Building a home QC lab... questions

[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).

 

[butterpants]

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).

Wow, ha. Thanks for eventually answering us

 

[butterpants]

Ok guys I've been messing around with this a bunch and am back for follow up Q's.

Putting cell counts and pitch rates into practice
Does this sound logical? I snagged a 7 month old Wyeast pack from the LHBS for free. Wanted to see if I could make this sucker viable for a session beer pitch in a limited amount of time. I was figuring maybe 10% viability in the pouch.

So I made a starter. 2L wort + stirplate in a 4L flask. Lots of room for gas exchange. 1.040 was the starter gravity + nutrient addition. 68F for 36 hours. Initially I was worried as there was no sign of kreusen at all, which is odd for my starters but considering the initial pitch rate, oh well. Around hour 30 it seemed to become substantially more turbid than before so I felt confident the log growth phase was progressing. Starter was cold crashed at 40F for 2 days and around 1600mL decanted and thin slurry was waiting for brew day.

MrMalty says for 5 gallons of 1.053 wort, I'll need 185 billion cells.

I remove a small sample of the 400ml thin slurry. Make a set of serial dilutions by adding 9ml of DI water to a few test tubes then 1ml of the slurry, then 1ml of the mix, etc eventually getting 1:10, 1:100 and 1:1000 samples. The 1:10 factor is what I used to count.

Using my hemocytometer, I count 170 cells in the 16 small (1/16th) squares. Phone app gives me 4.25e7 or 42.5 million cells per ml (1:10 dilution).

Then I add 1ml alkaline methyl violet solution to a 1ml sample of the 1:10 dilution creating a 1:20 dilution that I can assess for viability.

Again using the hemocytometer, I count 92 cells in 16 squares but this time with 9 staining purple (that weren't budding). Phone app gives me 2.30e7 or 23 million cells per ml (1:20 dilution). (92-9)/92x100= 90% viability.

Sooo the average pitch rate between the two counts without dilution factor is 442 million cells per ml. I have 400 ml of slurry. Multiply them together to get a pitch of 176.8 billion cells....of which 90% are viable, so 159 billion viable cells pitched, slightly under the MrMalty calculated rate.

Does this look correct? I hate math. Not a bad turn around though from a mega old yeast pack.

How are you guys decanting off starters? For some of the less flocculant strains, I don't think pouring off the top is effective. I'd like to siphon out of the flask but man that takes forever even with a 50 ml pipette, if you can even find a decent bulb for one (I can't). Is there some sort of sterile suction method/apparatus that will somewhat quickly remove large quantities of fermented starter wort from big Erlenmeyers?

Filling hemocytometers? I've been using a 100-1000ul pipetteman and make a mess every time. Gotta be a better way. Small glass pasture pipettes??

 

[WoodlandBrew]

I remove a small sample of the 400ml thin slurry. Make a set of serial dilutions by adding 9ml of DI water to a few test tubes then 1ml of the slurry, then 1ml of the mix, etc eventually getting 1:10, 1:100 and 1:1000 samples. The 1:10 factor is what I used to count.

Using my hemocytometer, I count 170 cells in the 16 small (1/16th) squares. Phone app gives me 4.25e7 or 42.5 million cells per ml (1:10 dilution).

Then I add 1ml alkaline methyl violet solution to a 1ml sample of the 1:10 dilution creating a 1:20 dilution that I can assess for viability.

Again using the hemocytometer, I count 92 cells in 16 squares but this time with 9 staining purple (that weren't budding). Phone app gives me 2.30e7 or 23 million cells per ml (1:20 dilution). (92-9)/92x100= 90% viability.

Sooo the average pitch rate between the two counts without dilution factor is 442 million cells per ml. I have 400 ml of slurry. Multiply them together to get a pitch of 176.8 billion cells....of which 90% are viable, so 159 billion viable cells pitched, slightly under the MrMalty calculated rate.

Does this look correct? I hate math.

It's great that you are doing cell counts. Personally, I would have been interested in what the viability was before the starter.

It sounds like what you counted is one "box." (this area fits nicely inside the field of view at 400x on most microscopes.) Each box is 4nl. Typically five boxes are counted for statistical reasons. 170 cells / 4nl is 42.5 cells per nl or 42.5 billion cells per liter. With the 10:1 dilution that means that the original sample was 425 billion cells per liter.

