Basic Cell Culture-Methods in Molecular Biology vol.290-Humana Press 3rd Edition.pdf

Basic Cell Culture-Methods in Molecular Biology vol.290-Humana Press 3rd Edition.pdf

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Unformatted text preview: METHODS IN MOLECULAR BIOLOGY TM TM Volume 290 Basic Cell Culture Protocols THIRD EDITION Edited by Cheryl D. Helgason Cindy L. Miller Basic Cell Culture Techniques 1 1 Culture of Primary Adherent Cells and a Continuously Growing Nonadherent Cell Line Cheryl D. Helgason Summary Cell culture is an invaluable tool for investigators in numerous fields. It facilitates analysis of biological properties and processes that are not readily accessible at the level of the intact organism. Successful maintenance of cells in culture, whether primary or immortalized, requires knowledge and practice of a few essential techniques. The purpose of this chapter is to explain the basic principles of cell culture using the maintenance of a nonadherent cell line, the P815 mouse mastocytoma cell line, and the isolation and culture of adherent primary mouse embryonic fibroblasts (MEFs) as examples. Procedures for thawing, culture, determination of cell numbers and viability, and cryopreservation are described. Key Words: Cell culture; nonadherent cell line; adherent cells; P815; primary mouse embryonic fibroblasts; MEF; hemocytometer; viability; subculturing; cryopreservation. 1. Introduction There are four basic requirements for successful cell culture. Each of these will be briefly reviewed in this introduction. However, a more detailed description is beyond the scope of this chapter. Instead, the reader is referred to one of a number of valuable resources that provide the information necessary to establish a tissue culture laboratory, as well as describe the basic principles of sterile technique (1–4). The first necessity is a well-established and properly equipped cell culture facility. The level of biocontainment required (Levels 1–4) is dependent on the type of cells cultured and the risk that these cells might contain, and transmit, infectious agents. For example, culture of primate cells, transformed human cell lines, mycoplasma-contaminated cell lines, and nontested human cells require a minimum of a Level 2 containment facility. All facilities should be From: Methods in Molecular Biology, vol. 290: Basic Cell Culture Protocols, Third Edition Edited by: C. D. Helgason and C. L. Miller © Humana Press Inc., Totowa, NJ 1 01/Helgason/1-12 1 8/26/04, 9:09 AM 2 Helgason equipped with the following: a certified biological safety cabinet that protects both the cells in culture and the worker from biological contaminants; a centrifuge, preferably capable of refrigeration and equipped with appropriate containment holders that is dedicated for cell culture use; a microscope for examination of cell cultures and for counting cells; and a humidified incubator set at 37°C with 5% CO2 in air. A 37°C water bath filled with water containing inhibitors of bacterial and fungal growth can also be useful if warming of media prior to use is desired. Although these are the basic requirements, there are numerous considerations regarding location of the facility, airflow, and other design features that will facilitate contamination-free culture. If a new cell culture facility is being established, the reader should consult facility requirements and laboratory safety guidelines that are available from your institution’s biosafety department or the appropriate government agencies. The second requirement for successful cell culture is the practice of sterile technique. Prior to beginning any work, the biological safety cabinet should be turned on and allowed to run for at least 15 min to purge the contaminated air. All work surfaces within the cabinet should be decontaminated with an appropriate solution; 70% ethanol or isopropanol are routinely used for this purpose. Any materials required for the procedure should be similarly decontaminated and placed in or near the cabinet. This is especially important if solutions have been warmed in a water bath prior to use. The worker should don appropriate personnel protective equipment for the cell type in question. Typically, this consists of a lab coat with the cuffs of the sleeves secured with masking tape to prevent the travel of biological contaminants and Latex or vinyl gloves that cover all exposed skin that enters the biosafety cabinet. Gloved hands should be sprayed with decontaminant prior to putting them into the cabinet and gloves should be changed regularly if something outside the cabinet is touched. Care should be