O’Neill Research Lab Introduction to Basic
Techniques.
Part Two: Cell and Tissue Culture
This is by far the most important basic technique that you will need to
master in order to efficiently carry out research in our lab. Everyone
working in my lab will be required to pass off tissue culture before they
can proceed. Learning these techniques are essential as a contamination
can be very costly in time, money, and effort. Each section must be
passed off by instructors some sections will require you observing three
different instructors to become proficient. Learn from each instructor
and make good notes. Become a professional.
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ASEPTIC TECHNIQUE
This section outlines the basic procedures in aseptic technique. This will help minimize the possibility of
contamination. Often times, carelessness, not incompetence, is the cause of contamination. The following
is presented as a foundation for aseptic technique, and does apply for research outside of the laminar flow
hood as well.
Laminar Flow Hood
Although primarily used for cell culture practices, the flow hood is NOT a sterile environment.
However, it creates a clean-air space, minimizing the possibility of air-borne dust and microorganisms
from entering into the experiment medium, cells, etc.
1. Turn off the UV light. You do not want to expose UV light onto your skin.
Note in some models, the “receptacle” button turns the UV light off.
2. Turn on the fluorescent light. Note that sometimes the aforementioned procedures may be done with
the same switch.
3. Open front shield about 7 inches or 8 inches.
4. Turn the Laminar Flow Hood on.
5. Spray the Laminar Flow Hood liberally with 70% ethanol. Spray deep inside as well as the outer rims
of the hood.
6. Wait at least 10 minutes before entering the hood. This will give the Laminar Flow Hood sufficient
time to create a good “clean-air” space in the hood.
I know how to start up a laminar flow hood.
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Cardinal Rules
1. Spray your hands and arms with 70% ethanol when introducing them into the hood. There are no
exceptions to this rule.
Note: You do not need to wear gloves in the hood unless working with hazardous material. Gloves
reduce the sensitivity of your touch and therefore you may brush a flask without feeling it. This can
lead to contamination.
2. Organize your space. This will minimize the likelihood of contamination.
3. Always work in the clean-air space. The hands should be placed deep in the hood, never near the front
where the air vent is located (keep elbows off vents).
4. Minimize contamination by avoiding contact with bottle/flask necks, caps and openings. If any item is
possibly contaminated (through touch, exposure outside hood, etc.), remove from hood, or spray
liberally with 70% ethanol.
Do not pass hands over any open containers. This will result in dust particles falling into the
containers.
I understand the importance of the cardinal rules.
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2
Media Bottles
1. Remove bottle from yellow fluorescence protection bag.
2. Warm the media bottle. This is necessary to avoid ‘cold shock’ when you are feeding your cells.
3. Handle bottles with care. Splashing inside the bottle will create unwanted air bubbles and you do not
want any media to touch the neck/cap of the bottle.
4. Lightly flame the neck of the bottle with a Bunsen burner. Process should never exceed 2-3 seconds.
This will remove dust particles from the outside of the bottle.
5. Spray bottle liberally with 70% ethanol.
6. Place the bottle in the hood.
7. Unscrew the bottle cap until it is open. Let the lid sit on the bottle mouth, this will give you quick
access to the bottle when needed.
NOTE: Never handle the neck of the bottles. Always handle bottles by the base to avoid
contamination.
8. Carefully lift the cap and tilt the bottle in one motion. Practice this technique as advised. Tilting of the
bottle will minimize contamination by particles falling into the bottle
9. Close the bottle firmly. Remember to close the bottle before removing from the Laminar Flow Hood.
10. Remove bottle from the hood. Be careful not to get media in the neck of the bottle while removing.
11. Spray with 70% ethanol, then replace in yellow fluorescence protection bag. This will minimize
contamination. Pay attention to how you handle the bottle, handling the neck of the bottle is most
common in this process.
Flasks
1. Handle flasks at the base. The neck of the flask should not be handled under any circumstance. Avoid
touching media to the neck of the flask. This will lead to contamination.
2. Tighten cap of the flask when removing from incubator. This will minimize contamination, but do not
leave flask unattended while sealed, as your cultures will die. When returned to the incubator the flask
caps should be loosened a ¼ turn to allow the passage of Co2
3. Lightly flame the neck of the flask with a Bunsen burner. Process should never exceed 2-3 seconds.
This will remove the dust particles from the bottle.
4. Spray flask with 70% ethanol.
5. Introduce into the Laminar Flow Hood.
6. Unscrew the flask cap until it is open. Let the lid sit on the flask mouth, this will give you quick access
to the bottle when needed.
7. Carefully lift the cap and tilt the flask in one motion. Practice this technique as advised. Tilting of the
flask will minimize contamination by particles falling into the bottle.
8. Close the flask firmly. Remember to do this before removing from the Laminar Flow Hood.
9. Remove flask from the hood.
10. Unscrew the cap of the flask to allow CO2 into the flask. The arrows should point upwards and
downwards, or cap should be fitted loosely.
