Building a home QC lab... questions

[butterpants]

Text books/Lab manual recommendation?

Brewing Microbiology by Fregus Priest was mentioned a few places I looked. Amazon has a 2013 softcover edition for 75$.

I'm the type of person who easily reads a text then can go put it into practice, so for me a lab manual is just in the cards...

 

[helibrewer]

Text books/Lab manual recommendation?

Brewing Microbiology by Fregus Priest was mentioned a few places I looked. Amazon has a 2013 softcover edition for 75$.

I'm the type of person who easily reads a text then can go put it into practice, so for me a lab manual is just in the cards...

Once you get yourself organized, write yourself some simple protocols to use everytime you do a particular task, including the aseptic techniques to use. For the hemacytometer make sure to note your counting techniques, i.e. which 2 sides of a square you count cells that are touching the line or not. We used the center section (25 small squares) and counted each corner square and the center square ignoring cells that touched the left and top lines, then averaged those 5 sections, which is why written protocols help even if it is just you doing the work.

 

[trentm]

First off guys, thanks for making this thread not only informative but a lively discussion as well. I was afraid initially there would be little interest. I'm glad that wasn't the case. I've found literally every post to be helpful, which is crazy because that never happens.

Ok so I'm actually buying stuff now! I've got a stupid long laundry list of items i'm thinking of. Hopefully you guys can help me narrow it down a bit....

The microscope was bought. Bausch&Lomb galen 3. Ebay 300$. I just couldn't pull the trigger on bargain optics even if the price point was appealing. I spent a lot of time behind a bolt gun and understand the value of glass. Doesn't have phase contrast (for now) which is a bummer but I can add that later.

Autoclave purchased as well.....American Sterlizer, again ebay 300$. 25quarts I believe and it's the model with its own heat source so you just plug it in. I can't wait to cram my Therminator in there.....soooo dirty

Hemacytometer and pipettes listed earlier from Cynmar. I already have an Acculab VIC123 so precision weighing is not a problem.

For the questions....

1) Are you guys using an incubator for plate growth? If not room temp is ok?

2) Thinking of getting a better pH meter while Cynmar is in my sights. The pen style I have from Hana keeps going tits up. Thoughts?

3) For staining/fixing are their certain slides that are preferred? What thickness of coverslips to get?

4) How are you cleaning your optics? Alcohol and lens paper?

5) Which housing tubes should I use for slants? How long as you guys getting away with freezing them at household fridge temps?

6) Glassware.....between borosilicate bottles for keeping sterile wort, beakers for mixing agar, flasks for propagation and graduated cyls what count and sizes are mandatory or need multiple of? Stoppers as well or only tinfoil? The biggest batches I ferment are 12 gallons.

7) Thinking of a stirplate with a heating element. Anyone doing this?

8) Do they make Bunsen burners that will run off compressed camp fuel bottles or even larger propane tanks? I really don't like alcohol lamps (if you've ever had a crazy ******* throw a Molotov cocktail at you, you'll know why).

9) Methylene Blue and gram stain kit....any other stains needed?

10) For the guys pouring their own plates.....into plastic or glass? I was initally thinking of just buying prepoured but the shelf life worries me. Think i'd rather have a bag of WLN/WLD and just be able to make them as I go since I have the autoclave. I'm assuming (hoping) the dry media has an extended shelf life.

That's it for now

Looks like you got a great deal on the microscope and sterilizer. I'll give my opinion on some of your questions:

1) For yeast culturing I use one of my temp controlled chest freezer's when possible. It just speeds things up a bit as ale yeast really like temps ~80F. However they will do fine at room temp; I try to find a place where the temp is consistent. It seems you are interested in using selective media to identify/isolate bacteria. If so, a controlled temp device becomes more important. It's fairly easy build your own "hot box", if you have a working environment that stays below your desired culturing temp as keeping the temp up is all you have to deal with.

2) A. J. response +1

3) For viability staining it's common to do this with your hemocytometer as you count you can tally the live and dead cells a derive a percentage. Fixing? I don't do this at home as often the chemicals required are very dangerous to work with and I haven't had a purpose, yet. That said, for observing fixed specimens, I think your standard slide and cover slip are adequate. I am curious about what specimens and why you are interested in fixing.

4) Could be a mistake, but I just wipe frequently with a soft cloth like one that comes with a new pair of eyeglasses, I think it's microfiber. Kimwipes with alcohol if necessary. Lens paper is probably better. And I keep the cover on my scope at all times it is not in use.

5) I use 15ml disposable centrifuge tubes (aka Falcon Tubes). This may not be the best choice for longevity of your culture because I suspect the the plastic may allow some air to enter the tube and air is one of my biggest concerns. I reslant every 6 months and my cultures have always been viable at that time (have gone as long a 9 months). And I have maintained the my first isolate in good condition since 2008 (although I have lost a couple along the way). The reasons I choose Falcon tubes is for their versatility, low cost, and ease of use. I also use them for; the first step of a yeast culture, isolating yeast from beer or wild yeast from fruit. Many are sold as sterile and some can be autoclaved. I have not tried freezing in a household freezer, although many have reported good results.

6) I like to use home canning jars for keeping sterile wort. they are relatively inexpensive, come in a variety of sizes, available at most grocery stores, and work great. For media, I use 500 ml "media bottles" available from Cynmar. While most instructions for preparing agar include boiling before autoclaving, the numerous labs I have been associated with skip this step and mix up the ingredients in a media bottle and autoclave. Certainly for wort agar and YPDA this process works fine, however, there may be some types of media that would benefit from pre boiling. If you choose media bottles, be sure to get the one with a pouring ring, the ring makes pouring much easier with less mess. BTW, 200ml of media will pour ~10, 100mm x 15mm plates. For a graduated cylinder, I have a 100ml and wish for a 500ml. I think tinfoil is the best for a flask closure for propagation. Your flask size will depend on your yeast yield and of course you batch size and gravity. Yeast strain may have an effect on yield. For my Belgian strains, in a well controlled environment (air and temp), I can produce ~300 million cells per ml. Keep in mind that lab produced yeast can be pitched at around half the recommended rate compared to re-pitched yeast. I use 400,000 to 750,000 cells/ml/degree plato for a final cell density as a rule of thumb and for my Belgian Strong Pales (~1.07 OG), I go with 600,000. So for my 10.5 gallon batches, I use ~1400ml culture. I believe I get better aeration of the culture by using 2, 2000ml flasks containing ~700ml each on a orbital shaker. I also believe this would be optimal on a stir plate as well. Aeration, in my opinion, is the key to healthy yeast and high yields.

7) I have often thought a heated stir plate would work well but not sure about temp control.

8) I get by with a wickless alcohol burner. Bunsen burners have definite advantages.

9) I only have experience with methylene blue. Works fine, but with lab produced yeast, it just reassures that the culture is >99% viable. If you are re-pitching, some type of viability stain would be valuable.

10) I have no experience with selective media. For wort agar and YPDA, I mix and pour my own using disposable petri dishes. These ingredients have a long shelf life and at 50 cents for a disposable dish and 14 cents per plate for agar, this is affordable. On the other hand, glass plates would allow you set the plates in you sterilizer and relieve the worry of contamination. Warthaug (aka Bryan) has a protocol for this.

 

[helibrewer]

Here is the WLN media:

And here is the WLD (selective) media:

You can see that the selective media does not grow S. cerevisiae so it is good if you are only looking for bugs.

We cleaned objective lenses with Windex and lens paper

 

[Warthaug]

1) Are you guys using an incubator for plate growth? If not room temp is ok?

