Cell Cultures

Introduction

A major advance in our knowledge of cells came about with the ability to maintain them in continous culture. Prokaryotes have been cultured for a relatively long time, but eukaryotic cultures were first accomplished in the early 1900's (Harrison, 1 Carrel 2), with major advances being made only in the past two or three decades

The procedures employed during this exercise will examine several types of prokaryotic cultures as well as a eukaryotic suspension culture, and establish a culture of embryonic fibroblasts. The prokaryotic cultures and the eukaryotic suspension culture are "established" while the embryonic culture will be a "primary" culture. Chick embryos are used for this latter procedure because of the relative ease of culture, ease of obtaining embryos, and relative simplicity of their nutritional needs. Highly differentiated cells would be more difficult to establish and in some cases not possible at all (given current technology).

Prokaryotes

Prokaryotes are cells without nuclei and are generally considered to be more primitive than eukaryotes (cells with true nuclei). Typically, prokaryotes are easier to culture in a laboratory because most of them have less stringent nutrient requirements. For this lab we will utilize bacterial cultures grown in nutrient agar, an environment in which human pathogenic organisms are extremely unlikely to grow.

Bacteria may be examined by observing the living, unstained microbes in a wet mount (phase contrast or dark field illumination), by observing dead cells stained with dyes under bright field illumination, or by observing cells prepared for electron microscopy. Our procedures will use "fixed" (i.e. dead) cells stained for standard light microscopy.

When cells are grown, they will have specific growth characteristics, depending on the media, the temperature and the strain of cells utilized. For bacteria (and some algae, fungi and other eukaryotic tissue culture) it is possible to measure the growth of cell populations by calculating cell number or mass. With modern equipment and the proper computer software, this can be a completely automated analysis, that would include the specific morphological data as well (size, shape and density of colonies or individual cells).

Since bacteria are small, they are difficult to count through direct visualization, but can be counted if one makes an assumption that each bacterium is capable of forming an individual colony. The mass of a bacterial suspension can be deduced from the optical turbidity of a suspension.

Eukaryotes

By contrast to the simple broth cultures of E. coli, the nutrient requirements of even the simplest eukaryotic culture are complex. Refer to Table 12.1 for a comparison of the ingredients of Nutrient Broth and Minimum Essential Media (MEM), a typical eukaryotic media. Eukaryotes also require supplemental sources of materials, most often in the form of blood serum. Fetal calf serum is used extensively for this purpose, since it is readily available, and the fetal nature of the serum limits the presence of antibodies, which might negatively effect cell growth.

Our first procedure will involve the simple transfer of an established culture from a suspension culture. An aliquot will be removed from a commercially available cell line and transferred to prepared transfer vessels. Each day students will observe these transfer cultures with an inverted phase contrast microscope, and remove aliquots for cell counting with a hemacytometer. Simultaneously, they will check on the viability of the cells through a dye exclusion technique.

The second procedure will be somewhat more complicated. Students will remove chicken embryos from the egg, trypsinize them to disaggregate the cells, and transfer the resulting cells to culture flasks. This procedure establishes a primary culture, or one that is a first generation growth from "in vivo" cells. Established culture lines are the result of long term selection for cells capable of growth under "in vitro" conditions. As such, they are more consistent clones, but often have genetic and structural alterations that differ significantly from the starting cell lines.

Table 12.1 Eukaryote and prokaryote culture media

As the cells grow in culture, we can observe three distinct phases. The first is a Lag Phase, usually no more than 1-2 days in length, and during which there is little or no increase in cell number. During this time, the cells are "conditioning" the media, undergoing internal cytoskeletal and enzyme changes and adjusting to the new media.

This is followed by a Log Phase. During this phase the cell number increases exponentially. This growth will continue as long as there is sufficient nutrient to support the increasing cell number. Eventually some critical nutrient will become limiting, however.

