DIY electroporation protocol
Nathan McCorkle
have mid-log phase E.coli in 3 hours (TAing a lab using CaCl2 for
transformation) and LB for recovery... I was thinking washing a few
times with water, then electroporating and adding LB to recover.
Anyone have any better quick/dirty ideas for this?
--
Nathan McCorkle
Rochester Institute of Technology
College of Science, Biotechnology/Bioinformatics
John Griessen
> quick and dirty electroporation?
Meredith was asking about that and making a melaminometer
with it. Meredith?
JG
PS She mentioned http://uwspace.uwaterloo.ca/bitstream/10012/844/1/j2grenie2006.pdf
Design of a MOSFET-Based Pulsed Power
Supply for Electroporation
by
Jason R. Grenier
A thesis
presented to the University of Waterloo
in fulfillment of the
thesis requirement for the degree of
Master of Applied Science
in
Electrical and Computer Engineering
Waterloo, Ontario, Canada, 2006
©Jason
as a starting point.
John Griessen
The parallel and series MOSFET-based pulsed power supplies are capable of
producing controllable square pulses with widths of a few hundred nanoseconds to dc and amplitudes
up to 1500 V and 3000 V, respectively. The load in this study is a 1-mm electroporation cuvette filled
with a buffer solution that is varied in conductivity from 0.7 mS/m to 1000 mS/m. The results
indicate that by controlling the circuit parameters such as the number of parallel MOSFETs, gate
resistance, energy storage capacitance, and the parameters of the MOSFET driver gating pulses, the
output pulse parameters can be made almost independent of the load conductivity.
Sounds like you get a DC HV supply, a bank of capacitance equal to holding the volts
and storing energy, and let it rip. Instead of transistors, quick and dirty wold imply
an insulated wire dangling and touched to make a loud cracking spark. Repeat while necessary.
8 uF was used in the Jason Grenier machine of 2006.
Warning: Keep fingers out of spark path -- your skin is no barrier at all to such Voltage.
JG
sgt york
simple...mix plasmid & bugs ~1:10 or less (pref less), set on ice ~15
minutes, shock, resuspend in media, incubate @37 shaking 1hr, plate.
If you have a richer media (TSB, SOC) that would be better for the
recovery phase.
This is far from optimal, but for a plasmid you know is high quality,
good size, etc, this should work just fine. About as well as a heat
shock technique.
If you don't have the equipment, you could probably do it with a power
supply, a capacitor, good insurance, and some very serious gonads.
You need a LOT of volts in a very small space of time. It's based on
the distance of the gap between your electrodes, but for a 1mm gap,
you'll need ~1k-10k volts delivered in a maximum of 1 millisecond.
Calculate those amps before you decide just how big your gonads are.
If you decide you do indeed have brass balls, just bear in mind they
are conductors ;). And for the record, this is tongue in cheek. I do
not recommend you do this.
For best results, you want to use highly washed DH5-alpha cells. I
used to prep them by growing them up to midlog in low salt LB (LB with
have the NaCl), then doing repeated washes in the centrifuge using
glycerol-water (I think it was 10%). Resuspend in 50% glycerol and
freeze @-80 in aliquots. It takes the better part of a day to prep the
cells. Good for ~6 months.
When you're ready, thaw on ice, mix in plasmid, shock, and resuspend
in 1mL SOC+glucose (it's a rich media, helps the bugs recover. I'm
sure you can find a recipe online). Let them recover ~1hr @ 37
degrees, shaking. Plate 100uL. Spin down, resuspend in 100uL (PBS,
saline, LB, whatever) and plate that.
Nathan McCorkle
and electro-cuvettes.... I know SOC is best, but I don't think I have
the time before the lab in 2 hours to prep any or look around for
some.... I was just thinking it would be fun because my lab is small
and there will be extra plates to compare results. So I am going to
assume I will only have water, MM294 E.coli grown in LB, fresh LB, and
the electro supplies
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Nathan McCorkle
washes with water unless I hear otherwise from y'all soon!
John Griessen
> You need a LOT of volts in a very small space of time. It's based on
> the distance of the gap between your electrodes, but for a 1mm gap,
> you'll need ~1k-10k volts delivered in a maximum of 1 millisecond.
A setup with parallel plate conductors would get most of the "bugs"
between exposed to the same zap strength.
