[DIYbio] Re: DIY electroporation protocol

On May 9, 2012 5:31 PM, "Nathan McCorkle" <nmz787@gmail.com> wrote:
> 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.
>

Ok, I added pics of the latest electroporators here:
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

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