This work has been made possible thanks to funding from the Howard Hughes Medical Institute (HHMI). We wish to thank our head of lab, Jue Chen, for reading the manuscript and for her continued support, Donna Tallent for her helpful edits and comments, and Jeff Hennefeld from the Information Technology Department at The Rockefeller University for his help with the video component of this manuscript.

1. Preparations 
<ol>
	<li>General
	<ol>
		<li>Wear a clean lab coat designated to be worn only in the cell culture room and no other parts of the laboratory. 
		NOTE: The lab coat does not need to be sterile.</li>
		<li>Wear new gloves that have not touched any other surfaces. Make sure the gloves fit tightly. Nitrile, powder-free gloves are best. 
		NOTE: The gloves do not need to be sterile.</li>
		<li>To prepare for work, spray the gloves, lab coat sleeves, and interior of the biological safety cabinet with 70% EtOH. Wipe the working surface and glass panel dry with a lint-free paper towel. 
		NOTE: Using 70% EtOH kills bacteria with the highest efficiency16.&#160;The paper towel does not need to be sterile.</li>
		<li>Keep water baths inside the cell culture room and only use these for warming culture media or thawing cells. Drain and wash the water baths once a week, following the manufacturer&#8217;s instructions for cleaning.</li>
	</ol>
	</li>
	<li>Inside the biological safety cabinet
	<ol>
		<li>Limit the number of items brought into the cabinet. Do not interrupt the airflow inside the biological safety cabinet by blocking the front or back grills. 
		NOTE: See Figure 1 for an explanation of how air flows inside a cabinet.</li>
		<li>Spray all the items placed inside the cabinet with 70% EtOH and wipe them dry. Begin by spraying the top of the media bottle and working down. Similarly, with a clean paper towel, wipe it dry, progressively working the way to the bottom. Do not go back up toward the cap.</li>
		<li>If the cabinet is large enough to accommodate serological pipettes, they can be placed inside, otherwise they can be stored in a receptacle mounted on the outside of the cabinet. Check either side of the encased pipette for holes, tears, or punctures in the packaging before use. Do not rip off the wrapping. Instead, gently peel the ends of the wrapping, insert the serological pipette into a pipette aid, and remove the wrapping in one fluid motion.</li>
		<li>Do not hover over open bottles or flasks; reach over open bottles or flasks, or open items over the tops of already opened items in the biological safety cabinet. 
		NOTE: The airflow inside the cabinet pushes down on the work surface, so any contamination present on the sleeve, for example, may enter the cell cultures.</li>
		<li>Do not pour liquids. Instead, add them using a serological pipette. After supplementing the media, mix the contents thoroughly and initial the bottle. Also, ensure to include a label for what the media was supplemented with.</li>
	</ol>
	</li>
</ol>
2. Working with adherent cell lines
<ol>
	<li>If a plastic flask is needed, spray the entire bag and place it inside the cabinet.</li>
	<li>Use autoclaved glass pipettes or disposable sterile plastic pipettes to aspirate the media or washing solutions. Carefully remove the metal cap from the storage container. Isolate one glass pipette by gently shaking the container at an angle. When reaching into the container, avoid touching any other pipettes. 
	NOTE: Handle the chosen pipette from one end only.&#160;</li>
	<li>Quickly replace the caps on the bottles as soon as possible. Place the caps on the work surface upside down so that the rim does not touch the work surface. Do not grab the cap from the top or bottom; instead, touch the caps from the sides.</li>
	<li>When aspirating liquids, use a vacuum trap flask located outside of the cabinet in a secondary container on the floor. 
	NOTE: Do not throw any liquid waste in biohazard waste bags as the bags will leak. Waste will be created while working inside the cabinet. Moving hands in and out of the cabinet too often will interrupt the airflow. Leave any waste inside the cabinet temporarily. Place it off to the side so it will not interrupt the work.</li>
	<li>Generously spray the gloves with 70% EtOH any time they become dry. Rub the hands together so the gloves are not dripping wet.</li>
	<li>If a serological pipette mistakenly touches something in the cabinet, do not hesitate to throw it out. Start anew with a clean serological pipette instead of using one that may be contaminated.</li>
</ol>
4. Checking and storing cells 
<ol>
	<li>Before placing cells in the incubator, check to see how they look under the microscope. If the cells have been thoroughly suspended, single cells should be observed.</li>
	<li>Do not speak, sneeze, cough, or breathe heavily into the incubators. Quickly open and close the incubator doors. Leaving doors open for longer than necessary may allow contaminants present in the air to enter the incubators. 
	NOTE: Wearing a mask while working in the cell culture room can help since mycoplasma may be present in the human mouth. Avoid the use of cell phones in the cell culture room, as talking is not recommended.</li>
	<li>Ensure caps on all the bottles are tightly closed before removing the bottles from the cabinet.</li>
	<li>Store cell culture media in the dark at 4 &#176;C when not in use since it is light sensitive.</li>
	<li>Spray the interior of the cabinet with 70% EtOH again after the cell culture work is complete and wipe the surface dry with a paper towel. Empty the biohazard trash waste bags. Repeat this process and replace the gloves when switching to a different cell line.</li>
</ol>
5. Working with suspension cell lines
<ol>
	<li>For suspension cells grown in glass flasks, ensure the gloves are sprayed thoroughly with 70% EtOH, then touch the aluminum foil with the wet gloves and spray only the bottom of the flask before placing it inside the cabinet.</li>
	<li>When taking a sample for cell counting, remove only one 1.5 mL tube from its container. Do not touch any other tubes. Place the cap upside down on the working surface. Do not touch the inside rim. Handle it with care from the sides and replace it once finished.</li>
	<li>Carefully remove the double-folded piece of aluminum foil that covers the entire neck of the flask. Handle the glass flask from the bottom only&#8212;do not touch it from the neck&#8212;once the foil is off. Use a 1 mL serological pipette to take a sample for cell counting.</li>
	<li>Do not let media drip down the side of the flasks; if it does, spray a paper towel with 70% EtOH and clean it up right away.</li>
	<li>Ensure caps and aluminum foil are tightened before removing bottles or flasks from the biological safety cabinets.</li>
</ol>
6. Cell incubation 
<ol>
	<li>Use separate incubators for different cell types to prevent cross-contamination of the different cell types.</li>
</ol>
7. Liquid waste collection
<ol>
	<li>Collect liquid waste in a vacuum trap flask located outside the cabinet on the floor in a secondary container labeled &#8216;Waste&#8217;. 
	NOTE: The hose is connected to a high-efficiency particulate absorbing (HEPA) filter, which is replaced on a monthly basis.</li>
</ol>
8. Cleanup
<ol>
	<li>Remove the biohazard waste bag and wash the glass flasks as soon as possible. Keep a glass-washing protocol by the sink. See Supplementary File 1 for the washing protocol.</li>
	<li>Autoclave the cell culture glassware instead of sending it to a glass washing facility. Keep it separate from glassware in the main area of the lab. 
	NOTE: SF-9 cells leave a rim of dead cells on the side of flasks if the glass is not scrubbed well. See Supplementary File 1 for the autoclave protocol.</li>
</ol>
9. Organization 
<ol>
	<li>Organize the cell culture room so that all the supplies are located in one area, thereby minimizing the need for lab members to leave the room in search of supplies.</li>
	<li>Label any plastic bottles used to harvest cells and reused in cell culture afterward as &#8216;For Cell Culture Use Only&#8217;. Use the designated cell culture bottles for one specific cell type. Store the bottles in the cell culture room for easy access.</li>
</ol>
10. Identifying bacteria, fungi, and mycoplasma contamination 
NOTE: Not following the workflow above can lead to bacterial, fungi, and mycoplasma contamination.
<ol>
	<li>Always before beginning work, observe flasks for heavy turbidity, extra growth in the form of fuzzy balls, and dense accumulations of cells on the side, all of which are indicators of contamination. 
	NOTE: An experienced eye can tell the difference between turbidity caused by microbial contamination versus the actual cells. While cells cause the normally clear media to appear cloudy, bacterial contamination causes heavy turbidity with white color.
	<ol>
		<li>Visually identify mold contamination by noting the appearance of round, fuzzy balls floating in the media (see Figure 2).</li>
		<li>Visually identify bacterial contamination if media is cloudy, white or turbulent (see Figure 2).</li>
		<li>Identify mycoplasma contamination by doing a monthly mycoplasma test. One PCR-based test instructs users to take a 1.5 mL sample of cells, perform a cell count using 10 &#956;L, dilute to 0.08 x 106 cells/mL, spin down the cells, lyse the cells using the buffer contained in the kit, and spin down one final time. The supernatant is incubated with primers that amplify a range of mycoplasma species. Run a DNA gel to visualize the bands. No bands mean that mycoplasma is not identified. 
		NOTE: This major contaminant can be present in the human mouth. It is good practice to test for mycoplasma contamination in cells after thawing them and before using them for experiments. Afterward, monitor the cells for mycoplasma contamination once a month. Many companies offer mycoplasma test kits. Pick one that is suitable.</li>
	</ol>
	</li>
</ol>

