This is an experiment you will do twice.  The first time is to become acquainted with working with a cell culture and the procedures for enumerating and then planting cultures.  The second time will be to test the skills you've learned from the first trial and improve on your skills.  First, however, some introductory material to guide you in working with materials with which you may not be familiar.


Hemocytometer Counting - This is probably the most convenient and practical method of counting cells in suspension culture or in dispersed monolayer culture. The hemocytometer consists of two chambers, each of which is divided into nine 1.0 mm squares. A cover glass is supported over the chambers and cell suspensions are introduced under the cover glass. The hemocytometer is placed on the microscope stage and the cell suspension is counted.

In counting cells, unless there is a valid reason to do otherwise, it is only useful to count those cells which are likely to be viable. Why count dead cells? A number of stains have been employed to distinguish between viable and nonviable cells. Trypan blue, Erythrosin B, and Nigrosin, are excluded by the membrane of "viable" cells. In dead or damaged cells, the stain enters the cytoplasm and the cells take up the stain. Although such methods have been seriously questioned, they have the virtue of simplicity. If more than 20% of the cells are stained, the cell suspension is most likely not to be a viable one.

Hemocytometer counts are, however, subject to the following sources of error:

(1) Non-uniform suspensions

(2) Improper filling of chambers

(3) Failure to adopt a convention for counting cells in contact with boundary lines or each other

(4) Statistical error

(1) Non-uniform suspensions: It is assumed that the volume of cell suspension placed into the chamber represents a truly random sample. This will not be a valid assumption unless the suspension is monodisperse and free of cell clumps. Distribution in the hemocytometer chamber depends on the number of particles, rather than particle mass. Thus, cell clumps will distribute in the same way as single cells, distorting the final result. Unless 90% or more of the cells are free from contact with other cells the count should be repeated with a fresh sample from the original culture. Clumping can be minimized by keeping the suspension in an ice bath in plastic tubes, and by using a diluent without calcium and magnesium. Always mix thoroughly before sampling!

(2) Improper filling of chambers: In order to fill properly by capillary action, a hemocytometer chamber must be scrupulously clean and this also applies to the Pasteur pipet used to fill the chamber. New pipets may be dry-heat sterilized. In a laboratory where the hemocytometer is in regular use throughout the day, following rinsing with water after use, it, and the coverslip, may be placed in a small beaker containing soapy water so that it is always ready for its next use.  The chamber and cover slip are cleaned first with distilled water, then by 70% ethanol and wiped dry with a Kimwipe. If the cell suspension does not flow in and fill the chamber immediately and smoothly when the drop from the pipet is placed in the depression under the coverslip, the chamber is dirty and should be recleaned.

(3) Failure to adopt a convention for counting cells in contact with boundary lines or with each other: See the discussion accompanying Figure 1.

FIGURE 1: One-half (i.e. one chamber) of a hemocytometer with Neubauer ruling. This diagram is what you would see when looking down onto the hemocytometer under the microscope at low power (i.e. 4-10X). There are two chambers in the hemocytometer. This diagram depicts one of the two chambers. A coverslip sits on top of the hemocytometer enclosing both chambers, creating a space into which is loaded the cell suspension. There is a "V" or notch at either end which is the place where the cell suspension is loaded into the hemocytometer. Here you can see the entire chamber with its nine 1.0mm x 1.0mm large squares separated from one another by the triple lines. The area of each is 1mm 2. Within each larger 1mm2 square are 16 smaller squares which are there to help orient you during counting to help avoid counting a given cell more than once. Some cells can settle on the border gridlines and so it becomes difficult to decide whether or not to count such a cell. Is it in or is it out? In order to determine what to count and what not to count, concerning a cell on a boarder, you should develop a convention in which you do not count half of the cells that touch a boarder. For example you may decide not to count cells if they touch the bottom or right boarder or the top and left boarder. You can decide on your own convention but whichever you choose, you MUST be consistent.

You count the total number of cells found in five (5) of the large squares, by counting the cells in the four corner squares (i.e. those divided further into sixteen small squares) and the one at the very center of the chamber (here shown with a circle).

(4) Statistical error: With careful attention to detail, the overall error can be reduced to between 10% and 15%.


Each of these chambers is divided into nine exactly equal squares each 1mm x 1mm. Of the nine squares in the top chamber and the nine squares in the bottom chamber, only five are used to count those cells which have landed in them. These five squares are the upper left and upper right, the lower left and lower right and the one directly in the center. Furthermore, there is the convention concerning which of the cells in the square are to be counted. Referring to the diagram of the hemocytometer chamber, you will note that the four corner squares are each bordered on all sides by triple lines. A convention one can use is that cells which land on, or touch, the outer perimeter lines ARE counted, while those which land on, or touch, the inner perimeter lines are NOT counted. In the central square, no cells touching any of the perimeter lines are counted. One should count the cells in the five squares of BOTH the upper and lower chambers for the most accuracy although, in many laboratories, for convenience, only the five squares of one of the two chambers are counted.


