2.1 DNA Restriction & Nucleic Acid Analysis Protocols
1. Click on the following links to view the sample protocol for conducting a Digesting DNA Reaction.
Digesting DNA Reaction (openwetware.org, HTML Document)
2. Click on the following links to see sample protocols for conducting a restriction enzyme digestion reaction and agarose gel electrophoresis.
Restriction Enzyme Digestion of DNA (userpages.umbc.edu, HTML Page)
Agarose Gel Electrophoresis (userpages.umbc.edu, HTML Page)
2.2 Nucleic Acid Amplification & Sequencing Protocols
1. OpenWetWare Protocol: Click on the following link to view the information for conducting PCR. When you have finished viewing the protocol, complete the theoretical lab assessment problem set.
Polymerase Chain Reaction (openwetware.org, HTML Document)
cDNA Synthesis & qPCR
cDNA Synthesis (openwetware.org, HTML Document)
Real-Time PCR (qPCR) Data Analysis (openwetware.org, HTML Document)
Sequencing DNA (openwetware.org, HTML Document)
2. UMBC PROTOCOL: Click on the following link to see sample protocol for PCR Amplification of DNA.
PCR Amplification of DNA (userpages.umbc.edu, HTML Page)
2.3 Nucleic Acid Hybridization & Expression Analysis Protocols
1. Click on the following links to view the introduction, on modern techniques used to detect the RNA and protein output of a cell. You will focus on the RNA information for Hybridization and Northern Blotting as it applies to this subset, however, you will revisit the text in the Proteins Techniques module to better understand how one can conduct enzymatic assays to analyze gene expression by measuring the protein product of a gene.
RNA Blot (openwetware.org, HTML Document)
2. Click on the following link to see a sample protocol for the Southern Blotting technique. The protocol also contains recipes for the solutions required to conduct the technique.
Southern Blotting (userpages.umbc.edu, HTML Page)
2.4 Molecular Cloning Protocols
1. Click on the following links to view the sample 1 protocol and reagents list respectively, for conducting a Ligation Reaction followed by a Bacterial Transformation. When you have finished viewing the protocol, complete the theoretical lab assessment problem set.
DNA Ligation (openwetware.org, HTML Document)
2. Click on the following links to see sample protocols for a restriction enzyme digestion reaction, a ligation reaction and bacterial transformation.
Protocol Samples 1-3:
(1) Restriction Enzyme Digestion: (userpages.umbc.edu, HTML Page)
(2) Ligation Reaction: (userpages.umbc.edu, HTML Page)
(3) Bacterial Transformation: (userpages.umbc.edu, HTML Page)
Recipes for Common Stock Solutions: (userpages.umbc.edu, HTML Page)
3. View the following document to see a visual sequence and protocol for plating out a bacterial transformation reaction.
Protocol Sample 5: “Plating Out Your Transformation” (PDF Document)
2.5 Preparation, Purification, and Quantitation of DNA & RNA Protocols
1. Click on the following links to view information for DNA purification and RNA extraction.
DNA Purification: (openwetware.org, HTML Document)
RNA Extraction: (openwetware.org, HTML Document)
2. Click on the following links to see sample protocols for the Preparation of genomic DNA from bacteria, Phenol/chloroform extraction of DNA, and Ethanol precipitation of DNA. The protocols also contain recipes for the solutions required to conduct the applications.
Preparation of genomic DNA from bacteria (userpages.umbc.edu, HTML Page)
Phenol/chloroform extraction of DNA (userpages.umbc.edu, HTML Page)
Ethanol precipitation of DNA (userpages.umbc.edu, HTML Page)
3.2 Protein Detection & Analysis Protocols
Western Blotting & SDS-PAGE
2. ELISA Protocol
ELISA Protocol (thermofisher.com, HTML Document)
3.3 Protein Purification Protocols
1. Protein Extraction ProtocolUNC Protein Extraction Protocol: (www.unc.edu, Multimedia Page)
Bradford Protein Assay (openwetware.org, HTML Document)
2. Protein Precipitation
California Institute of Technology: Protein Precipitation Protocol (caltech.edu, PDF Document)
3. Protein Assays
Protein assay techniques can be categorized into “Absorbance” assays and “Colorimetric” assays. The following links will direct you to a webpage outlining the protocol for each technique.
a. Absorbance Assays
• Absorbance Assay (280 nm): (www.ruf.rice.edu, Multimedia Page)
• Biuret Protein Assay: (www.ruf.rice.edu, Multimedia Page)
• Bradford Protein Assay: (www.ruf.rice.edu, Multimedia Page)
• Bicinchoninic Acid (BCA) Protein Assay: (www.ruf.rice.edu, Multimedia Page)
- Corning Guide for Identifying & Correcting Common Cell Growth Problems
Cell Growth Problems (catalog2.corning.com, PDF Document)
- Corning Subculturing Monolayer Cell Cultures
Monolayer Cell Cultures (catalog2.corning.com, PDF Document)
- Corning Cryogenic Preservation & Storage of Animal Cells Protocol:
Preservation & Storage Protocols (catalog2.corning.com, PDF Document)
MIT & UMBC Protocols
1. MIT Biological Engineering 2007 – Introduction to Cell Culture Click on the following link
Cell Culture (ocw.mit.edu, PDF Document)
2. MIT Biological Engineering 2006 – Tissue Culture Protocol Click on the following link
Tissue Culture Protocol (ocw.mit.edu, WORD Document)
3. UMBC – Applied Molecular Biology Viable Cell Counting Protocol (Using a Hemocytometer to Determine Total Cell Counts and Viable Cell Numbers, Reference: Sigma Catalogue). When accessing the webpage (referenced throughout section IVb. Content) scroll down to the end of the page and refer to step 7 for additional information on conducting Viable Cell Counts.
