Gel electrophoresis safety question: measuring deep UV exposure from a UV transilluminator
Mackenzie Cowell
--
+1.231.313.9062 / m...@diybio.org / @100ideas
Nathan McCorkle
http://cgi.ebay.com/Absorption-Longpass-Filter-GG400-w-AR-23-X-17-8mm-/160372850952?pt=LH_DefaultDomain_0&hash=item2556f78108#ht_485wt_754
or maybe even better:
http://cgi.ebay.com/Optical-Filter-435LP-10-1mm-dia-X-3mm-thick-/310294327118?pt=LH_DefaultDomain_0&hash=item483efbaf4e#ht_500wt_665
You added what, 5uL per lane? It comes as 10000X, a 3X solution will
work for post-staining alternatively. Generally I let a gel sit for
about 20 minutes in the 3X solution...
> --
> You received this message because you are subscribed to the Google Groups
> "DIYbio" group.
> To post to this group, send email to diy...@googlegroups.com.
> To unsubscribe from this group, send email to
> diybio+un...@googlegroups.com.
> For more options, visit this group at
> http://groups.google.com/group/diybio?hl=en.
>
--
Nathan McCorkle
Rochester Institute of Technology
College of Science, Biotechnology/Bioinformatics
Cory Tobin
phenixresearch.com site. It looks like you should be able to use a
UVB bulb instead. Wikipedia shows UVB as 280-315 which falls right in
the middle of the GelRed excitation.
-Cory
Nathan McCorkle
with a white light bulb)... otherwise a jacket and goggles will work.
Did you mix your DNA with 50% sucrose syrup? Or add bromophenol
tracking dye? Your DNA could be off the gel.
Get some of this to try??:
http://www.invitrogen.com/site/us/en/home/Products-and-Services/Promotions/DNA-Ladder-Sample.html
On Tue, Apr 5, 2011 at 12:44 AM, Mackenzie Cowell <m...@diybio.org> wrote:
Cory Tobin
> http://cgi.ebay.com/Absorption-Longpass-Filter-GG400-w-AR-23-X-17-8mm-/160372850952?pt=LH_DefaultDomain_0&hash=item2556f78108#ht_485wt_754
Even cheaper, acrylic plastic. That stuff absorbs UV.
-Cory
Mackenzie Cowell
Where is your UV-filter? You need a highpass of about 400nm
Simon Quellen Field
--
Mackenzie Cowell
Mackenzie Cowell
Cory Tobin
Ethidium bromide gels get picked up by university's hazardous waste
folks. Other gels go in the trash.
> Is it possible to melt them down and re-use them?
Yes. But cut out the sections with the loading dye otherwise your
second gel will be colored. Also, I only do it twice in a row. After
that it doesn't work nearly as well. I think due to the loss of water
each time I nuke it, but I don't know for sure.
-Cory
Cathal Garvey
The UVC is probably not any good for your eyes, so get some uv-filtering glasses or goggles pronto. Expect a transilluminator to output more uv close up than strong sun exposure, I've even seen people tweet about sunburn from overexposure during band excision.
Ideally you could avoid uv entirely, as it will dice up your DNA. At maker faire we experimented with Crystal Violet and Methyl Orange as stains, with good first-try results. There's a paywall protected paper allegedly detailing a more effective method with Crystal Violet, so hopefully I will be able to try it out again soon and report. The advantage is that the dye is visible without uv transillumination and isn't very hazardous or expensive. DIYbio win if it works!
jlund256
> I've become a bit paranoid about UV-C radiation after conversations with
> various biologists and hobbyists, so I've build an ad-hoc box from pink
feedback from UV exposure--sunburn. If you aren't getting any sunburn
you are OK. I've worked in the glow of UV boxes for minutes at a time
cutting out bands with no ill effects, all my sunburn has come the
natural way. :)
Jim Lund
Nathan McCorkle
> Also, what do you guys do with your used gels? I typically run 0.8%-1.5%
> agarose gels and just throw them away after one run (or two, if I have
> unused lanes I can use for a second run).
> Is it possible to melt them down and re-use them?
> On Tue, Apr 5, 2011 at 1:29 AM, Mackenzie Cowell <m...@diybio.org> wrote:
There might be a way to precip the agarose in solution using some
other solvent, but I'm not sure... maybe after your next gel is ready
to discard, liquify it and add some 91% isopropanol... if agarose is
insoluble in the alcohol, the alcohol should absorb some of the water
and then form two layers once saturated.
