You don’t need to be a biologist to accurately measure the count and viability of your yeast culture. Without sampling and counting your library yeast and starters, you have to rely on estimates and calculators from software which give really rough generalized estimates, but can be far off from reality depending upon the specific yeast strain, or other factors like how it’s been stored. It’s a lot easier to count yeast than you think, you just need a few pieces of equipment.

Here is a link to the yeast counting spreadsheet.

Equipment

Process

Prepare Sample

For reference, the vials will be referred to as vial #1 through #3 and will be used as follows:

  • Vial #1: Sample for measuring, this is used to count the yeast
  • Vial #2: Used for mixing stain
  • Vial #3: Stain for yeast, first dilution

1. Measure 9 mL distilled water into vial #1.

2. Add 10 to 20 drops of stain to vial #2, then add about 10 mL distilled water. Mix completely, then use the pipette to transfer 9 mL from vial #2 into vial #3.

3. Add 1 mL of yeast sample into vial #3. Mix completely and let stand for 30 seconds so that non-viable cells can absorb the stain.

4. Add 1 mL of vial #3 mixture to vial #1, make sure to mix vial #3 right before transferring so that it’s completely stirred up.

5. Set slip cover over hemocytometer chambers.

6. Get about 1 mL from vial #1 of the prepared yeast sample and carefully touch the edge of the hemocytometer slide, and allow the liquid to wick under the slip cover via capillary action. Gently use the pipette to drop a little bit until the hemocytometer troughs are mostly full.

Count Yeast

Now that the sample has been prepped, we can move on to the microscope to actually count some yeast.

1. Put the slide on the microscope and center one of the grids (the grids are in the top and bottom space of the “H” part of the slide). This is best done at the lowest power on the microscope.

2. Set the power to 40x and refocus. You should see a grid of lines and squares. Find the 25 squares (5×5) of smaller squares (we’ll call these grid-squares) that each contain 16 of the smallest squares (4×4) in them (we’ll call these count-squares). You will be counting 5 of the grid-squares, the four corners and the center one.

3. Open up the yeast counting spreadsheet and make a copy of your own. If you don’t want to use the spreadsheet, the formula for all calculations are included below for use elsewhere. Enter the volume of the full yeast culture that you pulled the 1 mL count sample from. Begin at the upper left grid-square (square 1) and count all of the viable (clear and unstained) vs. nonviable (stained) cells. Record these numbers in the spreadsheet. You must be consistent with counting rules. Count cells that overlap only the right or bottom border of a square. Move from left to right just like reading a book.

4. Once all cells in a square have been counted and recorded, move to the upper right grid-square (square 2). Repeat the count and record the numbers.

This is a thick sample. While it does’t hurt anything, it makes counting more tedious. The sample could be diluted more, and the calculations would need to be adjusted.
This sample is about the right dilution to make counting easy, but has some clumping which can make it harder to count, as well as skew accuracy and precision. If your sample is clumping, try placing it in a refrigerator for a few hours. and see if the cells separate.

5. Repeat the counting step for the center grid-square (square 3) and the left and right bottom grid-squares (squares 4 and 5). Once all the data has been recorded, the spreadsheet will calculate the viability and total yeast in the original yeast culture.

Conclusion

Counting yeast is not difficult and can give you a better way to ensure your pitch rates are accurate and that your yeast are healthy without a large investment in equipment. It can also be instrumental when using yeast that were kicked up from dregs of another beer or that came from an unknown source. This allows you to eliminate one more variable from your brewing process.

Data

  • Chamber height: 0.1 mm (slip cover creates this chamber height)
  • Grid-square dimensions:  0.20 mm x 0.20 mm
  • Grid-square area: 0.04 mm^2
  • Grid-square volume: 4 nL
  • Grid dimensions: 1 mm x 1 mm
  • Grid area: 1 mm^2
  • Grid volume: 0.1 nL
  • Dilution: 100 (diluted 1 mL yeast in a 10 mL solution, then diluted 1 mL of that in another 10 mL solution).
  • Record the total culture volume (of the starter or vial) that you took a sample from in mL. This will allow you to extrapolate the total amount of yeast in the culture based on your counting.
  • Record the number of viable and nonviable cells from each of the count-squares. Add them together for an overall total of viable and nonviable for each grid-square.

Formulae

  • G_V = grid-square viable = sum of all count-squares viable cells
  • G_N = grid-square nonviable = sum of all count-squares nonviable cells
  • T_V = total viable = sum of five grid-square viable cells (all GV)
  • T_N = total nonviable = sum of five grid-square nonviable cells (all GN)
  • A_V = average viable cells per grid-square = \frac{T_V}{5}
  • A_N = average nonviable cells per grid-square = \frac{T_V}{5}
  • V = calculated viable cells for grid = {A_V}\cdot{25}
  • N = calculated nonviable cells for grid = {A_N}\cdot{25}
  • viability % = \frac{V}{(V + N)}
  • CM = culture cell count per mL (in millions of cells) = \frac{1}{grid volume}\cdot{dilution}\cdot\frac{V + N}{1,000,000}
  • CT = culture total cell count (in billions of cells) = CM\cdot\frac{total culture volume}{1,000}p
  • b = number of budding events (how many yeast divisions) = 3.32 log_{10}(\frac{CT}{CT_0}) where CT is the final yeast cell count and CT_0 is the initial yeast cell count

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