Viable Cell Counts (userpages.umbc.edu, HTML Page)
Additional Guidelines: Cell Culture Techniques
Media for Cell Culture and Other Important Things From the ATCC web FAQ pages (www.atcc.org, HTML Page)
How should I subculture a monolayer? Answer: Refer to the product sheet included with your ATCC cell line for the subculturing procedure recommended for that particular cell line. Anchorage-dependent cell lines are usually subcultured by disaggregation of the cell sheet with proteolytic enzymes such as trypsin. Ethylenediaminetetraacetic acid (EDTA), a chelating agent, may be added to the dissociation solution to enhance the activity of the trypsin by removing calcium and magnesium from the surfaces of the cells. An appropriate solution for general use is a solution of 0.25% (w/v) trypsin to 0.03% (w/v) EDTA prepared in saline without divalent cations (such as calcium- and magnesium-free phosphate buffered saline). Rinse the cell sheet with the trypsin solution (e.g., 5.0 to 10.0 ml/75 sq. cm flask) and remove. Add a small amount of trypsin solution to cell sheet (2.0 to 5.0 ml/75 sq. cm flask) and observe until the cell sheet is dispersed (usually within 5 to 15 min). To avoid clumping, do not agitate the cells by hitting or shaking the flask while waiting for the cells to detach. Monolayers that are particularly difficult to detach can be placed at 37°C to facilitate dispersal. Once the cells lift off the surface of the flask, add growth medium containing serum to the flask. Serum contains proteins that will suppress the activity of the trypsin. It is not usually necessary to centrifuge the cells at this step. However, if either serum-free or low serum medium is used, centrifuge the cells at 125 x g for 5 min to remove the residual dissociation solution. Aspirate the cells by pipetting gently, and then disperse the cells to new flasks. The subculture ratio will vary from 1:2 to 1:20 or greater depending on the cell line. Consult the product sheet for the recommended dilution ratio. Trypsin can be damaging to the membranes of some cell types. Gently scraping the cell sheet from the surface of the flask into a small amount of medium and then aspirating the mixture to obtain a cell suspension may disperse these cells. Individually wrapped sterile cell scrapers, which are available from a number of commercial vendors, may be used for this purpose. For further detail on enzymatic solutions and the maintenance of cell lines, consult Chapter 10 in Culture of Animal Cells: A Manual of Basic Technique, 3rd edition, (1994) by R. Ian Freshney (Wiley-Liss, Inc., New York).
How often should I subculture a cell line? Answer: Subculture refers to the transplantation of cells from one vessel to another. The term subculture is synonymous with the terms passage and split. The subculture interval is the time between subsequent subcultures. The subculture number is simply the number of times a culture has been transferred from one vessel to another. Adherence-dependent cell lines are generally subcultured either at or near confluency. However, some cell lines never reach 100% confluency but grow in colonies or domes and should be subcultured when the cells appear to have reached a maximum density for that particular cell line and vessel. Contact-inhibited cell lines such as ATCC CCL-92 (3T3-Swiss albino) should be subcultured before they reach confluency to avoid gradual selection of contact-insensitive variants. Suspension cell lines should be subcultured before they reach their maximum saturation density (the number of cells per unit volume of medium). This usually varies between 3 x 105 and 2 x 106. The cells must be diluted to a lower cell concentration to allow sufficient nutrients for resumption of logarithmic growth. If the medium is simply replaced and the cell density is not decreased, cells will rapidly deplete the medium and die. If the cells are diluted below their minimum density, they will either enter into a lag phase and grow very slowly, or they will die. The saturation densities and hence subculture intervals for each suspension line will vary. Therefore, daily cell counts are the best way to monitor suspension lines.
What does plating efficiency mean? Your catalog description states that the plating efficiency is 5% for my cell line. Does this mean I can expect to lose 95% of my cells every time I subculture? Answer: For cell lines consisting of adherent cells, plating efficiency, or more precisely, colony forming efficiency, refers to the percentage of cells planted which give rise to colonies. When a cell line can be grown from single cells, the plating efficiency is determined at ATCC directly from a thawed ampule of cells by a simple dilution of the cell suspension and subsequent inoculation of dishes or flasks with a suitable number of cells (100 to 1,000 cells/75 sq. cm flask). The plating efficiency is determined by counting the number of colonies that form: (number of colonies formed/ number of cells inoculated) x 100. Some researchers use the term plating efficiency to refer to the percentage of cells plated that attach to the surface of the culture vessel after a short period of time. Plating efficiencies are usually quantitated at low or clonal cell densities. It can be as low as 1% or, in the case of some tumor lines, even zero. Some cell lines must be seeded at a high density to survive. All cell cultures received from ATCC will survive if seeded according to the instructions on the product sheet and if subcultured at the recommended split ratios.
How do I change the medium in (feed) a suspension culture? Answer: Suspension cell lines can be fed by the simple addition of fresh medium to the culture (if room is available) or by separating the cells from the old medium by centrifugation (125 x g for 5 min) with subsequent resuspension of the cell pellet in fresh medium. However, with most suspension cell lines, a simple addition of medium is the preferred method. In either case it is absolutely necessary to feed the cultures before the cells reach their maximum saturation density. This can range between 3 x 105 to 2 x 106 depending on the cell line and the culture conditions (stationary versus stirred, oxygenation levels, etc.). The cells must be diluted to a lower cell concentration to allow sufficient nutrients for resumption of logarithmic growth. If the medium is simply replaced and the cell density is not decreased cells will rapidly deplete the medium and die. If the cells are diluted below their minimum density they will either enter into a lag phase and grow very slowly, or they will die. The saturation densities and hence subculture intervals for each suspension line will vary. Therefore, daily cell counts are the best way to monitor suspension cell lines.
