Introduction to Basic Cell Culture
Molecular Biology Lab #5
Background:
Cell culture is used frequently in molecular biology laboratories. It is a reductionist
approach that removes cells from the complex environment of the body and
allows one to study them in an experimental environment that is well controlled. Cell
culture experiments are referred to as in vitro studies as opposed to in vivo or in situ
studies that examine cells in the complete organism.
The first step is to start a primary cell culture by dispersing cells from a living tissue. In
this lab you will study human epithelial cells that were originally isolated from the
cervix. The sample is digested with enzymes that dissociate the tissue and liberate the
individual cells. The cells are placed in a plastic cell culture dish and allowed to settle
down on the bottom. The cells attach tightly to the plastic substrate and start to grow as a
monolayer (one layer). After a variable period (several days to 3 weeks) the cells have
formed a confluent monolayer and they stop growing. This is because normal cells are
contact inhibited by touching adjacent cells. The situation is analogous to a wound. The
cultured cells think they are in an injured area and proliferate until the wound is closed
and cells touch. At that point they stop growing.
If the culture is to be maintained, the cells must be subcultured (also called splitting or
passing cells) by placing some of the cells in a new dish at low density. This is
accomplished by gently digesting the cells with trypsin, an enzyme that cleaves the
plasma membrane proteins that participate in cell-substrate attachment (integrins). After
several minutes, the trypsinized cells round up, they are removed from the monolayer and
they float in the medium. The released cells are resuspended in fresh medium and about
10% are transferred to a new dish. The cells settle down, attach, and start to grow again
as a secondary culture.
After human cells have grown in culture for about 60 population doublings (divisions)
their rate of growth slows down and they essentially run out of gas. Eventually, the cells
fail to divide and become arrested in the G1 phase of the cell cycle. This is a feature of all
human cells and it is called senescence. Senescent cells are not dead; they just do not
divide anymore. If the cells are treated with certain tumor viruses (papillomaviruses) or
oncogenes (genes that contribute to cancer) some of the cells become immortal. This
means that they can grow forever. The step of immortalization is often accompanied by a
period called crisis, when most cells in the culture become senescent. Only a small
percentage of cells become immortal. Once a cell is jmmortal it is called a cell line. This
is to distinguish it from a cell strain, which means any cell that has not yet reached
immortality (normal cells including primary or secondary cultures).
Regardless of whether you are culturing cell strains or cell lines, it is necessary to feed
the cultures with fresh medium every 2-3 days and to subculture the cells whenever they
become confluent. Most cell culture media formulations are purchased as basal medium.
This contains all the ingredients needed to maintain cells in culture (amino acids,
vitamins, salts, glucose, buffers, phenol red, etc ). The basal media is usually
supplemented with 5-10% fetal bovine serum as a source of growth factors needed to
allow cells to grow rapidly in culture. This is called complete medium- Serum is easy to
use and efficiently stimulates growth of most cells, however, it is also an undefined
substance. Different preparations (or lots) of serum vary in potency and most scientists
prefer to culture cells in a medium that is completely defined. There are a number of
serum-free media that are commercially available for specific cell types. The advantage
of these media is that the components are well defined and the scientist knows exactly
what is going into the cells. The disadvantage of serum-free medium is that it is difficult
to discover the right combination of growth factors that stimulate each different type of
cell. In addition, serum free media are often relatively expensive because one must
purchase all of the purified growth factors.
Cell culture medium usually has a red color. This is due to the presence of phenol red, a
pH indicator. At pH 7, the optimal pH for cell growth and survival, the dye appears
orange-red. At lower pH values, the dye turns yellow, and at alkaline values the dye turns
purple. The pH is buffered by adding bicarbonate to the medium. The bicarbonate in the
medium combines with CO2 in the cell culture incubator to balance the pH at 7. Usually,
the incubator injects CO2 into the atmosphere at 5% to accomplish this end. The
incubator also keeps the cells warmed to 37°C. Cells can survive well at lower
temperatures (although they will grow more slowly), but cells quickly die if the
temperature of the incubator exceeds 40°C. Finally, the incubator also provides humidity
to the cell culture to prevent the culture dishes from drying out.
Cultured cells can be cryopreserved by freezing in liquid nitrogen. This is useful if one is
working with a large number of cell lines. When a particular cell line is no longer needed
for the experiment it can be temporarily frozen so that you do not need to continually
feed and pass the cells. This saves time and money. Cells are routinely frozen by
resuspending in medium with 10% DMSO and placing cells in cryovials at -80°C. The
DMSO prevents formation of large ice crystals during the slow process of cell freezing.
After 24 hours, the frozen cells are moved to liquid nitrogen for long term storage (-80°C
is not cold enough). Cells are stable in liquid nitrogen for many years. They can be
returned to cell culture simply by retrieving a vial from the freezer, warming at 37°, and
adding cells to fresh medium.
Sterile technique is very important when performing routine cell culture. The process of
working with cells usually occurs in a laminar flow hood. This hood uses a stream of
filtered, sterile air to keep the working surface clean. Cell culture can also be performed
in a static hood. This hood has a covered area to work in but there is no sterile airflow.
Thus, it is important to flame sterilize objects that will come in contact with cells
(pipettes, medium bottles, instruments) when working in the static hood.
