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Copper for cell incubator to prevent contamination

Copper for cell incubator to prevent contamination


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For some reason the lab seems to have a problem with contamination every so often. It's virtually impossible to prevent bacteria, viruses, fungi, etc. from getting into the incubator every time you open the door. Our incubator is pretty old, we don't have one of the new fancier versions that's made with pure copper inside, which effectively kills microorganisms.

Would it be a good idea to buy a copper plate and place it on the shelf, on which we could put our culture flasks? Or would it disrupt the flow of heat/CO2 too much? Would it be better to buy copper foil and wrap the shelves in it, then poke holes into it where all the holes are in the shelf?

We use good cell culturing practices; I don't contamination is happening because people don't know what they're doing. It just happens over time with enough people working in a cell culture room. Could copper in the incubator help? We probably don't have the money to buy a brand new incubator that's made out of copper, so would this be a viable, economic alternative?


First thing I'd do is replace the HEPA filter. A copper plate/foil may help, and it certainly won't disrupt the flow of heat, as copper is incredibly conductive (that's why they make electrical wires out of it), and poking holes would help with the airflow, but a new filter will probably make the most difference. I've never used a full-copper incubator, and the few serious contaminations I've had to deal with were related to the HEPA filter and poor TC practices.

If you haven't already, I'd institute a good routine cleaning program as well. Change the water bath each week (make sure you're adding fungicide to the water), and wipe down the whole incubator with bleach followed by an alcohol rinse each month. Make sure you clean all the bleach off, as it corrodes metal if left on very long. At the same time, clean and autoclave all of the removable components like shelves, rails, and water pans.

Good luck!


Back to Basics CO2 Incubator: A Laboratory Staple

Research in the areas of cell biology, molecular biology, cancer, pharmaceuticals, etc. has made amazing strides during the past decade and the technology used in those areas has had to keep pace. The landscape (or labscape) of a typical life science laboratory has changed dramatically over the years but the CO2 incubator continues to be a staple in the research lab. Although the ultimate goal of maintaining cell culture stocks has not changed, the functioning and operation of CO2 incubators has become more accurate, more reliable and more convenient. CO2 incubators are designed with the user and his/her applications in mind. Three primary areas of concern when selecting a CO2 incubator are with reliability, contamination control, and ease of use. The following overviews several of the options that are available for CO2 incubators.


CO2 Incubator Humidity Control Q&A

In a recent Lab Manager webinar, we discussed in depth which types of CO2 incubator humidity control technologies to look for to provide optimal relative humidity and condensation control for cell culture work. If you didn&rsquot get a chance to attend this webinar, you can always view it on-demand here.

We received some great questions from attendees during the live webinar, and below is a summary for the benefit of all who need to control humidity and condensation for their work.

Will condensation impact the integrity of the RH sensor?

ignificant exposure to condensate can cause the accuracy of capacitive RH sensors (the type used in most CO2 incubators) to drift, and can lead to their failure, adding to increased maintenance costs, downtime, and the total cost of ownership of the incubator. This is why it is important to ensure that your incubator&rsquos control algorithm factors in all relevant variables to control humidity. For a review of these factors, read our RH and condensation control paper.

Which humidification system provides the best humidity recovery? Evaporative (water pan) or ultrasonic (nebulizer)?

While it is possible to attain good recovery times with a water pan, ultrasonic humidification offers the fastest recovery. As an example, Baker&rsquos Cultivo ® Ultra and Ultra Plus model CO2 incubators, which use a nebulizer to deliver humidity, recover humidity to set point in just 8 minutes after a 30-second door opening. The base model of Cultivo, which uses a water pan, recovers humidity to set point in just 10 minutes following a 30-second door opening. This is possible in part because of the water pan&rsquos large surface area. Most incubators that use smaller water pans will not recover that quickly.

Is it necessary to add copper sulfate as an antifungal?

This is not necessary if you use a copper water pan. See our contamination control technology paper for more details on how to control contamination in your CO2 incubator.

Are there any incubators that control oxygen tension?

Oxygen tension (the partial pressure of O2 dissolved within a liquid, such as blood plasma) can be controlled by managing the oxygen concentration (% O2) within the atmosphere of the incubated environment. A number of multi-gas incubators and controlled-oxygen workstations use gas mixing systems to control atmospheric O2 levels &ndash anywhere from 0.0% (anoxia) to 21% (ambient) O2, with varying ranges and stability.

Does the amount of water in the water pan affect the level of CO2?

No, the CO2 level in the chamber operates independently of the water level in the pan.

Are there stand-alone humidity sensors that could be placed inside an incubator to take recorded readings?

If your CO2 incubator does not have an embedded RH sensor, there are stand-alone RH sensors available that can be placed inside the chamber to monitor RH. However, an important advantage to using an incubator with an embedded RH sensor is that the data that is collected feeds immediately and automatically back to the control algorithm, which is programmed to make adjustments as needed to keep conditions inside the chamber stable. This is simply not possible with a stand-alone RH sensor.

Exactly how is the RH sensor calibrated?

Calibration procedures differ by brand. You can request a copy of the calibration procedure for your specific CO2 incubator from the manufacturer.

Could temperature or other gases affect the detection of humidity by the RH sensor?

As we discussed in the webinar, temperature does affect the accuracy of capacitive RH sensors. As temperature varies, RH sensor accuracy fluctuates from +/-1.8% to as high as +/-3.8%, with the least accurate RH detection occurring at the most common temperature condition, 37°C. This is why it is important that the incubator&rsquos control algorithm accounts for a fluctuating RH sensor accuracy when controlling RH.

The RH sensor used in Cultivo has been pre-qualified to function correctly in normal atmospheric conditions, as well as with high concentrations of CO2 (up to 20%). Baker does not recommend operating Cultivo outside those conditions, and the presence of other gases may affect the RH sensor&rsquos lifespan and accuracy. Check with your incubator manufacturer to learn more about the particular RH sensor used in your system.

What contamination control technologies are available in case you do have condensate?

The most common contamination control technologies in CO2 incubators are UV lights, copper interior components, and HEPA filters. A UV light is useful for controlling contamination in a water pan or reservoir, but there is a limit to its effectiveness. First, the effectiveness of a UV light is only as good as the intensity of the light&rsquos wavelength. This intensity diminishes over time, and because the bulb continues to operate long after its germicidal effectiveness has weakened, it is not always obvious when it is time to change the bulb. Second, only contaminants that come into contact with the UV light waves will be killed. Many UV lights make use of a hood or other partial container, which prevents the light waves from reaching most of the interior of the incubator.

Copper has intrinsic properties that can assist in destroying a wide range of microorganisms. As such, copper is well known for its ability to control contamination in CO2 incubators. Purchasing a unit with a copper interior or copper interior components may further mitigate the risk of contamination, should you experience excessive condensation. Some incubator suppliers use copper infused materials in construction and design (where a percentage of the material is copper), while others provide an interior that is 100% copper. The use of copper within an incubator is the subject of continuing study by academic research laboratories around the world. It is unclear whether either copper-infused or 100% copper is more or less effective. It is certaintly contingent upon the microorganism and its resistance to the antimicrobial properties of copper. Should you wish to pursue copper as a method of contamination control, the limitation of this technology is fairly obvious: any surfaces that are not manufactured from copper are available to harbor contaminants. This includes plates, sensors, instrumentation placed inside the incubator, and any other interior component that is not made of copper.