How do I grow hybridoma cell lines as ascites, especially rat-mouse hybridomas? Answer: ATCC propagates most hybridomas as cell cultures in vitro, but many can be grown efficiently as ascites by inoculation into the peritoneal cavities of mice. Some rat-mouse hybridomas are more problematical as the host mouse strain may develop anti-rat antibodies and eventually reject the hybridoma inoculum. To minimize this difficulty one can use athymic nude mice as hosts or immunosuppress host animals prior to inoculation. For a protocol see p.403 in Hybridomas: A New Dimension in Biological Analyses (1980), Kennett, R.J., McKearn, T.J., and Bechtol, K.B., eds. (Plenum Press, New York).
How can I adapt a monolayer cell line to grow in suspension culture? Answer: Not all cell lines can be adapted to suspension growth. In general, normal diploid anchorage-dependent (must be attached to a substrate to grow) cells cannot be adapted without the use of microcarrier beads to which they can attach. Lines such as L-929 (ATCC CCL-1) and HeLa (ATCC CCL-2) which are not anchorage-dependent can be adapted and variants that grow in suspension already exist. Patience, and the use of specially modified suspension culture medium, are the keys to adapting cells to suspension growth. Cultures are usually grown in a small 250 ml to 1,000 ml spinner cultures (with half that volume of actual medium), in glass vessels with a stirring paddle suspended inside that is driven at approximately 50 to 100 rpm by a magnetic stirrer. The spinner/suspension culture medium, such as Joklik's modified MEM medium (GIBCO BRL Cat. No. 22300; BioWhittaker Cat. No.12-126Q; Sigma Cat. No. M4767; Irvine Scientific Cat. No. 9489), usually omits calcium and magnesium ions to help prevent clumping and may require an antifoaming agent if serum is used to prevent foaming from the serum. The glass culture vessel is usually coated with a siliconizing compound to prevent the cells from sticking to the glass. Initially, most of the cells will plate out on the glass surface of the vessel or form large clumps with each other. At each passage, those cells that are in suspension are used to inoculate the next vessel. With time, a population of cells may be selected that does not self-aggregate or adhere to glass as readily as the parental line. Bear in mind that the newly selected line may have lost or acquired characteristics independent that are different from the original cell population. For further detail on suspension cell culture techniques, consult Chapter 12 in Animal Cell Culture Methods (1998), J. Mather and D. Barnes, eds. (Academic Press, New York); Chapter 10 in Culture of Animal Cells: A Manual of Basic Technique, 3rd ed., (1994) by R. Ian Freshney (Wiley- Liss, Inc., New York); Chapters 15 and 17 in Methods in Enzymology: Cell Culture, Vol. 58 (1979), W. B. Jacoby and I. H. Pasten, eds. (Academic Press, New York).
What are the recommended carbon dioxide (CO2) levels needed to grow a cell culture? Answer: While the levels of carbon dioxide in cell culture systems vary from that in ambient air (about 0.03%) up to 40% in air, generally either no added CO2 or 5% to 10% CO2 in air are the most frequently used. It is very important to adjust the concentration of sodium bicarbonate used in a medium to that required for equilibration with the level of CO2 used in the gas phase. Cells in culture produce CO2 and require small amounts of the compound for growth and survival. If no CO2 is added and mass cell cultures are being propagated, anhydrous sodium bicarbonate at 4 mM (0.34 g/L) can be used. However, the culture vessel should be kept sealed (caps on flasks should be tight). If 5% or 10% CO2 is desired, use 23.5 mM (1.97 g/L) or 47 mM (3.95 g/L) sodium bicarbonate, respectively, for an initial pH of about 7.6 at 37°C. Under these conditions unsealed (loose caps) culture flasks or dishes must be used to allow the gases to equilibrate. For more detail on the gaseous environment for the mammalian cell in culture, consult Chapter 5 by W.F. McLimans in Growth, Nutrition and Metabolism of Cells in Culture, Vol.1 (1972), G.H. Rothblat and V.J. Cristofalo, eds. (Academic Press, New York).
Can human, pig, or mouse interleukin-2 (IL-2) be substituted for rat spleen cell-conditioned medium (rat factor) in the media for IL-2-dependent cell lines? Answer: Generally, IL-2 is not species specific, and rat IL-2 is active in mouse IL-2 dependent cell lines. Other IL-2's (e.g., human or guinea pig) can be used with the CTLL-2 (ATCC Cat. No. TIB-214) cell line, but may not give the same degree of growth stimulation. We recommend using medium conditioned by concanavalin A-stimulated rat spleen cells for the growth of our mouse IL-2 dependent lines. Alternatively, purified rat or mouse IL-2 can be purchased from commercial suppliers. In our experience, the conditioned medium gives a better response than the purified IL-2 preparations.
