Volume 1 Number 2
Feature articles
Nature's palett - How humans and other animals produce colours
Margareta Wallin
Department of Zoology, University of Gothenburg, SE.
Read the article
Cheese and cheese making
Ulla-Kerstin Nilsson-Blom och Per-Olof Weréen
Ostfrämjandet, Falkenberg, SE.
Read the article
Viruses - A tool for gene therapy
Erik Nordenfelt
Professor emeritus, University of Lund, SE.
Read the article
Practical protocols
Green DNA
Simple isolation, restriction and electrophores of chloroplast DNA
Leighton Dann, Science and plants for School, Cambridge University, UK.
Amplification of mithocondrial DNA
John Schollar och Andy Harrison, NCBE, The University of Reading, UK.
Reviews
The human genomeJeremy Cherfas (Series editor John Gribbin) |
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First fruit - the creation of the Flavr SavrTM tomatoBelinda Martineau
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The blue planet- a natural history of the oceansNarrated by David Attenborough
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The seven daughters of EveBryan Sykes
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Biography of a germ Arno Karelen
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Genes, girls and GamowJames D. Watson
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The Schollar Test
Safer stains for DNA
Guest reviewer Dean Madden tests safer alternatives to ethidium bromide for staining DNA on electrophoresis gels.
The stains under test were:
- Methylene blue
- Nile blue sulphate
- Sigma BlueView (TM)
- Azure A
- CarolinaBLU (TM)
- Crystal violet
- Brilliant cresyl blue
Ethidium bromide, a potent mutagen
In research laboratories, ethidium bromide and similar fluorescent compounds such as Acridine Orange are normally used to visualise DNA on a gel. Unfortunately, ethidium bromide and its breakdown products are potent mutagens and carcinogens and therefore they should not be used in schools. Such dyes are often flat molecules with similar dimensions to DNA base pairs. When ethidium bromide binds to DNA, it slips between adjacent base pairs and stretches the double helix. This explains the dye's mutagenic effect - the 'extra bases' cause errors when the DNA replicates. In addition, short-wavelength UV light (which itself is harmful) is required for ethidium bromide to fluoresce and reveal the DNA. For reasons of safety and because UV light of this wavelength causes unwanted mutations in the DNA being studied, several researchers have sought alternative methods of revealing DNA.
Safer alternatives
Crystal violet binds to DNA in a similar way to ethidium bromide and although it is a mutagen, it is not thought to be as harmful as ethidium bromide. Because it can be viewed in normal daylight (avoiding the need for damaging UV light), some researchers have advocated its use where functional DNA is to be recovered from a gel.
Thiazin dyes
The most widely used alternatives to ethidium bromide are methylene blue and its oxidation products, such as Azures A, B and C, Toluidine blue O, Thionin and Brilliant cresyl blue.
These dyes are used individually or as mixtures (often in proprietary formulations). Although their exact mode of action is unknown, they are thought to bind ionically to the outside of nucleic acids (to the negatively-charged phosphate groups) and can therefore be used to detect both DNA and single-stranded RNA.
Such dyes are not as sensitive as ethidium bromide, and some of them colour the gel heavily. Consequently, prolonged 'destaining' may be necessary before the DNA bands can easily be seen. Several dyes also fade rapidly after use - methylene blue falls into both categories and is therefore, despite its popularity in school texts, not ideal for staining DNA on a gel.
All of the thiazin dyes may be used in aqueous solution at a concentration of about 0.02-0.04% and applied to the gel after it has been run. They may also be dissolved in mild alkaline solutions (e.g., running buffer; not over about pH 8). Destaining with dilute acetic acid or 0.2 M sodium acetate buffer, pH 4.7 may be necessary for alkaline solutions.
The age of the dye may have a considerable effect upon the results achieved. For example, old samples of methylene blue will almost certainly contain a proportion of other dyes (such as Azures A and B) and these breakdown products may be responsible for much of the staining. Dye solutions are best stored in glass bottles (some dyes will stain plastic containers), either wrapped in foil or kept in the dark.
Staining DNA on the move
Recently, several commercial products have emerged that enable the DNA to be seen as it moves across the gel. Suppliers seldom reveal their composition, but several of these stains contain Nile blue sulphate (also known as Nile blue A), a dye which had not previously been noted for its ability to stain DNA. Adkins and Burmeister (1996) give useful guidance as to its use as well as hints for identifying other dyes which may be useful for visualising DNA. Before it left the schools education market, Stratagene used to sell a product called 'Stratabloo', which was amixture of Nile blue sulphate and methylene blue.
All of the dyes used for staining 'mobile' DNA are cationic - that is, they are positively charged in the gel buffer, at pH 8. They move through the gel in the opposite direction to the DNA, latching onto the DNA molecules as they meet them. There exact mode of action is unknown, but, for example, Nile blue sulphate is thought to intercalate within the DNA double helix.
