Let me say up front that I DO NOT expect this to work well. However, if it does, culturing yeast will be a piece of cake for the future. Based on what I have read in the following two articles, it seems that I should be able to culture my own yeast, thus saving $30 or so per year for the yeast for my balsamic. I know, $30 is a minimal amount of money overall, but when you think about the total cost as a percent of the amount of balsamic you are making over several years of time, it is significant.
I am planning on taking two samples of 200ml of fermenting mosto right now and stick them both in the refrigerator. I will add 100 ml of fresh sterilized grape juice in 4 months, then 100 ml more in 4 more months, then in four more months I will pitch the yeast into a fresh batch of mosto. The idea of adding 100ml of juice is to keep the yeast as active as possible in the refrigerator and to stimulate growth to replace death from refrigeration.
If it doesn’t work, next year I will invest in a few tubes, some agar, and make a must/agar solution and transfer some yeast from the slant I buy that year to the new slants. Refrigerate and use the following year or two. Actually, I could buy one new yeast slant plus 4 or so unpopulated slants, transfer 1/4 of yeast to each of the other four slants and use them over 3 or 4 years and start over.
It has to be this way.
I found this too:
(Beer makers seem to be way out ahead of us balsamic makers on this issue. And where are the winemakers?!)
HOMEMADE STARTER TUBES, PLATES, AND SLANTS
If you culture yeast frequently, you will probably want to prepare your own starter tubes. Starter tubes are fairly easy to prepare, but slants and plates are more challenging (6). Capped glass tubes for starters and slants are available from scientific supply houses. You may be able to reuse empties from prepared commercial starter tubes.
After boiling starter wort for 10 min, let it cool and decant the clear wort. Wort can be injected into tubes with a clean plastic syringe. Steam the tubes in a pot of boiling water for 20 min. Cool, and make sure the caps are on tight before storage. Some sediment in the sterilized, cooled tubes is normal. Place these blanks in a sandwich bag and refrigerate. The starter solutions should remain clear over time. Any haziness or turbidity throughout the solution is a sign of contamination.
The wort can also be thickened with agar for use with slants and plates (2-3% [w/w] is common, but the ratio can vary widely with brand). Wide-bore syringes are useful for injecting the more viscous wort-agar mix into tubes. Slants must be steamed and then cooled at an angle to generate the necessary surface configuration. Store the same as the starter tubes.
If you have never worked with these items before, it would be good to start with the commercially prepared versions so that you have fewer variables to contend with.
Yeast is expensive – aside from the grapes themselves, no other annual expenditure is more. Its about $30 per year for a slant from my source. It would be great to culture your own yeast to both avoid this cost and to assure timely availability of the yeast.
I found a great article on the subject here: http://www.maltosefalcons.com/tech/MB_Raines_Guide_to_Yeast_Culturing.php
(coincidentally, it appears that it is on the site of the beer club that operates out of the same store that I get my winemaking supplies from www.homebeerwinecheese.com)
The process is described for home beer makers, but I suspect the process for balsamic vinegar makers (all two of us) is the same.
I am copying the important portion of it here for future reference in case the link above disappears:
Tips for handling yeast cultures
and starters at home
When I first started brewing I was amazed that very few people were culturing their own yeast. The main reason seemed to be that most homebrewers thought that it was necessary to have a glove box to successfully propagate yeast at home. I knew full well that this was a misconception since I had been culturing micro-organisms on the benchtop of research laboratories for years. This also seems to be common practice in brewery laboratories. One of the biggest misconceptions regarding microbiology is that bacteria can “crawl” into containers and infect it. Airborne bacteria do not have wings or legs. They get transported about by dust particles. The classic experiment illustrating this point was done by Louis Pasteur. He designed a gooseneck flask to be open at one end and contain sterile broth at the other. Although air could pass freely into the tube, dust particles became trapped in the curved portion. At no time have micro-organisms grown in this sterile broth. I’ve heard that this flask is still on display in the Pasteur Institute in France! This is also
Figure 6. Pasteur’s gooseneck flask. Only air and not dust particles can reach the broth. Therefore it remains sterile.
why plates can be left unsealed and stay sterile or why it is sufficient to cover sterile glassware with a sheet of heavy duty aluminum foil. In all of these cases it is nearly impossible for dust to get into the sample. Below are a few tips which can help create a bacteria-poor environment at home.
- Work in a draft-free clean area.All manipulations should be performed within a 2-4 square foot area. I’ve used a desk, coffee table, or dining table. The kitchen is not recommended since there tends to be a higher concentration of bacteria (especially lactobacillus) in that area. Clean the working area and your hands. The new antibacterial soaps on the market would be good for this. Sanitize the area and your hands by spraying them with 70% rubbing alcohol or ethanol before starting.