While a HEPA filter does help to control contamination, it doesn&rsquot help in the case of condensation, because it only captures contaminants in the air stream. However, we still recommend the use of a HEPA filter inside a CO2 incubator, preferably one with a large surface area for the most efficient filtration. Based on testing in our laboratory, the combination of a HEPA filter, UV light and copper interior components is the most effective at preventing contamination throughout the incubator.

We are working in a medical college laboratory with mainly undergraduates. How could we benefit from humidity control technology? Do you think it would be beneficial for the students?

In short, yes. The scientific method relies on systematic observation, measuring, and experimenting, as well as the formulation, testing, and modification of hypotheses. The tools, equipment, and instrumentation used to perform any test of a hypothesis are crucial links in that system, regardless of the level of education or complexity of research being performed. A hypothesis should be supported by results that are repeatable. As such, understanding or controlling the limitations of your current equipment, relative to the conditions your hypothesis requires, is crucial to the integrity of that experiment. Many tissues, cells, and microorganisms require precise environmental conditions to thrive. An incubator featuring control of temperature, CO2, and relative humidity can provide a learning tool to help students better understand the impact of one variable over a another on those cultures, affording them the opportunity to explore a wide array of research questions. In fact, relative humidity within an incubator chamber can impact these sensitive cell culture organisms, affecting their morphological and elastic properties, leading to a change in the structure of a cell wall and, therefore, growth rates.

What features should be considered for GMP environment application?

A CO2 incubator is a vital component of many bioproduction processes. Some incubators provide innovative features within the control software and user interface that are integral to the system, which do help certain customers achieve compliance with Good Manufacturing Practices (GMP). Such incubators are specifically designed to help users meet requirements for data logging, electronic recordkeeping, security, and permissions. Some incubators identify, track, and record any changes made to the operational parameters, as well as log any errors or events critical to the operation of the system. When integrated into quality and validation protocols for each organization, these features can help the end user validate the performance specification of the equipment itself, relative to the desired outcome of the research or work being performed. It is important to note, however, that it is the end user&rsquos responsibility for achieving compliance with all applicable regulatory guidelines, including GMP and FDA 21 CFR Part 11 compliance. Your incubator&rsquos operator&rsquos manual or your incubator supplier can provide you with more information on how your incubator may help you achieve compliance with these guidelines.

The Latest RH and Condensation Control Technology

For a more in-depth overview of the latest RH and condensation control technology, download our paper, &ldquoControlling Relative Humidity and Condensation in a CO2 Incubator&rdquo.


Heavy Metals

Some of the first chemical disinfectants and antiseptics to be used were heavy metals. Heavy metals kill microbes by binding to proteins, thus inhibiting enzymatic activity (Figure (PageIndex<3>)). Heavy metals are oligodynamic, meaning that very small concentrations show significant antimicrobial activity. Ions of heavy metals bind to sulfur-containing amino acids strongly and bioaccumulate within cells, allowing these metals to reach high localized concentrations. This causes proteins to denature.

Heavy metals are not selectively toxic to microbial cells. They may bioaccumulate in human or animal cells, as well, and excessive concentrations can have toxic effects on humans. If too much silver accumulates in the body, for example, it can result in a condition called argyria, in which the skin turns irreversibly blue-gray. One way to reduce the potential toxicity of heavy metals is by carefully controlling the duration of exposure and concentration of the heavy metal.

Figure (PageIndex<3>): Heavy metals denature proteins, impairing cell function and, thus, giving them strong antimicrobial properties. (a) Copper in fixtures like this door handle kills microbes that otherwise might accumulate on frequently touched surfaces. (b) Eating utensils contain small amounts of silver to inhibit microbial growth. (c) Copper commonly lines incubators to minimize contamination of cell cultures stored inside. (d) Antiseptic mouthwashes commonly contain zinc chloride. (e) This patient is suffering from argyria, an irreversible condition caused by bioaccumulation of silver in the body. (credit b: modification of work by &ldquoShoshanah&rdquo/Flickr credit e: modification of work by Herbert L. Fred and Hendrik A. van Dijk)

Mercury

Mercury is an example of a heavy metal that has been used for many years to control microbial growth. It was used for many centuries to treat syphilis. Mercury compounds like mercuric chloride are mainly bacteriostatic and have a very broad spectrum of activity. Various forms of mercury bind to sulfur-containing amino acids within proteins, inhibiting their functions.

In recent decades, the use of such compounds has diminished because of mercury&rsquos toxicity. It is toxic to the central nervous, digestive, and renal systems at high concentrations, and has negative environmental effects, including bioaccumulation in fish. Topical antiseptics such as mercurochrome, which contains mercury in low concentrations, and merthiolate, a tincture (a solution of mercury dissolved in alcohol) were once commonly used. However, because of concerns about using mercury compounds, these antiseptics are no longer sold in the United States.

Silver

Silver has long been used as an antiseptic. In ancient times, drinking water was stored in silver jugs. 8 Silvadene cream is commonly used to treat topical wounds and is particularly helpful in preventing infection in burn wounds. Silver nitrate drops were once routinely applied to the eyes of newborns to protect against ophthalmia neonatorum, eye infections that can occur due to exposure to pathogens in the birth canal, but antibiotic creams are more now commonly used. Silver is often combined with antibiotics, making the antibiotics thousands of times more effective. 9 Silver is also commonly incorporated into catheters and bandages, rendering them antimicrobial however, there is evidence that heavy metals may also enhance selection for antibiotic resistance. 10

Copper, Nickel, and Zinc

Several other heavy metals also exhibit antimicrobial activity. Copper sulfate is a common algicide used to control algal growth in swimming pools and fish tanks. The use of metallic copper to minimize microbial growth is also becoming more widespread. Copper linings in incubators help reduce contamination of cell cultures. The use of copper pots for water storage in underdeveloped countries is being investigated as a way to combat diarrheal diseases. Copper coatings are also becoming popular for frequently handled objects such as doorknobs, cabinet hardware, and other fixtures in health-care facilities in an attempt to reduce the spread of microbes.

Nickel and zinc coatings are now being used in a similar way. Other forms of zinc, including zinc chloride and zinc oxide, are also used commercially. Zinc chloride is quite safe for humans and is commonly found in mouthwashes, substantially increasing their length of effectiveness. Zinc oxide is found in a variety of products, including topical antiseptic creams such as calamine lotion, diaper ointments, baby powder, and dandruff shampoos.

Why are many heavy metals both antimicrobial and toxic to humans?


Protect yourself from cell culture contamination

Ranging from minor annoyance to major catastrophe, contamination causes loss of time, money, and effort spent in developing cultures and setting up experiments.

Contaminants can affect all cell characteristics (e.g. growth, metabolism, and morphology) and contribute to unreliable or erroneous experimental results. Cell culture contamination will likely create a need for experiments to be repeated, resulting in frustrating time delays and costly reagent wastage. Data derived from undetected contaminated cultures can end up published in scientific journals, allowing others to build hypotheses from dubious results. Cell culture contamination costs millions of dollars every year in the United States alone, and it seems to be only getting worse. While the problem will not and cannot ever be fully defeated, however, it can be fought.

