clonewell electrophoresis
Douglas Ridgway
of the ideas could be applied in DIY gel boxes:
Bryan Bishop
That's certainly a neat trick. But, since this is a logarithmic
effect, pre-specifying an end point would mean that you're sacrificing
accuracy, right? Normally, say you have an infinitely long gel, you
have your DNA running down it in different columns, and as the hours
roll by the distances increase nonlinearly. So if you cut it short
after one minute, that will hardly be the same result as a million
years later. So, can anyone confirm here my suspicions about
invitrogen's "just use a comb near the bottom where you'd like to
collect" ?
Since every gel has a finite length though, you might as well just use
these wells at the end, plus a webcam to track when precisely each
column of stained DNA hits. Some friends and I were talking about this
(or, rather, we're talking about this in the background at the
moment)- some extra ideas would be:
(1) four-directional control of DNA
(2) webcam for visual tracking of stained DNA (we can call it a
DNAlogger, just throw an arduino or PIC at it, right?)
(3) some way of using the refractive index of DNA to your advantage so
as to not have to have a nasty stain on your dna
http://en.wikipedia.org/wiki/Refractive_index
For protein gel electrophoresis + using the refractive index, see:
The instantaneous monitoring of polyacrylamide gels during electrophoresis.
http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=1164177
Real-Time Monitoring of Polyacrylamide Gel Electrophoresis by Schlieren Optics
http://jb.oxfordjournals.org/cgi/content/abstract/102/4/681
("A band containing as little as 0.3 µg of a protein could be detected.")
http://en.wikipedia.org/wiki/Phase_contrast_microscopy
DIY phase contrast (expensive)
http://www.gpmatthews.nildram.co.uk/microscopes/diyphase.html
http://www.nhm.ac.uk/hosted_sites/quekett/Technical/phase.html
"""
Phase contrast set-ups using the Zernicke method can be an expensive
option for the amateur, but home made phase contrast although
potentially cheap, at first seems a daunting prospect. Etching phase
plates accurately using hydrofluoric acid would be technically
difficult and hazardous. In 1953, Prof. Wilska, in a letter to Nature,
described the use of phase plates using a sooted pattern on glass.
This can either be directly on the back lens of the objective, or on a
glass plate, a cover slip for example, mounted close to the objective
focal plane. There is a useful Discussion in The Microscope journal
of 1953 describing the technique. This link also gives access to
transcripts of the correspondence in Nature.
I found that my Watson 30x and 40x parachromats had a ledge behind the
lens in about the right place to mount a phase plate as described in
an article on DIY Phase Contrast on the Quecket Microscopical Club
website, and having seen other correspondence on this technique, but
using a smoked stripe directly on the lens (although this also
referred to the use of a separate plate), decided to have a go. I
tried using a circular cover slip, applying soot from a candle and and
scraping it off to give a stripe, but found this difficult to do in a
controlled fashion. I then tried masking the coverslip with adhesive
tape before sooting, and this worked quite well, although one needs to
select a tape that leaves minimal adhesive residue once removed. The
substage stop was made by masking a transparent disc with PVC tape to
leave a stripe that aligned just inside the phase plate stripe when
the objective back-lens was viewed using a pinhole eyepiece.
"""
"""
I do make phase plates (and modulation plates) and that article was a
little ahead of what I intended to write about, and got sidelined from
(excuse the contorted syntax, but I'm sure you know what I mean!) My
method is not original - it's based on a method that was shown to me
by my old friend and mentor, the late Leslie Martin. Essentially the
phase plate is produced from wax/soot deposited by candle flame on a
coverslip, which is then placed as near as possible to the back focal
plane of the objective. Having found the rear focal plane by the zero
parallax method (you reminded me of this last time we met) I machine a
ledge on the inside of the objective (with great care!) to take a
brass holder to take a standard 6mm, 8mm or 10mm coverslip at the
required distance from the rear lens (of course modern objectives with
the rfp inside the elements are too difficult for the amateur, but a
Zeiss DD and Watson Para both have unscrewable front components to the
barrel and seem suitable). I make several of these holders at one time
so that I can experiment with different types with minimum effort. I
cement the coverslip before smoking. I then make the phase ring by
nipping the brass holder in a split chuck in my Unimat and using a
cocktail stick to wipe away the unwanted wax/soot.