- Perform all manipulations near a flame source.Good flame sources include propane torches or alcohol lamps (we don’t recommend using your stove since the flame source is too low to effectively perform your manipulations). The flame and heat from the torch keeps all the bacteria-containing dust particles well above your working space and out of your yeast and wort. Flaming the mouth of all jars, bottles, and fermenters will also help get rid of air-borne or surface contaminants. Flaming is a standard microbiological practice, and although it may not be necessary, many consider it as a prayer to the sterility gods.
- Keep containers open for as little time as possible.It is best to loosen all caps and rehearse exactly what is going to transpire before actually doing it. For example if you are going to transfer some yeast from one tube into another. First loosen the cap on the tube of yeast, then the cap on the broth. Both should be situated in a rack or area for easy access. Then figure out exactly what hand you are going to hold each tube in and how exactly you are going to make the transfer. Are you going to put the cap down or hold it with your little finger of the piping hand? Are both tubes going to be held in the same hand or is one going to be left in the rack? If you cannot handle, (or are not coordinated enough to hold) the caps while you are opening and closing the various tubes and vessels, it is okay to put the caps down on the table. Be sure to place them with the sterile side facing down and if possible, place them on a paper towel wetted with alcohol. (Special precautions however, should be taken to keep the flame source away from any alcohol since it could ignite). Know exactly what you want to do before starting. This will lead to smooth easy transfers. The more you do this the more adept you will get at it. In all cases it is necessary to avoid touching any part of the pipet to the outside of the tubes or the surface of the bench. Transfers should be done quickly but not rushed so as to drop things or make mistakes.
Methods of Yeast Maintenance
Maintaining and storing your own yeast stocks is both convenient and cost-effective. Three major things must be considered when choosing a method of yeast storage. These are yeast strain purity, viability and genetic stability. Each of these differ depending on the method of preservation. The one most suitable for homebrewers is somewhat controversial. Each method has its own advantages and disadvantages and depends on personal preference as well as access to specialized equipment.
Media preparation. It is important to point out that the media used for long-term storage should be sterile. That means all micro-organisms including spores are destroyed. This can be done by heating in an autoclave or pressure cooker for 15-30 minutes at 15 psi. If this equipment is not available the media can be sterilized by tyndallization. This is done by boiling the media for 15 minutes every other day for a week. Note that this is similar to canning where the media is immersed in a pot of boiling water and boiled. At least two to three successive boilings are necessary for complete sterilization. Propagation media such as that used for starters need not be sterilized but has to be sanitized. In this case it should be boiled for at least 15 minutes and used within two or three days of preparation. Propagation media which is stored for any extensive length of time should be sterilized by one of the methods described above.
Master stocks. In general, it is a good idea to keep two stock preparations of yeast; one which is referred to as a working stock and the other, a master stock. The working stock is for routine use such as initiation of yeast propagation. The master stock is used to preserve the integrity of the original yeast strain. It is only used to replace the working stock or to propagate new master stocks. New master stocks are prepared when viability of the current master stock may be diminished. When this needs to be done depends on the yeast strain and the method of storage.
Liquid Media. This is a common method of storage for homebrewers and has also been referred to as yeast ranching or parallel yeast culturing. The best media for this method is wort or wort-containing media. Yeast is inoculated into 10 – 20 ml of media and grown until it reaches the stationary phase of growth (approximately 3 days) then stored in the refrigerator as cold as possible (40 °F). That means don’t keep it on the door. Stocks should be made in duplicate; one to use for brewing, the other as a stock. Some homebrewers prefer to build the 10 ml culture upto a larger volume and then dispense it into 12 oz. bottles. Storage in culture tubes or small jars also works fine. If stored properly, these cultures are stable for up to 6 months and then must be recultured (preferably from the untouched master stock). There are reports that storage in 10% sucrose after growth in wort can increase the shelf-life of yeast to as long as 2 years. In this case, it seems to be necessary to remove all residual nutrients or wort since direct addition of sucrose to the stationary yeast leads to continued fermentation even at 40 °F. Other bona-fide non-fermentable sugars such as lactose or glycerol may be more suitable but have yet to be tested for improving yeast’s shelf-life. Yeast strains vary in their sensitivity to storage in liquid wort. In general, only a small percentage of the cells survive storage. Therefore, it may be necessary to store in volumes larger than 10 ml especially if longer storage periods are used. Culturing in wort has been extensively characterized by the National Collection of Yeast Cultures (NCYC). They have cultured yeast for periods of up to 60 years and find that the mutation rate can be high. Of 600 strains studied as many as 50% with specific nutritional markers had lost at least some of their specific markers after culturing for 10-25 years (that’s after 20-50 passages). This was for all types of yeast strains including brewing yeasts. 10% of the 300 brewing yeast strains tested showed changes in flocculation behavior after 10 years or 20 passages. Thus storage in liquid media is feasible, but it is not the method of choice for long-term storage since it can undergo considerable genetic drift from the original stock. It is not clear whether minimizing the number of passages will also reduce the overall mutation rate.
