Tissue culture was originally developed about 100 years ago as a method for learning about mammalian biology. The term tissue culture was coined because people were doing exactly that, extracting tissue and letting it live in a dish for a short time. Today, most tissue culture experiments are done using isolated cells rather than whole tissues. Much of what we know about cancer, heritable diseases, and the effects of the environment on human health has been derived from studies of cultured cells.
Normal Mouse Fibroblasts; Photographs courtesy of G. Steven Martin
Transformed Mouse Fibroblasts; Photographs courtesy of G. Steven Martin
What types of cells do people study, and where do they come from? Cells that are isolated from tissue are called primary cells, because they come directly from an animal. It is very difficult to culture primary cells, largely because primary cells that are placed in culture divide only a limited number of times. This limitation on the lifespan of cultured primary cells, called the Hayflick limit, is a problem because it requires a researcher to constantly remove tissues from animals in order to complete a study. Cell isolation processes can be quite labour-intensive, and also can complicate data analysis due to inherent animal-to-animal variation. To get around the first of these problems, people have studied cells that are immortal, which means that they can divide indefinitely. (Some inherent cell-to-cell variation still exists in such cells.)
One familiar type of immortalized cell is the cancer cell. Tumor cells continuously divide, allowing cancer to invade tissues and proliferate. Cancer cells behave the same way in culture, and under the right conditions, cells can be taken from a tumor and divide indefinitely in culture. Another type of immortalized cell is the embryonic stem cell. Embryonic stem cells are derived from an early stage embryo, and these cells are completely undifferentiated and pluripotent, which means that under the right conditions, they can become any mammalian cell type. Mouse embryonic stem cells have become a valuable research tool, and it is this cell type that we will be using for our current experimental module.
The art of tissue culture lies in the ability to create conditions that are similar to what a cell would experience in an animal, namely 37°C and neutral pH. Blood nourishes the cells in an animal, and blood components are used to feed cells in culture. Serum, the cell-free (and clotting-factor free) component of blood, contains many of the factors necessary to support the growth of cells outside the animal. Consequently, serum is frequently added to tissue culture medium, although serum-free media exist and support some types of cultured cells.
Cultured mammalian cells must grow in a germ-free environment and researchers using tissue culture must be skilled in sterile technique. Germs double very quickly relative to mammalian cells. An average mammalian cell doubles about once per day whereas a bacterium is able to double every 20 minutes under optimal conditions. Consequently, if you put 100 mammalian cells and 1 bacteria together in a dish, within 24 hours you would have ~200 unhappy mammalian cells, and about 100 million happy bacteria! Needless to say, you would not find it very useful to continue to study the behavior of your mammalian cells under these conditions!
Maintaining cultured cells
Good cell culture technique will simultaneously protect you from anything dangerous that might be living with the cells and protect the cells from contamination by you. In Module 1, you will be working with an established cell line unlikely to carry any agents that could harm you. Consequently, the guidelines here emphasize techniques for maintaining healthy and uncontaminated cells. Some points are particular to the 20.109 cell culture facility but most are common practice and will be good habits for any tissue culture work you do.
As cells grow and divide in a dish, they use up the nutrients provided by the media. Old media must be removed and the cells must be “fed” with some fresh media. This must be done every two or three days for most animal cell lines.
Cells growing in a dish begin to crowd each other and then stop growing. This crowded state is called “confluence” and to maintain cells, confluent cultures must be “split” and “reseeded” into new culture dishes at a lower density.
Every lab that works with cultured cells has a freezer stock of each cell line they study. The freezer stock is a critically important resource for the lab, storing lines that aren’t in use but are worth saving and also providing “back-up” cells if working cultures get contaminated.
- Wear gloves to protect yourself but also to prevent dry skin and micro-organisms from contaminating your samples.
- Swab down the work surface liberally with 70% ethanol. Start from the back and proceed forward. Swab during work if necessary.
