Hemocytometer protocol


Counting cells can’t be done directly from the flask because you don’t have an order of magnitude of the volume of cells you are seeing. You also don’t have a specific area that you can count every single time in the same way, and you don’t know if the portion you have counted is statistically significant or not. What I’m trying to tell you is: there’s a standard way.

It’s called hemocytometer. Hemo, for blood; cyto, for cell; meter, for measuring. So altogether: measuring blood cells.

“Hey but my cells are not blood cells!!”

Ok, ok, wait a sec. Now, when this hemocytometer was invented, the guy (Louis-Charles Malassez) was trying to count blood cells. But in the end, the purpose is to count cells that are in suspension, so any cells can be counted really (as long as you suspend them).

How does a hemocytometer work?

A hemocytometer is a square chamber carved into a piece of thick glass that has a specific depth. This big square has other little squares inside, following the pattern below:

Hemocytometer square

Each of the nine squares inside the big square is equal in surface. In turn, each of the sixteen little squares inside each of the four corner squares is equal in surface. What this tells you is basically, if you count cells in squares that are the same size, you will then be able to take an average of the counts you get, and that will be representative of the cell density of your culture. It is also important to note that if your cell concentration is very high, then you will go and count the cells in the smaller squares rather than the big ones.

“You’re going too fast, I’m lost.”

No worries, I didn’t get to the explanation of the protocol yet. Let’s get to the action first.

Counting cells with a hemocytometer

Before I start, I would like to recommend an iPhone app I have been using lately that has made my life a lot easier. It’s called HemocyTap (icon on the right) and it does most of the following itself: it helps you record your data, by acting as a double tally counter, and then performs all the calcs for you and saves the data so that you can review it and even send it to your email!

As I already said, you need to have a suspension sample of your cell culture to perform a cell count. If you are going to determine the viability as well, then you will need to add a viability dye (1:1 dilution usually works best). In any case, this will be your counting solution. Dilute further if your sample is very concentrated. If you need help choosing the basic materials to count cells with a hemocytometer, check our starter kit.

Now that you’re ready, take your hemocytometer, place a glass slide on top (making sure that it does not move; if it does, put some ethanol/water to stick it to the surface) and pipette 10-20μL of you count solution (dilute it if necessary, add viability dyes if you want to tell the difference between live and dead cells). Carefully introduce it in the space between the slide and the hemocytometer (it will go in by capillarity). When the space is filled, you’re done with that side. Repeat with the other big square of the hemocytometer if desired (improved accuracy). Now you’re ready to go to the microscope.

Place the hemocytometer under the microscope, in such a way that you see the first small square of the top big square in the middle of your field of view:

Hemocytometer first square

Start counting your cells. Yes, get your tally counters (or iPhone), feel like those air hostesses checking that everyone’s in, and don’t miss any of your cells! You should decide which two lines of your square you are going to discard. Because you can only count cells once, and some cells will be half-in half-out the square you are counting, it is common practice to choose on which two sides of the square you’ll be counting cells that are touching and on which ones you won’t. But keep these consistently throughout your count. For example:

Hemocytometer squares to count

Once you get the cell numbers in the first square, you go to the second (the one on the top right) and do the same thing. Remember, if you counted the cells on the top and right sides but not the ones on the bottom and left sides of the first square, then you must do the same thing with the second one. Keep counting squares until you have enough cells for your count to be statistically significant (big word! well basically, if you did another cell count, the difference between both counts would not be very big). It is recommended that you count at least 100 cells, so that should give you an idea of how many small squares you should count. Keep the counting symmetric (i.e., if you are going to count three small squares per big square, count the top left, bottom right and middle small squares in a big square). Also, if you are not using the app (they are saved for you while you count!), remember to note down your counts every time you finish with a square and reset your tally counter to zero, OR count all the squares and note down the total number, and then also write down how many squares you have counted, so that you can take the average. The advantage of counting all of them is that you don’t need to reset your tally counter to zero every time, but the downside of this is if you are not sure about one of the counts in one square, there is no going back, you’ll have to start again from the first square. This is not the case with HemocyTap, as it keeps the count of the cells in each square without you doing any extra effort of resetting the counters!I definitely recommend using HemocyTap, it saves you many steps and ensures you are not messing it up with the numbers. For a printable protocol, click here. Finally, go to Hemocytometer calculation for a detailed explanation on what to do with your data.

