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Is there a generally accepted maximum number of times you should split and reseed mammalian culture cells before restarting cultures from new stock?
This is a very good question and it highlights a very frequent misconception about cell culture and it is using the passage numbers as an indication of how well a cell is behaving. Although this can be a good barometer but it is misleading. Passaging can stress the cells if performed very frequently, although again that depends on how fast the cells are dividing.
You should not think of passage (and cell culture in general) in terms of the procedure itself but in terms of population doubling (PD) which can be calculated using the formula provided here (http://www.sciencellonline.com/site/techsupport.php). I personally use [Log10(harvesting cell No/seeding cell No)/Log10(2)]. This should allow you to characterise your cellular behaviour between every passage and allows you to draw a cumulative PD graph. Once the cumulative PD versus time becomes non-linear, then you would know that the cells have changed behaviour and it might be best to use a fresh stock.
The main advantage of this method is that you are not relying on guess work and deal with solid numbers and allows you to predict when you should expect to start a fresh batch for passage. Although passage numbers can do the same job if you assume per passage, the cells have done a certain PD but this is more of a guess work and you can not track how well your cells are doing at a given time point. For example one of the main problems with using a passage as a measure is that you can passage the cells at 98% confluence to a fresh flask and although that counts as a passage but it is pretty uninformative in terms of cell behaviour and gives you no solid numbers and you can do this many many times and although you would have done high number of passages but your cells would appear ok. Although such an exercise is pointless and will probably stress the cells but it is just to make the point that passage numbers are not necessarily a very informative and quantifiable measure of how well the cells are doing or behaving hence PD and cumulative PD should be used. Another advantage is that since you are dealing with actual cell numbers, if you changed your cell culture flask size from lets say T-75 to T-25, you don't have to be worried about changes to your frequency of passage or having to make further guess work on how that has effected your cell behaviour.
My main advice is that you should keep you frequency of passage the same at least in the beginning (although keep an eye on the cell density as you don't perhaps want to let cells to be too over-confluent or unconfluent when passaging, depending on your specific methodology and what has been established in your lab), but count your seeding and harvesting density (cell No) and calculate the PD and cumulative PD and draw a cumulative PD vs time graph and monitor the behaviour of cells. This should give you a good indication of you cellular behaviour under the conditions you are working with them since guidelines are just that, guides and they can never substitute experimental characterisation of your cells under your conditions and handling method!
Subculturing Suspension Cells
The following protocols describe general procedures for subculturing mammalian cells in suspension culture. Note that the procedure for passaging insect cells differs from that for mammalian cells on several crucial steps. For more information, refer to Notes on Subculturing Insect Cells.
For passaging your own cell line, we recommend that you closely follow the instructions provided with each product you are using in your experiments. The consequences of deviating from the culture conditions required for a particular cell type can range from the expression of aberrant phenotypes to a complete failure of the cell culture.
Passaging Suspension Cultures
Subculturing suspension cells is somewhat less complicated than passaging adherent cells. Because the cells are already suspended in growth medium, there is no need to treat them enzymatically to detach them from the surface of the culture vessel, and the whole process is faster and less traumatic for the cells. Replacement of growth medium is not carried out in suspension cultures instead, the cells are maintained by feeding them every 2 to 3 days until they reach confluency. This can be done by directly diluting the cells in the culture flask and continue expanding them, or by withdrawing a portion of the cells from the culture flask and diluting the remaining cells down to a seeding density appropriate for the cell line. Usually, the lag period following the passaging is shorter than that observed with adherent cultures.
