White Labs Microscope Kit (product # MA1400) includes:
-Methylene blue solution
-Fine tip glass pipets
-Hand held counter
You will need to provide: Microscope with 400X capability
1. Make certain hemocytometer is clean and dry before use. Hemocytometer can be easily cleaned with water. Chamber and cover slip may be scrubbed gently using a lint free towelette (kimwipe).
2. Dilute yeast sample to an appropriate concentration. Usually a 1:100 or 1:1000 works well for yeast slurry. It is best to have 100 or fewer yeast cells per microscope field at 400X (40x lens + 10x eyepiece). Sample can be diluted with deionized/distilled water or with 0.5% H2SO4 if cells clump excessively. Note your dilution factor.
3. Invert sample several times to mix, taking care not to introduce bubbles. Degas if necessary.
4. Mix 1ml of your diluted yeast sample with 1ml of methylene blue solution and allow to incubate for 1 - 2 minutes.
5. Invert sample 1-2 times, and take up sample by placing a glass pipet tip into the liquid mixture and letting it fill via capillary action (draw upwards automatically) or using a transfer pipette.
6. Fill chamber by placing 2 drops of sample onto center of counting grid.
7. Gently position cover slip so that glass covers both counting areas equally.
8. Carefully place hemocytometer on microscope stage. As you focus at each objective leading up to the 40X lens, note the distribution of yeast cells. If cells are well distributed then you can use the short cell count method. If cells are grouped or clumped together, you may need to prepare another sample or use the long cell count method.
Considerations for when cell counting
1. You will be counting squares within the 1mm2 ruled area centrally located on the chamber (see images above)
2. It is helpful to establish a counting protocol for all cell counts and stick with it! Consistency is key!
--For example, cells touching or lying on the top and right boundary lines are not counted, whereas cells touching or lying on the bottom or left boundary lines are counted (see image below).
3. Yeast cell buds emerging from mother cells are counted as a separate cell if the bud is at least one-half the size of the mother cell.
4. If performing viability counts, dead cells will stain dark blue. Non-viable cells do not have the metabolic capability to expel the intruding dye.
5. Do not count cells that are pale blue in color as dead. Some budding cells will also stain blue, do not count these cells as non-viable.
6. If you are performing a cell count and viability count simultaneously, it is best to count all cells on the hand held counter and record noted dead cells on a written tally.
Short method: For an evenly distributed sample
a. You will be counting cells within the 5 numbered squares
b. Estimating number of cells in total grid = # of cells counted in 5 numbered Squares x 5
c. Yeast cells/ml = Number of cells in total grid x dilution factor x 104 (or 10,000, depending on the volume of the chamber. This number is a constant.)
--Note: 106 are millions and 109 are billions. Volume of the chamber. This number is a constant.
Yeast Cell Counting and Viability Staining Calculations
Squares (if you count all 25 squares, you will not need to use this number in the calculation)
25 total/5 counted = 5 (multiply by this number to get the total number of squares in the grid)
200 (two 1:10 dilutions, then a 1:2 dilution of yeast and methylene blue)
Volume of chamber (constant)
= 220 x 5 x 200 x 104 = 2200000000 = 2.2 x 109 or 2.2 billion cells/ml
# Live cells/Total cells counted x 100= % Viability
**View step-by-step instructions for performing your own cell counting/viability testing, explained by Lab Operations Manager Kara Taylor.