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N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+culture+contamination.">Cell culture contamination.</a> Methods in Molecular Biology. 731, 79-91 (2011).</li><li>Young, L., Sung, J., Stacey, G., Masters, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+Mycoplasma+in+cell+cultures.">Detection of Mycoplasma in cell cultures.</a> Nature Protocols. 5 (5), 929-934 (2010).</li><li>Barth, O. 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The author does not have any conflicting interests.

While contamination is one of the primary concerns when performing cell culture work, the practices and techniques outlined in this manuscript will help mitigate the risks. The critical steps include wearing a clean lab coat, which is only used in the cell culture room, using clean, powder-free gloves that are sprayed with 70% EtOH often and which are changed when switching between cell lines, encouraging each individual to not share media bottles, cleaning the cabinet thoroughly prior to and after finishing work, neatly unwrapping serological pipettes, avoiding prolonged close contact with open bottles or flasks, not sharing media bottles between multiple cell lines, and quickly opening and closing incubator doors. Additionally, washing and autoclaving one&#8217;s own glassware ensures quality control, thereby reducing the probability of introducing outside contaminants. Quality control methods include using autoclave tape to indicate if the sterilizer has reached the proper temperature of 121 &#176;C, regular preventative maintenance of the equipment, and visually inspecting the flask to ensure it has been scrubbed properly18. Another important point is to use separate incubators for different cell types to ensure that contamination from one cell type is not be transferred to others and to also ensure cross-contamination with cells does not occur.
Bacterial and fungal infections are the two most common intruders in cell cultures. One of the major contaminants, M. orale, is the most common mycoplasma species in the human mouth and also &#8220;represents the single most common isolate accounting for 20%&#8211;40% of all mycoplasma infections in cell cultures&#8221;4. In other words, lab personnel are the major source of this contamination. It is always a concern in the cell culture room since mycoplasma is a small, slow-growing microorganism lacking a rigid cell wall that can pass through 0.45 &#956;m filters4. Studies show the incidence of mycoplasma contamination is 15%&#8211;35% of continuous human or animal cell lines4. Furthermore, statistics indicate that about 5% to 30% of the world&#8217;s cell lines are contaminated with mycoplasmas6. Unfortunately, mycoplasma is resistant to most antibiotics used for cell culture, and infection can affect cell physiology and metabolism4,6. Although contamination does not slow down cell metabolism, it may contaminate the final product6. Infections can stick around without lab members noticing cell damage. If contamination does occur, the cost of thawing new cells, the time spent propagating the new cultures, the expensive media that has been wasted, decontaminating incubators, and the amount of time colleagues spend waiting to begin their experiments again, is enormous. Our lab estimates the total lost time accounts for 2 weeks and the total cost associated with contamination of a typical human mammalian cell protein expression of 8 L is $1,400. The practices and techniques outlined in this manuscript offer good preventative measures to mitigate additional costs, lost cells, and downtime.
Modifications of these practices are not recommended. All lab members should be trained prior to working in the cell culture room and then receive refresher training every year7. Lab members should also organize the cell culture room so that all the necessary supplies are in one area, thereby minimizing the need for lab members to exit the cell culture room in search of supplies. &#160;
Limitations of the techniques presented are observed if contamination comes from external sources such as serum, incubators, malfunctioning autoclaves, dirty lab coats, or the source of the cells. Troubleshooting of the technique may be necessary if contamination occurs despite strict adherence to this protocol. Many external factors can contribute to this type of contamination. Lab members need to be prudent and check external sources. For instance, the lot of fetal bovine serum (FBS) purchased from the supplier may have been contaminated with mycoplasma. Compared to the 1950s, this is now a rare occurrence, but it can still happen6. All media bottles should be thrown out if any contamination is noticed, and the protocol restarted by thawing new cells and using new media. The contamination may have already been present inside the incubator if it is mold or bacteria; the other flasks inside the same incubator should be checked to see if bacterial or mold growth is observed. All the contaminated flasks should be thrown out and the incubator decontaminated. If no contamination is found, the autoclave should be checked for malfunctioning errors; the glassware may not have been autoclaved properly in the first place. The autoclave tape may change color despite the autoclave not reaching 121 &#176;C. Afterward, expiration dates on the antibiotic solutions used should be checked; new solutions may need to be purchased. Finally, the source of the cells should be considered; were they from a trusted lab supplier or borrowed from another lab? Cells should always be purchased from a trusted lab supplier and should never be borrowed from another lab. Finally, lab coats should be washed to ensure their cleanliness19. Decontamination of incubators should be considered whenever a flask has been contaminated. Bacteria cells or mold spores must not be allowed to multiply and continue spreading contamination.
Existing methods reflect upon the elimination of contamination. The methods outlined in this manuscript focus on prevention rather than the elimination of contamination. Several procedures exist regarding using antibiotics to remove mycoplasma20. However, the use of antibiotics is risky as they may not completely eliminate the infection and &#8220;may permit resistant organisms to develop&#8221;12. In fact, &#8220;72% of cultures grown continuously in antibiotics were shown to be mycoplasma-positive while only 7% grown in the absence of antibiotics were infected&#8221;12. This data emphasizes the idea that while the use of antibiotics is recommended and can be helpful, overuse or complete reliance on antibiotics is not beneficial. Furthermore, using antibiotics to remove contamination may only allow the problem to persist. Antibiotic solutions are not necessary; if the techniques in this paper are followed, contamination should be successfully prevented.
The practices and techniques outlined in this manuscript can be used in the future to maintain an aseptic environment when performing the monthly mycoplasma test and when making homemade competent bacterial cells. Both protocols require a clean environment, similar to cell culture work, so that the final product is not contaminated.&#160;

Cell culture has many uses in the research laboratory. Since the origins of cell culture in the early 20th century, cell lines have helped advance science. Cell lines have several advantages; various cell lines can help researchers study cell biology, produce baculovirus for further studies, or produce large quantities of a protein of interest, to name a few1. Some additional uses include studying tissue growth, helping to advance vaccine development, toxicology research, studying the role of genes in healthy organisms and diseased models, and the production of hybrid cell lines2,3. Cell lines can also enable drug production3. Proper aseptic techniques are necessary when working with cell lines; the practices and techniques outlined in this manuscript are applicable to research laboratories where cell culture work is performed. Other laboratory settings are not discussed.