One of the major problems in cell culture is contamination by microorganisms. Using
several antibiotics in the cell culture medium usually prevents potential contamination. If
your culture technique was perfect, antibiotics would not be needed. However, most labs
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use an anti-bacterial agent (penicillin or streptomycin) and an anti-mycotic solution
(fungizone for fungus and yeast) to protect against contamination.
Objectives:
The objective of this lab is to introduce you to basic information about
cell culture and to provide hands on experience in splitting and freezing a cell line.
Materials:
Laminar flow hood
CO2 incubator
Inverted microscope
Table top centrifuge
Bunsen burner
Vacuum flasks and tubing
Sterile pipettes (serological and Pasteur)
Cell culture medium
100 mm cell culture dishes
trypsin solution
phosphate buffered saline
rubber bulbs
70% ethanol and Clorox
cryovials
liquid nitrogen storage container
-70°C freezer
37°C water bath
cell cultures
frozen cultures
15 ml conical centrifuge tubes
cell freezing medium with DMSO
cell splitting medium with 5% bovine serum
Methods:
Hood clean up: Turn on the visible light and turn off the UV light (if available). Start by
cleaning the working surface of the hood. Wash the surface with 70% ethanol and dry
with a paper towel. Turn on the flame and vacuum. Place the vacuum hose in the bottle of
Clorox and suck about 50 ml through the hose to clean.
Cell feeding:
1. Remove a Pasteur pipette from the can and connect the end to the vacuum hose.
Remove the top of the cell culture dish and set upside down. Suck the old medium
out of the culture dish.
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2. Open the bottle of medium and place the cap upside down on the surface of the
hood. Open a can of sterile tissue culture pipettes and remove a 10 ml pipette.
Attach the pipette to a bulb and remove 4 ml of medium from the bottle. Add to
the empty dish.
3. Place used pipettes in the pail or pan.
Cell splitting:
4. Remove a Pasteur pipette from the can and attach to the vacuum tubing. Remove
the top of the cell culture dish and set upside down. Use the pipette to suck the old
medium out of the culture dish.
5. Open the bottle of phosphate buffered saline (PBS). Open a can of sterile tissue
culture pipettes and remove a 10 ml pipette. Attach the pipette to the bulb and
remove 4 ml of PBS from the bottle and add to the empty dish.
6. Rinse the dish for 10-20 seconds to remove serum or other proteins that inhibit
action of trypsin. Using another Pasteur pipette, remove the PBS. Add 2 ml of
trypsin solution to the dish and place the dish at 37°C in the incubator. Check the
dish under the inverted microscope every 2-3 minutes to see whether the cells
have started to round up and detach from the substrate. This process usually
requires 10 min.
7. When all of the cells have rounded up and are free of the substrate, add 2 ml of
serum-containing medium to neutralize the trypsin. Gently mix the cells and
medium using a 10 ml pipette and add contents (4 ml) to a 15 ml conical
centrifuge tube.
8. Spin the tube at low speed (setting 2) for 3-4 minutes to pellet the suspended cells.
6. Remove the tube from the centrifuge and suck away the medium using a Pasteur
pipette attached to the vacuum line. Resuspend the cell pellet in 4 ml of medium
and mix gently using a 10 ml pipette.
7. Add 0.4 ml of cell suspension to a new cell culture dish (60mm dish) containing
4 ml of medium. Move the dish to the incubator and rock back and forth at right
angles to mix up the cells and distribute them evenly throughout the culture. Label
with a piece of tape that says 'do not disturb' and place in the rear of the
incubator.
8. Place any used cell culture dishes or centrifuge tubes in the biohazard bag.
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Cell freezing:
1. Follow steps 1-5 for splitting cells.
2. Resuspend the pellet in 2 ml of cell culture medium. Add 100 l of cells to a new
60 mm culture dish in 4 ml of medium (if you wish to maintain the cells in
culture).
3. Add 2 ml of cell freezing medium to the centrifuge tube and mix with a pipette.
4. Add 1 ml of the cell suspension to a cryovial and attach cap. Label the cryovial
with date, your initials, and the name of the cell line.
5. Transfer the cryovial to the -80°C freezer for 24 hours to allow cells to freeze
slowly.
6. After 24 hours, place cryovials in the liquid nitrogen container. Put on goggles to
protect your eyes! Use a glove to remove a basket from the large nitrogen
container. Be careful that liquid nitrogen does not spill on you or others. Remove
one of the canes from the basket and snap the frozen cryovial into the metal
holders. Make sure it is snug, then return the cane to the basket and replace in the
storage tank.
7. To bring back a cell line that was previously frozen in liquid nitrogen, first put
on eye protection. Use the glove to remove a basket and cane from the storage
tank. Take one of the cryovials from the cane and immediately loosen the cap
(the vials can explode if they warm up and the cap is not loosened). Place the
frozen vial in a boat in the 37°C water bath for 2- 3 min to thaw.
8. Open the cryovial and briefly flame the top. Withdraw the cell suspension with a
1 ml pipette and place into a 60 mm cell culture dish with 4 ml of fresh medium.
Transfer to the incubator and label 'do not disturb'.
Hood clean up: After you have finished, suck about 50 ml of Clorox through the
vacuum line to clean. Turn off the gas and the vacuum. Wash the work surface of the
hood with 70% ethanol. Turn off the regular light and turn on the UV light (if available)
to sterilize the hood. If the large suction flask is full, please remove rubber stopper and
pour contents down sink. Clean up the area and put materials back in the refrigerator!
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