The final phase is the Plateau Phase. During this phase the number of cells remains constant (although not necessarily viable). Eventually, of course, the cells will die unless subcultured or fresh media is added. The final procedure in this laboratory exercise involves establishment of a primary culture from a chick embryo. Here, the cells are not established in culture, but must be disaggregated (detached from each other) and placed into a foreign environment (the culture media). Disaggregation in embryos is reasonably easy, but depends on the enzymatic dissolution of the cells glycocalyx, and the disruption of many plasma membrane structures and chemical elements. Consequently, the cells will take a longer time before they grow (that is, there is a long Lag Phase), and the selection process will favor cells which grow in contact with the culture vessel surface.

The cells will continue to grow in contact with the vessel and give rise to a "monolayer" culture. The cells will cease to divide when they reach confluency; they are said to demonstrate contact inhibition.

Consequently, growth curves are not measured by removing aliquots and determining cell concentration, but are measured by the density of cells growing on the vessel surface. This is accomplished through the use of an ocular grid inserted into an inverted phase contrast microscope. Cell density (cells/cm2) is then plotted on a log scale against time in culture.

Exercise 12.1 - Aseptic Cell Transfers

Level I

Figure 12.3 Flaming a wire loop and removing cap

Materials

• Bunsen burner
• Wire loop
• Petri plate or broth tube
• Bacterial culture
• Microscope slides
•  

Procedure

1. Pick up the inoculating loop and hold it pointed down into an open flame until the loop glows red. This process sterilizes the loop of wire and is known as "flaming" the loop. It will result in a sterile loop and will not contaminate your stock culture. If there are liquids already on the loop, the loop should be gradually placed into the flame to dry the loop. If the loop is rushed into the flame, the drop of liquid will splatter and spread bacteria over your work surface.

2. Pick up a broth culture in one hand, while holding the loop in the other. With the last two fingers of the hand holding the loop remove the cap from the culture and gently flame the top of the test tube (do not overheat). Insert the flamed inoculating loop into the test tube until it is submersed in the broth. The loop should be allowed to cool slightly before immersion. Retract a small quantity of the broth held in the loop and replace the cap on the culture.

3. Open the top of your transfer vessel (tube with water or nutrient agar) and flame the open top of the tube. Insert the loop into the tube and into the liquid in the tube. Withdraw the inoculating loop slightly from the liquid, blot gently on the inside of the tube and completely remove from the tube. Replace the top on the tranfer tube.

4. Immediately flame the inoculating loop.

Exersise 12.2 - Examination of Bacterial Colonies

LEVEL I

Materials

Petri plate cultures of various bacteria

Procedure

1. Obtain an established culture. Note that the bacteria grow in clearly defined groups, known as colonies. In most cases, each colony is the outgrowth from an individual cell, although they may overlap if excessive numbers of cells were plated.

2. Visually examine the individual colonies of bacteria and describe them according to the following characteristics:

Size. Pinpoint, small, medium, or large, based on the relative differences between the largest and smallest colonies seen.

Shape and Margins. Round, regular or irregular.

Elevation. Flat, convex or rounded, umbonate (flat on margins and raised in the center - like a fried egg), craterlike (with depressed center).

Consistency. Shiny or rough.

Color. Describe the color as accurately as possible, distinguishing between different types of gray or white, yellows, and red. If the pigment appears to diffuse into the surrounding medium, rather than coloring only the colony, it is a water-soluble pigment.

  3. Determine and record the identity of your colonies.

Exercise 12.3 - Gram Stain (+\-)

Figure 12.4 Typical bacterial shapes

Materials

• Colonies of bacteria from Exercise 12.2
• Toothpicks
• Crystal violet
• Gram's iodine
• 95% ethanol
• Safranin
• Microscopes with oil immersion 
•  

 Procedure

1. Before staining the individual colonies, you should first practice the technique by observation of the gram positive micro- organisms normally found in the gum linings of your mouth.

2. Use a clean toothpick to rub along the gingival crevices (area between tooth surface and gums) of your mouth.
Rub lightly!

3. Mix the scrapings with a drop of water previously placed on a clean slide, spread in a thin film over the center of the slide and allow to air dry.

4. Fix the smear to the slide by passing the slide (smear side up) quickly through a flame three times. If the slide is held directly in the flame, it will heat up too rapidly and break. The trick is to gently dry the smear without overheating the slide.

