Published: November 1st, 2016 Last Modified: September 18th, 2020
Earlier I made a post about decontaminating solutions and how to homebrew them for cheap. I had one more patent to talk about but I felt kinda burnt out (and a bit guilty that I was bullying the decontaminating solution manufacturers association (DSMA) so much ), I ended up letting it sit for a bit. But this horse isn’t quite dead yet, as I think there’s some very useful information to be gleaned by examining the solution known as DNA Zap, from Thermo.
It’s a bit of an oddity, a TWO part cleaning solution meant to destroy nucleic acids, which you apply to an area one after another. Pretty…how you say, le fancy, non? Taking a peek at the MSDS will tell you that Part A contains copper sulfate, which I hadn’t encountered in decontaminating solutions before. WHY??? WHY DO YOU NEED COPPER SULFATE??? What magic is in the mix that can possibly improve on your standard soapy/bleachy/NaOH-y goodness?
Go read the patent, titled “METHOD OF MAKING A FORMULATION FOR DEACTIVATING NUCLEIC ACIDS” it’s fascinating. It’s written in legalese, but the amount of figures/tables/DATA proving the efficacy of this mixture is astounding! Hats off to the people who did all the work, what a nice bit of optimizing 🙂 It’s a pretty neat little system they’ve thought up.
Part A consists of copper sulfate (2 mM) and hydrogen peroxide (3%). This solution in and of itself will destroy nucleic acids on contact. They do not comment on the mechanism on which this works, though. Interestingly, the peroxide based inactivation has similarities to patents by Qiagen and the University of Montreal, which we covered in the last article.
Part B is a pH 9.3 solution of 0.6% hypochlorite (so basically 10% bleach? the authors used CLOROX which is ~6%), 90 mM Sodium Bicarbonate, 0.015% SDS and 0.0075% 2141-BG which is some weird proprietary fragrance which helps with solubility AND smells good? This is more in line with the classic RNase Away solutions. One important improvement to RNase-away-like mixes is sodium bicarbonate, which inhibits the corrosiveness of the bleach on lab equipment. Having watched bleach corrode the crap out of certain materials, the corrosion inhibition seems like a good call. The SDS and fragrance are wetting and emulsifying agents, respectively. The SDS needs the emulsifying agent to stay in solution. Both help the bleach/bicarbonate to make good contact with whatever mess you made (Fiillllthy, FILLLTTTHHHYYYYYYY!!!). The author notes that the bleach/bicarb solution itself is enough to destroy nucleic acids if you don’t have a high organic load.
So, why the two reagents together? Well, one reason is if you absolutely, positively HAVE to have 100% degradation of nucleic acid on the surface. If one of your reagents is diluted or absorbed by your mess, the second reagent acts as backup to ensure you have complete nucleic acid degradation. The dual action also seems to speed the effect of nucleic acid degradation, requiring no soak period. Wax on, wax off.
The inventors actually had labs compare the efficacy of 50% bleach (which is allegedly used in medical assays) VS Part A + Part B VS Only Part B. Turns out that while Part A + B are 100% effective in destroying nucleic acids, Part B by itself is still 99.3%. Now, how effective was 50% household bleach? 99.1%!!! They even did statistical tests to show that there is no difference in efficacy between the two part formulation and 50% bleach with a P value = 0.11.
So what can we take away from all this?
-50% bleach will destroy all nucleic acids on a surface within a minute
-bleach/bicarbonate pH 9.3 is equally effective, and is much less corrosive
– Adding SDS/emulsifier helps the already potent solution clean MORE BETTER by helping make better contact and removing organic load (Shmoo).
– Using a fragrance to hide the scent of bleach is a good idea, especially when that fragrance also keeps the SDS in solution
-If you must have all nucleic acids dead, DEAD, a solution of peroxide and copper will finish the job that your bleach didn’t.
– The authors don’t mention RNase activation, but looking at the previous patents this mixture would likely destroy most nucleases
So yeah, just throw some bleach on it, no worries.
Update: Here’s the decontamination solution recipe I’ve settled on. 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)
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. ****
Thanks Pipette Jockey! My advisor said something to the same effect, about just throwing bleach on it, but it’s always nice to have another source. You have saved me lots of money, stress, and time!
I use a spray bottle too, but I just refill it from the keg, it also allows me to flush hoses and anything else without any problems.
Because, before its invention, lab protocol called for scientists to pipette with their mouths, much like you would a straw.
I use a spray bottle too, but I just refill it from the keg, it also allows me to flush hoses and anything else without any problems.
Fantastic post! Thank you for that. I am looking for a cheap solution to make used 1.5 mL tubes nuclease-free. I would imagine soaking them in this solution, followed by a rinse in Millipore water and autoclaving should do the trick.
Hey! Glad you enjoyed it! As for sterilizing 1.5 mL tubes, the solution would certainly do the business, you would likely not even have to autoclave, just wash thoroughly with millipore water. If you did autoclave, I wouldn’t do it more than 1 or 2 times, the plastic degrades and gets foggy/brittle and likely leaches more into your solutions.