I don't use an incubator. They can accelerate growth, but they can also alter gene expression in undesired ways.

3) For staining/fixing are their certain slides that are preferred? What thickness of coverslips to get?

Other have mentioned this already, but fixation is rarely needed. The only time I do anything like it is if I am trying to get a good count on yeast vs. bacterial numbers, or am trying to get a really good look at morphology. In these cases I rely on heat fixation, which is dead-simple:

1. Place a few drops of a liquid culture on a slide. If working from a plate, place a few drops of water on a slide, then mix in a colony of yeast using a sterile loop
2. Spread out the suspension until the middle 3rd or so over the slide is covered
3. Let sit until completely dry; don't rush it - letting it sit too long is harmless, not letting it sit long enough will ruin it
4. Once dry, very quickly pass the slide - yeast/bugs side down - through the flame of your burner. When I say quick I mean quick - less than a second. This direct exposure to flame will bind your sample to the slide; any longer and the sample gets cooked.
5. Stain/image/etc as you require

Obviously, you don't do that for viability staining

4) How are you cleaning your optics? Alcohol and lens paper?

I would recommend isopropyl and lens paper. If using an oil immersion lens, first dab off the oil before cleaning with alcohol. In either case, lightly whet the lens paper with alcohol and wipe with that - never pour/spray alcohol on the lens.

5) Which housing tubes should I use for slants? How long as you guys getting away with freezing them at household fridge temps?

I freeze my yeast, but not in a lab situation so the methods are not really comparable. When I used to slant I preferred the 10-15ml screw-top glass tubes; reusable, you can prep your slants by autoclaving the media right in the tube, and they are big enough to give you enough media volume to allow for long-term storage.

Yeast can be kept stably on slants for several years using a simple protocol:

1. Grow yeast on slant until there is a good layer of yeast
2. Seal the tube and place in the fridge for 2-3 days. This will induce the formation of trehalose, which will aid the yeast's long-term survival
3. Fill the tube with sterile mineral oil, and re-seal the top *(e.g. with vinyl electrical tape). There are reports of yeasts stored like this being recovered after 60+ years - although most people would recommend re-culturing every 3-5 years.

Obviously, if you're using the yeast a lot the above is not a preferred method. You can also overlay a slant with sterile water; yeast last about a year like that, or you can tightly seal the caps - good for 3-6 months. There are lots of other yeast-storage options other than slants as well.

6) Glassware.....between borosilicate bottles for keeping sterile wort, beakers for mixing agar, flasks for propagation and graduated cyls what count and sizes are mandatory or need multiple of? Stoppers as well or only tinfoil? The biggest batches I ferment are 12 gallons.

For 12 gal batches you'll probably want a 1, 2 and 4L flask for starters. Sterile wort can be kept in plain-ol' mason jars (can in a pressure cooker like any other preserve). Mason jars also make great beakers for mixing media - no need to buy the expensive lab stuff. I use foil, but there are foam stoppers that are a little more convenient.

7) Thinking of a stirplate with a heating element. Anyone doing this?

Seems risky, in terms of overshooting temps and what not. In the lab we certainly would not do this.

8) Do they make Bunsen burners that will run off compressed camp fuel bottles or even larger propane tanks? I really don't like alcohol lamps (if you've ever had a crazy ******* throw a Molotov cocktail at you, you'll know why).

I have seen propane models that run off of the little compressed tanks, but they seem to be few and far between.

9) Methylene Blue and gram stain kit....any other stains needed?

I've found viability dyes to not be overly useful; because they rely on enzymatic action dead cells can be missed to to residual activity. I've had a few cases where cultures that stained with many viable cells were unable to grow.

10) For the guys pouring their own plates.....into plastic or glass? I was initally thinking of just buying prepoured but the shelf life worries me. Think i'd rather have a bag of WLN/WLD and just be able to make them as I go since I have the autoclave. I'm assuming (hoping) the dry media has an extended shelf life.

I use both, but prefer glass. Glass can be reused, making it cheaper, and you can prepare media right in the plate (no need for sterile pours). Dry media will last years on the shelf; especially if you seal the containers tight enough to limit water entry.

Bryan

 

[orangehero]

How about one of these?

http://www.amazon.com/dp/B004U8CP88/?tag=skimlinks_replacement-20

The piezo is junk but it's actually a really nice burner for less than $6.

 

[butterpants]

^^Totally forgot about a light camp stove. I own a jetboil too....Good call, it's perfect.

Oh wow lots of good information here. Thanks guys for tracking with my questions! I was interested in fixing slides only for showing others (like my kid). For the stirplate/hotplate combo my thoughts were on heating mixtures of things while actively stirring not in any shape or form of incubation.

Ordered a pile of stuff. I just made ebay some fat cash this week. In a few weeks I should have my basement workspace converted and culturing. Exciting. I have more questions....

1) When you're doing cell counts for density and vitality what dilution range do they have to be in for quick counts to be accurate as possible? For instance lets say I have a 1.8L starter and I decant it down to 200mL. I take 1mL of the starter slurry and add it to 99mL of sterile water to get a 1:100 dilution. Will this be sufficient density of cells to count or in your practice would another, or multiple sets of serial dilutions be made? When is best to add viability dye during this procedure? How are you accounting for the added volume of dye?

2) Is there an inexpensive yet feasible (don't want to burn down the house) alternative to anaerobic "packs" for fermenting your plates without the presence of oxygen?

3) I've searched yet haven't been able to find a good deal on a small laminar flow hood, so I got to thinking what if I construct a work space out of laminated countertop with a partial plexiglass shield that contains 2 sources of light...a fluorescent bulb AND a UV bulb? I could work within the enclosed space with the UV lamp on and a small flame keeping the wild bug population very low. It could be left on for extended periods to sterilize all interior surfaces and no dealing with filters/air flow calculations. The downside would be personal exposure to the radiation so gloves and long sleeves would need to be worn but the plexiglass (Lexan, Home Depot...cheep) would filter out almost all of the UV escaping the pseudo fume hood. I'm thinking you might need some sort of swan neck exhaust port just for the hot gasses from the bunsen burner.... thoughts on any of this? I'll attach a couple of pictures I found on google...just imagine them without any air filtration, only UV sterilization.

4) So really all I need to test, select and isolate for lacto, pedio, brett + sacro would be WLN vs WLD (+ ana vs aerobic) while observing colony morphology and then a growth media such as UBA to propagate once isolated? How many plates (100mmx15) do you get from a 500gr bottle of dry media?

5) If you have large glassware and need to sterilize it in your oven BUT it's borosilicate glass, can you crank up the temp and do it faster than the 250F/2 hours? I'm thinking about a 5L flask or odd shaped things. Just wondering

6) Any recommended apps/spreadsheets for making cell counts and the calculations after for pitch rate easier? Found a few in the Play store. Just looking for an endorsement or warning to avoid. Couldn't find the one mentioned earlier in this thread.

7) Can you autoclave Bell mason jars with their lids on and loose? If not do you just take the lids completely off?

8) So if I'm mixing up lets say WLD and WLD agar. I put it in a mixing vessel then transfer the liquid to media bottles. Lightly cap and autoclave. I can wait for it to cool a bit then pour plates or I can just park the bottle on the shelf a few weeks....but what is the preferred method of reheating the mixture for pouring? Is there a limited number of times it should be heated and cooled before performance degrades?

9) For DI water are you guys just using distilled bottled water from a grocery store in some kind of lab friendly dispenser or sterilizing it first?