5. Place the slide on a staining rack. Apply the stains on the fixed smear as follows:

• Flood the slide with crystal violet for 30 sec.
• Rinse with water.
• Flood with Gram's iodine for 60 sec.
• Rinse with water.
• Decolorize with 95% ethanol.
• Rinse with water.
• Counterstain with safranin for 60 sec.
• Rinse with water and blot dry (no rubbing!).
• Examine under oil-immersion objective lens. 3
•  
• Gram-positive bacteria retain crystal violet after washing with 95% ethanol, while gram-negative bacteria lose the purple dye after washing with 95% ethanol. The positive or negative reaction is a measure of the presence or absence of specific polysaccharide components of their cell walls. Safranin is used as a pink counterstain, so that Gram - cells can be visualized. In practice then, the distinction is made between purple cells (+) and pink cells(-).
•  

6. Determine the basic cell shape of the bacteria.

7. Add the information on Gram stain and cell shape to the work done in Exercise 12.2.

Exercise 12.4 - Prokaryote Cell Number by Dilution Plating

LEVEL II

Figure 12.5 Colony counter

 

Materials

• Broth culture of E. coli
• Tubes of nutrient broth (9.9 ml each)
• Nutrient agar plates
• Sterile transfer pipettes (1.0 ml)
• Quebec colony counter (Optional)
•  

Procedure

1. Obtain a broth culture of E. coli and carefully mix the contents to ensure equal suspension of the bacteria.

2. Obtain four test tubes each containing 9.9 ml. of nutrient broth. These will be used to produce a serial dilution of the stock culture.

3. Using sterile pipettes, remove 0.1 ml of well suspended cells from the stock culture and transfer these aseptically to one of the waiting test tubes of broth. This tube now contains a dilution of 1/100 or 10. Gently but thoroughtly mix the contents and label this tube at 10.

4. Repeat this process, but now aseptically remove 0.1 ml of culture from the 10 tube and place it into a new broth tube, which now becomes a 1/10,000 or 10 dilution. Mix the contents and label as 10.

5. Repeat this procedure twice more to produce respectively a 10 and a 10 dilution. Be sure to mix thoroughly and label each tube.

6. You should now have a serial dilution of the stock culture with tubes at 10, 10, 10, and 10 . The original stock culture will almost invariably be too high a population for the next step, so you will only use the four dilutions that you have produced.

Using four separate petri plates containing 15 ml. of nutrient agar each and four separate sterile pipettes, transfer 1.0 ml of each dilution broth suspension onto the surface of a petri plate. Carefully label the plates and place them in an incubator for 24 hours at 37° C.

7. At the conclusion of incubation, remove each of the four petri plates and count the number of colonies formed on the plates. For proper statistical analysis, the plate containing between 30 and 300 colonies will give the most accurate results.

The colonies can be more easily counted by using a Quebec Colony Counter which allows proper illumination, a grid overlay and by slight magnfication of the plate surface.

8. Multiply the number of colonies counted by the dilution factor to obtain the population density of the original broth culture.

Notes

A growth curve can be established by repeating this procedure every two hours. Since the number of bacteria can be large, it will be necessary to plate cultures serially diluted. Count the number of colonies for each dilution and average the results.

Exercise 12.5 - Cell Mass by Measurement of Turbidity

LEVEL II

Materials

• Trypticase soy broth culture of E. coli
• Tubes of trypticase soy broth (9.9 ml each)
• Tubes of trypticase soy broth (5.0 ml each)
• Nutrient agar plates
• Sterile transfer pipettes
• Spectrophotometer and cuvettes 4
•  

Procedure

1. Obtain a trypticase soy broth culture of E. coli. Trypticase broth is better than nutrient broth, since it is inherently less light absorbent.

2. Prepare trypticase soy broth dilutions of 10 and 10 of an appropriate bacteria culture, as described in Exercise 12.4.