11. Spray with 70% ethanol, then place back into incubator.
I understand how to work with bottles and flasks.
Sign____________________________ Super____________________ Date__________
Pipette Canisters
1. Handle the bottom of the pipette canister. Do not handle the canister near its opening (neck).
2. Lightly flame the neck of the pipette canister with a Bunsen burner. This process should never exceed
2-3 seconds. This will remove the dust particles from the bottle.
3. Spray liberally with 70% ethanol.
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4. Place pipette canister in the hood.
5. Shake the pipette canister upside-down gently. This will slide the pipettes inside towards the lid of the
canister, making the pipettes more accessible.
Note: excessive shaking will crack the pipettes inside the canister.
6. Slowly remove the lid of the pipette canister. Be sure that the pipettes inside do not touch the edges of
the canister lid.
7. Angle the pipette canister upwards. Use the lid of the canister to stabilize. Pipettes should be sticking
out of the canister.
8. When finished, close lid on canister.
Note: When working in the Laminar Flow Hood, keep the pipette canister in an area free of any
obstruction. Hitting the pipettes can not only crack them, but also potentially contaminate your
cultures.
Pipettes
1. Pipettes should be projecting out from the pipette canister. This will have the pipettes readily
accessible.
2. Slowly slide one pipette outwards. Do not slide the tip along other pipettes. Grasp the pipette below
the notch and above the graduations. This is the only area which you should handle these pipettes.
Usually, there are brand names, professor names, or department names labeled on the pipette in this area.
3. Use an underhand grasp, and not an overhand grasp.
Insert into Pipet-Aid firmly. Some twisting of the pipette might fit the pipette deeper into the Pipet Aid.
4. Orient graduations towards yourself. This will insure correct measurement and minimize
contamination.
5. Angle the bottle/flask. This will minimize the area for dust particles to potentially fall into the
container.
6. Pipettes should be dipped into the bottles/flasks without touching anything. Be cautious of pipettes
touching the inside walls of the bottles/flasks. This will lead to immediate contamination. If the pipette
touches any surface, remove and start with a new pipette.
7. Siphon/expel the contents of the pipette. If the pipettes are correctly angled when expelling contents,
formation of air bubbles will be minimized. Be cautious to not froth the cell medium. Consult instructor if
necessary.
8. Remove the pipette from the Pipet-Aid.
9. Place pipettes in the Clorox soak columns outside the Laminar Flow Hood. Tips of the pipettes should
be pointing downwards in the columns, as they soak in Clorox but when washing the tips should be
pointing up to ensure the pipettes are filling.
Note: Place any unused pipettes in appropriately labeled drawers. This will bypass the washing
procedure and the pipettes will go immediately to sterilization.
10. Store empty pipette canisters in specified locations.
Pipette-Aid
1. Spray liberally with 70% ethanol.
2. Introduce into Laminar Flow Hood. Do not let a Pipette Aid rest on its side, the stand is attached for a
reason. Do not touch the opening of the Pipette-Aid (where the pipette will be inserted).
3. Adjust the speed of the Pipette-Aid ( s, m, f ) by turning the analog buttons. The upper button controls
siphoning speed, lower button controls expelling speed. Adjust the speed of the siphoning/expelling
through depressing the analog buttons.
Note: The amount of depression controls the speed of siphoning/expelling.
4. After use remove Pipette-Aid from Laminar Flow Hood. The UV light from the flow hood
will damage the Pipette-Aid when left for extensive periods of time.
5. DO NOT siphon into the Pipette-Aid. If you do, immediately report it to an instructor
Pipettors
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Be sure that you have selected the appropriate pipettor for the amount desired to pipette.
Pipettor Volume
P5 Under 5.0 µL
P20 1.0 µL -20.0 µL
P200 10.0 µL -200.0 µL
P1000 100.0 µL -1000 µL
1. Spray liberally with 70% ethanol.
2. Introduce into Laminar Flow Hood.
3. Select the appropriate amount to siphon by adjusting the turn knob on the pipettor.
4. Depress pipettor to first point of resistance to siphon, depress past the first point of resistance to expel.
Avoid depressing and releasing the button on the pipettor too rapidly; this can result in liquid being
siphoned up into the pipettor.
5. Always siphon pipettor straight down. Volumes will slightly change if siphoned at an angle.
Pipette Tips
1. All pipette tips must be sterilized in the autoclave prior to work in the Laminar Flow Hood.
2. Spray the pipette tip container liberally with 70% ethanol.
3. Introduce into the Laminar Flow Hood.
4. Lift the top of the container off carefully without touching the pipette tips inside. Contact with the
outside of the container will contaminate your pipette tips.
5. Push the pipettor down firmly on the pipette tip. Pressing excessively hard will result in cracked
pipettors. Not enough pressure and the tip may fall off.
6. Note: The pipette tip containers are not meant to keep the tips sterile once they are opened. If in doubt,
autoclave the pipette tips again before use. They are usually safe as long as they are kept in the hood.
