Cell Culture Protocols and Notes
1.0 Preparing Necessary Reagents
1.1 Media Components and Other Necessary Solutions
1.1.1
Heat inactivated fetal bovine serum (FBS)
We purchase heat inactivated fetal bovine serum which should be
sterilized and tested for mycoplasm contamination. This solution
will arrive frozen on dry ice. Thaw the solution in the water bath
and then separate into 50mL aliquots (large falcon tubes). The
aliquots can be refrozen.
1.1.2
Penicillin streptomycin
We purchase the penicillin streptomycin from Sigma. This
antibiotic is used to prevent bacterial growth in the media.
Penicillin-G prevents the final formation of the bacterial cell wall.
Streptomycin binds to the 30S subunit and cause misreading in the
of bacterial DNA. The solution will arrive frozen on dry ice. Thaw
solution in the water bath and then separate into 5mL aliquots
(small falcon tubes). The aliquots may be refrozen until needed.
The stock solution (P0781 Sigma Aldrich) contains 10,000 units of
penicillin-G, and 10mg of streptomycin per mL. The final
concentration in our media is ~ 100U/mL penicillin and 100ug/mL
streptomycin.
1.1.3
L-glutamine
L-glutamine is an essential amino acid required for most if not all
cell culture media formulations. The glutamine serves as an energy
source for rapidly diving cells, and those which must synthesize
large quantities of nucleic acids and proteins. L-glutamine should
be in the DMEM which we order, however it does degrade over a
period of 2-6 weeks at 4ºC, and with exposure to light. If you are
not going through media very fast it will be necessary to
supplement your media with fresh L-glutamine after about 4
weeks, or if you note a large change in the rate of growth of your
cells. Ammonia is a byproduct of the L-glutamine degradation, and
old media should be discarded (> 6 weeks) to avoid harming the
cells.
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1.1.4
Non-essential amino acids (NEAA)
Non-essential amino acids are added to media to increase the
nutrients available to the cells. The stock solution is very acidic
(pH 1.5 – 1.7), and thus if added to a non-buffered solution you
may want to check the pH to ensure it is not too acidic. The
solution contains glycine, L-alanine, L-asparagine, L-aspartic acid,
L-glutamic acid, L-proline and L-serine.
1.1.5
HEPES
HEPES is used to keep the pH stable between 7.2 – 7.6. We add
HEPES to media which is used for imaging or when cells are
removed from the incubator for extended periods of time.
Excessive cellular metabolism or rapidly growing cultures may
acidify quickly and small amounts of HEPES can help stabilize the
cells.
1.1.6
0.25% Trypsin solution
Stock solutions of trypsin are purchased in 2.5% strength, and kept
in the freezer. To prepare working solutions we typically add 1.0
mL of the trypsin to 9.0mL of PBS (-) (contains no Ca2+/Mg2+).
These solutions can be prepared in the small 15mL falcon tubes
and placed in the freezer. When needed thaw the solution in the
water bath.
1.1.7
Citric saline solution
Some cell lines are much less adherent than the CHO cells, and
thus require less coaxing in order to passage them. Citric saline
solution works well for HEK293T and AD293 cell lines. We begin
by first making a 10X stock solution of citric saline and then
diluting to a 1X working solution. The following recipe is for
500mL of the 10X solution:
1.35M KCl (MW 74.5 g/mol  50 g)
0.15M sodium citrate (294 g/mol for trisodium salt  22g)
Dissolve the above in 500mL of pure Millipore water. Sterilize the
solution by autoclaving. The stock solution can be kept for up to 3
months at room temperature. Dilute this solution by placing 5mL
of the 10X solution in 45mL of pure sterile water. This solution
can be kept in the fridge.
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1.1.8
90% FBS/10% DMSO freezing media
Prepare this solution by thawing a 50mL aliquot of the FBS.
Remove 5.0mL of the media, and replace it with 5.0mL of DMSO.
We use DMSO to prevent ice crystal formation, which would
result in damage to the cells during the freezing process. Caution:
DMSO can be toxic to some cells lines, it is also possible to use
10% glycerol in these cases.