Cell passage: How to correctly dilute and split cultured cells
There are many different cell culture techniques. Utilizing correct cell passaging methods is important to keep cell line in exponential growth curve, therefore making it a good model as a host for transfection experiments. “Cell passage” is a term used by other scientists to demonstrate the following process:
- Wash cells with PBS
- Detach cells from flask by trypsinization
- Resuspend in complete media (contains FBS) to neutralize trypsin
- Transfer appropriate dilution to new flask containing ample media
The correct passage dilution is cell line specific and depends on factors such as doubling time and intended use of the cells. The important aspect to remember is that the split ratio is determined from the total volume of trypsin and media from steps 2 and 3 above. As an example for a T75 flask, if 1 mL of trypsin is used to detach the cells and 9 mLs of complete media is used to neutralize the trypsin, then the total suspension volume is 10 mL. Depending on the cell line, here are examples of split ratios a researcher might follow:
- View cultures using an inverted microscope to assess the degree of cell density and confirm the absence of bacterial and fungal contaminants. Harvest cells in the log phase of growth. For adherent cell lines harvest cells as close to 80 - 90% confluency as possible.
- Bring adherent and semi adherent cells into suspension using trypsin/EDTA as described previously and re-suspend in a volume of fresh medium at least equivalent to the volume of trypsin. Suspension cell lines can be used directly.
- Remove a small aliquot of cells (100-200μL) and perform a cell count. Ideally, the cell viability should be in excess of 90% in order to achieve a good recovery after freezing.
- Centrifuge the remaining culture at 150 x g for 5 minutes.
- Re-suspend cells at a concentration of 2-4x10 6 cells per mL in freeze medium.
- Pipette 1mL aliquots of cells into cryoprotective ampoules that have been labelled with the cell line name, passage number, lot number, cell concentration and date.
- Place ampoules inside a passive freezer e.g. Nalgene Mr. Frosty Freezing Container. Fill freezer with isopropyl alcohol and place at -80 °C overnight.
- Frozen ampoules should be transferred to the vapor phase of a liquid nitrogen storage vessel and the locations recorded.
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Nalgene is a registered trademark of Thermo Fisher Scientific or its subsidiaries
Liquid Nitrogen Safety Considerations
General Safety Issues
It is important that staff are trained in the use of liquid nitrogen and associated equipment including the storage vessels which need to be vented safely and containers which may need to be filled. As with all laboratory procedures personal protective equipment should be worn always whilst handling nitrogen, including a full-face visor and thermally insulated gloves in addition to a laboratory coat and preferably a splash proof plastic apron. Proper training and the use of protective equipment will minimize the risk of frostbite, burns and other adverse incidents.
Risk of Asphyxiation
The single most important safety consideration is the potential risk of asphyxiation when escaped nitrogen vaporizes and displaces atmospheric oxygen. This is critical since oxygen depletion can very rapidly cause loss of consciousness, without warning. Consequently, liquid nitrogen refrigerators should be placed in well-ventilated areas in order to minimize this risk and be subject to planned preventative maintenance. Large volume stores should have low oxygen alarm systems.
Primary Cell Lines
Primary cell lines are derived directly from excised tissue and cultures either as an explant culture or following dissociation into a single cell suspension by enzyme digestion. Such cultures are initially heterogeneous but later become dominated by fibroblasts. The preparation of primary cultures is labor intensive and they can be maintained in vitro only for a limited period of time. During their relatively limited lifespan primary cells usually retain many of the differentiated characteristics of the cell in vivo. Important Note: Primary cultures by definition have not been passaged, as soon as they are passaged they become a cell line and are no longer primary.
Assessing Cell Growth
Accurate cell counting is of paramount importance when assessing cell growth in cultures, as faulty counts can lead to mistaken conclusions about cell health. Cell counts can be determined using a hemocytometer. However, automated cell counters that utilize the Coulter principle where cells flow, one by one, through an aperture within an electrical sensor, yield the most precise cell counts.
Figure 2. Scepter™ 2.0 Handheld Automated Cell Counter
Freezing and Thawing Cells
How to Freeze Cells
- cultured cells
- cell growth media
- solutions for detaching cells from the plate if using adherent cells (e.g. balanced salt solution, trypsin/EDTA)
- tissue culture grade dimethylsulfoxide (DMSO)
- fetal bovine serum (FBS)
- cell freezing chamber
- –80°C freezer
- liquid N2 tank for long-term storage.
