Cell and organ cultures are used to maintain living animal cells and
outside the body (in vitro). With separate, living cell cultures, it is
and study the behavior of animal cells in greater detail than when they
the animal (in vivo). Cell culture also frees the cells from some of the
controls that normally regulate their activities. Cells, or tissues, are
varying periods of time, at times undergoing repeated divisions over
Cells grown and cultured for study have been taken from a wide variety
species, such as humans, monkeys, mice, dogs, cats, frogs, insects,
others. The cultures have come from a number of organs—heart, lungs,
kidney, blood, skin, etc. In cancer research, it is common practice to
from normal and cancerous tissues to compare their properties. In fact,
cellcultures have become one of the best means of testing potential
utilizing cell cultures is more cost-effective and faster than
animals and, with this method, isolated human cancerous tissue can be
General Principles and Techniques
Tissue fragments used in the preparation of cell cultures must be
the laboratory to avoid microbial contamination; sterile, or aseptic,
technique must be employed at all times. All instruments, culture
come in contact with the cells or medium must be sterile. The tissues
37°C and suspended in a physiologically balanced salt solution.
small amount (1%–15%) of blood serum helps protect the cells during
preliminary manipulations. Antibiotics may also be added as well as pH
phenol red. Tissues are cut into very small fragments, called
explants, which are put into vessels and bathed with nutrient medium.
Sometimes the fragments are attached to the surface of the vessels by
attraction or with blood plasma, which is then allowed to clot. In the
the cultures of this type, cells migrate from the explanted tissue into
and undergo division to produce a “halo” of outgrowth around the
Cell cultures are also started by treating tissue fragments with
(enzymes such as trypsin, or chelating agents such as versene) to
into a suspension of single cells. These cells are then placed in a
and a portion of the suspension is put into a suitable culture vessel.
culture vessel is incubated without being disturbed, which allows the
settle out from the suspension, attach to the wall of the vessel, and
complete sheet of cells covering the vessel wall is called a monolayer.
“Cell lines”, which are capable of continuous growth, are usually grown
monolayers. To transplant such cultures, the cells are either scraped
the glass with a rubber spatula or the medium is taken off and the cells
removed with a chemical (enzyme or chelating agent). The cells are then
suspended and counted, and the suspension is diluted with fresh medium
the number of cells desired.
Students will demonstrate one of the methods used to initiate primary
cultures from fresh embryonic tissue and observe patterns and rate of
in a mixed culture (the culture prepared representing a mixture or many
types). Students will also become acquainted with the initiation and
tissue cells in in vitro culture and with fixation and staining
the field of cytology, and will clearly differentiate between nuclear
material and cytoplasmic material in animal tissue cells.
A. Establishment of the Primary Cell Culture
- 2 curved dissecting forceps
- 1 sterile culture tube
- 1 sterile Petri dish per group
- 25 sterile pipettes
- 10 sterile culture flasks
- 1 sterile glass rod
- 2 sterile versene tubes
- 1 sterile medium tube 199
- 5 sterile alcohol pads
- 1 staining jar
- 1 bottle of hanks balanced salt solution, 100 mL
- Methanol 30 mL
- Hematoxylin stain 30 mL
- Eosin stain 30 mL
- Isopropyl alcohol 99% 100 mL
- Piccolyte II mounting medium 15 mL
- Fertile hen’s egg, incubated for 7 days
- Isopropyl alcohol, 70%
- Compound microscope
- Safety goggles
- Lab aprons
- Pre-Lab preparation
Seven days before the experiment, obtain a fertile hen’s egg, and
according to instructions accompanying your incubator.
B. Propagation of Chick Cells onto Cover Slips
Though the embryo is relatively underdeveloped, working with a chick
be a sensitive issue for some students. If this is the case, you may
1 through 12, involving maceration of the embryonic tissue, in advance.
fertile egg, incubated for 7 days, provides sufficient material for 5
groups. Do not remove sterile materials from their protective packages
ready to use them.
- Obtain a fertile hen’s egg, which has been incubated for 7 days. Candle
the egg to verify the presence of a developing embryo and locate its position.
