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  Section: Biotechnology Methods » Tissue Culture Techniques
 
 
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Culture and Maintenance of Cell Lines

 
     
 
Content
Tissue Culture Techniques
  Tissue Culture Methods
  Plant Tissue Culture
  Plant Tissue Culture (Cont.)
  Many Dimensions of Plant Tissue Culture Research
  What is Plant Tissue Culture?
  Uses of Plant Tissue Culture
  Plant Tissue Culture demonstration by Using Somaclonal Variation to Select for Disease Resistance
  Demonstration of Tissue Culture for Teaching
  Preparation of Plant Tissue Culture Media
  Plant Tissue Culture Media
  Preparation of Protoplasts
  Protoplast Isolation, Culture, and Fusion
  Agrobacterium Culture and Agrobacterium — Mediated transformation
  Isolation of Chloroplasts from Spinach Leaves
  Preparation of Plant DNA using
  Suspension Culture and Production of Secondary Metabolites
  Protocols for Plant Tissue Culture
  Sterile Methods in Plant Tissue Culture
  Media for Plant Tissue Culture
  Safety in Plant Tissue Culture
  Preparation of Media for Animal Cell Culture
  Aseptic Technique
  Culture and Maintenance of Cell Lines
  Trypsinizing and Subculturing Cells from a Monolayer
  Cellular Biology Techniques
  In Vitro Methods
  Human Cell Culture Methods

Purpose
Cell and organ cultures are used to maintain living animal cells and groups of cells outside the body (in vitro). With separate, living cell cultures, it is possible to see and study the behavior of animal cells in greater detail than when they are in the animal (in vivo). Cell culture also frees the cells from some of the controls that normally regulate their activities. Cells, or tissues, are kept alive for varying periods of time, at times undergoing repeated divisions over many generations.

Cells grown and cultured for study have been taken from a wide variety of species, such as humans, monkeys, mice, dogs, cats, frogs, insects, fish, and many others. The cultures have come from a number of organs—heart, lungs, liver, kidney, blood, skin, etc. In cancer research, it is common practice to grow cells from normal and cancerous tissues to compare their properties. In fact, cellcultures have become one of the best means of testing potential anti-cancer drugs; utilizing cell cultures is more cost-effective and faster than experiments using animals and, with this method, isolated human cancerous tissue can be tested.


General Principles and Techniques
Tissue fragments used in the preparation of cell cultures must be handled with care in the laboratory to avoid microbial contamination; sterile, or aseptic, technique must be employed at all times. All instruments, culture vessels, etc., that come in contact with the cells or medium must be sterile. The tissues are kept at 37°C and suspended in a physiologically balanced salt solution. Addition of a small amount (1%–15%) of blood serum helps protect the cells during these preliminary manipulations. Antibiotics may also be added as well as pH indicators, such as phenol red. Tissues are cut into very small fragments, called explants, which are put into vessels and bathed with nutrient medium. Sometimes the fragments are attached to the surface of the vessels by opposite charge attraction or with blood plasma, which is then allowed to clot. In the case of the cultures of this type, cells migrate from the explanted tissue into the medium and undergo division to produce a “halo” of outgrowth around the original tissue.

Cell cultures are also started by treating tissue fragments with chemicals (enzymes such as trypsin, or chelating agents such as versene) to dissociate them into a suspension of single cells. These cells are then placed in a nutrient medium and a portion of the suspension is put into a suitable culture vessel. The culture vessel is incubated without being disturbed, which allows the cells to settle out from the suspension, attach to the wall of the vessel, and grow. The complete sheet of cells covering the vessel wall is called a monolayer.

“Cell lines”, which are capable of continuous growth, are usually grown as monolayers. To transplant such cultures, the cells are either scraped carefully from the glass with a rubber spatula or the medium is taken off and the cells are removed with a chemical (enzyme or chelating agent). The cells are then suspended and counted, and the suspension is diluted with fresh medium to obtain the number of cells desired.


Objectives
Students will demonstrate one of the methods used to initiate primary cell cultures from fresh embryonic tissue and observe patterns and rate of cell growth in a mixed culture (the culture prepared representing a mixture or many cell types). Students will also become acquainted with the initiation and subculture of tissue cells in in vitro culture and with fixation and staining techniques used in the field of cytology, and will clearly differentiate between nuclear material and cytoplasmic material in animal tissue cells.