92 cells / 4nl is 23 billion per liter. factor in the 20:1 dilution and you have 460 billion cells per liter. About 10% variation. That's pretty typical for what is essentially box to box variation.

Your viability assessment is correct, but as mentioned before, most people count 5 boxes.

 

[butterpants]

It's great that you are doing cell counts. Personally, I would have been interested in what the viability was before the starter.

It sounds like what you counted is one "box." (this area fits nicely inside the field of view at 400x on most microscopes.) Each box is 4nl. Typically five boxes are counted for statistical reasons. 170 cells / 4nl is 42.5 cells per nl or 42.5 billion cells per liter. With the 10:1 dilution that means that the original sample was 425 billion cells per liter.

92 cells / 4nl is 23 billion per liter. factor in the 20:1 dilution and you have 460 billion cells per liter. About 10% variation. That's pretty typical for what is essentially box to box variation.

Your viability assessment is correct, but as mentioned before, most people count 5 boxes.

So my counts are off by a factor of 100?

170 cells counted in 1 small square come out of my app at 4.25e7. I would read this as 42,500,000 (42 million). Multiply by 10 for the dilution factor and you get 425 million cells per ml. You say 425 billion per ml.

If so then I pitched 159 trillion cells? That doesn't seem right. I can't imagine propagating that many in such a short time from that gimped source. Please help!

I'll attach a hemocytometer app screenshot from my phone....one sec

 

[butterpants]

So I chose option #3 and enter 170 cells in 1 box.

 

[butterpants]

And I was interested in the original pack viability but to be honest the starter was prepared at 2am after a long day at work and I had no desire to do lab stuff at that late hour. . A moment of weakness, HA

 

[ajdelange]

The number of squares you count is determined by the accuracy you want. The coefficient of variation in your estimate is 1/sqrt(n) where n is the number of cells you counted. If you count 100 cells you will have Cv = 1/sqrt(100) = 0.1 and what ever you determine that to be, dependent on how many boxes of what size it took to get 100 cells, the estimated error will be ±10%. So, suppose you go into that central area where the boxes are defined where triple lines intersect and count 19, 20, 21,18,18 and 24 cells in the top row (out of the total of 25 in that area). That's 100 cells. If the central area is 1 mm x 1 mm (typical) and the depth 0.1 mm (also typical) you would have 100 cells in 5*2E-4*2E-4*1E4) = 20*E-12 m^3 = 20*E-6 cc for a density of 100/20E-6 = 5E6 cells/mL ± 10%, If you counted two rows (10 boxes) and got a total of 200 cells your Cv would go down by a factor of sqrt(2) to 1/sqrt(200) = 0.0707 or 7%. You counted twice as many cells in twice the volume so the densty is now 200/40E-6 which is still 5E6 cells per mL but the standard error is now 7% of that or ±350,000 cells/mL. Dilution factors would, of course, need to be applied. To halve your accuracy you have to double the number of cells counted.

 

[butterpants]

Thanks for the reply AJ. Yes next time, more counting, less error. My app gave me a 15% error margin so I was tracking it just didn't talk about it here. My fault.

You didn't touch on Woodlands comments and the discrepancy of 100 fold in his vs my numbers. Need to get this remedied....maybe it's just something innocuous I'm missing

 

[ajdelange]

I'm sure the count discrepancies are due to incorrect volume assumptions on someone's part. If the central area is 1 mm x 1mm x .1 mm then it corresponds to 1E-3*1E-3*1E-4 = 1E-10 m^3 which is 1E-4 cc. That area is subdivided into 25 boxes (each of which has 1 or 2 triple ruled lines) and they are (1E-4)/25 = (100E-6)/25 = 4E-6 cc (4 nL). Those boxes are in turn subdivided into 16 subsquares each of which contains (4E-6)/16 = (40E-7)/16 = 2.5E-7 cc (0.25 nL)

 

[WoodlandBrew]

If someone else ran the numbers I would feel more confident, but it seem that you just misread the units that I wrote. We are getting the same result. 425 billion cells per liter (not billion per milliliter) is what I came up with for the first cell count, and 460 billion per liter for the second. 400ml of 425 billion per liter would be 170 billion cells. 400ml of 460 billion per liter would be 184 billion cells. At 90% viability that's 153 and 166 billion cells.