Can I use a different medium from the one recommended in the ATCC catalog or listed on the ATCC product sheets? Answer: ATCC generally lists in the catalog and product sheet either the medium recommended by the originator of the cell line or a standard medium formulation that has been found to be effective otherwise. However, other unspecified media may also be entirely satisfactory. It may be prudent to prepare a seed stock using the originally prescribed medium and other conditions before attempting a substitution. Two approaches can be used. The simplest method is to simply change the medium to the desired alternative formula and then pass the cell line 3 to 5 times to force it to adapt to the new conditions. A more gentle approach is to subculture the cells at a 1:2 split ratio into two identical culture vessels with one vessel containing the original medium and the other containing a mixture of 50% original medium and 50% new medium. The effects of the new medium formulation on the cells can then be easily observed by comparing the growth in the cultures with the medium mixture with the culture containing only the original medium formulation. Once the cells in the mixed culture have become confluent, the process can be repeated by again splitting the mixed culture at a 1:2 ratio into two identical vessels. This time one vessel should contain 50% original medium and 50% new medium while the second culture has 25% original medium and 75% new medium. Once this second culture has become confluent it can be subcultured into 100% new medium.
How much Earle's salts is required with Eagle’s Minimum Essential Medium? How much nonessential amino acids? Do I need glutamine? Sodium bicarbonate? Answer: Minimum essential medium (MEM) was developed by Eagle as a “bare bones” or very simple medium and is commercially available in many different formulations. IT IS VERY IMPORTANT TO READ CATALOG DESCRIPTIONS AND MEDIA BOTTLE LABELS CAREFULLY SINCE MEDIA MIXUPS ARE A LEADING CAUSE OF CELL CULTURE PROBLEMS. These formulations usually incorporate either Earle’s or Hanks’ balanced salts to provide appropriate buffering of the medium. Most commercially available liquid media usually have the appropriate bicarbonate levels (2.2 g/L for Earle’s formulations and 0.35 g/L for Hanks’ formulations) added during manufacturing and do not require additional bicarbonate. Powdered media require sodium bicarbonate when the medium is reconstituted. Follow the manufacturer's directions for the recommended amounts. Earle’s salts are recommended when carbon dioxide (approximately 5%) is added to the gas phase of the incubator. This medium should either be used with unsealed culture vessels in a CO2 incubator, or in a sealed vessel that has been gassed with 5% CO2 after filling. Hanks' salts have a lower bicarbonate level and less buffering capacity than Earle’s salts and are designed to be used without addition of carbon dioxide to the gas phase. Thus, a sealed culture vessel should be used with this buffering solution to maintain pH. Because MEM is a simple medium, it is often recommended that additional supplements or higher levels of serum be used to improve cell growth. One such supplement is a solution of nonessential amino acids, which can be prepared or purchased separately as a sterile stock (often 10 mM; 100X), that is aseptically added to the medium for a final concentration of 0.1 mM each. MEM can also be purchased with nonessential amino acids already added. Sodium pyruvate, another frequently recommended supplement, can be purchased as a sterile stock and added to a final concentration of 1 mM. L-glutamine (2 mM) is always required and must be added if it is not already included in the formulation purchased. Sodium bicarbonate (2.2 g/L for Earle's formulations and 0.35 g/L for Hanks’ formulations) is also required and must be added if not already included. In addition, there are formulations that eliminate calcium and magnesium to facilitate the growth of cells in suspension. Check ATCC’s Web site to search our media formulations or contact media manufacturers for complete formulations of the most common cell culture media.
Does my ATCC cell culture require L-glutamine in the medium? Why isn't L-glutamine listed on the information sheet? How much L-glutamine should I add? Answer: L-glutamine is an essential amino acid required by virtually all mammalian and invertebrate cell lines. It is an ingredient of all media formulations currently used for cell culture development, growth and maintenance. Because L-glutamine is somewhat unstable in liquid media, it is often omitted from commercial liquid media preparations but included in powdered formulations. If it is not included in the original preparation, it must be aseptically added to the medium prior to use. Additional L-glutamine can also be added to media to extend its shelf life. Liquid L-glutamine can be purchased from most commercial vendors of cell culture reagents. L-glutamine concentrations for mammalian cells can vary from 0.68 mM for Medium 199 to 4 mM for Dulbecco's modified Eagle's medium. Invertebrate cell culture media such as Schneider's Drosophila medium may contain as much as 12.3 mM L-glutamine. For more detail on the utilization of amino acids by cell cultures, consult Chapter 6 by M.K. Patterson, Jr. in volume 1 of Growth, Nutrition and Metabolism of Cells in Culture, 1972, edited by G.H. Rothblat and V.J. Cristofalo (Academic Press, New York).
Why is supplemental insulin required for ATCC CRL-1794 (13C4)? Answer: Rarely do donors of cell lines determine if there is an absolute need for each and every component of the medium they use. Often, a medium or additives are chosen simply because they have proven in the past to be successful with similar cell lines. Insulin is a hormone that has multiple functions including the stimulation of glucose transport and utilization, the uptake of amino acids, and the maintenance of differentiation. It has often been found to increase the growth of cultured cells, especially hybridomas. ATCC's Hybri-Care Medium (ATCC Cat. No. 46-X, 47-X) contains insulin and oxalacetic acid and will support the growth of 80% to 90% of the hybridoma cell lines distributed.
