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					Brianna Payne                           1/20/2010                                 F6 & 11.5


F6 – Aseptic Technique
6.1 Objectives of Aseptic Technique: Through aseptic techniques, we aim to reduce
contamination to a minimum. Contamination comes from multiple sources: bacteria, yeast,
fungal spores, yourself, the air, work surfaces, etc. Once in a culture, contamination easily
spreads to others, making complete asepsis the only option! Or at least that is what we will aim
for. Five steps can help minimize contamination catastrophes:
   1). Check cultures frequently, particulary when handling them. Apparently special checks are
   required for mycoplasmal infection. Will we need to worry about that? What is it?
   2). Maintain cultures without antibiotics for at least some period of time
   3). Check reagents for sterility
   4). Do not share media across labs or cell lines. Not sure how much this applies to our work.
   5). Maintain a high standard of sterile technique.
   Maintaining Sterility – all elements coming into contact with the culture must be sterilized.
   And Aseptic technique is mandatory. Learn it! Use it! Because it employs a combination of
   different procedures, several must fail before contamination has a chance. Fortunately, once
   you get in the habit of taking those precautions, sterility is “not difficult to maintain”. Here’s
   hoping I get into the habit fast! Though equipment has improved, labs are often more
   crowded, busier places. Aseptic technique is still vital!

6.2 Elements of Aseptic Environment
   Quiet Area – free from air currents via doors and windows and passerby’s or equipment that
   causes air flow. Non-sterile activities (such as sample processing, staining, or extracting)
   simply should not occur in the tissue culture area.
   Work Surface – Clear and cleaned with 70% alcohol. Bring and keep only the items
   necessary on the surface, cleaning between procedures. Try to arrange your items so you
   easily obtain what you require as you work without compromising sterility. Laminar-flow
   hoods only work properly when they are clear. I wonder if we overdo it when we pour plates
   for FB. We haven’t had sepsis issues, yet it gets pretty crowded under there by the time we
   are done. I might split up the work between the two hoods next year. Wipe up spills with
   alcohol. You can minimize spills and other “oops” moments by working within your field of
   vision and arm’s reach. Clean the surface down with alcohol when you are done.
   Personal Hygiene – Wash your hands to remove dry skin and other microorganisms that
   might be hanging out there so that they do not get in your culture. Gloves may be helpful,
   but must also be washed and can lower sensitivity. Tie back long hair and refrain from
   talking while working at the bench. Yeah…..I don’t know about that. Since we have a
   vertical hood and a barrier….I can TRY to keep it to a minimum….but man! Avoid culture
   work at the height of an infection if ill.
   Reagents and Media – if obtained from commercial suppliers, they ought to be sterile. Swab
   unwrapped bottles with 70% ethanol when you bring them out of the fridge or a water bath.
Brianna Payne                            1/20/2010                                 F6 & 11.5


   Cultures – Quarantine new cell lines and handle them separately from others until proven
   innocent of contamination! Because they may suppress but not eliminate contaminations,
   avoid using antibiotics.

6.3 Sterile Handling
   Swabbing – Always wipe the work area down with 70% alcohol before, in between and after
   tasks are completed. As mentioned above, swab bottles from cold storage or water baths. Will
   need to label it in a creative manner, as sharpie will not work….nor lab tape I’m guessing.
   Capping – Deep screw caps are the way to go. “Wadless” polypropylene without the rubber
   liners is preferable.
   Flaming – On open benches, flame sterilize pipettes, bottle-necks and caps after opening and
   closing. If you do not hold the cap in your fingers, keep the open side of caps down and
   sterilize the bottom before replacing them. Flames are not recommending with laminar flow
   hoods, as they disturb the air flow and may constitute a fire hazard. We use them when
   working on FB prep fall semester. Should we not?
   Handling Bottles and Flasks – Hold bottles at a shallow angle to prevent contamination,
   exercising caution to not spill either. In the hood but never on the bench should they be
   vertical and open!
   Pipetting – Much preferred to syringes. Have a variety of size ranges available and
   sanitized. Fast flow can compromise accuracy. Avoid mouth pipetting. Does anyone really
   do this anymore? Why would you? And that would be compromising to sterility! An
   instrument should be selected that does efficiently sucks up and expels solution, as carryover
   is not desired. This technique often boils down to a compromise betwixt speed and accuracy.
   A good rule of thumb is to use the smallest pipette compatible with your manipulations. To
   keep a glass pipette sterile during use, insert a cotton plug into it’s top. If it becomes wet, get
   a new pipette. Plastic pipettes are already plugged.
   Pouring – Avoid it when possible, unless you plan to transfer all the contents of one sterile
   container into another in one foul swoop. Pouring, because it creates a bridge between liquid
   in the sterile bottle and that in the external environment, it begs contamination.
6.4 Laminar Flow – Flow can be horizontal or vertical. Horizontal flow provides the most
sterility to the culture, while vertical flow hoods grant the user more protection (they also
recirculate the air and extract 20% of it). Special hoods are required for potentially hazardous
material (some use carbon filter traps or have pathogen traps on the vent). For sterility, the
pressure drop across the filter must be at least 0.4 m/s. Hmm…do we have any way of measuring
flow on ours? Is it up to par? The HEPA filter should be checked (preferably by a professional)
for holes and airflow every six months. When was the last time ours was checked? Always clean
the work surface with alcohol (70% with 5% phenolic disinfectant) before, after and during
work. Ideally, flow should be running continuously. Occasionally, ultraviolet lights are used to
sterilize hoods. If such be the case, wear protective equipment.
Brianna Payne                           1/20/2010                                F6 & 11.5