Protect yourself from cell culture contamination

The first step in avoiding cell culture contamination lies in being aware of potential sources, and building practices that reduce the risk of contamination from those sources.

Before being allowed to work in a tissue culture facility, laboratory personnel should be given full practical training in aseptic cell culture techniques by an experienced staff member. While each laboratory will have their own standard operating procedures related to use of incubators, autoclaving, labeling of cultures, media storage, and waste disposal, guidelines typically include the following tips:


Cell culture contamination.

Sooooo I have my first contamination of my cell culture. I'm really kicking myself because I am usually so on top of things. I am not sure where it came from. I found one contaminated bottle of media which i threw out so I hope that I got rid of the source. I'm thankful it was bacterial and not fungal at least. Now I am throughly cleaning everything in the room. I'm going to clean the incubator but we don't have a big enough autoclave to fit the shelves inside. we have one of those tiny bioclave things which holds practically nothing. How effective would it be to use UV and ethanol? my PI suggested using these clorox wipes and then ethanol as well. I would really appreciate some advice so I don't have to worry when I revive my cells!

First off - contaminations happen to everyone occasionally. It's OK, just remember to prevent it from happening in the future and move on. Don't freak out, it happens to everyone.

Don't trust the UV, use bleach and ethanol to wipe the shelves down. I'll expound on this more later.

The suggestions /u/kroxywuff gave are all good ones, and might help you track down your contamination. Remember, though, that it will tell you where you contamination is, not where it came from.

Trying to trace your contamination to the source will likely cost you more time and money than just throwing everything away and starting over. If it happens once, no big deal. If it keeps happening, that's when you need to start investigating. It sounds to me like this is your first one and you're overreacting. It also sounds like your media became contaminated.

I have some suggestions, and it's combined with a bit of my opinion and some preachy-type stuff that I've developed after working with tissue culture for the last 10 years, so bear with me, and take what I say with a grain of salt if you think any of it doesn't make sense, but I will give you my honest opinions:

Contamination comes from technique error. Something that wasn't supposed to be in the culture got in. That's it. Now, maybe a package or a container was cracked or compromised, but 99% (not a real statistic) of contaminations are caused by human technical error. We avoid those errors by using proper aseptic technique.

Presumably you're doing your culture in a biosafety cabinet, which has been serviced within the last year. The BSC is the most essential piece of equipment and it's the one that is most abused.

Remove everything from the cabinet, and clean it. You might even consider taking the tray out (if possible) and cleaning underneath it (consult the user manual, and use proper PPE!) You can use a 10% bleach solution for a few minutes, followed by 70% ethanol. Remember to rinse everything with ethanol because long-term chlorine exposure will damage stainless steel after a while.

When working in the BSC, do not leave items inside the cabinet for long periods of time. Remove items and wipe them down before and after use. If you keep boxes of pipet tips in there, throw away the ones you have now and get new ones. Keep the work area clear of anything you don't need. Never block the grills! I see people do this all the time with notebooks, paper, tools, bottles, ect. Blocking the airflow will compromise the sterile environment of the cabinet. It is very bad. Consult videos online and ask your institution for training in proper aseptic technique.

UV lights are source of contention. I don't like UV in BSCs for several reasons. The light only disinfects things on the outside and only the side facing the light. They take several minutes to several hours to effectively disinfect, depending on the organism and the power of the UV. UV bulbs also decrease in effectiveness over time. I don't have data but I've heard different numbers that range from a few weeks to a few months. It's likely your bulb is only replaced once a year or when it burns out so it might not be as effective as youɽ like. Also, UV destroys plastic, it will bake-on any spills that you don't clean up and leave a stain on the cabinet surface, and if you leave the light on while you're in the room, it's bad for your skin. I never use the UV, personally, but some people swear by them. If you do use them don't leave plastics in the BSC while it's on. The CDC recommends against using UV in BSCs. At any rate, the UV is no substitute for a bleach/ethanol cleaning. info: https://cfo.asu.edu/node/2667?destination=node%2F2667

For the incubator, you can use the same cleaning procedure on the shelves, although I would advise against spraying bleach inside the warm environment of the incubator, for those you could use wipes and ethanol. Remove all the old water and replace it with new DI (sterilize the DI if you can).

Remember: an incubator is made to grow things. The instant you open it you expose it to the outside air and all the microorganisms in the environment. Unless it is kept in a clean room, it's likely the dirtiest thing in your laboratory, along with the water bath. Make a schedule and clean them at least quarterly. I recommend monthly or weekly for the water bath. You can use chemicals and algicides in the water if you wish, but remember that they might select for more precocious organisms, so nothing can substitute for a good cleaning and water change.

As long as your supplies are sterile and not compromised, they should be fine. The flasks should seal appropriately and filter the air so that nothing gets in while it's sitting in the incubator. I've heard people with PHDs talk about what I would describe as magic of bacteria 'somehow' getting into closed containers in the incubator, but don't go chasing ghosts because you will just drive yourself crazy.

The BEST thing you can do is throw away ALL the reagents you used for that culture and start fresh. If that's not possible, sterile filter your reagents and complete media with a 0.22um filter before use. It's a good practice to sterile filter all media once it's made.

A good technician using proper aseptic technique and sterile reagents and supplies should have greater than 90% contamination-free cell culture without the use of antibiotics. So make sure your technique is good and you'll be fine. Don't sweat 1 contamination because life is too short.

TLDR: Don't worry about 1 contamination, Use good technique, use bleach & ethanol for cleaning, don't trust UV, toss or filter reagents, technique technique technique!


Laboratory contamination: Preventing contamination

In the second part of this laboratory contamination series, experts from Thermo Fisher Scientific offer their top tips on the prevention of laboratory contamination.

Part two, presented by Mary Kay Bates, global cell culture specialist and Douglas Wernerspach, business director, Thermo Fisher Scientific, is entitled: Tips for preventing contamination.

In the first part of this series, we discussed types of biological contaminants and learned that laboratory personnel and procedural errors can be the greatest contamination sources, but what can we do about it?

“Contamination most frequently occurs through avoidable procedural errors

Modern laboratories are busy environments with personnel sharing equipment across overlapping workstations that may be near high-traffic areas and busy instruments.

Practicing good aseptic technique is critical to maintaining the purity of cell cultures as well as a safe lab environment.

Some of the most basic laboratory procedures are the most important, including using proper aseptic technique, wearing clean lab coats and washing hands in order to reduce the risk of introducing microorganisms into mammalian cell cultures.

Below we discuss other techniques that are not as basic - good lab design, consistent culturing procedures with regular identity testing to document the authenticity of cell lines, and cleaning regimes.

These procedures may not currently get the attention they deserve.

Every lab should have a dedicated area for cell culture that is separate from high-traffic lab areas.

Only essential personnel should have direct access to the cell culture area. The placement of water and HVAC units should also be considered carefully.

For example, since tap water can be a source of microbial contamination, sinks and water baths should be located far enough away to avoid splashing onto sterile workstations.

Also, the air diffusers of HVAC units should not be installed directly above incubators since mould spores and other contaminants could be blown into the workspace.

Small labs need to be particularly careful about lab design because as they grow and new instruments are introduced, the layout of the lab can become more vulnerable to contamination.

