Proper Haemocytometer Usage

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Huff360

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I am getting ready to start doing some cell counts. I have the Chinese improved nebular Haemocytometer. The only references I can find online for the proper way to do a count use the fancier triple line slide.

I am curious about which cells that lay on the perimeter to count.

Would this be correct?

Cell.PNG
 
I am getting ready to start doing some cell counts. I have the Chinese improved nebular Haemocytometer. The only references I can find online for the proper way to do a count use the fancier triple line slide.

I am curious about which cells that lay on the perimeter to count.

You need to decide what your cutoffs are, then count it the same way every time. I use a 50% cutoff, if half the cell is within the line of the grid area that I am counting, I count it.

Your picture is a tad confusing, but you should count the ones marked X, that are within the 4x4 square.

Here's a useful link:
http://www.animal.ufl.edu/hansen/protocols/hemacytometer.htm
 
Your picture is a tad confusing, but you should count the ones marked X, that are within the 4x4 square.

I can see how that would be. :eek:

I ran into this picture a lot when looking up the 'standard' protocol for counting.

Counting_of_cells_01.png

Basically, it says count anything that is touching the middle line on the top and left, but do not count anything touching the middle line on the bottom or right. I modified that a bit, cause to me not counting the top felt better. From: http://amrita.vlab.co.in/?sub=3&brch=188&sim=336&cnt=2

BUT - I was not sure what to do about the double line vs triple line.


You need to decide what your cutoffs are, then count it the same way every time.

Surely there is a SOP, otherwise me and you sharing a cell count would be pretty worthless. If I decided anything touching the line and you said anything 50% in, that could make a pretty huge difference . Or is it all within the margin of error anyway?
 
I was not sure what to do about the double line vs triple line.

Surely there is a SOP, otherwise me and you sharing a cell count would be pretty worthless. If I decided anything touching the line and you said anything 50% in, that could make a pretty huge difference . Or is it all within the margin of error anyway?

Ha funny, posting the same picture at the same time...:mug:
And yes sorry, I see the line question now.

There is a ton of variability of cell counts based on the counter, this is why the same person is used for all cell counts in a particular experiment. There is also a pretty large margin of error, which can be reduced when more cells are counted.
 
Seems to me that you would count cells touching the outer lines on the upper/left sides, but not count those touching the outer lines in the lower/right sides. However, I'm not a biologist and would never play one on TV.
 
I don't think it matters too much so long as you are consistent and the area you are counting is correct. That is, upper left/lower right whatever doesn't matter. It's the same as rounding numbers...you can use whatever system you like so long as you apply it consistently. There are varying conventions that people use (even/odd) but the most important thing is that it is unbiased and consistent.
 
Seems to me that you would count cells touching the outer lines on the upper/left sides, but not count those touching the outer lines in the lower/right sides. However, I'm not a biologist and would never play one on TV.

Hmmm, so if the cell is 100% outside the grid, but touching the outside line, you would count it?

Refer to the 2nd image I posted. Left side, bottom cell with a green check mark. That would be counted.

Basically, you are saying follow that 2nd image I posted and ignore the fact that there are only 2 lines instead of three.
 
I don't think it matters too much so long as you are consistent and the area you are counting is correct. That is, upper left/lower right whatever doesn't matter. It's the same as rounding numbers...you can use whatever system you like so long as you apply it consistently. There are varying conventions that people use (even/odd) but the most important thing is that it is unbiased and consistent.

I guess I was trying to learn what is consistent with the 'standard' method of counting a 2 line. I have found most everyone uses the same method for the 3 line, so I would feel confident that 2 people would look at the same picture and get the same count.
 
If you understand how the hemacytometer is ruled it is pretty clear what you should do but it is not clear from the diagram how one is ruled. In the attached photograph I drew a rectangle over one of the center squares and then dragged it to its current position. From this it is clear that it is the center line of the triplet that defines the square in which the cells are to be counted. If your rule is that any cell that touches, even if tangentially, a left or upper square boundary is considered within the square and that any cell that touches, tangentially or otherwise, a lower or right square boundary is not within the square then you will, assuming a uniform distribution of cells, count each cell on an interior boundary only once (in the square that is above or to the left of the line on which it lies) and include all cells that are on the larger (4 x 4 small cell) square boundary at top or left assigning them to the 4 x 4 array you are counting and exclude all cells that are on the right or lower boundary effectively assigning them to the 4x4 square to the right of or below the one you are counting. In this way, you will, on average, exclude as many as you include and, on average, will get the 'correct' count. The most important thing about this is to understand that it is the middle line that defines the boundaries. The rectangles and squares defined by the inner lines are smaller than the standard squares.

