Published: May 6th, 2016 Last Modified: September 18th, 2020
There are some products from large molecular biology companies that I think are definitely worth the hefty price tag. Q5 polymerase from NEB and Superscript III from Thermofisher are two which give awesome results. The amount of R&D involved to mutate your pedestrian polymerases and give them cool and useful features must be astronomical. Not to mention paying fellow pipette jockeys to grow up huge batches, purify them to a high degree and distribute them. Worth it, take my money, hats off for making a good product.
Now, you want me to pay 35 bucks for half a liter of “ultrapure water” or for a simple buffer? There are situations where purchasing these could be called for, perhaps where traceability is absolutely necessary. For your average Joe/Jane Shmoe lab doing basic research? Naaaaaaahhhhhhh, comon buddy. One reagent I’ve come across that’s expensive to buy but costs very little to manufacture are your RNase/DNase/RNA/DNA decontaminating solutions, such as RNase away (100-400$/L) and RNaseZAP (250$/L). Why not make your own for less than 20$/L? (Up to date recipe at bottom of post)
You use these solutions by spraying them on your bench, pipettes, what have you, wait a bit and in theory the surfaces should be nuclease free. Let’s not debase ourselves by discussing whether you really NEED to do this, plenty of labs and researchers get by just fine by following good RNA practices, yada yada, have a cookie. Some people still buy the stuff to assuage their paranoia and you know what? That’s okay too! We all have our superstitions. But let’s try to save a buck, eh?
My usual go-to for reverse engineering recipes are patents. You have to navigate some legalese but there’s definitely lot’s of good hints to be gleaned and even figures sometimes! Oh my! I searched pretty thoroughly for patents on RNase away and RNaseZap and came up empty handed. If they patented these solutions then the jig would be up, so I can understand why they didn’t. MSDSs are the next best source for a recipe, however something to keep in mind is that the laws vary by locality and time as to what the manufacturer HAS to disclose is in their product. For RNase Away several SDSs suggest that RNase away is nothing more than 0.5-1% NaOH and water. Due to it’s foamy nature though, I highly suspect that the other ingredient in RNase away is likely 5% SDS or sodium lauryl sulfate (SLS). So, SDS to denature proteins and breakup aggregates, and NaOH to hydrolyze proteins? Possible, but the NaOH concentration seems a bit low for that purpose, so if that is the recipe it likely doesn’t work great. RNaseZap MSDS revealed more recipe information. RNaseZap is composed of 1-5% SDS (5% probably), Sodium dichloroisocyanurate (0.5-1%) and (took a bit of digging) NaOH (?%, SDS states pH 7-8.5). Sodium dichloroisocyanurate is the interesting ingredient here, which in water will form hypochlorus acid (bleaching agent) that will destroy proteins, DNA and RNA.
I could’ve probably stopped there and been happy, but after a bit more digging found that Perkin Elmer manufactures AbSolve glassware cleaner/RNase decontamination reagent. No patent, but the SDS is very descriptive disclosing NaOH (2.5-10%), sodium hypochlorite (bleach, 2.5-10%), “alkylsulfonate steol” (<=2.5%), potassium citrate (2.5-10%) and 1-Ethyl-2-pyrrolidone (<=2.5%). The bleach, like before, destroys proteins and nucleic acids, the “steol” seems to be a trade name for SLS, and 1-Ethyl-2-pyrrolidone appears to help dissolve stuff…more better. I think the citrate could be used as a chelating agent to stabilize the bleach, or perhaps to balance pH.
Sometimes you strike reverse engineering gold. Qaigen filed a patent recently for “Permanent inactivation of nucleases “. Essentially their mixture is a simple solution of 5% SDS, 2% H2O2 and 6 mM EDTA, pH’d to 4.0 (pH’d with peracetic acid?). They likely imitated an older patent by the University of Montreal, “Synergistic detergent and disinfectant combinations for decontaminating biofilm-coated surfaces”, which used 1% EDTA, 5% H2O2, 1-2% SDS, 0.1% cetylpyridinium chloride and 1% peracetic acid. The Qaigen patent goes into great detail as to the rationale for the ingredients as well as some lovely gel images! Yay!