1.1.9
4% paraformaldehyde
This solution is used to fix cells for imaging purposes. The 4%
refers to 4g per 100mL of pure water. The mixture will be very
cloudy, heat the solution carefully on a low setting (3) with the cap
loose swirling occasionally until all the powder has dissolved.
Take care not to breath in fumes as they are toxic. This solution
should also be kept frozen. When preparing paraformaldehyde
avoid breathing in the powder, wear protective gloves, lab coat,
safety goggles.
1.1.10 Geneticin (G418 Sulfate)
Geneticin is an aminoglycoside antiobotic which is toxic to yeast,
bacteria, protazoans and mammalian cells. It is possible to
introduce a genetic resistance to cells (on transposons Tn601(903)
and Tn5). The antibiotic interferes the function of the 80S
ribosomes and disrupts protein synthesis in non-resistant cells.
According to some sources potency is achieved at concentrations >
450 g/mL, these higher concentrations can be used for selection,
while lower concentrations around 200 g/mL are suitable for
maintenance of an already stable cell line. For our CHO (+) media
we add 0.235g of the Geneticin to the media yielding a total
concentration of 370g/mL.
1.1.11 0.1M Bicarbonate Buffer
This buffer is used in for reactions between succinimidyl esters and
amines forming amide bonds. The basic pH of this buffer is used to
keep the -amino group in the non-protonated form during the
reaction. You can not use buffers which contain amine groups (ie.
TRIS) for this type of reaction because the buffer itself will
compete with the amine and react with the activated dye.
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The following recipe will yield you a 0.2M bicarbonate buffer with
a pH ~9.2. Depending on the type of ester you will need to adjust
the pH using the pH meter and the 1.0M HCl solution.
Obtain sodium carbonate (Na2CO3) and sodium bicarbonate
(NaHCO3) from the shelf. You will actually prepare two different
stock solutions and them mix them together to obtain the buffer. It
is wise to make extra stock solution so that you do not have to do
this step each time.
Preparation of the 0.2M Na2CO3 stock solution:
MW = 105.99 g/mol
Vol = 0.500 L
C = 0.2 mol/L
n = 0.1 moles
mass needed = 10.56 g
Preparation of the 0.2M NaHCO3 stock solution:
MW = 84.11 g/mol
Vol = 0.500 L
C
= 0.2 mol/L
n
= 0.1 moles
mass needed = 8.41 g
Preparation of final buffer mixture:
Mix 92.0 mL of NaHCO3 with 8.0 mL of Na2CO3 to yield a
final volume of 100.0 mL.
Adjust the pH using HCl to obtain the desired pH level.
This solution should be made fresh every month.
1.1.12 LB Broth for Maxi Preps
We purchase LB broth powder in small pre-weighed packages.
Each package is suitable for preparing 500mL of media. Each maxi
prep requires about 300mL of LB broth. Once prepared sterilize
the media using the programmed “liquids” setting on the autoclave
leaving the cap slightly un-screwed. Once autoclaved media can be
stored at room temperature for up to 3 months, as long as it
remains un-opened.
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1.1.13 X
1.2 Media for Cell Culture
We are now using washable and reusable glass bottles to prepare our media and
our PBS solutions. The media bottles have tops which are compatible with the
disposable sterile filters from VWR. These bottles should be washed with bleach
and water (3x with bleach and 9x with water), then dried and autoclaved. After
being autoclaved you can spray the outside with EtOH and bring into the laminar
flow cabinet. I would recommend opening the bottle and placing the lid upwards
on the bottom of the cabinet and turn on the UV light for 10 minutes just to be
sure.
1.2.1
CHO K1 (-) Media
This media is suitable for non-transfected CHO K1, or transiently
transfected CHO cells which do not have any antibiotic resistance.
To prepare the media will need the following:
1. 430 mL of DMEM
2. 50 mL of heat inactivated FBS
3. 5 mL of Pen/Strep
4. 5 mL of Non-essential amino acids
5. 10 mL of L-glutamine
Mix all reagents together and run through a sterile filter.