Why is oxalacetate used in cell culture? Can oxalacetic acid be used as a substitute for oxalacetate? Answer: Oxalacetic acid is an essential component of the Krebs cycle where it combines with acetyl-coenzyme A to yield citric acid. Thus, it is a necessary intermediate in the intercyclic pathway between the Embden-Meyerhof cycle and the Krebs cycle. The decarboxylation of oxalacetic acid gives pyruvic acid and CO2. The prime function of these cycles is the production of energy and the shunting out of these acids effectively slows the cycle down. Therefore, keto acids, such as pyruvate and oxalacetic, are frequently added to tissue culture media formulations to maintain maximum cell metabolism. Oxalacetic acid also provides a carbon skeleton for transamination reactions. As components of tissue culture media, the terms oxalacetate and oxalacetic acid are used interchangeably
What is the purpose of heat-inactivating serum, why is it recommended for some cell lines, and how do I do it? Answer: Heat-inactivation (heating to 56°C for 30 min) is done to inactivate complement, a group of proteins present in sera that are part of the immune response. This is sometimes important for cells that will be used to prepare or assay viruses, used in cytotoxicity assays or other systems where complement may have an unwanted influence. Heat has also been used to destroy mycoplasma in serum. Because most serum suppliers filter through 0.1 µm filters to remove mycoplasma before distribution, this is not usually necessary. Heat-inactivation is also recommended for growing embryonic stem cells [p. 75 in Rudnicki, M.A., and McBurney, M.W. (1987) Teratocarcinomas and Embryonic Stem Cells - A Practical Approach (IRL Press Ltd., Oxford)] as well as for many insect cell lines (Weiss, S.A., et al. (1995). Meth. Mol. Biol. 39:65). Heat inactivation will reduce or destroy serum growth factors, and should only be done when there is a compelling reason. The following procedure can be used to heat-inactivate serum:
- Thaw serum following directions in the next question.
- Preheat water bath to 56°C. There must be sufficient water to immerse the bottle above the level of serum.
- Mix thawed serum by gentle inversion and place serum bottle in the 56°C water bath. (The temperature of the water bath will drop.)
- When the temperature of the water bath reaches 56°C again, continue to heat for an additional 30 min. Mix gently every 5 min to insure uniform heating.
- Remove serum from water bath and cool. Place at recommended storage conditions. ATCC stores the serum at -70°C. We recommend that you store at -70°C if possible, otherwise at -20°C.
What is the best way to thaw fetal bovine serum? Answer: Remove the serum from frozen storage and place it overnight in a refrigerator at 2°C to 6°C. Transfer the bottles to a 37°C water bath. Agitate the bottles from time to time in order to mix the solutes that tend to concentrate at the bottom of the bottle. Do not keep the serum at 37°C any longer than necessary to completely thaw it. Thawing serum in a bath above 40°C without mixing may lead to the formation of a precipitate inside the bottle. We don't recommend thawing the serum at high temperature.
Why is my serum cloudy after thawing? Answer: The procedures used to prepare both ATCC and other brands of serum may retain some fibrinogen. Since external factors may initiate the conversion of fibrinogen to fibrin, flocculent material or turbidity may be observed after thawing. Testing of serum after this has happened indicates that it does not alter its ability to function as a supplement for cell culture media. If the presence of flocculent material or turbidity is a concern, it can be removed by filtration through a 0.45 µm filter. A precipitate can form in serum that is incubated at 37°C for prolonged periods of time. Electron microscopy and X-ray microanalysis indicate that the precipitate may include crystals of calcium phosphate. The formation of a calcium phosphate precipitate does not alter the performance of the serum as a supplement for cell culture
Are there any alternatives to using fetal bovine serum? Answer: Due to the fluctuating and increasing cost of fetal bovine serum many cell culture scientists are trying to find serum substitutes. In addition, many specialized cell types do not grow or function well in serum-containing medium. As a result a substantial amount of research as been done to develop serum-free media and serum substitutes. In some cases it is possible simply to reduce the concentration of fetal bovine serum without altering cell line growth or other properties. For example, the human colon cancer line ATCC CCL-227 (SW620) proliferates equally well in DMEM supplemented with 1%, 5%, or 10% fetal bovine serum. Alternatively, it may be possible to propagate many lines in bovine calf serum or iron-supplemented bovine calf serum instead of fetal bovine serum. If successful, either method will reduce serum use and costs significantly. The addition of insulin, selenium, and especially transferrin to culture media may substantially reduce the amount of serum required for optimal proliferation (Barnes, D., and Sato, G. (1980) Anal. Biochem. 102:255.). Many preparations are now commercially available with these and other ingredients (e.g., linoleic acid, serum albumin) to permit cell growth at low serum concentrations. For further detail on growing cells in serum-free media, consult Chapter 2 in Animal Cell Culture Methods (1998), J. Mather and D. Barnes, eds. (Academic Press, New York); Chapter 7 in Culture of Animal Cells: A Manual of Basic Technique, 3rd ed., 1994, by R. Ian Freshney, (Wiley-Liss, Inc., New York); Chapter 6 in Methods in Enzymology: Cell Culture, Vol. 58, (1979) W. B. Jacoby and I. H. Pasten, eds. (Academic Press, New York).
Can I use HEPES buffer in my cell culture medium? Answer: HEPES and other organic buffers can be used effectively with many cell lines (see Shipman, C. (1969) Proc. Soc. Exp. Biol. Med. 130: 305). However, be aware that the compound can be toxic, especially for some differentiated cell types, so its effects should be evaluated before routine use (People, C.A., et al., (1982) In Vitro 18: 755). HEPES has also been shown to greatly increase the sensitivity of media to the phototoxic effects induced by exposure to fluorescent light. [Zigler, J.S., et al. (1985) Analysis of the cytotoxic effects of light-exposed HEPES-containing culture medium. In Vitro 21: 282. Spierenberg, G.T., et al. (1984) Phototoxicity of N-2-hydroxyethylpiperazine-N-ethanesulfonic acid-buffered culture media for human leukemic cell lines. Cancer Research 44:2253.