taken to ensure that anything coming in contact with the cells of interest, or the reagents needed to culture and passage them, is sterile (either autoclaved or filter-sterilized). The biosafety office associated with your institution is a valuable resource for providing references related to the discussion of required and appropriate techniques required for the types of cells you intend to use. A third necessity for successful cell culture is appropriate, quality controlled reagents and supplies. There are numerous suppliers of tissue culture media (both basic and specialized) and supplements. Examples include Invitrogen ( ), Sigma–Aldrich ( ), BioWhittaker ( ), and StemCell Technologies Inc. ( ). Unless otherwise specified in the protocols accompanying your cells of interest, any source of tissue-culture-grade reagents should be acceptable for most cell culture purposes. Similarly, there are numerous suppliers of the plasticware 01/Helgason/1-12 2 8/26/04, 9:09 AM Basic Cell Culture Techniques 3 needed for most cell culture applications (i.e., culture dishes and/or flasks, tubes, disposable pipets). Sources for these supplies include Corning (www. corning.com/lifesciences/), Nunc ( ), and Falcon (www. bdbiosciences.com/discovery_labware). Two cautionary notes are essential. First, sterile culture dishes can be purchased as either tissue culture treated or Petri style. Although either can be used for the growth of nonadherent cells, adherent cells require tissue-culture-treated dishes for proper adherence and growth. Second, it is possible to use glassware rather than disposable plastic for cell culture purposes. However, it is essential that all residual cleaning detergent is removed and that appropriate sterilization (i.e., 121°C for at least 15 min in an autoclave) is carried out prior to use. If the three above-listed requirements have been satisfied, the final necessity for successful cell culture is the knowledge and practice of the fundamental techniques involved in the growth of the cell type of interest. The majority of cell culture carried out by investigators involves the use of various nonadherent (i.e., P815, EL-4) or adherent (i.e., STO, NIH 3T3) continuously growing cell lines. These cell lines can be obtained from reputable suppliers such as the American Tissue Type Collection (ATCC; ) or DSMZ (the German Collection of Microorganisms and Cell Cultures) ( mutz/mutzhome.html). Alternatively, they can be obtained from collaborators. Regardless of the source of the cells, it is advisable to verify the identity of the cell line (refer to Chapters 4 and 5) and to ensure that it is free of mycoplasma contamination (refer to Chapters 2 and 3). In addition to working with immortalized cell lines, many investigators eventually need or want to work with various types of primary cells (refer to Chapters 6–21 for examples). Bacterial contaminations, as a consequence of the isolation procedure, and cell senescence are two of the major challenges confronted with these types of cell. The purpose of this chapter is to explain the basic principles of cell culture using the maintenance of a nonadherent cell line, the P815 mouse mastocytoma cell line, and adherent primary mouse embryonic fibroblasts (MEF) as examples. Procedures for thawing, subculture, determination of cell numbers and viability, and cryopreservation are described. 2. Materials 2.1. Culture of a Continuously Growing Nonadherent Cell Line (see Note 1) 1. P815 mastocytoma cell line (ATCC, cat. no. TIB-64). 2. High-glucose (4.5 g/L) Dulbecco’s Modified Essential Medium (DMEM). Store at 4°C. 3. Fetal bovine serum (FBS) (see Note 2). Sera should be aliquoted and stored at –20°C. 01/Helgason/1-12 3 8/26/04, 9:09 AM 4 Helgason 4. Penicillin–streptomycin solution. 100X stock solution. Aliquot and store at –20°C (see Note 3). 5. L-Glutamine, 200 mM stock solution. Aliquot and store at –20°C. 6. DMEM+ growth medium: high-glucose DMEM (item 2) supplemented with 10% FBS, 4 mM glutamine, 100 IU penicillin, and 100 µg/mL streptomycin. Prepare a 500-mL bottle under sterile conditions and store at 4°C for up to 1 mo (see Note 4). 7. Trypan blue stain (0.4% w/v trypan blue in phosphate-buffered saline [PBS] filtered to remove particulate matter) or eosin stain (0.14% w/v in PBS; filtered) for determination of cell viability. 8. Tissue-culture-grade dimethyl sulfoxide (DMSO) (i.e., Sigma) stored at room temperature. 9. Freezing medium, freshly prepared and chilled on ice, consisting of 90% FBS and 10% DMSO (see Note 5). 