For simplicity, in this document all information given will apply to NIH 3T3 cells. Many procedures will be the same or similar for other cell lines but it is best to check with someone who has cultured them before in the lab or to check the relevant literature for additional information.
NIH 3T3 mouse fibroblasts are a cell line used in the lab for many basic assays because they are easily transfectable and also easily maintained. However, we have many other lines from several species (mouse, rat, dog, human), cell types (fibroblast, epithelial) as well as cells that are derived from tumors or that have been transformed by the introduction of various oncogenes. Our available inventory of cell lines can be found on the computer in the FileMaker Pro file “Where the cells are”. This inventory is a record of the contents of the liquid nitrogen tanks we have in the lab for long term storage of cell lines. A stock cell line does not have any additional genes transfected into it. Different cell lines can be derived from stock lines by stably transfecting plasmids into them to form NIH 3T3 that express H-Ras 61L (NIH 3T3 H-Ras 61L), for example.
Cell lines will have a preferred set of conditions in which they grow best. This will generally entail a specific growth media, serum to provide additional growth factors, antibiotics to prevent spurious bacterial growth, and a percentage of CO2 at which the cells are happiest. Some cell lines also require supplements such as non-essential amino acids (NEAA) or glutamine, for example. All mammalian cells in the lab are grown in incubators set to 37° C. Some, but not all, cell lines have their culture condition information included with their entry in the liquid nitrogen inventory. If this is not included, check with the source of the cells (ATCC, UNC Tissue Culture Facility, Lab of origin), someone who has grown them before, or the literature (note that some conditions for the same cells will vary with the lab of origin or the type of experiment they were used in).
Culture Conditions for NIH 3T3 cells:
DMEM-Dulbecco’s Modified Eagle Medium, High Glucose
(sometimes labeled DMEM-H, we use Gibco Brand)
10% GCS-Gibco Calf Serum
to make 10% serum, add 50 ml GCS to a bottle of DMEM (500 ml)
to make 1% P/S, add 5 ml 100X P/S to a bottle of DMEM
DMEM + 10% GCS, 1% P/S is also referred to as ‘complete’ media since it has all of the components in it to grow NIH 3T3 cells. Maintain the cells at 37° C in 10% CO2.
For experiments that call for serum free, antibiotic free media, use DMEM with no additions (no GCS, no P/S). Use the appropriate media for other types of cells.
For experiments that call for starving NIH 3T3 cells, generally we use media with 0.5 % serum (so, 2.5 ml of GCS per 500 ml DMEM) and 1% P/S. Growth overnight in low serum should be sufficient to starve NIH 3T3 cells. The time required and the percentage of serum will vary with the cell line. The reduced amount of serum will reduce the amount of growth factors that are available to the cells. This is useful if you want to see a stimulatory effect of a particular growth factor or drug on your protein of interest, if you want to reduce background noise from growth factors in the media for a particular readout, or if you want to see if your protein of interest reduces the need for growth factor supplementation (by producing its own growth factors).
Most types of growth medium contain phenol red as a pH indicator (this is why they are reddish-pink in color). Bright pink media usually indicates low CO2, if your cells are not supposed to be in low CO2 but the media is pink when you open the incubator door, this may indicate that there is a problem with the incubator (alert someone and check the tanks to see if any are empty). Your media will also be pink after cells have been sitting out of the incubator for a while during work in the hood, or as your media bottle becomes emptier (it should return to a more reddish shade in the incubator). Yellow-orange media usually indicates that the cells are overgrown on the dish (usually with transformed cells).
Passaging and Plating Cells
Most cells (unless transformed) will stop growing when they fill up a plate, thus they need to be passaged, or split, when they get confluent to get more cell growth. In our lab, most cells that are frozen are done so at a “low” passage number (right after selection stops for stable lines or right after they were received from another lab or purchased). This is not technically true as some lines have been in culture for years and even decades and will have a correspondingly high passage number. Some labs keep track of this number since cells drift genetically over time in culture (this is why NIH 3T3 cells from one lab will not be exactly the same as NIH 3T3 cells from another lab). For simplicity, we generally don’t keep track of historical passage number and instead keep track of passage number from the time they were thawed, thus the first split after thaw will be passage one (P1), etc. Generally NIH 3T3 cells are kept in culture for 10-12 passages before a new tube is thawed. Our lab manager will usually have a plate of NIH 3T3 stock cells in culture for the lab to use, you can request plates for transfection in advance by signing up on the sheet in the tissue culture room. However, it is sometimes more convenient to passage and plate them yourself (and of course you will have to do this yourself if you are using cells other than NIH 3T3 stock cells).
Split cells when they are 80-90% confluent, if possible. Confluency is a relative measure of how many cells are on the plate, for example, 50% confluent means that in any given field you view under the microscope, about half of the space will be taken up by cells. Cells that are confluent will be very crowded with little or no space between cells. At this point the cells can get stressed or stop growing (or start dying or grow all over each other, depending on the cell line). Try to avoid this if passaging cells.
General protocol for passaging cells:
Generally stock cells that are being grown for plating or passaging are grown in 100 mm dishes (p100). If you are not using many cells, it may make sense to grow them in smaller dishes.
1. Aspirate the media from the cells, wash once with PBS (generally with the same volume as the media that was on the cells). This step is necessary to remove traces of serum from the cells, which can inhibit trypsin.