1.2.2
CHO K1 (+) Media
This media is suitable for CHO K1 cells which stably express fluorescent
proteins and are resistant to geneticin.
To prepare the media you will need the following:
1. 400 mL of DMEM – Low Glucose
2. 50 mL of FBS
3. 5 mL of Pen/Strep
4. 5 mL non-essential amino acids
5. 10 mL of L-glutamine
6. 0.235g of geneticin
The geneticin is not immediately soluble in the DMEM. It is
recommended that you add the geneticin first to the warm DMEM to
increase the solubility and maximize the concentration of geneticin in the
sample.
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1.2.3
AD293T Media
These cells are variants of HEK 293 cells which are slightly more
adherent. To prepare this media you will need the following:
1. 445 mL DMEM - High Glucose
2. 50 mL FBS
3. 5 mL Pen-Strep
If this media is for the AD291 NK1 cells which stably express the NK1
receptor reduce the amount of DMEM by 3 mL and add 3 mL of G418
antibiotic.
If we do not have high glucose media you can supplement low glucose
media with 1.750g of D-glucose to achieve the necessary amount of
glucose.
1.2.4
HEK293 Media
To prepare this media you will need the following:
1. 445 mL DMEM – High glucose
2. 50 mL FBS
3. 5 mL Pen-Strep
If we do not have high glucose media on hand you can supplement low
glucose media with 1.750g of D-glucose to achieve the necessary amount
of glucose.
1.2.5
HeLa Media
HeLa media is the same as HEK293 media described above. HeLa cells
can be easily contaminated, and have been shown to contaminate other
cell lines. For this reason it is advised that if you have both HEK293T and
HeLa cells going, separate the media into two separate bottles to avoid
cross-contamination of the cell lines. It is also possible to use CHO(-)
media with the HeLa cells again, be very careful to avoid contamination of
the HeLa cells
1.3 Imaging Medias
Growth media contains phenol red as an indicator allowing us to observe large
changes in the pH of the media. Phenol red does have both substantial
autofluorescence and can act as a screen preventing the fluorescence from
other proteins from being observed. This is less problematic when examining
the basal membrane of adherent cells using an inverted microscope, but in
general it is good practice to image the cells in media which is phenol red free
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(-). Because we often remove the cells from the tightly controlled CO2
environment we want to prevent the buildup of toxic metabolites which will
limit the life of the cells for imaging. We do this by adding HEPES to the
buffer at low concentrations (~ 20 – 25 mM). We often add radical scavenger
such as ascorbic acid to help reduce general phototoxicity. And finally,
imaging media will often be FBS (-) free. Cells are often more responsive to
stimulation with drugs when they have been deprived some FBS for at least 1
hour (but up to 12 hours is often ideal). This is particularly true for the EGFR
containing cells. In order to get a nice dispersion of EGFR on the membrane
we will starve the cells (of FBS) for 12 hours prior to imaging.
1.3.1
For CHO K1 EGFR-GFP cells
EGFR can cluster in response to light stimulation. The addition of
N-acetyl-L-cystein (NAC) can prevent this from occurring.
However we have recently discovered that for transactivation
studies this component shuts off the necessary pathways and thus
for transactivation work you can omit the NAC. The imaging
media is made up of the following components.
The HEPES and other sterile solutions can be aliquoted into 1
large falcon tube (50mL) in the laminar flow hood and then added
to the Hanks Balanced Salts Solution. The media can be prepared
outside the laminar flow cabinet because you will need to adjust
the pH to 7.4. Once the pH has been adjusted use a filter and clean
media bottle to sterilize the media.
1.
2.
3.
4.
5.
6.
1.3.2
400 mL of Hanks Balanced Salts Solution
10.6 mL of HEPES
5.0 mL of PenStrep
5.0 mL of MEM NEAA
0.2125g ascorbic acid (shelf)
0.3329g N-acetyl-L-cysteine (fridge)
For AD293/HEK cells
We can prepare an imaging media from phenol red free (-) high
glucose media. For imaging purposes we once again omit the FBS,
but will add the Pen-strep to prevent bacterial growth. Importantly,
we will have to add HEPES buffer.