Materials
  • 2 curved dissecting forceps
  • 1 sterile culture tube
  • 1 sterile Petri dish per group
  • 25 sterile pipettes
  • 10 sterile culture flasks
  • 1 sterile glass rod
  • 2 sterile versene tubes
  • 1 sterile medium tube 199
  • 5 sterile alcohol pads
  • 1 staining jar
  • 1 bottle of hanks balanced salt solution, 100 mL
  • Methanol 30 mL
  • Hematoxylin stain 30 mL
  • Eosin stain 30 mL
  • Isopropyl alcohol 99% 100 mL
  • Histoclear
  • Piccolyte II mounting medium 15 mL
  • Fertile hen’s egg, incubated for 7 days
  • Isopropyl alcohol, 70%
  • Incubator
  • Compound microscope
  • Safety goggles
  • Lab aprons
  • Pre-Lab preparation
    Seven days before the experiment, obtain a fertile hen’s egg, and incubate according to instructions accompanying your incubator.

A. Establishment of the Primary Cell Culture
    Procedure Notes
    Though the embryo is relatively underdeveloped, working with a chick embryo may be a sensitive issue for some students. If this is the case, you may perform steps 1 through 12, involving maceration of the embryonic tissue, in advance. One fertile egg, incubated for 7 days, provides sufficient material for 5 lab groups. Do not remove sterile materials from their protective packages until you are ready to use them.

    Procedure
    1. Obtain a fertile hen’s egg, which has been incubated for 7 days. Candle the egg to verify the presence of a developing embryo and locate its position. To candle the egg, use a box containing a 150-watt bulb, with a 2”-diameter hole cut in the box. The box should be in a dark room. Place the egg, large end up, over the hole. If the egg is fertile, the developing blood vessels will be visible.
    2. Place the egg, large end up, in an egg carton. The large end contains the air sac. Wipe the egg thoroughly with a sterile alcohol pad to sterilize the surface.
    3. Sterilize a pair of forceps by passing them slowly through an open flame; allow to cool before using. Once used, items are no longer considered sterile.
    4. Gently crack the shell over the air sac. Remove the shell surrounding the air sac with the sterile forceps, taking care not to rupture the shell membrane.
    5. Tear off the shell membrane. Do not let bits of shell fall into the egg.
    6. Carefully pour the contents of the egg into a sterile Petri dish. With 2 pairs of sterile forceps, remove the membrane from around the embryo. Be very careful when removing the membrane; the embryo could explant. Materials can be damaged easily if pressed too tightly or pulled too hard.
    7. Place the embryo in the sterile culture tube. Cap the tube.
    8. With a sterile pipette, add 1 mL versene solution to the explant material. Discard the pipette. Set aside the remainder of the versene for use in part B of the investigation.
    9. Gently grind the explant material and versene with a sterile glass-grinding rod. Replace the cap on the culture tube and set aside for approximately 20 to 30 minutes. You may also macerate embryonic tissue with a sterile. Syringe. The process is similar to grinding, with less chance of contamination. Add 1 mL versene solution to the tissue in the culture tube. Place the embryo and versene in a syringe with at least a 50-mL capacity. Reset the plunger, position the syringe over the culture tube, and press, using one fluid motion. Pushing the tissue through the syringe will facilitate the action of the versene.
    10. With a sterile pipette, draw the tissue up and expel it from the pipette several times to homogenize it further. Discard the pipette.
    11. With a sterile pipette, add 1 tube of Medium 199 to the culture tube. Discard the pipette.
    12. Shake to mix the suspension well. Allow it to stand for 5 minutes.
    13. With a sterile pipette, carefully transfer 3 mL of the cell suspension to a sterile culture flask. Do not draw up large particulate matter. Cap the flask.
    14. Label the flask with the date of the culture and the tissue source. Incubate the flask with its largest, flat surface facing downward at 37°C (99 °F).
    15. Once the tissue has been incubated, invert the flask and place it on the stage of a compound microscope. The cells will be growing on the surface that was facing down during incubation. Examine the cells at 100X. At this magnification, an individual cell will appear to be about the size of a grain of rice. Focus carefully to view the nucleus within each of the cells. You should be able to observe a network of cells. Since there will be a mixture of cell types, some cells will be elongated and others compact. On occasion, some cells may regenerate into a large organized mass.
    16. If your cultures turn yellow (acidic) or the fluid becomes cloudy, it is an indication that they have become contaminated; they should be discarded. Disposed-of biologic materials should be autoclaved. If an autoclave is not available, place the unopened cultures in a pan of water, bring to a vigorous boil for 20 minutes, then discard. Instruments used in the investigation should likewise be sterilized by either autoclaving or boiling.
B. Propagation of Chick Cells onto Cover Slips
Once tissue cells have been established in primary culture (first isolate), they can usually be maintained for some time by serial subculture. However, primary cultures, as developed in part A, can usually be transplanted, or subcultured, only 2 or 3 times. When a cover slip is added to the culture medium in which the cells are suspended, some of the cells will settle out and grow on its surface creating a monolayer that can be viewed under a microscope.