Now, I’m not sure what your specific reason to sterilize 1.5 mL tubes is, but if it’s in terms of cost savings factor in the time to do the decontamination/washing/autoclaving etc (Those tubes are wee, and if you’re doing it in bulk how will you make sure every one is deconned/washed out). Whether it’s worth it depend on how much labor you have and how much you pay for it vs. how much you can get a box of sterile tubes for.
We wash/autoclave 15 mL and 50 mL centrifuge tubes and then use them for applications that don’t require absolute sterility or for balancers etc.
Quick question re: this mix.
Sodium hypochlorite is a strong base / weak acid salt and so the pH of bleach is already ~12.
Bicarbonate buffers the bleach do to a pH around its pKA ~10.
Why the need for NaOH? Is that to bring the pH up to a higher level?
Obviously you can’t eliminate all the corrosion since bleach is still a strong oxidizing agent, but I wonder if leaving out the NaOH might help some.
It’s a good question! You’re probably right that having such a basic solution isn’t necessary for RNase inactivation, considering that the RNAzap patent doesn’t mention using a solution as basic as my recipe.
The recipe I have right now is an amalgamation of MSDS’s and patents that I’ve read, and the closest commercial product would be Perkin Elmer’s AbSolve glassware cleaner, which is bleach, NaOH, soap, and a few other ingredients. Basic enough that it can strip a few molecules of glass and anything stuck to the glass. Essentially the nuclear option of decontamination, I’m not sure there’s many proteins out there that can withstand bleach, soap and extremely alkaline conditions.
I think you could take out some, if not all of the NaOH, however it may not clean as aggressively.
When I get a bit of a breather from my PhD I will be testing the efficacy of the mix, just so I can give a bit more proof that the mix works other than “Just trust me, bro”. Turns out that testing decontamination mixes is a pretty interesting subject by itself. Most standards use small stainless disks with a small amount of contaminants deposited on its surface, so I’ll be doing something of that nature and I will absolutely test an NaOH free mix.
Thank you so much for all your hard work to save us all some money! You said that the solution will corrode aluminum and iron/cheap stainless steel at high concentrations and when treating for long periods of time, but good quality stainless hold up fine. Is it okay to use on pipettes? I’m wondering if the metal tip ejector part would corrode, or if it’s considered “good quality stainless”?
Thank you for the kind words, makes me happy to help out 🙂 Yes, it’s fine on the metal part of the pipette, our workhorse is the Gilson Pipetteman and I just wipe it down with decon mix on a towel or kimwipe. I also cleanse the inside of the plastic tip, you can leave that part submerged in the mix for a while, then I rinse with water and ethanol. I would avoid soaking the plunger/o-ring/spring assembly though, I do a quick wipe of the metal shaft (without the o-ring) with decon mix, then wipe it away with 75% ethanol, no corrosion so far.
Hello, thank you for your great work! They are amazing! Can I ask if there are any relevant literature for DNAZap? Thank you!
Sure, my recipe is based in part on the following patent: http://www.google.ca/patents/US8765652
Also, there’s some details to be gleaned from the msds: https://research.amnh.org/molecular/msds_folder/dnazap.pdf
Great post! Thank you very much. Do you think it’s OK to wash my micropipettes using this solution? We usually use “just bleach” (actually a 1% bleach solution) on all removable parts, then wash THOROUGHLY, with water (dd) and finally with EtOH 70%; but having a sort of “soapy water” along with the bleach and the bicarbonate sounds like a better approach.
Yep, absolutely! I use it on our Gilsons and Eppendorfs no problem. The only thing to watch out for is to not let the decon solution touch any aluminum, anodized or otherwise…it’ll strip it to bare aluminum real quick.
So yeah, wipe down the body of the pipette with decon followed by water/etoh. Soak the easily detachable plastic bits with decon and make sure its washed away with good water and ethanol after. Metal bits (pistons that come in contact with o rings and seals) I usually wipe down with decon and wash it clean and dry fairly quickly to avoid corrosion. Exact protocol will vary on your pipettes…some of the newer eppendorfs have grease on the inside, not very user serviceable, like fine clockwork watches you know?
Any one used pfusso7d for site directed mutagenesis . If any one used please share the protocol
You can use almost any high fidelity enzyme for SDM, really the challenge is to generate the PCR product. After that I use the following protocol, it’s essentially NEB’s KLD mastermix. Grabbed the recipe from reddit posted by username msr2009:
1ul PCR product
1ul 10X T4 DNA ligase buffer
1ul T4 PNK
1ul T4 DNA ligase
1ul DpnI
5ul H2O
1 hour at RT, transform
Works awesome! Been using it for years now.
Thank you for reply,
Any simple method for determine enzyme unit activity per mg . All commercial companies are using radiolabeled, not possible in academic and non profit organization . What is difference between unit activity and specific activity per mg
Hi sorry for the potentially dumb question but do you just dilute the decon in water?
Not a bad question at all. For best results I think deionized/distilled is best, from my reading hard water can speed the decomposition of the bleach.