2.2. Culture of Primary Mouse Embryonic Fibroblasts 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. High-glucose (4.5 g/L) DMEM (see Subheading 2.1.). FBS (see Subheading 2.1.). Penicillin–streptomycin solution (100X) (see Subheading 2.1.). MEF culture medium. DMEM supplemented with 10% FBS and 1X (100 IU penicillin and 100 µg/mL streptomycin) antibiotics. Dulbecco’s Ca2+- and Mg2+-free PBS (D-PBS). D-PBS can be purchased as 1X or 10X stocks from numerous suppliers or a 1X solution can be prepared in the lab as follows: Dissolve the following in high-quality water (see Note 6): 8 g/L NaCl, 0.2 g/L KCl, 0.2 g/L KH2PO4, 2.16 g/L Na2HPO4·7H2O; adjust pH to 7.2. Filter-sterilize using a 0.22-µm filter and store at 4°C. 0.25% Trypsin–0.5 mM EDTA (T/E) solution (see Note 7). Store working stocks at 4°C. Freezing medium (see Subheading 2.1.). Timed pregnant female mouse (see Note 8). 70% Ethanol solution or isopropanol. Two sets of forceps and scissors; one set sterilized by autoclaving at 121°C for 15 min. Fine forceps (sterile) (Fine Science Tools, cat. no. 11272-30). Small fine scissors (sterile). 18-Gage blunt-end needles (sterile) (StemCell Technologies Inc.). 3. Methods Prior to the initiation of any cell culture work, it is essential to ensure that all equipment is in optimal working condition. Moreover, if cell culture is to become a routine technique utilized in the laboratory, scheduled checks and regular maintenance of the equipment are required. A partial checklist of things to consider includes the following: check to ensure that the temperature and CO2 levels in the incubator are at the desired levels; check to be sure that the 01/Helgason/1-12 4 8/26/04, 9:09 AM Basic Cell Culture Techniques 5 water pan in the incubator is full of clean water and that it contains copper sulfate to inhibit bacterial growth; check to ensure that the water bath is at the required temperature and contains adequate amounts of clean water; check to ensure that the biological safety cabinet to be used is certified and operating correctly; ascertain that the centrifuge is cleaned and decontaminated. 3.1. Culture of a Continuously Growing Nonadherent Cell Line 3.1.1. Thawing Cryopreserved P815 Cells 1. In the biological safety cabinet, prepare one tube containing 9 mL of DMEM+ growth medium warmed to at least room temperature. 2. Remove one vial of cells from the storage container (liquid nitrogen or ultralow temperature freezer) (see Note 9). 3. Transfer the vial of cells to a 37°C water bath until the suspension is just thawed (see Note 10). 4. In the cell culture hood, use a sterile glass or plastic pipet to transfer the contents of the vial slowly into the tube containing the growth medium. 5. Centrifuge the cells at 1200 rpm (300g) for 7 min to obtain a pellet. 6. Aspirate the supernatant containing DMSO and suspend the cell pellet in 10 mL of DMEM+ growth medium (see Note 11). 7. Transfer the cells to a tissue culture dish (100 mm) and incubate at 37°C, 5% CO 2 . 8. Examine cultures daily using an inverted microscope to ensure that the culture was not contaminated during the freeze–thaw process and that the cells are growing. 3.1.2. Determination of Cell Number and Cell Viability Every cell line has an optimal concentration for maintaining growth and viability. Until sufficient experience is gained with a new cell line, it is recommended to check cell densities and viability every day or two to ensure that optimal health of the cultures is maintained. 1. Gently swirl the culture dish to evenly distribute the cell suspension. 2. Under sterile conditions, remove an aliquot (100–200 µL) of the evenly distributed cell suspension. 3. Mix equal volumes of cells and viability stain (eosin or trypan blue); this will give a dilution factor of 2. 4. Clean the hemocytometer using a nonabrasive tissue. 5. Slide the cover slip over the chamber so that it covers both sides. 6. Fill the chamber with the well-mixed cell dilution and view under the light microscope. 7. Each 1-mm2 square should contain between 30 and 200 cells to obtain accurate results (see Note 12). 01/Helgason/1-12 5 8/26/04, 9:09 AM 6 Helgason 8. Count the numbers of bright clear (viable) and nonviable (red or blue depending on the stain used) cells in at least two of the 1-mm2 squares, ensuring that two numbers are similar (i.e., within 5% of one another). Count all five of the 1-mm2 squares if necessary to ensure accuracy (see Note 13). 