I understand how to work with pipettes. pipettes aids, and pipetters.
Sign____________________________ Super____________________ Date__________
Epindorph (microcentrifuge) tubes
1. Spray the outside of the epindorph tube liberally with 70% ethanol.
2. Introduce into the Laminar Flow Hood.
Make use of the test tube racks found in the lab to hold the epindorph tubes.
Tissue Culture Waste Container
1. It is important to have at least one waste container in the corner of the hood. Usually two are required,
one is for liquids (spent media) and the other for solids (pipette tips)
2. To introduce into the hood, spray liberally with 70% ethanol. Keep in mind to spray ALL surfaces
exposed to air. This includes both inside and outside surfaces of the container.
3. Introduce into the Laminar Flow Hood. When finished it can be removed from the hood and the waste
disposed of appropriately.
4. Expel contents down the drain. Do not expel down the drain if it is a radioactive substance or other
controlled substance.
5. Rinse container with tap water.
6. Dry the container and spray lightly with 70% ethanol.
I understand how to use and dispose of waste in the hood
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Cleaning Up the Laminar Flow Hood
1. The hood needs to be cleaned after each use and thoroughly at least once a week.
2. When finished with the hood, wipe any liquids with a paper towel. Any liquids on the surface of the
hood will dry and potentially contaminate the entire Laminar Flow Hood.
3. Spray liberally with 70% ethanol. Be sure to spray deep inside hood, as well as the edges and front
shield glass.
4. Turn off the airflow.
5. Lower and close the front shield.
6. Turn off fluorescent light and turn on the UV light.
I understand how to clean and shut down the flow hood. .
Sign____________________________ Super____________________ Date__________
Autoclaving
Sterilization
Note: not all autoclaves in the building operate the same make sure you become familiar with the
autoclave we use in the lab.
Key Pad Code for autoclave door is Room___________ Code__________
1. Turn the autoclave on. The knob on the autoclave needs to be turned in a clockwise direction.
2. Wait 5 minutes for the autoclave to warm up.
3. Place the items that need sterilization into the autoclave. Items that need to be sterilized should be
placed in a metal tray. Be sure that nothing touches the walls of the autoclave.
4. Close door and seal. Turn the wheel on the door clockwise until the CLOSE light turns on.
5. Set sterilization and drying times. Drying time not applicable for liquid items.
6. Press either DRY GOODS (dry cycle), or WET GOODS (liquid cycle.) to start sterilization.
7. Check to see if STERILIZATION light turns on.
General Autoclave Times
Item Time Dry Time Cycle
Biohazard Bags 25 min Liquid
Glassware 45 min 30 min Dry
Pipette Tips 35 min 30 min Dry
Canned Pipettes 75 min 45 min Dry
Media, Liquids 25 min Liquid
Octapipette 25 min Liquid
Removing from the autoclave
1. Always wear the heat resistant gloves and bring a cart.
2. Be sure that the sterilization process is complete. The DONE light should be on.
3. Slowly turn the wheel counter clockwise to unseal the autoclave door. This process should be done
slowly and carefully, the steam from the chamber can cause serious burns.
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4. Let the steam gradually come out by opening the autoclave door in small increments. Since the inner
chamber has been pressurized, the contents may burst if the autoclave door is opened rapidly.
5. After opening the autoclave door, remove the contents with caution and place on metal cart. Carrying
the sterilized contents will be dangerous, as items will be at a significantly higher temperature.
6. Cool down sterilized items prior to use. Pipettes are usually cooled down in an oven (in the lab) and
further dried for 24 hours.
I understand how to work the autoclave.
Sign____________________________ Super____________________ Date__________
Pipette Washing
Note: It is very important to do your part to maintain the supply of pipettes. Without pipettes the
lab could not function. Please make sure that you do not hoard pipettes and that you help in the
production of sterile pipettes. This job must be undertaken regularly by all in the lab.
Step One: Preparation
1. DO NOT ALLOW THE PIPETTES TO DRY. This is a cardinal rule in pipette washing. When the
pipettes dry, the contents will attach on to the inner wall of the pipettes, making it very difficult (at times
impossible) to clean.
2. Remove the pipettes from the columns located next to Laminar Flow Hoods.
Removal process should be performed in a sink.
3. Remove the cotton from the pipettes. Either use tweezers to remove cotton, or use air to blow cotton
out. All cotton is recommended to be removed prior to soaking, or it creates a huge mess.
4. Insert the pipettes into a column insert, all tips facing upwards. Pipettes will not be removed from
the crates throughout the washing process. The entire crate will move between the following wash steps.
Step Two: Soap Soak
5. Insert the pipette column insert into the SOAP SOAK. Slowly insert, as the soap solution will be
gradually introduced into the pipettes.
6. The pipettes will need to soak for 30-40 minutes. Pipettes can be left in soap soak for extended periods
of time if time does not permit for step three.
7. Lift the pipette column insert and drain liquids. Once again, slowly drain the liquids from the pipettes
as much as possible.