1.  Each of the nine squares making up the hemocytometer chamber is: 1mm x 1mm = area of 1mm2

2.  With the coverslip on the chamber, the coverslip sits 0.1mm over the chamber. Therefore, the volume of each square, contained under the coverslip is:
1mm x 1mm x 0.1mm = 0.1mm3 or 10 -4cc
(10mm = 1cm; 1mm = 0.1cm; 1mm 3 = 10-3 cubic centimeters (cc) or ml; 0.1mm 3 = 10-4cc or ml (the vol. of 1 square)]

3. Since the volume of one square = 10-4ml, we take the average number of cells counted (the total number of cells in the upper left and right and bottom left and right and center, divided by 5, the number of squares counted) and multipy by 104 which = # cells/ml in the suspension used to charge the hemocytometer. If this represents the number of cells per ml of the resuspended cells from the culture flask, that is the correct number per ml.  If, however, the suspension were diluted prior to counting, then then one would also multiply by the dilution factor to get the # cells /ml in the culture flask .

Two calculation conventions:
A. Counting the ten squares of the hemocytometer (i.e. both chambers):
    1. Avg. # cells counted x 104 = # cells/ml (x diln. if any)
    2. Total # cells counted x 103 (because here we counted TOTAL cells not ending up with the number per square) = # cells/ml (x diln. if any).

B. Counting 5 squares only (i.e. one chamber):
    1. Avg. # cells counted x 104 = # cells/ml (x diln. if any)
    2. Total # cells counted x (2 x103) = # cells/ml (x diln. if any).

Average count of 10 squares = 50 cells. Therefore: 50 x 10,000 = 5 x 10 5 cells/ml
Total count of 500 cells in 10 squares. Therefore 500 x 1,000 = 5 x 10 5 cells/ml
Total count of 250 cells in 5 squares. Therefore 250 x 2,000 = 5 x 10 5 cells/ml


The purpose of this laboratory exercise is to acquaint you with some of the fundamentals of culturing cells in vitro.

In this exercise you will: II. MATERIALS

1.  Cultures of KB cells--1 T-75 flask and 1 T-25 flask at or near confluence for each person in each group. (In the future, each "lab" will have, and must maintain on a regular basis, a particular cell culture.)

2. Culture medium: EBME90 ABS10, 100mg/ml gentamycin

3. Pipettes - 1 ml, 5 ml, and 10 ml.

4. T-25 culture flasks - 9 per student.

5. Hemocytometer.

6. Hand tally counter.

7. Medium disposal system.

8. Trypsin (0.05%)/Versene (0.02%) solution.

9. Non-sterile test tubes for cell counting.

10. Pasteur pipettes.

11. Centrifuge tubes (15.0 ml).

12. Calcium and magnesium-free phosphate buffered saline (CMF-PBS).

13. Microscope.

14. Test tube racks.

15. Trypan blue.


To trypsinize cells grown in a monolayer:

  1. Each person in each "lab" will have a KB cell culture. Examine its macroscopic appearance and observe the culture microscopically to determine cellular morphology, appearance of the culture (whether or not cells look "healthy" or are granular, sloughing, etc.) and whether or not confluency has been reached. Record your observations.
  2. Thaw (where necessary) and warm the trypsin-Versene, CMF-PBS and medium solutions to 37C.
  3. Aseptically remove the spent culture medium as completely as possible into the vacuum flask.
  4. Add 5.0 ml of warm CMF-PBS to the side of the flask opposite the cells (Why?) and gently tilt flask to wash the monolayer and adjacent sides of the flask, taking care not to splash the solution into the neck of the flask.
  5. Remove the CMF-PBS into the vacuum discard flask.
  6. Add 3.0ml (for a T-25 flask, 5.0ml for a T-75 flask) trypsin/versene solution to the side of the flask opposite the cells then rotate to bring the entire monolayer in contact with the solution.
  7. Remove most of the trypsin-versene (that is, when aspirating off the solution, you do not have to be quantitative and remove every drop.), bring the residual solution in contact with the monolayer and then place the cultures in the incubator until the monolayer detaches. (It is helpful to observe the cells frequently during this stage because you will then have the opportunity to observe how the cells come off the plastic.) You can tell when this stage is reached by holding the flask up to the light and observing that the monolayer has become opaque. By bringing the residual solution up and over the disintegrating monolayer and shaking or gently hitting the flask against your hand, you can see the monolayer break up and flow down the plastic.
  8. When all of the cells have detached, add 5.0 ml of growth medium to the T-25 flask; 10.0 ml to the T-75 flask (this time to the very side on which the cells were attached) so as to wash the cells down to the bottom of the flask. With the same pipette, draw up the cell suspension and expel the solution back against the side or bottom of the flask. Try not to discharge the suspension into the liquid in the flask or foaming will occur. The pulling up of the solution and then blowing it back into the flask is known as trituration. Continue this trituration process 5-7 times in order to obtain a well dispersed cell suspension. Be sure the cells are well dispersed at this point by examining under the microscope and looking to see if the cells are separate or in clumps. If a significant number of cell clumps or aggregates are present, resume trituration and reexamine.
  9. Place 0.5 ml trypan blue into a small nonsterile tube. Then place 0.1 ml of cell suspension into it. ( What level of dilution of the cells does this effect?) Triturate with a Pasteur pipette and charge one side (one chamber) of a hemocytometer. Triturate once again and charge the other chamber of the hemocytometer. Count the cells in both chambers. The total number of cells in both chambers x the dilution in trypan blue x 1000 (hemocytometer factor used when counting both chambers [x 2000 if counting only one chamber] = no. of cells/ml of the cell suspension in the flask. Multiplying the number of cells/ml of the cell suspension by the number of mls of the cell suspension = the total number of cells in the monolayer of the bottle trypsinized for this count. If the hemocytometer count exceeds 2-5 x 105 cells/ml (20-50 cells per square of the hemocytometer) a further dilution of the cell suspension in CMF-PBS should be made because counting larger numbers of cells than this tends to be inaccurate. Thus, if you routinely dilute in the trypan blue to the same degree (i.e. 0.1ml cells + 0.5ml trypan blue) and always count one chamber of the hemocytometer, you would multiply the number of cells in the five squares by 2,000 x the dilution to get the number of cells/ml of the cell suspension
 10. A. From the T-75 flask, label and seed three T-25 flasks each as follows*:
                                                                 Label: 3 Flasks #1, 3 Flasks #2 and 3 Flasks #3  
3 #1 Flasks 
3 #2 Flasks 
3 #3 Flasks 
Label = 3 x 105 cells 
Label = 6 x 105 cells 
Label = 1 x 106 cells 
Label = 24, 48, 72hrs.
Label = 24, 48, 72hrs.
Label = 24, 48, 72hrs.