Copper conductors may get reacted into the saline solution
and be toxic CuCl2 to your bugs.
Some ni-chrome heater wire or stainless steel welding wire
flattened on an anvil might make good less reactive electrodes.
Or search around for some thin sheet stainless steel...and tin snips...
Use a plastic rod to do the switching -- duct tape the spark wire onto it.
Don't allow any arm to arm possibility of conduction or any distractions.
To ensure that, duct tape the spark wire onto it
and don't pick up the plastic rod until after changing any settings on anything,
and lay it down before touching anything else after doing the zap contacting.
Shouldn't need a super thrill seeker mentality with those precautions.
JG
sgt york
Do you have access to milliQ, DI or RO water? A few washes in one of
those would probably be better than sparking in LB, but I'm not sure.
Shriveled or cooked....rough on the cells either way, I guess.
If you have some glycerol, use that as an osmotic agent in milliQ
water (18 megaohm, ultrapure, whatever your lab calls it; the kind you
use for HPLC). RO or DI if you don't have the milliQ. Not perfect, but
it's something. Anything that doesn't give you tons of ions and isn't
toxic to the cells would work.
If my quick & dirty calculations are right, ~2% (v/v) glycerol should
give you a good osmotic gradient & not increase the conductance too
much. Make up some 2% glycerol in as pure water as you have. Wash the
cells 3-5 times in that stuff & use that. Just keep them cold; do
everything prechilled, on ice, etc.
Nathan McCorkle
TAing (by myself this week, and I'm an undergrad!), as well I
performed the quick and dirty electroporation too. It worked and my
results were about 4 times as many transformants than the CaCl2 method
(2 plates CaCl2 (17, 8 colonies) 2 plates electroporated (52, 53
colonies) )!
The transformation (CaCl2) failed for the Monday class I don't TA (
and with a different prof teaching the lab too), and this was a good
experiment to try and ensure we'll have transformed cell lines to use
next week!
I would have tried the piezo sparkers, but they were packed in boxes
due to a recent lab move. Another time for sure!
My procedure was simple:
~1.5mL mid-log phase MM294 E.coli (filled microfuge tube with cultured
broth), spin for 10 minutes at 2700-G
decant supernatant and resuspend in sterile water (I used NanoPure, 18
Mohm or so)
spin again for 10 mins at 2700-G, resuspend in 0.5ml sterile water
add 5-10uL ligation reaction (I can't remember now)
transfer solution to electroporation cuvette (if you need it)
electroporate (Bio-Rad genepulser, first transferred the to a 2mm path
electroporation cuvette)
as fast as possible get them some recovery broth (I used SOC and its
preferred, LB prob works with lower efficiency, though not sure) (in
my case lid got stuck and it took prob 20 - 30 seconds from shocking
them to getting them )
place in incubator for 45-60 minutes (can be shaking up to 50 RPM)
plate on selective media gel
Somebody should wiki this or something!
Nathan McCorkle
CaCl2 transformation used 15mL of E.coli culture, whereas the
electroporation experiment used only 1.5ml, so I guess that would mean
40 times better efficiency. (4 times better * (15/1.5))
J. S. John
> On 01/05/2011 10:39 AM, sgt york wrote:
>>
>> You need a LOT of volts in a very small space of time. It's based on
>> the distance of the gap between your electrodes, but for a 1mm gap,
>> you'll need ~1k-10k volts delivered in a maximum of 1 millisecond.
>
> A setup with parallel plate conductors would get most of the "bugs"
> between exposed to the same zap strength.
>
Back when we were talking about DIY electroporation, I found an old
article similar to what you described. From what I remember, there
were 2 metal blocks and a layer of the cell solutions was placed in
between. I don't know the exact setup but I may be able to find it. If
you can wait until the 18th, I may be able to scan and send you the
article from the library. I'll look for the citation but IIRC, it was
not online
Nathan McCorkle
'electroporation and electrofusion (chang chassy saunders sowers)"
its got the low down on everything with graphs.... parallel plates are
already made in one-time use cuvette form, about $3 a piece... and
they can probably be reused with alcohol or/and HCL soaking for
sterilisation
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John Griessen
> electroporate (Bio-Rad genepulser, first transferred the to a 2mm path
> electroporation cuvette)
What volts and pulse width?
Nice write ups for your lab students.