 

[helibrewer]

I found 1:1000 dilutions worked best for counting. I don't do viability on a hemacytometer, the stain will eventually effect your hemacytometer.

One easy way to do anerobic growth is to use slants and stab into the media, anerobic stuff will grow in the "stabbed" area within the media.

Do not work under a powered UV light, use the light before you start work then turn it off and use standard aseptic techniques.

For the autoclaved media you want to pour your plates when the media is "can touch your cheek" warm...just don't actually touch your check, that WLD is toxic.

For Mason jars, place the lid on them then the ring, but tighten the ring only enough to hold the lid in place. A well sealed lid after cooling indicates that you have a good sterile sample.

Sterile water and distilled water are different. For DI, just use store bought.

You don't have to be too fancy with your flame source, just something that can flame your loop and create a slight air current away from your working area.

Let your prepared plates sit for several days (media side up) before using them. It's much better to know if they are contaminated before you spend all the time streaking them.

 

[Warthaug]

1) When you're doing cell counts for density and vitality what dilution range do they have to be in for quick counts to be accurate as possible?

The answer is "it depends" - the dilution you need is proportional to the yeast density, which can be quite variable. 1:100 often works for fermenting wort (e.g. in a starter or a fermenter), 1:1000 to 1:10,000 is more typical for slurry or anything with a lot of bacteria in it.

As a "rule", for your yeast counts to be accurate you need to be counting 100-300 cells on the hemocytomoeter, at your selected dilution. Less than that and counts can be off due to sample variability, more than that is hard to count.

2) Is there an inexpensive yet feasible (don't want to burn down the house) alternative to anaerobic "packs" for fermenting your plates without the presence of oxygen?

Yep, an overlay plate. Its pretty simple:

1. Prepare a plate as you normally would, but with ~1/2 the normal amount of media
2. Sterilize more medium, and cool to just above its gelling point (40C or so). You want an excess amount of this
3. Streak out your sample on the first plate, than immediately overlay with the cooled (but not yet solidified) agar. Fill the plate completely & cap while still liquid (a bit of gel should leak around the edges)
4. The gel should set quickly, once set seal the edges of the plate with vinyl tape (electricians tape)

This is not completely impervious to air, but is sufficient for all but the strictest of anerobes - anything that would grow in anaerobic-phase beer would do fine in this plate.

Harvesting bugs from these plates is a bit more challenging; generally you recover colonies by using a stab or punch.

3) I've searched yet haven't been able to find a good deal on a small laminar flow hood

Don't waste your money. Laminar flow hoods are generally not used for microbiology applications - standard aseptic techniques are more than sufficient. In essence, they are a crutch for poor technique (or, in the case of pharmaceutical applications, a guard against mistakes). The only other thing to use them for is to protect yourself from pathogens, and the design you've picked wouldn't do that - it would blow them in your face.

And, FWIW, UV sanitation is pretty weak; a few seconds exposure to starsan or bleach has a grater effect than several minutes of UV.

4) So really all I need to test, select and isolate for lacto, pedio, brett + sacro would be WLN vs WLD (+ ana vs aerobic) while observing colony morphology and then a growth media such as UBA to propagate once isolated? How many plates (100mmx15) do you get from a 500gr bottle of dry media?

You don't need anaerobic for any of those - both pedio and lacto are areotolerant anaerobes, which means they don't need oxygen but also don't give a s**t if its around. Depending on the species, the impact of oxygen on their growth will range from a mild growth impediment to a mild growth enhancement. I culture lacto and pedio all the time without any anaerobic media or chambers without issue. The standard for commercial preparation of both of those organisms (e.g. for yoghurt, sausage, probtiotic pills, etc) is non-aerated, but otherwise air-exposed fermenters.

The medium you choose is upto you. Commercial media are expensive, but can simplify work-flow through the inclusion of selective agents. Beer-wort and potato-dextrose agar are cheap, but either require colony identification or the addition of selective or differential agents such as antibiotics and pH indicators to achieve the same thing as commercial media.

In terms of volumes, the answer is "it depends". A typical 100 mm plate should contain ~30 ml of media. How far a bottle goes depends on the amount of medium required to make the needed volume of gel, and how much you waste (some waste is inevitable).

5) If you have large glassware and need to sterilize it in your oven BUT it's borosilicate glass, can you crank up the temp and do it faster than the 250F/2 hours? I'm thinking about a 5L flask or odd shaped things. Just wondering

Yep - sterilization time is proportional to temperature:
http://www.cdc.gov/hicpac/disinfection_sterilization/13_10othersterilizationmethods.html

6) Any recommended apps/spreadsheets for making cell counts and the calculations after for pitch rate easier? Found a few in the Play store. Just looking for an endorsement or warning to avoid. Couldn't find the one mentioned earlier in this thread.

I like yeastcalc.co - free, and is accurate. Its a re-write of the former yeastcalculator page.

Cell counts don't really need an app - (# cells counted per quadrant) * dilution * hemocytometer conversion factor (usually 10,000). With a bit of use you'll probably just do the math in your head.

7) Can you autoclave Bell mason jars with their lids on and loose? If not do you just take the lids completely off?

Lids on & loose work just fine.

8) So if I'm mixing up lets say WLD and WLD agar. I put it in a mixing vessel then transfer the liquid to media bottles. Lightly cap and autoclave.

Don't bother with the mixing vessel; weigh out the medium and put into the flask/jar; measure out the water and pour in. Swirl flask/bottle to mix, cap lightly (or with foil if using a flask) and autoclave. No need to make extra dishes

I can wait for it to cool a bit then pour plates or I can just park the bottle on the shelf a few weeks....but what is the preferred method of reheating the mixture for pouring? Is there a limited number of times it should be heated and cooled before performance degrades?

Repeated re-heats can degrade some of the medias (e.g. those containing antibiotics or dyes), but two or three reheats should be OK as you're sticking to basic medias. Simply place the bottle of media into a pot of water, and heat on your stove until melted.

9) For DI water are you guys just using distilled bottled water from a grocery store in some kind of lab friendly dispenser or sterilizing it first?

Don't bother with it for media - tap water is fine. Heating will drive off any chlorine, and the trace minerals in the water should not otherwise impact your media. Even in a research lab, we use tap water for most of our bacterial culture media. DI is for more sensitive cells - like them wussie mammalian cells.

Bryan

 

[ajdelange]

1) When you're doing cell counts for density and vitality what dilution range do they have to be in for quick counts to be accurate as possible?

The accuracy depends on the square root of the number of cells you count. The coefficient of variation is its reciprocal. Thus if you count 100 cells (total in all squares) your Cv = 1/sqrt(100) = 0.1 and the standard error in your count is Cv times what you counted or, in this example (0.1)(100) = 10 cells. Or just looking at the Cv expressed as a percentage 10%. If you count 200 cells your Cv= 1/sqrt(200) = 0.0707 or 7%. I would think either of those levels acceptable but if you want tighter accuracy you will have to count more cells - lots more. For 1000 counted Cv = 3.2%.

 

[butterpants]

Thread keeps getting better and better, thanks again guys.

100-200 cells counted and an accuracy range from 7-10%. AJ I can live with that, thanks. 1k no way. Diminishing returns on the loss of my mental sanity. Heli, if you're not using a hemocytometer to count stained cells, how are you determining viability....or is that just not a concern for you? Bryan I checked on the FDA site and it looks like 340F@60min will get my big pyrex items sterilized.

