Freezing Brettanomyces

Eureka, another Brettanomyces post. This time about a feasibility study if you can freeze Brettanomyces like any other Saccharomyces strain. I would hereby like to discuss my latest results.

All started by preparing some Brettanomyces strains I either bought or isolated for cryo storage like described in a previous post of mine concerning freezing yeasts. I put the following Brettanomyces strains in my -20°C (-4°F) freezer in August/September 2012:

Brettanomyces isolated from WY3191 Berliner Weisse blend
Brettanomyces isolated from Girardin Gueuze
Brettanomyces isolated from 3 Fonteinen Gueuze
Brettanomyces isolated from Cantillon Kriek (3 strains)
Brettanomyces isolated from Cantillon 2007 Lou Pepe Gueuze (2 strains)
Brettanomyces bruxellensis (WY5526)
Brettanomyces lambicus (WY5112)

Some isolates consisted of more than one strain which were separated during trial runs with bromocresol green (not published). All these different strains were frozen separately.

In mid November 2012, the Brettanomyces were taken out of the freezer and transferred into fresh YPD media. After two weeks some of the yeasts showed signs of growth such as turbid media and gas production. In the end all media showed signs of activity and formed off-white coloured sediments. Both yeasts isolated from the Cantillon beers even showed signs of pellicle formation (not shown). Although activity could be observed it still has to be evaluated if the activity originates from the yeasts and not any contamination. Due to lack of time the yeasts remained in the YPD media for nearly two months until further experiments could be conducted.


Some micrographs showing the yeasts from YPD liquid cultures before freezing and afterwards.


Fig 1: Brettanomyces from Cantillon Kriek before freezing

Typical elongated cell shape of Brettanomyces visible (Fig 1 and 2). Even some hyphae formation (Fig 2). Somehow the colonies in Fig 1 look smaller than the ones in Fig 2 although both pictures were taken with the exact same setup.


Fig 2: Brettanomyces from Cantillon Kriek after freezing


Fig 3: Brettanomyces from Cantillon Lou Pepe after freezing

Again some hyphae formation (Fig 3).

Concluding from the micrographs shown (Fig 1-3), Brettanomyces yeasts could be found in the YPD media after reviving them. Although Brettanomyces yeasts could be observed in the microscope observations still does not prove that the yeasts are viable. Liquid cultures were first streaked on some Sabouraud agar plates and incubated at room temperature until colonies were visible. Colonies were then picked from the Sabouraud plates and streaked on Sabouraud agar with an addition of bromocresol green.

Some yeasts had different morphologies like WY5526 B. bruxellensis on bromocresol green containing agar media (Fig 4). Some colonies grew as green, others as white ones. For the next agar platings, each a white and green colony were picked.


Fig 4: WY5526 B. bruxellensis on bromocresol green agar

One strain of Cantillon’s Kriek and the strain isolated from a 3 Fonteinen Gueuze grew in different morphologies (white and green colored colonies) as well and were treated separately for the next agar platings. Please further notice that WY5112 Brettanomyces lambicus was not streaked on Sabouraud Bromocresol green due to a mold contamination on the first Sabouraud plate. However, typical colonies of B. lambicus could be observed (not shown).

Agar plate results

All the revived Brettanomyces strains formed colonies on Sabouraud Bromocresol green agar (Fig 5-7).


Fig 5: Brettanomyces on Sabouraud agar after six days of incubation. Left: Cantillon Kriek_green colony (B04_green); Top: Cantillon Gueuze 2007 (B05); Right: WY5526 B. bruxellensis colony 1; Bottom: WY5526 B. bruxellensis colony 2

All the yeasts grew as white colonies expect the one known to grow as green colonies (B04_green) (Fig 5). In the case of WY5526 B. bruxellensis, the two picked colonies from Fig 4 showed the same morphology again. Both grew as white colonies (Fig 5).