Culturing procedures

Practicing good aseptic technique will help advance research from the first passage procedure to genomic analysis to understanding what drives the disease.

The first step in an aseptic culturing procedure is to avoid cross-contamination by only working with one cell line at a time.

Accidental switching of cell lines and cross-contamination between cultures often occurs and can result in erroneous or misleading data.

Biological safety cabinets (BSCs) and work areas should be stringently cleaned and disinfected between culture sessions.

Advanced protocols in a modern cell culture lab require frequent testing.

Testing for mycoplasma is recommended on a monthly basis. To avoid cross-contamination and mixing of cell lines, work with only one cell type at a time.

Finally, avoid routine antibiotics in cell culture media, since antibiotics can mask an underlying contamination which is not targeted by the antibiotics.

A clean, well-organised lab that has multiple users with different experience levels should also employ Good Pipetting Practice (GPP) to assure both accuracy and sample safety.

The first step is to ensure that lab employees avoid accidentally touching the pipette tip to non-sterile media and surfaces.

If such an error does occur, that tip should be discarded.

Next, cell culture processes from growth and passage through analysis all require secure pipetting systems with precise reproducibility.

Thermo Scientific’s F1-ClipTip Pipetting System prevents the pipette tip from falling off mid-application.

Having a secure seal for each sample with exact sample volumes makes ClipTip an invaluable tool to accelerate discovery.

Use of high quality consumables, culture media and reagents from reputable manufacturers is an easy way to ensure consistent performance in the lab.

Consumables and sterile containers should be made from resins that are free from additives that could leach into samples or reagents and negatively impact research results.

Cleaning procedures

To maintain safe and sterile work surfaces, a standard operating procedure for properly maintaining, cleaning and disinfecting equipment, work surfaces and non-work surfaces needs to be implemented.

Lab cleaning should include any high flat surfaces, such as the tops of refrigerators, freezers and incubators, which can collect dust and other potential contaminants.

In a similar way that a vault protects your valuables, protection of cell lines starts with a reliable BSC.

Choose a BSC that will ensure safe air filtration and prevent sample contamination which are designed for user simplicity and sample security.

Culture media should only be opened in the BSC and lab personnel should avoid clutter in the BSC workspace.

Use gentle arm movements to reduce disturbances in the clean downflow air while working.

Filter and airflow integrity should be tested yearly by a certified specialist. Inner surfaces should be cleaned and disinfected after each working session and all surfaces, including the tray under the working surface, should be cleaned and disinfected monthly with 70% ethanol or another appropriate disinfectant.

Furthermore, procedures need to be optimised to minimise the number of times the user removes their hands and arms from the BSC work area while manipulating samples.

The work should be planned so that the BSC can be loaded with the samples and supplies needed for the procedures and then to remove waste and clean-up after the procedure.

During the procedure the BSC operator only needs to work with items already inside the BSC work area. Regular cleaning procedures must be followed and spills must be cleaned immediately.

Today’s new procedures and innovative protocols require a CO2 incubator with advanced features that help prevent contaminants from becoming established within the incubator or invading cultures once inside.

Understanding the fundamental features of CO2 incubators and assay requirements for successful experimentation is important to making an informed decision when purchasing an incubator.

Because microorganisms are our constant companions and they can enter any incubator each time the door is opened, investing in leading-edge technology incubator design is imperative to combating cell culture contamination.

Look for an incubator that works with you to control and combat contamination.

The Thermo Scientific Forma incubator, for example, can offer HEPA filtration that scrubs the entire chamber air volume every 60 seconds to establish ISO 5 cleanroom conditions within five minutes after every door opening.

This feature is available in either a water-jacketed or a direct heat format. Direct heat incubators can offer additional protection with an automated high temperature cycle that saves time by eliminating the need to autoclave internal parts.

The Thermo Scientific Heracell CO2 incubator provides a 100% pure copper internal chamber and parts, as well as high temperature decontamination.

Also available today are advanced, precise microprocessor features for CO2 sensors, humidity control and onboard data monitoring that allow researchers to perform their best work in a trouble-free environment.

Key features for an incubator include:

  • Easy-to-clean surfaces
  • Copper interior chambers
  • Good air filtration - HEPA filters
  • Automated high-temperature disinfection cycle
  • Solid-glass, divided, gas-tight inner door
  • External water source
  • Data logging and touch screen controls
  • Advanced CO2 and O2 sensors

Contamination most frequently occurs through avoidable procedural errors.

Without good aseptic practices and consistently applied cell culture management procedures, chronic contamination problems have a higher probability of occurrence.

A properly managed laboratory will provide researchers with an environment that reduces the chances of accidents and errors and leads to more consistent and reliable performance.

The third and final part of the series will provide useful advice on how to choose a membrane and filtration product.


Incubator-Clean™

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© 2021 VWR International, LLC. All rights reserved.
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Protocol

1. Prepare a Sterile Workspace

Before starting any sterilization procedures in your work area, wash your hands thoroughly with antiseptic soap and warm water.

Be sure to re-wash your hands any time you suspect you have contamination from your experimental manipulations.

Clear away all materials cluttering your work area on the laboratory bench. Remove a pre-moistened disinfectant wipe from the canister and wipe down the entire area. Allow the disinfectant to evaporate - do not wipe dry!

Use disinfectants such as alcohol (isopropanol or 70% ethanol) or phenolic compounds (o-phenylphenol) .

To prevent aerosolization, or the production of a fine mist containing bacterial cells, and spread of microbial contaminants, avoid dispensing disinfectant from a squeeze bottle.

Desiccation of microorganisms is one of the most effective ways to decontaminate surfaces.

Even if someone has recently used the laboratory bench and the bench top was wiped down with disinfectant, ALWAYS begin your laboratory time by wiping down the bench.

After the disinfectant has dried completely, use an igniter to light the Bunsen burner. Adjust the flame so that a blue cone can be seen in the middle of the flame. The flame is now producing an updraft, or air convection currents in which warm air rises up and away from the flame (Figure 1). As heat rises, microorganisms and dust particles are forced upward and away from the immediate work area. Work slowly, carefully, and deliberately at all times within this area created by the Bunsen burner, referred to as a sterile field. Keep the Bunsen burner on during the entire procedure.

The tip of the blue cone is the hottest part of the flame.

Be careful not to disturb the updraft by rapid movements that dramatically change the air currents around the laboratory bench. Creating an updraft with the Bunsen burner minimizes the possibility of microorganisms and dust falling onto the bench or into open bottles, tubes or flasks in the work area.

Arrange all the supplies needed for the procedure on the laboratory bench near the sterile field. Make sure all the materials are properly labeled.

Supplies may include serological pipettes and micropipettors, sterile culture tubes, sterile flasks, media bottles containing broth, sterile microcentrifuge tubes, micropipettor tips, racks for tubes, bacterial cell cultures, and phage stocks.

Liquid media should be sterilized in an autoclave at 121 ଌ for at least 15 minutes on the liquid setting. Larger volumes of media (> 1L) require longer autoclave times. Labware should be sterilized in an autoclave at 121 ଌ for at least 30 minutes on gravity (dry) setting.

In general, sterile solutions may be stored at 4 ଌ for up to 5 months. Note that storage time is significantly reduced for solutions containing unstable components such as antibiotics - always check the manufacturer's recommendations.

