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. You will be working with a mouse embryonic stem cell line, which is unlikely to carry any agents that could harm you. Consequently, the guidelines here emphasize techniques for maintaining healthy and uncontaminated cells.




Maintaining cultured cells
Feeding
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.
Splitting
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.
Freezing
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.

Hood preparation
1. Wear gloves to protect yourself but also to prevent dry skin and micro-organisms from contaminating your samples.
2. Swab down the work surface liberally with 70% ethanol. Start from the back and proceed forward. Swab during work if necessary.
3. Swab any instruments that will be used in the hood with 70% ethanol, particularly the pipettes, which will often be used above biological samples.
4. 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.
5. 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).
6. 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.
7. 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.

Sterile handling
1. Spray gloves with 70% ethanol as often as necessary.
2. The indicator stripes on the autoclave tape should turn black if an object has been properly autoclaved.
3. 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).
4. 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.
5. Avoid working above an open bottle or dish in vertical laminar flow. Always work around them unless they are capped or covered.
6. 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.
7. 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.
8. Mop up any spills immediately and swab with 70% ethanol to prevent the growth of microorganisms.
9. 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.
10. Withdraw the pipette so that it slides through the sterile interior of the wrapper without touching the outside of the wrapper.
11. 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.
12. 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.
13. 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.
14. 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.
15. 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.

Cleaning up
1. Cap bottles tightly before removing them from the hood.
2. Swap down the work surface liberally with 70% ethanol.
3. Turn off the vacuum, if used.
Source from: openwetware.org

Disinfection

Methods designed for the disinfection/decontamination of culture waste, work surfaces and equipment represent important means for minimizing the risk of harm.
The major disinfectants fall into four groups and their relative merits can be summarized as follows:

• Hypochlorites (e.g. Chloros, Presept) Good general purpose disinfectant
  Active against viruses,Corrosive against metals and therefore should not be used on metal surfaces e.g. centrifuges. Readily inactivated by organic matter and therefore should be made fresh daily Should be used at 1000ppm for general use surface disinfection, 2500ppm in discard waste pots for washing pipettes, and 10,000ppm for tissue culture waste and spillage
• Phenolics (e.g. Sudol, Hycolin)
  Not active against viruses
  Remains active in the presence of organic matter
• Alcohol (e.g. ethanol, isopropanol)
  Effective concentrations 70% for ethanol, 60-70% for isopropanol
  Their mode of activity is by dehydration and fixation
  Effective against bacteria. Ethanol is effective against most viruses but not nonenveloped viruses. Isopropanol is not effective against viruses.
• Aldehydes (e.g. glutaraldehyde, formaldehyde)
  Aldehydes are irritants and their use should be limited due to problems of sensitization
• Glutaraldehyde may be used in situations where the use of hypochlorites is not suitable e.g. cleaning of centrifuge bowls or materials constructed of stainless steel that may be attacked or corroded by using hypochlorite solutions.

Waste Disposal

Any employer has a ‘duty of care’ to dispose of all biological waste safely in accordance with national legislative requirements. Given below is a list of ways in which tissue culture waste can be decontaminated and disposed of safely(especially the solid waste, such as flasks, centrifuge tubes, contaminated golves etc). One of the most important aspects of the management of all laboratory-generated waste is to dispose of waste regularly and not to allow the amounts to build up. The best approach is ‘little and often’. Different forms of waste require different treatment.

Tissue culture waste (culture medium) - Inactivate overnight in a solution of hypochlorite (10,000ppm) prior to disposal to drain with an excess of water
Contaminated pipettes should be placed in hypochlorite solution (2500ppm) overnight before disposal by autoclaving and incineration
Solid waste, such as flasks, centrifuge tubes(such as 15ml Centrifuge Tube, 50ml Centrifuge Tube), contaminated gloves, tissues etc. should be placed inside heavy duty sacks for contaminated waste and autoclaved prior to incineration. These bags are available from Bibby Sterilin and Greiner.
If at all possible waste should be incinerated rather than autoclaved.

Source: Sigma-Aldrich

Given below are a few of the essential "do’s and don'ts" of cell culture. Some of these are mandatory e.g. use of personal protective equipment (PPE). Many of them are common sense and apply to all laboratory areas. However some of them are specific to tissue culture.

The Do’s

1. Use personal protective equipment, (laboratory coat/gown, gloves and eye protection) at all times. In addition, thermally insulated gloves, full-face visor and splash-proof apron should be worn when handling liquid nitrogen.
2. Always use disposable caps to cover hair.
3. Wear dedicated PPE for tissue culture facility and keep separate from PPE worn in the general laboratory environment. The use of different colored gowns or laboratory coats makes this easier to enforce.
4. Keep all work surfaces free of clutter.
5. Correctly label reagents including flasks, medium and ampules with contents and date of preparation.
6. Only handle one cell line at a time. This common-sense point will reduce the possibility of cross contamination by mislabeling etc. It will also reduce the spread of bacteria and mycoplasma by the generation of aerosols across numerous opened media bottles and flasks in the cabinet.
7. Clean the work surfaces with a suitable disinfectant (e.g. 70% ethanol) between operations and allow a minimum of 15 minutes between handling different cell lines.
8. Wherever possible maintain separate bottles of media for each cell line in cultivation.
9. Examine cultures and media daily for evidence of gross bacterial or fungal contamination. This includes medium that has been purchased commercially.
10. Quality Control all media and reagents prior to use.
11. Keep cardboard packaging to a minimum in all cell culture areas.
12. Ensure that incubators, cabinet, centrifuges and microscopes are cleaned and serviced at regular intervals.
13. Test cells for mycoplasma on a regular basis.
    

The Don’ts

1. Do not continuously use antibiotics in culture medium as this will inevitably lead to the appearance of antibiotic resistant strains and may render a cell line useless for mmercial purposes.
2. Don’t allow waste to accumulate particularly within the microbiological safety cabinet or in the incubators.
3. Don't have too many people in the lab at any one time.
4. Don't handle cells from unauthenticated sources in the main cell culture suite. They should be handled in quarantine until quality control checks are complete.
5. Avoid keeping cell lines continually in culture without returning to frozen stock.
6. Avoid cell culture becoming fully confluent. Always sub-culture at 70-80% confluency or as advised on ECACC's cell culture data sheet.
7. Do not allow media to go out of date. Shelf life is only 6 weeks at +4ºC once glutamine and serum is added.
8. Avoid water baths from becoming dirty by using Sigma Clean (Prod. No. S5525).
9. Don’t allow essential equipment to become out of calibration. Ensure microbiological safety cabinets are tested regularly.

Source: Sigma-Aldrich ECACC Handbook