Counting Cells with a Hemocytometer

The hemocytometer used in cell culture is almost always the improved Neubauer pattern. It is a thick glass slide with two polished counting platforms, each carrying an identical ruled grid, separated by moats. Raised shoulders on either side hold the coverslip exactly 0.1 mm above the ruled surface, and that fixed depth is what turns a two-dimensional count into a volume measurement.

The ruled area is 3 mm x 3 mm = 9 mm2, divided into nine 1 mm x 1 mm squares:

• The four corner squares are each subdivided into 16 squares of 0.25 x 0.25 mm. These are the squares used for mammalian cell counting.
• The centre square is subdivided into 25 medium squares of 0.2 x 0.2 mm, each of which is further divided into 16 small squares of 0.05 x 0.05 mm — 400 small squares in total. This finer ruling is intended for small, numerous particles such as erythrocytes and platelets.

The volumes follow directly from area x 0.1 mm depth:

The 1 x 10^-4 mL figure is the origin of the famous x 10^4 factor. A count of cells in one corner square is a count in one ten-thousandth of a millilitre, so multiplying by 10,000 converts it to cells per mL. There is nothing else to the factor, and it changes if you count in a chamber of a different depth — some disposable chambers are 0.02 mm or 0.1 mm deep and carry a different multiplier, which is printed on the packaging.

On the improved Neubauer the boundaries of the 1 mm2 squares are drawn as triple lines. The middle line of the triple is the boundary for counting purposes. This is a common source of quiet error: counting to the outer line inflates the area, and therefore the count, by a few percent.

1. Clean the chamber and coverslip with 70% ethanol and dry with a lint-free tissue. Use the thick, ground coverslip supplied with the hemocytometer, not an ordinary microscopy coverslip — an ordinary coverslip bends and the chamber depth is no longer 0.1 mm.
2. Seat the coverslip. Moisten the shoulders very lightly by breathing on them, then press the coverslip down firmly. When it is seated correctly, Newton's rings — faint interference fringes — appear where glass meets glass. Their presence is the only reliable confirmation that the chamber depth is correct.
3. Produce a genuine single-cell suspension. Clumps make the count meaningless. Trypsinise adequately, pipette gently to disperse, and pass through a cell strainer if the line is prone to aggregation.
4. Mix immediately before sampling. Cells settle within a minute. Pipette up and down or invert the tube several times, and take the aliquot straight away from mid-depth.
5. Dilute with trypan blue. The standard is a 1:1 mix — for example 50 microlitres of cell suspension plus 50 microlitres of 0.4% trypan blue, giving a dilution factor of 2. Mix gently and count within 3-5 minutes: trypan blue is cytotoxic, and if the mixture stands, viable cells begin to take up dye and viability reads artificially low.
6. Load the chamber. Take about 10 microlitres, touch the pipette tip to the notch at the edge of the coverslip, and let capillary action draw the sample in. Fill each chamber in one continuous action.
7. Check the fill. The ruled area should be evenly covered with no bubbles. If liquid has run into the moats, the coverslip has been floated off its shoulders and the depth is wrong — clean, dry and reload. Under-filling leaves dry patches and biases the count the other way.
8. Let the cells settle for about 1 minute before counting, so they lie in a single focal plane.

For mammalian cells in culture, count the four 1 mm2 corner squares. Some laboratories add the centre square and count five. Either convention is valid; what is not valid is changing between them, because the two give different sampling statistics.

The boundary-line rule exists because a cell lying on a ruled line would otherwise be counted twice, once in each adjacent square, or missed entirely. The rule is simple:

Count cells touching two adjacent sides of the square — an "L" shape — and exclude cells touching the other two.

The commonest convention is to include the top and left lines and exclude the bottom and right; the mirror-image convention, including top and right, is equally correct. Choose one, write it into your SOP, and apply it identically in every square and by every operator. A cell is judged by where the majority of its body lies relative to the middle line of the triple ruling.