Fig 6: Brettanomyces on Sabouraud agar after six days of incubation. Left: Cantillon Kriek (B04_2); Top: Cantillon Kriek (B04_1); Right: Girardin (B01); Bottom: Cantillon Geuze 2007 (B05 dark_1)

The two colonies that grew differently on the first bromocresol green agar from the Cantillon’s Kriek isolate grew again as white colonies (Fig 6).


Fig 7: Brettanomyces on Sabouraud agar after six days of incubation. Left: Cantillon Geuze 2007 (B05 dark_2); Top: Newly isolated Brett (nothing to do with this experiment); Right: 3 Fonteinen (B02_2); Bottom: 3 Fonteinen (B02_1)

The same is true for the different morphologies from a 3 Fonteinen isolate (Fig 6, 7). The Brettanomyces strain(s) isolated from WY3191 Berliner Weisse blend formed colonies as well (not shown).

In all cases, the bromocresol agar media turned from a blue color to yellow indicating the secretion of acid. Some plates even had a strong acetic acid smell. An un-streaked bromocresol agar media was included as a control and the color remained blue throughout the whole experiments (not shown).

Summary/ Conclusion of agar platings

It could be shown that all the frozen Brettanomyces strains formed colonies on Sabouraud agar. Some of the isolated yeasts grew in different forms (white and green colonies) but such a differentiation could not be observed after a second streak on agar media. This is not the case for the yeast strain isolated from Cantillon’s Kriek (B04 green) which grew in green colonies on every plating.

The differentiation between Brettanomyces and Saccharomyces based on bromocresol green and its issues will be covered in a future post. One could already observe in these platings that some of the yeast colonies grew as green colonies in a first run but grew as white colonies in a second run again.

Micrographs of the colonies

At last some micrographs of the colonies.


Fig 8: WY5226 Brettanomyces bruxellensis (5526_1)


Fig 9: WY5226 Brettanomyces bruxellensis (5526_2)

The two different samples of WY5226 B. bruxellensis look very similar (Fig 8, 9). The differences in color appearance on the bromocresol green might be due to some issues of bromocresol green as an indicator for Brettanomyces and wild yeasts. As mentioned already, more about that in a future post.


Fig 10: Brettanomyces from Girardin Gueuze (B01)

Typical Brettanomyces cells visible in the Girardin isolate (Fig 10).


Fig 11: Brettanomyces from 3 Fonteinen’s OudeGueuze (B02_1)


Fig 12: Brettanomyces from 3 Fonteinen’s OudeGueuze (B02_2)

Both colonies from 3 Fonteinen isolate seem to be Brettanomyces (Fig 11, 12). Hard to tell based on the morphology of the cells if the two samples are the same or not.


Fig 13: Brettanomyces from Cantillon’s Kriek (B04_1)

Not a lot of elongated cells were visible in B04_1 (Fig 13) like in B04_2 (Fig 14). Maybe these two samples are not the same strain of Brettanomyces. Maybe B04_1 is not a Brettanomyces strain. Further studies are necessary.


Fig 14: Brettanomyces from Cantillon’s Kriek (B04_2)


Fig 15: Brettanomyces(?) from Cantillon’s Kriek (B04_green)

Some elongated cells visible in B04_green (Fig 15). Yet a lot of the cells looked like Saccharomyces cerevisiae. Might be a mixture of S. cerevisiae and Brettanomyces.


Fig 16: Brettanomyces from Cantillon’s Kriek (B04_white_1)

Lots of hyphae visible in one of the isolates from Cantillon’s Kriek (Fig 16). Very typical for Brettanomyces.


Fig 17: Brettanomyces from Cantillon’s Kriek (B04_white_2)

On the other hand, in the second sample of B04_white not that many Brettanomyces cells visible form hyphae as shown in Fig 16 (Fig 17). Yet again, maybe these two samples are not the same strain of Brettanomyces. Further studies necessary.


Fig 18: Brettanomyces from Cantillon’s Lou Pepe 07 Gueuze (B05)

Lots of elongated, boat-shaped cells in Cantillon’s Lou Pepe isolate B05 visible (Fig 18).