"""
Anyway, a phase-contrast webcam would be fantastic (and allow for
ignoring #4, below). For defractive lenses, you can print out a
spiral-like pattern on your printer and then make the non-printed
portions transparent, and then use this as a defractive lense. But
this isn't the same, though still worth investigating some more.
(4) some way of electrically detecting when DNA hits a well (built
into the bottom plate), such that your gel would block it until you
use your comb to dig into it, such that when the DNA drains into it,
you can have a timestamp (wire it up to a microcontroller).
Finding an alternative to staining would be really, really awesome.
Norman Wang
you have 8 different sized DNAs you have to wait 16 times (fill water
into 2nd well when you wait till DNA reaches reference line, wait
another 1-2 minutes for it to go into well, extracting it). For these
pre-cast gels you'll also need to "pre-run" it for 2 minutes, and top
off all the empty wells with water. Be ready for lots of pipetting and
wasted plastic tips...
Also when your gel runs in a smiley pattern, same sized DNA doesn't
reach the wells at the same time. However, with these gels you won't
get your hands wet in the wet lab. It's $840 for the startup kit.
On Feb 2, 9:50 am, Douglas Ridgway <ridg...@dridgway.com> wrote:
> Here's an interesting commercial electrophoresis system -- maybe some
> of the ideas could be applied in DIY gel boxes:
>
Bryan Bishop
> They may look convenient, the downside is that you have to wait... If
> you have 8 different sized DNAs you have to wait 16 times (fill water
> into 2nd well when you wait till DNA reaches reference line, wait
> another 1-2 minutes for it to go into well, extracting it). For these
> pre-cast gels you'll also need to "pre-run" it for 2 minutes, and top
> off all the empty wells with water. Be ready for lots of pipetting and
> wasted plastic tips...
I wonder though: how much of a difference can you make in the gel
process by changing the general shape and gel cast? For instance, does
anyone have some numbers on what level of accuracy you can get with
gel casting, and can this be used to our advantage in other ways than
just extra wells at the end?
Lora
re-filling wells/tip thing is not the major problem. The iBase unit
does all the run times for you, you just push a button. Nor do the
things run in a bow-shape pattern, although on occasion they don't run
at all because the embedded electrodes in the things do corrode over
time, just sitting in the package.
The problem is that while you very much hope your PCR primers are
going to amplify exactly the right size DNA, they don't *always*.
Especially with the Invitrogen cloning systems, which have these long
goofy overhangs that tend to shrivel up and do wonky things to the
start/end points of your insert. And you can't do anything convenient
like digest or alk-phos polish the ends to neaten them up prior to
ligation, either, the ends are a part of the cloning system. And then
your precious DNA is stuck in the gel and you can't get it back and
you have to start all over.
Even when things work the way they are supposed to, the yields are
still pretty bad. If you have to do something like a sequential
digestion, you won't have enough recoverable DNA at the end of the
second digest to do a good ligation. With a Qiagen kit and homemade
agarose, I get about 0.2mg/ml DNA back, but with the iBase/E-Gel
system it's more like 0.02 mg/ml. Now I only use it for quality
control check, I won't use it for cloning.
Anyway, I vaguely recall seeing a poster somewhere that said DNA
rotated sufficiently detectable amounts of light at 90 degree crossed
polarizing filters, but, hmm, if I was building my dream gel
filtration & visualization unit, there would be Raman spec involved...
> 1 512 203 0507
Bryan Bishop
> Anyway, I vaguely recall seeing a poster somewhere that said DNA
> rotated sufficiently detectable amounts of light at 90 degree crossed
> polarizing filters, but, hmm, if I was building my dream gel
> filtration & visualization unit, there would be Raman spec involved...
I recall the original Raman spectroscopy paper had an easy
do-it-yourself version that could be used. There are also a number of
typical spectroscope designs on the web for using a cereal box and CD
or DVD.
