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Non-hazardous substitute for ethidium bromide? - (Aug/15/2012 )

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Is anyone using a non hazardous substitute for ethidium bromide (EtBr)?

I am interested in hearing what others are using.

Our lab is required to treat used EtBr (in gels and running buffer) as hazardous waste. This is expensive ($100 per pickup, 25 L pail of used gels), time consuming, paperwork, hastle, etc. To remove the EtBr from the used running buffer we stir overnight with "tea bags" from Amresco. Afterward we pour the running buffer down the sink. After 50 uses we discard the used tea bags with our used gel waste.

I've tested EZ Vision (Amresco), Sybrsafe (Invitrogen) and GelRed (Biotium).
I'm not plugging any product here. I only care about what works without any bias as to where I buy it from. We only have the camera filter for EtBr. EZ Vision and Sybrsafe require a the Sybrgreen camera filter ($900 - yikes!). However, GelRed works well with the EtBr camera filter.

Although Sybersafe and EZ-Vision appear to work well, the long exposure required (EtBr camera filter) causes too much background fluorescence. I would like to test the EZ-Vision with a Sybersafe camera filter because the safety data on this product appears to be very thorough. I think EZ-Vision can actually be discarded as a truly non-hazardous substance in regular garbage (double bag the used gels and pour used running buffer down sink with running tap water).

For time being I opted to pursue GelRed further because it uses EtBr filter.

At first I tried using GelRed in the gel but the PCR bands were too strong with smeary frown. I suspect the problem of heavy smeary frown may be due to overloading the gel. However, the smeary frown was still a problem, even after I tested serial dilutions of the DNA and different concentrations of GelRed in the gel.

In this whole experiment I tested a variety of PCRs between 100 bp and 1,000 bp.

I decided to try instead using GelRed in the loading buffer. I used gels and running buffer without any dye. This appears to work much better.

I tried the following amounts of GelRed in loading buffer:
1ul/100ul, 1ul/200ul, 1ul/500ul, 1ul/1ml, 1ul/2ml, 1ul/5ml

I added 5ul of GelRed loading buffers to 25 ul PCRs and loaded 10 ul into gel.
The gel and running buffer (1x TAE) do not contain any dye.

1/100 and 1/200 GelRed in loading buffer gives too strong staining for most of our PCRs, which requires us to titrate how much PCR to load on gel. It also gives some background fluorescence.

1/500, 1/1,000 and 1/2,000 GelRed in loading buffer appear to give optimal staining for most of our PCRs, which does not require us to titrate how much PCR to load on gel.

Although 1/5,000 GelRed in loading buffer works, it may give too weak staining for some of our PCRs.

See photo.

Several years ago we stopped using agarose gels and instead use agarose-composite polysaccharide gels. These gels are stronger, more flexible, more clear (less background fluorescence) and have wider separation range than agarose. At first we used Visigel from Stratagene, but they discontinued the product in 2007. They kindly gave me the recipe. If anyone is interested I will post my recipes. It consists of 0.7% agarose and 1% clarified high molecular polysaccharide from Locust Bean Gum (aka, carobe bean). As far as I'm aware the only folk that make cHMW-LGB is Diversified Biotech, (Agarmate, AGM-200).

Attached ImageIf anyone is interested I will post my recipes.


Please post your recipes.


Basically anything that binds to DNA has the potential to be hazardous, so it is unlikely that you will find something that is completely safe. There are a range of dyes that work with blue light illumination which cuts down on the UV exposure, and has the added bonus of not damaging the DNA when trying to see your bands prior to gel extraction.


How much gelred did you use in your gels?

I add gelred to my agarose at a dilution of about 1/40 000, although they recommend 1/10 000.


just don't buy RedSafe. It's terrible.


GelGreen Biotium. We use 83 uL of 10000x in 250 mL of 100 mM NaCl as a staining bath. We also kept the visualization system from EtBr.



Though I agree to bob1's reply that all chemicals that bind to DNA can be hazardous (if they can penetrate the skin), I'd say Silver staining is a quite save alternative that also has a very good characteristics (sharp bands, high sensitivity).
Anyway the other chemicals usually involved (formaldehyde, acids) and the silver itself (silver nitrate) not everyone might like. And the dying protocol is more laborious.
Here are some other alternatives (somewhere posted this pdf earlier in an other thread, too): Attached File


I'll reply in the order of posted replies.

RE: "phage434", I will post my recipe for composite agarose-cHMWLGB gels in my next reply.

RE: "bob1", I agree. All DNA binding agent must be handled safely. Same applies for anything that stains cells (precautionary principle). Wear gloves and change them regularly. I am agnostic regarding any product - as long as it works. I like the literature for EZ-Vision DNA stain (Amresco). It appears to be the closest thing to being non-hazardous. The US gov't toxicology program seems to support this.

RE: "leelee", "Curtis", "ascacioc" and "hobglobin", I work in a government diagnostic lab that uses PCR to routinely test for infectious diseases of livestock animals. So post run staining is not an option that will work for us because we have to produce results in a timely manner. That's why I'm looking for a non-hazardous alternatives to EtBr. One which works at least as well as EtBr and will be more efficient for our lab staff.

How terrible is RedSafe? Also, really how safe is it? That's a problem I have with chemical companies. So many new chemicals are developed and produced every day, yet apparently with minimal safety standards. Oh well.

Back to GelRed. So far I've tried several concentrations of in the gels, as follows:
- 1/10,000 (as recommended by manufacturer)
- 1/20,000
- 1/40,000
- 1/80,000

All resulted in smeary frown bands (remind me and I can post example photos). The problem is not in our gels itself because the 100 bp ladder looks fine with all , albeit fainter at lower . However, GelRed does work very well when used only in the loading buffer and not in the gel.