There you go, stop buying that over-priced stuff and make it yourself.
Update: Here’s the recipe I’ve settled on. Check out Just Bleach It for the rationale behind it. I use it on basically any surface I want to clean/decontaminate, watch out for your eyes though! It’ll burn them clear out of your eyeholes!
Decontamination Solution (v1.4)
10% Store bought bleach (2L per 20L)
1% NaOH (200g per 20L)
1% Sparkleen or similar powdered detergent (200g per 20L)
Instructions for use:
- For most applications (Wiping down countertops, equipment, pipettes) the decon solution can be diluted 2-3X, wiped on, allowed to soak for several minutes and then rinsed off with distilled water and towels. More stubborn messes can be hit with undiluted mix.
- Glass and parts can be set to soak in straight or diluted (2-3X) decon mix, then washed as normal and rinsed with distilled water.
- Don’t let the decon mix come in contact with anodized aluminum, it will take the color right off!
Changelog:
- Latest, easiest decon mix. Dropped the addition of sodium bicarb since sparkleen contains a good bit of it and it provides the detergent for the wetting action. Thank you to the reader who suggested it!
Decontamination Solution (v1.3) (Old, legacy version)
10-15% Store bought bleach (100-150 mL/L)
1% NaOH (10 g/L)
1% Alconox/Sparkleen/dish soap (10 g/L) *
90 mM sodium bicarbonate (7.5 g/L) **
* Commercial versions use SDS, but at higher concentrations (=>1%) the SDS will tend to crash out. Unless you have the 2141-BG fragrance/emulsifier, either use a lower concentration of SDS (<0.1%) or use the above detergents *
** Sparkleen and Alconox have sodium bicarbonate already in it in high concentrations, up to ~40% for Alconox, so the addition of bicarbonate may not be necessary.
Assuming Sparkleen has 30% bicarbonate, 10 grams of sparkleen has 3 grams bicarbonate, which would make a final solution that has 36 mM bicarbonate, which could still provide corrosion inhibition, depends how strongly you want to believe the 90 mM from the DNAzap patent. **
*** This decon mix will corrode aluminum and iron/cheap stainless steel at high concentrations and when treating for long periods of time. Good quality stainless hold up fine. ***
**** This mix is awesome for cleaning glassware, let a beaker sit in 0.5X or 1X decon mix for a while, the longer the better. It will sparkle after you rinse it! The high NaOH content is reminiscent of base baths used by chemists to etch a nice clean layer on their glass. ****
For more background on how I arrived at this recipe check out my second post on the topic, Just bleach it
Good post. I absolutely love this site. Thanks!
Great post. Just wonder whether there is any way to test the efficiency of the mix.
Thank you 🙂 The rigorous way would be to treat DNA, RNA, and protein (DNA could be plasmid, maybe some RNA from the back of the fridge, BSA?) with the solution and compare it to the control . The patent is interesting as it goes into the intricacies of testing efficacy…stuff like, how soiled is the surface? how long do you leave it on the surface? I’m gonna post a little article about another cleaning solution in a bit, It’ll include link to a patent that does a LOT of efficiency testing.
Correct me if my idea or concept is wrong… But I plan to verify real world situations by smearing three areas of my bench with my pure RNase solution and allow it to dry. The first area is the no decon area, the second is Pipette Jockey’s solution and third a proprietary solution. Then I will use a wet swab and wipe it across the test areas and place it my known RNA sample. To save time I will just verify the effectiveness by comparing the conc. and quality of the RNA by nanodrop using the 280/260 260/230 modes… I once did this to verify the impurity of a colleague’s DNase… Of course she bought a normal vial of DNase… Not the RNase free one…
It’s an awesome idea, using the nanodrop for getting an estimate of RNA integrity cuts out running some gels, which is nice. The only gotcha in this situation is that the decon solution will itself degrade nucleic acids on contact. So while the decon solution will degrade the RNases, the RNA will still be degraded by the decon solution, giving you a false negative. You may have to take your swab, soak it, and use a dilution of that on your RNA, such that the decon solution is at a low enough concentration.
