Specialized equipment is needed in order to carry out research with biohazardous agents. This equipment may include Biosafety Cabinets (BSCs), autoclaves, glove boxes, incubators, centrifuges, syringes, pipettes, and mixers.
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4.1 Biosafety Cabinets
In order to reduce the need for respirator use in laboratories, it is recommended that a biological safety cabinet should be used for all procedures that might generate aerosols or droplets and for all infected cell-culture manipulations of BSL2 and higher organisms.
The Principal Investigator is responsible for the provisions, maintenance, and upkeep of biosafety cabinets. Information on the proper use of equipment in relation to specific biohazardous materials shall be written and readily available to all employees with exposure potential. When temporarily inoperable, the handling or manipulation of the microbiological agent must be suspended until repairs are affected. A lapse in required certification will be cause to consider the equipment inoperable. The Biosafety Officer will periodically review authorized laboratories to assure that this equipment is operating as expected.
4.2 Biosafety Cabinet Certification
The National Research Council Committee on Hazardous Biological Substances in the Laboratory, requires the following:
- Class I and II biological safety cabinets shall be tested and certified in situ at the time of installation within the laboratory, at any time the cabinet is moved, and at least annually thereafter.
This annual certification program shall be continued for active BSCs that are used for microbiological agents or toxins requiring Biosafety Level 2 precautions. This is to assure proper functioning of this equipment. This program will protect both students and employees of the University.
Certification of biosafety cabinets shall be completed by outside contractors who employ appropriately trained technicians. The certification testing must be done in accordance with the specifications of the National Sanitation Foundation Standard #49.
All BSC HEPA filters need to be replaced at intervals varying from two to ten years, depending on cabinet use and the conditions of the laboratory environment. If filter replacement or repair work is required, the individual researcher is responsible to underwrite these expenses. Until the repair or filter replacement is completed, the BSC will not be certified and the cabinet in question shall be labeled by the PI as out of service for use with any infectious agents.
4.3 Procedures for Centrifugation
All centrifugation shall be done using centrifuge safety buckets or sealed centrifuge tubes in sealed rotors. If a small centrifuge is used and centrifuge safety cups are not available, the centrifuge should be operated in the biological safety cabinet. Each person operating a centrifuge should be trained on proper operating procedures. Keep a log book detailing operation records for centrifuges and rotors to assist in determining service requirements.
The following procedures for centrifugation are recommended:
- Examine tubes and bottles for cracks or stress marks before using them. Discard damaged tubes.
- Fill and decant all centrifuge tubes and bottles within the biological safety cabinet. Wipe outside of tubes with disinfectant before placing in safety cups or rotors.
- Never overfill centrifuge tubes as leakage may occur when tubes are filled to capacity. The maximum for centrifuge tubes is 3/4 full.
- Always cap tubes before spinning.
- Place all tubes in safety buckets or sealed rotors. Inspect the “O” ring seal of the safety bucket and the inside of safety buckets or rotors. Correct rough walls caused by erosion or adhering of matter and remove debris from the rubber cushions.
- Wipe exterior of tubes or bottles with disinfectant prior to loading into rotor or safety bucket.
- Never exceed safe rotor speed.
- Stop the centrifuge immediately if an unusual condition (noise or vibration) begins.
- Wait five minutes after the run before opening the centrifuge. This will allow aerosols to settle in the event of a breakdown in containment.
- Decontaminate safety carriers or rotors and centrifuge interior after each use.
4.4 Syringes and Needles
The hypodermic needle is a dangerous instrument. To lessen the chance of accidental injection, aerosol generation, or spills, the use of syringes are prohibited when alternate methods are available. The following practices are recommended for hypodermic needles and syringes in the laboratory:
- Use the syringe and needle in a biological safety cabinet only and avoid quick and unnecessary movements of the hand holding the syringe.
- Examine glass syringes for chips and cracks, and needles for barbs and plugs. This should be done prior to sterilization before use. Use needlelocking syringes only, and be sure that the needle is locked securely into the barrel. Replace glass syringes with plastic disposable syringes whenever possible.