http://heybryan.org/~bbishop/docs/Raman_spectroscopy_1928.pdf
DIY versions:
http://www.cs.cmu.edu/~zhuxj/astro/html/spectrometer.html
http://www.chemguide.co.uk/analysis/masspec/howitworks.html
http://www.kosi.com/raman/resources/tutorial/
http://ioannis.virtualcomposer2000.com/spectroscope/toyspectroscope.html
http://ull.chemistry.uakron.edu/analytical/Mol_spec_equip/
http://www.physicsforums.com/archive/index.php/t-128052.html
http://mass-spec.lsu.edu/mswiki/index.php/Mass_spectrometry
- Bryan
Bryan Bishop
> http://www.cs.cmu.edu/~zhuxj/astro/html/spectrometer.html
> http://www.chemguide.co.uk/analysis/masspec/howitworks.html
> http://www.kosi.com/raman/resources/tutorial/
> http://ioannis.virtualcomposer2000.com/spectroscope/toyspectroscope.html
> http://ull.chemistry.uakron.edu/analytical/Mol_spec_equip/
> http://www.physicsforums.com/archive/index.php/t-128052.html
> http://mass-spec.lsu.edu/mswiki/index.php/Mass_spectrometry
Please excuse me. Some of those are for mass specs, not photospec. My
mistake. The first one still certainly applies.
http://www.cs.cmu.edu/~zhuxj/astro/html/spectrometer.html
http://www.kosi.com/raman/resources/tutorial/
http://www.thespectroscopynet.com/Index.html?/History_1.html
http://www.scienceofspectroscopy.info/edit/index.php?title=Wiki
fenn
buffer for cooling, but not too much buffer or all the electricity
goes through the buffer instead of the gel. What if instead each lane
went in its own capillary tube filled with agar? Each tube would have
its own electrodes at each end. This would raise the current density
tremendously, adding significant heat per unit volume, but also allows
faster electrophoresis rates with less current. So, how about
submerging one of these capillary tubes in ice water and running DNA
and current through it?
The neat thing is once we have confined the DNA to a repeatable
location like a rigid tube (instead of somewhere in a big floppy
rubber thing) we can use inexpensive electronic sensors to measure its
characteristics, such as for example absorbance at 260nm, and this
means we dont have to resort to nasty tricks like phase interference
or polarization. (but we could if we wanted)
I am picturing a little device, basically two heatsinks with a groove
down the middle with that you clamp over a UV-transparent plastic
tube about 3mm in diameter, upon clamping the ends are cut off by two
electrode-blades which simultaneously seal the ends. Somewhere near
the opposite end of the tube DNA was loaded into is a UV LED and
appropriate photodiode. A small microcontroller watches the output and
plots absorbance vs time (vs current) and spits out a length spectrum
over USB. It could be programmed to pause the gel at a certain length,
for extraction. Then you just break off the very end of the tube to
extract the band of interest. I suppose you'd need good control over
the agar composition for repeatability, or maybe you could just throw
a standard length ladder in with the DNA you want to measure and
subtract it out in software.
if someone with access could look at this paper and see what the
absorbance at 400nm is like vs 260nm
http://www.jstor.org/pss/3575456
it would help determining the immediate feasibility of this idea, as
400nm LED's cost about $0.10 - they make LED's in UV-C but after a
quick google it seems that they cost around $200. Perhaps a backpack
water filter UV lamp would work with the appropriate color filters.
oh, and hi, I'm fenn
-ben lipkowitz
> 1 512 203 0507
Meredith L. Patterson
>
> Right now we use big flat rectangular gels which are submerged in
> buffer for cooling, but not too much buffer or all the electricity
> goes through the buffer instead of the gel. What if instead each lane
> went in its own capillary tube filled with agar? Each tube would have
> its own electrodes at each end.
Consider that more viscous substances, e.g. agarose solution, will
require a wider capillary tube than the sort one would normally use
for, say, blood. How about a thin, clear drinking straw? You could use
paperclips for electrodes.
This could also be a really neat trick for doing PAGE, since those
gels are usually run vertically, with a little well dug in the top. A
drinking straw would provide external support for the gel.
Cheers,
--mlp
Meredith L. Patterson
> Then you just break off the very end of the tube to
> extract the band of interest.
Or, with a tube that was flexible like a drinking straw, snip the band
of interest out with scissors and slit the plastic open with a razor
blade.
I love this mailing list so much.
Cheers,
--mlp
Tito Jankowski
a 9v battery.
Tito
Splicer
http://en.wikipedia.org/wiki/Capillary_electrophoresis
-Splicer
> 400nm LED's cost about $0.10 - they make LED's in UV-C but after a
> quick google it seems that they cost around $200. Perhaps a backpack
> water filter UV lamp would work with the appropriate color filters.