Fig 19: Brettanomyces from Cantillon’s Lou Pepe 07 Gueuze (B05_dark_1)

The second strain from Cantillon’s Lou Pepe looks different from the first one shown in Fig 18 (Fig 19). Hard to tell if the second sample B05_dark_2 is another strain than B05_dark_1 or not.


Fig 20: Brettanomyces from Cantillon’s Lou Pepe 07 Gueuze (B05_dark_2)

Summary/ Conclusion of micrographs

Brettanomyces were visible in most of the micrographs shown above. However the shape of Brettanomyces can differ significantly. The experiment strongly suggest that it is possible to freeze Brettanomyces and successfully revive them. In addition, it could be shown that some of the frozen samples might contain further strains of yeasts. Additional experiments are necessary to further look into this possibility.

Unfortunately, the yeast isolated as Brettanomyces from WY3191 Berliner Blend looked very similar as Saccharomyces cerevisiae cells (not shown). It might be possible that the yeast isolated from the blend wasn’t a Brettanomyces strain in the first place. But beside the S. cerevisiae colonies were some smaller colonies visible as well (Fig 21).


Fig 21: Isolated cells from WY3191 Berliner Weisse blend

The cells shown in Fig 21 are no yeast cells. Theses cells look like Lactobacillus. Interestingly, these bacteria cells were in the freezer as well (therefore possible to freeze Lactobacillus like yeast cells as well) and they grew on bromocresol green containing agar.

Enough with the experimental part. I am very happy about theses results. Took me some time to do all the platings, micrographs but I have the feeling it was all worth the efforts. At least now I know that I can easily freeze all my Brettanomyces strains I have (well over 20) without any worrying. The only thing to keep in mind here is that theses yeast might take some additional time for reviving than normal S. cerevisiae strains. Some preliminary results even suggest that it is possible to freeze Lactobacillus like you would any yeast cells.

The next post will be about another big yeast experiment. Stay tuned and thanks for commenting!


Yeast basics: Check yeast viability

Eureka, its time for another yeast basics post. This post is all about yeast viability. Viability describes how many living cells there are in a cell population. The higher the viability the more living cells you have. Lets apply this definition to homebrewing. A high viability is very important in brewing since only living yeast cells can ferment wort. Pitching the right amount of yeast cells is important. However, viability is important to consider in the pitching rate calculations as well. Pitching a yeast with a low viability is basically the same as underpitching. Not only is it important to pitch the right amount of yeast but having viable yeast cells as well. Let me walk you through some of the most important facts concerning yeast viability and homebrewing.

How do you determine the yeast viability?

I would like to show you one example how you can determine the yeast viability by use of a microscope. If you do not have a microscope or are not interested in how it is done, please skip this chapter.

The easiest way to determine the viability is to stain yeast cells with methylene blue and count the yeast cells as described in a previous post about yeast counting. Methylene blue stains the dead yeast cells which then appear as dark blue cells. This could look like shown in Fig 1. The dark blue cells are dead, the other ones still alive.

Fig 1: Methylene blue stain of yeast sample

To perform a viability count, mix equal parts of your yeast solution with a 0.1% (w/v) methylene blue solution. Mix well and let it react for one minute then count the cells by use of a counting chamber. Concerning the methylene blue solution. Mix 0.1 g of methylene blue and dissolve it in 100 mL of distilled water.

First, the cell counting is done the same way as you would do it without the methylene blue staining. The only difference, you count the dead cells in parallel in the different squares of the counting chamber. If you like further information about yeast counting, please have a look at a previous post of mine concerning yeast counting. Now, count all the yeast cells including the dead (dark blue) ones. Lets say you counted 105 cells in total. Second, you count all the dead ones. Lets assume you counted 15. Next, you calculate the viability:

Viability [%] = (Total counted cells – total counted dead cells) / total counted cells x 100

In our example, you get a viability of 86%. Meaning that 86% of the yeast cells are still alive. 14% are dead. For further information about the methylene blue staining please consult and the book called Yeast: The Practical Guide to Beer Fermentation” written by C. White and J. Zainasheff.