JG
Nathan McCorkle
> On 01/06/2011 11:31 PM, Nathan McCorkle wrote:
>>
>> electroporate (Bio-Rad genepulser, first transferred the to a 2mm path
>> electroporation cuvette)
>
> What volts and pulse width?
http://nathanmccorkle.com/projects/biorad1.JPG
http://nathanmccorkle.com/projects/biorad2.JPG
2.5kV (I used a 2mm path), 2.5msec time constant (not sure what this
means exactly)
>
> Nice write ups for your lab students.
>
> JG
>
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Nathan McCorkle
> So I ended up doing the CaCl2 transformation along with the class I am
> TAing (by myself this week, and I'm an undergrad!), as well I
> performed the quick and dirty electroporation too. It worked and my
> I would have tried the piezo sparkers, but they were packed in boxes
> due to a recent lab move. Another time for sure!
before though... got 2 colonies on the circumference of the agar, so
it could be contam from the incubator... subcultured them but waiting
on the results... will still need to induce with arabinose too I think
pics of the ghetto-porators
https://picasaweb.google.com/109403794341975968814/DropBox?authuser=0&authkey=Gv1sRgCIew8dDku-y-ugE&feat=directlink
Procedure (really crappy, not worth repeating in this exact way):
HB101 cells from a healthy streak plate were scraped with a sterile
toothpick, about 1cm drag of the tip.
Toothpick w/cells was beaten/swirled in 500uL sterile water in a 1.5mL
eppendorf tube
This was repeated 6 times with a fresh toothpick and a fresh
tube/water, each labelled numerically in sequence
lyophilized pGLO plasmid (bio-rad) was suspended in TE buffer to
80ng/uL, 10uL of this was added to each eppendorf with cells in it
one tube didn't have cells (control for reagent contamination), one
tube didn't get electroporated (control for anomalous growth not due
to electroporation procedure)
2 methods of electroporation (ghettoElectroPoration, or ghettoPoration)
piezo with wires soldered on, and end of wires coiled in circles
piezo with wires soldered on, and end of wires covered in conductive
aluminum foil tape (to increase surface area)
recorded which tubes i gave 1 click, 3 clicks, or 5 clicks... after
done clicking the piezo sparker, I quickly added 500uL LB broth
between tubes, I immersed the wet areas of the wires in 100% ethanol
for about 10-20 seconds, then dried with kimwipe (small paper towel)
and twirled the leads between my fingers to spin the ethanol off them
After about 2 hours of 'incubating' the benchtop, I plated 500uL per
plate (2 plates for each tube) on LB+amp
OVERALL:
I think the whole process was too dilute cell-wise.
Next I'll try LB-broth mid-log liquid culture with plasmid added
directly to it... to avoid using a centrifuge. I think I'll also try
something using the aluminum foil on a microscope slide and thin line
of cell+plasmid solution between them (to increase concentration from
a plate scraping, and adjust the spark gap.
Avery louie
--A
Nathan McCorkle
Yes pGLO is amp resistant, not sure what you mean by satellite colonies
Cory Tobin
protein gets secreted into the media surrounding the amp/carb
resistant colony and breaks down the antibiotic in the surrounding
region. This means that non-resistant colonies can form in a ring
around the resistant colony. This usually happens when you let the
plate incubate for more than a day.
-cory
Nathan McCorkle
Ahh, to be clear there were only two colonies on 12 non-control plates, and those two were at edge so they might just be contamination. I doubt it has anything to do with satellite colonies
John Griessen
> I think the whole process was too dilute cell-wise.
of concentration you mention fresh toothpicks. 6 times.
Does a fresh toothpick have any cells on it, or sterile?
Not understanding all of it.
6 shaking dilutions with the *same* toothpick would be a huge dilution factor --
PPM I'd guess.
John
Nathan McCorkle
drag along a lawn of E.coli, each drag got swirled in its own tube
that had 500uL sterile water
What I meant re 'too dilute' was that I think the solution's cell
concentration needs to be higher. I could just try to scrape 6cm of
cell law per toothpick, which would increase the concentration of
cells 6X. My reasoning is that there just weren't enough cells to get
in the way of the DNA particles crashing towards the positive + side
of the circuit during the pulse, basically like an all electro-liquid
gene gun. I could also increase the concentration of the DNA, but the
cells are easier and cheaper to get than the plasmid.