Step Three: Water Wash
8. Insert the pipette column insert into the WASH column.
9. Turn the water on. Be sure that the water is at a constant flow rate. Turning the water on too fast will
result in an overflowing column.
10 Rinse the pipettes for 60 minutes.
Step Four: Distilled Water Wash
11. Fill the distilled water wash column with distilled water. Fill to 3/4 full.
12. Slowly dip the pipette column insert into the distilled water. Wait for all air to escape from the
pipettes.
13. Lift the pipette column insert to drain distilled water contained in the pipettes. Be sure to drain all
remaining water contained in the pipettes.
14. Repeat above procedure a minimum of 5 times.
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15. Immediately drain all water from the wash column after one complete wash.
Step Five: Drying
16. Empty pipettes from pipette crate into a steel pan. Remember to put the tips pointing upwards.
17. Place in oven to dry for 24 hours. An option would be to air dry as well – however this takes longer.
Step Six: Preparation for Autoclave
18. Remove the pipettes from oven and cool.
19. Stuff the ends of the pipettes with cotton using the cotton-stuffing machine.
20. Start filling pipette canisters with pipettes. Fill the same types of pipettes
It is important to check each canister for light leaks as this may be a source of contamination. An
easy way to check for leaks is to hold the canister top or bottom to your eye then look towards the
light and look for any light breaking the seals.
Place (5mL, 10mL, 25ml etc…) into their own pipette canisters.
Never fill the pipette canisters more than half full one-third is about ideal.
21. Label the pipette canisters appropriately with autoclave tape. The tape should label the contents of the
pipette canister (5mL, 10mL, 10mL, etc…) and be placed over the opening of the pipette canister. Fold
over the tape to make a tab for easy removal of the tape. This will serve as a seal to insure sterility.
Step Seven: Autoclave Sterilization
22. Place all full pipette canisters into an autoclave tray.
23. Autoclave pipettes. Use autoclave gloves to protect yourself from the heat.
24. Remove pipette canisters and cool down before use. Use autoclave gloves to protect yourself from
the heat. Pipettes are usually cooled for at least 24 hours prior to use.
I understand how to wash and sterilize pipettes.
Sign____________________________ Super____________________ Date__________
Media
Complex media will serve as a source of carbohydrates, vitamins, and proteins for the cells in your
culture. In addition, it will provide undefined growth factors, a pH buffering system, and a specific salt
concentration to match osmolarity. Appropriately calculate amounts of each component to be added to the
base medium.
Note that it may be necessary to take out some media from your base medium to make the appropriate
concentrations in the provided containers. All preparations should be done in the Laminar Flow Hood. Be
extremely careful with your aseptic techniques, as contamination of the media will lead to contaminated
cultures.
Base Media
There are many different types of base medium available. Usually contained in 500mL bottles, the
container will be used to comprise our complex media. All supplements are added to this bottle. Some
cell types prefer one type of medium to another, be sure to know all this information ahead of time.
If you are unsure as to the type of media to use for your cells look up the ATCC website and check
their recommendations.
Recipe
1. Base Medium 500mL or 1000mL
2. 10% serum or 20% serum
3. 1.5g/L Sodium Bicarbonate (NaHCO3)
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4. 2.05mM L-Glutamine
Note: Some cells will require fetal calf serum others bovine calf serum. There is a big difference.
Fetal calf is much more expensive than bovine calf. Make sure you check which one you need to
use. We have also Cosmic Calf, which is supplemented bovine calf and is useful for certain types of
cells. There are other types of serum please make sure you use the correct bottle. Make sure it is not
left sitting out on the bench and that when opened you put your name and date on the bottle.
Serum
Serum is the supernatant of clotted blood. Many undefined growth factors are contained, including many
proteins and metal ions. Although Fetal Bovine (Calf) Serum is most commonly used in publications,
there are alternatives for economic purposes.
Types of Sera
Bovine Calf Serum Standard sera used in our lab, a cost efficient alternative to FBS.
Fetal Calf Serum Most commonly used sera in literature. However, FBS is more expensive than BCS.
Cosmic Calf Serum BCS supplemented with extra nutrients, provided exclusively through HyClone.
Equine Serum Serum from horses. Some select cell types prefer this serum.
Example Calculation for 10% supplemented media
1. 435mL RPMI 1640
2. 50mL Bovine Calf Serum
3. 5mL L-Glutamine (200nM stock solution)
4. 10mL Sodium Bicarbonate (7.5% stock solution)
Types of Base Medium
Dulbecco’s Modified Eagle’s Medium Low Glucose/High Glucose
Iscove’s Modified Dulbecco’s Medium
Medium 199 Earle’s
Minimum Essential Medium (MEM)
Nutrient Micture F10 (F12) Hams
Roswell Park Memorial Institute 1640 (RPMI1640)
Dulbecco’s Phosphate Buffered Saline
Earle’s Balanced Salt Solution (EBSS)
Hank’s Balanced Salt Solution (HBSS)
Making up Media
1. Select appropriate medium for cell culture. RPMI 1640 is our lab standard. Unopened media is found
in the walk-in refrigerator.