- Whenever possible use safer needle systems.
- Wear latex gloves for all manipulations with needles and syringes.
- Fill the syringe carefully.
- Expel excess air, liquid and bubbles from a syringe vertically into a cotton pledget moistened with an appropriate disinfectant, or into a small bottle of sterile cotton.
- Do not use the syringe to forcefully expel a stream of infectious fluid into an open vial for the purpose of mixing. Mixing with a syringe is condoned only if the tip of the syringe is held below the surface of the fluid in the tube.
- If syringes are filled from test tubes, take care not to contaminate the hub of the needle, as this may result in the transfer of infectious material to the fingers.
- When inoculating animals, position the hand that is holding the animal “behind” the needle or use a pair of forceps to hold the animal in order to avoid puncture wounds.
- Be sure the animal is properly restrained prior to the inoculation and be on the alert for any unexpected movements of the animal.
- Before and after injection of an animal, swab the injection site with an appropriate antiseptic.
- Discard syringes into a sharps container. DO NOT bend, shear, recap or otherwise manipulate the needle.
4.5 Pipettes
The following practices are recommended for pipettes in the laboratory:
- Never suction or pipette by mouth; always use some type of pipetting aid when pipetting infectious materials. Preferably, all activities should be confined to a biosafety cabinet.
- Pipetting of toxic chemicals should be performed in a chemical fume hood.
- Infectious or toxic materials should never be forcefully expelled from a pipette. Mark-to-mark pipettes are preferable to other types because they do not require expulsion of the last drop.
- Infectious or toxic fluids should never be mixed by bubbling air from a pipette through the fluid.
- Infectious or toxic fluids should never be mixed by alternate suction and expulsion through a pipette.
- Discharge from a pipette should be as close as possible to the fluid or agar level, and the contents should be allowed to run down the wall of the tube or bottle whenever possible, not dropped from a height.
- Pipettes used for transferring infectious or toxic materials should always be plugged with cotton, even when safety pipetting aids are used.
- Avoid accidentally dropping infectious or toxic material from the pipette onto the work surface. Place a disinfectant dampened towel or other absorbent material on the work surface, and autoclave before discard or reuse.
- Contaminated pipettes should be placed horizontally into a pan or tray containing enough suitable disinfectant, such as hypochlorite, to allow complete immersion of the pipettes.
- Place discard pans for used pipettes within the biosafety cabinet.
- After suitable contact time, remove disinfectant. The pan and pipettes can be autoclaved together, and replaced by a clean pan with fresh disinfectant.
4.6 Blenders, Mixers, Sonicators, and Cell Disruption Equipment
Hazardous aerosols are created by most laboratory operations involving blending, mixing, stirring, grinding or disrupting biohazardous materials. Even the use of a mortar and pestle can be a hazardous operation. Other devices that may produce aerosols are ball mills, colloid mills, jet mills, tissue grinders, magnetic mixers, stirrers, sonic cleaning devices, ultrasonic cell disintegrators, and shakers.
The laboratory practices generally required when using equipment that may generate aerosols with biohazardous materials are as follows:
- Operate blending, cell disruption, and grinding equipment in a biological safety cabinet.
- Use safety blenders designed to prevent leakage from the rotor bearing at the bottom of the bowl. In the absence of a leakproof rotor, inspect the rotor for leakage prior to operation. A preliminary test run with sterile water, saline, or methylene blue solution is recommended prior to use.
- If the blender is used with infectious material place a towel moistened with an appropriate disinfectant over the top of the blender. Sterilize the device and residual contents promptly after use
- Glass blender bowls are undesirable for use with infectious material because of the potential for glass bowls to break.
- Before opening the safety blender bowl, permit the blender to rest for at least one minute to allow settling of the aerosol cloud.
- Grinding of infected tissues or materials with any open device is best done within a biological safety cabinet.