Experimental pH manipulation of a yeast starter

[ldave]

I've been researching the limitations of yeast starters. In particular, why a starter, innoculated with a given amount of yeast cells, has an upper limit to it's size. Thus, the whole step-up thang.

So, it appears that yeast health suffers when you pitch a small innoculation into a large starter. Apparently, yeasts create a low pH during fermentation as part of the process to ensure cell wall permeability. If the initial pH is high, they have to work hard to do this. If the initial wort is strongly buffered at a high value, they have to work really hard to do this. So, when you underpitch a starter, so few cells are acidifying the wort that the pH never really gets down to where it needs to be and the cells wear out and the tires fall off of that jelopy.

Now, boiled worts have a pH around 5.0-5.3 or so. And finished beer has a pH 4.1-4.5 or so. Suppose you know that with a given specific gravity of a given DME boiled starter wort you get a pH of, say, 5.1. And suppose, by trial, you establish that a given strain of yeast lower that pH to, say, 4.4 during fermentation. So, here's my question: why not take the boiled starter wort and acidify with phosphoric acid to pH=4.4 to begin with? That's what the yeast want to do immediately anyway. Why not save them the energy expenditure? With this obstacle removed, wouldn't any size starter from any size innoculation be possible?

 

[ldave]

To develop this idea further, I researched mead fermentations. It seems that sluggish fermentations are often a result of must pH becoming out of bounds for the yeast. Ideal pH window: 3.7 to 4.6. So, for instance, if you add ingrediants that are very acidic like, say, raspberries, to the must, the fermentation can reduce pH to below 3.7 and the fermentation stops. Solution: add base to raise ph into the window. So, it appears that yeast continues to produce acids even when they are in their window and can continue to acidify their environment until they are out of window.

Assuming that the ideal pH window is 3.7 to 4.6, it would be reasonable to conclude that there is an ideal pH target for fermentation, just as there is an ideal pH target for mashes. What would be useful, then, is increasing the buffering capacity of the must or wort solution at that ideal pH point. This is what the product '5.2' does for mashes: it buffers at 5.2, resisting pH change in either direction. Could it be that a similar product, for fermentation, buffered at, say, 4.1 could produce consistant fermentations irrespective of innoculation levels? I'm sure that such a product would massively and negatively affect flavor profile, so it wouldn't be good for making beer. But, in a starter, where we decant the 'beer', this would be irrelevant. If all this were true, it would then be possible to pitch a 1 ml aliquot of yeast into a 2.5 L starter and generate enough yeast to decant and pitch into 10 gal of wort. A single tube of White Labs yeast would be an entire yeast bank for a homebrewer, with only a large single step starter necessary for each 1 ml aliquot.

 

[flars]

When you say a starter 'has an upper limit to it's size'. Do you mean OG or volume?
I have been looking for information similar to this. Do you have a reference?

 

[ldave]

The OG part of the question is easy: no.

OG appears to have a sweet spot. Most references say between 1.030 and 1.040 for non-stressed yeast. The BrewersFriend Yeast Calculator pin points it at 1.036. 'Yeast', White and Zainasheff, references 1.030 to 1.040 for non-stressed yeast and 1.020 for stressed yeast. So, here, OG is best thought of in terms of 'optimized', not maximized. That's all about osmotic pressure, which you shouldn't overdo with too high of a concentration. Too low a concentration and you're justing wasting flask space.

The short version of the volume question is: yes. But.

The volume question needs some clarification and, at the same time, can get a little muddy. First, to clarify, as 'Yeast' points out, you can't really talk about volume of a starter without some reference to how much yeast you've innoculated with. So the volume question really boils down to innoculation rate: (how much you've innoculated with)/(volume of starter). It's only when you've standardized the innoculation amount that you can really just refer to volumes. So, if you always pitch the same ole White Labs tube, then the innoculation amount is always the same, and the question then is: how much starter volume can I increase to in one step without getting bad effects. And here's where we go a little muddy. So, increasing starter volume is equivalent to decreasing yeast innoculation, which, at some point, becomes 'under pitching'. That this effect exists is not questioned, but putting a number on it appears to be a bit elusive. Looking again at the BrewersFriend Yeast Calculator, when you select a non-agitated starter, there is an upper limit on the yeast growth of 6, which is a simple multiplier starting with the innoculation amount. This factor is probably taken directly from the figure 5.8 in 'Yeast', p142. Translation: given a yeast amount, a starter OG, and a growth cap of 6, you arrive at a maximum starter volume. All of this logic in the calculator does not apply when you select the 'braukaiser-stir plate' method of calculation. With this method, there is no upper limit to yeast growth currently, but the documentation notes that this aspect should be forthcoming down the road pending more research. Commonly accepted wisdom for starters is the 'rule of tens', where each step is capped at 10 times the size of the last. So, practical people in practical jobs in the industry are essentially saying that there is a limit and, for good results, that number is x 10. 'Yeast' is a killer treatment of yeast and I would call it, in total, 'my reference' for much.