2. Yellow fluorescent protection bag is needed during storage. Only take the bottle out of the bag during
media use.
3. Remove seal on the media bottle in the hood. (Only for first time use). Flame the lid. Make sure you
label the bottle with your name and date opened.
4. Select appropriate sera. Bovine calf serum and Cosmic Calf Serum are the usual standards for our
laboratory. Sera are found in the freezer in room 863 WIDB Code for door is 12345
5. Thaw in 37ºC water bath. Stir occasionally, the sera may form insoluble
clusters if you do not pay attention. Let sera thaw slowly in the water bath.
Rapid thawing will denature the proteins in the sera. Do not heat above 37oC
6. Remove seal on the serum bottle in the hood. (Only first time use, label bottle with name and date).
Flame the lid.
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Sodium Bicarbonate
This will act as a buffer for pH changes in your culture due to metabolic processes.
1. Stock 7.5% NaHCO3 is found in the refrigerator. Our stock Sodium BiCarb is currently a non-sterile
powder found on the shelf.
L-Glutamine
Is an essential amino acid that needs to be added to the media because it doesn't store very well long term.
A purchased stock solution of L-Glutamine is usually available, but when comprising this from the
powder form, remember to sterilize the solution before use. Do not heat, sterile filter.
1. Stock 205mM L-Glutamine should be stored in –20ºC freezer. This will be diluted down to 2.05mM
L-Glutamine when added into medium.
2. Thaw in 37ºC water bath.
3. Vortex solution until the precipitate dissolves. Leftover precipitate will lead to an altered
concentration.
4. Note that the half-life of L-Glutamine is 2-3 weeks. Reconstitute L-Glutamine by adding 0.5% (of
remaining media volume) every 2-3 weeks.
Every time you make up media you should carry out a media test. This is done by placing 1ml of
the freshly made up complete media in a 15ml tube, screwing on the top tightly, carefully labeling
with your name and date and placing in an incubator. Every time you look at your cells, look at the
tube and check that it has not changed color or become cloudy. When you need to do another media
test, simply take the tube into the hood and add a fresh 1ml from the newly made up media. Make
sure you change the date. This media test tube will last you for the time you are in the lab. Media
should be made up and tested two-three days before it will be used.
Media Test
You will need to perform media test each time you constitute complex media. This will save time and
efforts, as contaminated media will lead to contaminated cell cultures.
1. Examine your media. Look for any turbidity, or color change in media. You can
examine the media under the microscope if desired.
2. Incubate your media. Pipette some media into a 15mL conical vial. Loosen the
cap and let incubate for 24 hours.
3. Agar streak test. You can streak your media onto an agar plate, let sit in
incubator for 24 hours and observe for growth.
I understand how to make up media and perform a media test.
Sign____________________________ Super____________________ Date__________
Cell culture
Suspension Cells
In general terms cultures derived from blood (e.g. lymphocytes) grow in suspension. Cells may grow as
single cells or in clumps (e.g. HL-60’s grow as single cells, as Raji’s like to cluster). These subcultures
are relatively easy to grow, and are well suited for many studies as they are easy to run in assays. For
lines that grow in clumps it may be necessary to bring the cells into a single cell suspension
by centrifugation and re-suspension by pipetting in a smaller volume before counting.
1. View cultures using an inverted phase contrast microscope. Cells growing in exponential growth phase
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should be bright, round and retractile. Hybridomas may be very sticky and require a gentle knock to the
flask to detach the cells. EBV transformed cells can grow in very large clumps that are very difficult to
count and the center of the large clumps may be non-viable.
2. Do not centrifuge to subculture unless the pH of the medium is acidic (color = yellow) which indicates
the cells have overgrown and may not recover. If this is so, centrifuge at 150g for 10 minutes, re-seed at a
slightly higher cell density and add pre-warmed media to the flask.
3. You may check for cell concentration and viability (see Cell Quantification
section).
Note: Cells generally double every 24hrs therefore they will need daily attention. Some cells can be
left for two days however, if diluted back too far the calls can take a long time to come back. Try
where possible to look at your cells every day. NEVER dilute suspension cell concentration to less
that 250,000 cells/mL, as cell line may not recover at such low concentrations. Generally cells need
cell-cell interaction to survive which is why it is a bad idea to dilute to less than 250,000 cells per ml
Adherent
(Monolayer) Cells
Most cells found in the body are not in suspension. Cell Adhesion Molecules (CAMs) and desmosomes
adhere some cells to the surfaces of the culture flasks. Since adherent cells grow in one layer (monolayer),
cell morphology is easy to observe, and treatments applied to these cultures are homogenous.
