Same lesson, with the figures in motion.
Pro Cell Counting
Why count cells?
Allow half an hour, and expect the handling to feel awkward for the first few attempts. It makes a good first lab to practise on: the sample is thrown away afterwards, so there is nothing here you can ruin.
1. Dilute in Two Steps
Yeast slurry is far too dense to count — the cells sit in clumps. Dilute tenfold, twice over: 10 g sample + 90 g water, mix, then take 10 g of that + 90 g water. The result is a 1:100 dilution.
Why not 1 g + 99 g in a single step? Because the scale weighs to 0.1 g. One gram is then measured with a ten percent error — exactly as large as the whole counting error you're about to work hard to keep down. Ten grams is measured to one percent. Same dilution, but the error source disappears.
Yeast sinks in seconds. Swirl the tube right before you draw the next sample, every time — otherwise you move water in one step and sediment in the other.
1:100 is a starting point, not a law. Thick, freshly harvested slurry may need 1:200. You'll only know how much is needed once you're at the microscope: cells packed into one solid crowd means you diluted too little; barely finding any means you diluted too much.
The sample gets thrown away afterward — nothing goes back into the beer. Only what you dip into the mother jar needs to be clean. Don't worry about the tube, pipettes, or chamber.
2. Stain & Count in Time
Add a drop of methylene blue to the diluted sample and wait a couple of minutes. Living cells reduce the dye to a colorless form and stay clear. Dead cells lack that ability and stain blue.
The clock starts ticking the moment the drop goes in. You want to be at the microscope within five minutes — so load the chamber right away, since some of that time goes toward letting the cells settle there.
Leave the sample much longer and even living cells eventually stain, making the yeast look worse than it is.
Trust the number with a grain of salt. Methylene blue overestimates viability — the share of living cells — most when it's actually low. On healthy yeast — around 90% and up — it works fine. Get a noticeably lower reading and you should read it as something is wrong with this yeast, not as an exact figure.
3. Load the Chamber
The haemocytometer is a thick glass slide with an etched grid. Place the coverslip over it and press it down until you see Newton's rings — faint rainbow-colored bands where the two glass surfaces meet.
The rings are the whole measurement. They prove the coverslip sits exactly 0.1 mm above the grid, and that depth is what turns the chamber into a measuring instrument. Get the glass wrong and every number afterward is wrong, no matter how carefully you count.
Touch the pipette to the edge of the coverslip — capillary action pulls the liquid in on its own. Don't overfill. If it runs into the moats around the grid, the depth is no longer accurate: rinse, dry, and start over. Then let it sit a couple of minutes so the cells settle into one plane.
The chamber should be clean, but not sterile. Dried-on yeast from last time becomes ghost cells in your count, and debris under the coverslip lifts it off its depth. Hygiene here is about measurement error, not contamination.
4. The Five Squares
Find the central grid at 400× magnification. It's one square millimeter divided into 25 medium squares. You don't count them all — the four corners and the centre are enough.
Five squares is chosen for speed, not because five is inherently correct. What decides accuracy is how many cells you add up: around 100 cells gives ±10%, 400 cells gives ±5%. In practice you're aiming for 10–40 cells per square — five squares with twenty each is the hundred you want. If you only get about forty total, count more squares instead of settling, and dilute less next time.
The grid described here is for a Neubauer improved, the common type. If you have a different chamber the grid looks different, and the formula in the last step won't apply either.
5. The Edge Cases
Some cells sit right on a line. Count them in both squares they touch and you get too many; skip them in both and you get too few.
Count cells touching the top and left line.
Skip cells touching the bottom and right.
Two of the four sides count, so every cell lands in exactly one square. Which two doesn't matter — only that it's the same two every time.
And yeast buds. A bud only counts as its own cell once it's grown to at least three quarters of the mother cell — smaller than that and it counts with the mother. Here too, consistency matters more than the exact threshold: keep the same rule every time, or your own measurements won't be comparable to each other.
6. Do the Math
Tally the clear ones and the blue ones separately — then add them together. The sum gives you the density; the two numbers apart give you viability.
× 25
× dilution factor (100)
× 10,000
= cells per mL
All cells, not just the living ones. Viability is applied at the next stage — take it off here as well and it gets deducted twice, and you over-pitch.
The number 10,000 isn't magic. The grid is 1 mm² and the coverslip sits 0.1 mm above it, so the box you're looking down into holds 0.0001 mL. Getting from there to a full milliliter means multiplying by 10,000. The chamber measures its own volume — that's why it doesn't matter how precisely you dropped in the sample.
living / (living + blue) × 100%
Now you have a measured number to plug into the pitch calculator, instead of a guess at how many billion cells are sitting in the jar. Open the Measured mode →
Want to practise the rules first – in the game Counting Chamber?
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