--
Mac Cowell
Nathan McCorkle
the used up buffer and other junk that may have accumulated...
http://www.amazon.com/Nasco-Dialysis-Membrane-Tubing-10/dp/B001DBH42C
you'd have to clamp/tie the tubing on one end, pour in liquid agarose,
tie/clamp the other end, and place in warm warm (enough to keep
agarose liquid) overnight with a stir-bar to mix it (or similar
setup).
Then you could evaporate off the water, to get dry powder. If the
agarose isn't damaged by the electrical current (oxidised, reduced,
etc...), this should basically allow recycling until transfer losses
accumulate (agarose stuck to inside of dialysis tubing, decreasin your
recovery yield).
--
Cathal Garvey
Cathal Garvey
Cathal Garvey
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 Onlinehttp://research.bmn.com/tto
This article is an extended version of one from 'Illuminating DNA' by Dean Madden
Cathal Garvey
Cathal Garvey
From: Cathal Garvey <cathal...@gmail.com>
Date: 6 April 2011 01:05
Subject: Re: Gel electrophoresis safety question: measuring deep UV exposure from a UV transilluminator
To: diy...@googlegroups.com
Wait, there's life in me yet!
Finally, this might be useful but I haven't read it: http://www.ncbe.reading.ac.uk/NCBE/PROTOCOLS/illuminating.html
That's it for me, sleep beckons.
Ruediger Trojok
since the descriptions I red were quite ok.
Now I am little worried if it is a good solution after reading this
here...
On 6 Apr., 02:07, Cathal Garvey <cathalgar...@gmail.com> wrote:
> Great, can't include either attachment due to group attachment caps. Will
> mail them on privately to anyone interested (default Mac).
>
> ---------- Forwarded message ----------
> From: Cathal Garvey <cathalgar...@gmail.com>
> Date: 6 April 2011 01:05
> Subject: Re: Gel electrophoresis safety question: measuring deep UV exposure
>
> from a UV transilluminator
> To: diy...@googlegroups.com
>
> Wait, there's life in me yet!
>
> <http://www.sciencedirect.com/science?_ob=ArticleURL&_udi=B8JCW-4PNDBS...>And
> to increase sensitivity. Although, that's the protocol we used in Maker
> Faire, and it wasn't that great first-go.. early days yet. Mostly, I'd just
> rather it were an in-gel protocol rather than a stain/destain protocol.
>
> Attached also is a decent picture of the result. To do it justice, it's
> slightly more visible in RL, but not much. Although, illumination was by my
> tablet rather than a proper white-light box or scanner, so perhaps it's an
> unfair representation. It would have been a lot clearer without the loading
> dye too; perhaps it's wise to formulate my own cationic dye so it'll flow up
> rather than down with the DNA?
>
Cathal Garvey
Didn't bother trying, all accounts suggest that it's really poor sensitivity (although resolution of what it does pick up is good) and you can't do in-gel staining so it requires a long stain/destain each time.
For teaching purposes it's great, when you're doing a PCR or miniprep with loads of DNA, and safety is paramount. But for routine use it sucks. That's why I've been exploring crystal violet: I believe a marriage of the high sensitivity methyl-orange costain and the in-gel method could make a great routine gel staining method that anyone can use with minimal issue (as long as they work with pre-prepared methyl orange solution to avoid the small toxicity risk).
Sent from my Phone
www.twitter.com/onetruecathal
www.indiebiotech.com
On 6 Apr 2011 10:36, "Ruediger Trojok" <ruedi...@googlemail.com> wrote:
What were your experiences with methylene blue? I just ordered it,
since the descriptions I red were quite ok.
Now I am little worried if it is a good solution after reading this
here...
On 6 Apr., 02:07, Cathal Garvey <cathalgar...@gmail.com> wrote:
> Great, can't include either attac...
> From: Cathal Garvey <cathalgar...@gmail.com>
> Found that 1996 reference on using Crystal Violet:http://www.sciencedirect.com/science?_ob=ArticleURL&_udi=B8JCW-4PNDBS...
> Date: 6 April 2011 01:05
> Subject: Re: Gel electro...
>
> <http://www.sciencedirect.com/science?_ob=ArticleURL&_udi=B8JCW-4PNDBS...>And
> attached is a paper describing the use of Crystal Violet with Methyl Orange
> to increase sensitiv...
> Also promising is Ethyl violet:http://www.sciencedirect.com/science?_ob=ArticleURL&_udi=B6W9V-4YM7FM...
>
> On 6 April 2011 00:50, Cathal Garvey <cathalgar...@gmail.com> wrote:
>
> > Finally, this might b...