Some more questions/discussion points:

1) Watched a White Labs YouTube video not too long ago on making yeast starters (which I have been doing since I began brewing but I'd consider them an authority on yeast and was looking for refinement of technique)....it appeared to be around a 1.5-2L starter in a 4L Erlenmeyer flask with stirbar. Nice amount of room for gas exchange....BUT the presenter then took the aluminum foil cover on top of the flask and cranked it down sealing it all around....something I've never done, infact just the opposite. I leave it loose for gas exchange. Thoughts on oxygen/stirplate starters/oversized flasks/lid sealing? You guys are the yeast whisperers right? What say you

2) Do you recommend actually mixing up my own differential media from its raw individual components (WLD/WLN i'm thinking) instead of buying it in a dry premixed form? Significant cost savings? My worry is that some of the trace products (bromocresol green, cyclohexamide) I'll have to purchase in quantities I'll never use making it no longer financial advantageous.

3) Why would one use "beer" agar over "wort" agar....from a plating or growing perspective? If making wort agar is it important to obtain a hot break and rupture starch granules before adding the wort constituent or can you just weigh out DME?


Packages have started arriving

 

[ajdelange]

100-200 cells counted and an accuracy range from 7-10%. AJ I can live with that, thanks. 1k no way.

That reminds me. A very important part of the kit that hasn't been mentioned is one of those counter thingies that theater ticket takers use. Its a VR counter that increments once each time you push the little lever. You can count like the wind with one of those, never have to write anything down and don't have to remember anything except which squares you have counted.

 

[trentm]

I don't use an incubator. They can accelerate growth, but they can also alter gene expression in undesired ways.

I haven't heard of this, could you elaborate. Seems odd to me, isn't an incubator just a way of keeping a consistent temperature? How would the environment in a incubator at 75F be different from a room temperature environment at 75F?

 

[trentm]

Don't waste your money. Laminar flow hoods are generally not used for microbiology applications - standard aseptic techniques are more than sufficient. In essence, they are a crutch for poor technique (or, in the case of pharmaceutical applications, a guard against mistakes). The only other thing to use them for is to protect yourself from pathogens, and the design you've picked wouldn't do that - it would blow them in your face.

And, FWIW, UV sanitation is pretty weak; a few seconds exposure to starsan or bleach has a grater effect than several minutes of UV.

Bryan, I have great admiration for your microbiological knowledge but I am surprised by this statement. I am going to give an alternate view with all due respect.

My profession is Plant Pathology and as I am sure you know it is the study of plant disease. My area is very applied and I do a lot of work in the field and greenhouse, however, lab work is required in order to isolate and produce inoculum to make plants sick and to look at the effects of the disease. Over my 15 years in the field, I have worked in 7 different labs and collaborated with dozens of other labs. Often these labs are fairly primitive, base in field locations, others are at major Universities and USDA/ARS research stations. Without exception all Plant Pathology labs I have worked in or visited are equipped a laminar flow hood and use these hoods for all microbiological work when there is the possibility of contamination. Within my experience, in a lab situation, working with open cultures or media outside a hood is a thing of the past.

I do agree that basic work required for the advanced homebrewer can be achieved without a hood. I have had great success for over 5 years in my home (garage) lab without one. As for a crutch, nothing, in my opinion will forgive poor sterile technique including a hood. In Plant Path we work with highly contaminated plant material including roots and soil from field conditions. Isolating fungal or bacterial pathogens from these samples can be very challenging, and just as isolating yeast from a field sample a laminar flow hood is a great tool. Personally, I find many other uses for my hood, none of which include guarding against mistakes. I believe the $200 I spent on my hood was a great addition to my lab. On the other hand, spending $2000 - $10,000, the price of a new hood, would be extravagant.

+1 on the UV sterilization. Well equipped hoods do use UV light, but the idea is to use the light when the hood is not in use to maintain sterility within the hood environment between uses.

 

[butterpants]

That reminds me. A very important part of the kit that hasn't been mentioned is one of those counter thingies that theater ticket takers use. Its a VR counter that increments once each time you push the little lever. You can count like the wind with one of those, never have to write anything down and don't have to remember anything except which squares you have counted.

Yep I snagged one of those... just forgot to list it

 

[trentm]

^^Totally forgot about a light camp stove. I own a jetboil too....Good call, it's perfect.

Oh wow lots of good information here. Thanks guys for tracking with my questions! I was interested in fixing slides only for showing others (like my kid). For the stirplate/hotplate combo my thoughts were on heating mixtures of things while actively stirring not in any shape or form of incubation.

Ordered a pile of stuff. I just made ebay some fat cash this week. In a few weeks I should have my basement workspace converted and culturing. Exciting. I have more questions....

1) When you're doing cell counts for density and vitality what dilution range do they have to be in for quick counts to be accurate as possible? For instance lets say I have a 1.8L starter and I decant it down to 200mL. I take 1mL of the starter slurry and add it to 99mL of sterile water to get a 1:100 dilution. Will this be sufficient density of cells to count or in your practice would another, or multiple sets of serial dilutions be made? When is best to add viability dye during this procedure? How are you accounting for the added volume of dye?

2) Is there an inexpensive yet feasible (don't want to burn down the house) alternative to anaerobic "packs" for fermenting your plates without the presence of oxygen?

3) I've searched yet haven't been able to find a good deal on a small laminar flow hood, so I got to thinking what if I construct a work space out of laminated countertop with a partial plexiglass shield that contains 2 sources of light...a fluorescent bulb AND a UV bulb? I could work within the enclosed space with the UV lamp on and a small flame keeping the wild bug population very low. It could be left on for extended periods to sterilize all interior surfaces and no dealing with filters/air flow calculations. The downside would be personal exposure to the radiation so gloves and long sleeves would need to be worn but the plexiglass (Lexan, Home Depot...cheep) would filter out almost all of the UV escaping the pseudo fume hood. I'm thinking you might need some sort of swan neck exhaust port just for the hot gasses from the bunsen burner.... thoughts on any of this? I'll attach a couple of pictures I found on google...just imagine them without any air filtration, only UV sterilization.

4) So really all I need to test, select and isolate for lacto, pedio, brett + sacro would be WLN vs WLD (+ ana vs aerobic) while observing colony morphology and then a growth media such as UBA to propagate once isolated? How many plates (100mmx15) do you get from a 500gr bottle of dry media?

5) If you have large glassware and need to sterilize it in your oven BUT it's borosilicate glass, can you crank up the temp and do it faster than the 250F/2 hours? I'm thinking about a 5L flask or odd shaped things. Just wondering

6) Any recommended apps/spreadsheets for making cell counts and the calculations after for pitch rate easier? Found a few in the Play store. Just looking for an endorsement or warning to avoid. Couldn't find the one mentioned earlier in this thread.

7) Can you autoclave Bell mason jars with their lids on and loose? If not do you just take the lids completely off?

8) So if I'm mixing up lets say WLD and WLD agar. I put it in a mixing vessel then transfer the liquid to media bottles. Lightly cap and autoclave. I can wait for it to cool a bit then pour plates or I can just park the bottle on the shelf a few weeks....but what is the preferred method of reheating the mixture for pouring? Is there a limited number of times it should be heated and cooled before performance degrades?

9) For DI water are you guys just using distilled bottled water from a grocery store in some kind of lab friendly dispenser or sterilizing it first?

Sorry for the delayed reply. I'll comment on what has not been covered or where I think I have something to add.

1) You may find counting before you allow the yeast to settle and just after you remove the culture from the stir plate to be the most accurate. After the yeast have settled, the cells have clumped together and it is sometimes hard to get a homogeneous suspension. I make sure to keep up with the size of the culture by measuring and tallying each step. If you find you need to count the slurry, you may have to do serial dilutions until you find an appropriate concentration. It may be however somewhere between 1:100 and 1:1000.