>
> oh, and hi, I'm fenn
>
> -ben lipkowitz
>
> On Feb 3, 2:56 pm, Bryan Bishop <kanz...@gmail.com> wrote:
>
>
>
> > On Tue, Feb 3, 2009 at 1:33 PM, Norman Wang wrote:
> > > They may look convenient, the downside is that you have to wait... If
> > > you have 8 different sized DNAs you have to wait 16 times (fill water
> > > into 2nd well when you wait till DNA reaches reference line, wait
> > > another 1-2 minutes for it to go into well, extracting it). For these
> > > pre-cast gels you'll also need to "pre-run" it for 2 minutes, and top
> > > off all the empty wells with water. Be ready for lots of pipetting and
> > > wasted plastic tips...
>
> > I wonder though: how much of a difference can you make in the gel
> > process by changing the general shape and gel cast? For instance, does
> > anyone have some numbers on what level of accuracy you can get with
> > gel casting, and can this be used to our advantage in other ways than
> > just extra wells at the end?
>
> > - Bryanhttp://heybryan.org/
>
> - Visa citerad text -
ben lipkowitz
> That sounds a bit like capillary electrophoresis:
>
> http://en.wikipedia.org/wiki/Capillary_electrophoresis
Not exactly. In capillary electrophoresis you're relying on the molecule's
affinity for the walls of the capillary, so it has to be really small (say
50 microns) and this makes it no good for cloning DNA. Typically you'd
have 20 microliters of reaction solution from your PCR, so this would be a
2.5 meter long band! vs a 2mm band in a 3mm diameter tube. Also by using
standard agarose it should behave the same as in a regular gel, which
makes people feel warm and fuzzy inside.
Aaron Hicks
What was nice was that the sample eluted very "cleanly," allowing a UV LED (?) to screen the samples- very similar to HPLC with a UV detector in that sense. 3D-CE also allowed a broad range of samples- anything that moved with electromotive force, in fact.
-AJ
Meredith L. Patterson
--mlp
Nick Taylor
>> Then you just break off the very end of the tube to
>> extract the band of interest.
>
> of interest out with scissors and slit the plastic open with a razor
> blade.
>
> I love this mailing list so much.
The other day I did a little micro-rant about how great this site is...
http://www.arvindguptatoys.com/toys.html
not because it resembles a cheap way of keeping kids entertained so much as a compendium of ultra-cheap, ultra-low-tech building blocks for alternative technologies.
Drinking-straw electrophoresis belongs in there somewhere I think.
JonathanCline
> I'm gonna try this tonight with a straw, agar, and food coloring, and
> a 9v battery.
>
> Tito
figured that the edge effects would kill the ability of DNA to run
(??). (Otherwise, wouldn't the microfluidics guys already be all over
this?) Anyway, I'm interested in the results, since the current
requirement to run a gel (and hence power supply requirement) is
directly proportional to the gel's physical dimensions, and there
would be significant advantages for cooling.
Also one thing to keep in mind is that electronics technology has far
surpassed original experiments done on agar even in the 90's. 2-D (X-
Y axis) electrophoresis (previously alluded to by Brian as "4-way")
has been done previously, and these days would be very easy to do
(cheap microcontrollers, etc). So if, for example, a capillary
agarose gel can work but needs something strange, like pulsed electric
field in order to reduce edge effects, or rotation, or whatever, this
would not be a problem to prototype or build these days. All prior
research into gels has to be put into a moore's law perspective.
Tap plastics has acrylic (or whatever) tubes to try as well, either
circular or square, afair. In case plastic straws prove too thin.
> On Tue, Feb 3, 2009 at 5:38 PM, Meredith L. Patterson
>
> > On Wed, Feb 4, 2009 at 2:23 AM, fenn <f...@sdf.lonestar.org> wrote:
>
> >> Right now we use big flat rectangular gels which are submerged in
> >> buffer for cooling, but not too much buffer or all the electricity
> >> goes through the buffer instead of the gel. What if instead each lane
> >> went in its own capillary tube filled with agar? Each tube would have
> >> its own electrodes at each end.
>
> > Consider that more viscous substances, e.g. agarose solution, will
> > require a wider capillary tube than the sort one would normally use
> > for, say, blood. How about a thin, clear drinking straw? You could use
> > paperclips for electrodes.
>
> > This could also be a really neat trick for doing PAGE, since those
> > gels are usually run vertically, with a little well dug in the top. A
> > drinking straw would provide external support for the gel.
>
> > Cheers,
> > --mlp
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