Serva offers DNA Stain G, they also claim that it's much less hazardous compared to EtBr....we got a free sample but didn't test it so far...Perhaps you give it a try with a sample...


Below is my recipe for Agarmate + Agarose Composite Gels, courtesy of Agilent-Stratagene, 2009. This replaces Stratagene Visigel (as of 2007, no longer available).
Modified from Perlman et al. 1987. Improved Resolution of DNA Fragments in Polysaccharide-Supplemented Agarose Gels. Analytical Biochemistry. 163: 247-254.

Agarmate-Agarose slurry: Updated May 25, 2012
Supplies for 250 mL of slurry:
- clean, dry 500 mL wide mouth plastic screw cap bottle (ex/ empty ultrapure diH2O bottle)
- 250 mL Absolute 100% ethanol
- 30 grams Agarmate, Diversified Biotech cat# AGM-200 (200 gm, High Molecular Weight Clarified Locust Bean Gum )
- 25 grams Agarose, Bio-Rad cat# 161-3101 (125 gm, Molecular Biology Agarose)

1. Add the 250 mL ethanol, then 30 grams Agarmate and 25 grams Agarose into the 500 mL bottle.
2. Close the lid, then shake the bottle for 10 sec. Label the date and contents. Store at room temp for 1 year.
Pouring a gel:
- clean, dry 1 L Erlenmeyer flask; Tuttle flask cover (Fisher, VWR, etc)
- clean 15 cm x 25 cm gel tray, top edges are marked for even spacing of combs and marked with red dots at "top end" and red dots at "bottom end"
- gel casting stand; 7 plastic 20 well gel combs (glued to homemade 1 cm x 1 cm x 20 cm acrylic plastic supports)
- 20 mL Agarmate-Agarose slurry
- 350 mL 1x Tris-Acetate EDTA (TAE) buffer, pH 8.3 (without ethidium bromide )
- 25 µL of 0.2 µg/µL EtBr (optional, can pour gels without any dye and use instead DNA dye in loading buffer)
- gel box containing just enough 1x TAE (with EtBr or without if using dye in loading buffer) to barely submerge the raised central platform
- o-ring with support stand
- microwave oven set to high; silicone oven mitts; safety goggles
- if using loading buffer containing DNA staining dye, we recommend using GelRed (Biotium) at 1 in 2,000 in 6x loading buffer
- 6x loading buffer (1x TAE, 50% w/v sucrose, 0.1% bromophenol blue, 0.1% xylene cyanole), store in 1.5 mL tubes at - 20*C

1. Centre the gel tray in the casting stand, then lock the assembly.
2. Level the gel tray assembly in a fume hood. Stand the o-ring at the back left corner of the gel tray.
3. Vigorously shake the bottle of Agarmate-Agarose slurry. (Until thoroughly mixed/resuspended.)
4. In a polypropylene 25 mL cylinder immediately measure 20 mL of the slurry, then pour it into the Erlenmeyer flask.
5. Add 330 mL of 1x TAE to the flask, swirl for 10 sec. Rinse the cylinder with the remaining 20 mL TAE and pour this into the flask.
6. Place the Tuttle flask cover on the flask and microwave 3 min.
7. Carefully remove the flask from the microwave, lightly weigh down the Tuttle cover, swirl gently until no more steam.
8. Swirl the flask vigorously for 10 sec.
9. Microwave for 30 sec.
10. Repeat steps 7 -9 two more times, except microwave 15 sec or until Tuttle cover can be heard jingling.
11. Repeat steps 7 and 8. Immediately transfer the flask into the fume hood while the molten gel is still hot.
12. Remove the Tuttle cover. Only if using EtBr in the gel: immediately pipet 25 µL of EtBr onto the centre of the molten gel. Otherwise do not add any dye.
13. (This step only if using EtBr in the gel. Replace the Tuttle cover. Swirl the flask vigorously for 20 sec to evenly distribute the EtBr in the molten gel.)
14. Pour the molten gel into the gel tray. Put the flask in the o-ring to drain the remaining molten gel.
15. Immediately insert seven gel combs. Center them, evenly spaced and in parallel.
16. Let the gel solidify for 1 hour. (If > 1 hour the gel will need longer to rehydrate before removing the combs.)
17. Unlock the gel assembly. Remove the gel tray with the combs still intact.
18. Place gel tray with combs in the gel box (red|red, black|black). Adjust volume of 1x TAE (with EtBr) to barely submerge the gel.
19. Let sit in the TAE for 1 minute to allow the wells to rehydrate. (Add 1 minute for every extra hour after pouring.)
20. Carefully tap/tip each comb. (This allows TAE to enter the wells and lubricate the combs.)
21. Gently remove each comb. Tug|wiggle alternating left|right up no more than 2 mm at a time.
22. Replace the gel box lid. Cover the gel box under a shopping basket (to block out light and prevent photo bleaching).
23. Store the gel box with gels and 1x TAE in the walk-in cold room. (This prevents growth of yeast in the TAE).
24. Wash the entire flask with hot tap water and Sparkleen detergent (Fisher). Rinse several times with hot tap water.
25. Rinse and rub the combs under running warm tap water. (Occasionally clean with warm tap water and Sparkleen).

Store gels and 1x TAE at 4*C when not in use. Example, on a cart in walkin fridge. If using EtBr in gel and in running buffer, store in dark.
Example, we cover gel box on cart with one of those heavy plastic shopping baskets which can be purchased at large supermarkets.

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