Unfortunately, growth curves are very difficult to perform, as it is hard to take a sample culture
without disrupting the cells. Adherent cell lines will keep growing until they have covered the surface
area available, or until the medium is depleted of nutrients. At this point the cell lines should be subcultured in order to prevent the culture from dying. The first step in the subculture of these cells is to
bring them into suspension. The degree of adhesion varies from cell line to cell line but in the majority of
cases proteases (trypsin), are used to release the cells from the flask. However, this may not be
appropriate for some lines where exposure to proteases is harmful or where the enzymes used remove
membrane markers/receptors of interest. In these cases cells should be brought into suspension into a
small volume of medium mechanically with the aid of cell scrapers.
Trypsinization of Adherent Cells
1. Aspirate the medium and discard.
2. Wash cells with pre-warmed PBS or HBSS, aspirate and discard. The volume of PBS of HBSS should
be approximately the same as the volume of medium used for culturing cells. This will remove the
chelating ions from serum that will inhibit trypsin enzymatic activity.
3. Repeat above procedure.
4. Add enough 1x trypsin-EDTA solution (pre-heated to 37oC) to cover the monolayer and rock the
flask 4-5 times to coat the monolayer. (Depending on the size and type of flask the amount of trypsin
required will vary, in a 25ml flask usually 2-3 mls are required, after adding and rocking the surplus can
be removed). Our stock solution is 10x trypsin-EDTA solution. Dilute to 1x concentration using PBS or
HBSS.
5. Loosen the flask cap and place the flask in the incubator for 1-2 minutes.
6. Remove flask from incubator, tighten flask cap and firmly rap the side of the flask with palm of hand
to assist detachment. If cells have not dislodged, loosen the flask cap and return the flask to the incubator
for a few minutes. If the cells dislodge then aspirate them gently using a pipette (25ml, or 10ml) to
separate the cells.
Note: do not leave the cells in trypsin for extended periods of time as trypsin
will kill the cells.
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Overly confluent cultures, senescent cells and some cell lines may be resistant to trypsin. Some cells lines
may detach when repeatedly pipetteing the cells up and down in media. This should be done as gently as
possible to avoid damaging the cells.
7. Once dislodged, resuspend the cells in pre-warmed growth medium containing serum. The
magnesium and calcium ions found in the serum will deactivate the trypsin.
8. Remove about 100µL of the cell culture and perform a cell count (see Cell Quantification section).
9. If your cell concentrations are low or if you are only seeding one flask, centrifuge the cells using a
15mL conical vial at 1000rpms for 10 minutes.
10. Transfer the required number of cells to a newly labeled flask containing pre-warmed medium. Refer
to online references for recommended seeding densities, such as the ATCC.
11. Incubate the cell culture.
Troubleshooting
1. Some cultures while growing as attached lines adhere only lightly to the flask, thus it is important to
ensure that the culture medium is retained and the flasks are handled with care to prevent the cells
detaching prematurely.
2. Although most cells will detach in the presence of trypsin alone, the EDTA is added to enhance the
activity of the enzyme.
3. Trypsin should be neutralized with serum prior to seeding cells into new flasks otherwise the cells will
not attach.
Placing the flask directly on the incubator may dislodge the cells faster.
I understand how to dilute cells suspension and monolayer.
Sign____________________________ Super____________________ Date__________
Note: Do not place cultures on the top shelf of the incubator this is where the Co2 comes in and can
cause air turbulence. This can lead to contamination. Place your cultures on the appropriate shelf
so that the air will flow through the shelf holes and over you flasks in an even manner.
Cell Quantification
Trypan Blue Exclusion
In order to find the concentration of cells in our flask we take a representative sample. From that
representative sample we are able to calculate cell concentration and viability. This will give an overview
of the trypan blue exclusion method, which is a standard in manual cell counting. A trypan blue solution
is added to the cell culture, and dead cells will stain blue. The live cells will take up the dye, then
pump it out through the membrane immediately, hence do not stain blue.
Trypan Blue Constitution
1. 0.4% Trypan Blue, 0.002% Sodium Azide in double distilled water, or PBS.
2. Prepare a 50mL conical vial with a Whatman filter.
3. Weigh out 0.125g of Trypan blue and add to top of the filter.
4. Weigh out 0.015g of sodium azide and add to top of filter
5. Slowly add 50 mL of ddH2O to the filter.
6. Centrifuge at 1300 rpm for 10 minutes when filtration is complete.
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Trypan Blue Exclusion Method
1. Siphon and expel media in the flask two to three times. This will make your cell culture homogenous
throughout. Be careful not to froth the cell media.
2. Pipette out less than 0.5 mL of your cell culture into an epindorph tube. This should be performed in a
Laminar Flow Hood to insure the flask is protected from contamination.
3. Seal the epindorph tube, put culture flask back into the incubator.
4. Remove epindorph tube from the Laminar flow hood.
5. Cut out a small strip of parafilm.
6. Vortex the epindorph tube.
7. Using a P20/P200 pipettor, pipette out 20µL of culture and expel on a strip of parafilm. Discard pipette
tip into normal trash receptacle.