Contamination is often the primary concern when performing cell culture work. In the context of this paper, contamination generally refers to fungi and bacteria. The overall goal of the method outlined in this paper is to thoroughly describe the best practices for avoiding contamination. All lab members should adhere to these practices when working in a research laboratory&#8217;s cell culture room. Laboratories should ensure all workers are active participants in using these best practices to prevent contamination. The knowledge of the correct practices and techniques will help ensure cell cultures remain viable, healthy, and free from contamination. The development of this technique is based on research of the literature, seven years of experience working with cell cultures, and the need for a method that both novices and experienced cell culture workers can refer to on an annual basis.
There is a need for a clear, standardized technique that all research cell culture laboratories should follow. Much of the literature on cell culture contamination discusses the detection of mycoplasma, aseptic techniques, sources of contamination, elimination of contaminants, and prevention by use of antibiotics and regular testing4,5,6,7,8. While this information is helpful, there are no videos present in the literature that demonstrate the proper cell culture techniques one should follow. The advantage of the practices presented over alternative techniques is a focus on preventing contamination before it happens, rather than detecting and correcting mistakes later. Moreover, a thorough demonstration of aseptic techniques, a discussion about preventing fungi and bacterial growth, and information regarding biosafety cabinet airflow are valuable for both novice and experienced cell culture workers alike.
Bacteria and fungi are the two most common types of contaminants. Within the bacteria category, mycoplasma is a major concern due to its small size and ability to proliferate while remaining unnoticed. They are self-replicating organisms with no rigid cell wall that rely on eukaryotic cells to grow. They have reduced metabolic capabilities and can multiply greatly while remaining unrecognized in the routine visual inspection of cell cultures and regular microscopic analysis, although transmission electron microscopy can detect mycoplasma9,10. Moreover, they can pass through microbiological filters10. Cell culture medium provides mycoplasma with nutrients, although unfortunately supplementing media with antibiotics does not affect mycoplasma10. One should note that, in general, it is not necessary to supplement media with antibiotics; proper techniques should suffice in keeping contamination at bay. Infection with mycoplasma does not lead to immediate cell death, but it is concerning to researchers as it affects data reproducibility and quality.&#160;
All lab personnel should strictly adhere to good cell culture practices. Cultures should be tested for mycoplasma after they&#8217;ve been newly purchased, while they are currently being grown, before cryopreservation, and when thawed from liquid nitrogen2,10. Different tests are available on the market using polymerase chain reaction (PCR), enzyme-linked immunosorbent assay (ELISA), or immunostaining.3 The literature indicates that &#8220;human isolates represent a large percentage of the mycoplasma contaminants found in cell culture&#8221;5. Although more than 200 mycoplasma species have been described, about six of these account for most infections. These six species are M. arginini, M. fermentans, M. hominis, M. hyorhinis, M. orale, and Acholeplasma laidlawii10. As with other types of contamination, air and aerosols bring these into cell cultures5. This is echoed in other papers since the &#8220;human operator is potentially the greatest hazard in the laboratory&#8221;7. Although this is done through human error, the risk can be eliminated if a standard procedure is followed. Shedding from personnel is not restricted to only mycoplasma contamination; cell cultures in one lab are usually infected with the same mycoplasma species, indicating that contamination spreads from one flask to another due to improper cell culture techniques10.
The prevention of cross-contamination is also another reason why proper cell culture techniques should be followed. It is noted that at least 15%&#8211;18% of cell lines worldwide may be cross-contaminated or misidentified11,12. In addition to testing cell lines for mycoplasma contamination, they should also be tested for cross-contamination10. For human cell lines, cell line authentication by an inexpensive DNA-based technique called short tandem repeat (STR) profiling is the current international reference standard, as it&#8217;s an easy way to confirm cell line identity2,10,13,14. STR can identify mislabeled or cross-contaminated cell lines, but it cannot detect incorrect tissue origin10,13,14. The validity of research data can be compromised if cell lines are mislabeled, wrongly identified, or contaminated13.&#160;Similar to other types of contamination, cross-contamination can occur due to poor technique causing aerosols to spread, mistaken contact leading to the wrong cell type entering a flask, or using the same media bottle and reagents with different cell lines10. No sharing of media bottles should occur; sharing one bottle of media between two different cell lines can allow those cell populations to be mixed, leading the faster growing cell type to completely take over the flask. This replacement is not noticeable and leads to mislabeling and misidentification2. A cell line can also be mistaken for another if cultures are confused during handling or labeling10. Careful attention should be paid to keeping reagents, media, and flasks separate from one another. Each lab member should have their own media bottles; no sharing should occur between lab members. Cell lines themselves should be purchased from a qualified cell bank and provider. Laboratories should not share cells. Studies show that, although STR and mycoplasma tests are regularly used, many research papers in the literature have already used misidentified or contaminated cell lines15. Sifting through the research to find these problematic papers and retroactively inform readers about this matter is cumbersome. Prevention is the best way to ensure this problem does not occur in the first place.
The simple action of spraying items with 70% EtOH can kill organisms; 70% EtOH works by denaturing proteins and dissolving lipids in the most commonly contaminating organisms, including bacteria and fungi16. Studies have shown that 70% is the most effective concentration; surface proteins do not coagulate rapidly with 70% EtOH so they can enter the cell, while the water it contains is necessary for the denaturing process of proteins. Due to the concentration difference of water and alcohol on either side of the cell wall, 70% EtOH enters the cell to denature both enzymatic and structural proteins. If mold growth is observed in flasks, the entire incubator must be decontaminated by first spraying it with 70% EtOH and wiping it dry, followed by a 16 h overnight incubation at 60 &#176;C17. This kills most mold and any bacteria.
The main advantage of prevention practices over alternative techniques of eliminating contamination after it occurs is that by preventing contamination early on, laboratory workers can be sure their cell cultures are healthy and there will be no high costs associated with the decontamination of incubators or discarding of cell cultures. The elimination of mycoplasma contaminants after, for instance, is not efficient7. Taking the time early on to ensure laboratory personnel are properly trained, the cell culture room is self-contained, and a standard procedure is used will save time and money.

