Berglund:Cell culture guide
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Understanding proper cell culture technique is key. (taken from http://www.sci.sdsu.edu/classes/chemistry/chem467l/mardahl/basiccell.html and moderately edited-AEM 2012)
Aseptic or sterile technique is the execution of tissue culture procedures without introducing contaminating microorganisms from the environment. In doing tissue culture work, 70% of the problems are due to a lack of good sterile technique. Microorganisms causing the contamination problems exist everywhere, on the surface of all objects and in the air. A conscious effort must be made to keep them out of a sterile environment. Because many and sometimes awkward manipulations are required for various techniques, tissue culture media used are often supplemented with antibiotics. Antibiotics do not eliminate problems of gross contamination which result from poor sterile technique or antibiotic-resistant mutants. Autoclaving renders pipettes, glass—ware, and solutions sterile.
Nutrient medium cannot be autoclaved. The compounds in nutrient medium are destroyed by the heat of autoclaving. Medium must therefore be sterilized by passing it through a sterile filter small enough in pore size to hold back bacteria and mycoplasmas (Millipore Sterivex - GS 0.22u disposable filter units). Here are some rules of thumb to follow to keep your medium, cultures, and glassware from becoming contaminated:
1. Wipe your work area and hands with 70% ethanol before starting.
2. Never uncover a sterile flask, bottle, petri dish, etc., until the instant you are ready to use it. Return the cover as soon as you are finished. Never leave it open to the environment.
3. Sterile pipettes should never be taken from the wrapper until they are to be used. Keep your pipettes at your work area. Sterile pipettes do not have to be flamed. Pipetting your cells with a hot pipette will kill them.
4. When removing the cap from a bottle, flask, etc., do not place the cap with the open end upright on the lab bench. Do not hold the opening straight up into the air. If possible, tilt the container so that any falling microorganisms fall onto the lip.
5. Be careful not to talk, sing, or whistle when you are performing these sterile procedures.
6. Do not draw from a different bottles with the same pipette. Because such a pipette has been exposed; the chance for contamination is too great; use sterile pipette for each bottle -- especially when pipetting medium.
7. Techniques should be performed as rapidly as possible to minimize contamination.
You may find yourself involved in a procedure which these sterile technique "rules-of-thumb" do not cover. Therefore, you must constantly be aware that microorganisms are everywhere and take proper steps to keep them out of your cultures. When first developing your aseptic technique you must always be thinking of sterility. Eventually it will become second nature to you. Mastering good aseptic technique will save you considerable frustration in the labs to follow.
Furthermore, the same principles for good aseptic technique also minimize biohazard risk to the investigator when infectious organisms or dangerous chemicals are used.
EYEBALLING THE CULTURES
Before doing anything with a culture, its general "health" and appearance should be evaluated. This can be done quickly and quantitatively by making the following observations:
1. Check the pH of the culture medium by looking at the color of the indicator, phenol red. As a culture becomes more acid the indicator shifts from red to yellow-red to yellow. As the culture becomes more alkaline the color shifts from red to fuchsia (red with a purple tinge). As a generalization, cells can tolerate slight acidity better than they can tolerate shifts in pH above pH 7.6.
2. Cell attachment. Are most of the cells well attached and spread out? Are the floating cells dividing cells or dying cells which may have an irregular appearance?
3. Percent confluency. The growth of a culture can be estimated by following it toward the development of a full cell sheet (confluent culture). By comparing the amount of space covered by cells with the unoccupied spaces you can estimate percent confluency.
4. Cell shape is an important guide. Round cells in an uncrowded culture is not a good sign unless these happen to be dividing cells. Look for doublets ordividing cells. Get to know the effect of crowding on cell shape.
5. Look for giant cells. The number of giant cells will increase as a culture ages or declines in "well-being." The frequency of giant cells should be relatively low and con—stant under uniform culture conditions.
6. One of the most valuable guides in assessing the success of a "culture split" is the rate at which the cells in the newly established cultures attach and spread out. Attachment within an hour or two suggests that the cells have not been traumatized and that the in vitro environment is not grossly abnormal. Longer attachment times are suggestive of problems. Nevertheless, good cultures may result even if attachment does not occur for four hours.