Why is it important to limit exposure of cell culture media to fluorescent lights? Answer: An important but often overlooked source of chemical contamination results from the exposure of media containing riboflavin or tryptophan to normal fluorescent lighting (see references below). These media components are photoactivated by UV radiation emitted from most fluorescent lights and give rise to hydrogen peroxide. This generates free radicals that are toxic to cells; the longer the exposure the greater the toxicity. HEPES (an organic buffer commonly used to supplement bicarbonate based buffers) appears to increase the phototoxic exposure effects (2, 3). Sodium pyruvate has been shown to reduce or eliminate this phototoxicity (3). Short-term exposure of media to room or hood lighting when feeding cultures is usually not a significant problem. However, leaving media on lab benches for extended periods, storing media in walk-in cold rooms with the lights on, or using refrigerators with glass doors where fluorescent light exposure is more extensive will lead to a gradual deterioration in the quality of the media.
- In Vitro 12:19.
- Zigler, J.S., et al. (1985) Analysis of the cytotoxic effects of light-exposed HEPES-containing culture medium. In Vitro 21: 282.
- Spierenberg, G.T., et al. (1984) Phototoxicity of N-2-hydroxyethylpiperazine-N-ethanesulfonic acid-buffered culture media for human leukemic cell lines. Cancer Research 44: 2253.
Why do some cell lines require sodium pyruvate? How much should I add to the medium? Answer: Pyruvate is an intermediary organic acid metabolite in glycolysis and the first of the Embden Myerhoff pathway that can pass readily into or out of the cell. Thus, its addition to tissue culture medium provides both an energy source and a carbon skeleton for anabolic processes. Its addition may help in maintaining certain specialized cells, may help when cloning and may be necessary when the serum concentration is reduced in the medium (Culture of Animal Cells: A Manual of Basic Technique, 3rd edition, (1994) by R. Ian Freshney (Wiley- Liss, Inc., New York). Sodium pyruvate may also help reduced fluorescent light-induced phototoxicity (Spierenberg, G.T., et al. (1984) Phototoxicity of N-2-hydroxyethylpiperazine-N-ethanesulfonic acid-buffered culture media for human leukemic cell lines. Cancer Research 44:2253). Usually sodium pyruvate is added to give a final concentration of 0.1 mM. Sodium pyruvate is commercially available as a 10 mM (100X) stock solution.
Are there tissue culture media that do not require using a CO2 incubator? Answer: Some cell lines may be maintained satisfactorily on an alternative medium such as CRCM-30 (Macy, M.L., and Shannon, J.E. (1977) TCA Manual 1: 3), L-15 medium, or CO2-independent medium (GIBCO BRL Cat. No. 18045) which do not require CO2 in the gas phase. You can usually determine if a medium is satisfactory by using it with the cell line in question for 3 to 5 passages. However, cultures established at very low concentrations (e.g., cloning) usually require CO2 in the gas phase. An alternative to using a CO2 incubator is to have a 5% CO2 gas tank at your work site and stream filtered CO2 into the gas phase above the medium prior to sealing them (p. 80 in Culture of Animal Cells: A Manual of Basic Technique, 3rd edition, (1994) by R. Ian Freshney (Wiley-Liss, Inc., New York).
Can antibiotics and/or antimycotic agents be added to the cell culture medium? Answer: Most cell culture technologists avoid using antibiotics for routine culture work. Antibiotics may mask contamination by susceptible bacteria and fungi while permitting mycoplasma to flourish unnoticed. Antibiotics may interfere with the metabolism of sensitive cells in culture. However, one may elect to introduce antibiotics for short periods to primary cultures or as a safeguard while propagating specific valuable stocks (e.g., cells obtained directly from ATCC) to produce working stocks. Typical concentrations are 50 to 100 units penicillin G, 50 to 100 µg gentamicin sulfate or 2.5 µg amphotericin B/ml of culture medium. See Chapter 7 in Methods in Enzymology: Cell Culture, (1979) Vol. 58, W. B. Jacoby and I. H. Pasten, eds. (Academic Press, New York).
When should I use a feeder layer with my cultures and do you offer feeder layer cultures? Answer: Cultures of irradiated human or mouse cells (feeder cells) have been used for years to promote proliferation, particularly with low-density inocula (7). High-energy irradiation can completely suppress cell division long before general metabolism is appreciably affected. Since such an irradiated "feeder" cell population continues to metabolize actively, the non-multiplying cells provide diffusible and short-lived growth plus conditioning factors to the medium. These in turn stimulate non-irradiated, fastidious, proliferative cells added to the co-culture system. Numerous studies have utilized feeder cells effectively with normal (1,5,8) and tumorigenic epithelia (9), glioma (3), teratocarcinoma (6), B lymphoblast (2), hybridoma (4) and other populations. ATCC offers three cryopreserved feeder populations, MRC-5 (human diploid lung, ATCC Cat. No. 55-X); 3T3 (mouse, contact sensitive, ATCC Cat. No. 48-X) and STO (mouse embryonic fibroblast, ATCC Cat. No. 56-X). These feeder layer cells are available as frozen suspensions in 1 ml portions, each containing a sufficient cell population for seeding up to 225 cm2 of culture vessel surface. They have been characterized and shown to be mitotically arrested. They are treated by gamma irradiation, which suppresses cell division yet allows active metabolism. These feeder layer cells are available as frozen suspensions in 1 ml portions, each containing a sufficient cell population for seeding a culture vessel with up to 225 cm2 of surface area. Irradiated populations of other designations may be provided on a custom-quote basis. For additional information on feeder layers refer to the references below:
What methods does the ATCC use to detect bacterial contamination in cell cultures? Answer: ATCC tests all sera and media used plus all cell culture freezes prepared for bacteria, fungi, and mycoplasma. Blood agar, thioglycollate broth, trypticase soy broth, BHI broth, Sabouraud broth, YM broth, and nutrient broth with 2% yeast extract are used to test for bacteria and fungi. The direct culture method and Hoechst staining are used for mycoplasma. Detailed protocols are given in the manual ATCC Quality Control Methods for Cell Lines, available by order (ATCC Cat. No. 82-X). See also Chapter 2 in Methods in Enzymology: Cell Culture Vol. 58 (1979), W. B. Jacoby and I. H. Pasten, eds. (Academic Press, New York).