1. 400 mL of phenol red free high glucose media
2. 10.6 mL of HEPES
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3.
4.
5.
6.
5.0 mL of PenStrep
5.0 mL of MEM NEAA
0.2125 g ascorbic acid
0.3329 g N-acetyl-L-cysteine
2.0 Thawing
Cells are typically frozen in 90% FBS and 10% DMSO freezing media. When cells
are removed from either the -80 or the N2(l) freezer you must thaw the cells quickly.
In order to prevent damage to the cells it is wise to place 8.5 mL of the appropriate
media (warmed to 37ºC) in a 15 mL falcon tube. Thaw the cells at 37ºC in the water
bath until about ½ of the sample has melted. Wipe down the vial with ethanol and add
to the 8.5 mL of media. Place the cap on the falcon tube and invert to mix. Now there
are one of two options.
1) Place the tube in the centrifuge and spin down for 5
minutes on setting 3. Suck out the media and resuspend the cells in 10 mL of the appropriate media.
Divide the 10mL of suspended cell solution into two
small culture flasks. Make sure to label the flask with
the passage number, date and cell line.
2) Divide the 10mL of suspended cell solution into two
culture flasks. Make sure the label the flask with the
passage number, date and cell line. Place flasks in the
incubator.
Notes about these options: Some people are very concerned that the DMSO can harm
the cells, and thus spinning down the cells and removing the media helps the cells
recover from freezing faster. It has been my experience that the second method
actually works better for our cells.
After 24 hours check the dishes to see whether the cells have become adherent. If
there is a substantial amount of dead cells in the flask it is a good idea to switch the
media to avoid poisoning the cells which are viable. The cells can take up 1 week to
recover from the thawing process. Depending on how fast the cells grow you may
need to passage the new cultures in a few days, or you may simply need to keep
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changing the media every few days. Changing the media helps ensure that while the
cells are growing and dividing there is sufficient nutrients, and limited build up of
toxic metabolites in the media.
3.0 Freezing
Cells should be frozen when at low passage number, healthy and confluent. When
you thaw cells check to see how many tubes we have of that cell line, if we have less
than 2 tubes you must freeze new cells as soon as possible.
To freeze we grow the cells up in the large 75cm2 culture flasks. During normal
passaging we add 5mL of freshly diluted cells to the flask and add 20mL of the
appropriate media. Monitor the cells and freeze the cells when they are ~80 – 90%
confluent. Some protocols suggest changing the media about X hours before you wish
to freeze to ensure all the cells are in the X phase of growth. Once the cells are ready
for freezing use the following protocol.
1. Warm PBS (-), appropriate media and freezing media in water bath.
2. Pre-label 5 tubes with the following information:
a. Cell line and any expressing proteins (ie. CHO K1 EGFR-GFP)
b. Passage number
c. Date
3. Have waiting a small cooler full of dry ice. Place an empty styrofoam 15 mL
falcon tube holder in the dry ice and pack dry ice around the styrofoam. This will
create a way for the cells to freeze slowly and uniformly over a 1 hour period.
Some people suggest flash freezing the cells, but we find this method is suitable
for preparing the cells for nitrogen freezing that same day.
4. Remove the media from the flask using suction.
5. Rinse with ~ 5 mL of PBS (-).
6. Add about 1 mL of trypsin enough to cover the whole bottom of the dish. Remove
half of the trypsin immediately and place the cells in the incubator for 5 minutes
until they detach. For HEK or AD293 cells use citric saline, and do not place the
cells in the incubator.
7. Add 10mL of appropriate media to suspend cells. Using the 10mL pipette transfer
all the suspended cells into a small 15mL falcon tube.
8. Centrifuge at setting 3 for 5 minutes.
9. There should be a large palette at the bottom of the tube.
10. Suck up 9mL of the freezing media and add it to the tube forcefully enough to
dislodge the pellet at the bottom. Mix well by sucking up and dispensing the
solution a few times.