I thought that nanodrop only measures the concentration of RNA, but not the integrity of the RNA. Can it be used for checking the quality of RNA?
You’re right… Nanodrop does not check the “integrity” of RNA… A denaturing electrophoresis is the basic way to go… But I use it to compare a RNase affected sample to a sample with known concentration of RNA and see if there’s a reduction…. But then again a gel run or a bioanalyzer is the definitive and proper way to go to check for RNA quality
I think bleach doesn’t destroy nucleic acids but rather chlorinate them and make them undetectable during PCR and any enzymatic reaction
i fuckin love this site.
A fluorescent nuclease substrate kit (Ambion RNase Alert, for example) could be used as the basis of an experiment to determine efficacy.
One (maybe dumb) question. Would you autoclave the solution, for example the 5% SDS, 2% H2O2 and 6 mM EDTA, pH’d to 4.0, before adding the H2O2??
As long as you control for the SDS boiling a bit then you should be okay autoclaving that. Guess the question would be why would you want a sterile decon solution when it should (theoretically) be sterile when you mix it? Assuming nothing decomposed then it probably won’t hurt though.
If possible, use a mix of 1% NaOH, 1% SDS (or any detergent really), 80 mM sodium bicarbonate and 10-15% bleach. I covered that in my most recent decon solution post. That stuff does not need to be sterilized before use because quite frankly it will burn your eyes right out of your eyeholes, not a problem (goggles are a good idea). I’ve got a 50 liter bottle full of it, throw it on pretty much everything, makes glassware shine!
Thanks for the recipe Pipette Jockey… I have prepared your solution, but what concerns me is that the 10% to 15% bleach in the solution is unstable. Normally diluted bleach 10% is effective for only 24 hrs. Therefore, I was wondering if the sodium bicarbonate and NaOH in the recipe helps to maintain stability (I have read literature that sodium hypochlorite is best stabilized by alkaline conditions but at a pH of 11!). Or would you advise that we replenish the bleach component daily (or if alkalinic) weekly? Thanks for your insight…because you are saving us much needed cash!
P.S. It’s also great that you pointed out that RNAseZap may use sodium dichloroisocyanurate as an ingredient… And as far as I know sodium dichloroisocyanurate is much more stable than sodium hypochlorite because it decomposes much slower and is primarily affected by exposure to air. So this may explain the longevity of these solutions as well as the less pungent odor of said products. 🙂
With 1% NaOH you should be well above pH 11 in your solution. Check out this paper “Stability and Bactericidal Activity of Chlorine Solutions”, looks like 20% bleach will stay good for more than a month if diluted with just tap water and stored in closed bottles. 10-15% bleach will have a slightly lower pH but we’re compensating with the NaOH + bicarbonate. Also, very concentrated sodium hypochlorate solutions decompose rapidly, but we’re not approaching that level here. I usually make a batch of 25 liters at a time and go through that in a month pretty quickly.
Thanks for the advice.. You’re right; the solution is relatively caustic… Haven’t checked the pH… But wouldn’t be surprised if it’s pH 11. Oh…Ive read that some would use 70% alcohol to wipe away the residual bleach decon solution… But then chemically it creates chloroform… Not sure if the amounts are toxic enough but I’m thinking of using plain old ddH20…
Pardon my ignorance, but when you say 10-15% bleach do you mean already diluted 10% bleach or that of the total decontamination solution 10-15% should be bleach (i.e., much stronger starting concentration of bleach)? If the higher concentration, where can you get it in such a high concentration and/or not already in solution?
Thanks!
No worries about ignorance, we all put our pants on one leg at a time, I’ll try to make it more clear in the post.