9 comments on “Hemocytometer protocol

  1. […] amount of growth medium (10-20mL should be enough) and take a sample to do the cell count (click here for an […]

  2. I’m having some problems training some folks here on counting techniques. We work with some very sticky cultures, and I’ve directed them to discard their counts if the left and right sides differ by more than 20%, as an estimate of CoV. Can you direct me to a resource that can do a good job of explaining this method? We will often need to repeat this count multiple times to get numbers that are within 20%, and for the sake of time, I’ve directed them to stop at three repeats, and go back and average all counts. Incorrect statistically, but should be reasonably close.

    • Hi Stu,
      I haven’t come accross this problem so far but I’ve done some research for you:

      CV of cell counts according to concentration in 19 cell lines

      in here they mention the coefficient of correlation (a good technique might be setting R^2 = 90% and checking if your calculated R^2 for that count falls lower than that), see page 3 table 1

      your RHS count falling in the range of the average count in LHS +- 2 times the standard deviation (see equation 1 here

      Let me know if that helped!



  3. Hi

    I am Vishwas, I am planning to work out a protocol to disassociate the brain tissue and make a cell suspension and stain it with DAPI stain and count the number of neurons to estimate the overall neurons in the brain tissue. There are some publications which came out in the last couple of years, unfortunately there is not enough information to start something from scratch, at least people like me who don’t know much about the haemocytometer.

    Currently, I had fixed brain tissue, I mechanically mashed up and centrifuged. But for some reason I dont know how to make up/ determine the intial volume?. I understand that I need to have original volume and concentration, but I dont know how I make up the original volume and concentration. I appreciate if you could help me out, or point me out in the right direction to find a good protocol, which I can follow.

    Looking forward to hear from any one of you.

    Thank you

    • Hi Vishwas,
      In my understanding, the initial volume is the volume of PBS or counting fluid you resuspend you cells in. Let’s say you have dssociated your cells and you now have a pellet after centrifuging, then just remove the supernatant, add a known volume of PBS or whatever solution is appropriate for your cells, mix and count.
      For a video protocol on how to dissociate brain cells, see here.
      For a hemocytometer protocol for brain cells, see the last slide of this presentation.
      For a protocol on how to use DAPI with brain cells and a fluorescence microscope, see here.

      If you combine all three, you should be able to design your own protocol for processing and suspending brain cells, and counting DAPI stained cells with the hemocytometer

      Hope that helped! Cheers,


      • Thank you Maria for the details, I will try out some of these and let you know how it goes.

        • No prob! Good luck :)

  4. How is it that the solution is diluted with the dye? if the cells are taking in the dye then the dye cannot be the diluent because its volume is being altered.
    I was always taught to dilute with saline or phosphate buffer and then add dye. the dye was never used as the diluent.
    am I missing something here?

    • Hi Heather,
      You do have to dilute with PBS if your cell density is very high (prior to adding the dye) and then you add the dye. You need to measure how much dye you’re adding, because you’re altering the volume of your counting solution. Unless you’re adding a drop in 1mL+ of counting solution (which would be wasting a lot of cells – hence the use of smaller volumes), the drop in smaller volumes (such as 10uL) will matter and you do have to take that into account. The way you do that is you measure how much dye you add (say, 10uL) and account for the dilution you have created due to the addition of the dye (in this example, 10uL:10uL or 1:1 dilution).

      Was that clear? Let me know otherwise what are your specific doubts :)



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