<table><tbody><tr><td>DPBS</td><td>Gibco</td><td>14-190-144</td><td></td></tr><tr><td>DMEM F-12 Media</td><td>ATCC</td><td>30-2006</td><td></td></tr><tr><td>Glass Baffled Flask</td><td>Pyrex&amp;nbsp;</td><td>09-552-40</td><td></td></tr><tr><td>Glass Pipettes</td><td>Fisher&amp;nbsp;</td><td>13-678-6B</td><td></td></tr><tr><td>Pipette Aid</td><td>Drummond</td><td>13-681-15A&amp;nbsp;</td><td></td></tr><tr><td>Serological Pipette</td><td>Corning</td><td>07-200-573</td><td></td></tr><tr><td>T75 flask</td><td>Corning</td><td>07-202-004</td><td></td></tr><tr><td>Trypsin</td><td>Gibco</td><td>25-300-054</td><td></td></tr><tr><td>*Items may vary because this video is about general cell culture techniques</td><td></td><td></td><td></td></tr></tbody></table>

cell culture techniques and practices to avoid contamination by fungi and bacteria in the research cell culture laboratory

Cell culture is a delicate skill necessary for growing human, animal, and insect cells, or other tissues, in a controlled environment. The goal of the protocol is to emphasize the correct techniques used in a research laboratory to prevent contamination from fungi and bacteria. Special emphasis is placed on avoiding mycoplasma contamination, a major concern in the cell culture room due to its small size and resistance to most antibiotics used for cell culture. These same techniques ensure continuous growth and maintain healthy cells. For new and experienced cell culture users alike, it&#8217;s important to consistently adhere to these best practices to mitigate the risk of contamination. Once a year, laboratories should review cell culture best practices and follow-up with a discussion or additional training if needed. Taking early action to prevent contamination in the first place will save time and money, as compared to cleaning up after contamination occurs. Universal best practices keep cell cultures healthy, thereby reducing the need to constantly thaw new cells, purchase expensive cell culture media, and reducing the amount of incubator decontamination and downtime.&#160;

