As discussed in my last article, I am now managing a yeast bank on behalf of the London Home Brewers Guild. In the previous article I outlined the methods we use, and the rational behind using those methods over other conventional methods. In this post, I will outline how the bank itself is run, with detailed protocols for the clean-up, freezing and “withdrawal” of the yeasts.
- Preparing & freezing the yeast
- Checking for contamination
- Secondary cleanup
- How to perform a withdrawal
- Counting yeast
Preparing & Freezing the Yeast:
Sterile media with antibiotics (left),
fresh White Labs yeast and washed
yeast (tubes in racks) as well as
micropipettors, in a biosafety cabinet
Prepare 50ml of 1.035 to 1.040 wort from DME, in a 250ml flask. Cap with foil and boil for 10 min or autoclave in wet cycle to sterilize. Once cooled, add antibiotics at the desired concentration.
- Prepare a solution of 30% glycerol in 1.035 to 1.040 wort. Autoclave to sterilize. A large volume of this can be prepared in advance and stored in the fridge; only 2ml is required per banked yeast. For 100ml, add 30ml of glycerol to 70ml of water and 9.8g of DME.
- In a biosafety cabinet, add yeast sterilely to the 50ml flask of wort, using the following guidelines:
4 strains on the shaker
If starting with a yeast slurry (i.e. yeast from a White Labs tube, washed or top-cropped yeast, etc), suspend the slurry in an approximately equal volume of water or wort. For white lab yeasts, simply mix the sedimented yeast into the liquid contents of the tube. Add 500ul (0.5ml) of this slurry to the wort.
- If starting with an active liquid culture (i.e. starter, yeast cultured from a bottle, wyeast smack-pack), transfer 5ml of the active culture to the wort.
- If starting with a dried yeast, add 5-10 prills to the wort.
- Place on a shaker, 20-24C, at 200-250RPM, for 18 to 24 hours. This will let the yeast grow to a density of 5 to 10 million/ml. This is the point where we want to harvest the yeast, as they are minimally stressed; longer incubations will produce larger numbers of yeast, but the increasing alcohol concentration will stress the yeast, leading to poorer recovery after freezing.
Transfer the culture to a sterile 50ml centrifuge tube. Spin for 10 min at 500g – this is sufficient to pellet all but the least flocculant of strains; if a large pellet is not obvious by the end of the centrifugation period, re-spin at 750g for 10min. If possible, avoid the higher g-forces as these can damage the yeast.
Remove the supernatant and suspend the yeast pellet in 2ml of 30%glycerol/wort. Sterially transfer to two sterile cryovials, 1ml per vial.
– If desired, remove 10ul to test for contamination (see next section)
- Freeze at -18C to -80C. Store the duplicate vials in different freezers to insure against loss.
Checking for Contamination:
|S-05 Yeast (large cells in middle) in a culture heavily|
contaminated with acetobacter (small, rod-shaped cells). Yeast
are out-numbered 100:1 by bacteria in this infection
- Pipette 10ul of yeast-containing solution onto centre of the slide. For concentrated sources (i.e. pure yeast slurry), you may need to dilute 1:10 (tap water is fine) before this step.
- Place cover glass over droplet, allowing yeast solution to spread between the cover glass and the slide.
- Image under the microscope. Yeast will appear as spheres or buds (see above image), contaminating bacteria will be much smaller, and either spherical or rod-shaped in morphology. Debris will appear as black dots – do not confuse these with bacteria! For a comparison, an equal volume of the sterile media used to grow the yeast can be viewed.
- View at least 10 fields; if there is less than 1 bacteria per field (40x) or less than 1 bacteria per 2 fields (60x) your stock is clean enough for most uses.
|S-05 from the infected culture (see above),|
after clean-up method 1. No contaminating
bacteria are present.
- Put 5ml of sterile 1.035 to 1.040 wort into a sterile culture tube.
- Add antibiotics at 2X the normally used concentration.
- Inoculate a small volume (10-20ul) of the contaminated yeast.
- Grow for 24 hours, room temperature, with continuous shaking to oxygenate and suspend the yeast.
- Transfer 10-20ul of this solution into a new tube of 5ml 1.035 to 1.040 wort + antibiotics, and culture for another 24 hours.
- Check for infection; if clean use the 1ml of the culture prepared in step 5 to inoculate a 50ml flask (with antibiotics). Culture and freeze as per usual.
- Prepare a sterile agar plate of YPD yeast media, or sterile plate of 2% agar/1.035 wort, with antibiotics at their normal concentration. DO NOT double the antibiotic concentration for this method.
- Streak 20-50ul of the contaminated yeast onto the plate.
- Culture at room temperature until visible colonies form (generally 24 – 48 hours)
- Prepare 4-5 culture tubes containing 5ml 1.035-1.040 wort + antibiotics, for each tube pick a single colony off of the plate and add to the tube.
- Grow 24 hours (shaking, etc) at room temperature and check for infection. Tubes showing a pure-yeast culture can be pooled, and 1ml used to inoculate a 50ml flask (with antibiotics). Culture and freeze as per usual.
How to Perform a ‘Withdrawal’:
to pitch into a
Withdrawals are easy – a small amount of yeast are removed from the frozen stocks, placed into 5 to 7 ml of sterile media, and grown for 24 to 36 hours. This tube of yeast – containing anywhere from 25 million to 700 million yeast – can be pitched into a 250ml starter, then into a 1.5L starter, providing the ~100 billion cells needed for the average 5 gallon ale (an additional step-up is needed for larger volumes or lagers). Because the freezing media, growth media and starters are antibiotic free, there will be no residual antibiotics going into the beer. The withdrawal protocol will only cover the actual withdrawal process – stepping-up will be a topic of a later post.
- Using sterile methods, transfer 5 to 7ml of 1.035 to 1.040 wort into a sterile culture tube
- Using a flame, flame and cool a transfer loop. Use this to collect 10-20ul (one loops worth) of yeast from the frozen vial. Be sure to cool the loop as to not kill the yeast – to be certain, touch the loop to the inside of the cryovial before contacting the yeast. Never touch the outside of the cryovial (or any other potentially contaminated surface) before putting the loop into yeast.
- Swirl the yeast-filled loop in the culture tube of media
- If desired, check for contamination, then pitch into a 250ml starter.
- Assemble the hemocytometer by placing a cleaned cover glass onto the cleaned hemocytometer.
- Inject 10ul of yeast between cover glass and the hemocytometer
- It may be necessary to dilute the yeast sample 1:10 or 1:100 to get countable numbers
- Count the number of yeast in the four corners of the hemocytometer grid (i.e. the four areas containing the largest-sized grid).
- The number of yeast per ml is = (number of yeast cells counted/4) x 104 x dilution