2. Transferring Liquids Using Serological Pipettes

Serological pipettes come in many sizes and options: plastic or glass, disposable or reusable, plugged or unplugged. These are calibrated to deliver volumes ranging from a 0.1 ml to 25 ml.

Common sizes for serological pipettes are 5 ml, 10 ml and 25 ml and should be used for aseptic liquid transfers of 0.1 ml or more (panel A of Figure 2). There also are larger serological pipettes that can deliver volumes up to 100 ml however, the focus of this protocol is on the more common, smaller sized pipettes.

Pre-sterilized pipettes with a cotton wool plug are needed for microbiology and tissue culture experiments. The plug should not be removed from the top of the pipette it is designed to function as a barrier to overfilling the pipette.

Different applications call for plastic versus glass serological pipettes. Glass is needed for organic solvents. Either can be used when performing BSL-1 experiments on the laboratory bench top. Only plastic may be used when working in a Biosafety cabinet with BSL-2 organisms where a Bunsen burner cannot be used. It also is recommended that plastic be used for applications involving transfer of molten agar.

Serological pipettes are of two types: TC ("to contain") or TD ("to deliver"). TC pipettes deliver all the volume, including the tip, and must be "blown out" or rinsed to get the specified volume. TD pipettes are calibrated to leave a tiny bit in the tip that should not be delivered. Be sure to check the label on the body of the pipette near the top to ascertain which type it is (Figure 3). The most commonly used are TD pipettes, which are marked with double rings at the top.

Take a sterile plastic serological pipette (also called a volumetric transfer pipette) and carefully remove the paper sleeve at the end with the cotton wool plug by peeling it away like the skin of a banana - do not remove the entire sleeve, protecting the tip of the pipette that will come in contact with the liquid to be transferred. Touch only the top of the pipette (above the graduation marks) with your hands.

Never go into a sterile solution with a used pipette, even if great care has been taken to keep it sterile.

Glass serological pipettes are typically stored in metal canisters (panel B of Figure 2). Loosen the top of the canister then carefully remove the cap, and flame the open ends of the cap and canister. Place the cap down, on its side, on the disinfected bench. Remove one pipette from the canister by holding it horizontally and gently shaking it so the tops of one or two pipettes stick out about an inch and can be easily grasped. Lay down the canister on its side and remove one pipette, but be cautious not to touch the other pipettes in the container. Do not touch the bottom tip of the pipette with your hands, and avoid contact of the tip with other non-sterile surfaces.

Affix a pipette aid such as a bulb, pump, or gun to the top end of the serological pipette. Remove the paper sleeve from the plastic pipette. Hold the pipette aid in your right hand.

If using a glass pipette, pass the bottom third of the pipette through the blue cone in the Bunsen burner flame for 1-3 seconds. Rotate the pipette 180° as it passes through the flame. Plastic pipettes and tubes cannot be flamed.

If left-handed, hold the pipette aid in your left hand and conduct subsequent manipulations of culture bottles and tubes with your right hand.

Contamination tends to occur with plastic pipettes when withdrawing the final inches of the pipette from the sleeve because the sterile tip comes into contact with the part of the sleeve touched by your hands.

Remove the cap of the bottle containing sterile media. Do not place the cap on the laboratory bench but hold it between your ring finger and palm of the right hand while manipulating the pipette aid with the thumb, index and middle fingers of the same hand (Figure 4). Holding the bottle at a 45° angle, pass the rim of the bottle through the flame of the Bunsen burner, creating a sterile field around the open bottle.

Although best avoided, if you must put the cap down, place it face down on a disinfected surface. With a cap that faces up, there is a greater chance of contamination from movements of objects or hands, creating air currents that cause microorganisms and dust particles to descend to the inside surface of the cap.

The purpose of flaming is not to sterilize but to warm the opening of the bottle and create air convection currents up and away from the opening (i.e., updraft). The warm, rising air helps prevent dust particles and other contaminants from entering the bottle.

Keep the sterile container open for as little time as possible. It is important to keeping the points of entry of airborne microorganisms to a minimum throughout the procedure.

Avoid coughing, sneezing, talking, and other inadvertent movement while sterile containers are open.

Never pass hands and fingers over the top of a sterile field (i.e., open bottles or flasks, the inside of tubes and bottle caps) once they have been passed through the Bunsen burner flame.

Always work with an open flame when opening sterile tubes or bottles. Never have more than one tube, bottle, or flask open on the bench at a time. Flaming should be done immediately upon opening and just before closing tubes, bottles and flasks.

Place the tip of the serological pipette into the bottle containing the sterile media then aspirate, or draw the sample aseptically, from the bottle. Use the pipette aid to control the flow of the sample into the pipette. Precisely read the volume drawn into the pipette by aligning the meniscus formed on top of the liquid column to the graduation marks on the calibrated pipette (Figure 5).

DO NOT MOUTH PIPET! Always use a pipet aid (pump, bulb, or gun).

Pay attention to the sequence of numbers when determining the volume aspirated. The numbers may be printed tip to top, or vice versa, or often times in both directions.

When reading the volume, always hold the pipette vertically, perpendicular to the ground, and view the liquid meniscus dead-on at eye level.

Serological pipettes are only as accurate as the smallest marked increment, which is typically 0.1 ml for 5 ml and 10 ml pipettes and 0.2 ml for 25 ml pipettes. If greater precision is needed, a serological pipette may be used in combination with a micropipettor.

Pass the rim of the bottle through the Bunsen burner flame once again, then place the cap back on the bottle. Set the media bottle aside.

Don't burn yourself with the Bunsen burner in a rush to close the bottle.

Hold a test tube or flask in your left hand. Remove and hold the cap as described in step #4 above. Create a sterile field by flaming the rim of the tube or flask in the Bunsen burner.

Dispense the media in the pipette into the tube or flask. Control the flow of the sample so it does not splash out of the tube or flask.

Volumes may be measured such that the entire volume is delivered and the pipette drains completely, or a specific volume is achieved by doing a point-to-point delivery (one volume marking to another).

Pass the rim of the tube or flask through the Bunsen burner flame once again, then replace the cap. Set the tube or flask aside. Remove the pipette aid, and discard the pipette into the proper waste receptacle.

Plastic serological pipettes are disposable, while glass serological pipettes can be sterilized and used again. Proper disposal requires plastic pipettes be placed in a designated sharps container (rigid box lined with plastic disposal bag) while glass pipettes initially should be immersed in a container with 10% bleach solution to disinfect the inside and outside surfaces. Then the glass pipettes should be thoroughly washed with laboratory detergent, rinsed with distilled water, and sterilized in an autoclave.

These same steps should be followed when inoculating media with a bacterial culture or phage stock or when performing serial dilutions.

3. Transferring Liquids Using Micropipettors

Precisely measuring and dispensing minute volumes can be accomplished using micropipettors (also referred to as Pipetman Panel A of Figure 6). These instruments come in different sizes each with a specific volume range: P2 for 0.2-2 μl , P10 for 1-10 μl, P20 for 2-20 μl, P200 for 20-200 μl, and P1000 for 200-1000 μl.

