by Herman Gordon, Dept. of Cell Biology & Anatomy, Univ. of Arizona
E-mail comments, updates, etc. to flash @ arizona.edu.
0. Introduction
This protocol derives from one originally written by Evelyn Ralston and circulated privately in the late `80s.
1. Raw materials
DMEM Preparation
Over the course of scientific history, poor quality DMEM has too
often been responsible for the demise of C2 cells.* The
quality control problems appear to be the result of calcium precipitates
that get lost during the filtering of the DMEM. For this reason, care should be taken to thoroughly mix the DMEM in
water and then to use only a high capacity filter to
sterilize it.
* Symptoms include diversification in cell morphologies in GM followed by a high rate of cell death on switching to DM.
Growth medium (vol. for 500 ml bottle; final concentration):
Differentiation medium:
Cell preservation medium (CPM): 10% DMSO + 10% FBS in DMEM (bought pre-made from Gibco)
Sulfate-free Differentiation medium:
2. Incubator conditions
3. Thawing cells
Thaw cells quickly by holding vial in 37oC water bath. Wipe vial with 70% EtOH. Use a 1 or 2 ml
pipet to transfer cells to a 15 ml conical tube containing 10
ml growth medium. Centrifuge for 3 min
at 600 rpm and then remove the medium with a Pasteur pipette connected
to a vacuum line. Tap the
cells to get them to disperse in the remaining droplet of medium,
and then add 10 ml growth medium
and plate into a 100 mm dish. Feed the next day with fresh growth
medium.
4. Plating the cells on the coverslips
By plating the cells directly onto coverslips in 6 well plates, we were not able to get the cell concentration even on the coverslips. Another problem was that we would have cells growing all over the well rather than only on the coverslips. In order to solve these problems:5. Splitting cells
When you grow C2s with the purpose of establishing a cell stock or any other goal that requires that the cells remain healthy over several consecutive passages, it is important to not let them grow too dense and to feed them at the same time every day. On the other hand, they do not like to be passed often at low density. (Presumably, there is a self-conditioning component to their growth.) Best is to keep them below 60% confluence and split no more than 1:10. Once you have that figured out, they are lovely, not-too-finicky cells.
Split the cells when they are about 60% confluent, using 0.05% trypsin-0.02% EDTA in saline (STV).
Aspire to expose the cells to trypsin for only 30-60 seconds. It is our belief that overexposure to trypsin contributes to poor cell health in subsequent generations.
To split a surface of about 25 cm2:
For a larger/smaller surface, increase/decrease the volume of STV accordingly:
Replate in a new flask or dish at a dilution of 1:20 to 1:5. Feed the cells daily.
6. Differentiating the wild C2
C2s will grow on and attach to uncoated, clean glass coverslips.
For better attachment, we use
collagen, gelatin, Vitrogen, or any similar product. Of course,
tissue culture plastic dishes are fine.
Plate about 5,000 cells / cm2 (that means, for example, 10,000 cells per well of a 24-well
plate) on day 1.
Do not feed them the next day. On day 3, remove the medium and
replace with differentiation medium.
Change the medium daily until you decide that the myotubes are
good enough for your experiment. We have found that 3 ml DM / well of a 6 well plate works well and that primo cultures can be achieved by exchanging only 1.5 ml DM each day.
In general, fusion becomes apparent 24 hours after switching to
differentiation medium (day 4), with
good myotubes on the next 2 days (days 5 & 6). If the tubes start
contracting, they are really healthy.
Unfortunately, they also have a tendency to come off the dish.
Ara C (10 uM; 1 mM =100X stock) can be used to get rid of unfused
myoblasts. Add for 48 hours as
soon as fusion is apparent and then feed with medium without Ara
C thereafter.
7. The perfect culture
Differentiated cultures of C2 cells are very dense and are only optimal for a few days. Even after switching to DM, C2 cultures contain ca. 50% myoblasts that continue to divide. Within a few days, the myoblasts overgrow the culture and push the myotubes off.
With a little extra effort, it is possible to achieve cultures
of isolated myotubes that have the potential to last for a
long time. Grow cultures through to 2 days in DM, so that many
myotubes have begun to form. Trypsinize &
resuspend in GM. Allow to settle for 2 min. The myotubes settle
more quickly than do the myoblasts, so pipet
off the bulk of the myoblasts in the supernatant. Replate the
myotubes at low density. They will pull themselves
out o/n. The residual myoblasts will eventually overgrow this
culture as well, but they can be inhibited by ara C
as under 6 above. Another trick for promoting long term cultures
is to replace only 1/2 of the medium each day.
We suspect that this allows for the accumulation of self-conditioning
factors.
8. Freezing the wild C2 for posterity
Label cryovials (Corning, Nunc, etc.) with cell line, passage
number, date, and your initials. Trypsinize and
count the cells. Spin down, remove as much medium as possible,
tap to disperse cells, and resuspend the pellet
with CPM (0.2 ml/ 2 x 105 cells). 200,000 to 250,000 cells / cryovial is ideal. Aliquot
in freezing
vials (0.2 ml/vial). Wrap in a blue diaper inside of a styrofoam
box, & leave to slowly freeze in the -80oC
overnight. Transfer to liquid N2 for long-term storage.
9. Troubleshooting
The most common problem is that cells seem to grow reasonably well in GM, only to fall off in droves when