Procedure
  1. Obtain a culture flask that has been incubated for 6 to 7 days.
  2. Pour off the medium, taking care not to contaminate the flask opening or its contents. Replace the cap. Discard the medium.
  3. With a sterile pipette, add 1 mL versene solution to the flask. Swirl the flask and pour off the versene. Replace the cap. Discard the pipette.
  4. Place the flask on a flat surface so the solution completely covers the layer of cells. Leave at room temperature for 15 minutes. Mark the side of the culture flask on which the cells are growing.
  5. The cells will be loosened by the versene solution and will float off the surface. If necessary, you may extend the incubation period, or you may agitate the culture flasks vigorously to help dislodge the cells.
  6. With a sterile pipette, draw up and eject the suspension several times to break up any larger cell clumps that may be present.
  7. Draw up the full amount of cell suspension in the pipette and transfer it to the unused vial of Medium 199. Pool the cells from 5 flasks into 1 fresh medium vial. Discard the pipette.
  8. Swirl the vial gently to mix the transferred cells with the fresh medium.
  9. Obtain a sterile vial containing a cover slip.
  10. With a sterile pipette, add 2 mL of the cell suspension to the vial. Cap the culture vial tightly. Discard the pipette.
  11. Label the vial with the name of the culture, its original initiation date, and the subsequent transfer.
  12. Place the vial in the incubator; stand it upright to ensure the cover slip remains perfectly flat on the bottom of the vial. Incubate the culture vial at 37°C (99 °F) for 4 to 5 days. The cells should settle and attach to the cover slip within a few hours.
C. Fixation and Staining of Chick Cell Monolayer
Cells, which are normally colorless, are fixed and stained for viewing under a microscope. Fixing is a process that stabilizes the chemical and structural characteristics of the cells. Cells are then stained with a dye or combination of dyes to highlight structural details. Different fixation and staining procedures highlight different structural features. The fixative methanol is good for monolayers. Hematoxylin, a powerful stain commonly used by cytologists, stains nuclear material bright blue to dark blue. Hematoxylin does not stain cytoplasmic material, however, so another stain must be employed, such as eosin, which stains cytoplasmic material light pink to red.

Procedure
Safety: Wear safety goggles at all times. Use care in handling stains This procedure should be performed 3 to 4 days after preparing the monolayer tissue culture.
  1. Heat Hanks balanced salt solution to 37°C (99 °F.) Place in a staining jar.
  2. Carefully remove the cover slip from the culture vessel: tip the cover slip away from the bottom of the vessel and gently remove it with forceps. The cover slip is extremely thin; be very careful when handling it with forceps.
  3. Immediately place the cover slip in warm Hanks balanced salt solution in the staining jar. Orient the cover slip so the side with cells attached is easily identified.
  4. Taking care to keep the cover slip in the staining jar, pour off and discard the used balanced salt solution. Immediately add fresh, warm balanced salt solution for a second washing. Repeat, for a total of 3 rinses in the salt solution.
  5. Pour enough methanol into the staining jar to just cover the cover slip. Allow cells to fix for 7 to 10 minutes. Agitate the staining jar occasionally to ensure thorough fixation. Be careful when using methanol; it is poisonous if taken internally.
  6. Pour off and discard the methanol. Rinse the cover slip with water to remove excess methanol.
  7. Add enough hematoxylin stain to completely submerge the cover slip. Allow the cells to stain in this solution for 10 minutes.
  8. Pour off and discard the stain. Rinse the cover slip with water.
  9. Add balanced salt solution and set aside for 2 minutes.
  10. Pour off and discard salt solution. Rinse the cover slip with tap water.
  11. Add enough eosin stain to completely submerge the cover slip. Allow to the cells to stain for 5 minutes.
  12. Pour off and discard the stain. Rinse the cover slip in 99% isopropyl alcohol 3 times. Allow 1 minute for each rinse, agitating the staining jar occasionally.
  13. Add Histoclear to the staining jar and allow to set for 2 minutes, agitating occasionally.
  14. Pour off and discard the Histoclear. Add fresh Histoclear and set aside for 2 minutes.
  15. During these 2 minutes, clean a glass microscope slide with a lint-free cloth.
  16. Just before the end of the 2-minute period, add 1 drop of Piccolyte II mounting medium to the center of the clean microscope slide.
  17. Remove the cover slip from the Histoclear and place it cell-side down onto the center of the microscope slide. Be sure not to trap air bubbles beneath the cover slip. This can be avoided by placing one side of the cover slip against the microscope slide and gently lowering the cover slip onto the slide. If an air bubble is trapped, it can be forced out by gently pressing on the cover slip with a sharp instrument.
  18. Allow the slide to dry for at least 24 hours before handling it.
  19. Examine the slide under 100X magnification. Note the pattern and rate of cell growth. The increase in cell rate can be roughly calculated by selecting a representative microscope field and counting the number of cells in that field. Mark the field by circling it with a glass marking pencil.
 
     
 
 
     




     
 
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