Treat micropipettors with care, as they are precision instruments. Do not leave them lying on the laboratory bench unattended where they can be knocked off and damaged. Do not allow pipettes to come into contact with corrosive chemicals.

For volumes greater than 1000 μl, use a serological pipette.

Although working within the sterile field created by the Bunsen burner, do not flame micropipettors, tubes and plastic tips. The tubes and tips should be pre-sterilized. The micropipettors may be wiped down with a pre-moistened disinfectant wipe prior to use.

A numeric volumeter showing the dispensed volume can be set by turning adjustment knob. Adjust the volume before proceeding to step #3.

NEVER TURN THE ADJUSTMENT KNOB ABOVE THE INTENDED RANGE!

To obtain maximum accuracy when decreasing the volume setting on the micropipettor, slowly dial down the thumb wheel, making sure not to overshoot the graduation mark.

To obtain maximum accuracy when increasing the volume setting on the micropipettor, dial the thumb wheel up, passing the desired graduation mark by 1/3 of a turn. Then slowly dial down the thumb wheel to reach the intended volume, making sure not to overshoot the graduation mark.

The volumeter shows three numbers. Depending on the micropipettor, the numbers are interpreted differently. Note that each micropipettor is only as accurate as the smallest graduation mark.P2: For volumes between 0.2-2.0 μl. The top number denotes volume in microliters. The second number indicates tenths of a microliter (0.1 μl), and the third number represents hundredths of a microliter (0.01 μl). Each graduation mark equals an increment of two one-thousandths (0.002 μl) of a microliter.P10: For volumes between 1.0-10.0 μl. The top number is for tens of microliters this usually is set at "0" and should only be set at "1" with the other two numbers set at "0" when dispensing 10.0 μl. The middle number denotes volume in microliters. The third number indicates tenths of a microliter (0.1 μl). Each graduation mark equals an increment of two one-hundredths (0.02 μl) of a microliter.P20: For volumes between 2.0-20.0 μl. The top number in black is for tens of microliters this should only be set at "2" with the other two numbers set at "0" when dispensing 20.0 μl. The second number in black denotes the volume in microliters. The third number in red indicates tenths of a microliter (0.1 μl). Each graduation mark equals an increment of two one-hundredths (0.02 μl) of a microliter.P200: For volumes between 20.0-200 μl. The top number is for hundreds of microliters this should only be set at "2" with the other two numbers set at "0" when dispensing 200 μl. The middle number indicates the dispensed volume in tens of microliters, and the third number denotes volume in microliters. Each graduation mark equals an increment of two one-tenths (0.2 μl) of a microliter.P1000: For volumes between 200-1000 μl. The top number is for thousands of microliters this usually is set at "0" and should only be set at "1" with the other two numbers set at "0" when dispensing 1000 μl. The middle number is for hundreds of microliters. The bottom number is for tens of microliters. Each graduation mark equals an increment of two (2 μl) microliters.

Performance check: These instruments should be calibrated annually, ensuring accuracy and precision are maintained to stay within ± 5% of specifications. Use an analytical scale to measure water, making sure the minimum and maximum settings correspond to the intended volume. For example, use a P1000 to transfers 200 μl of water to a weigh dish on the scale. Since water has a density of 1, then 1 ml of water is equivalent to 1 gram (g). Thus, 200 μl (0.2 ml) of water should be equal to 0.2 g. Also, make sure the tip does not leak and can maintain the desired volume until dispensed using plunger system.

Micropipettors must be used with plastic disposable tips at all times. Fit a tip tightly onto the end of the barrel of the micropipettor. Press down and twist slightly to ensure an airtight seal.

Tips are usually packed into plastic boxes that can be sterilized by autoclaving. Open the tip box to retrieve a tip, then close the tip box to minimize contact with contaminants in the air.

Some tips have filters similar to the cotton wool plugs on serological pipettes. These tips are often more expensive than regular tips and thus are used for specialized applications. For instance, when measuring volatile chemicals such as chloroform or radioactive liquids such as 32 P-labeled DNA, using filter tips helps prevent the barrel of the micropipettor from getting contaminated.

Hold the micropipettor in a vertical position.

Keeping the micropipettor upright will prevent liquids from running inside and contaminating the barrel of the micropipettor.

The micropipettor has three positions: (1) Rest position, (2) First stop, and (3) Second stop (Figure 6, panel B). The instrument has a two-stop plunger system. The first stop has two functions. The first is to draw in the desired volume of liquid into the tip when releasing the plunger from the first stop to the rest position. The second function is to dispense the majority of liquid from the tip when depressing the plunger from the rest position to the first stop. Further depressing the plunger to the second stop dispenses whatever liquid remains in the tip. Depress the pushbutton on the plunger from the rest position to the first stop. Air equal to the volume of the setting will be displaced.

Immerse the tip into the liquid while holding down the pushbutton to the first stop.

Do not touch the micropipettor itself to the sides of bottles, tubes and flasks otherwise the inside surfaces of these vessels will become contaminated. Only the tips are sterile.

Release the pushbutton slowly to aspirate the liquid into the tip. Stop once the pushbutton is back to the rest position. Wait a moment so liquid can be drawn into the tip.

The volume of liquid in the tip will equal the volume of the setting of the micropipettor.

Viscous liquids such as those containing glycerol require more time to enter the tip.

Remove the tip from the liquid, and visually inspect the tip to confirm that the liquid drawn up has reached the expected level in the tip and there are no air bubbles in the tip.

If necessary, expel the liquid and manually tighten the tips onto the micropipettor. Draw up the liquid and check again.

Place the tip at an angle (10 ° to 45 °) against the wall of the tube receiving the liquid. To expel the liquid, slowly depress the pushbutton on the plunger to the first stop. Wait a moment then press the pushbutton to the second stop to expel any residual liquid in the tip.

Depressing the plunger too quickly may cause the liquid being expelled to splatter or will produce undesirable bubbles in the tube.

Before releasing the plunger to the rest position, remove the tip from the liquid.

Discard tips into designated sharps waste container by pressing the ejection button on the micropipettor.

4. Cleaning Up The Work Space

When finished with an experiment requiring use of aseptic technique, turn off the Bunsen burner, then put away all supplies and reagents. Wipe down the outside surfaces of labware (bottles, micropipettors, pipette tip boxes) with a pre-moistened disinfectant wipe to ensure contaminants are not transferred to the storage location.

Place contaminated glassware and hazardous waste materials into the proper disposal receptacle. Laboratory waste includes labware such as gloves, pipettes, tips, and tubes. Non-infectious hazardous waste is generated when performing experiments with non-pathogenic organisms (BSL-1) while infectious hazardous waste is generated when using pathogenic organisms (BSL-2 or above). Infectious waste must be autoclaved or disinfected before it is discarded. Follow laboratory safety guidelines described in BMBL (5 th Ed.) as well as those provided by OSHA and institutional Environmental Health and Safety departments.

Wipe down the entire work area on the laboratory bench with a pre-moistened disinfectant wipe from the canister, once again allowing the disinfectant to evaporate.

Wash hands thoroughly with antiseptic soap and warm water before leaving the laboratory.