Include viability in the pitching rate

Assume you want to ferment a 20 L (5.3 gal) ale batch with a gravity of 12°P (OG 1.048). In this case you need to pitch: 20000 mL x 12°P x 0.75E6 cells mL-1 °P-1 = 180E9 yeast cells. You can calculate the pitching rate with a calculator as well if you want. Lets assume you made a 1 L starter, diluted the sample by 1:100 and counted a total cell concentration of 1.3E9 cells per mL (including the dead yeast cells). To get the appropriate amount of yeast for pitching you need 138 mL of the starter (180E9 cells divided by 1.3E9 cells mL-1 = 138 mL). We also determined the viability in parallel to be 86%. In this case you would need around 160 mL of the yeast starter to have the right amount of yeasts (138 mL divided by 0.86 = 160 mL). You can easily see, the lower the viability is the more volume of the yeast starter is necessary.

How to determine the viability without counting yeast cells?

I assume most of the readers here do not have a microscope, therefore have no counting chamber and no methylene blue. How could you determine the viability without a microscope? As far as I know, there is no other way to do it. Well there are ways but these methods are even more expensive than a simple microscope, a counting chamber and methylene blue. The easiest way to do it is using assumptions. A few things to get good assumptions:

1. Using commercial liquid yeasts: As far as I know there is a production date on every liquid yeast package you buy from the main two yeast suppliers (Wyeast, White Labs). It is therefore possible to estimate the yeast viability based on the manufacturing date if the yeast is stored in a refrigerator:

Fig 1: Viability as a function of storage time for yeast stored in a refrigerator

Every day, you lose viability (Fig 1). Storing the yeast for a month, you already lost about 20% of the living yeast cells! Another month and you already lost nearly half of the living yeast cells! You can calculate the viability with J. Zainasheff’s (aka mrmalty) yeast calculator and the date printed on the yeast package. This works if you pitch the yeast from the package/vial directly. But please avoid pitching any yeast with a viability less than 90-80%. I recommend to make a yeast starter if the yeast sample has a low viability.

Please keep in mind, we are speaking about assumptions here. I am sure the viability decrease is dependent on the yeast strain, storage conditions, storage temperature, pH, oxygen level etc. etc. etc. However, this does not change the fact that yeast viability only decreases over time and therefore not recommendable to store yeast for longer than a couple of months.

2. Using yeast from a starter: If you do a yeast starter, only the viable cells will divide. The dead ones remain and decrease the viability as they are still present in the starter. If you do a starter from a Wyeast Activator package being 40 days old, the viability of the yeast is around 70% (Fig 1). Meaning, only 70% of the yeast cells are still alive. If you now do a 1 L starter with this yeast, the 30% dead cells will remain. The maximal viability you can obtain with this 1 L starter-step will be around 80% (Fig 2).

Fig 2: Maximal yeast viability obtained by a 1 L starter

However, after the 1 L starter, you have approximately 125E9 viable yeast cells in the starter (Fig 3). A Wyeast package originally contains about 100E9 cells. Meaning, if you store your Wyeast package for 40 days, the viability is at 70% (Fig 1). From the original 100E9 cells, only 70E9 will be still viable. Then do a 1L yeast starter and your viability afterwards will be around 80% (Fig 2) and you have 125E9 viable yeast cells in total (Fig 3). Let me explain how the chart in Fig 3 must be read to get to the 125E9 cells. As mentioned a Wyeast Activator package has roughly 100E9 viable yeast cells at the beginning. Then store it for 40 days and do a 1 L yeast starter. Use Fig 3 to get from the 40 days to the 125% (or the equation). For the equation, use 40 days as x to get the 125% (as y). This means, from the original 100E9 viable cells you now have an increase of 125%, which is equal to 125E9 viable yeast cells in total.

Please notice, Fig 3 only works for Wyeast Activator packages and White Labs vial with an original yeast amount of 100E9 cells and a 1 L starter size. However, the equation from Fig 3 does not work for other yeast samples and other starter sizes. For these cases, use the calculator mentioned below.