1. Passage Cells
Healthy, actively growing cells should be used for cryopreservation. I always try to freeze large numbers of cells that have not been passaged in culture too many times.
I passage the cells 1–2 days prior to freezing to ensure that they are in a log rhythmic phase of growth at the time of freezing.
Some labs recommend changing the growth medium 24 hours prior to freezing if the cells have not been passaged at that time.
2. Prepare Cells
Remove cells from the dishes following the usual method for passaging adherent or suspension cells. Pool all cells and centrifuge.
3. Resuspend Cells in Freezing Medium and Aliquot to Cryovials
Freezing cells can be lethal. To avoid the damage that can be caused by, for example, ice crystal formation, osmotic stress, or membrane damage, a cryoprotectant is used to lower the cells’ freezing point.
DMSO, as a 10% stock solution, is the most commonly used cryoprotectant.
Caution: Wear gloves when working with DMSO as it easily penetrates the skin.
The most common freezing medium is 90% FBS/10% DMSO. For less finicky cells and for tissue culture on a budget, 10% DMSO in cell growth medium can also be used.
After centrifugation, resuspend the cell pellet in 1 mL of freezing medium per cryovial.
Make sure you have cryovials designed for liquid N2 storage. I typically plan on 1 100-mm dish of cells per cryovial. Vials should be labeled using a lab marker that will withstand alcohol and liquid N2.
Be careful to include passage/lot numbers when labeling cryovials.
4. Freeze Cells
To allow water to move out of the cells before freezing, freeze cells slowly. This is accomplished using a cell freezing chamber. Pricey freezing chambers pulse in liquid N2 periodically to control the freezing rate.
Less expensive options include chambers that use room-temperature isopropanol. Vials are placed in the chamber, isopropanol is added, and the chambers are placed at –80°C for at least 4 hours.
Those of us on a tight budget (like me!) use homemade chambers. I keep the Styrofoam™ racks that come with 15-mL conicals, place a cryovial in each hole, cover with a second rack, tape the racks together, and place at –80°C.
5. Remember To Move Your Cells to the Liquid N2 Tank
Probably one of the hardest things in this protocol is remembering to move your frozen cryovials to the liquid N2 tank! I usually allow my cells to freeze overnight and then quickly place the cryovials into the liquid N2 tank.
How to Thaw Cells
1. Remove Cells From the Tank and Thaw
For the greatest cell viability, it is important to freeze the cells slowly. The opposite is true for thawing—thaw quickly! Remove cryovials from the liquid N2 tank and immediately place them in a 37°C water bath.
Keep the cryovials in the water bath until just the tiniest ice crystal is left in the cryovial.
2. Transfer, Spin (?), and Plate
Immediately transfer the cells to a large volume of pre-warmed cell growth medium (
10 mL/1-mL aliquot of cells). For the next step, you have a decision to make.
There are two schools of thought on whether the cryoprotectant should be immediately removed from the cells:
- In some labs, the cells are centrifuged and the cell pellet is resuspended in fresh cell growth medium prior to placing cells in a culture dish. This is the way I first learned to thaw cells.
- However, some data suggest that cells are extremely fragile after thawing and that centrifugation may increase cell death. Therefore, it is recommended that the cells are plated and that the growth medium is changed at a later time.
For adherent cells, you can change the medium as soon as the cells have attached to the dish.
I switched to this method and now I generally take my cells out just before leaving the lab for the night and change the medium first thing the next morning.
3. Check Your Cells
About 24 hours after thawing, I look my cells over carefully under the microscope to make sure they are healthy and behaving normally.
Freezing and thawing cells effectively will save you the embarrassment of asking a colleague for yet another aliquot of their cells or the expense of purchasing new cells.
