?
PCR products produced using colored polymerases have been analyzed on FBM gels. No difficulties are reported by our customers to date. With these products, a separate loading medium usually is not added to the PCR reaction prior to loading on a gel. Note, however, that the higher salt content of an undiluted PCR reaction will not produce the highest resolution band separation when compared to a lower-salt sample.
 

11  Are FBM conductive media compatible with specialty agaroses?

LB, SB, and LA perform nicely using a standard low EEO molecular biology grade agarose such as FBM Tap-Out™ low-EEO agarose. Some special-purpose agaroses, such as low melting temperature agaroses, have also performed well with our low-conductance media.

Some forms of specialty agarose will require a high salt environment to retain their strength and optimal optical properties, and are not suitable for low-conductance media. The composition of such agaroses are generally proprietary; it has therefore not been possible to predict in advance which of the more expensive agaroses will prove most suitable for use in high-voltage electrophoresis.


   
    technical properties   
12  What is the buffering capacity of LB or SB?
LB and SB will maintain the pH near 8 as well or better than TBE and much better than TAE - before, during, and after your electrophoretic run.
 

13  Can I recirculate LB or SB during a run?
YES, although the delayed electrolyte exhaustion rate of LB and SB as compared to TBE and TAE generally will make recirculation unnecessary.
 

14  Sometimes I re-use my TBE and TAE. Will LB and SB allow me to do this?
 

YES. LB and SB have an improved electrolyte stability as compared with tris-containing and acetate-containing media.
 

15  Where does orange G migrate?
 

Orange G is the dye provided in our loading media. When photographed, it will not obscure DNA bands as will bromphenol blue and some other dark dyes.  Orange G co-migrates in agarose gels near the 100 bp DNA fragments, although this varies somewhat with agarose concentration.
   
    other questions we have received       

16  I ran a 10 cm SB slab gel at 500V, and it got warm. Isn't SB supposed to run cool?

LB and SB run cooler. You can be certain that at 500 volts, your TBE or TAE gel would have boiled over.


17  In adjacent lanes, I compared a sample in my old familiar loading buffer with the same sample in the SB loading medium. The DNA migrated differently, with the lane containing the older loading buffer running a bit farther and every band a bit curved like a smile. I also compared ladders from different companies, and similar-sized bands don't exactly migrate side-by-side. What is going on? 

The differing migration distances are due to differing salt contents in the loaded samples. When one has a higher salt content in a particular sample, the DNA in the same lane will migrate faster and often migrate farther in the middle of the lane (bending), perhaps due to local heat generation caused by a locally increased conductance. This is true of all electrophoretic systems, irrespective of whether one analyzes proteins or polynucleotides. It is advisable to try, as best one can, to match the salt content of the loaded sample to the salt content of the reference samples and, preferably also, to the salt concentration of the gel. It is for this reason that restriction digestion products and PCR products can often run at the "wrong" size when compared to a standard ladder. Most older recipies for loading buffer have a salt content about four times or more that of the LB and SB loading media. In contrast, the LB and SB loading media are matched closely to the salt content of the  gels, at 10 mM cation concentration. If one does not match the salt contents of the reference and analytic lanes, one cannot determine the relative sizes of DNA with any high degree of confidence. In a brief attempt to quantitate the influence of salts on 1X LB and SB gels, migration was quite notably affected when the sample exceeded 50 mM salt, and the resolution was worse when the predominant sample cation was tris. Luckily, most users do not need to determine absolute fragment lengths very often. For those that do, the matching of salt content is critical, or ladders can be mixed with the analytic sample and loaded into a single well to provide a salt-independent internal reference marker for each lane.


18  I like to load restriction digests directly on my gel. How can I handle the high salt load that I know is in my samples? 

For most restriction digests, salt will not be a major problem. Usually, you can mix one part of the 5X FBM loading medium with 4 parts of your unpurified restriction digest to form your sample that you will load on the gel.
 
Some restriction digestion buffers, however, are very high in salt. Reading the answer to FAQ 18 confirms that you may experience altered migration distances and fuzzy bands if you load solutions containing over 50 mM salt and high amounts of tris in a low-salt electrophoretic system. Tris gels are a high-salt system, which explains why you may not have noticed the salt effects in the past. To use the new electrophoretic media with your saltier restriciton buffers, try the following ideas. Suggestion A produces the highest quality separations, while suggestion C is the most rapid.

  A  Consider purifying the DNA by precipitation or a column. Or to keep the time demands down...

  B  Try diluting the sample. For example, we like to add (by pipetting up and down a couple of times to mix) 2 parts of an unpurified restriction digest to a well containing 2 parts water and one part 5X loading medium in a U-bottom microtiter plate (or use 3 parts of an appropriately diluted loading medium, which is essentially the same thing). One can do 8-12 samples at a time and load them onto a gel without changing tips. Or even simpler...

  C  Try using a 2X concentration of FBM media in your gels and reservoirs to reduce the discrepancy in salt content of your samples and the gel. Keep the loading medium at 1X. You can run a 2X gel faster than a tris gel, although not as fast as with a 1X FBM medium.