For an un-concentrated culture containing ~300 million cell/ml a 20:1 dilution works well for me. At that concentration and dilution you will be counting ~60 cells/small square if you are counting the center square, for a total of ~300 cells.

You can you use the viability dye as your last diluent. I make my last dilution using 0.5 mL of 0.01% methylene blue to 0.5 mL of the sample. Remember to calculate the final dilution.

3) For me, a laminar flow hood is a great tool but not a requirement. The advantage to working in a hood is you have a sterile environment that allows you to work with open cultures without the concern of environmental contamination. Unless your environment is completely sterile, you will still need to work around a flame and pray you are in the updraft.

4) I have no experience with selective media. Of course the number of plates you can pour will depend on how much media is required. Some have said ~30 mL per plate. I shoot for 20 mL.

6) The “Hemocytap” app is only available from Apple. I mentioned earlier it was free but it is actually $1.99.

7) As mentioned, leave the rings loose but tighten them after you remove them from your sterilizer. The rings need to be tight to ensure the lids seal properly.

 

[trentm]

Thread keeps getting better and better, thanks again guys.

100-200 cells counted and an accuracy range from 7-10%. AJ I can live with that, thanks. 1k no way. Diminishing returns on the loss of my mental sanity. Heli, if you're not using a hemocytometer to count stained cells, how are you determining viability....or is that just not a concern for you? Bryan I checked on the FDA site and it looks like 340F@60min will get my big pyrex items sterilized.

Some more questions/discussion points:

1) Watched a White Labs YouTube video not too long ago on making yeast starters (which I have been doing since I began brewing but I'd consider them an authority on yeast and was looking for refinement of technique)....it appeared to be around a 1.5-2L starter in a 4L Erlenmeyer flask with stirbar. Nice amount of room for gas exchange....BUT the presenter then took the aluminum foil cover on top of the flask and cranked it down sealing it all around....something I've never done, infact just the opposite. I leave it loose for gas exchange. Thoughts on oxygen/stirplate starters/oversized flasks/lid sealing? You guys are the yeast whisperers right? What say you

2) Do you recommend actually mixing up my own differential media from its raw individual components (WLD/WLN i'm thinking) instead of buying it in a dry premixed form? Significant cost savings? My worry is that some of the trace products (bromocresol green, cyclohexamide) I'll have to purchase in quantities I'll never use making it no longer financial advantageous.

3) Why would one use "beer" agar over "wort" agar....from a plating or growing perspective? If making wort agar is it important to obtain a hot break and rupture starch granules before adding the wort constituent or can you just weigh out DME?


Packages have started arriving

1) I like the idea of a large headspace because it allows the stir plate to do an adequate job of stirring which allows for gas exchange and creates a large surface area. In my opinion the large head space is not so much for the air present in the space before the yeast start to grow, as the space will quickly become saturated with CO2 if sealed, but to allow more surface contact with the air you that can coax into the flask throughout the fermentation. Due to the conical shape of an E. flask, the surface area is diminished the fuller the flask.

A loose fitting foil cover is the best in my opinion because, as you know, microbe laden dust does not fall up but air can move in and CO2 out, due to action of the vortex. Foam or cotton plugs are another option and can keep out crawling insects but you may have more of a blow off issue if your system creates a large krausen.

2) Sorry, again no experience with differential media. My guess is buy the premixed form.

3) No knowledge on “Universal Beer Agar”. Here is a link to a fact sheet: http://www.neogen.com/Acumedia/pdf/ProdInfo/7574_PI.pdf. For culturing with the intention to brew, wort agar is said to be best because the yeast get used to the environment they are in. I noticed in the recipe that Glucose is the sugar source and yeast can get lazy on Glucose. For wort agar, I brew up a batch of wort for starters and media, remove as much break material as possible and can it. I like to make my own because I pitch the entire starter. Even though I have removed much of the break, after canning more break appears. When making solid media I decant the wort from the break because I like clear plates and slants. When using it for a starter, I add at least some of the break because I have heard that yeast benefit from small amounts of break material. I don't know about “rupturing starch granules”.

 

[Warthaug]

Bryan, I have great admiration for your microbiological knowledge but I am surprised by this statement. I am going to give an alternate view with all due respect.

My profession is Plant Pathology and as I am sure you know it is the study of plant disease. My area is very applied and I do a lot of work in the field and greenhouse, however, lab work is required in order to isolate and produce inoculum to make plants sick and to look at the effects of the disease. Over my 15 years in the field, I have worked in 7 different labs and collaborated with dozens of other labs. Often these labs are fairly primitive, base in field locations, others are at major Universities and USDA/ARS research stations. Without exception all Plant Pathology labs I have worked in or visited are equipped a laminar flow hood and use these hoods for all microbiological work when there is the possibility of contamination.

Outside of non-bacterial cell culture, in the health sciences side of things we don't use hoods. In fact, they are counter-indicated for BSL 1 and BSL 2 organisms (i.e. our permits explicitly recommend against using hoods for those organisms). Its not until we hit BSL 3 organisms where we use a hood - to protect ourselves, not our samples - and those hoods are dedicated strictly to that work.

Within my experience, in a lab situation, working with open cultures or media outside a hood is a thing of the past.

Than you plant biologists are weird :cross: I work in a large microbiology department, have visited microbiology departments across the world, been part owner of 3 of biotech startups, worked briefly in industry, and in none of those environments I have never seen a hood used strictly for bacterial/fungal work, outside of BLS3/4 organisms.

Bryan

 

[Warthaug]

I haven't heard of this, could you elaborate. Seems odd to me, isn't an incubator just a way of keeping a consistent temperature? How would the environment in a incubator at 75F be different from a room temperature environment at 75F?

Sorry, by incubator I thought you meant growing at 30-37C. Regardless, many genes are regulated by environmental factors such as temperature, osmolarity, pH, etc. Meaning that yeast cultured under conditions different from than of beer may express different genes, and thus produce different fermentation profiles, flavour compounds, viability, etc. I've done some experimentation with higher-temp starters to accelerate growth, and found (in general) that going more than 3-4 degrees above the recommended fermentation range produces yeast which ferment more slowly and throw more off-flavours than do yeast grown within the recommended fermentation range, despite the beer itself being in the recommended range.

Bryan

 

[Warthaug]

1) Watched a White Labs YouTube video not too long ago on making yeast starters (which I have been doing since I began brewing but I'd consider them an authority on yeast and was looking for refinement of technique)....it appeared to be around a 1.5-2L starter in a 4L Erlenmeyer flask with stirbar. Nice amount of room for gas exchange....BUT the presenter then took the aluminum foil cover on top of the flask and cranked it down sealing it all around....something I've never done, infact just the opposite. I leave it loose for gas exchange. Thoughts on oxygen/stirplate starters/oversized flasks/lid sealing? You guys are the yeast whisperers right? What say you

It doesn't matter all that much - the amount of O2 actually entering the starter is rather small (I think Kai did some experiments on this end). The main benefit of stirring is removal of CO2, which will have no trouble getting past a tightly "sealed" foil cap.

2) Do you recommend actually mixing up my own differential media from its raw individual components (WLD/WLN i'm thinking) instead of buying it in a dry premixed form? Significant cost savings? My worry is that some of the trace products (bromocresol green, cyclohexamide) I'll have to purchase in quantities I'll never use making it no longer financial advantageous.

If you're making enough on your own than it is far cheaper to make it from scratch, but if only using small amounts it may be cheaper to buy a small bottle of pre-mixed.