My pH references are a bit weaker, however. 'Yeast' offers almost none. My mead pH references are from 'The Compleat Meadmaker', Schramm. My reference for how fermentation pH proceeds in beer is, unfortunately, from an internet reference that I failed to save and have deleted my browsing history. I am trying to refind it because it was an interesting read and should be 'in the folder'. I find it curious that so much attention is focussed on mash pH, with some focus on the boil, and then, when the wort leaves the kettle, nobody really measures or cares very much about what happens to pH in the fermenter. Even 'Principles of Brewing Science', Fix (the bible, as far as I'm concerned), only cursorily mentions 'acidification' when discussing the lag phase of fermentation.

 

[ldave]

I found my 'net references for pH in fermentation. It originates in Kai Troester's Braukaiser series. Link:

http://braukaiser.com/wiki/index.php?title=How_pH_affects_brewing

Scroll to section 'Nutrient Uptake By Yeast'.

He solidly references this text in this section:

[Briggs, 2004] Dennis E. Briggs, Chris A. Boulton, Peter A. Brookes, Roger Stevens, Brewing Science and Practice, Published by Woodhead Publishing, 2004

And I don't have it. Figures.

 

[GuldTuborg]

Interesting thread. I'll be following this.

 

[ldave]

Yeast S. cerevisiae (brewers yeast) is used to produce alcohol for fuel. Study of bioreactors that are used to continously produce alcohol focus on maintaining yeast health to maintain high productivity. There are a varieity of stress factors. Of those tested, pH=3.7 is the number one bad one, followed by alcohol=9.5%. pH=4.5 is the number one protector from bad effects from all other stresses. Looks like, in this business, if you don't control pH, you lose, no matter what.

I'm beginning to think that if you've got a fermentation problem, one of the first things you should do is measure the pH.

pH relevant is highlighted p 5:

View attachment 2013 - pH- fuel ethanol fermentation-yeast inhibition factors and new perspectives to improve th.pdf

 

[RM-MN]

You're starting to question conventional wisdom........I like it. Please keep us informed on your findings.

 

[Warthaug]

Yeast S. cerevisiae (brewers yeast) is used to produce alcohol for fuel. Study of bioreactors that are used to continously produce alcohol focus on maintaining yeast health to maintain high productivity. There are a varieity of stress factors. Of those tested, pH=3.7 is the number one bad one, followed by alcohol=9.5%. pH=4.5 is the number one protector from bad effects from all other stresses. Looks like, in this business, if you don't control pH, you lose, no matter what.

I'm beginning to think that if you've got a fermentation problem, one of the first things you should do is measure the pH.

pH relevant is highlighted p 5:

This is becoming an interesting discussion. The paper provided by ldave was not the actual science done to show the effect of pH on fermentation & viability; they were reviewing the work of others. The paper they reference can be found here:
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2289159/pdf/jc1152289.pdf

Unfortunately, it appears that they have referenced the wrong paper; this paper does no experiments exploring pH and viability; instead it is looking at the processing of the yeast's H+-ATPase (which helps regulate intracellular pH).

I think, digging through their references, this is what they meat to reference:
ERASO, P; GANCEDO, C. Activation of yeast plasma membrane ATPase by acid pH during growth. FEBS Lett., v. 224, n.1, p.187-192, 1987.

I have access to it via a paid subscription service, so I cannot post it here. That said, I'm not sure it would support the OP's idea - according to that study, activation of the yeast H+ATPase can be via glucose or acidity. Glucose is a much more potent activator than pH, by a factor of >4, so the only time they saw significant pH effects was in glucose-poor media. The more important thing this paper found was that the combined glucose + pH ATPase activity was able to control intracellular pH down to about pH 4.5 (although it was lower at 4.5 than at higher pH). At pH's lower than that, yeast undergo a continuing decrease in their intracellular pH.