19  How good is the resolution in a critical situation, such as multiplex PCR?

Multiplex PCR is a perfect example of the increased resolution of low-conductance media. See our PDF comparing TAE and SB in multiplex methylation-specific PCR.

20  How does the toxicity of FBM media compare to TBE?

FBM media are less toxic than TBE. The major toxicant is boric acid (and borate). Although few reports exist, it appears that the acute ingestion of 30 grams of boric acid can be lethal if untreated. To reach this lethal dose, one would need to ingest 2.7 liters of 1X TBE, or 12 liters of 1X SB or LB.  Boric acid and borate are not significantly absorbed through skin.

Lithium is somewhat less toxic than the borate, given the amounts used in FBM media. Indeed, lithium used to be a component of 7-Up and some lithia beers. Ingesting 12 liters of 1X LB would be expected to provide moderate toxicity from the lithium alone (but a potentially fatal quantity of boric acid). The same dose of lithium, supplied by LA, would require the acute ingestion of 24 liters of a 1X solution.  These are not likely scenarios.

Considering the above numbers, any realistic danger of toxicity would have to come from acute ingestion of a few hundred milliliters of concentrated solutions of these conductive media (for example, stock solutions of 10X TBE or 20X SB, LB, or LA).


21  I like to control the current in my electrophoresis. Can I crank the current?

No.  FBM media are intended to dramatically reduce the current drawn by an electrophoretic gel at a given voltage. This allows one to increase the voltage, and thus the speed at which DNA migrates. Increasing the current by cranking the amperage settings at the power supply would engender a tremendous increase in heat generation, ruining your gel. Still, setting a maximal current or maximal wattage is good practice in DNA electrophoresis, for those with more sophisticated power supplies, as it can permit the use of maximal voltage with less danger of reaching excessively high gel temperatures.

22  How do I use FBM's borate media to resolve single-strand DNA in agarose and without chemical denaturants?

This application was published in BioTechniques in October, 2004. It takes advantage of the ability of low-salt solutions to retard the speed of DNA hybridization after melting has been accomplished. In our publication, we used a matched loading medium to melt the sample and to apply it to a well of a horizontal agarose submarine gel. We made fresh a special 5X loading medium that contained 0.5X LB, 50% glycerol, and only trace orange G for the slightest coloration (the dye can be omitted to further reduce salt). 40 ng of each oligo were added to the loading medium and water to make a 1X sample containing 0.1X LB, 10% glycerol. The DNA of the sample was melted at 70 C for 5 minutes. The melted sample was loaded onto a 3% agarose gel that had been pre-heated to 37 C in an incubator. Electrophoresis was done at 29 V/cm. Lower temperatures of gel did not work, presumably due to reduced stringency. (Other methods of achieving sample melting would presumably be permitted, such as non-ionic conditions or the use of denaturants in the sample. We have used formamide samples for standard polyacrylamide gels using 6M urea, for example.)


23  FBM loading media produce a light yellow band on my gels. How can I get a darker dye marker?

You have a number of options, and most of them will perform quite well.  For many types of DNA samples, any dye will work and any loading media available to the researcher - from any source - usually will perform at least adequately.

FBM uses orange G in the loading media. We utilize orange G because it will not obscure the visualization and photography of DNA bands as will bromphenol blue (BPB) and some other dyes. In order to optimally match the loading media to our 1X low-conductive solutions, we have set the concentration of the orange G rather low. We do this because all of the migrating dyes for electrophoresis are anions, usually supplied as a sodium salt.  In order to provide maximal resolution, to minimize the effects of salt on DNA migration speed, and to provide maximal flexibility for users whose samples might have a bit too much salt (such as when analyzing unpurified PCR products), we minimized the salt in the loading medium and provide a sample of the FBM loading medium matched to each type of conductive media you use (LB, SB, or LA). We indeed developed our new conductive media using a standard TAE-based loading buffer containing BPB; while it worked adequately, we were more satisfied after developing our current set of matched loading media.

In short, here are some simple options for you. 1) Try any loading buffer with which you are comfortable. 2) Add additional dye solution of your choice to our matched loading media - just take care not to add so much that you get undesirable salt effects in your lanes. 3) Use FBM loading media at a 1.5X or a 2X concentration. Our 1X solutions have 10% glycerol, which allows considerable leeway in acceptable concentrations. If you use option 3, you will likely have a darker migrating marker as desired, and still have considerably less salt in your loaded sample than if you use a standard, older-style loading buffer.


24  Do FBM media have an expiration date?

There is no known basis for the products to expire. Since the invention of these novel conductive media in mid-2003, no precipitation or other evidence of degradation has been observed. We continue to store and re-test our initial samples, but have not yet detected a change in performance.


25  How do I request a certificate of analysis?

All FBM media offered for sale are filtered at 2 micron and have passed three tests at the time of manufacture. 1 - an assay of conductivity, 2- a determination of pH, 3 - a functional test based on the resolution of a DNA ladder in agarose electrophoresis. For facilities needing a Certificate of Analysis, date of manufacture, or other reference information, please email us or contact us using the online form at this link. Include the lot number found on the container label and your email address. We will attach the relevant Cert. of Anal. to our return email. Certificates can also be sent by mail upon request.