Two further rules matter in practice:

• Both chambers. Load and count both sides of the slide. The two counts should agree within roughly 10%. If they do not, the suspension was not properly mixed, the chamber was unevenly filled, or cells are clumping — reload rather than averaging a bad pair.
• Density window. Aim for roughly 25 to 100 cells per 1 mm2 corner square, which corresponds to a suspension of about 2.5 x 10^5 to 1 x 10^6 cells/mL after dilution. Below about 25 per square the count is dominated by sampling noise; above about 100 cells overlap and are systematically undercounted. If the suspension is too dense, dilute further and record the extra factor.

The precision limit is statistical, not human. Cell counting follows Poisson statistics, so the coefficient of variation is approximately 1/sqrt(N) where N is the total number of cells counted. Counting 100 cells gives a CV of about 10%; counting 200 gives about 7%; counting 400 gives about 5%. No amount of care with a single square of 30 cells will beat this — the only way to a tighter count is to count more cells.

The formula is:

Cells/mL = (total cells counted / number of squares counted) x dilution factor x 10^4

The dilution factor is the total volume divided by the volume of cell suspension in it. A 1:1 mix with trypan blue is a dilution factor of 2, not 1. This is the single most frequent arithmetic error in cell counting.

Worked example. A T-75 flask of HeLa has been trypsinised and resuspended in 10.0 mL of complete medium. 50 microlitres of that suspension is mixed with 50 microlitres of 0.4% trypan blue (DF = 2) and 10 microlitres loaded. The four corner squares give:

Viable cell concentration

84.0 x 2 x 10^4 = 1.68 x 10^6 viable cells/mL

Total (viable + dead) concentration

(84.0 + 4.5) x 2 x 10^4 = 1.77 x 10^6 cells/mL

Total viable cells harvested

1.68 x 10^6 cells/mL x 10.0 mL = 1.68 x 10^7 viable cells

Viability

336 / (336 + 18) x 100 = 94.9%

Note that the dilution factor cancels in the viability calculation, so it does not appear there. Note too that 336 cells counted gives a CV of about 5.5%, so the concentration is properly reported as 1.7 x 10^6 cells/mL — quoting it as 1.68 x 10^6 implies a precision the method does not have.

Each corner square held between 76 and 91 cells, inside the 25-100 window, so no further dilution was needed.

Trypan blue is an anionic diazo dye that cannot cross an intact plasma membrane. Cells with compromised membranes take it up and appear distinctly blue and often swollen; viable cells exclude it and appear bright and refractile with clear borders. The standard working solution is 0.4% (w/v) in saline or PBS.

What this measures is membrane integrity, and that is narrower than "viability" in ordinary use. A cell that has committed to apoptosis but still has an intact membrane counts as viable by trypan blue. A cell that has been mechanically damaged during trypsinisation counts as dead even though the population it came from is healthy. Trypan blue exclusion is the right tool for a routine harvest check and the wrong tool for measuring the outcome of a cytotoxicity experiment, where an apoptosis-specific or metabolic assay is needed.

Practical points:

• Time matters. Count within 3-5 minutes of mixing. Trypan blue is itself toxic and prolonged exposure kills cells, dragging the viability figure downward.
• Protein interferes. Trypan blue binds serum protein, which stains the background and makes discrimination harder. If the background is heavily blue, wash the cells into PBS before counting.
• Expect above 90% from a healthy log-phase culture harvested cleanly. A result of 70-85% points to over-trypsinisation, an over-confluent flask, harsh pipetting, or a culture problem worth investigating before the cells are used.
• Debris is not dead cells. Small, irregular blue fragments are membrane debris and should not be counted. Only count objects of plausible cell size and shape.

The count is only useful if it is turned back into a volume to pipette. Continuing the worked example above, the suspension is 1.68 x 10^6 viable cells/mL.

Task: seed a 6-well plate at 2 x 10^5 cells per well in 2 mL of medium.