If the proper cell culture techniques and practices outlined in this paper are not followed, contamination by fungi and bacteria may occur in the research cell culture laboratory. Figure 2 shows flasks containing contamination in both the suspension and adherent cultures.
When not following aseptic techniques, mold contamination may occur 2&#8211;3 days later. Round fuzzy balls floating in the media are noticeable in suspension cells, while mold growth in attached cells can be observed as large, irregular, white, or green patches.
For bacteria, contamination is observed the following day. The media is turbulent, white, and cloudy. The white color is typical of bacterial cells, which multiply much more rapidly than cell lines. An experienced eye is able to tell the difference between non-contaminated media and contaminated media. For attached cells, one can compare a bottle of unopened media with a flask to check if any turbulence is seen in the flask.
Inside the biological safety cabinet, the number of items should be kept at a minimum. Avoid placing items on the back and front grills (see Figure 3). In this cabinet, a set of pipettes, tips, autoclaved glass pipettes, a pipette aid, and markers are inside. The working area in the middle is clear. Keeping cabinets organized in this way is a good idea. In addition, the operator should wear a clean lab coat and gloves before beginning work. A spray bottle with 70% EtOH should be kept nearby so the operator can spray their gloves often. The skin is covered by gloves or a lab coat. The operator should adjust their chair so their arms are at a 90&#176; angle when working inside the cabinet, and items should be within easy reach inside the cabinet (see Figure 4).
Many species of mycoplasma can be reliably identified using a PCR-based assay. Figure 5 shows the results of a negative mycoplasma test. The band on the left shows the molecular weight standards for DNA. The four bands on the right are positive controls. No bands appear under the tested cell types because mycoplasma was not detected.
If the media contains pH indicators, it will be red for the optimal pH value of cells at 7.4. Once the cells grow, the media will change color from red to yellow3. This color change can also occur if bacteria take over the flask and overgrow (Figure 6). The yellow color indicates the pH is low. Observing a new, unopened bottle of media next to a flask is an objective way to observe contamination in attached cells. For suspension cultures, the user can closely observe the flask for any growths floating in the media or whether a thick ring of overgrown bacterial cells is present around the inside of the glass flask. For both cell types, a small sample can be taken and observed under the microscope. If other growths or cell shapes are observed, especially if the cells are moving, then this is an indicator of contamination (Figure 7). A thorough cleansing of the hemocytometer should be performed prior to cell counting, as this type of debris may be present only on the hemocytometer and not in the cell cultures themselves.&#160;
The smell may be another indicator of contamination in an incubator. Bacterial overgrowth has a typical smell that an experienced cell culture user will notice. The smell always coincides with a contaminated flask, although one infected flask may not always cause the entire incubator to smell.
Mold contamination tends to be more prevalent in HEK 293 S cells grown in suspension. Bacterial contamination is more common in SF-9 cells. This may be because RIC cells are grown with humidity, thereby leading to moisture accumulation in the incubators. SF-9 cells are grown without humidity, so the environment is drier. The rate of contamination in adherent cultures is less than the rate of contamination in suspension cultures. This may be due to the smaller flask size, the non-reusable nature of the flask, or the vented cap instead of the use of aluminum foil.
Mycoplasma cannot be observed with the naked eye nor with a regular light microscope, although specialized transmission electron microscopy can detect mycoplasma. A mycoplasma identification kit should be used to test the cell cultures monthly. Many species of mycoplasma can be reliably identified using a PCR-based assay. A brief description about how this PCR test is performed can be found in the protocol section, and more information about mycoplasma can be found in the discussion section.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64769/64769fig01.jpg" /> 
Figure 1: How air flows in a biosafety cabinet. Biological safety cabinets pull contaminated air from the room itself and from the cabinet through the front and back grills. This air goes under the metal working surface, toward the back of the cabinet, and up to the top of the unit where a HEPA filter is located. There, the air passes through the filter and gets filtered. This clean air pushes down on the work surface. Due to how the flow of filtered air is pushed down inside the cabinet, it is good practice to not hover. For example, it is undesirable to have a sleeve on top of an open bottle and risk having any potential contaminants be pushed into the media. The number of items brought inside the cabinet should be kept at a minimum, and items should not be placed on the front or back grills in order not to interrupt the air flow. Moving arms in and out of the cabinet too quickly can also disturb the airflow. Created for NuAire, Inc., by Jeff Kaphingst, Jeff the Designer, LLC. Used with permission. <a href="https://www.jove.com/files/ftp_upload/64769/64769fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64769/64769fig02.jpg" /> 
Figure 2: Non-contaminated suspension and adherent cells and cells contaminated with mold or bacteria. The first image on the left contains insect cells (SF-9 cells) that are not contaminated. The second image shows another flask of these cells contaminated with mold. The third flask was contaminated by bacteria, as can be noted by the thick, white, cloudy appearance. The second and third flasks were contaminated because none of the proper cell culture techniques and practices were followed. All flasks were prepared on the same day. Growth was observed the next day for bacterial contamination and 2 days later for mold contamination. Non-contaminated adherent cells are shown (Hek293 cells) along with mold and bacterial contamination in adherent cultures. Mold contamination is shown in the second line in a round Petri dish. The photo is taken from the top of the dish. <a href="https://www.jove.com/files/ftp_upload/64769/64769fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64769/64769fig03.jpg" /> 
Figure 3: Cell culture cabinet organization. Inside the biological safety cabinet, the amount of items should be kept at a minimum. Placing items on the back and front grills should be avoided. In this cabinet, a set of pipettes, tips, autoclaved glass pipettes, a pipette aid, and markers are inside. The working area in the middle is clear. <a href="https://www.jove.com/files/ftp_upload/64769/64769fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64769/64769fig04.jpg" /> 
Figure 4: The correct way for an operator to work under the flow hood. An operator should wear a clean lab coat and gloves before beginning work. A spray bottle with 70% EtOH should be kept nearby so the operator can spray their gloves often. The skin is covered by the gloves or lab coat. The operator should adjust their chair so their arms are at a 90&#176; angle when working inside the cabinet. Items should be within easy reach inside the cabinet. <a href="https://www.jove.com/files/ftp_upload/64769/64769fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64769/64769fig05.jpg" /> 
Figure 5: A negative mycoplasma test result. Many species of mycoplasma can be reliably identified using a PCR-based assay. The band on the left shows the molecular weight standards for DNA. The four bands on the right are positive controls. No bands appear under the tested cell types because mycoplasma was not detected. <a href="https://www.jove.com/files/ftp_upload/64769/64769fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/64769/64769fig06.jpg" /> 
Figure 6: Normal media color change from red to yellow. Cell culture media changes the color from red to yellow if pH indicators are present. The yellow color indicates the pH is low and the media should be replaced. <a href="https://www.jove.com/files/ftp_upload/64769/64769fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/64769/64769fig07.jpg" /> 
Figure 7: Contaminants observed under the light microscope. If other growths or cell shapes are observed under the light microscope while performing cell counts, then this may be an indicator of contamination. It should be noted that a thorough cleansing of the hemocytometer should be performed prior to cell counting as this type of debris may be present only on the hemocytometer and not in the cell cultures themselves. <a href="https://www.jove.com/files/ftp_upload/64769/64769fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary File 1: Appendix <a href="https://www.jove.com/files/ftp_upload/64769/Supplemental Materials- Appendix .docx" target="_blank">Please click here to download this File.</a>