11. Add 1.8mL of the suspended cells in freezing media to each of the labeled Nunc
cryotubes. Place the cap on tightly.
12. Place each tube in the empty styrofoam holder in the cooler of dry ice.
13. Allow to freeze until solid. This should take between 1 – 2 hours.
14. Once frozen the cells can be stored in N2(l). To do this you must first wrap each of
the tubes in cryotubing.
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15. Prepare the cryotubing by cutting the long tubing into appropriate lengths and
sealing one end.
Diagram: Cutting the cryotubing
Diagram: Sealing one end of the cryotubing
16. Place a frozen Nunc tube inside the cryotubing which as been sealed at one end.
17. Holding the sealed end (with fingers or with tweezers), quickly pass the body of
the tube over the flame to allow the tubing to more or less suck into the Nunc tube
getting rid of as much air as possible. Then pass the open end of the cyrotubing
over the flame and seal with tweezers. Be careful not to melt the frozen cells,
which may cause damage.
18. Immediately place the cells back into the dry ice until all tubes have been sealed.
19. Place the sealed tubes into the N2(l) dewer. Note the canister number and tube
number on a freezing sheet.
4.0 Passaging and Maintaining a Culture
General Notes:
1. Cells are passaged when they reach 70 – 90% confluency (density). We do not
want the cells to become 100% confluent, this can lead to detachment of the
culture (in the case of HEK/AD293 cells), reduced protein expression
(particularly for fluorescent constructs and non-native proteins), or death of the
culture. Additionally, we do not want to passage the cells when they are
2. Ideally we want to carry out experiments on cells with low passage numbers.
Over time the protein expression levels within a cell line can change, which may
affect the outcome of the experiment. Sudden decrease in the growth of the
culture may suggest that the passage number is getting too high to use reliably.
Additionally, high passage numbers may dramatically reduce the efficiency of a
transfection.
3. General notes about getting ready to passage cells and work in the laminar flow
cabinet:
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a. Place all the necessary reagents in the water bath and warm to 37ºC. Take
care to only warm the solutions for a short period of time. Excessive light
and heat will degrade many of the reagents in the media. Do not leave
media in the water bath for more than 30 minutes at a time.
b. Turn on the light and blower on the face of the laminar flow cabinet.
Spray the cabinet with EtOH and wipe with a paper towel.
c. Spray all bottles and supplies with 70% EtOH liberally and transfer into
the laminar flow cabinet.
d. Loosen all caps and prepare area for working taking care to ensure that all
materials are spread out around the hood to avoid passing your hands and
arms over top of anything.
e. When finished remove all unnecessary materials, tightly cap all reagents
and replace in the fridge/freezer. Spray down the hood with 70% EtOH.
4.1 Passaging adherent cell lines
For adherent cells (CHO K1 and HeLa) a 0.25% solution trypsin is used to detach
the cells from the plastic culture flask. In order to passage these cells you will
need the following: appropriate media, PBS (-) and 0.25% trypsin.
1. Remove old media using vacuum filtration.
2. Rinse flask with 2mL of PBS (-), and remove with vacuum filtration
3. Add 1.0mL of 0.25% trypsin to flask, swirl gently and immediately remove
0.8mL of the trypsin.
4. Place the cap back on the flask tightly and place in the incubator for up to 5
minutes. Take care not to over-tryprinize the cells as this may cause
irreversible damage to the culture.
5. Check the cells after 3 minutes to determine how many cells have detached. If
the cells are having trouble detaching you may need to tap the dish on the heel
of your palm to encourage them to pop up. Once the cells are round and
popped off the flask transfer the flask back into the laminar flow cabinet.
6. Add 10mL of fresh media to the cells. Suck up this media and dispense a few
times to dislodge any remaining cells from the surface of the flask. This will
also help to adequately disperse the cells to avoid clumping in the new flask
(or MatTekimaging dishes, this step is critical for preparing samples for
imaging purposes).