How can I tell if my culture is contaminated with another cell line? How can contamination be avoided? Answer: Morphological examination will occasionally suffice but this is a notoriously poor method for identification. Fluorescent antibody staining, isoenzyme, cytogenetic and/or DNA analyses can be used to detect interspecies and intraspecies cross contamination. The best means to guard against cross contamination include some common sense procedures: working with only one cell line at a time, using separate aliquots of media for each, and taking great care in labeling. ATCC publishes a Quality Control Manual for Cell Lines, available by order (ATCC Cat. No. 82-X). Also consult Chapters 3, 4 and 11 in Animal Cell Culture Methods (1998) J. Mather and D. Barnes, eds. (Academic Press, New York); Chapter 16 in Culture of Animal Cells: A Manual of Basic Technique, 3rd edition, (1994) by R. Ian Freshney (Wiley-Liss, Inc., New York); or Chapter 2 in Methods in Enzymology: Cell Culture (1979) Vol. 58, W. B. Jacoby and I. H. Pasten, eds., (Academic Press, New York).
How are cells screened for mycoplasma contamination? Answer: Cell lines are screened for mycoplasma contamination by direct cultivation and by indirect methods. For example, the fluorochrome Hoechst DNA stain will bind to the DNA of mycoplasma and the organisms can be detected easily when examined using a microscope equipped with appropriate fluorescence optics. The ATCC Mycoplasma Detection Kit (ATCC Cat. No. 90-1001K) provides a very sensitive PCR-based mycoplasma detection method. For additional information refer to the following references: Cell culture contamination: sources, consequences, prevention and elimination, by C.K. Lincoln, and M.G. Gabridge, in Animal Cell Culture Methods, (1998), J. P. Mather and D. Barnes, eds., p. 49 (Academic Press, San Diego); Chapter 16 in Culture of Animal Cells: A Manual of Basic Technique, 3rd edition, (1994) by R. Ian Freshney (Wiley-Liss, Inc., New York). The direct culture method requiring both broth and agar will permit isolation of cultivable strains as apparent by appearance of characteristic mycoplasma colonies on the agar medium. Both of these techniques are used several times while a cell line is expanded for distribution. The methods employed are described in detail in the publication ATCC Quality Control Methods for Cell Lines, available by order (ATCC Cat. No. 82-X).
Can a cell line be cured of mycoplasma contamination? Answer: Yes, several lines in our collection have been cured of mycoplasma (e.g., ATCC CCL-229, ATCC HB-175). However, this process is time consuming and does not always work; discarding the culture and starting over is always the preferred method. As with other microbial infections, one should first identify the contaminant and select a suitable antibiotic, preferably by testing the contaminating mycoplasma for its antibiotic sensitivity. The cells are cultured for 1 to 2 weeks in the presence of the antibiotic, and then cultured without antibiotic for 1 to 2 months. At this point, the line is retested to make sure that the culture is clean. A very sensitive testing method should be used . The ATCC Mycoplasma Detection Kit (ATCC Cat. No. 90-1001K) is highly recommended for this purpose. Periodic retesting is necessary to make sure that the contaminant does not reappear. Since many antibiotics are more or less toxic to cells, a selected population that no longer exhibits qualities of the parental line may result. It may be necessary to examine the cured culture to determine if it is sufficiently similar to the original line. For more detailed information see: 1. Culture of Animal Cells: A Manual of Basic Technique, 3rd ed., (1994) by R. Ian Freshney (Wiley-Liss, Inc., New York). 2. Cell culture contamination: sources, consequences, prevention and elimination, by C.K. Lincoln, and M.G. Gabridge. In Animal Cell Culture Methods (1998), J. P. Mather and D. Barnes, eds., pp. 49-65 (Academic Press, San Diego). 3. Antibiotic treatment of mycoplasma-infected cultures. In Molecular and Diagnostic Procedures in Mycoplasmology Vol. II (1996), S. Razin and J.G. Tully, eds., p. 439 (Academic Press, San Diego).
Can a frozen ampule of cells be put back into liquid nitrogen and stored for any length of time? Answer: In many cases, an ampule shipped in dry ice (–70°C) can be placed back into liquid nitrogen and the population recovered by rapid thawing at a later date. However, the viability may be reduced by such treatment, and for some sensitive cell lines, this may make recovery more difficult. The phenomenon is thought to be due to a change in the ice crystal structure within cells that occurs during the temperature shift. For this reason, we recommends that cells be thawed and placed into culture as soon after receipt as possible. It is best to minimize storage time at –70°C; that is, to use this temperature only for shipping. Recovery of frozen ATCC cultures is guaranteed, and replacements will be provided without charge within 30 days of their receipt.
What safety precautions are necessary for thawing ampules that have been stored in liquid nitrogen? Answer: A glass ampule or plastic vial that has been submerged in liquid nitrogen can explode upon removal if it has not been properly sealed. Resulting glass or plastic fragments fly at high force in all directions creating a hazard. Thus a face guard and protective gloves and clothing must be worn whenever an ampule is removed from liquid nitrogen. ATCC does not routinely store cells in the liquid phase. For reconstitution, the ampule should be agitated in a covered water bath at 37°C until its contents have thawed completely. To recover the cell suspension from a glass ampule, the neck is nicked with a small file, the ampule is washed with 70% ethanol, wrapped between several folds of a sterile towel or gauze and snapped open. The ampule contents can then be removed with a sterile 1 ml pipette (needles and syringes should be avoided whenever possible).
