One of the key factors in a healthy and timely fermentation is knowing how much yeast you are sending into your fermentation. Pitching too little runs the risk of stalled fermentations or increased ester production. Too much yeast and you increase your chances of having older yeast due to decreased growth rates. By counting your yeast before you pitch it you know how much you are sending into a tank and can also provide early warning signs of bad yeast.
Before you can start counting yeast you’ll need a few pieces of equipment.
Figure 1: A comparison of high quality and low quality haemocytometers
- Microscope with at least a 40X objective lens and movable stage. A stationary stage will work, but moving a counter by hand is a little tricky.
- A quality Haemocytometer - While you can get haemocytometers for cheap from Amazon they are of dubious quality and have very poor contrast.
- A Clicker - Depending on your density you can be counting upwards of 400 cells within your counting regions. It’s a simple solution that has the physical & audio feedback. Alternatively there are phone apps.
- Methylene Blue or Trypan Blue to dye the cells. Both will work, however Trypan Blue is a stronger stain that makes it easier to differentiate between live and dead cells.
- Pipette Pump & Tips - We use micropipette pumps which are nicer to work with, since working at smaller scales can make it easier to perform dilutions and reduce costs. However, using 10mL and 1mL disposable pipettes will also work.When setting up a yeast quality control program, it may be something to invest in to reduce labour and costs down the road.
- Kimwipes to clean up your haemocytometer after you are done counting.
- Plastic or glass tubes for sample collection and dilutions. This will depend on your operational demands, as well as the scale you plan to operate at (micropipettes vs serial dilutions). You may re-use your dilution tubes, but keep in mind this means samples are not fit for additional quality assurance checks due to contamination risks and should be limited to only performing counts.
Collecting a Sample
Once your QC lab is up and running, and the proper equipment has been purchased, you can begin performing basic QC work on harvested yeast samples. After your yeast has been harvested in your brink/bucket/carboy/jar, aseptically collect a sterile sample of the yeast near an open flame. You’ll want to ensure your sample is as homogenized as possible otherwise you will get inaccurate numbers, so shake/mix it well before you sample.
When collecting your sample do whatever you can do help protect the integrity of the yeast to reduce the risks of contamination. If your yeast is in a sealed vessel such as a brink you can simply use CO2 to push out the yeast from your pitching hose. Spray the surface down with sanitizer (we like isopropanol, but there are plenty of options) and collect a sample. You can also create an aseptic environment by creating a controlled flame using a Bunsen burner, and using a disposable 10-25mL pipette to collect a sterile sample (you can also get re-usable glass pipettes, in which case you would want to wash with caustic/PAA and autoclave if possible).
Calculating Dilution Ratios
Now that the yeast is harvested, and a sample has been collected, you need to perform some tests to ensure its quality. Since the slurry is dense, you will need to dilute it in order to count it on the haemocytometer. A dilution is calculated by taking your slurry volume and dividing it by the sum of your slurry and solvent (usually water).
For example, to create a 1:100 dilution of yeast slurry you can perform the following based on available equipment:
Add 100 µL of yeast slurry to 9.9mL of water using micropipettes (our preferred method)
Add 1mL of yeast into 99mL of water using 10-25mL pipettes.
Perform a serial dilution to reach a 1:100 dilution ratio by performing multiple steps using 10-25mL pipettes:
STEP 1: 1 mL slurry to 9 mL water (1:10 dilution ratio), followed by
STEP 2: 1 mL of that thinner slurry added to 9 mL of water (1:10 x 1:10 = 1:100).
Preparing Yeast for Counting
After the dilutions are successfully prepared, a small amount of the 1:100 dilution is then mixed with an equal amount of dye in a 1:1 ratio (Methylene Blue, Trypan Blue, etc.) to facilitate identification of dead yeast cells. This results in a 1:200 final dilution, which will be important later when calculating the density of the yeast slurry. A small sample of the dyed yeast is then added to the haemocytometer chamber via capillary action. This is where having a micropipette would be useful to have, as most haemocytometers have a sample capacity of ~10µL. If you do not have a micropipette, an eyedropper or disposable 1mL pipette dropper could be used. Check out this video on how to properly load a haemocytometer.