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Cell Culture Techniques and Practices to Avoid Contamination by Fungi and Bacteria in the Research Cell Culture Laboratory

This protocol presents essential cell culture techniques and practices to be used in the research cell culture laboratory to avoid contamination by fungi and bacteria. Within the category of bacteria, special emphasis will be placed on preventing mycoplasma contamination.&#160;&#160;

Cell culture is a delicate skill necessary for growing human, animal, and insect cells, or other tissues, in a controlled environment. The goal of the ...

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Nanyang Technological University

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JoVE Journal

Biology

13.4K Views.  The Rockefeller University. This protocol presents essential cell culture techniques and practices to be used in the research cell culture laboratory to avoid contamination by fungi and bacteria. Within the category of bacteria, special emphasis will be placed on preventing mycoplasma contamination.  

Video: Cell Culture Techniques and Practices to Avoid Contamination by Fungi and Bacteria in the Research Cell Culture Laboratory

Culturing Human Pluripotent and Neural Stem Cells in an Enclosed Cell Culture System for Basic and Preclinical Research

This paper describes how to use a custom manufactured, commercially available enclosed cell culture system for basic and preclinical research. Biosafety cabinets (BSCs) and incubators have long been the standard for culturing and expanding cell lines for basic and preclinical research. However, as the focus of many stem cell laboratories shifts from basic research to clinical translation, additional requirements are needed of the cell culturing system. All processes must be well documented and have exceptional requirements for sterility and reproducibility. In traditional incubators, gas concentrations and temperatures widely fluctuate anytime the cells are removed for feeding, passaging, or other manipulations. Such interruptions contribute to an environment that is not the standard for cGMP and GLP guidelines. These interruptions must be minimized especially when cells are utilized for therapeutic purposes. The motivation to move from the standard BSC and incubator system to a closed system is that such interruptions can be made negligible. Closed systems provide a work space to feed and manipulate cell cultures and maintain them in a controlled environment where temperature and gas concentrations are consistent. This way, pluripotent and multipotent stem cells can be maintained at optimum health from the moment of their derivation all the way to their eventual use in therapy.

Here is a protocol to grow pluripotent stem cells (PSC) and neural stem cells (NSC) in an enclosed cell culture system that permits maximum sterility and reproducibility, replacing the traditional biosafety cabinet and incubator. This equipment meets clinical good manufacturing practice (cGMP) and clinical good lab practice (cGLP) guidelines.

This paper describes how to use a custom manufactured, commercially available enclosed cell culture system for basic and preclinical research. Biosafety ...

Developmental Biology

Aseptic Laboratory Techniques: Volume Transfers with Serological Pipettes and Micropipettors

Microorganisms are everywhere - in the air, soil, and human body as well as on inanimate surfaces like laboratory benches and computer keyboards. The ubiquity of microbes creates a copious supply of potential contaminants in a laboratory. To ensure experimental success, the number of contaminants on equipment and work surfaces must be minimized. Common among many experiments in microbiology are techniques involving the measurement and transfer of cultures containing bacterial cells or viral particles. To do so without contacting non-sterile surfaces or contaminating sterile media requires (1) preparing a sterile workspace, (2) precisely setting and accurately reading instruments for aseptic transfer of liquids, and (3) properly manipulating instruments, cultures flasks, bottles and tubes within a sterile field. Learning these procedures calls for training and practice. At first, actions should be slow, deliberate, and controlled with the goal being for aseptic technique to become second nature when working at the bench. Here we present the steps for measuring volumes using serological pipettes and micropipettors within a sterile field created by a Bunsen burner. Volumes range from microliters (&mu;l) to milliliters (ml) depending on the instrument used. Liquids commonly transferred include sterile broth or chemical solutions as well as bacterial cultures and phage stocks. By following these procedures, students should be able to: &#8226;Work within the sterile field created by the Bunsen burner flame. &#8226;Use serological pipettes without compromising instrument sterility.&#8226; Aspirate liquids with serological pipettes, precisely reading calibrated volumes by aligning the meniscus formed by the liquid to the graduation marks on the pipette. &#8226;Keep culture bottles, flasks, tubes and their respective caps sterile during liquid transfers. &#8226;Identify different applications for plastic versus glass serological pipettes. &#8226;State accuracy limitations for micropipettors. &#8226;Precisely and accurately set volumes on micropipettors. &#8226;Know how to properly use the first and second stop on a micropipettor to aspirate and transfer correct volumes.

When working in a laboratory, it is imperative to minimize sources of contamination. Aseptic technique refers to procedures that permit transfer of cultures and reagents while avoiding contact with non-sterile surfaces. Serological pipettes and micropipettors are used to measure precise volumes without compromising sterility of solutions used in experiments.

Microorganisms are everywhere - in the air, soil, and human body as well as on inanimate surfaces like laboratory benches and computer keyboards. The ...