       B. From the T-25 flask, seed 1 T-75 flask with 1.5 X 10 6 cells. This flask will be used for the repeat of this experiment next week.
*Determine the volume of cell suspension to be added to each flask and then add the cells. Next, add the correct amount of medium to each flask to bring the total volume to 5 ml and agitate the flask to disperse the cells. Be sure each flask is appropriately labeled with the number of cells added, the name of the cell line the date and sample time and your initials.

11. Incubate the cultures in your CO2 incubator with the caps loose if the caps are solid, or tight if they have a filtered cap. (Under what conditions might the caps have to be kept tightly closed?). Refrigerate any remaining liquids (i.e. medium, trypsin, etc.)

12. Observe and count the cells after 24, 48, 72 hours.

13. Record the following at each interval: a. Appearance of the cells (presence of floating cells, morphology of cells, intracellular granulation).

  1. Estimate the percentage of the growing surface occupied by the cells each day by examining under the microscope. Then count the cells to determine the number of cells in the flask. Prepare a table demonstrating the growth of the cells over time.
  2. At the end of the 72 hour period, answer the following question: What cell number would you seed/plant, if you wanted to use the cells at a 3/4 monolayer in two days? In one week?


 A. Freezing Cells

The objective in freezing cells is to do so relatively slowly (i.e. approximating 1o C per minute) in the presence of the cryoprotectant DMSO (dimethylsulfoxide), to minimize the formation of ice crystals inside the cytoplasm. At that rate of freezing, most of the ice crystals form outside of the cytoplasm. However, as the ice crystals form outside of the cell, the intracellular water moves to the outside leaving the cells in a plasmolyzed state. Without the cryoprotectant, which binds water keeping a sufficient amount inside the cells to maintain cell viability, the cells would not survive the process.

1. Remove 2 x 106 cells from the T-75 suspension of cells remaining after completing the setup of the exercise above and centrifuge, at 600 rpm for 5 min.

2. Remove (as demonstrated) the supernate and replace with 2 ml of 1x cryoprotective medium (EBME85, DMSO5, FBS10 , 100 mg/ml Gentamycin).

3. Resuspend cells and add 1 ml per freeze vial.

4. Imediately place sealed vials in freezing apparatus and adjust depth of vials properly as demonstrated.

5. Place the freeze unit in a nitrogen freezer that is at least one half full of liquid nitrogen.

6. After 3 hr (or overnight) place the vials on canes and place the canes in the appropriate freezer cans.

    B. Recovery of Frozen Cells - at the next lab period
    1. Remove a vial from the nitrogen freezer and rapidly thaw at 37C.
    2. Using a pipette carefully layer the contents of the vial onto 10ml of prewarmed, properly pH’d medium contained in a 15ml centrifuge tube. Centrifuge at 4-600 rpm for 5 minutes. Remove the medium carefully so as not to disturb the pelleted cells at the bottom and resuspend the pellet in the quantity of prewarmed medium to be used for the size flask in which the cells will be planted. (5ml for a T-25; 15ml for a T75, etc.)
    3. Incubate the cells in a CO 2 incubator. Once the cells have attached to the flask, usually within 4-6 hours but usually by the next day, one can remove the medium containing any dead cells and possible residual DMSO and add fresh growth medium. Depending upon the cells you are using this may or may not be necessary--examine your cells and decide if they are exhibiting signs of toxicity.

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