When I say 10-15% bleach I mean 10-15% of your decontamination solution should be store bought bleach. Since household bleach is 6-8% sodium hypochlorite, the final concentration in your decon mix will be somewhere in the neighborhood of 0.6% to ~1%ish. You could in theory buy sodium hypochlorite from sigma, but concentrated bleach degrades significantly faster than dilute bleach, not worth the hassle.
Hi!
Can you suggest some stabilizers for these decontamination solution to increase the shelf-life??
Hmm,good question, I’d keep the bottle tightly capped and away from the light if possible. Beyond that I’d look into the literature for stabilizing bleach, it’s the ingress most likely to lose strength over time. The very basic conditions should help.
I feel a little silly for asking this, but when I attempted to make the decontamination solution, I keep getting a wispy precipitate that clouds up the whole solution. I have a feeling that I may have added the SDS improperly – I interpreted 1% SDS as 10 grams of SDS powder per liter of decontamination solution – so if that is incorrect, how should I fix my mistake?
Hmm, how much solution are you making? I would dissolve the SDS and other powder components in a glass beaker with stirring before adding it to the larger volume. Are you getting a white precipitate after you combine everything? I’m wondering if it’s a problem of not enough mixing or is the sds coming out of solution?
Alternatively, substitute the SDS for another detergent like sparkleen or regular dishwashing soap. I favor those because having airborne SDS isn’t the nicest thing in the world.
I ended up with a liter of solution, and here’s what I put in:
100 ml of bleach
10 g of sodium hydroxide
10 g of SDS
6.72 g of sodium bicarbonate
Add the above to a small volume of water to stir and dissolve, then filling bottle up to the 1-liter mark
When I first made the solution and kept it stirring for a few hours, everything dissolved as expected, but overnight a clear-ish precipitate fell out of solution (which I *think* is the SDS). Upon attempting to stir the solution again, I ended up with little more than a snow globe. If something’s wrong with my amounts, that’s easy enough to fix, but if not, I think I’ll swap out SDS for Alconox.
Yeah, sounds like the SDS crashing out. The original recipe uses a fragrance/emulsifier that also keeps the SDS in solution better, 2141-BG, but frankly it’s probably easier just to use Alconox as you said, sparkleen, dish soap vs trying to optimize the recipe. Really the detergent is there to help solubilize organic material so the bleach and NaOH can do its job, so it’s not critical which one.
On my end I’m sorry about the precipitation in your solution, I think the 1% SDS is too ambitious for the recipe. I’ll replicate your results and then try 0.1% and 0.015% SDS, which is more in line with the 2-part RNaseZap solution. In the meantime I’ll add a warning about SDS crashing out in the recipe. Thank you for your input! 🙂
I’ve got the same as Alex. I mixed NaOH pellet, sds and sodium bicarbonate with distilled water, it formed clear solution. But when added with 150ml sodium hypochlorite@ bleach it became cloudy. Hope you can advise me on this? Thank you
I make up 20L at a time and I also notice that at the end of the barrel I get some crystals, but nothing major. I usually dilute the mix 1:3 or so before using it, and it dissolves any crystals and cleans well.
If the precipitate bothers you, you can try dialing back the amount of sodium bicarbonate, that’s where I would start first.
Ideally we would figure out a substitute for the emulsifier used in the original decon mix.
One comment on chemistry: 1% NaOH and 80 mM sodium bicarbonate will react to produce sodium carbonate and water. Actually, it will reduce effective NaOH concentration. At the same time, Fisherbrand Sparkleen-1 detergent already have 10-25 % of sodium carbonate in it (don’t know about other detergents). So I guess sodium bicarbonate can be omitted, if you use Sparkleen-1 detergent.