A haemocytometer contains numerous cell counting regions for varying sizes of cells such as blood (quite large) and yeast (small-ish). For counting yeast, there’s a smaller counting region within the center of the counting chamber that looks like Figure 3 below. It contains 25 regions, and for counting yeast (or any evenly-distributed sample), only the four corners and middle regions will be counted. Within each of these regions (pink highlighted regions in Figure 3) is 16 squares, all of which will be filled with cells and counted to obtain the correct cell density. From this count, we can estimate how many yeast cells are in our sample.
Figure 3: Zoomed out view of yeast counting regions on Haemocytometer, with the pink squares indicating which regions are to be counted. Each region contains 16 sub-regions, all of which are to be included in the counting process.
Please note that counting at the magnification shown above is nearly impossible and not recommended, so zooming in is necessary to be able to identify the difference between a live and dead cell. Often, this is achieved by using the 40x objective lens (400x total magnification) so it is important to source a microscope capable of reaching this level of magnification.
Which yeast to count? When is it dead or alive?
It is important to count both the alive and dead yeast within the counting regions, in order to know the viability and overall health of the yeast slurry. When a yeast cell is dead or dying, the cell is unable to process the dye and thus the cell remains blue. It is safe to assume that any tinge of blue in the cell means the yeast won’t contribute in any significant way to your fermentation, and should be added to the “dead yeast count”.
Figure 4: Zoomed in view of a single counting region, with the arrows showing blue, dead yeast cells.
Another important aspect of counting is which areas of the grids should be counted. As seen in Figure 5, you do NOT want to count all of the yeast on every edge (along the border, displayed as three closely grouped lines) of the counting area. Doing so results in artificially inflated and inaccurate counts. Instead, two adjacent edges should be chosen (we like to choose the top and left edges, which also are the edges used in Figures 4 and 5) and constantly use the same ones for all counts you perform. While not required, it is best that all staff who perform counts use the same regions and edges to provide more consistent and accurate results.
Figure 5: Proper counting technique for consistent and accurate counts following the Top and Left edge rule. Note that only cells counted on the top and left edges are counted, while cells on the bottom and right edges are not. Additionally, all the white cells would also be counted in this example (but were not illustrated to emphasize the importance of the edges).
Finally, once all the samples have been prepped and the counts have been conducted, some math has to be done to calculate the final cell density and overall yeast viability. The following equation will help you calculate both density and viability:
For example, a sample with a dilution ratio of 1:200 (if you followed our dilution method above) yielded a count of 134 live cells and 14 dead cells. What is the cell density and viability?
Lastly, we can use the viability and density information to effectively and accurately estimate how much yeast is needed for the next brew (and this is where the cost-effective savings come in!!). Let’s determine how much yeast would be required for the following scenario:
The cellaring team has harvested two full 30L yeast brinks of slurry that has a density of 1.34 billion cells / mL and a viability of 95%. The head brewer wants to use the yeast for a 15hL batch at 18°P. How much yeast was harvested and is there sufficient yeast for a pitching rate of 1.5 million cells / mL / degP?
By using this simple equation and performing basic quality control measures, not only did we determine how much yeast was needed for the brew, but we also were able to control the pitch rate rather than use both yeast brinks in a single brew - preventing off-flavours and additional labour while increasing production capacity!! We know that pitching rates can be tricky, so we built a straightforward tool to simplify the process. We also recommend setting up a spreadsheet to track cell counts, pitch rates and yeast generations. Regular cell counting and viability tracking can improve fermentation consistency and help identify when yeast cultures need to be replaced.
Controlling yeast pitching rate helps improve consistency, increases yeast efficiency and reduces overall costs. It is very possible that a single yeast harvest is excessive for the next batch to be brewed, meaning that a single harvest can often be split between multiple subsequent tanks. Thus, proper yeast cell counts can vastly improve the reuse of yeast and increase the value out of each yeast generation. Proper pitching rates also ensure healthy yeast growth and reduces the average age of the yeast cells in the slurry.