And if all else fails, remember to freeze slowly and thaw quickly! For more on cryopreservation, check out our article on preserving microorganisms. Let us know about your top tips for freezing and thawing cells!
Originally published April 3, 2013. Reviewed and updated May 2021.
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I’m working on CHO suspension cells, in our institution there is no place in -196 to store freezed cells. So
1.Can we store freezed CHO cells at -80 ?
2. What is the appropriate number of cells to freeze?
How long can we store CHO suspension cells at -80?
Depends on whether you have a cryoprotectant. But generally they can last for a very long time.
Any info on how many cells should be plated on a flask of a particular size?
It really depends on the type of cells you are using. They can be very different sizes.
Sometimes you can get an idea by looking at the ATCC website and look at the culture conditions.
When freeze thawing, I put as many cells in to the flask as were in the flask when I harvested.
If you think you cells will not survive freezing well, you can put them in to a smaller flask.
I was taught by some guy who made transgenic mice to slowly add DMSO seperately to your freezing media. This allows the cells to get used to the DMSO, since it is toxic for cells, especially at higher temperatures. I now use 2 parts of medium to freeze my cells in. I use normal medium (DMEM, 10% FCS, Gentamycin) to resuspend my cells in and 2x freeze medium (DMEM, 30% FCS, 20% DMSO, Gentamycin). After centrifugating my cells, I resuspend my cells in half the end volume in ice-cold normal medium and then I gradually add the ice-cold 2x freeze medium, swirling the cells between each addition. I keep the cells on ice to keep them cool until they go into the -80C (in whatever container that is available).
For the thawing part, I thaw the cells in a 37C water bath and put them on ice once the ice in the tube is almost gone. Then is put the cells into an ice-cold 14ml Falcon tube on ice and gradually add ice-cold DMEM, 10% FCS, Gentamycin, again swirling between every addition. (I usually use ¬6ml to add, but you can add more if you want). After spinning my cells down, I resuspend my cells in room temperature DMEM, 10% FCS, Gentamycin and put them into the incubator.
It may seem overly complicated (for some cell lines it certainly is), but if you have to wait every timeuntil your cells are healthy to do experiments with, it may just be worth the effort. Good luck!
REAGENTS AND SOLUTIONS
HEPES buffer, 50 mM (pH 8.0)
- 238.3 g HEPES (free acid e.g., Sigma-Aldrich, cat. no. H3375)
- 800 ml Milli-Q water
- Adjust pH to 8.0 with 10 N NaOH
- Bring volume to 1000 ml with Milli-Q water
Store at room temperature for up to 1 year
The solution prepared is a 1 M HEPES stock solution. Before use, dilute 20× in Milli-Q water to prepare a 50 mM solution.
PEG 8000, 50%
- 250 g PEG 8000 (e.g., Sigma-Aldrich, cat. no. 89510-250G-F)
- 250 ml Milli-Q water
- Heat to 80°C to dissolve PEG 8000
- Allow solution to cool
- Store at room temperature for up to 2 months
PEI, 1 µg/µl
- 1. Dissolve PEI (e.g., Polysciences, cat. no. 23966-1) by swirling using sterile water heated to 80°C and using ∼90% volume required to reach a final concentration of 1 g/L (1 µg/µl).
- 2. Cool to room temperature.
- 3. Adjust pH to 7.0 with HCl.
- 4. Add sterile water to a final concentration of 1 µg/µl.
- 5. Filter using a 0.22-µm membrane filter, and divide into aliquots. Store at −20°C for up to 1 year.
- 6. Thaw in a 37°C water bath until precipitates have fully dissolved.
7. Store thawed aliquot at 4°C for up to 1 month.
Redissolve precipitates by incubating at 37°C before use.
It’s tempting to get right into experimentation the moment your cells arrive. In reality, it can take some time to get a good feel for how your cells are likely to behave. Not every aliquot is made equal, even if the source is the same. It’s worth taking the time to implement every strategy necessary to keep your work as consistent and accurate as possible.
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