7. Obtain a fresh 25-cm flask from the cubbord and transfer to the laminar flow
cabinet
4.2 Passaging non-adherent/semi-adherent cell lines
This includes HEK 293 and AD293 cells. Passage as you would the adherent cell
lines (detailed in section 4.1), using citric saline instead of trypsin. One other
modification is that once you add the citric saline you typically do not need to
place the dishes in the incubator to help the cells detach.
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5.0 Transfecting Cells
5.1 Transfections using Lipofectamine 2000
Lipofectamine 2000 is used here to introduce siRNA or plasmid DNA by a
process called lipofection. This process makes the cell membrane porous and
allows the DNA to cross into the cytoplasm. For transfections we typically use
OPTI-MEM which is a serum free media. The presence of serum in regular media
can bind with the DNA and reduce the efficiency of the overall transfection.
1. For transfection grow cells to 80% confluency in a 6 or 12 well plate. Conditions
below are for the 6 well plates. For 12 well plates divide all the volumes and
masses by ½.
2. Transfections are done in the laminar flow cabinet so take care to preserve the
sterile environment.
3. 2 hours prior to transfection remove the growth media (DMEM) from each of the
wells. Replace this with 800L warm OPTI-MEM solution. By serum starving the
cells for a short period before introducing the plasmid DNA you will improve the
efficiency of the transfection.
4. For each well you have you will need two small sterile eppendorf tubes, labeled
tubes A and tubes B.
5. Put 100uL of OPTI-MEM in each of the tubes
6. In tubes A aliquot the necessary volume of DNA (0.5 – 1.0 g works best),
however you may add as much as 4.0g of DNA to a 6 well plate (1.6g of DNA
to a 12 well plate). Mix well by tapping the bottom the tube to ensure adequate
distribution of the DNA
7. In tubes B you will place the necessary volume of lipofectamine transfection
reagent. For each 1g of DNA in tube A you will need 5L of lipofectamine. An
example is given below for calculating the amount of lipofectamine required.
Importantly, you should not leave the lipofectamine in the OPTI-MEM for more
than 5 minutes before mixing with the DNA in tube A.
a. Example: If you have 0.75 g of DNA in tube A you will need to add
3.75 L of lipofectamine solution to your tube B
8. Mix the lipofectamine well in the 100L of OPTI-MEM and transfer to tube A
(DNA solution). Add dropwise to the DNA solution and then mix well with the
pipette.
9. Leave the DNA-lipofectamine solution for 20 – 25 minutes at room temperature
in the laminar flow cabinet. Do not leave the solutions more than 25 minutes.
10. Add the contents of 1 of the tubes to 1 of the wells in your dish dropwise. Do this
for all the wells you wish to transfect. Place the lid on the dish and swirl gently
with your hand.
11. Place the dish in the incubator for 4 hours.
12. After 4 hours suck out the media and replace with geneticin free media (CHO (-)).
This is very important, cells will be very sensitive after the transfection and the
presence of geneticin will result in almost complete death of the cells even if they
have a plasmid which does encode for geneticin resistance. It will take the cells
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some time before they produce enough copies of this DNA to protect themselves
from the antibiotic.
5.2 Transfections using CaCl2 Phosphate Kit
6.0 Plating Cells for Imaging
6.1 Plating cells on fibronectin coated MatTek dishes
Fibronectin (FN) is an extracellular matrix protein which contains a sequence
of amino acids which interact with integrins allows the cells to form strong
adhesions. The concentration of the fibronectin needed will depend on your
specific experiment. Higher concentrations of FN result in stronger adhesions
and thus if you are interested in studying migration you will want to use small
amounts (1 – 2 g/mL), and if you want your cells to adhere very quickly you
will use higher concentrations (5 - 10g/mL). For most experiments we
simply use 2g/mL. We purchase the protein dissolve it in water and then
aliquot it into small volumes suitable for 1 or 2 experiments. This is done to
avoid multiple freeze thaw cycles and degredation of FN. Stock solutions
(1mg/mL) are kept in the freezer in 200L eppendorff tubes with the label FN
on the top. The fibronectin is sterile and thus can be used in the laminar flow
cabinet.