Solid media (agar). The standard method for maintaining yeast and bacteria is on some type of solid media either in the form of plates or slants. Agar is typically used as a solidifying agent and is added at a concentration of 1.5-2% (1.5-2.0 grams per 100 ml liquid). The base media can be wort or one of the laboratory media described above. Agar is insoluble in wort or media and needs to be boiled for a few minutes to dissolve. After pouring plates or slants it is important that they be sufficiently dried at room temperature (2-5 days) before using them. Otherwise condensation may form on the sides of the tube or petri plate during storage which can lead to contamination especially by mold and fungus.
Agar Plates– Agar plates are made by pouring a sterile agar solution into pre-sterilized glass petri dishes or disposable petri dishes. Once solidified and air dried, yeast can be applied to the plate. This is done with a sterile inoculating loop. A small amount of yeast is added to the plate at one end and then spread across the plate. If performed properly, the yeast will be diluted to a point where a single yeast cell will be deposited. After growing for 3 – 5 days that single yeast cell will develop into a small round white mound of cells on the plate. This growth is referred to as a colony or clone. Colonies originating from single cells are round since they grow and expand outwards from the center. They do not have arms and legs and cannot move around on the plate. So nice distinct round colonies on a plate represent one yeast cell from the original yeast and should be free of bacteria and other contaminants. Bacteria and other contaminants may exist in other areas of the plate depending on the quality of the yeast used. Streaking on plates is classically used to purify yeast away from contaminants. In this case, a single colony is removed with an sterile inoculating loop and transferred to a fresh plate, slant, or tube of liquid media.
There are a number of procedures used to for spreading or streaking yeast. The so-called quadrant technique consistently produces single colonies and is depicted in Figure 4. This method involves dragging lightly (like writing with a pencil) an inoculating loop containing yeast over one quarter of the plate (quadrant) using a back and forth motion. The inoculating loop is then resterilized, cooled and used to transfer a small amount of the spread sample into the next quadrant. Again this diluted sample is then spread back and forth in the upper area of the quadrant. The inoculating loop is resterilized, cooled and used to transfer some diluted sample from the second quadrant into the third quadrant. After spreading over the third quadrant, the inoculating loop is resterilized and cooled, then used to transfer diluted sample from the third quadrant into the fourth and last quadrant. This sample is spread back and forth and should fill in any remaining unstreaked area on the plate. By sterilizing the loop between each transfer to a new quadrant, a dilution is made such that finally a single yeast will be deposited on the plate. After incubation at room temperature for 3-5 days, these single cells will divide many times and finally form a dull white mass of growth on the agar surface. Streaked plates are incubated inverted (agar side up) since condensation can form and drip onto plate and disperse the colonies. Plates are sealed with electrical tape, saran wrap, or parafilm and store inverted in the refrigerator.
|Figure 7. Quadrant streak method for applying yeast to an agar plate. Schematic diagram showing the 4 areas or quadrant over which yeast is applied and the approximate number of back and forth streaks to be used in each quadrant. Note that the inoculating loop is sterilized between streaking of each quadrant and therefore should dilute the yeast such that a single yeast is applied.|
The main advantage of agar plates allow separation of yeast from possible contaminants. Also the large working area is readily visible and easily accessed. The disadvantage is that its accessibility makes it less reliable. Not only is it much more susceptible to mold contamination, condensation can also be a problem and if it isn’t, then the plates may dry out. The larger surface for air (oxygen) exposure appears to diminish the shelf life of the yeast as well. There have been claims of yeast being stored on plates for a year or longer, but usually they are only stable for a few months. The shelf-life may actually vary depending on whether glass or plastic plates are used, how dry they are, and how well they are sealed. Many homebrewers prefer agar plates over slants, yet plates are not even considered as a method of storage by professionals. Because of the reliability factor, I prefer to use plates primarily for purification of single yeast colonies and not for yeast maintenance.
Agar Slants– Agar slants or slopes are made usually by adding a molten agar solution to glass culture tubes, sterilizing them, then allowing them to solidify at a 30°-45° angle. Yeast is applied with an inoculation loop and is streaked back and forth from the bottom up. The culture tube is incubated upright at room temperature (3-5 days) with the caps turned back one-half turn. Once a nice lawn of yeast is present the tubes are tightly capped and stored in the refrigerator.