Fig 3: Increase of (viable) yeast cells from Wyeast Activator package/ White Labs vial after a 1 L starter step vs. yeast age

Things to remember: The viability of yeast decreases over time (Fig 1). On the other hand, you cannot reach a 100% viability of an old yeast sample since the dead cells remain (Fig 2). The older a yeast sample is, the harder it is to increase the viability. And to estimate the viable yeast cells of a yeast sample after a 1 L yeast starter, use Fig 3.

Lets make a final example to show you how to use the charts:
Assume you want to ferment a 20 L (5.3 gal) ale batch with a gravity of 8.8°P (1.035). For this batch you need to pitch 132E9 cells. Assume you have a Wyeast package or White Labs vial being 30 days old. Now, use Fig 1 to determine the viability which is 76%. The Wyeast Activator/White Labs vial therefore only has 76E9 viable cells left. So not enough for a direct pitch. You therefore need to do a 1 L yeast starter and you will have 132E9 yeast cells at the end (Fig 3). That is just enough yeast to pitch. You therefore can pitch the yeast from the whole 1 L starter and have the right amount of yeast.

Because these charts only work for 1 L starters, use the yeast calculator from J. Zainasheff ( for any other batch sizes, starter sizes etc. In this particular calculator all the assumptions and such are already implemented. I really like programs where you can fill in numbers and you get a result (such as the yeast calculator). However, now you know some of the equations behind the yeast calculator and how it maybe work. I am not related to J. Zainasheff or have any financial benefits from referring to his site. I do so because I use his calculator for some time now. I wrote my own calculator based on my own assumptions a few years back and luckily for me, the results from my calculator are very close to the one from J. Zainasheff. So no need for me to improve my calculator any further.

I like to emphasize again, all the charts shown in this post are based on assumptions. I would not be surprised if someone comparing the calculated viability done with the graphs above and a real cell count encounters differences. In addition, I would not be surprised either if differences between yeast strains exist as well. Even a cell count done with a counting device is not 100% correct. Luckily for us, brewing beer is not like rocket science and does not have to be very precise. Cheers on that!

Yeast banking – #3 Isotonic sodium chloride

Eureka, today I proceed with the second technique to bank yeast at home (or in a lab). I am sorry for the delay of this post. I am currently very busy and writing such a post is very time-consuming. The introduction to the banking yeast series can be found here. The post about the first technique (bank with agar plates) can be found here. Lets proceed with another technique, isotonic sodium chloride solutions (or any other sterile solutions).

Description of the technique

One way to store yeasts over a period of time is storing them in sterile solutions such as isotonic sodium chloride solutions. Isotonic in this case refers to solutions which have the same osmotic pressure as the cells itself. The osmotic pressure mainly depends on the salt concentration of a liquid. If you have a liquid with a high salt concentration, the osmotic pressure of this solution is high. On the other hand, distilled water (low to no salt concentration) has a low osmotic pressure. If two liquids with different osmotic pressures are connected by a membrane, the two pressures can equalize: Water from the low pressure solution passes the membrane and flows into the high pressure solution. The water flows until the two pressure potentials of the two solutions are equalized.

Lets get back to the topic. Yeast cells have a membrane as well. It basically surrounds the whole cell. If you store your yeasts in a solution with a higher osmotic pressure than the yeast cells (lot of salts and other organic compounds are in a cell), your yeast cells will eventually die due to dehydration (loss of water). Losing water is not ideal for a cell. Think about humans, losing water can lead to severe health problems as well. On the other hand, storing yeast in distilled water could eventually burst the cells since water thrives into the yeast cell (into the high osmotic pressure solution). This is not ideal as well. The key is to prevent an osmotic pressure difference between the yeast cell and the surrounding liquid. You do so by using liquids with the same salt concentration as in the cell. You therefore have the same osmotic pressures in the cell and the surrounding liquid. And this is called isotonic. If you dissolve 9 g of sodium chloride (a.k.a table salt) in one liter of distilled water, the solution is isotonic. For other salts/compounds, there are lists are available to look up the amounts you need to get an isotonic solution. Sodium chloride seems to be in every kitchen and why not use it to store your yeasts. I will therefore talk only about isotonic sodium chloride from now on.