I have just received a human tumor cell line. What are the biohazards associated with this line? How should I work with it? Answer: It is not feasible to test every cell line for the presence of every possible adventitious agent. It is strongly recommended that all human and other primate cell lines be handled at the same biosafety level as a cell line known to carry HIV or hepatitis virus. At the very minimum all cell manipulations should be carried out using mechanical pipetting devices in a vertical laminar flow biosafety cabinet and all contaminated material should be decontaminated before washing or disposal. Detailed discussions of laboratory safety procedures are provided in the references below: 1. Caputo, J.L. Biosafety procedures in cell culture. (1988) J. Tissue Culture Methods 11:223. 2. U.S. Government Publication, Biosafety in Microbiological and Biomedical Laboratories Human Health Service Publication No. (CDC) 93-8395. U.S. Dept. of Health and Human Services; 3rd Edition, U.S. Government Printing Office Washington D.C.). This publication is available in its entirety in the Center for Disease Control Office of Health and Safety's Web site at www.cdc.gov/od/ohs. 3. Fleming, D.O., et al. (1995) Laboratory Safety: Principles and Practice, 2nd ed. (ASM Press, Washington, DC).
What is the difference between the passage number of a cell line and its population doubling level? Answer: The passage number simply refers to the number of times the cells in the culture have been subcultured, often without consideration of the inoculation densities or recoveries involved. The population doubling level refers to the total number of times the cells in the population have doubled since their primary isolation in vitro. This is usually a very crude estimate rounded off to the nearest whole number. A formula to use for the calculation of population doublings is as follows: n = 3.32 (log UCY - log l) + X, where n = the final PDL number at end of a given subculture, UCY = the cell yield at that point, l = the cell number used as inoculum to begin that subculture, and X = the doubling level of the inoculum used to initiate the subculture being quantitated. A more complete discussion of this subject was provided by L. Hayflick (1973) Tissue Culture Methods and Applications, P.F. Kruse, Jr. and M.K. Patterson, Jr. eds., p. 220 (Academic Press, New York).
Since we do not have a programmable freezer, how can we achieve the optimum –1°C/min freezing rate when preserving cells? Answer: One can avoid the expense of a programmable freezer by purchasing devices that can be used for cryopreservation in conjunction with mechanical or liquid nitrogen freezers. Alternatively, ampules and vials can be placed in a small Styrofoam™ box having wall and cover thickness of about 15 to 20 mm. When placed in a mechanical freezer at –70° to –90°C the internal cooling rate will approximate that required. Thermocouples inserted into dummy ampules can be used to check cooling rates if desired. See the ATCC Quality Control Manual for Cell Lines for more information.
Can mechanical freezers be used for cryogenic storage of cultured cells? Answer: Mechanical freezers can be used for storage if temperatures of –135°C or lower can be maintained. Viability will decline at higher temperatures. Backup freezers are needed in case of mechanical failure.
How long can frozen cells be kept in liquid nitrogen or dry ice without affecting recovery? Answer: Cells that are properly frozen using an effective cryoprotective agent can be stored in liquid nitrogen indefinitely without affecting recovery. Cells of CCL-1 were cryopreserved in February 1962. After decades in liquid nitrogen or its vapor, their viability has not significantly declined. Cell lines maintained on dry ice or in –70°C mechanical freezers, however, often lose their viability very quickly. The loss of viability will, of course, vary from cell line to cell line. Studies done at ATCC show some mouse lines dropping to 0% viability within six months on dry ice and some human lines were non-viable in only four months. Thus, it is recommended that ampules and vials be thawed as soon as possible upon receipt. If the cells cannot be thawed and cultured immediately, the ampules and vials should be stored at temperatures below –135°C, preferably in liquid nitrogen vapor. From the ATCC web FAQ pages
Reference (www.atcc.org, HTML Page)
1. UMBC – Applied Molecular Biology Procedure for Transfection of Mammalian Cells (Lipofection): Procedure Link (userpages.umbc.edu, HTML Page) In addition, refer to the details provided in Promega Transfection Guide. An additional protocol for Lipofection will be provided in the next subset IV-d. Expression Analysis, in one of the MIT protocol links that will be provided. The MIT protocol combines both transfection and expression analysis so it will appear in subset IV-d.
Overview of Transfection Methods: The inside scoop – evaluating gene delivery methods, Laura Bonetta (Nature Methods 2, 875-883 (2005)).
Journal Article (www.nature.com, HTML Page)
Bioluminescence Protocol: The Luciferase Assay – Promega (www.promega.com) The following link will take you to Promega’s webpage providing information about the Luciferase Assay System. The link below will take you to a page displaying an abstract for the Luciferase Assay System and both a detailed Complete Protocol (pdf file) and a Quick Protocol (pdf file).
The Luciferase Assay (www.promega.com, Multimedia Page)
1. Ultrasound Enhances Reporter Gene Expression After Transfection of Vascular Cells In Vitro. (Circulation 1999; 99;2617-2620).
Journal Article 1 (circ.ahajournals.org, PDF Document) 2. Normalizing Genetic Reporter Assays: Approaches and Considerations for Increasing Consistency and Statistical Significance. (Cell Notes, Issue 17, 2007, www.promega.com)
Journal Article 2 (www.promega.com, PDF Document)