>
> John
>
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Nathan McCorkle
Seems that one of the two colonies has pGLO! So overall the protocol works, but I'll do some more tuning to increase the efficacy
Pic added to the Picassa link I posted earlier
Meredith L. Patterson
Brilliant! I brought this up on the NPR Weekend Edition interview I just got back from, so I'm especially glad it worked ;) (You and Cathal and the mailing list all got shout-outs.)
--mlp
Cathal Garvey
Cathal Garvey
Nathan McCorkle
I did that control, I must've forgot to mention
I'm gonna repeat again in next few days
Cathal Garvey
Thanks Nathan, awesome work and an encouraging outcome!
I wonder if you can improve efficiency with some buffer jiggery-pokery, or by altering spark-gapping..
Hmm! So, things confirmed you can electroporate with a gas lighter: E.coli, Mice. :P
Nathan McCorkle
Mice!!?? Where?
Cathal Garvey
It involved subdermal plasmid delivery in mice using a piezoelectric
sparker, with fair results! (Don't try this at home, etc.)
www.indiebiotech.com
twitter.com/onetruecathal
joindiaspora.com/u/cathalgarvey
PGP Public Key: http://bit.ly/CathalGKey
Cathal Garvey
Nathan McCorkle
Looks like this is it
http://www.sciencedirect.com/science/article/pii/S0378517301006081
Electroporation of the skin to deliver antigen by using a piezo ceramic gas igniter
Pramod Upadhyay ,
National Institute of Immunology, Aruna Asaf Ali Marg, New Delhi 110067, India
Received 26 October 2000. Revised 5 January 2001. Accepted 27 January 2001. Available online 30
March 2001.
International Journal of Pharmaceutics
Volume 217, Issues 1–2, 17 April 2001, Pages 249–253
Nathan McCorkle
Meredith, will that NPR weekend edition be audio or print? Can you post when we can listen where we can read?
Meredith L. Patterson
Audio, this Saturday. I'll post a link to the recording when I know the address :)
--mlp
Nathan McCorkle
> OVERALL:
>
> I think the whole process was too dilute cell-wise.
>
> Next I'll try LB-broth mid-log liquid culture with plasmid added
> directly to it... to avoid using a centrifuge. I think I'll also try
> something using the aluminum foil on a microscope slide and thin line
> of cell+plasmid solution between them (to increase concentration from
> a plate scraping, and adjust the spark gap.
>
https://picasaweb.google.com/109403794341975968814/DropBox?authuser=0&authkey=Gv1sRgCIew8dDku-y-ugE&feat=directlink
Electroporators:
Aluminum adhesive-backed foil tape for electrodes
($7.58, Home Depot, Nashua Tape 322 1-57/64 in. x 150 ft,
http://www.homedepot.com/h_d1/N-5yc1v/R-100030120/h_d2/ProductDisplay?catalogId=10053&langId=-1&keyword=foil&storeId=10051)
Glass microscope slides were cleaned by dunking in 70-100% ethanol,
gripping the wet slide with a microscope slide holder (springy wide
pliers) and passing it through a bunsen burner to sterilize and dry.
(CAUTION alcohol on slide will catch on fire)
I made one and my friend Chris made one. I used two pieces of foil
with the factory cut edges facing each other with a 1cm gap, with the
piezo wires wrapped around the end of the glass slide, being covered
by the foil. My friend tried using a single piece first, then cutting
and peeling out a channel, but there was significant adhesive residue
that would be hard to clean off.
The only thing he did different was, instead of a 1cm gap, he used
2.1cm gap, and he taped his wires on instead of wrapping them under
the main foil layer.
We flamed the aluminum covered slides in the bunsen burner, then while
warm drew two lines perpendicular to the aluminum electrode edges. We
tried using Sharpie, but a wax pencil (or crayon) worked a lot better.
My lines were about 0.2cm apart, my friends were 0.3cm apart.
Protocol:
Make overnight cultures of HB101 from 9pm to 5pm (20 hours), a shaking
tryptic soy (TS) broth tube and a streaked MacConkey agar petri dish
(LB agar would be fine too).