Hemacyt.jpg
 
Basically, you are saying follow that 2nd image I posted and ignore the fact that there are only 2 lines instead of three.
Yes, taking into account ajdelange's advice about the "ruling" of the slide, and adjusting appropriately to use the inner or outer line as your limit.
 
- Drags out scope and cheap Chinese Haemocytometer-

It looks as though the double line ones omit the inner set of lines, making all the blocks the same area, and not creating rectangles.

So one should treat the inner line on a double line as if it were the middle line on a triple line unit. Which actually makes the first graphic I posted correct also. Right?

2013-01-29_17-12-04_333.jpg
 
There is also a pretty large margin of error, which can be reduced when more cells are counted.

It turns out this is very simple to quantify. The coefficient of variation (multiply by 100 to get percentage error) is simply 1/sqrt(number of cells you count). This if you count 1200 cells total your Cv is 2.88%. If you only count 300 it's twice this.

More details at http://www.wetnewf.org/pdfs/hemocytometer.html
 
It turns out this is very simple to quantify. The coefficient of variation (multiply by 100 to get percentage error) is simply 1/sqrt(number of cells you count). This if you count 1200 cells total your Cv is 2.88%. If you only count 300 it's twice this.

More details at http://www.wetnewf.org/pdfs/hemocytometer.html

This is by far the absolute best explanation of how and why to decide when and where to count a cell that I have seen.

Thank you very much for taking the time to write that up.
 
Good points. Consistency is key in counting. The rule I use is that if it is touching the top or left box boarder it gets counted. If it is touching the bottom or right it does not. I also have an inexpensive hemocytometer and these lines are the ones that made the boxes the most square.

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. keep in mind that dilutions and homogenization are also critical. 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.

A little acidic acid helps with un clumping the cells, but seems to make staining difficult. Glycine work well for declumping and seems to work better with stains.
 
A little acidic acid helps with un clumping the cells, but seems to make staining difficult. Glycine work well for declumping and seems to work better with stains.

I'm fighting that right now. I got a sample of 002 from a 60bbl conical. It is extremely thick and clumped worse than cottage cheese! Based on Kai's site I did the first dilution with wort. That seemed to help some, but after 20 minutes the cells were still clumping. I dropped a few grains of PBW into the tube and agitated. That solved the clumping issue immediately! But Kai says PBW interferes with staining.
 
I'm fighting that right now. I got a sample of 002 from a 60bbl conical. It is extremely thick and clumped worse than cottage cheese! Based on Kai's site I did the first dilution with wort. That seemed to help some, but after 20 minutes the cells were still clumping. I dropped a few grains of PBW into the tube and agitated. That solved the clumping issue immediately! But Kai says PBW interferes with staining.

That's the same place I started when looking for ways to un-flocculate the cells! Kai has some very good information on his site. Top notch.

The other day I did a count on some WLP002. It clumps more than all the yeasts I have seen.

See here for pictures:
http://woodlandbrew.blogspot.com/2013/01/wlp002-lab-results.html

I did the viability count with MB diluted with water, and the cell density count with a 5% acetic acid solution. (actually just distilled white vinegar)


Glycine seems to do the trick and doesn't seem to interfere with the staining, although I haven't tried WLP002 with it. I just got some off Amazon here: Glycine
 
I did the viability count with MB diluted with water, and the cell density count with a 5% acetic acid solution. (actually just distilled white vinegar)

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
 
I am tempted to do the viability with them clumped and just kinda wing it. For a count though, the PBW worked like a charm. I added about 10 grains into a test tube with a 1:500 diluted sample. Gave it a couple seconds on the vortex mixer and it immediately went from chunky to just a nice haze.

I am redoing it now with the PBW in the first 1:10 dilution so hopefully the serial dilution gets more accurate.
 