Long story short, I think the OP's experiment is worth trying, but I wouldn't expect much. As Kai's page discusses, pH is important for yeast uptake of nutrients. But, at least according to his source, simply pitching yeast at an appropriate density is sufficient. The above papers appear to put a limit on pH ranges; 4.5 being the bottom of "safe", 6.5 the 'top', and at the lower end (down to 4.5) there may be some protection against high alcohol & other stressors (which shouldn't be an issue in a starter).

Bryan

 

[RM-MN]

This is becoming an interesting discussion. The paper provided by ldave was not the actual science done to show the effect of pH on fermentation & viability; they were reviewing the work of others. The paper they reference can be found here:
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2289159/pdf/jc1152289.pdf

Unfortunately, it appears that they have referenced the wrong paper; this paper does no experiments exploring pH and viability; instead it is looking at the processing of the yeast's H+-ATPase (which helps regulate intracellular pH).

I think, digging through their references, this is what they meat to reference:
ERASO, P; GANCEDO, C. Activation of yeast plasma membrane ATPase by acid pH during growth. FEBS Lett., v. 224, n.1, p.187-192, 1987.

I have access to it via a paid subscription service, so I cannot post it here. That said, I'm not sure it would support the OP's idea - according to that study, activation of the yeast H+ATPase can be via glucose or acidity. Glucose is a much more potent activator than pH, by a factor of >4, so the only time they saw significant pH effects was in glucose-poor media. The more important thing this paper found was that the combined glucose + pH ATPase activity was able to control intracellular pH down to about pH 4.5 (although it was lower at 4.5 than at higher pH). At pH's lower than that, yeast undergo a continuing decrease in their intracellular pH.

Long story short, I think the OP's experiment is worth trying, but I wouldn't expect much. As Kai's page discusses, pH is important for yeast uptake of nutrients. But, at least according to his source, simply pitching yeast at an appropriate density is sufficient. The above papers appear to put a limit on pH ranges; 4.5 being the bottom of "safe", 6.5 the 'top', and at the lower end (down to 4.5) there may be some protection against high alcohol & other stressors (which shouldn't be an issue in a starter).

Bryan

That study references the yeasts reaction to glucose. I don't think I have much glucose in my beer. Does the finding of this study parallel what happens in wort or is it more aligned to what happens in bread where I add table sugar?

 

[Warthaug]

Wort should have some glucose; starch is, afterall, chains of glucose. Amylase breaks the starch down into glucose plus short chains of glucose (2 glucose = matlose, 3 glucose = maltriose, 4+ glucose = dextrins). Table sugar is a disaccharade of glucose + fructose. So you actually have more free glucose in your beer (some % of the total fermentables, exact amount depending on DME source/mash schedule) than you do with table sugar (no free glucose)

That said, the yeast break down maltose, maltriose and sucrose into their component sugars before metabolizing them; as such I would suspect that all of these "complex" sugars would activate the ATPase is a similar fashion. After all, the ATPase doesn't know where the glucose came from - it only sees the glucose*.

*The assumption here being that it is intracellular glucose that triggers the ATPase; if it is extracellular glucose than you would see this ATP response with wort, but not with table sugar.

Bryan

 

[NeverDie]

It seems that the OP hasn't logged back in since 2015, and I'm left hanging in suspense because no one reported back, at least not on this thread. Did anyone ever test the OP's theory that he so eloquently puts forth above, and if so, what were the results?

At the very least, it seems the OP was correct for the corner case of storing yeast. According to this reference, "All media used to store brewing yeast (slants, plates, etc.) should contain yeast nutrients, some malt, and should have an acidic pH. " https://www.maltosefalcons.com/tech/yeast-propagation-and-maintenance-principles-and-practices

 

[NeverDie]

I thought part of the reason for the upper limit on size, referred to by the OP, might have to do with the yeast's ability to outcompete and dominate competing microorgranisms. If the ratio of 1ml compacted yeast to 10ml wort is maintained, then the desired yeast will, almost by definition, dominate. On the other hand, if it were 1ml to 1000ml ratio, that leaves a lot of room for competing microorganisms to thrive without facing competition.

 

[IslandLizard]

I was curious to see if you could awake this topic. It holds a good promise...