Add 15uL sterile water to a sterile 1.5mL tube, scrape petrified dish
about 1cm with sterile stick, twirl stick in 15uL of water to suspend
the cells. Add to the now ~15uL of E. coli water 5uL pGLO plasmid
(80ng/uL). Pipette this solution (~20uL total) onto the prepared
electroporator capillary, moving the tip back and forth from one
terminal to the other, to ensure the path between the electrodes is
completely wet.
Add 100uL sterile LB broth to sterile 1.5mL tube.
Spark electroporator 3 times, with about 4 seconds between each pulse.
Take up the liquid from the electroporator, and dispense into the tube
containing 100uL sterile LB broth.
My friend used his device (2.1cm gap) with a bit different protocol.
He mixed 45uL sterile LB broth, 45uL overnight culture, and 10uL pGLO
plasmid (80ng/uL), sparked 3 times, then transferred the liquid from
the electroporator to a tube containing 20uL sterile LB broth.
CONTROLS were prepared, one of 120uL overnight culture (to test the
ampicillin in the plates), and one of 100uL LB + 15uL overnight
culture + 5uL pGLO plasmid.
We incubated them in a shaker for 60-75 minutes at 37 C (except the
pure overnight 120uL subculture), then plated all 120uL from each tube
onto separate LB+ampicillin agar plates using sterile cell spreaders,
then incubated at 37 C upside down.
They're incubating now, I'll update in a day or two!
-Nathan
Nathan McCorkle
the slide off with alcohol and a paper towel, then redrew with wax,
and flamed one last time before re-using.
There were no cells today, they're still incubating so I'll give them
another day or two.
Nathan McCorkle
toxicity (probably not according to a paper I found), if I need higher
field strength (non-log phase cells have thicker membranes), maybe the
pulse time is too short, maybe the field isn't being setup correctly
(I can try with a commercial cuvette).
"E. coli cells in the logarithmic phase were more sensitive to PEF
[Pulsed Electric Field] treatment when compared to cells in the
stationary and lag phases (Pothakamury and others 1996)."
(supposedly from)
U. S. Food and Drug Administration
Center for Food Safety and Applied Nutrition
June 2, 2000
Kinetics of Microbial Inactivation for Alternative Food Processing Technologies
Pulsed Electric Fields
http://altered-states.net/barry/rife/pulsedelectricflds.htm
High efficiency electrotransfection with aluminum electrodes using
microsecond controlled pulses
U. Friedricha, N. Stachowiczb, A. Simmc, G. Fuhrd, K. Lucase, U. Zimmermann
Bioelectrochemistry and Bioenergetics
Volume 47, Issue 1, November 1998, Pages 103–111
http://www.sciencedirect.com/science/article/pii/S0302459898001639
John Griessen
> Again today, no cells... I'm wondering if the Aluminum caused acute
> toxicity (probably not according to a paper I found), if I need higher
> field strength (non-log phase cells have thicker membranes), maybe the
> pulse time is too short, maybe the field isn't being setup correctly
> (I can try with a commercial cuvette).
and now they're slow...
Things that are affecting field strength in the zone include
the spark generator's ability to drive,
The load impedance --> (path length and width and thickness and concentration
of cells in electrolyte),
randomness of sparks,
connections (and gaps) of wires to electrodes (if under sticky tape).
I'd look at the path. 2 and 3 are less likely.
1. You could try shortening it, all else the same.
2. You could look at the conductivity of your cells in medium.
3. You could increase the stripped wire length and contact it
to the bare aluminum surface of the top of the tape that is your electrode.
You could switch to strips of stainless steel instead of Al tape since aluminum
might oxidize more and more and conductivity drop with repeated sparking.
John
Nathan McCorkle
> On 05/14/2012 03:11 PM, Nathan McCorkle wrote:
>>
>> Again today, no cells... I'm wondering if the Aluminum caused acute
>> toxicity (probably not according to a paper I found), if I need higher
>> field strength (non-log phase cells have thicker membranes), maybe the
>> pulse time is too short, maybe the field isn't being setup correctly
>> (I can try with a commercial cuvette).
>
>
> The first time only one sample grew if I remember correctly,
> and now they're slow...
time the colony came from a plate where the electrodes were ~8mm
coiled braided coming out of the end of an insulated wire, the
electrodes were about 8mm apart. So maybe it was higher field
strength???