Fig 1: Glass tube for yeast banking

First of all, you have to think about a containment for your yeast-sodium chloride solution. I used small glass tubes with a screw cap first (volume around 5 mL) as shown in Fig 1. Any other tube will do the job as well. Just keep the sterility in mind. It is best to have a sealed containment to avoid any contaminations. In addition, any containment which can be sterilized in a pressure cooker is even better.

Fig 2: Isotonic chloride ampule

Another way to go is using ampules filled with sterile sodium chloride solution (Fig 2). These ampules are used in hospitals very frequently and can be bought in many pharmacies in Europe. Can’t tell it this is true for any other country though. You only need a sterile syringe and a cannula and inject the yeasts in the ampule. Done!

I used the tubes (Fig 1) first but went with the ampule technique later on because of the volume. I first thought 5 mL of yeast solution are not enough for my purposes. Now I know, most of the ampules still contain about 95% of the original liquid… If you have the same concerns, just fill additional tubes with the same strain you use more often.

If I would consider going into the isotonic solution method again, I would probably use the glass tubes with a volume of approximately 5 mL, fill them with 4- 5 mL of isotonic sodium chloride solution and sterilize them in a pressure cooker. With the ampules you need syringes and cannulae in addition.


– Containment for the yeast-sodium chloride solution (i.e. glass tubes with screw cap)

– Sodium chloride. Common table salt will do the job.

– Distilled water

– Sterile syringes and cannulae to transfer the yeast into the containment


Isotonic sodium chloride solution: Dissolve 9 g of sodium chloride (a.k.a table salt) in 1 L of distilled water. Depending on the water quality, even tap water would work it is low in salts. However, get yourself one liter of distilled water if you can. 1 L of the sodium chloride solution will last for many, many tubes.

Depending on your containment:

– If you can sterilize your containments, fill them with the isotonic sodium chloride solution and sterilize them in a pressure cooker or in boiling water for approximately 15 min. They are ready to go. You can even store the sterilized tubes at room temperature for nearly forever. Just make yourself a batch of tubes and the yeast banking can begin.

– If you can’t sterilize your containments (either because they are made out of plastic and melt during the sterilization process or you do not have an opportunity to sterilize them) you need to sterilize the sodium chloride solution by boiling it and then transfer it into the containments later on. However, don’t forget the sterility of the containment itself. If you buy them as pre-sterilized, well good enough. If you buy something not sterile, disinfect it somehow. Either use diluted Javelle water or Vodka. Just remember a disinfection is not the same as a sterilization. Some microorganisms will survive the disinfection process. Either way, I would not recommend this method. Get yourself some sterilisable tubes and all the worries are gone.

Bank the yeast

Now as the tubes are filled and sterilized, its time to bank the yeast. The easiest way in my opinion is to get the yeast directly from the source such as a fresh Activator package from Wyeast or a vial from White Labs. Use a sterile syringe and get yourself approximately 1 mL of yeast slurry for 5 mL of isotonic sodium chloride solution from the package or vial respectively and transfer the volume into the sodium chloride solution. You are basically done. You could even pour some of the yeast slurry from the package or vial into the sodium chloride solutions directly. Whatever works.

Fig 3: Wyeast’s 3942 Belgian Wheat yeast in sodium chloride solution

On the other hand, you could harvest yeast from Kräusen and transfer them into the isotonic solution. In this stage the yeast cells are very viable and vital. Using harvested yeast could work as well though the viability/vitality could be an issue here. I would recommend doing a small starter with the harvested yeast first (around 10- 50 mL), decant as much of the supernatant off the yeast as possible and transfer the yeast slurry into the isotonic sodium chloride solutions.

In addition, you could transfer a colony from an agar plate into a sodium chloride solution.