Unlike plates, slants tend to be less susceptible to mold contamination and to drying out, and therefore are more reliable. They are slightly more difficult to work with since you have to deal with caps and you can’t see the yeast as well. The major advantage is that yeast can readily be maintained on slants for 1-2 years. However, reculturing every year is recommended. The quality of the tube and the how well a seal the cap forms may be a determining factor in the shelf-life of yeast on agar slant. A sterile overlay of mineral oil has been reported to extend the shelf-life of a slant by up to a year but this is can be messy. Presumably the oil overlay helps keep the air out and prevents the yeast from oxidizing.
Stabs– Stabs are upright tubes or bottles of semi-solid agar media. They contain only 0.7-1.0% agar. These are best prepared in screw-cap container with rubber inserts in the caps such as bijou, McCartney or Universal bottles but standard culture tubes may also be used. Yeast is applied with an inoculation loop and is inserted or stabbed into the agar all the way to the bottom of the tube or bottle. The stab is incubated and stored similar to a slant. Stabs are a common method used for long term storage of bacteria. Like plates, I have not seen any reference to this technique for yeast. The main advantage of this procedure is that it minimizes exposure to air which appears to be the primary limiting factor for shelf-life and stability. The disadvantage is that they are difficult to work with since the yeast is embedded in the agar and difficult to see. Also a fair amount of agar is usually picked up during transfer. It is unclear what the shelf-life is for stabs but my guess is that it would be at least 2-4 years (using tight-sealing bottles). Stabs therefore are a good for long-term storage, but not appropriate for routine propagation.
Dried yeast- Yeast is spotted onto a 1 inch square of sterile filter paper (Whatman 3MM) or a thick paper towel; wrapped in foil and dried (by desiccation) in the refrigerator for 2-3 weeks. Yeast can be spotted in growth media but it is better to add yeast that is suspended in condensed skim milk! Just let the yeast settle out of suspension, decant off the majority of liquid and resuspend in a small amount of evaporated skim milk from the grocery store. Stored the dried yeast in an envelope in the refrigerator. Although yeast maintained by this method is stable for 3-6 years, their manipulation during drying and resuscitation makes them more susceptible to contamination. Dried yeast can be resuscitated by placing in liquid media or on a plate. In general it should be streaked out on plates prior to use. This method is commonly used for genetic strains of yeast. I have successfully used this technique on brewing yeast, although the shelf-life and stability has yet to be determined. Supposedly the Siebel Institute is exploring this issue. In any case it is a great way to send yeast around world.
With regard to the dry yeast packets available at most homebrew shops, not all brewing yeast are amenable to the propagation and drying procedures used in the dry yeast industry. These yeast are typically grown to mass quantities in a dextrose/molasses mixture supplemented with nutrients at high temperatures (85°F). These adverse conditions may induce mutation or alter their performance in wort.
Freezing– Yeast can be frozen if a cryoprotectant is added. Glycerin and sucrose are commonly used and should be added to exponentially growing yeast at a final concentration of 5-15% (5-15 g/100 ml). Yeast stored at ultra cold temperatures (in liquid nitrogen or -112 °F) are stable for almost indefinitely (at least over 5 years) with over 99% of the cells surviving freezing. Freezing at higher temperatures (6.4 °F) yields shorter shelf-lives and less viability. It is important that once you freeze your yeast that, it does not thaw. This requires a really good quality non-frostfree freezer which maintains temperatures at or near 6.4 °F. Some homebrewers place their tubes of in is some denatured alcohol which has a supercooling effect and helps stabilize the temperature. Others imbed the tubes in ice. If you’re going to use this method freeze small aliquots (1-5 ml), then just thaw your yeastsicle and pitch it into a starter.
Homebrewers are faced with a variety of options on maintaining their yeast (summarized in Table below). The method of choice depends solely on the needs of the individual and their equipment. We are fortunate that there is an ever increasing number of inexpensive commercial sources of yeast so long-term storage by the homebrewer is not the necessity it once was. No matter what source of yeast or how it is stored, further propagation along with adequate aeration and fermentation at the correct temperature are sure to improve the quality of the beers you make at home.
Table 6. Summary of methods for yeast storage.
|Liquid media||0.5||Convenient but low viability and stability, questionable purity|
|Agar plate||0.2-1||Pure cultures but unreliable shelf-lives.|
|Agar slant||1-2||Easy, reliable, but moderate shelf-life|
|Agar stab||2-4||Easy, reliable, good shelf-life, but messy.|
|Dried||3-6||Inconvenient, requires purification.|
|Frozen||>5||Need special freezer or liquid nitrogen|