7. Keep in mind that some cells will show oriented growth patterns under some circumstances while many transformed cells, because of a lack of contact inhibition may "pile up" especially when the culture becomes crowded. Get to recognize the range of cells shapes and growth patterns exhibited by each cell line.
SUBCULTURING PROTOCOL FOR ANCHORAGE-DEPENDENT CELLS:
(Use Sterile Technique Throughout!!!)
1. Decant supernatant fluid from culture into a waste collection jar, applying the principles of sterile technique.
2. Rinse by adding 7 ml of cold versene buffer to the T-25, then decant into waste collection jar.
3. Rinse again by adding 3 ml of cold trypsin to the T-25, then decant into waste collection jar. This will remove the proteins present from the old media that tend to decrease the ability of the trypsin to act on the cells.
4. Now add 3 ml of cold trypsin and examine under low magnification for 3-7 minutes with the phase contrast inverted scope. When it appears that most of the cells have rounded up, but have not yet completely detached, whack the T-25 gently. Then immediately return to the hood and pipette the cells up and down several times to help break up the clumps of cells.
5. Remove 0.3 ml of cells in trypsin and place in microfuge tube for cell counting.
6. Add 9.1 ml fresh MEM media to three new flasks (if you are doing a 1:3 split). Using the same pipette, and as continuation of the same process, draw up the cell suspension and quickly dispense a 0.9 ml aliquot into each T-25 flask containing 9.1 ml of media in the new flask giving a total volume of 10 ml.
7. Incubate at 34C. Monitor periodically, beginning 30-45 minutes after inoculation. Rapid attachment (within 1 hour or so) is indicative, but not proof, that all went well.
8. Start making observations, such as those described in the handout "Eyeballing the Culture."
HEMOCYTOMETER COUNTING AND CELL VIABILITY
The importance of accurate enumeration of cells at time of inoculation, at the termina—tion of an experiment, and throughout the course of many experiments is self-evident. Cell enumeration with a hemocytometer is the most widely used method, and it continues to have a place in all laboratories, including those equipped with electronic cell counters (like us). This laboratory exercise will serve to introduce the use of a hemocytometer as adjunct to a subculturing exercise. The following review information may assist you in proceeding through the exercise.
1. Cell populations are usually expressed as # cells/ml or # cells/culture.
2. When viewed with a compound microscope with a 10x ocular and 10x objective (total magnification 100x), one large square of the hemocytometer (1 mm x 1 mm) will fill the field.
3. Each of the four large corner squares and the large center square of a Neu—bauer type hemocytometer usually are counted. When the cell count is low (less than 10 cells/square) all nine squares should be counted.
4. Each large square measures 1 x 1 mm and is 0.1 mm deep. Hence, each large square has a volume of 0.1 mm3 or 0.001 cm3 or 0.0001 ml (10-4 ml).
5. The final calculation takes into account:
a. how you wish to express your count (cells/ml or cells/flask)
b. the dilution (# of mls of saline or dye or medium into which your cells have been suspended
c. the number of squares counted
Hence the number of cells/ml of sample is calculated as follows:
total # cells counted x dilution x 104 number of squares counted
1. Proceed to make a 1:3 split of a culture following the procedure described above. In order to determine the number of cells which were harvested from the flask.
2. Aseptically remove 0.5 ml of cell suspension in trypsin and place in a microfuge tube on ice. Sterility need not be main—tained.
3. Remove a few drops of the cell suspension with a Pasteur pipette and load both chambers after putting the hemocytometer coverslip in place.
4. Count the total number of cells in each of the four corner and central squares.
1. Count 10 squares and then multiply that number by 1,000 and that gives us the number of cells per ml from your solution of cells in trypsin.
N = Sum of 10 Squares X 103
2. This gives you the number of cells per ml, you should now adjust this for any dilution you are doing. That is, you have a T-25 flask that you have just added 3.3 ml of trysin to, this is our total volume of cells and you have calculated the number of cells per ml. You have removed 0.3 ml (or whatever volume you have used), so you have 3.0 ml of cells left that you are going to add to a T-25 with 9.0 ml of fresh media. This gives you a 1:10 dilution of the cells you have just counted.