Hey! Thanks for the chemistry input, I appreciate it. I’ll add the bit about the Sparkleen having bicarbonate in it. I would caution against removing bicarbonate from the solution altogether, the patent calls for 90 mM bicarbonate to inhibit corrosion. Could you get by with just whats in a good helping of Sparkleen? I’m thinking probably, but I’m having a hard time finding any literature on the corrosion inhibition effects of bicarbonate, so I’m not sure how low can you go. There’s a few patents for some light reading, if you’re so inclined:
https://www.google.com/patents/US8105531
https://www.google.com/patents/US20080300160
Fantastic article as usual! While we’re still talking about bleach, take a look at this article that makes denaturing agarose gels super easy! Forget about expensive and toxic buffers, just bleach it!
https://www.ncbi.nlm.nih.gov/pubmed/22222980
Well, I’ll be damned, who knew you could add bleach to agarose gels? And up to 5% too, wow. Thanks for letting me know about this! 🙂 I’ll have to give this a go, I always wondered just how much the smearing of RNA of the gel had to do with RNases in the TAE/agarose.
Hi there, thanks for all of the work you put into this. Probably a bit of a stupid question but for the household bleach do you think there would be any difference using thick bleach instead of standard thin bleach. I would like to try this decontamination solution myself but I can only find thick bleach. I assume it should be fine, but just wanted a second opinion.
Also what would you say would be the rough shelf-life of this stuff?
Thanks!
Not a problem, glad you enjoyed the read!
Yeah, assuming your thick bleach has a hypochlorite concentration between 5-6% (Or even more), then it should be functionally the same, you’re diluting it a fair bit.
As for shelf life, your Biosafety officer may throw out a value like one month, but I’d say that unless you have it in direct sunlight it should last for 3-6 months pretty easily. Hypochlorite is stabilized by the basic conditions of the mix so it’s not really going to degrade that fast on you. At any rate, the stuff will bleach your clothes and solublize skin months after it’s made. We tend to go through 20L of this stuff every month or two, so it works out.
When I get a bit of a breather from my PhD I will be doing some testing on the actual decontamination activity of the mix as a third party validation, there’s a lot of cool protocols to follow to validate decon mixes!
Hey thanks for the recepie.
Just wanted to know if it will work as efficient as RNAase zap?
I think RNase zap will be gentler, this stuff will strip glass pretty easily, so it’s probably overkill as it is. If you wanted RNase zap then you could follow the patent.
I followed the recipe for the RNAse zap and 1) it’s super viscous with the 5% SDS and 2) very opaque. Is it supposed to be translucent?
Use 1% Sparkleen/detergent or 0.1% SDS…yeah, 5% will start precipitating out. Even 1% SDS is a bit much. The commercial stuff has an emulsifier in it which helps keep stuff in solution.
Pipette Jockey I am extremely happy to find this space so interesting since I try to be very cost-effective in some plastics for laboratory, my question is this: if I want to wash a plastic box to make analysis of quantification of DNA and RNA and reuse it Sometimes I want to use this mix I put it on the surface of the box I let it act a few minutes and then I must rinse the box with miliQ water? or just do not need to rinse it ?, finally I have to worry about the DNAses and the RNAses after using my mix or are they already eliminated? thanks!
Hey! I’m glad you enjoy the content! Yes, I frequently use the decon mix to sterilize plastics, almost everything that can take it, even electroporation cuvettes (albeit for short bursts). Leave the mix on for at least 5-10 minutes.
Yes, you want to wash out the container with milliQ water and let it dry, residual decon will degrade your nucleic acids if it gets into your reaction.
The mix should do a very good job of degrading proteins, it can strip glass of organic matter! It’s reminiscent of base baths that chemists use. However, my claims of effectiveness vs RNases etc are based on patents of cleaning solutions, I haven’t done an in depth study of my mix.
I will be testing the effectiveness of the mix in future articles, testing decon mixes is a complicated subject by itself.
Hi! Very good post! I was wondering, which concentration is your Store bought bleach? Here in Argentina it comes 25 g/L or 54 g/L. Which should I buy? Thanks! Analía
Hello Analia, glad you liked it! Purchase the 54g/L stuff, that’s what we would call store bought here (~5% sodium hypochlorite). Stronger flavors exist, up to 8% or so, sold under “concentrated” bleach.