I did the viability count with MB diluted with water, and the cell density count with a 5% acetic acid solution. (actually just distilled white vinegar)


Rough volume? Did you just do all your dilutions into vinegar instead of water?
 
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/2012/11/counting-yeast-cells-to-asses-viability.html

And White Labs sugestion for a Glycine MB solution:
http://www.whitelabs.com/beer/alkaline.html
 
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.
 
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.

..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.

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.
 
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.

Kai
 
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.
 
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?
 
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
 
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?
 
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.
 
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.

Remember what I said in #23. The standard deviation will be 1/sqrt(number of cells you count) so for 10,000,000 cells/mL and a large square volume of 0.0001 mL you'd expect to have 1000 cells in a large square and the standard deviation between large squares would be sqrt(1000) = 31.6. An estimate of your overall error is 100/sqrt(total_cells_counted) so that if, in this example, you counted one large square your estimated error is 100/sqrt(1000) = 3%. If you counted 3 large squares and got a total of 2998 then your estimated error is 100/sqrt(2998) = 1.8%. These are your counting errors only. If you are making dilution errors those add to the uncertainty. If you find variances between counts more than sqrt(cells_counted) then you should suspect that you are not properly doing the dilutions.
 
As a practical matter, generally how consistent are your counts?

Sometime I just need a ball park number and might only count 100-300 cells. These have an average box to box standard deviation of 6%.

If I take my time and do everything right and count 500 cells or more the standard deviation is 3% on average.

It is quite possible to have a very low standard deviation but have the numbers be way off. Recently I had some alcohol left in a tube from cleaning and didn't realize that it had not completely evaporated. My viability was much lower than expected.

It's important to know what to expect. This might be from taking day to day measurements or you running the tests in duplicate or triplicate.
 
Sometime I just need a ball park number and might only count 100-300 cells. These have an average box to box standard deviation of 6%.
1/sqrt(100) = 10%; 1/sqrt(300) = 5.77%. Ain't science grand.

If I take my time and do everything right and count 500 cells or more the standard deviation is 3% on average.

To cut your 300 count population sd in half you'd have to count 4*300 = 1200 cells. It's entirely possible that you get, by luck of the draw, a sample standard deviation less than the population sd but that won't happen very often. I question 3% in the long term. Yes, it can happen but it's unlikely.

It is quite possible to have a very low standard deviation but have the numbers be way off. Recently I had some alcohol left in a tube from cleaning and didn't realize that it had not completely evaporated. My viability was much lower than expected.
That's called a 'bias error'. If I snuck into your lab and replaced your 0.0001 mL heomocytometer with one that had a volume of 0.00009 mL all your counts would be 10% low but your variances would be little changed.
 
It might have to do with some post processing of the data. I throw out the high and low of both the dead and live cell counts. The data includes 21 cell counts of more than 500 cells.
 
It might have to do with some post processing of the data. I throw out the high and low of both the dead and live cell counts. The data includes 21 cell counts of more than 500 cells.

Do you have a reason to suspect that the high and low counts are biased somehow? If you don't, then you're really just hurting your estimate by doing this.
 
Remember that the mean isn't that robust an estimator. I'd say that if he has readings that are more than a couple of sigmas out he should toss them as the probability of getting a reading like that without some manipulation error like counting a square more than once, or not counting a square or doing a dilution wrong or waiting too long after pipetting before filling the slide (so that cells settle to the bottom of the pipet) is very small.
 
Good points. The equation I use in excel just tosses out the highest and the lowest counts regardless of the deviation. I'll have to rethink this.
 
Remember that the mean isn't that robust an estimator. I'd say that if he has readings that are more than a couple of sigmas out he should toss them as the probability of getting a reading like that without some manipulation error like counting a square more than once, or not counting a square or doing a dilution wrong or waiting too long after pipetting before filling the slide (so that cells settle to the bottom of the pipet) is very small.

Well, so that's why I was asking what the reasoning was behind dropping the highest and lowest counts. If there aren't 'outliers' then the total number of counted cells will give you the minimum-variance unbiased-estimator for a poisson count. Testing for outliers depends a lot on the actual data, but if the counts in each square are large enough that a gaussian assumption can be made then an ANOVA could give you some guidelines for dropping the counts for particular squares.
 
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