Yeast Banking II: Methods to Manage the Bank
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
15 thoughts on “Yeast Banking II: Methods to Manage the Bank”
Hi Bryan, I’ve been following your guide for the yeast bank and its been working great. But there is one piece that is lacking. The regeneration. Do you do it? How often? What is the most practical way of regrowing a large number of samples at once?
Part 4, please? 🙂
The answer is “it varies”. I have access at work to a scientific -80C freezer, which is where I keep my main stocks. At -80C, these stocks are good (in theory) for over 50 years and do not require regeneration.
In a home freeze (-20C), assuming you’ve kept them in an insulated container to reduce the impact of defrost cycles, they should be good for 3 years. For these, you simply need to use the old culture to start a new culture that you re-freeze.
Unfortunately, there is no way to improve “throughput” if dealing with a large number of samples. Probably the best bet would be to have a schedule on which you regrow and refreeze 5 or so samples at a time, spreading the work out over several weeks. It’ll be time consuming, but at any one time you have a managable number of cultures.
I do have a new video series in the works, and one of the videos will be on managing a yeast bank. I have no ETA on that though, as life right now leaves me with very little free time.
I have been reading your articles carefully and watching your videos. I would like to start a yeast bank of my own, but I am looking for a way to do it with a modest investment in equipment. I was wondering if instead of an orbital shaker a stir plate could be used? Also, what is the reason for using the centrifuge? Could a couple of days in the fridge do the same thing?
I managed to find some inexpensive orbital shakers, but finding centrifuges with 50ml capacity for a reasonable price has been difficult. I could find 20ml centrifuges for a reasonable price. Could I just split the 50ml of wort from the shaker into two 20 ml tubes and spin that? Then you would not have to split the yeast from the 50ml tube when moving to the 2ml cryo-tubes. What do you think?
I have a video entitled “freezing yeast” on my youtube page that shows a more home-friendly version that doesn’t need centrifuges or shakers. That may be a better entry point for you: https://www.youtube.com/watch?v=o0zluMPcwrY
That said, you don’t need a centrifuge or a shaker. In place of a tube & shaker you can use a 150 ml flask + mini-stirbar + stirplate to grow up 50 ml cultures. These flasks will fit into an instapot-type electric pressure cookers, so you can ensure proper sterilization of the wort and flask. Grow on your stirplate as per normal. In lieu of a centrifuge, you can simply transfer the culture from the flask to a test tube (or more than one if you don’t have ones big enough). Place in the fridge for 1 to 2 days to let the yeast settle – no centrifuge required. You can then resuspend the pellet in freezing medium and put into a cryotube.
Hi Bryan, does the denatured protein in the 10% wort that coagulates during autoclaving present a problem? Is it ok to spin it down together with the yeast, or is it better to be removed prior to incubation?
Its not an issue, other than it may lead you to over-estimate the amount of yeast you have. If you are concerned, you can filter it out using a coffee filter, and then re-sterilize the medium. I often do this for medium I am preparing for agar plates, as the lack of precipitate in the plates makes it easier to find small colonies.
Hi Bryan, thanks for sharing your experience. I found your blog when looking for wild yeast capturing and I must say it is a great source of information.
Recently, my friend gave me some frozen yeast cultures in 1.5 ml tubes. Because I have only one tube of each strain and I have no backup I decided to multiply so that I have at least two tubes of one strain in my freezer and one backup tube in my parent’s house.
How shall I proceed? Use inoculating loop and inoculate 7ml wort and then to 250ml and after that divide the culture into four 50ml tubes (or one 50ml tube and use the rest for starter) and proceed as you wrote. Or inoculate the 50ml directly?
I also have to consider that I have no orbital shaker just stir plate. Will it be sufficient?
I would use a loop to inoculate 5-7 ml of wort with the yeast (be sure to return your original vial to the freezer ASAP), and then use that to inoculate larger volumes. There is a higher risk of infection going from a loop of yeast (which is typically 0.01 to 0.1 ml in volume) to a larger volume, so its best to start with a smaller volume, and then use that to inoculate larger volumes.
For the small tube, I wouldn’t worry about mixing it – just get the yeast stirred in with the loop and then let it sit somewhere warm until growth is done. You can then inoculate your larger volume, and use the stirplate to maximise growth.
Damn your quick! 😊 Thank you so much for the answer.
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Thanks very much for your reply! I was worried about the soot coming from meths. Although now I see on e-bay there are some pretty cheap high % ethanol solutions I'll pick up (it's hard to get ethanol in the UK, it's only allowed for research purposes) :(.
Denatured/methylated spirits would work fine. In fact, the earliest alcohol lamps used by microbiologists burned pure ethanol – i.e. you could use everclear…but that would be expensive!
Hi, would denatured alcohol/methylated spirits be a usable fuel for an alcohol lamp or would fondue fuel be a better source?
Many thanks in advance for your time,
I use a mix of penicillin and streptomycin; although honestly, only the streptomycin is needed. I use a pre-mixed 100X solution, which in the prepared plates/media works out to 100U/ml penicillin and 100ug/ml streptomycin.
Hi Bryan. Thanks for spending time on writting this. It has been helpful.
Could you please explain a little more about the concentration of antibiotics you use for wort or agar?