8. Using a P20/P200 pipettor, pipette out 20 µL of Trypan blue and mix with the cell culture on the
parafilm. Mix carefully by pipetting up and down without producing any bubbles.
Note: Use from the top layers of the stock trypan blue, the bottom 1/4 usually contains a significant
amount of garbage.
9. Wait 2-3 minutes. This will allow the dead cells to stain, and the live cells to pump out the dye.
10. Pipette mixture onto a hemocytometer. Slowly pipette the mixture in, and let the capillary action fill
the hemocytometer. Filling in too much will change the volume of your sample, and alter your
calculations.
11. Count cells manually by using the cell counter. Dead cells will stain blue, while live cells should be
translucent.
12. Calculate the amount of cells per milliliter. The volume within the four squares of the hemocytometer
is 0.4mm3 (0.4mL).
Also remember to double the actual count, due to the 50/50 dilution of the culture with trypan blue.
13. When finished, rinse hemocytometer and glass cover slip with distilled water and dry with Kim
Wipe. Be careful not to break the cover slips.
14. Trypan blue is toxic and is a potential carcinogen. Protective clothing, gloves and face/eye protection
should be worn. Do not breathe the vapor.
15. The central area of the counting chamber is 1mm2. This area is subdivided into 25 smaller squares
(1/25mm2). Each of these is surrounded by triple lines and is then further divided into 16 (1/400mm2).
The depth of the chamber is 0.1mm.
16. There are several sources of inaccuracy:
a. The presence of air bubbles and debris in the chamber.
b. Overfilling the chamber such that sample runs into the channels or the other chamber
c. Incomplete filling of the chamber.
d. Cells not evenly distributed throughout the chamber.
e. Too few cells to count. Centrifuging the cells, resuspending in a smaller volume and recounting can
overcome this.
f. Too many cells to count. This can be overcome by using a higher dilution factor in trypan blue e.g.
1:10
I understand how to count cells using Trypan blue.
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Sign____________________________ Super____________________ Date__________
Some Useful Calculations
Total time = end of exponential à start of exponential.
N = concentration at end of exponential
N0 = concentration at beginning of exponential
X= number of generations
N = N02x
Log N = log N0 + x log 2
Generation time = Total time / number of generations
Cryopreservation of cells.
Cell Freezing
Cryopreservation of cultured cell lines is essential in the cancer research laboratory. The aim of
cryopreservation is to enable stocks of cells to be stored to prevent the need to have all cell lines in culture
at all times. It is invaluable when dealing with cells of limited life span. The other main advantages of
cryopreservation are:
a. Reduced risk of microbial contamination
b. Reduced risk of cross contamination with other cell lines
c. Reduced risk of genetic drift and morphological changes
d. Work conducted using cells at a consistent passage number
e. Reduced costs (consumables and staff time)
There has been a large amount of developmental work undertaken to ensure successful cryopreservation
and resuscitation of a wide variety of cell lines of different cell types. The basic principle of successful
cryopreservation is a slow freeze and quick thaw.
Standard Freezing Medium
1. 92% fetal calf serum and 8% sterilized DMSO
Cell Freezing
1. Check that cells are healthy, not contaminated, and have the correct morphological characteristics.
2. Change the medium 24 hours prior to freezing the cells. Both monolayers and suspension cells should
be at about ~500,000 cells per milliliter. To freeze cells down concentrate them to about 5 million per ml
of freeze down media.
Cells must be in exponential growth phase when frozen.
3. For monolayers, trypsinize cells, resuspend in medium containing serum, pellet the cells by
centrifugation at 200G for 10 minutes, and resuspend the cells in freezing medium.
4. Transfer 1mL of cells into each cryotube. Label the vials with name of cell line,
date, and growth medium.
5. Place the freezing vials into a polystyrene box and freeze at the rate of 1~3ºC/min. The box is placed
in the refrigerator for 30-40 minutes, and then placed in the –20ºC freezer for 1.5-2 hours. Then place the
cells in the –80ºC overnight before transferring them into the liquid nitrogen.
Note: cells need to be frozen slowly (so ice crystals don't form and fracture the cells, because flash
freezing induces crystal formation whereas slow freezing prevents crystal formation)
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Troubleshooting
1. The most commonly used cryoprotectant is dimethyl sulphoxide (DMSO) however, this is not
appropriate for all cell lines e.g. HL60 where DMSO is used to induce differentiation. In such cases an
alternative such as glycerol should be used. WEAR GLOVES WHENEVER YOU USE DMSO.
2. It is essential that cultures are healthy and in the log phase of growth. This can be achieved by using
pre-confluent cultures (cultures that are below their maximum cell density) and by changing the culture
medium 24 hours before freezing.
3. The rate of cooling may vary but as a general guide, a rate of between –1ºC and –3ºC per minute will
prove suitable for the majority of cell cultures.