Aseptic Laboratory Techniques: Plating Methods

Microorganisms are present on all inanimate surfaces creating ubiquitous sources of possible contamination in the laboratory. Experimental success relies on the ability of a scientist to sterilize work surfaces and equipment as well as prevent contact of sterile instruments and solutions with non-sterile surfaces. Here we present the steps for several plating methods routinely used in the laboratory to isolate, propagate, or enumerate microorganisms such as bacteria and phage. All five methods incorporate aseptic technique, or procedures that maintain the sterility of experimental materials. Procedures described include (1) streak-plating bacterial cultures to isolate single colonies, (2) pour-plating and (3) spread-plating to enumerate viable bacterial colonies, (4) soft agar overlays to isolate phage and enumerate plaques, and (5) replica-plating to transfer cells from one plate to another in an identical spatial pattern. These procedures can be performed at the laboratory bench, provided they involve non-pathogenic strains of microorganisms (Biosafety Level 1, BSL-1). If working with BSL-2 organisms, then these manipulations must take place in a biosafety cabinet. Consult the most current edition of the Biosafety in Microbiological and Biomedical Laboratories (BMBL) as well as Material Safety Data Sheets (MSDS) for Infectious Substances to determine the biohazard classification as well as the safety precautions and containment facilities required for the microorganism in question. Bacterial strains and phage stocks can be obtained from research investigators, companies, and collections maintained by particular organizations such as the American Type Culture Collection (ATCC). It is recommended that non-pathogenic strains be used when learning the various plating methods. By following the procedures described in this protocol, students should be able to: 
 
&#9679; Perform plating procedures without contaminating media. 
&#9679; Isolate single bacterial colonies by the streak-plating method. 
&#9679; Use pour-plating and spread-plating methods to determine the concentration of bacteria. 
&#9679; Perform soft agar overlays when working with phage. 
&#9679; Transfer bacterial cells from one plate to another using the replica-plating procedure. 
&#9679; Given an experimental task, select the appropriate plating method.

When working with media and reagents used to culture microorganisms, aseptic technique must be practiced to ensure contamination is minimized. A variety of plating methods are routinely used to isolate, propagate, or enumerate bacteria and phage, all of which incorporate procedures that maintain the sterility of experimental materials.

Microorganisms are present on all inanimate surfaces creating ubiquitous sources of possible contamination in the laboratory. Experimental success relies ...

Complementary Use of Microscopic Techniques and Fluorescence Reading in Studying Cryptococcus-Amoeba Interactions

To simulate Cryptococcus infection, amoeba, which is the natural predator of cryptococcal cells in the environment, can be used as a model for macrophages. This predatory organism, similar to macrophages, employs phagocytosis to kill internalized cells. With the aid of a confocal laser-scanning microscope, images depicting interactive moments between cryptococcal cells and amoeba are captured. The resolution power of the electron microscope also helps to reveal the ultrastructural detail of cryptococcal cells when trapped inside the amoeba food vacuole. Since phagocytosis is a continuous process, quantitative data is then integrated in the analysis to explain what happens at the timepoint when an image is captured. To be specific, relative fluorescence units are read in order to quantify the efficiency of amoeba in internalizing cryptococcal cells. For this purpose, cryptococcal cells are stained with a dye that makes them fluoresce once trapped inside the acidic environment of the food vacuole. When used together, information gathered through such techniques can provide critical information to help draw conclusions on the behavior and fate of cells when internalized by amoeba and, possibly, by other phagocytic cells.

Complementary Use of Microscopic Techniques and Fluorescence Reading in Studying Cryptococcus-Amoeba Interactions

This paper details a protocol for preparing a co-culture of cryptococcal cells and amoebae that is studied using still, fluorescent images and high-resolution transmission electron microscope images. Illustrated here is how quantitative data can complement such qualitative information.

To simulate Cryptococcus infection, amoeba, which is the natural predator of cryptococcal cells in the environment, can be used as a model for ...

Immunology and Infection

Microfluidic Tools for Probing Fungal-Microbial Interactions at the Cellular Level

Filamentous fungi are successful inhabitants of soil and play a major role in soil ecosystems, such as in the decomposition of organic and inorganic matter, as well as regulation of nutrient levels. There they also find numerous opportunities to interact with a variety of other microbes such as bacteria or other fungi. Studying fungal interactions at the cellular level, however, can be challenging owing to the black box-like nature of soil. New microfluidic tools are being developed for the study of fungal interactions; two platforms designed to study bacterial-fungal and fungal-fungal interactions are highlighted. Within these microchannels, fungal-microbial interactions can be monitored in controlled physico-chemical environments at higher temporal and spatial resolution than previously possible. Application of these tools have yielded numerous novel biological insights, such as the observation of bacterial polar attachment to hyphae or revealing uncharacterised fungal-fungal antagonisms. A key feature of these methodologies regards the ease of use of this tool by non-experts, yielding highly translatable technologies for use in microbiology labs.

Owing to the opacity of soil, interactions between its constituent microbes cannot easily be visualised with cellular resolution. Here, two microfluidic tools are presented, which offer new opportunities for investigating fungal-microbial interactions. The devices are versatile and simple to use, enabling high spatiotemporal control and high-resolution imaging at the cellular level.

Filamentous fungi are successful inhabitants of soil and play a major role in soil ecosystems, such as in the decomposition of organic and inorganic ...

Bioengineering

Microbial Control and Monitoring Strategies for Cleanroom Environments

Microbial Control and Monitoring Strategies for Cleanroom Environments and Cellular Therapies

A well-validated and holistic program that incorporates robust gowning, cleaning, environmental monitoring, and personnel monitoring measures is critical for minimizing the microbial bioburden in cellular therapy manufacturing suites and the corresponding testing laboratories to ensure that the facilities are operating in a state of control. Ensuring product safety via quality control measures, such as sterility testing, is a regulatory requirement for both minimally manipulated (section 361) and more than minimally manipulated (section 351) human cells, tissues, and cellular and tissue-based products (HCT/Ps). In this video, we provide a stepwise guide for how to develop and incorporate the best aseptic practices for operating in a cleanroom environment, including gowning, cleaning, staging of materials, environmental monitoring, process monitoring, and product sterility testing using direct inoculation, provided by the United States Pharmacopeia (USP&lt;71&gt;) and the National Institutes of Health (NIH) Alternative Sterility Testing Method. This protocol is intended as a reference guide for establishments expected to meet current good tissue practices (cGTP) and current good manufacturing practices (cGMP).