To candle the egg, use a box containing a 150-watt bulb, with a 2”-diameter
hole cut in the box. The box should be in a dark room. Place the egg, large
end up, over the hole. If the egg is fertile, the developing blood vessels will
- Place the egg, large end up, in an egg carton. The large end contains the
air sac. Wipe the egg thoroughly with a sterile alcohol pad to sterilize the
- Sterilize a pair of forceps by passing them slowly through an open flame;
allow to cool before using.
Once used, items are no longer considered sterile.
- Gently crack the shell over the air sac. Remove the shell surrounding the
air sac with the sterile forceps, taking care not to rupture the shell
- Tear off the shell membrane.
Do not let bits of shell fall into the egg.
- Carefully pour the contents of the egg into a sterile Petri dish. With 2 pairs
of sterile forceps, remove the membrane from around the embryo.
Be very careful when removing the membrane; the embryo could explant.
Materials can be damaged easily if pressed too tightly or pulled too hard.
- Place the embryo in the sterile culture tube. Cap the tube.
- With a sterile pipette, add 1 mL versene solution to the explant material.
Discard the pipette.
Set aside the remainder of the versene for use in part B of the investigation.
- Gently grind the explant material and versene with a sterile glass-grinding
rod. Replace the cap on the culture tube and set aside for approximately
20 to 30 minutes.
You may also macerate embryonic tissue with a sterile.
Syringe. The process is similar to grinding, with less chance of
contamination. Add 1 mL versene solution to the tissue in the culture tube.
Place the embryo and versene in a syringe with at least a 50-mL capacity.
Reset the plunger, position the syringe over the culture tube, and press,
using one fluid motion. Pushing the tissue through the syringe will facilitate
the action of the versene.
- With a sterile pipette, draw the tissue up and expel it from the pipette
several times to homogenize it further. Discard the pipette.
- With a sterile pipette, add 1 tube of Medium 199 to the culture tube.
Discard the pipette.
- Shake to mix the suspension well. Allow it to stand for 5 minutes.
- With a sterile pipette, carefully transfer 3 mL of the cell suspension to a
sterile culture flask. Do not draw up large particulate matter. Cap the flask.
- Label the flask with the date of the culture and the tissue source. Incubate
the flask with its largest, flat surface facing downward at 37°C (99 °F).
- Once the tissue has been
incubated, invert the flask and place it on the
stage of a compound microscope. The
cells will be growing on the surface
that was facing down during
incubation. Examine the cells at 100X. At this magnification, an
individual cell will appear to be about the size of
a grain of rice. Focus carefully to
view the nucleus within each of the cells.
You should be able to observe a
network of cells. Since there will be a mixture
of cell types, some cells will be
elongated and others compact. On occasion, some
cells may regenerate into a large
- If your cultures turn yellow (acidic) or the fluid becomes cloudy, it is an
indication that they have become contaminated; they should be discarded.
Disposed-of biologic materials should be autoclaved. If an autoclave is not available,
place the unopened cultures in a pan of water, bring to a vigorous boil for 20
minutes, then discard. Instruments used in the investigation should likewise be
sterilized by either autoclaving or boiling.
Once tissue cells have been established in primary culture (first
usually be maintained for some time by serial subculture. However,
cultures, as developed in part A, can usually be transplanted, or
only 2 or 3 times. When a cover slip is added to the culture medium in
cells are suspended, some of the cells will settle out and grow on its
creating a monolayer that can be viewed under a microscope.
C. Fixation and Staining of Chick Cell Monolayer
- Obtain a culture flask that has been incubated for 6 to 7 days.
- Pour off the medium, taking care not to contaminate the flask opening or
its contents. Replace the cap. Discard the medium.
- With a sterile pipette, add 1 mL versene solution to the flask. Swirl
flask and pour off the versene. Replace the cap. Discard the pipette.
- Place the flask on a flat surface so the solution completely covers the
of cells. Leave at room temperature for 15 minutes.
Mark the side of the culture flask on which the cells are growing.