So that sufficient dye remains in the gel, it is added to both the gel and the buffer above it. However, a far lower concentration (1-3 µg per ml) of dye is necessary for this method than for post-electrophoresis staining. This is because too much dye will neutralise the negatively-charged DNA fragments, slowing their movement and reducing the resolution or even preventing the DNA from moving at all. Consequently, there is a compromise to be struck between visibility and resolution. Better results are usually achieved by staining the DNA after the gel has been run, rather than staining during the run.
Drying gels
It is also possible to dry a gel after the dye has been applied, and thereby to concentrate the dye in bands which would otherwise be difficult to see. So that the gel dries evenly, it is advisable to place the wet gel on a sheet of good-quality writing paper, and to place this on several sheets of filter paper. Moisture from the gel soaks into the filter paper, while the writing paper layer stops too much of the dye from soaking out of the gel. Gels should be dried at room temperature.
Safety
Although several dyes that can be viewed in normal daylight are thought to be relatively safe, they have not been as intensively studied as the fluorescent dyes for long-term toxic effects. Some of these visible dyes, apparently, intercalate DNA like ethidium bromide so they too have a potential for mutagenesis and, depending on absorption and metabolism, a potential for carcinogenesis. As with all laboratory chemicals, suitable safety precautions should be exercised when handling any dyes, particularly when they are in dry, powdered form.
Further reading
Methylene blue
Yung-Sharp, D. and Kumar, R. (1989) Protocols for the visualisation of DNA in electrophoretic gels by a safe and inexpensive alternative to ethidium bromide. Technique 1 (3) 183-187.
Flores, N. et al (1992) Recovery of DNA from agarose gels stained with methylene blue. Biotechniques 13, 203-205.
Brilliant cresyl blue
Santillán Torres, J. and Ponce-Noyoia, P. (1993) A novel stain for DNA in agarose gels Trends in Genetics 9 (2) 40.
Nile blue sulphate
Adkins, S. and Burmeister, M. (1996) Visualization of DNA in agarose gels as migrating colored bands: Applications for preparative gels and educational demonstrations Analytical Biochemistry 240 (1) 17-23. http://www-personal.umich.edu/~steviema/blueDNA.html
Crystal violet
Rand, N. (1996) Crystal violet can be used to visualise DNA bands during gel electrophoresis and to improve cloning efficiency Technical Tips Online http://research.bmn.com/tto
This article is an extended version of one from 'Illuminating DNA' by Dean Madden.
Image lab
Great balls of yeast
Yeast cells immobilised in calcium alginate
Image courtesy: Department of Chemical Engineering, University of Birmingham
Guy says:
I don't know what possessed me to try my hand at making wine, but I'm pretty sure that it had something to do with one of our Chancellors of the Exchequer.
Although my initial attempts produced nothing more than a sweet, murky soup, I can now produce something of which at least my local Berni Inn would be proud.
It was during a visit to the NCBE at The University of Reading that I first heard of immobilising yeast cells. The process basically involves mixing your chosen yeast with a 2% sodium alginate solution and then adding this, a drop at a time via a syringe, to a 1.5% solution of calcium chloride. This traps the microbes inside tiny beads, which allow sugar, alcohol and carbon dioxide to pass through freely, but contain the yeast throughout the fermentation. When the fermentation is complete one simply draws off the wine, leaving the yeast balls in the demi-john.
And so, armed with my bottles of sodium alginate and calcium chloride solutions, I returned home with thoughts of those tiny beads floating up and down to the rhythmic 'plop-plop' of the air lock and a perfectly clear wine every time. Back in the kitchen I set about preparing the living quarters of my yeast whilst my filter kit and finings looked on disapprovingly.
The chosen wine was apple as this had proved successful in the past; and the yeast - it had to be a Sauterne type. The juice was extracted from about 10 lbs of apples (with a few pears thrown in). Enough granulated sugar was added to bring the total sugar content of my gallon of liquid to about 2.5 lb - which should produce a wine of around 12.5% alcohol. The acidity was measured by titration and adjusted until it was about 3.5 ppt. Finally, a yeast nutrient was added followed by my yeast balls - which immediately sank to the bottom of the jar and stayed there. After a couple of days the fermentation was well under way, however, and the yeast balls were floating merrily around on the surface.
I must warn you at this point that the initial fermentation can be quite vigour ous and even a slight shake of the demi-john will cause the 'wine' to come frothing out. It is therefore prudent to allow a good air space in the demi-john for the first week or two.
To my disappointment, once the yeast balls were floating they refused to budge, even towards the end of the fermentation. Nevertheless, when complete, the task of racking off the wine from the yeast was greatly simplified and could almost have been carried out with a sieve but for the fruity sludge at the bottom.
And the wine? Well, it's too early to tell yet, but it certainly looks promising. Oh, and by the way, my yeast balls have been transferred to another demi-john of fruit juice and seem content to sit there all day converting more sugar into alcohol for me.
Guy Madden
This article first appeared in the NCBE Newsletter in 1991.