3) Why would one use "beer" agar over "wort" agar....from a plating or growing perspective? If making wort agar is it important to obtain a hot break and rupture starch granules before adding the wort constituent or can you just weigh out DME?

I don't think there is a difference; commercially it is usually sold as wort-agar, but I treat the terms as interchangeable. That said, beer-agar may indicate a hopped product, which would have a weak selectivity against gram positive bacteria. DME + water + nutrient is more than enough - conversion, hydration, etc, are already complete with DME (there is no starch in DME; its all converted). I take mine through the hot-break, simply because that is inevitable if boiling long enough to sanitize the wort.

Bryan

 

[trentm]

Outside of non-bacterial cell culture, in the health sciences side of things we don't use hoods. In fact, they are counter-indicated for BSL 1 and BSL 2 organisms (i.e. our permits explicitly recommend against using hoods for those organisms). Its not until we hit BSL 3 organisms where we use a hood - to protect ourselves, not our samples - and those hoods are dedicated strictly to that work.



Than you plant biologists are weird :cross: I work in a large microbiology department, have visited microbiology departments across the world, been part owner of 3 of biotech startups, worked briefly in industry, and in none of those environments I have never seen a hood used strictly for bacterial/fungal work, outside of BLS3/4 organisms.

Bryan

Of course one would not use a simple laminar flow hood for working with human pathogens and that's not the issue here. To my knowledge the "hood" you are referring to for BLS3/4 organisms is known as a biosafety cabinet.

I'm not sure about Plant Biologist being weird, I've only worked with a few but you are right if you are referring to Plant Pathologists as weird, we are, but it not because we choose to use the best tools available.

 

[podz]

Source for lab odds and ends...

I've bought my entire collection of glass from second-hand stores such as Red Cross and Fida. I pop into them now and then and they don't usually have any lab glass, but now and then I find a gem.

My most recent find was a 10000 ml (10 litre) pyrex cylindrical flask with a pouring nose. That was 10 EUR, an extremely good bargain.

I picked up a 5000 ml erlenmeyer flask for 10 EUR as well.

Countless other odds and ends: petri dishes, measuring cylinders, boiling flasks, etc.

 

[Warthaug]

Of course one would not use a simple laminar flow hood for working with human pathogens and that's not the issue here. To my knowledge the "hood" you are referring to for BLS3/4 organisms is known as a biosafety cabinet.

Biosafety cabinets are but one form of the many types of laminar flow hoods that exist. But again, in any area of microbiology in which I have had experience (which spans a pretty broad range of the field - although obviously not to plant microbiology) no one uses any form of laminar flow hoods outside of high-risk pathogen work and preparation of pharmaceutical materials*.

* EDIT: and mammalian cell culture

I'm not sure about Plant Biologist being weird, I've only worked with a few but you are right if you are referring to Plant Pathologists as weird, we are, but it not because we choose to use the best tools available.

But are they the best tools available, or is it a case of bad or unnecessary methodology being propagated over time? This happens a lot, in many scientific fields.

Just for comparison, in my lab we prepare somewhere in the neighbourhood of 10,000 - 20,000 bacterial cultures a year, and aside from the 1-2 contaminated samples that inevitably come with training a new grad student, we never suffer contamination. Primary clinical samples (blood, tissues, etc) and handled in a hood; the remaining 99% (which includes post-isolation clinical samples) are handled on a bench using a flame & loop.

B

 

[Warthaug]

Source for lab odds and ends...

I've bought my entire collection of glass from second-hand stores

There are a lot of used lab ware companies out there, but ironically one of our better local sources here is a surplus store. Ebay is a good place to look as well.

Bryan

 

[trentm]

It doesn't matter all that much - the amount of O2 actually entering the starter is rather small (I think Kai did some experiments on this end). The main benefit of stirring is removal of CO2, which will have no trouble getting past a tightly "sealed" foil cap.

I don't think there is a difference; commercially it is usually sold as wort-agar, but I treat the terms as interchangeable. That said, beer-agar may indicate a hopped product, which would have a weak selectivity against gram positive bacteria. DME + water + nutrient is more than enough - conversion, hydration, etc, are already complete with DME (there is no starch in DME; its all converted). I take mine through the hot-break, simply because that is inevitable if boiling long enough to sanitize the wort.

Bryan

Kia's work shows a significant increase in yeast growth with increased stir speed and in his discussion he attributes the increase in growth to an increase in air: http://braukaiser.com/blog/blog/2013/03/25/stir-speed-and-yeast-growth/. Here is an article by a well established brewers yeast expert that illustrates the importance of "aerating" a yeast starter:http://www.pivarstvo.info/forum/files/yeast_propagation_and_maintenance_607.pdf. Finally, here is a journal article that proves air is taken into a flask on an orbital shaker. However, it does not address air intake in a stirred flask but one could draw a conclusion based much evidence and common understanding that it does happen.

I think the post was referring to Universal Beer Agar. As posted earlier, here is a link to a fact sheet on UBA: http://www.neogen.com/Acumedia/pdf/ProdInfo/7574_PI.pdf

 

[Warthaug]

Kia's work shows a significant increase in yeast growth with increased stir speed and in his discussion he attributes the increase in growth to an increase in air: http://braukaiser.com/blog/blog/2013/03/25/stir-speed-and-yeast-growth/.

I was actually thinking of this one, where he found minimal differences between varying levels of aeration, so long as there was some minimal exchange of air:
http://braukaiser.com/blog/blog/2013/03/19/access-to-air-and-its-effect-on-yeast-growth-in-starters/

Here is an article by a well established brewers yeast expert that illustrates the importance of "aerating" a yeast starter:http://www.pivarstvo.info/forum/files/yeast_propagation_and_maintenance_607.pdf

Figure 1 of this article shows that stirring has ~8X the effect of aeration, which reflects what I wrote earlier - e.g. expulsion of CO2 being a more important effect of stirring than O2 introduction. Retention of CO2 in solution is a major issue in commercial microbiology, as it is an enzymatic product produced during energy metabolism, and thus its presence inhibits the rate of metabolism through inhibiting reactant flow through the final stages of glycolysis/P5P/TCA/etc pathways (example).

Finally, here is a journal article that proves air is taken into a flask on an orbital shaker. However, it does not address air intake in a stirred flask but one could draw a conclusion based much evidence and common understanding that it does happen.

Please point out where I said that air entry does not occur into stirred cultures. You cannot, as I never once made that claim. My point was the opposite - i.e. that worries about maximizing air entry is unfounded as the primary effect of stirring in a starter is the removal of CO2, not the introduction of O2. Both one of your links and another I provided above highlight the relative impact of oxygenation versus CO2 degassing, and clearly show the later to be more important than the former.

You seem to have gotten your back up about this, so lets lay down the science...at least as it pertains to saccharomyces.

It has been well established for many decades (specifically, since 1928) that Saccharomyces makes minimal use of O2 during exponential-phase growth. Due to a phenomena called the crabtree effect, Saccharomyces does not use oxygen for energy production or growth when sugar sources are plentiful; ergo, the amounts of O2 required for maximal grow are quite minimal. For example, R. H. De Deken (J Gen Micro, 1966) demonstrated that yeast in an oxygenated culture preferentially underwent fermentation over oxidative metabolism - preferring fermentation 50:1 to 250:1 over oxidative metabolism (depending on the strain). Moreover, these Saccharomyces consumed minimal O2 (later studies showed the portion consumed was almost entierly used for FFA and sterol synthesis). Specifically, Saccharomyces consumed from 0 ul/10^7 yeast cells (e.g. the amount off O2 dissolved in the media prior to the experiment was sufficient) to a meagre 4.8 ul O2 per 10^7 yeast cells. To put the high end into context, for your average 1L starter (250 billion cells), that's equivalent to the amount of O2 in ~120ml of headspace in your flask, assuming 5ppm dissolved O2 at the start.