- The cells will be loosened by the versene solution and will float off
surface. If necessary, you may extend the incubation period, or you may
agitate the culture flasks vigorously to help dislodge the cells.
- With a sterile pipette, draw up and eject the suspension several times to
break up any larger cell clumps that may be present.
- Draw up the full amount of cell suspension in the pipette and transfer
to the unused vial of Medium 199. Pool the cells from 5 flasks into 1
medium vial. Discard the pipette.
- Swirl the vial gently to mix the transferred cells with the fresh medium.
- Obtain a sterile vial containing a cover slip.
- With a sterile pipette, add 2 mL of the cell suspension to the vial. Cap the
culture vial tightly. Discard the pipette.
- Label the vial with the name of the culture, its original initiation date, and
the subsequent transfer.
- Place the vial in the incubator; stand it upright to ensure the cover
remains perfectly flat on the bottom of the vial. Incubate the culture
at 37°C (99 °F) for 4 to 5 days. The cells should settle and attach to
cover slip within a few hours.
which are normally colorless, are fixed and stained for viewing under a
microscope. Fixing is a process that stabilizes the chemical and
the cells. Cells are then stained with a dye or combination of dyes
highlight structural details. Different fixation and staining procedures
different structural features. The fixative methanol is good for
Hematoxylin, a powerful stain commonly used by cytologists, stains
material bright blue to dark blue. Hematoxylin does not stain
material, however, so another stain must be employed, such as eosin,
stains cytoplasmic material light pink to red.
Wear safety goggles at all times. Use care in handling stains
This procedure should be performed 3 to 4 days after preparing the
- Heat Hanks balanced salt solution to 37°C (99 °F.) Place in a staining jar.
remove the cover slip from the culture vessel: tip the cover slip
away from the bottom of the vessel and gently remove it with forceps.
The cover slip is extremely thin; be very careful when handling it with
- Immediately place the cover slip in warm Hanks balanced salt solution
the staining jar. Orient the cover slip so the side with cells attached
- Taking care to keep the cover slip in the staining jar, pour off and
the used balanced salt solution. Immediately add fresh, warm balanced
salt solution for a second washing. Repeat, for a total of 3 rinses in
- Pour enough methanol into the staining jar to just cover the cover
Allow cells to fix for 7 to 10 minutes. Agitate the staining jar
to ensure thorough fixation.
Be careful when using methanol; it is poisonous if taken internally.
- Pour off and discard the methanol. Rinse the cover slip with water to
remove excess methanol.
- Add enough hematoxylin stain to completely submerge the cover slip.
Allow the cells to stain in this solution for 10 minutes.
- Pour off and discard the stain. Rinse the cover slip with water.
- Add balanced salt solution and set aside for 2 minutes.
- Pour off and discard salt solution. Rinse the cover slip with tap water.
- Add enough eosin stain to completely submerge the cover slip. Allow to
the cells to stain for 5 minutes.
- Pour off and discard the stain. Rinse the cover slip in 99% isopropyl
alcohol 3 times. Allow 1 minute for each rinse, agitating the staining
- Add Histoclear to the staining jar and allow to set for 2 minutes, agitating
- Pour off and discard the Histoclear. Add fresh Histoclear and set aside for
- During these 2 minutes, clean a glass microscope slide with a lint-free
- Just before the end of the 2-minute period, add 1 drop of Piccolyte II
mounting medium to the center of the clean microscope slide.
- Remove the cover slip from the Histoclear and place it cell-side down
the center of the microscope slide.
Be sure not to trap air bubbles beneath the cover slip. This can be
placing one side of the cover slip against the microscope slide and
the cover slip onto the slide. If an air bubble is trapped, it can be
forced out by
gently pressing on the cover slip with a sharp instrument.
- Allow the slide to dry for at least 24 hours before handling it.
- Examine the slide under 100X magnification. Note the pattern and rate
cell growth. The increase in cell rate can be roughly calculated by
a representative microscope field and counting the number of cells in
field. Mark the field by circling it with a glass marking pencil.