Author Spotlight: Microbial Control and Monitoring Strategies for Cleanroom Environments and Cellular Therapies  

The protocol summarizes the best practices to minimize microbial bioburden in a cleanroom environment and includes strategies such as environmental monitoring, process monitoring, and product sterility testing. It is relevant for manufacturing and testing facilities that are required to meet current good tissue practice standards and current good manufacturing practice standards.

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The Cultivation, Growth, and Viability of Lactic Acid Bacteria: A Quality Control Perspective

Lactic acid bacteria (LAB) are essential dairy starter cultures that are significantly employed for the manufacture of fermented dairy products such as yogurt and cheese. LAB predominantly produce lactic acid as a major end product of fermentation, and they synthesize important metabolites that impart the organoleptic characteristics of fermented food products. LAB are fastidious bacteria that thrive in many environments when adequate nutritional requirements are fulfilled. The demand for superior LAB dairy starter cultures for fermentation applications in the food and dairy industry, has resulted in the need to provide viable and active cultures for all bioprocessing operations. The development of a standard protocol for ensuring the viability and enhanced functionality of LAB cultures in the laboratory as well as dairy processing environments is thus very critical. In addressing concerns linked to resuscitating weak, stressed, and injured LAB culture cells, a protocol that vividly outlines salient steps to recover, enhance cell regeneration, and improve metabolic functionality of LAB strains is of the utmost importance. The maintenance of culture purity, functionality, and viability for LAB starter cultures is likewise critical. Therefore, adherence to a unique protocol guideline will result in the promotion of fermentation performance for many LAB strains dedicated to fermentation and biotechnology processes. As a result, the Food Microbiology and Biotechnology Laboratory at North Carolina Agriculture and Technical State University has developed a standard protocol for the activation and quality control of selected LAB strains that has resulted in highly functional and viable LAB culture strains employed for fermentation research. The adaptation and recommendation of a protocol such as this for use in the dairy and food industry&#160;will help to ensure LAB viability and functionality for many applications.

The quality control assessment of lactic acid bacteria (LAB) cultures has been confirmed as an effective way to enhance the viability and functionality of LAB strains for fermentation procedures. To buttress this assertion, we developed a protocol that elucidates how LAB cultures are activated and cultivated for fermentation and bioprocessing procedures.

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Isolation of Culturable Yeasts and Molds from Soils to Investigate Fungal Population Structure

Soil is host to an incredible amount of microbial life, with each gram containing up to billions of bacterial, archaeal, and fungal cells. Multicellular fungi such as molds and unicellular fungi, broadly defined as yeasts, fulfill essential roles in soil ecosystems as decomposers of organic material and as food sources for other soil dwellers. Fungal species diversity in soil is dependent on a multitude of climatic factors such as rainfall and temperature, as well as soil properties including organic matter, pH, and moisture. Lack of adequate environmental sampling, especially in regions of Asia, Africa, South America, and Central America, hinders the characterization of soil fungal communities and the discovery of novel species. 
We characterized soil fungal communities in nine countries across six continents using ~4,000 soil samples and a protocol developed in the laboratory for the isolation of yeasts and molds. This protocol begins with separate selective enrichment for yeasts and the medically relevant mold Aspergillus fumigatus, in liquid media while inhibiting bacterial growth. Resulting colonies are then transferred to solid media and further processed to obtain pure cultures, followed by downstream genetic characterization. Yeast species identity is established via sequencing of their internal transcribed spacer (ITS) region of the nuclear ribosomal RNA gene cluster, while global population structure of A. fumigatus is explored via microsatellite marker analysis. 
The protocol was successfully applied to isolate and characterize soil yeast and A. fumigatus populations in Cameroon, Canada, China, Costa Rica, Iceland, Peru, New Zealand, and Saudi Arabia. These findings revealed much-needed insights on global patterns in soil yeast diversity, as well as global population structure and antifungal resistance profiles of A. fumigatus. This paper presents the method of isolating both yeasts and A. fumigatus from international soil samples.

This protocol is an effective, speedy method of culturing yeasts and the mold Aspergillus fumigatus from large sets of soil samples in as little as 7 days. The methods can be easily modified to accommodate a range of incubation media and temperatures as needed for experiments.

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A Flame-Free Method for Sterilizing C. elegans Picks, Spatulas, and Scalpels

This article describes a method for sterilizing worm picks, spatulas, and scalpels using a micro-incinerator in place of an open flame.

Caenorhabtidis elegans (C. elegans) is an optimal model organism for research and education at primarily undergraduate institutions. Undergraduates can ...

Aseptic Technique in Environmental Science

Source: Laboratories of Dr. Ian Pepper and Dr. Charles Gerba - The University of Arizona 
Demonstrating Author: Luisa Ikner
Aseptic technique is a fundamental skill widely practiced in the field of environmental microbiology that requires a balance of mindfulness and practice in the laboratory. Proper use of this technique reduces the likelihood of bacterial or fungal contamination of reagents, culture media, and environmental samples. Aseptic technique is also vital to ensure data integrity and maintain the purity of culture libraries that may be comprised of very rare and difficult to culture isolates. Sources of contamination in the laboratory environment include airborne microorganisms (including those adhering to dust and lint particles), microbes present on the laboratory bench workspace or on unsterilized glassware or equipment, and microbes transferred from the body and hair of the researcher. The use of aseptic technique is also a safety measure that lowers the potential for the transmission of microorganisms to researchers, which is particularly important when working with pathogens.

Cell Culture Techniques and Practices to Avoid Contamination by Fungi and Bacteria in the Research Cell Culture Laboratory

Summary

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Aseptic Laboratory Techniques: Volume Transfers with Serological Pipettes and Micropipettors

Aseptic Laboratory Techniques: Plating Methods

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Microfluidic Tools for Probing Fungal-Microbial Interactions at the Cellular Level

Microbial Control and Monitoring Strategies for Cleanroom Environments and Cellular Therapies

The Cultivation, Growth, and Viability of Lactic Acid Bacteria: A Quality Control Perspective

Isolation of Culturable Yeasts and Molds from Soils to Investigate Fungal Population Structure

A Flame-Free Method for Sterilizing C. elegans Picks, Spatulas, and Scalpels

Aseptic Technique in Environmental Science