Not all yeasts undergo the crabtree effect - brett, for example, does a 50:50 mix of fermentation and acetic acid production (to my knowledge, no one has extensively studied brett production versus O2 availability). Obviously, none of this applies to lacto or pedio who don't give a rats rear-end if O2 is around or not.

B

 

[trentm]

I was actually thinking of this one, where he found minimal differences between varying levels of aeration, so long as there was some minimal exchange of air:
http://braukaiser.com/blog/blog/2013/03/19/access-to-air-and-its-effect-on-yeast-growth-in-starters/


Figure 1 of this article shows that stirring has ~8X the effect of aeration, which reflects what I wrote earlier - e.g. expulsion of CO2 being a more important effect of stirring than O2 introduction. Retention of CO2 in solution is a major issue in commercial microbiology, as it is an enzymatic product produced during energy metabolism, and thus its presence inhibits the rate of metabolism through inhibiting reactant flow through the final stages of glycolysis/P5P/TCA/etc pathways (example).


Please point out where I said that air entry does not occur into stirred cultures. You cannot, as I never once made that claim. My point was the opposite - i.e. that worries about maximizing air entry is unfounded as the primary effect of stirring in a starter is the removal of CO2, not the introduction of O2. Both one of your links and another I provided above highlight the relative impact of oxygenation versus CO2 degassing, and clearly show the later to be more important than the former.

You seem to have gotten your back up about this, so lets lay down the science...at least as it pertains to saccharomyces.

It has been well established for many decades (specifically, since 1928) that Saccharomyces makes minimal use of O2 during exponential-phase growth. Due to a phenomena called the crabtree effect, Saccharomyces does not use oxygen for energy production or growth when sugar sources are plentiful; ergo, the amounts of O2 required for maximal grow are quite minimal. For example, R. H. De Deken (J Gen Micro, 1966) demonstrated that yeast in an oxygenated culture preferentially underwent fermentation over oxidative metabolism - preferring fermentation 50:1 to 250:1 over oxidative metabolism (depending on the strain). Moreover, these Saccharomyces consumed minimal O2 (later studies showed the portion consumed was almost entierly used for FFA and sterol synthesis). Specifically, Saccharomyces consumed from 0 ul/10^7 yeast cells (e.g. the amount off O2 dissolved in the media prior to the experiment was sufficient) to a meagre 4.8 ul O2 per 10^7 yeast cells. To put the high end into context, for your average 1L starter (250 billion cells), that's equivalent to the amount of O2 in ~120ml of headspace in your flask, assuming 5ppm dissolved O2 at the start.

Not all yeasts undergo the crabtree effect - brett, for example, does a 50:50 mix of fermentation and acetic acid production (to my knowledge, no one has extensively studied brett production versus O2 availability). Obviously, none of this applies to lacto or pedio who don't give a rats rear-end if O2 is around or not.

B

My "back is not up about this", I am just trying to respectfully disagree.

Both of Kai's articles appear to me to be championing the advantages of O2 in a starter and neither even mention the advantage of evacuating CO2. That said, I agree it is important.

Dr. Raines-Casselman's article does not mention the advantage of evacuating CO2. However, I agree with you, it is important.

I did not mean to indicate that you said air entry does not occur. I apologize for the misunderstanding. However, I think you do not feel it is not important and that's where we disagree. If you will look at the conclusions drawn by both authors you will see that Kia and Raines-Casselman both agree the effect of increased yeast growth is due to exposure to air.

The crabtree effect is not at issue here as it refers to respiration as opposed to fermentation. Due to the crabtree effect S. cerevisiae cannot undergo respiration (at least it is very limited) when it exposed to an environment where sugar is present at greater than 0.2 percent. It is true that if we could present that environment we could achieve much greater yields but it is not feasible for a home lab to grow yeast under these conditions as it requires a fed batch system. This system is used to produce dry yeast at an industrial level.

Here is my view: We produce beer under anaerobic fermentation because exposure to O2 during fermentation causes problems with the beer. The challenge is keeping the yeast healthy under these conditions. We know that yeast need healthy cells walls and permeable cell membranes in order to bud and process nutrients and waste material. We also know the components that make healthy yeast are synthesized in the presence of O2. Once that is depleted as happens very quickly in beer fermentation, very few of these components can be synthesized. Now these components are shared with each new cell, so with each doubling, yeast health is diminished. Therefore at the end of anaerobic fermentation, yeast health is the poorest when we need it the most. So our job is to pitch yeast with highest level of health possible, enabling the yeast to complete the job. Under aerobic fermentation (fermentation in the presence of air and not to be confused with respiration) as we do with starters each new yeast cell can synthesize the components needed for vitality. If provided with air throughout propagation our yeast will be fully ready to make the great beer we strive for and for reasons I don't understand our yield will increase. Whether it is evacuation of CO2 or the presence of O2 that increase the yield, we know it is the presence of O2 that makes our yeast healthy.

The way I see it, aeration of a starter is highly important for 2 reasons: 1) to produce healthy yeast, 2) to obtain high yield. I do agree that removal of CO2 from the starter is important. It is fortunate that our best method (stir plate or shaker) for increasing O2 also removes CO2. I suppose it's OK if you see it the other way around.

 

[ajdelange]

The way I see it, aeration of a starter is highly important ....

In my brewery it certainly is. If I don't oxygenate the starter doesn't start. I had a situation where the starter (3 gal) wasn't up to snuff after a couple of days so I declared 'bad yeast' and got two more packs (making sure they were from a different lot). When they didn't start either I declared 'Ah, sub lot from the same bad main lot' and got two tubes of the same strain from the other manufacturer. When those didn't start I said 'Must be something else.' I may be slow but eventually I catch on.

'Something else' turned out to be that the O2 flow meter had cracked at the outlet port so that most of the O2 was going into the room even though the pith ball showed 1 LPM flow. Enough went through the stone to put some froth on the wort. I supposed I should have noticed that it wasn't a normal amount but I didn't.

Replaced the flow meter and everything was fine again.

I'll also mention that that in experiments in which I oxygenated wort and pitched yeast a DO meter indicated the O2 was completely consumed in about half an hour. Those growing yeast for certain want oxygen whatever M. Pasteur and Mr. Crabtree may have said.

 

[Warthaug]

I did not mean to indicate that you said air entry does not occur. I apologize for the misunderstanding. However, I think you do not feel it is not important and that's where we disagree.

In that assumption you would be wrong, however yeast biology dictates that the removal of CO2 will have a greater impact on yield than oxygenation. As the De Deken study showed, for most yeasts the amount of O2 required is the amount dissolved in a properly oxygenated starter; no additional (or minimal benefit) is gained from additional O2 entry.

Your link too showed the same effect - aeration with an airstone (which would provide better oxygenation than stirring), but without stirring, provided a minimal benefit. Stirring by itself was far superior; despite (in theory) providing much poorer aeration.

My point was never that oxygenation was not important, but rather it is unnecessary to take extra measures to ensure high levels of air exchange (i.e. tight foil is fine).

If you will look at the conclusions drawn by both authors you will see that Kia and Raines-Casselman both agree the effect of increased yeast growth is due to exposure to air.

