Passaging and counting

Why it matters

Cells in culture keep dividing until they run out of space or nutrients. Left too long, adherent cells become over-confluent and change behaviour, and suspension cultures exhaust their medium. Passaging (also called splitting or subculturing) moves a fraction of the cells into fresh medium and vessels. To do this reproducibly, and to seed experiments with the same number of cells every time, you count cells. Getting the count wrong by a factor of two or ten quietly ruins experiments and production runs.

When to passage

• Adherent cells are usually passaged at about 70 to 90 percent confluence (the fraction of the surface covered), judged under the microscope, before they fill the flask.
• Suspension cells are passaged when they reach a set density, for example a maximum cells/mL in the cell line's datasheet.
• Every passage increases the passage number. Cell lines can drift genetically over many passages, so labs work within a defined range and go back to frozen stocks from a cell bank.

How adherent cells are passaged (overview)

1. Remove the old medium.
2. Rinse with a calcium- and magnesium-free salt solution, which loosens cell attachments.
3. Add a dissociation enzyme (such as trypsin) briefly at 37 °C until cells round up and detach.
4. Add medium containing serum (or an inhibitor) to stop the enzyme.
5. Mix gently to break clumps, then take a sample to count.
6. Seed the right number of cells into new vessels with fresh medium, label them (cell line, passage number, date, initials) and return them to the incubator.

You do this only in a biosafety cabinet in a teaching or employer lab, after training and under supervision.

Counting with a hemocytometer

A hemocytometer is a thick glass slide with an etched grid. Each large square is 1 mm x 1 mm, and with the coverslip the chamber is 0.1 mm deep, so each large square holds 0.1 uL, or 1/10,000 of a mL.

To tell live from dead cells, the sample is mixed with trypan blue: live cells with intact membranes exclude the dye and look bright; dead cells take it up and look blue.

cells/mL = (cells counted / number of large squares) x dilution factor x 10,000
viability (%) = live cells / (live + dead cells) x 100

Worked example. You mix 10 uL of cells with 10 uL of trypan blue (dilution factor 2) and count four large squares: 52, 48, 55 and 45 live cells (mean 50), and 10 dead cells in total.

live cells/mL = 50 x 2 x 10,000 = 1,000,000 cells/mL (1.0 x 10^6)
viability     = 200 / (200 + 10) x 100 = 95.2 %

To seed a flask with 5 x 10^5 cells: volume = 500,000 / 1,000,000 = 0.5 mL of suspension, topped up with fresh medium to the flask's working volume.

Counting rules that keep counts consistent: count cells touching the top and left borders of a square but not the bottom and right; aim for roughly 20 to 100 cells per square (dilute further if there are more); and count at least four squares. Many labs now use automated counters, which apply the same arithmetic.

Split ratios

For routine maintenance, labs often use a split ratio instead of an exact count. A 1:5 split puts one fifth of the cells into a new vessel of the same size. Experiments that compare conditions almost always need an actual count, so every well starts with the same number of cells.

Common mistakes

• Forgetting the trypan blue dilution factor (a 2x error).
• Counting too few cells, so random error dominates.
• Letting cells settle in the tube before taking the counting sample.
• Leaving trypsin on too long, which damages cells and lowers viability.
• Not recording the passage number, so a culture drifts outside its validated range.

Quick recap

• Passage adherent cells at about 70 to 90 percent confluence; track passage number.
• Hemocytometer: cells/mL = mean count per large square x dilution x 10,000.
• Trypan blue: live cells exclude it; viability = live / total x 100.
• Seeding volume = cells wanted / cells per mL.

Try it

Open the Do tab. A Python counter is written except for the cells-per-mL formula.