4. When freezing in the –80ºC freezer it is important to have an allocated section for cell line freezing so
that samples are not inadvertently removed. If this happens at a crucial part of the freezing process then
viability and recovery rates will be adversely affected.
Make sure you close the door tightly on the -80oC freezer
I understand how to freeze cells down both suspension and monolayer.
Sign____________________________ Super____________________ Date__________
I have frozen the following cells down:
Cell type
Date/ name of
Researcher
Location Tank,
stack, and box
Media cells
frozen in
Date cells
brought up
and tested.
Sign
Cell Thawing
Many cultures obtained from a culture collection, such as ATCC, will arrive frozen and in order to use
them the cells must be thawed and put into culture. It is vital to thaw cells correctly in order to maintain
the viability of the culture and enable the culture to recover more quickly. Some cryoprotectants, such as
DMSO, are toxic above 4ºC therefore it is essential that cultures are thawed quickly and diluted in culture
medium to minimize the toxic effects.
1. Run tap water to 37°C and place a beaker under the tap, or use water bath set at 37oC
2. Remove a vial of frozen cells from liquid nitrogen, and place in beaker containing 37°C. Wear
protective goggles and gloves when thawing vials that have been stored in liquid nitrogen. Vials may
explode when removed from liquid nitrogen.
3. As soon as the solidified cell mass in the cyrotube begins to liquefy, immediately remove from water
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bath. Leaving the cells in the freezing medium (containing DMSO) is toxic, and it will kill the alreadyweakened cells.
4. Spray and wipe the outside of the vial with 70% ethanol.
5. Prewarm a suitable amount of medium to 37°C in an appropriately sized flask. The size of flask
depends on the cell type as well as the desired cell density.
6. Slowly pipette the cell suspension into the prewarmed medium. Mix during addition of the cells to the
medium. Note: Immediate removal of DMSO may sometimes be necessary, especially for suspension
cells. Following the addition of the thawed cell suspension to the prewarmed medium, the cells
should be centrifuged (1000rpm for 10 minutes) and resuspended in fresh, prewarmed medium.
7. Incubate cells at 37°C overnight, and then change medium.
I understand how to bring cells out of cryogenic storage both suspension and monolayer.
Sign____________________________ Super____________________ Date__________
Consent Form
I, ___________________________ hereby state that I have read and understood all of the presented
material in this manual. I will abide by the safety standards and practices set forth by this manual, and I
will be fully responsible for all my actions
Signature __________________________________________________
Date _______________________________________________________
Pass off:
In order to pass of this module you will be required to
1) Have completed the manual to this point.
2) Observed three labbies doing tissue culture for about one hour each.
A. I observed _____________________ on date__________________ as he/she performed tissue
culture.
I feel that ______________________ has understood the basics of tissue culture and has closely observed
me of about one hour.
Super ________________________ Date___________________
B. I observed _____________________ on date__________________ as he/she performed tissue
culture.
I feel that ______________________ has understood the basics of tissue culture and has closely observed
me of about one hour.
Super ________________________ Date___________________
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C. I observed _____________________ on date__________________ as he/she performed tissue
culture.
I feel that ______________________ has understood the basics of tissue culture and has closely observed
me of about one hour.
Super ________________________ Date___________________
3. Produce a growth curve carried out over 48hrs. You will need to set up cells count then time zero and
then count them on a regular basis throughout the next 48hrs then plot a growth curve of cell count
against time.
Cells type used: ______________________
Media Type (full details of media required)
_____________________________________________________________________________________
_____________________________________________________________________________________
________________________________________________________
Initial cell count ___________________ Date_____________ Time_____________________
Cell count ________________________ Date______________ Time_____________________
Cell count ________________________ Date______________ Time_____________________
Cell count ________________________ Date______________ Time_____________________
Cell count ________________________ Date______________ Time_____________________
Cell count ________________________ Date______________ Time_____________________
Cell count ________________________ Date______________ Time_____________________
Cell count ________________________ Date______________ Time_____________________
Cell count ________________________ Date______________ Time_____________________
Cell count ________________________ Date______________ Time_____________________
Graph of growth curve attached:
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4. Cryogenic preservation.
On (date)________ I froze the following cell line down
Cell line__________________________ Cell count per ml____________________
Number of cells in vial______________ Freeze media_____________________
_________________________________________________________________
Time in Fridge______________ Time in -20oC freezer________________
Time in -80oC freezer __________________
Transferred to Cryogenic freezer on (date) __________ Freezer number _______
Stack number _________ Box number__________ Number in box_______
Logged in catalogue as _________________________________________________
5. Cryogenic retrieval
On (date)___________ I retrieved the following cells from the liquid nitrogen and established a culture.
Cell type _____________________ From Cryogenic freezer on (date) __________
Freezer Number _______Stack Number _____ Box Number___Number in box_______
Logged out of catalogue as _________________________________________________
I have verified that ________________________________ removed and properly established a culture of
sterile cells from the cryogenic storage, and that I have observed them over the past week and found them
to be contamination free.
Date_____________ Super ________________________________
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