5. Representative Results

A sample application for using serological pipettes to transfer liquids is shown in Figure 7. These pipettes often are used in the microbiology laboratory to prepare media for inoculation with bacterial cultures. For example, sterile flasks first are filled with a specified volume of culture broth, in this case Luria Broth (LB), then a small number of cells (such as E. coli) are added to the media. Using a serological pipette, first the broth must be aseptically transferred from the media bottle to the flask. In this case, 25 ml of LB was added to a 125 ml sterile flask using a 25 ml serological pipette. Next, the broth must be inoculated with E. coli cells. Here, 10 μl of cells were transferred aseptically using a P20 micropipettor from a previously growing culture flask to the 25 ml of fresh LB. The flask is incubated in a growth chamber for a particular amount of time, allowing the cells to replicate (for this example, the E. coli cells were incubated overnight at 37 ଌ on a shaking platform). The result is a turbid bacterial cell culture that can be used for subsequent experiments.

Serological pipettes also may be used to transfer media originally supplied in a bottle to test tubes, or between test tubes, as is done when making dilutions of a bacterial culture. If aseptic technique is not maintained throughout these types of media manipulations, then cultures will become contaminated, and subsequent experiments using those cultures will be delayed because fresh, uncontaminated cultures will need to be prepared. Errors occur because a sterile field is not maintained throughout the procedure. For instance, you may forget to disinfect the laboratory bench or flame the rim of a culture bottle or tube. You may touch the tip of the pipette or set the cap of a bottle or test tube on the bench instead of holding it in your hand. Proper procedure is critical for keeping contamination of media and cultures to a minimum. Figure 8A provides an example of a pure versus contaminated culture of E. coli in a tube containing 5 ml of LB. The left panel shows a culture displaying uniform fine turbidity typical of a pure E. coli culture. In contrast, the right panel shows a contaminated culture in which the growth characteristics deviate from those expected for this bacterial strain.

Technical errors may occur when manipulating serological pipettes resulting in transfer of incorrect volumes of media between test tubes. For instance, you may read the volume on the pipette incorrectly (i.e., top versus bottom of the meniscus) or you may expel the media completely from a TD pipette, which was designed to leave a tiny bit in the tip not to be delivered. When performing a point-to-point delivery of media, you may use the wrong calibration marks and dispense the incorrect volume. Figure 8B shows an example of test tubes with correct versus incorrect volumes of media. The tube on the left contains 3.5 ml of LB measured with a 5 ml serological pipette. The student conducted a point-to-point delivery of the media in which LB was drawn up to the 5.0 ml graduation mark and dispensed to the 1.5 ml mark. The tube on the right contains 2.5 ml of LB measured with a pipette of the same size because the student who performed the point-to-point delivery of media incorrectly dispensed it from the 5.0 ml mark to the 2.5 ml mark. This mistake will result in a bacterial culture that will be at a higher concentration than planned, causing subsequent dilutions to be incorrect. This propagation of errors can result in a failed experiment that would need to be repeated with the correct cell concentrations.

A sample application for using micropipettors to transfer liquids is shown in Figure 9. These pipettors are used for a variety of experiments in molecular biology and microbiology including preparing samples for PCR and gel electrophoresis or inoculating sterile media or buffer with small volumes (less than 1.0 ml) of bacterial cells or phage particles. In the example provided, the student transferred 12.5 μl of TE buffer into a 1.8 ml microcentrifuge tube (left tube in panel A note that dye has been added to the buffer to facilitate visualization of the liquid inside the clear microcentrifuge tubes). This procedure required the student first to select the correct micropipettor, in this case a P20, and next to set the volumeter to the correct volume (panel B). A tip was used that contains a cotton wool plug at the end to prevent possible contamination that could be expelled from the barrel of the micropipettor from reaching the buffer sample in the tip. This precaution is not necessary if care is taken when aspirating liquids into the tips, depressing the plunger slowly so the liquid does not splash into the pipettor barrel. Technical errors may occur that result in transfer of incorrect volumes. For example, you may select the wrong micropipettor for the job or set the volumeter on the correct micropipettor to an incorrect volume. Before immersing the tip into the buffer, you may push the plunger past the first stop, causing an excess of buffer to be drawn into the tip when releasing the plunger. Alternatively, you may not immerse the tip far enough into the buffer, so air is drawn into the tip instead of buffer. You may forget to push the plunger to the second stop when dispensing buffer into the microcentrifuge tube causing less than the desired volume to be released from the tip. The right tube in panel A of Figure 9 shows a microcentrifuge tube containing the incorrect volume of buffer relative to the tube on the left. Instead of dispensing 12.5 μl of buffer, the student dispensed 125 μl. In this case, although the numbers are set identically on the volumeter, the wrong micropipettor was selected for the job (the student used a P200 instead of a P20 panel B) resulting in the delivery of a substantially larger volume of buffer. If this solution was being used to prepare a mixture of reagents for an application such as PCR, then this mistake will change the final concentration of all reagents subsequently added to the same tube. Consequently, it is unlikely that the experiment will be successful, since molecular biology procedures such as PCR require all components to be at specific concentrations for the reaction to work properly.

Because it is not always feasible to ensure micropipettors (especially the inside of the barrel) are sterile, stock solutions can become contaminated causing even troubleshooting efforts to fail when performing experiments. If using micropipettors to transfer sterile solutions, it is strongly recommended that aliquots of stock solutions (media, buffer, water) be made using aseptic technique with serological pipettes. It is common to maintain working stock solutions in 15 ml or 50 ml sterile conical tubes. These are often easier to manipulate while operating a micropipettor and can be replaced with a fresh aliquot of the stock solution if contaminated during volume transfers.

Figure 1. Sterile field created by updraft of Bunsen burner flame. To minimize contamination of sterile solutions and cultures, it is critical that all manipulations be conducted within the sterile field. The rims of glass culture tubes and flasks should be passed through the tip of the blue cone, the hottest part of the flame. Plastic tubes and tips cannot be flamed - these should be pre-sterilized by alternative methods prior to use.

Figure 2. Serological pipettes used for aseptic transfer of liquids. (A) Shown from left to right are drawings of 25 ml, 10 ml, and 5 ml pipettes. (B) Serological pipettes may be plastic or glass. Plastic pipettes are disposable (one-time use) and typically are individually wrapped in paper and plastic sleeves in which all inside surfaces are sterile (left side). Glass pipettes can be used multiple times provided they are cleaned and sterilized between uses these typically are stored in metal canisters (right side).

Figure 3. Serological pipettes are of two types: TC ("to contain") or TD ("to deliver"). Shown is the explanatory label of a TD 5 ml pipette.

Figure 4. Aseptic technique. When aspirating liquids from a bottle, flask, or tube with caps, never place the cap on the bench. Instead, hold the cap in the same hand as pipette aid while manipulating the vessel containing the liquid with the opposite hand as shown.

Figure 5. Meniscus formed when drawing liquid into serological pipette. The volume corresponds to the graduation mark on the pipette where the bottom of the meniscus aligns. In this example, the meniscus aligns with the 2.5 ml graduation mark.

Figure 6. Single channel micropipettor. (A) Shown is a sample micropipettor with a plastic tip attached to the bottom of the barrel tip holder. Indicated are the locations of the volumeter, the thumb wheel for changing the volumeter setting, the barrel tip holder, the tip ejector button, and the pushbutton for the plunger. (B) Two-stop plunger system on a micropipettor.