3. Now, for each of the two dilutions, set up a second series of dilutions, this time diluting by 1/2 each time. That is, transfer 5.0 ml of the 10 culture to 5.0 ml of fresh broth and mix thoroughly. Use 5.0 ml of this and transfer to 5.0 ml of fresh broth to produce a 1/4 dilution. Repeat for 1/8, 1/16, and 1/32 dilutions. Repeat the entire 1/2 dilution series for the 10 dilution.

You should now have twelve tubes, six for each of the 10 and the 10 dilutions. These are then diluted 1/2, 1/4, 1/8, 1/16, and 1/32.

4. Use 1.0 ml of each of the twelve dilutions and plate on nutrient agar plates to perform a colony count as in the preceeding section.

5. Set a spectrophotometer for 686 nm wavelength and be sure it is turned on and functioning properly. Adjust the dark current to 0% T.

6. Place a cuvette containing trypticase soy broth into the spectrophotometer and adjust the reading to 100% T. This is the blank for all subsequent measurements.

7. Transfer the remaining contents of the 12 dilutions to spectrophotometer cuvettes and measure the %T for each. Compute the absorbance for each sample. 5

8. Count the number of colonies formed for each sample after 24 hours. of incubation.

9. Plot the absorbance of each sample against the plate count for that sample.

Once this has been accomplished, you will have a value for computing the cell population number directly by measuring the absorbance of a suspension. This can be more readily measured on a continuous basis. To do this, grow suspensions directly in cuvette tubes and measure the A at timed intervals. Record a growth curve. By connecting a recorder to a spectrophotometer and keeping the chamber at 37° C, a continuous growth curve can be automatically recorded (assuming care is taken to ensure proper suspension of the bacteria throughout the time period involved). Lacking such sophisticated equipment, the tube can be removed from an incubator at intervals, gently suspended, measured for absorbance and returned immediately to the incubator. Plot cell growth against time to produce a growth curve.

Exercise 12.6 - Transfer of Eukaryote Suspension Cultures

LEVEL II

Materials

• Fibroblast suspension culture
• Tissue culture laminar flow hood
• Media appropriate to culture line used
• Disposable pipettes (10 ml and 1.0 ml)
• Disposable culture flasks
•  

Procedure

1. Obtain a culture of mouse fibroblast cells in suspension culture. This will be a simple culture with minimal requirements, and one selected for excellent growth characteristics. The transfer procedure will be similar to that for prokaryotes, with a few major changes. First, all transfers will be done in a tissue culture hood in order to maximize asepsis. Secondly, the cells will be transferred with siliconized pipettes rather than wire loops. The silicone prevents adherance of the cells to the glass wall of the pipette.

A tissue culture hood is a device that has air moving in layers and under positive pressure. Since the air is filtered, it contains minimal numbers of bacteria or fungal spores, and since it is under a positive pressure, those particles that are present are blown out of the hood. The layering prevents airborne organisms from settling on the work surfaces.

2. Arrange the materials in front of you, easily accessible through the opening of the tissue culture hood. Ensure that any alcohols and wrapping paper are kept clear of the bunsen burner. Pre-sterilize the hood before use, and use disposable sterile gloves. Loosen the cap of a tissue culture flask and the cap of a stock bottle of tissue culture media. 6

3. Insert the tip of a sterile pipette into the stock bottle and remove 10 ml of media. Transfer the media to the tissue culture flask.

4. Open the top of the suspension culture and use a sterile 1.0 ml transfer pipette to remove a 1.0 ml sample of the culture. Transfer it to the fresh media in the culture flask. Secure all caps that have been loosened.

5. Place the new cultures in an incubator at 37°C.

Exercise 12.7 - Viability Cell Count

Materials

• Suspension culture of cells
• Sterile transfer pipettes
• Stock 0.2% (w/v) Trypan blue
• Hemacytometer and microscope
•  

Procedure

1. Gently swirl a suspension culture to distribute the cells evenly. Aseptically remove a small sample (0.1 ml) of cells from the cultures. Place the sample in a separate test tube (it need not be sterile).

2. Dilute 4 parts of stock Trypan Blue with 1 part of 5X saline and add 0.1 ml of the diluted dye to your sample. Mix gently.