1. Target concentration. 2 x 10^5 cells in 2 mL = 1 x 10^5 cells/mL.
2. Total volume needed. Six wells x 2 mL = 12 mL. Prepare 13 mL to cover pipetting losses.
3. Total cells needed. 13 mL x 1 x 10^5 cells/mL = 1.3 x 10^6 cells.
4. Volume of stock suspension. 1.3 x 10^6 / 1.68 x 10^6 cells/mL = 0.774 mL = 774 microlitres.
5. Make up. Add 774 microlitres of cell suspension to 12.23 mL of pre-warmed complete medium, mix gently, and dispense 2 mL per well.
6. Distribute evenly. Rock the plate front-to-back and then side-to-side before placing it in the incubator. Swirling in a circle drives cells into the centre of each well.

The same arithmetic drives every downstream use of the count: expansion into flasks at a set cells/cm2, cryopreservation at a set cells/mL per vial, and seeding assay plates. Keep the concentration in cells/mL as the working number and convert once at the end.

Clumps. The most damaging failure, because it biases the count without looking like an error. Clumped cells are undercounted, seeding densities come out low, and results are irreproducible. Causes are incomplete trypsinisation, DNA released from lysed cells, and inadequate mixing. Fix by extending dissociation slightly, adding DNase to the neutralising medium, pipetting gently to disperse, or straining. Do not attempt to count a clumped suspension; re-prepare it.

Liquid in the moats. The coverslip has floated and the chamber is no longer 0.1 mm deep, so every number derived from it is wrong. Clean, dry, reseat the coverslip until Newton's rings appear, and reload.

Too few cells per square. Below about 25 per corner square the Poisson error swamps the measurement. Concentrate the suspension, or count more squares and adjust the divisor accordingly.

Too many cells per square. Above about 100 per corner square cells overlap and are undercounted. Dilute and multiply the extra factor into the calculation. Records should carry the full dilution chain, not just the final trypan blue step.

Forgetting the trypan blue dilution factor. Halves every result. If a seeding density comes out at exactly half or double what was expected, check the dilution factor first.

Counting the same chamber twice and calling it a replicate. Reloading and recounting the same chamber tests your eyes, not the suspension. Genuine replication means both chambers, from a freshly mixed sample.

Cells settling in the tube during a long count. The aliquot loaded second is more dilute than the aliquot loaded first. Mix immediately before each load.

Inconsistent boundary rule between operators. Produces a systematic offset between people counting the same suspension, typically of several percent. Write the convention down.

Dirty or scratched chamber. Debris in the ruling is counted as cells by tired eyes and by some automated systems. Clean with ethanol and a lint-free tissue, never with abrasive material — the ruling is etched and can be damaged permanently.

The hemocytometer survives because it is cheap, needs 10 microlitres, requires no calibration, and lets the operator see the sample. That last point is not trivial: clumping, debris, contamination and unexpected morphology are all visible during a manual count and invisible in a number returned by an instrument.

Its weaknesses are throughput and operator dependence. It takes several minutes per sample, and different operators applying different boundary conventions and different judgements about what counts as a cell will disagree by more than the statistical error.

Automated image-based counters run the same trypan blue chemistry in a disposable slide and apply a fixed algorithm, which removes operator bias and takes under a minute per sample. They are the right choice for routine counting at volume. They inherit the hemocytometer's blind spots — they also miscount clumps — and they need to be validated against a manual count for each new cell type, because the size and circularity gates are set for typical mammalian cells.

Impedance (Coulter-principle) counters count many thousands of particles per run and give excellent precision on total particle number, but they discriminate by size, not by membrane integrity, so they give no viability figure and count debris of the right size as cells.

Flow cytometry with a viability dye gives per-cell, multiparameter data and can distinguish apoptotic from necrotic populations. It needs more sample, more setup and more analysis, and absolute concentrations require either counting beads or a volumetric instrument. Use it when the question is about the state of the cells, not simply how many there are.