>
> Things that are affecting field strength in the zone include
> the spark generator's ability to drive,
> The load impedance --> (path length and width and thickness and
> concentration
> of cells in electrolyte),
> randomness of sparks,
> connections (and gaps) of wires to electrodes (if under sticky tape).
>
with a third wire to make sure there was a conductive path and not
insulated by the adhesive backing of the foil
From the oscilloscope traces Simon sent out a while ago:
https://groups.google.com/d/msg/diybio/eJeUIsCYPGo/zcmihRzecNAJ
its an oscillatory waveform that completes one period in 40
microseconds, so each pole gets ~20 microseconds, and peak to peak
(not including the 200MHz noise) is about 20kV. I read a paper earlier
that showed 200 microseconds of a DC exponential decay gave highest
efficiency at 13.5kV/cm, with mid-log phase cells. I read another
paper that said mid-log phase cells have thinner membranes, so they
need less field strength. I used stationary or lag phase cells, so the
membranes might be thicker, so maybe these cells need a higher field
strength? I also read that oscillatory pulses kill less cells than
exponential pulses, so I imagine that means it takes a higher field
strength to get the same effect with an oscillating wave.
So, assuming each pole of the 20kV p-p is 10kV, and you need >18kV/cm
for stationary cells, and assuming we want 200 microseconds of
pulse... I think next I'll try 5 pulses at 0.5cm, 10 pulses at
0.5cm... plated on both selective media and plain, to get an idea of
cell survival vs non-electroporated cells on plain media.
> I'd look at the path. 2 and 3 are less likely.
>
> 1. You could try shortening it, all else the same.
> 2. You could look at the conductivity of your cells in medium.
> 3. You could increase the stripped wire length and contact it
> to the bare aluminum surface of the top of the tape that is your electrode.
> You could switch to strips of stainless steel instead of Al tape since
> aluminum
> might oxidize more and more and conductivity drop with repeated sparking.
>
> John
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John Griessen
> so each pole gets ~20 microseconds, and peak to peak
> (not including the 200MHz noise) is about 20kV.
" you need >18kV/cm
for stationary cells," [jg] OK, if you want that and path length
was 0.8cm, 18kV/cm * X = 10 kV, solve for X = .55cm instead.
" and assuming we want 200 microseconds of
pulse"
for your next experiment.
John
PS keep the path skinny and solution only slightly conductive,
or the volts will go down with current load.
Cathal Garvey
exponential-phase cells. I gather it's far more than a 100-factor drop
in efficiency, but I don't have numbers on it.
Also, expect transformed cells to form colonies a little bit slower than
"normal" cells anyways, no matter the method. A lag of 2-4 hours would
be normal, I'd estimate. Part of this is down to the additional stress
of being transformed, but also because normal colonies are formed as
much from small clumps of cells as individual ones. Whereas, when you
transform cells, only one cell in a clump may survive. With an E.coli
doubling time of 20 mins at 37C, a clump of 16 cells will form a colony
of a given size 80 minutes before a single cell from a successful
transformation.
Nathan McCorkle
> On 05/15/2012 12:46 AM, Nathan McCorkle wrote:
>>
>> so each pole gets ~20 microseconds, and peak to peak
>> (not including the 200MHz noise) is about 20kV.
>
>
> Is 10kV peak higher than needed for a 1cm gap?
>
> " you need >18kV/cm
> for stationary cells," [jg] OK, if you want that and path length
> was 0.8cm, 18kV/cm * X = 10 kV, solve for X = .55cm instead.
membranes are thinner. Stationary phase (i.e. 20 hour culture growth)
will have thicker membranes, so they /should/ need a higher field
strength.
Nathan McCorkle
> From what I know of EP, you can't skimp on the requirement for
> exponential-phase cells. I gather it's far more than a 100-factor drop
> in efficiency, but I don't have numbers on it.
beginners will want results in worst-case situations. Even for me, not
being in the lab everyday, its a bit annoying to have to subculture
for 6-8 hours then come in and do some work... easier to throw a petri
dish in the incubator and come back where there are cells growing.
Also Mega had tried a transformation using a petri dish scrape, which
I hadn't thought of before (coming from academic labs where we always
pay attention to growth phase, and have liquid media, and centrifuges
to rinse and concentrate cells)... and I'm not sure what petri dish
cells experience as far as growth rates.
If I can't get any results with a piezo sparker in these conditions,
well, then we'll have to come up with some other ideas!