Pipette Jockey thank you for your good and helpful information!
I would like to make your decontamination solution using dish soap. Here in Portugal de dish soap is liquid so I was wondering wich volume do you think I could try to make 1 liter solution?
Howdy! Yeah, I’ve used dish soap before, I’d probably use it at about 1% concentration. You only need it to break the surface tension of the liquid and help the decon mix flow better.
Hey Alex,
Thanks for the latest update on v1.4, without the Sodium Bicarb.
Love the mix and they are good at cleaning lab benches for sure, sparkly clean! 😀
As a side note, the smell of the decon mix definitely smell very similar to RNaseZap or RNaseAway but way more corrosive.
Ryan,
No prob bro, there’s a video or post coming about testing the decon mix myself, long term goal is showing degradation of DNA, RNA and protein. Right now I’m testing to see what the decon can do to a dried spot of bacteria, seeing if I can pick up any DNA after with random primers. There’s a bit of literature about standards of testing the effectiveness of cleaning solutions…that’s what I’m trying to replicate.
Yeah, lab coat will protect your clothes from being ruined, then eye protection is a must. When you’re making the stuff a face shield or atleast splash proof goggles is a very good idea, you don’t want it in your eye.
I usually dilute the decon mix to a 1:3 or 1:4 dilution for everyday cleaning. I soak glassware for a few hours and they sparkle.
Sparkle cuz of Sparkleen !!
Would this decontamination solution (ver 1.4 or 1.3) be suitable to cleaning pipettors (basically the outside)? I’m thinking it would be too harsh on the plastic. What would you say?
Thanks!
We end up with a white residue on EVERYTHING. So we need to follow the spray down with a milliQ wash too?
You can reduce the white residue somewhat by using a diluted (2-3x) version of the mix when wiping down surfaces, however you will still get a whiteish haze. Normally you want to follow up with a wipe down with some distilled water, Milli-Q if you’re feeling fancy. I’ll put up some more explicit instructions for use, I don’t want anyone having less than pristine work surfaces if your local friendly public health agency comes by 🙂
Can alcanox be used instead of Sparkleen? Can it be used on plastic materials? I am trying to use this to decontaminate 384 well plate covers (EZCover 384). Does this need any adjustments for this specific purpose?
Alconox is a perfect substitute, and the mix is safe on plastics provided you dont leave them for days at a time. I use it to decon 96 well black plates for fluorescence readings, works a treat. Worth testing on one plate before bathing your entire labs stock…just in case 🙂
I tend to wipe the equipment with 70% ethanol after wiping with RNaseAWAY (air-dried), will there be any chemical reactions between the ingredients of RNaseAWAY and ethanol that will affect DNA/RNA reactions? Thanks!
I’m in a microbial ecology lab and without explaining too much, I’m re-using tubes that haven’t had samples in them but I wouldn’t say are 100% DNA free either. I’d soak them in bleach, but would you recommend auto-claving after? Or would that just re-introduce nucleic acid through the steam in the autoclave (there are always water droplets in my tubes after auto-claving)? Or should I just leave them to dry after washing residues off with MilliQ water in a fume hood with the UV light on?
Anyway, thank you for this, you’re truly doing some revolutionary work.
In theory some decon and washing with milli q should do the business, autoclave if you’re feeling fancy. I know your fear about steam depositing salts/nucleic acids on stuff you autoclave but I would say thats unlikely, I’m not sure if DNA would be carried around in steam like that. Then again, I’m also paranoid about autoclaving media and tips together. Autoclaving 1.5ml tubes is commonly done, so I think youll be okay. Good luck!
I was looking at the Sparkleen SDS and it mentions that one of the incompatible materials is oxidizing agents, and there are some powder detergents (i.e. Haemo-Sol) that explicitly call out sodium hypochlorite. With that in mind, how safe is it to mix Sparkleen or other detergents with bleach?
So far I haven’t noticed any sort of reaction with Sparkleen or Alconox, those are the two I’ve tried.