3. Set up a hemocytometer and cover slip. Immediately place a drop of the stain/culture combination on the hemocytometer (remember to use both sides of the hemocytometer) and wait one minute.

4. Observe the cells with low power microscopy. Count the total number of cells, and the number of stained cells.

5. Compute the concentration of viable cells per ml. of culture.

Notes

Trypan Blue is a stain that is actively extruded from viable cells, but which readily enters and stains dead cells. Therefore, the cells which are blue are dead. The difference between the total number of cells and the number of dead cells would be the number of viable cells in a given aliquot of your culture. Trypan Blue actually significantly overestimates the number of viable cells, but is sufficient for purposes of this lab.

Approximately 30% of the cells measured as viable with Trypan Blue will not be able to continue growth beyond a 24 hour period.

Exercise 12.8 - Computation of Transfer Aliquots

LEVEL II

Materials

• Suspension culture of cells
• Media appropriate to culture
• Culture flasks
• Transfer pipettes
• Hemacytometer and microscope
•  

Procedure

1. Obtain 3 transfer culture vessels. The vessels may already contain fresh culture media, or you may be asked to transfer your own.

2. Obtain a suspension culture and count the number of cells/ml of culture using the procedure listed in Exercise 12.7.

3. Compute the volumes of the suspension culture needed so that when added to 25 ml of fresh media, the final concentrations will be:

• 1 X 10 cells/ml
• 5 X 10 cells/ml
• 1 X 10 cells/ml
•  

For example: If you have 25 ml of fresh media, you will need to add 25 X 10 cells to obtain a final concentration of 1 X 10 cells/ml If your suspension culture contains 5 X 10 cells/ml, you will need to transfer 5 X 10 ml or 0.5 ml of culture to the fresh media (25 x 10 cells divided by 5 x 10 cells/ml).

Use the formula:

                                                    Number of cells to be transferred 
Aliquot to be transferred  =     -------------------------------------  
                                                         Culture concentration 

4. Seed three flasks each containing 25 ml of fresh media to a final concentration of 10 cells/ml, 5 X 10 cells/ml and 10 cells/ml.

5. Label and place your fresh cultures in the tissue culture incubator at 37° C.

Lay the culture flasks on their sides in order to maximize the air exchange surface of the culture and to prevent the cells from drowning. An alternative would be to incorporate mechanical shaking of some type.

Exercise 12.9 - Eukaryote Growth Dynamics

LEVEL II

Materials

• Suspension cultures set up from Exercise 12.8
• Sterile transfer pipettes
• Materials for viability counting (Exercise 12.7) '
•  

Procedure

1. After 12 hours, aseptically remove 0.1 ml from each of the three cultures, add 0.1 ml of trypan blue and count the total number of cells and the number of blue cells. Compute the number of viable cells/ml.

2. After 24 hours (from the time of seeding), repeat step 1.

3. Continue to repeat step 1 at 24 hour periods (i.e. daily) until there is no change in the number of cells/ml of culture.

4. Plot cell concentration on a log scale vs time of culture. Identify and label the Lag, Log and Plateau phases for your culture.

5. Select a period of time during the Log Phase and compute the doubling time for your culture. That is, the time required during the Log Phase to exactly double the number of cells/ml.

Exercise 12.10 - Establishment of a Primary Culture

LEVEL III

Materials

• Chick embryo (approximately 8 days old)
• 70% (v/v) ethanol for swabbing
• Sterile scissors, forceps and probes
• Sterile petri plates
• Phosphate buffered saline (PBS)
• Trypsin, cold sterilized in a 125 ml sterile erlenmeyer containing a magnetic stirring bar
• Minimum Essential Medium
• Fetal Calf Serum
• Clinical centrifuge with sterile capped centrifuge tubes
• Culture flasks
• Inverted phase contrast microscope (Optional)
•  

Procedure ⁸

1. Candle an 8 day old egg to ensure that it is alive. This is easily accomplished by holding the egg in front of a bright light source; the embryo can be seen as a shadow. Circle the embryo with a pencil.

2. Place the egg in a beaker with the blunt end up, and wash the top with a mild detergent, followed by swabbing with ethanol.