> twitter.com/onetruecathalhttp://www.indiebiotech.com
>
> --
> letters.cunningprojects.com
> twitter.com/onetruecathalhttp://www.indiebiotech.com
--
You received this message because you are subscribed to the Google Groups "DIYbio" group.
To pos...
Matías Gutiérrez
Regarding the weird migration of ladder with Gel Red, the problem
is common. There are two things you can do to get rid of it,
1)Load 5 times less ladder
2)add the gel red to your sample loading buffer and ladder stock
instead of staining the gel.
here is a good protocol made by a lab a while ago
http://malooflab.openwetware.org/loading_dye.html
hope it helps.
Matías
On Apr 6, 8:23 am, Cathal Garvey <cathalgar...@gmail.com> wrote:
> Didn't bother trying, all accounts suggest that it's really poor sensitivity
> (although resolution of what it does pick up is good) and you can't do
> in-gel staining so it requires a long stain/destain each time.
>
> For teaching purposes it's great, when you're doing a PCR or miniprep with
> loads of DNA, and safety is paramount. But for routine use it sucks. That's
> why I've been exploring crystal violet: I believe a marriage of the high
> sensitivity methyl-orange costain and the in-gel method could make a great
> routine gel staining method that anyone can use with minimal issue (as long
> as they work with pre-prepared methyl orange solution to avoid the small
> toxicity risk).
>
> Sent from my Phonewww.twitter.com/onetruecathalwww.indiebiotech.com
>
David Hosfield
Sent from my iPhone
Cathal Garvey
Yea, glycerol is fine up to some concentration, I think 20%. Possibly, if you add too much it'll behave like excess salt and accelerate bands. I reckon if this is so it'll be equally true for ficoll though.
Good pipette technique can go a long way too, so you don't need much glycerol, only enough to make the deposited liquid stay put while you lay more on top.
Sent from my Phone
www.twitter.com/onetruecathal
www.indiebiotech.com
On 6 Apr 2011 17:21, "David Hosfield" <hosf...@gmail.com> wrote:
Could glycerol be used instead of ficol 400?
Sent from my iPhone
On Apr 6, 2011, at 4:58 AM, Matías Gutiérrez <matiasgu...@gmail.com> wrote:
> Hi Evryone,
>
...
Nathan McCorkle
sucrose for Orange-G, and glycerol for bromophenol blue (see the GE
manual I posted a while ago, page 14)
Also, in regards to regenerating Agarose, I see there is an enzyme
that digests agarose, called agarase... so I think the agarose can
actually degrade with time, but purification for a while should be OK
for someone looking to save cash.
> --
> You received this message because you are subscribed to the Google Groups
> "DIYbio" group.
> To post to this group, send email to diy...@googlegroups.com.
> To unsubscribe from this group, send email to
> diybio+un...@googlegroups.com.
> For more options, visit this group at
> http://groups.google.com/group/diybio?hl=en.
>
--
Mackenzie Cowell
Nathan McCorkle
> Folks,
> Thanks for all the incredible feedback. I'm going to do my best to catalog
> it a bit more so that it may perhaps be useful to other folks trying to set
> up gel rigs with cheap and / or old equipment.
> I've run a couple more gels and contemplated the results. The good news is
> that I can electrophorese DNA in my gel and seem to be able to amplify DNA
> with PCR. Tonight I tried amplifying 16s rDNA fragments using standard DNA
> barcoding primers from non-clonal microbial samples I collected from a
> friends cheek and from some 1-month-old chili I found in the back of a
> fridge :)
> The PCR seems to have worked... but my bands are even worse than before.
> While I expected some smearing in the PCR lanes, my ladder also smeared.
> Two thoughts:
> 1) I'm using tap water. Ions and other impurities could be messing up the
> gel somehow. (but how?)
Dude, get some distilled from the store, it will be way better, I'm
pretty sure.... you (or someone in the future) could even try running
4 small gels, one with dH20, one with tap, one with dH20 through a
fresh Brita filter, one with tap through a fresh Brita filter
The paper Cory Tobin was talking about earlier, something about
Borate/Borax formulations, talks about how certain ions can hook up to
the DNA backbone and cause weird conformations that either slow or
speed up the migration... with water you'd get a lot more variety of
ions, so maybe it contributes to a general smear... either than or you
have serious nuclease contamination in your process somewhere. What
kind of stuff are you mixing and pouring agarose with/into? To get rid
of RNases (so most probably DNases) the NEB catalog says baking
glassware dry for 3 hours at some 140-160 C, and for plasticware like
agar setups soak for 10 minutes in Hydrogen Peroxide. What about your
pipettor or tips, are they nuclease free/have you pressure cooked
them/anything? Are you wearing gloves?