2. Aspirate the PBS, add Trypsin-EDTA (T-EDTA) to the plate. Generally, use 1-2 ml, although smaller volumes can be used in smaller dishes. Shake the plate gently to evenly distribute the T-EDTA. This step severs the adhesions that the cells have made to the plate, allowing them to be released into solution.
3. Allow cells to sit in the incubator for 1-5 minutes (note: some cells are more resistant to the action of T-EDTA than others and this may take longer). Shake or tap the plate gently with the side of your hand to release the cells from the plate. You should be able to see the floating cells in the liquid on the plate.
4. To passage NIH 3T3 cells, split them either 1:10 or 1:20, meaning if 1 ml of T-EDTA was used, transfer either 100 µl (1:10), or 50 µl (1:20) to a new plate with fresh complete media on it (10 ml for a p100). Pipet up and down to disrupt adhesions between cells and get a good single cell suspension; this is more of a problem for highly confluent cells or cells that grown in clusters such as some epithelial cells. Rock the plate gently to distribute the cells evenly in the media before returning to the incubator.
In general for NIH 3T3 cells that were 70-80% confluent, a 1:10 split will be ready to passage again the second day after they were plated (split/plate Monday, grow Tuesday, split/plate Wednesday). A 1:20 split will be ready the third day after they were plated. NIH 3T3 cells will tolerate being split further (1:50) but keep in mind that some cells are unhappy if they are too sparse (even 1:10 is too sparse for some) and some cells grow slowly so even a 1:10 split will take a long time to grow back. The doubling time for NIH 3T3 cells is 22-24 hours.
General protocol for plating cells:
Plating cells, as opposed to passaging them, refers to putting a specific number of cells on a plate to have them ready to transfect a plasmid into them for expression of your gene of interest or to get an even number of cells ready across several lines for a particular assay, etc. When plating cells, generally we put cells on a plate the day before or two days before an experiment to allow them to sit down and adhere to the plate. The density is determined by the experiment. For most transfections, a density is used that will give 70-80% confluency the day of transfection (see table below). However, if transfecting for immunofluorescence or live cell imaging for example, a lower density will often be used.
Plating NIH 3T3 Cells
|Plate size||# of cells 1 day before||# of cells 2 days before||Media volume|
|p35 (6-well plate)||0.8 x 105||0.4 x 105||2 ml|
|p60||2 x 105||1 x 105||3 ml|
|p100||4.5-5 x 105||2.5 x 105||10 ml|
1. Determine the number of cells you need and the total volume of media you need. For 10 p60 dishes 1 day before transfection you will need 20 x105 cells (or 2 x106 cells) in 30 ml of complete media. A 70-80% confluent p100 of NIH 3T3 cells should give you around 3-4 x106 total cells.
2. Trypsinize cells in 1ml of T-EDTA as for passaging cells. Resuspend trypsinzied cells in 9 ml of complete media to give about 10 ml total volume. Pipet up and down to get a good single cell suspension.
3. Count cells by moving 10 µl of cell suspension to a hemocytometer, pipet the cells in the groove on one of the sides of the hemocytometer so they flow under the rectangular cover glass. This tool helps estimate how many cells you have per ml of suspension. Under the 4X objective, you should see a square with 9 smaller square boxes inside of it, each with more boxes or lines through them. Under the 10X objective, focus on the 4 corner boxes of the nine (each containing 16 smaller boxes). Count the cells in each of these four larger boxes. Divide the total by 4 to get an average. Multiply this number by 1 x 104 to get an estimate of how many cells you have per ml. For example if your average number of cells was 20 then you would have 20 x 104 cells/ml or 2 x 105 cells/ml.
4. For 10 p60s use 10 ml of the cell suspension to get 2 x106 cells, dilute this with 20 ml of complete media to give 30 ml total. Pipet up and down to get a good suspension. Aliquot 3 ml of cell/media mix to each of 10 p60 dishes. Gently rock the dishes to cover the bottom surface evenly. Return cells to incubator and transfect the next day.
Make sure you split some cells out before you dilute the trypsinized cells for counting so you have a plate to continue passaging cells with if needed.
Quick Protocols: Basic Culture of NIH 3T3 cells
Complete Medium-(DMEM-H + 10% GCS, 1% P/S)
1 bottle of DMEM-H (500 ml)
50 ml GCS
5 ml 100X P/S
Starving or Low Serum Medium-(DMEM-H + 0.5% GCS, 1% P/S)
1 bottle of DMEM-H (500 ml)
2.5 ml GCS
5 ml 100X P/S
Passaging Cells-(for a p100 plate)
1. Aspirate media from cells, wash once with PBS
2. Aspirate PBS, add 1 ml T-EDTA to plate, rock gently to cover plate
3. Incubate at RT 1-5 min
4. Tap plate gently to loosen cells, pipet up and down to suspend cells
5. Pipet 50-100 µl (1:20 to 1:10 split) of cell suspension onto a new p100 plate containing 10 ml complete media
6. Rock plate gently to distribute cells evenly, return plate to incubator
1. Determine how many cells and how much media you need for your experiment
2. Trypsinize plate(s) as for passaging cells (steps 1-4)
3. Resuspend trypsinized cells in 9 ml complete media, remove 10 µl to hemocytometer, and count cells to determine the number of cells/ml
4. Dilute the appropriate number of cells in the appropriate amount of media for your experiment, aliquot it evenly to your plates, return plates to incubator