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Old 01-31-2013, 02:51 PM   #21
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Originally Posted by Huff360 View Post
Rough volume? Did you just do all your dilutions into vinegar instead of water?
The WLP009 slurry that I was working with was about 1/3 thick settled slurry to beer. (which is pretty typical)

For viability I did 0.6ml homoginize slurry, diluted with 10ml of water. Then 0.6 ml of that with 0.6ml of 0.03%MB/water solution.

For cell density I used 0.6mm of the 0.6ml+10ml diluted sample with 0.6ml of 5% actic acid.

Take this for what it is. Simply the way one person does it. It's roughly based on what White Labs suggests with some things I've found other places and what I've experianced.

Here is my procedure:
http://woodlandbrew.blogspot.com/201...viability.html

And White Labs sugestion for a Glycine MB solution:
http://www.whitelabs.com/beer/alkaline.html
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Old 01-31-2013, 02:59 PM   #22
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Thanks for that. I didn't think of using vinegar.

Lab standard seems to be sulfuric acid but I'd like to avoid that.

Kai

Thanks for the valuable information that you pour out to the homebrew community!
I never thought I would stumble on something that you hadn't thought of.

Sulfuric acid makes me cringe. I'm not even sure I want to use phosphoric acid. I also considered using lactic acid because I have that for adjusting mash pH.
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Old 01-31-2013, 03:42 PM   #23
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Originally Posted by WoodlandBrew View Post

AJ's statistical analysis is a good way to decide how many boxes to count. I also look at box to box standard deviation and make sure it is reasonable.
The average number of cells in a square is N*p where N is the number of cells you pipeted and p is the probability that a cell winds up in the square you are looking at which is the the ratio of the volume of the square to the volume you pipet. If you have a sample with 10,000,000 cells/mL, a large square has volume (which it typically does) of 0.0001 mL and you pipet 0.1 mL then N = 1,000,000 cells; p = 0.0001/0.1 = 0.001 and you expect to see 1,000,000 * 0.001 = 1000 cells. The standard deviation in the number of cells in a square is sqrt(N*p*(1-p)) which, as p is 0.001, is close to sqrt(N*p). Thus, as long as the amount you pipet is larger relative to a square size the standard deviation is closely approximate to the square root of the average cell count. For the example of 10,000,000 cells/mL with an average large square count of 1000 the standard deviation between large squares would be sqrt(1000) = 31.6.

The same applies to the small squares except that p is even smaller and the approximation even better. My hemocytometer has 16 small squares per large (many have 25) so I'd expect 1000/16 = 62.5 cells per small square and the standard deviation in small square count should thus be sqrt(62.5) = 7.9.

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Originally Posted by WoodlandBrew View Post
..keep in mind that dilutions and homogenization are also critical.
It should be made very clear that the uncertainty of 1/sqrt(cells_counted) uncertainty I mentioned in the earlier post is in addition to other uncertainties. Each time you dilute you introduce uncertainty not just from the uncertainties in the measurements of the volumes (though those are the major factor no doubt) but because you are making a random draw from a larger population of suspended yeast cells. There is a cell density in the fermentor - that's what we are trying to determine. The cell density in a hydrometer sample, for example, is not the same. Its expected cell density is but its actual cell density is a random variable with standard deviation dependent on the ratio of the sample size to the full volume. The smaller the sample the larger that deviation.

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One thing that I have found recently is that a pipette can retain 5% of the previous sample clinging to it's walls. So if you wash your pipette and then pull a sample it could be 5% diluted.
There are two types of pipet - blow down and non blow down. In a blow-down pipet you squeeze the pipetter bulb to force air through the pipet in order to expel all (or most all) of any liquid retained. Pipets are marked with either a double band or single band at the top to tell you which type you have.
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Old 01-31-2013, 04:17 PM   #24
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Thanks for elaborating on these points AJ,

That is all excellent information.

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Old 02-01-2013, 09:00 PM   #25
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One thing that I have found recently is that a pipette can retain 5% of the previous sample clinging to it's walls. So if you wash your pipette and then pull a sample it could be 5% diluted.
That's why you should rinse the pipette with the sample solution. In practice this means pulling in a sample, pushing out that sample and pulling in another sample.

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Old 02-01-2013, 09:51 PM   #26
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That's why you should rinse the pipette with the sample solution. In practice this means pulling in a sample, pushing out that sample and pulling in another sample.

Kai
Yes, thanks for taking that to conclusion.
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Old 02-01-2013, 10:22 PM   #27
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That's why you should rinse the pipette with the sample solution. In practice this means pulling in a sample, pushing out that sample and pulling in another sample.

Kai
The lady on YouTube said I should do that 3 times! (I'm having to get all my chem lab teachings from the internet as I had very little of it in school.)

One thing I am not quite sure on -

During a serial dilution, when I pipette my 1mL of sample into 10mL of diluent, should I then suck up some the diluted sample and rinse the pipette with it?
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Old 02-01-2013, 10:40 PM   #28
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During a serial dilution, when I pipette my 1mL of sample into 10mL of diluent, should I then suck up some the diluted sample and rinse the pipette with it?
yes, you want all the the solution in the pipette to have the same cell density.

properly mixing the diluted sample is also important. But don't shake it and create lots of foam. Some yeast strains like to aggregate in the foam and you may end up removing cells from the suspension.

When counting you should at least fill both sides of the hemocytometer with different samples. If you mixed correctly both counts should be close. If not there is a problem. Others recommend to fill the hemocytometer three times and count 3 times.

Kai
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Old 02-01-2013, 10:49 PM   #29
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Quote:
Originally Posted by Kaiser View Post
yes, you want all the the solution in the pipette to have the same cell density.

properly mixing the diluted sample is also important. But don't shake it and create lots of foam. Some yeast strains like to aggregate in the foam and you may end up removing cells from the suspension.

When counting you should at least fill both sides of the hemocytometer with different samples. If you mixed correctly both counts should be close. If not there is a problem. Others recommend to fill the hemocytometer three times and count 3 times.

Kai
As a practical matter, generally how consistent are your counts?
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Old 02-02-2013, 12:21 AM   #30
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Quote:
Originally Posted by Huff360 View Post
The lady on YouTube said I should do that 3 times! (I'm having to get all my chem lab teachings from the internet as I had very little of it in school.)

One thing I am not quite sure on -

During a serial dilution, when I pipette my 1mL of sample into 10mL of diluent, should I then suck up some the diluted sample and rinse the pipette with it?
Pipetting the mixture multiple times ensures a homogenous suspension (well mixed), as Kaiser said. When performing serial dilutions, you want to pipette 1ml of sample into 9ml of diluent for a 1:10 dilution. Ideally, you would want to change pipettes, although probably not required here. Then, you remove 1ml of the diluted mixture and go down the line. If diluted enough, you can view that sample on the hemocytometer for a count compensating for the dilution.
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