6.5 Standard Procedure – Keep it clean and keep it clear. Do as much as you can before hand
to minimize the time cultures are out of the incubator. Proper preparation will allow you to work
smoothly and efficiently. Pay heed to sterile and non-sterile surfaces; may they never meet!
   Pages 80 – 83 outline two protocols (6.1 and 6.2) that integrate the concepts highlighted
   above, one for working in vertical Laminar flow and another for the open bench.
   Petri Dishes and Multiwell Plates – these equipment are particularly prone to
   contamination for a variety of reasons; a large exposed surface area when open, risk of
   touching rims to non-sterile surfaces (your fingers, the work surface, etc) while working, a
   shallow height makes it possible for media to “slop” between the lid and dish if not careful
   during handling, and a high risk of contamination in the humid atmosphere of a CO2
   incubator. To avoid these risks, do not leave them open for long, transport with extreme care
   and enclose in a transparent box for incubation. Remember to swap the box when you remove
   it for work! If media lodges between the lid and the base, throw away the lid, wipe the base
   rim with 70% alcohol and place a new lid on.
   Pages 83 – 84 contain a protocol for work with dishes and plates.
6.6 Apparatus and Equipment – Clean all equipment in the lab regularly. Pay careful attention
to clean any replacement items before they enter the lab. While aseptic work is occurring,
equipment transfer should wait.
   Incubators – Humid ones are a major source of contamination. Clean regularly (weekly,
   monthly, etc) by washing down all interior surfaces with a non-toxic detergent and then
   wiping down with 70% alcohol. Placing a fungicide in the incubator is only of limited help
   and cannot beat regular cleanings.
   Boxed Cultures – Always swab boxes down, inside and out, before use and let them dry. I
   assume let them dry in the Laminar Hood. Swab them down upon removal from incubators
   before removing and working with the cultures placed therein. Swab again before
   replacement.
   Gassing with CO2 – Slackened or special gas-permeable caps are often used to allow for
   gaseous equilibration, but can increase the risk of contamination. Purging flasks with sterile,
   premixed gas prior to sealing can be a good solution to this problem.

11.5 Sterilization of Media
Autoclavable Media – Enjoys a higher success rate than filtration while happily being less work
and more cost-effective. Some components of media may need to be added (Read FILTERED)
sterile post-autoclaving.
Sterile Filtration – Via 0.1-0.2 μm filters, which come made from many different materials.
Often only used for heat-labile solutions. Can be accomplished by positive pressure from such a
container (faster) or a peristaltic pump (can be switched on and off). To high of pressure can
“clog” the filter and does not work well for highly viscous material. Negative pressure filtration
is also available (may cause pH rise due to CO2 evolution).
Brianna Payne                            1/20/2010                                 F6 & 11.5


        Disposable filters – syringe tip for low volumes, intermediate sizes can be used in
conjunction with peristaltic pumps or vacuum filtration. Large capacity filters typically involve
in-line + pressure from a reservoir. Though more expensive, these filters are more efficient in
terms of use and success rates.
        Reusable filters – sterilized by autoclaving. Typically operate under + pressure (recall
via a reservoir or peristaltic pump). Vacuum lines allow for negative pressure filtration, which
typically has a lower risk for contamination. For larger volumes, employing a peristaltic pump
for direct filtration via an inline membrane or cartridge filter from reservoir to receptacle fitted
with a bell is a good option. Protocols employing positive pressure allows larger volumes to be
filtered (up to 50 L). Praise God I have never had to sterilize so much!
Protocols for sterile filtration via syringe (11.11) and vacuum filters (11.12) are given on pg 163
and 164-165. On page 166 -167, protocol for sterile filtration with small in-line filter (11.13) is
given, while 167-168 give a protocol for a large in-line filter (11.14).
Serum – Due to its viscosity and particulate material, serum is notoriously difficult to filter. It is
also expensive, so unnecessary waste ought to be avoided. It is best to simply buy sterile sera,
but it can be prepared in the lab. A graded series of fiters remove the particulate matter before
the serum is finally filtered trough 0.1 μm at low pressure.
Protocol 11.15, pgs 168-170, outlines sterile procedure for collection and sterilization of serum.
        Small-scale serum processing – for less than 1 L, scale down the given protocol and
        centrifuged small volumes at 10,00 gs after clot retraction before filtering though a series
        of disposable filters.
        Storage – Freeze sera (to -20 C) as rapidly as possible after sterilization in aliquots that
        can be used in 2-3 weeks after thawing. Frozen serum should be used in 6 -12 months.
        Human serum – outdated plasma from a blood bank can be used. Just “titrate out
        heparin or citrate anticoagulant with Ca2+”, allow the blood to clot, then separate and
        freeze.
        Quality contol – possibility for viral infection when using serum, but use of 0.1 μm
        filters should eliminate this threat. Bottom line, try to get your serum from clean
        sources.
        Dialysis – constituents sucha s AAs, glucose, nucleosides, etc, can be removed by
        dialysis.
Protocol 11.16 on pg 171 outlines procedure for Dialysis of Serum
Perparation and Sterilization of Other Reagents – Most are sterilized by filtration if heat
liable and autoclaving if heat stable.

Questions
What should be used to clean your work surface?
Brianna Payne                            1/20/2010                                F6 & 11.5


For sterility, what must the pressure drop across a filter in a laminar flow hood be?
What makes serum difficult to filter? What are the basic steps to accomplish this feat?
What pore size of filter must you eventually use if you want to avoid viral contamination?
Why might you want to avoid pouring liquid?

				
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