1. Spray down bottle for the PBS and the aliquot of FN and transfer into
the laminar flow cabinet.
2. Determine the number of MatTek dishes you wish to coat with FN.
For nice even coating it is recommended that you use 1.5mL of
solution for each dish.
3. Dilute the FN to achieve the final desired concentration necessary for
your experiment.
4. Add 1.5mL of the diluted FN solution to each dish. Take care to
ensure that the solution actually does cover the glass portion of the
dish. Most of the time the FN solution will prefer to stay on the plastic
portions of the dish and you will need to gently swirl the dish to coat
the glass portion.
5. Place the dishes in the incubator for 1 hour.
6. After 1 hour remove the dishes and rinse 2x with PBS.
7. The dishes are now ready for cell plating. Do not let the FN dishes go
completely dry. If you have dishes which are ready but your cells are
not ready to go onto the dish pace a small amount of sterile media in
the dish until you are ready to add the cells.
8. Determining how many cells to place in each dish will depend on
when you are going to image the cells, and the specific conditions of
your experiment. CHO cells grow much faster than HEK cells and thus
need to be plated at lower densities to ensure the dish is suitable for
imaging. Dishes which are too dense are difficult to examine and thus
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it is best to err on the side of caution. The following guideline I use
when I prepare dishes and image them 48 hours later.
i. Add 1.5mL of fresh media to each dish.
ii. Add 0.2mL of cells (about 3 drops) to each dish.
9. Carefully place the cells back in the incubator. Ensure that no media is
spilling over the edges. Since media has ideal growth conditions we do
not want media all over inside the hood, allowing for the potential
growth of contaminants.
6.2 Plating cells on collagen coated dishes
7.0 Imaging Cells
7.1 Fixed Cells
Cells which are fixed with 4% paraformaldehyde should be kept moist. This
can be done by adding about 1 mL of PBS to the dish and keeping it in the
fridge until ready to image.
7.2 Live Cells
Imaging of live cell imaging should be done in media containing HEPES, and
no phenol red. When transporting cells from chemistry lab to the physics lab
make sure you give the cells about 1 hour of time to recover from the journey.
This will ensure that your cells are well adhered and “happy”.
Light can cause production of reactive oxygen species (ROS), which leads to
cellular damage and may interfere with sensitive pathways under
investigation. You can minimize this damage by lowering the laser power of
the Argon 488 to about 0.5 – 1%, and minimizing the exposure of the cells to
the Hg lamp.
8.0 General Lab Maintanance
8.1 Cleaning media bottles for cell culture purposes
Avoid using soap to clean the media bottles which will be later used for media
and other cell culture related solutions. Left over soap residue in the media
would be toxic to the cells. Instead remove the tape from each bottle and rinse
it 3x with the bleach and water solution (50:50) in the squirt bottle. Then rinse
the bottles 9x with DI water. Allow the bottles to dry upside down fully. The
bottles and caps will need to be autoclaved.
When autoclaving bottles place the cap on the bottle but do not tighten
completely. Place a piece of tinfoil over the bottle top and then a small piece
of autoclave tape. Autoclave empty bottles using the preset “empty” mode.
After the bottles have cooled to the touch, using a glove tighten the cap on the
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bottles (leaving the tinfoil over the cap). Place the bottles on the shelf above
the water bath in the cell culture room. When preparing media discard the
tinfoil and spray the bottles with EtOH before transferring them into the
laminar flow cabinet. Once in the cabinet remove the lid and place the lid
upwards on the floor of the cabinet. Using the UV light sterilize the bottle and
inside of the lid for 10 minutes prior to using.
8.2 Autoclaving glass pasture pipettes
Remove the old autoclave tape from the empty metal pipette holders. Fill the
holders with the 9” pipettes and seal one side of the box with new autoclave
tape. Autoclave on the preset “packs” mode. Once cooled transfer to the
drawer below the water bath in the autoclave room.
8.3 XX
9.0 Troubleshooting
10.0 Notes
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