- Swab any instruments that will be used in the hood with 70% ethanol, particularly the pipettes, which will often be used above biological samples.
- Keep sterile pipette tips in “Hood Only” boxes that are opened only in a sterile environment. Swab the exterior of the box with 70% ethanol.
- Bottles should always be tightly capped when outside the hood (i.e., they should have been tightly capped the last time they were in the hood).
- Dry bottles thoroughly if they have been taken out of the water incubator. Swab them with 70% ethanol, especially at the neck and the bottom, and place them directly into the hood. Avoid shaking them vigorously during handling.
- Bring only the items you need for a particular procedure into the hood to prevent cluttering your working space. Having a clear working space will significantly reduce the chance of contamination! Ensure easy access to items in the hood and maintain plenty of clear space in the center of the hood to work in.
- Spray gloves with 70% ethanol as often as necessary.
- The indicator stripes on the autoclave tape should turn black if an object has been properly autoclaved.
- Never block the negative pressure zone (also the frontal non-sterile area) of the vertical laminar flow hood with objects (i.e., notebooks, pipetteman handle).
- Avoid working too close to the front of the hood. Keep working area at the center or towards the back. Keep the objects needed for the current procedure within reach; keep the others in the back.
- Avoid working above an open bottle or dish in vertical laminar flow. Always work around them unless they are capped or covered.
- Avoid leaving bottles, dishes, and flasks open when they are not in use. If the cap must be laid down, place it face-up/face-down towards the back of the hood where there is less traffic and less chance of being touched or crossed over. Correct cap placement has been debated. Having a cap facing up can potentially introduce airborne particles and drive non-sterile lid liquid onto the interior face of the cap, where contaminations can fall into the bottle upon recapping. If face-down placement is preferred, then make sure to swab the area specifically and thoroughly before the cap is placed down there. Conversely, if hood surface sterility cannot be absolutely guaranteed due to high traffic or cluttering, then face-up is a better option. The best placement, however, is to place the cap on its side and towards the back of the hood. This way the interior is not in contact with the air flow or with the work surface. However, this is not possible with dishes. Therefore, exercise good judgment in light of individual operating style and the hood setup.
- Never pour from one sterile container to another. Pouring will generate a liquid path to introduce infection from the outside to the inside. Always pipette or use filters when transferring from one bottle to another.
- Mop up any spills immediately and swab with 70% ethanol to prevent the growth of microorganisms.
- Withdraw a pipette from its wrapper at the center of the work area, tilt it so the tip (bottom end) is pointing away from the frontal non-sterile area and away from other objects in the hood.
- Withdraw the pipette so that it slides through the sterile interior of the wrapper without touching the outside of the wrapper.
- Avoid contact between the tip of the pipette and the mouth of the bottle. The mouth and neck of the bottle (both inside and out) present a potential source of contamination.
- When working with Pasteur pipettes, do not reach into the box to remove it. Instead, shake the box gently to cause the pipettes to slide out slightly, and then withdraw a pipette without touching the other pipettes or the tube interior.
- To keep the hood from being cluttered, do not leave any trash in the hood. Immediately discard uncontaminated wrappers in the regular trash. Put all pipette tips and biologically contaminated sharps in the sharps biohazard waste container. Put all biologically contaminated tissue culture plates, flasks, and other non-sharps in the non-sharps biohazard waste container. However, an effort to minimize entry/exit from the hood should be made to minimize disturbances in the laminar flow at the entrance, which may create the potential to waft in contaminants.
- Handle the pipette with a steady hand. Avoid large motions and do not let the tip touch anything non-sterile. Keep the tip away from the front and far above the objects in the hood.
- Do not fill a dish/flask so full or swirl it such that the medium spills over the edge. This will introduce a path of infection via liquid and may cause cross-contamination.
- Cap bottles tightly before removing them from the hood.
- Swap down the work surface liberally with 70% ethanol.
- Turn off the vacuum, if used.