> 2) tonight I let my gel cool for perhaps 20 minutes longer than I meant to.
> Its viscosity seemed normal when I poured it, but perhaps the cooling then
> agitation as I poured it prevented it from casting in a totally homogenous
> way.
Meh, that's probably OK, I would have redissolved if it was chunky.
> By the way, I picked up a UV haze filter at a local camera store and have
> been using it. I think I need like 6 more :). The gel in the picture was
> exposed for 2 seconds at f5.6.
So did you try with and without the filter, see an improvement or
stark difference? I linked to that ebay one... could try that as the
bandpass should definitely be above your excitation signal
wavelengths, it seemed cheap enough. What did you pay?
> The 1.5% gel is made from 0.45g of "low EEO" agarose mixed with 30 mL of 1x
> TBE, stained with 3 uL 10,000x GelRed, and run in 300 mL of 1x TBE at 120v.
Sounds good, I think the Biotium GelRed data sheet says 5uL per 50mL
gel, though they recommend post-staining with a 3X solution. I've done
both, and it seems like so far I've saved some gel-red doing the
post-staining, though I'm not counting (but probably because I've been
doing big gels lately with 150mL agarose at 11mm height).
Mackenzie Cowell
To post to this group, send email to diy...@googlegroups.com.
To unsubscribe from this group, send email to diybio+un...@googlegroups.com.
For more options, visit this group at http://groups.google.com/group/diybio?hl=en.
--
Mackenzie Cowell
Where is your UV-filter? You need a highpass of about 400nm
http://cgi.ebay.com/Absorption-Longpass-Filter-GG400-w-AR-23-X-17-8mm-/160372850952?pt=LH_DefaultDomain_0&hash=item2556f78108#ht_485wt_754
or maybe even better:
http://cgi.ebay.com/Optical-Filter-435LP-10-1mm-dia-X-3mm-thick-/310294327118?pt=LH_DefaultDomain_0&hash=item483efbaf4e#ht_500wt_665
You added what, 5uL per lane? It comes as 10000X, a 3X solution will
work for post-staining alternatively. Generally I let a gel sit for
about 20 minutes in the 3X solution...
Mackenzie Cowell
--
You received this message because you are subscribed to the Google Groups "DIYbio" group.
To post to this group, send email to diy...@googlegroups.com.
To unsubscribe from this group, send email to diybio+un...@googlegroups.com.
For more options, visit this group at http://groups.google.com/group/diybio?hl=en.
--
Mackenzie Cowell
Ruediger Trojok
Methylene Blue:
http://bitesizebio.com/articles/ethidium-bromide-the-alternatives/
and http://www.carl-roth.de/media/_en-de/usage/0648.pdf (see 2nd page
for english)
Seems like folks have very different experiences. Could also depend
very much on the gel you are making,
agarose quality, concentration....
I ordered methylene blue anyway and will probably get some crystal
violet as well, soon, so I can come up with some direct comparison.
On 7 Apr., 07:23, Mackenzie Cowell <m...@diybio.org> wrote:
> Last October "Byrne" kindly mailed me a square section of red welding
> curtain. He suggested it might be a good diy post filter for gels. I told
> him whenever I could get software for my old UV/Vis spec (Uvikon XL; needs
> some software "LabPower3000" or "LabPowerJr"), I'd check out it's absorbance
> profile.
>
> Still haven't found the software, but I did just test the sheet in my
> gelbox. It works. Very cool.
>
> .
> > >> > On Apr 5, 2011, at 9:24 AM, Cathal Garvey <cathalgar...@gmail.com>
> > --
> > Nathan McCorkle
> > Rochester Institute of Technology
> > College of Science, Biotechnology/Bioinformatics
>
>
> 2011-04-06_P1030413_run2_with_welding_sheet_Vis.JPG
> 1663KAnzeigenHerunterladen
>
> 2011-04-06_P1030412_run2_with_welding_sheet_UV.JPG
> 970KAnzeigenHerunterladen
Nathan McCorkle
I actually meant 5uL per 50mL gel... so you're right on... the Biotium
folks definitely recommend post-staining, but only because they say
some people notice it changes migration when in the gel... folks here
do it both ways and seems fine (though the folks that say its fine
that I know of are using LB buffer, not TBE, could make a difference).
I would wait to add the gel red until the flask is comfortable to
touch, also I would not recommend putting it in the buffer, it seems
like it would just get reacted with at an electrode.