Figure 7. Using serological pipettes to transfer media into sterile 125 ml flasks. The left flask has 25 ml of media only (LB), while the right flask is a culture of E. coli resulting from inoculating LB with cells then incubating overnight at 37 ଌ. Note how the media in the flask on the right is turbid due to cell growth.

Figure 8. Using serological pipettes to transfer media into sterile test tubes. (A) The left tube contains 5 ml of a pure E. coli culture, while the right tube contains 5 ml of a contaminated bacterial cell culture. Note the differences in growth characteristics between the two cultures. Although both are turbid, the culture on the right has been contaminated with a fungus or other airborne microorganisms giving the culture a different color and consistency from that expected for E. coli cells. (B) The left culture tube contains 3.5 ml LB while the right tube contains only 2.5 ml LB. This volume difference resulted from a mistake made while conducting a point-to-point delivery of media to the tubes.

Figure 9. Using micropipettors to transfer buffer into sterile microcentrifuge tubes. (A) The left microcentrifuge tube contains only 12.5 μl of TE buffer, while the right tube contains 125 μl. Note that a dye has been added to the buffer to facilitate visualization of the liquid inside the clear microcentrifuge tubes. (B) The left volumeter is from a P20 micropipettor, while the right volumeter is from a P200 micropipettor. A common mistake is selecting the wrong micropipettor. Although the numbers are set identically on the P20 and P200 volumeter, selection of the wrong micropipettor results in transfer of incorrect volumes.

Figure 10. Laminar flow hood used to prevent contamination of solutions and cultures. Shown is a biosafety cabinet approved for work with BSL-2 organisms.


Introduction

These days, the application of cells in research laboratories (1, 2), regenerative medicine (3, 4) and biotechnological productions is growing extensively. Cells are used in wide-ranging activities from studies on cell proliferation to the production of biologically active substances. Due to restrictions on the use of laboratory animals by animal protection laws, the use of cell cultures will continue to increase in the future. In order to achieve reproducible results from cells, good cell culture conditions are vital. Despite the importance of bacterial and fungal contaminations in cell culture, they are not such a serious problem because they are usually obvious and easily detected. The most serious problem is mycoplasma infection since these microorganisms are subtle (5, 6).

Mycoplasma has generated considerable interest due to its ability to contaminate cell lines used in research as well as in manufacturing bioproducts (7-13). The lack of a cell wall in mycoplasmas besides their adherence to the cell surface makes them invisible to the naked eye. In the past, the use of animal sera in cell cultures was the main source of Mycoplasma arginini, Mycoplasma hyorhinis or Acholaeplasma laidlawii. Pipetting through mouth is the source of Mycoplasma orale, Mycoplasma fermentans, Mycoplasma salivarium and Mycoplasma pirum (14, 15). Culture supernatants and cell membranes are suitable for the growth of mycoplasma. Mycoplasmas are resistant to commonly used antibiotics and they cannot be detected visually by turbidity of fluid or under the inverted microscope. The frequency and impact of mycoplasma contamination in cell culture have been extensively discussed (12, 15-17). The incidence of single mycoplasma contamination is still high, being 15 to 35% worldwide with extreme incidences of 65 to 80%, whereas incidences of multiple mycoplasma infections with two or more mycoplasma species are between 7 and 60% (9, 10, 18, 19). Mycoplasma contamination can be persistent and difficult to detect for the affected lab (10). Between 5 and 35% of cell cultures are infected with the species Acholeplasma laidlawii, M. arginini, M. fermentans, M. hyorhinis and M. orale (12). It is estimated that about 5 to 30% of the world's cell lines are contaminated with mycoplasmas [5-16%, (14) 5-87%, (15) 25.7% (20) 29% (21) and 23% (22)]. Mycoplasmal contamination influences almost every parameter within the cell culture system (8, 23).

The use of contaminated cells endangers almost all aspects of cell physiology, and often leads to erroneous results or causes the loss of unique cell lines (24, 25). Even though the mycoplasma contamination does not slow down cell metabolism, it may contaminate the final product (such as a vaccine) resulting in the loss of the batch (26). Mycoplasmal infection of cell cultures might often linger for an extended period of time without noticeable cell damage (7, 23, 27). Therefore, it is important to use efficient detection methods to inspect mycoplasma contamination (12, 5, 28, 29). Usually, investigation of mycoplasma is carried out by direct and indirect detection methods (30). The direct method detects the colony growth of mycoplasma on agar however, the indirect detection technique consists of measuring a gene product that is linked to mycoplasmas rather than to the mammalian cells in culture. Moreover, DNA staining of mycoplasma after inoculation of suspected cell onto indicator cells is another type of indirect detection method of mycoplasma (31, 13).

The first source of contamination is usually contaminated media or their components (7, 32). Most cell culture media are not autoclavable therefore, filtration of the media through proper filters to remove mycoplasma is very important to protect cell culture lines. The prevention of mycoplasma contamination can be divided into three categories: cell culture facility, cell culture procedures, and operator technique (33). To prevent mycoplasma effectively, it is recommended that a good aseptic technique be used, accidents in the laboratory be reduced, the laboratory be kept clean, and the positive cultures be discarded (34- 37). In the case of valuable and unique cultures, it is possible to eliminate the contaminants effectively. Therefore, treatment of mycoplasma-positive cell cultures has become a feasible option (18, 27).

What are mycoplasmas?

The name of mycoplasma was chosen because of its mycelated fungi-like structure with a flowering plasma-like structure (38, 39). Mycoplasma is a kind of bacteria. One of the differences between mycoplasma and the other bacteria is the absence of cell wall and their flexible membrane in mycoplasma which results in taking different shapes and consequently difficulties in identifying even under a high powered electron microscope. The small size of mycoplasma (0.15-0.3µm) is the main reason for their escape through filtering systems and also their growth in high concentration in mammalian cell cultures without any turbidity or other obvious symptoms (40). However, there is some exception in the size of mollicutes. Recently it has been shown that different nutritional conditions can affect the size of Acholeplasma laidlawii. Folmsbee et al. showed Acholeplasma laidlawii cultures in tryptic soy broth (TSB) cannot penetrate 0.2µm rated filters to the same degree in other media such as mycoplasma broth or TSB supplemented with 10% horse serum (41).

Ways in which cells are contaminated by mycoplasma

Mycoplasmas can bind to their host cells using special tip organelles. These tip organelles have a high concentration of adhesins, to attach to eukaryotic cells and penetrate the host cell. The lack of a stiff wall in mycoplasma may help it to fuse with the membrane of the host cell and exchange its membrane and cytoplasmic components (38, 42, 43).

Frequency and sources of mycoplasma species

There are a number of different sources for mycoplasma contamination in cell cultures associated with human, bovine and swine species. Personnel in the laboratories are the main sources of M. orale, M. fermentans, and M. hominis. These species of mycoplasmas account for more than half of all mycoplasma infections in cell cultures and physiologically are found in the human oropharyngeal tract (44). M. arginini and A. laidlawii are two other mycoplasmas contaminating cell cultures and originate in fetal bovine serum (FBS) or newborn bovine serum (NBS). Trypsin solutions provided by swines are a major source of M. hyorhinis (45). Figure 1 is a diagram showing the normal host and frequency of different species of mycoplasma occurring in cell culture.


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