3. Carefully puncture the top of the egg with the point of a pair of sterile scissors and cut away a circle of shell, thus exposing the underlying membrane (the chorioallantois).

4. With a second pair of sterile scissors, carefully cut away and remove the chorioallantoic membrane, exposing the embryo.

5. Identify and carefully remove the embryo by the neck, using a sterile metal hook or a bent glass rod, and place the embryo in a 100mm petri dish containing phosphate buffered saline (PBS). Wash several times with PBS by transferring the embryo to fresh petri plates. After removal of all yolk and/or blood, move the embryo to a clean dish with PBS.

5. Using two sterile forceps, remove the head, limbs, and viscera. Be sure to remove the entire limb by pulling at the proximal end. Move the remaining tissues of the embryo to yet another dish and wash with PBS.

7. Mince the embryo finely with scissors and transfer the minced tissue to a flask containing PBS. Allow the tissue pieces to settle.

8. Remove the PBS with a sterile pipette and add 25 ml of trypsin, a proteolytic enzyme. Stir the solution gently at 37° C for 15-20 minutes.

9. Allow the larger, undigested tissue pieces to settle and decant the supernatant into an equal volume of Minimal Essential Medium (MEM) + 10% Fetal Calf Serum (FCS). FCS contains protease inhibitors which will inactivate the trypsin.

10. Centrifuge the cells in MEM at 1000 rpm for 10 minutes in a standard clinical centrifuge. Remove the supernatant and resuspend the pellet in 25 ml of fresh MEM + 10% FCS.

11. Remove 0.1 ml of the culture and determine cell concentration and viability as directed in the previous section.

12. Seed two 25 cm plastic culture flasks containing 25 ml of MEM + 10% FCS to a final concentration of 10 cells/ml.

13. Label and place your cultures in the tissue culture incubator at 37° C and examine daily for cell density and morphology.

14. Note any changes in the color of the media. Tissue Culture media has a pH indicator (Phenol Red) added in order to check on the growth of cells. The media initially is a cherry red (with slight blue haze) and turns orange and then yellow as the cells grow, thereby reducing the media. Should this color change occur within 24 hours, the culture is most likely contaminated and should be disposed of.

15. Examine the cultures using an inverted phase contrast microscope. This will allow observation of the cells without opening or disturbing the growth.

16. Make cell density determinations at 10 X magnification using a square ocular grid, as explained in Chapter One for the determination of area.

17. Plot the cell density on a log scale vs. time of culture.

18. Diagram the shape of the cells at each phase.

Notes

The cultures will develop differently than the suspension cultures. The viable cells will grow out of the trypsinized pieces of tissue and will remain in contact with the bottom of the culture flask. They will continue to divide and migrate until the entire bottom of the flask is covered with a single layer of cells (contact inhibition and the formation of a monolayer).

Chapter 12: Cell Cultures - Endnotes

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1. Harrison, R.G. (1907) Observations on the living developing nerve fiber. Proc. Soc. Exp. Biol. Med. 4:140-143.

2. Carrel, A. (1912) On the permanent life of tissues outside the organism. J. Exp. Med. 15:516-528.

3. One of the continuing differences between microbiologists and microscopists is the lack of a coverslip when viewing bacteria. Convenience causes microbiologists to skip the process of placing mounting media and a coverslip on their slides. This causes microscopists to cringe at the thought!

4. If the cultures are to remain aseptic, the cuvettes can be sterilized and plugged. Alternatively, culture flasks with cuvette side arms can be used.

5. Absorbance may be read directly if the spectrophotometer is equipped with digital display. Absorbance is more difficult to interpret on an analog display.

6. It is assumed that the media has been pre-mixed, with serum and other additives put into the media. The media should be in small aseptic containers for student use.

7. From: Barbara B. Mischell and Stanley M. Shiigi. "Selected Methods in Cellular Immunity". W.H.Freeman & Co. San Francisco, 1980, p. 17.

8. Modified from Freshney, R. Ian. Culture of Animal Cells: A Manual of Basic Technique. Alan R. Liss, Inc. New York, 1983.