To summarize, you could basically use every source of yeast possible. Just keep the vitality/viability in mind. You do not want to bank unhealthy yeasts.


If possible, store all the yeasts in the sterile solutions at around 6°C (43°F). In general, cold temperature would be fine. Just don’t freeze them. They probably won’t survive. If cold storage is no option, store them at a cool and dark place. After a short time, the yeast forms a nice sediment at the bottom of the tube/ampule etc.


To get from the banked yeast to a yeast starter. Collect 1 mL of the liquid from the vial with a sterile syringe (shake before removal to get the yeast back in solution) and transfer to around 100 mL of a 10°P sterile starter wort made with dry malt extract. To get a 10°P starter just add 10 g of any dry malt extract, some yeast nutrients, dissolve in 100 mL of water and sterilize it with a pressure cooker if possible. I use 500 mL Schott bottles for this purpose. Any mason jar will do just fine as well. Just sterilize it in either a pressure cooker or in some boiling water. This is a very crucial step because the yeast cells from the isotonic sodium chloride solutions might be quite slow growers at the beginning. Any contamination in the starter will outgrow the yeast for sure. And don’t use too large starters for this step.

Let the fermentation go for some days (up to seven days if necessary). A small layer of yeast will form. Then increase the volume up to 1 L in total (add 900 mL of freshly sterilized 10°P wort on top or transfer the 100 mL starter with the yeast to 900 mL of fresh wort). After the 1 L starter, there should be roughly the same amount of cells as in a fresh Wyeast activator package (100E9 cells). This may vary between yeast strains. Use a counting chamber to determine the exact cell concentration and cell amount if possible or estimate the cell count from the yeasts volume.

My experiences with this method

I use(d) this method for quite a while and all my yeast from my yeast library are/were in sodium chloride solutions at one point. My oldest strains are in the ampules since Summer 2010 and I could reanimate them successfully in Summer 2012. In my experience, the yeasts can be stored this way for at least two years without any problems (refrigerated at 6°C). This holds true for several different yeast strains. However, I lost some of the strains due to an infection.

 Advantage Disadvantage
Rather easy method  Needs space
No maintenance work  Contaminations invisible
Not a lot of equipment necessary  Long term storage?
Viable > 2 years (@ 6°C)  Storing yeast/bacteria mixtures

All in all, this is a really easy and cheap method in my point of view. I had a hard time to find disadvantages for this method. I do not bank my yeasts with this method anymore because of the space it needed. Please consider that we are talking about 40 different strains in my case. My refrigerator is basically filled with ampules… Some of them in two ampules… Another disadvantage is the observation of contamination. Other techniques (such as the agar plate method) are easier to identify any contamination. However, I would not expect any contaminations to occur if you work cleanly and with sterile equipment. The only unsolved question remaining is “how long can you store the yeast with this method”. From my experience, yeasts can be banked for two years at least. Without any maintenance work.

Fig 4: Part of Eureka Brewing’s yeast library

Another disadvantage of this method is storing yeast or yeast/bacteria mixtures. Storing mixtures with this method might change the ratio between the yeasts or yeasts and bacteria strains. On the other hand, there is no useful method in storing mixtures after all. Even if you manage to keep the ratio between the different strains constant during the storage, during the reanimation the ratios might change again… If you want to store mixtures, the only way would be to first determine the ratio of the different strains present and then separate them and bank them separately as well. Then create the mixture from the banked cultures again. Very labor intensive and not easy (been there). This even goes way beyond the topic of this post.

To summarize, banking yeasts in sterile solutions is a rather easy and cheap method. Not only is it less labor intensive than banking with agar plates but less expensive as well. From my point of view, this is a method where you can bank yourself yeasts for at least two years in a rather easy way and rather low in maintenance work. If I have to recommend a banking technique, I would recommend banking yeasts with sterile isotonic sodium chloride solutions. In addition, you can even easily trade your yeasts with other homebrewers.

The next post will be about banking yeasts with agar slants. Stay tuned!