DNA Electrophoresis

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DNA Electrophoresis

Postby lrwilson » Sep 12 2003 9:18 am

Hello.

I have been mulling over something and I was wondering if someone could help me out.

Under most circumstances, when you run an agarose gel (normal, run-of-the-mill agarose) using a buffer such as TBE or TAE, would it be considered denaturing? That is, would your DNA strands be double or single-stranded? In this case, I am also doing no heat treatment prio to loading and using a typical loading buffer with dye and glycerol. My common sense would say, no. That this would not be denaturing and the DNA would be double-stranded, but I really would value a second opinion.

Ok, (sorry to get so long-winded) but then I was wondering, what would be the purpose to running DNA single-stranded? Would it be faster run time through the gel? When would you do denaturing DNA gels?

ANY help would be greatly appreciated. And please, do not merely tell me to do a lit search or look in Maniatus. If anyone has any answers they would be wiling to post, I would be extremely grateful.

Thanks! :-)
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Postby tfitzwater » Sep 12 2003 9:27 am

Agarose gels (TBE or TAE) are not denaturing.

Denaturing gels are typically run to examine single stranded DNA or RNA that might form secondary structures or hybridize to other fragments and therefore run aberrantly on a native gel. They are useful for analysis of PCR amplifications when the products appear as a smear on native gels due to the formation of multiplexes during the plateau phase of amplification. Denaturing gels are also used for single stranded nucleic acid:protein band shift experiments.
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Postby lrwilson » Sep 12 2003 11:06 am

Thanks for the second opinion and all the additional information.

The reason I began mulling over this question was because I am doing human identity testing where we denature our PCR products (it is a multiplex) with formamide and heating/snap cooling followed by capillary electrophoresis. One of the primers of the primer pair is fluorescently labeled so we are, therefore, only detecting one of the strands. We also use an internal sizing standard which is comprised of various fragments of DNA that are single stranded. One of these fragments, once it is in the polymer, will form a hairpin and doesn't not run true to size. Something in the polymer must allow this secondary structure to occur (I am guessing)which makes me think it is not a denaturing polymer or environment. Ultimately, I got to wondering why we denature the PCR products at all in this situation. Any ideas on what would be the reason?

Sorry, this post has once again gotten away from me. I suspect that I will probably call the company, but I wanted to see what the scientific community thought first. Thanks again.
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Postby tfitzwater » Sep 12 2003 12:10 pm

It can be very difficult to maintain complete denaturation of some DNA or RNA sequences. This is presumably due to minor variations in handling the samples and the particular sequences involved. Samples that are not denatured may travel as smears rather than discreet bands.

For example, DNA ladders that are composed of direct repeats such as a 10, 20, 30, etc. or 100, 200, 300 etc. base pairs will sometimes behave as single-stranded markers after standard denaturation procedures, but more frequently will appear as bands and arcs between bands due to partial renaturation even on standard denaturing gels. For this reason, DNA ladders prepared by restriction digests are better for denaturing gels.

High concentrations of urea (7 to 8.3 M) and formamide loading buffers help to reduce secondary structure. Each 1% formamide reduces the Tm of nucleic acid hybrids by approximately 0.6°C, therefore, 2x Formamide dye that is 97% formamide is 48.5% formamide at 1x and reduces the Tm by about 29°C. (Dysion, N. J. 1991. In Essential Molecular Biology: A Practical Approach. T. A. Brown (ed) Vol. 2, Oxford University Press, Oxford.) Some formamide dye recipes are only 84% formamide at 2x, or 42% at 1x for a Tm reduction of 25°C. 6 M urea also reduces the Tm by about 30°C. Monovalent cations in gel samples should be avoided, as the Tm increases 16.6°C for each one-fold increase in monovalent cations, between 0.01 and 0.40 M NaCl. Running gels at 45 to 55°C helps reduce structure problems. Running gels above 55°C may result in broken gel plates and may increase smearing of the oligo.

Oligonucleotides with significant secondary structure will travel faster than expected for a similar single stranded oligo of the same length.

For maximum denaturation (and more effort), try an 8% acrylamide/7 M urea/30-40% formamide gels (from the Sequenase protocol manual and 1990 USB Comments 17 (1) 31.) Combine 20 mL of 40% acrylamide:bis at 19:1, 42 g of urea, 10 mL of 10x TBE, 30-40 mL of formamide, and adjusted to 100 mL water. Warming to 35-45°C may be required for complete dissolution. (Optional, dissolve the acrylamide, bis, 10x TBE and urea in 40 mL of formamide by stirring a Parafilm-covered beaker in a 65°C waterbath until dissolved. This takes 30-60 min and the mixture reaches a temperature of about 50°C. Adjust volume to 100 mL and cool to < 30°C) Cool completely to room temperature before pouring gel, or the gel solution will polymerize before you can finish pouring the gel and insert the comb. Add 1 mL of 10% APS and 100-150 µL TEMED to initiate polymerization. The gel solution is extremely viscous and may require pouring with the gel plates at a nearly vertical angle. The gel should polymerize within 30 min. Formamide-containing gels may resolve very strong compressions. They will require higher running voltage and run slower than urea-only gels in order to run at the same power (wattage) as regular gels.
Gel will require fixing in 20% methanol/5% acetic acid: 0.2 mm gels/5 minutes, 0.4 mm gels/15 minutes and gradient 0.4-1.2 mm gels/60 minutes in order to prevent swelling.

Another option in the literature is a 10% acrylamide/formamide gel. Combine 25 mL of 19:1 40% acrylamide:bis premix, 0.37 g of Diethylbarbituric acid, 80 mL of formamide, 1.5 mL of 10% APS, and 240 µL of TEMED. Adjust to pH 9.0 with 1 M HCl and add formamide to a final 100 mL volume. The gels contain approximately 98% formamide. When diethylbarbituric acid is used in gels, the electrophoresis buffer is 20 mM NaCl in water.
When the diethylbarbituric acid is replaced with 250 µL of 1 M Na2HPO4 (final pH adjusted to 9.0) or 250 µL 1 M NaH2PO4 (to a final pH of 6.0), the buffer is 10 mM sodium phosphate in water adjusted to pH 9.0 or 6.0. In both cases, use 100 µL 36% (w/v) APS to compensate for the added water.
98% formamide has been used in the buffer reservoirs in place of water in some reports. (Caution: formamide is a teratogen. Formamide causes irritation to skin, eyes and mucous membranes. Vapor or mist may cause irritation. Can be absorbed through the skin. Ingestion and inhalation may cause headache, nausea, vomiting and stomach pain. May cause reproductive/teratogenic effects according to animal studies. Chronic exposure may lead to liver and kidney damage.)

Loading denaturing gels while the power supply is in operation will reduce the time the sample spends in the well and subject to partial renaturation as the formamide loading buffer is diluted in the sample well. Appropriate care must be taken in loading live gels. This technique is also useful for gel shift assays.
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