In Kai's case, he lacks the data to separate the two effects. Raines-Casselman's data runs counter to his conclusion.

The crabtree effect is not at issue here... is true that if we could present that environment we could achieve much greater yields but it is not feasible for a home lab to grow yeast under these conditions as it requires a fed batch system

Its the predominant form of metabolism yeast will go under in our starters, so it is very much germane. It explains why the differences in yield observed by people like Kai and Rains-Casselman are so small; yield factors for respiring yeast would be around 35; a much larger increase over the levels observed by Kai under anoxic versus aerated conditions (yield factor changed from 1.5 -> 2.5).

How the yeast are metabolizing also tells us a lot about what we can (practically) do to improve our starters. A minimal degree of aeration is required; else we end up with poor-quality yeast lacking sterols and UFAs. But exceeding that level has minimal benefit in terms of yields or yeast health. If trying to optimize your starter there are better places to focus your attention - be it improving CO2 removal, increasing the proportion of fermentables versus dextrans, improving micronutrient levels, etc. I agree batch-feeding is not something we can practically do, and based on the statements and writings of White, it sounds like it produces yeast unfavourable for beer production anyways.

Here is my view: We produce beer under anaerobic fermentation because exposure to O2 during fermentation causes problems with the beer.

I clipped the rest of this, because I agree with what you wrote absolutely. But what I think you are doing is over-estimating the amount of oxygen required for the synthesis of sterols and UFAs. I referenced the De Deken paper for a reason - it quantified that requirement under conditions in yeast undergoing logrithmatic growth while growing under crabtree conditions. And the answer is the amount of O2 required is minimal - a few ul of gas per 10^7 cells - AKA roughly the amount of O2 in the wort if you oxygenate it properly before placing it on the stirplate.

Bryan

 

[butterpants]

While I truly do appreciate the academic conversation, I don't want this to devolve into a pissing argument over journal article nuances or unpublished and non-peer reviewed off the cuff studies.

Cynmar boxes arrived today. SCIENCE BONER

 

[trentm]

While I truly do appreciate the academic conversation, I don't want this to devolve into a pissing argument over journal article nuances or unpublished and non-peer reviewed off the cuff studies.

Cynmar boxes arrived today. SCIENCE BONER

Agreed. See a doctor if it last more than 4 hours!

 

[podz]

Not all yeasts undergo the crabtree effect - brett, for example, does a 50:50 mix of fermentation and acetic acid production (to my knowledge, no one has extensively studied brett production versus O2 availability). Obviously, none of this applies to lacto or pedio who don't give a rats rear-end if O2 is around or not.

B

Not wanting to quote the entire long post, let's just say that I liked this post and if I could like it 50 times by myself then I would.

 

[butterpants]

Recommendations on the membrane filtration apparatus? Reusable preferred... for the Nalgene all I found were disposable.

What type filter material? 45um pore diameter right? Hand vacuum pump ok?

 

[butterpants]

Anyone care to help me develop a yeast propagation protocol (slant to pitch) for the volumes I can work with?

Slant->30ml->300ml->3L->Decant/Pitch

Growth timeline for the individual volumes?

When/if counting cells and determining viability?

Liquid growth media makeup?

While I'm very familiar with growing starters... that's always been from a large pitch packet or vial... and I never much cared about specific pitch rate numbers. Everything got a 2L starter.

Pics of basement lab to come soon!

 

[Warthaug]

Recommendations on the membrane filtration apparatus? Reusable preferred... for the Nalgene all I found were disposable.

What type filter material? 45um pore diameter right? Hand vacuum pump ok?

What filter you need (if you need one) depends on your purpose. These days reusable filters are rare, you may want to look at second hand stores for them. Whatman still makes membranes for a range of reusable units.

Anyone care to help me develop a yeast propagation protocol (slant to pitch) for the volumes I can work with?

Slant->30ml->300ml->2L->Decant/Pitch

I would recommend a smaller step for the first step - 7-10ml is what I normally use for my first step. This is your highest risk of infection, so its best to keep the step small to limit risk. I will go from that to 250ml, and from there upto 3L*. A good rule of thumb is stepping up by 10's, but obviously I don't follow that religiously.

By upto 3L I mean I'll use it in final-stage starter upto 3L in volume, not that I do everything in 3L final steps. For most beers a 2L final stage is sufficient. 3L final stages are good for lagers and big ales.

When/if counting cells and determining viability?

I only count at the end of the last step, and to be honest, I rarely do viability stains. For various reasons they're not all that accurate with yeast, and more to the point, if you've started with a loop of yeast from a slant and end up with 2L of high-density yeast, by definition your yeast is quite viable.

Liquid growth media makeup?

1.035 to 1.040 SG wort (DME or made from grain), plus double the recommended amount of quality yeast nutrient by volume. I experimented a bit with potato-dextrose media (its dirt cheap), but found that the yields of yeast I got were poor. Its a great medium for growing yeast on a plate, but as a liquid medium its not the best.

Bryan

 

[butterpants]

Copy, thanks for the quick reply.

For the filters, I plan on using sterile disposable... but for the holding/vacuum apparatus, there's where I was looking for something reusable. Not the membrane. Sorry for the confusion. Far as the membrane goes I was wondering if a specific material composition and pore diameter was preferable, that's all. Planning on running 100ml of bottled beer through then taking the filter and growing it on selective media to see if non sacc yeast or bacteria grow.

 

[Warthaug]

I realize you were not expecting to reuse the membrane, I was simply pointing out that Whatman still makes the (single-use) membranes for the reusable filter cartridges/apparati. If you look at their webpage you may be able to track down a manufacturer. We use a similar system in one of the undergrad labs I run (for filtering water to look for coliforms); I'll see if I can dig up the manufacturer - ours are quite old, so I don't know if they're still made.

Pore-size wise, you want either a 0.45 or 0.22 um filter; bacteria range from 0.1-~2 um, yeast can be as small as 1 um, so a 45 um filter won't do the job.

Bryan

EDIT: the filters we use in the undergrad labs are Whatman, Cellulose Nitrate, 0.45um pore size, 47mm Diam.

 

[butterpants]

gracias

 

[dstar26t]

If you want to go with a bunsen burner, you can use a Home Depot camping cylinder, a Camco 57626 regulator (~$20 on Amazon) for fuel/control and I use an EISCO burner (CH0094LP) and hose (CH0100A), both from Amazon. You have to pull the snap ring and slide the threaded fitting off the regulator outlet and put the hose on with a hose clamp. It's the same as the one Avogadro sells except they add a 5/16" barbed fitting for $25.

I found Bromocresol Green on ebay for a reasonable price which you can use for selection of Sacch vs. Brett but it's not foolproof. Wallerstein media and cycloheximide are just too pricey. You can make a Potato Lactose media which should favor bacterial growth over Sacch/Brett.

Has anyone used antibiotics to suppress bacteria on a plate? I found Chloramphenicol as Viceton tablets which is a dog antibiotic and pretty cheap but you need a prescription. Fish penicillin seems easily available but I'm not sure how appropriate it is and at what concentration to use it. Chloramphenicol-N by Chevita is a powder for pigeons and is cheap but I can't find what the actual composition of the powder is.

Found some Cycloheximide at a fairly low price ($36) but it's for 100mg which should treat 10-25L of media so maybe not too bad.

Nate

 

[helibrewer]

Me 3 on the T handle! I have the same pressure cooker fwiw.

Just saw this, sorry for the delay. I added the "T" handle back when I was experimenting with steam injection in my mash tun...Now I use it just to monitor and vent (for non liquid items like my tools and plate chiller).