9. Calculate the numbers of viable and nonviable cells, as well as the percentage of viable cells, using the following formulas where A is the mean number of viable cells counted, B is the mean number of nonviable cells counted, C is the dilution factor (in this case, it is 2), D is the correction factor supplied by the hemocytometer manufacturer (this is the number required to convert 0.1 mm3 into milliliters; it is usually 104). Concentration of viable cells (per mL) = A × C × D Concentration of nonviable cells (per mL) = B × C × D Total number of viable cells = concentration of viable cells × volume Total number of cells = number of viable + number of dead cells Percentage viability = (number of viable cells × 100)/total cell number 3.1.3. Subculture of Continuously Growing Nonadherent Cells Maintenance of healthy, viable cells requires routine medium exchanges or passage of the cells to ensure that the nutrients in the medium do not become depleted and/or that the pH of the medium does not become acidic (i.e., turn yellow) as a result of the presence of large amounts of cellular waste. 1. View cultures under an inverted phase-contrast microscope. Cells growing in exponential growth phase should be round, bright, and refractile. If necessary, determine the cell density as indicated in Subheading 3.1.2. 2. There is no need to centrifuge the cells unless the medium has become too acidic (phenol red = yellow), which indicates the cells have overgrown, or if low viability is observed. 3. Transfer a small aliquot of the well-mixed cell suspension into a fresh dish containing prewarmed DMEM+ growth medium (see Note 14), ensuring that the resulting cell density is in the optimal range for the particular cell line. 4. Repeat this subculture step every 2–3 d to maintain cells in an exponential growth phase. 3.1.4. Cryopreservation of Continuously Growing Nonadherent Cells Continuous culture of cell lines can lead to the accumulation of unwanted karyotype alterations or the outgrowth of clones within the population. In addition, continuous growth increases the possibility of cell line contamination by bacteria or other unwanted organisms. The only insurance against loss of the cell line is to ensure that adequate numbers of vials (i.e., at least 10) are cryopreserved for future use. For newly acquired cell lines, cryopreservation of stock (master cell bank) vials should be done as soon as possible after the cell line has been confirmed to be free of mycoplasma (see Chapters 2 and 3). 01/Helgason/1-12 6 8/26/04, 9:09 AM Basic Cell Culture Techniques 7 1. View the cultures under a phase-contrast inverted microscope to assess cell density and confirm the absence of bacterial or fungal contamination. 2. Remove a small aliquot of the cells for determination of cell numbers as outlined in Subheading 3.1.2. Cells for cryopreservation should be in log growth phase with greater than 90% viability. 3. Prepare the cryopreservation vials by indicating the name of the cell line, the number of cells per vial, the passage number, and the date on the surface of the vial using a permanent marker (see Note 15). 4. Prepare the required volume of freezing medium as outlined in Subheading 2.1. and chill on ice. 5. Centrifuge the desired number of cells at 1200 rpm (300g) for 5–7 min and aspirate the supernatant from the tube. 6. Suspend the cells to a density of (1–2) × 106 cells/mL in the freezing medium. 7. Quickly aliquot 1 mL into each of the prepared cryovials using a pipet. Care is required to ensure that sterility is maintained throughout the procedure. 8. Place cryovials on dry ice until cells are frozen and then transfer to an appropriate ultralow temperature storage vessel (freezer or liquid-nitrogen tank) for longterm storage (see Notes 16 and 17). 3.2. Culture of Primary Mouse Embryonic Fibroblasts 3.2.1. Isolation of MEF 1. In order to obtain embryos at the desired stage of development set up female and male mice 14 d prior to the anticipated harvest date. On the following morning check for copulation plugs and remove the mated females to a separate cage. The day the plug is found is designated d 1. 2. On d 13 of pregnancy, sacrifice the females according to institutional guidelines. Spray or wipe the fur on the abdominal cavity of the dead mouse with 70% ethanol or isopropanol to reduce contamination risk and prevent fur from flying about. 3. Expose the skin of the abdominal cavity by cutting through the fur using a pair of scissors and forceps (sterility is not critical at this step). 4. Using the sterile scissors and forceps, cut through the abdominal wall and remove the uteri containing the embryos into a dish containing D-PBS. 5. In a biosafety cabinet, place the uteri into a sterile 100-mm dish. Dissect the embryos away from the yolk sac, amnion, and placenta using the sterile scissors and forceps. 6. Transfer the embryos to a clean dish and wash thoroughly to remove any blood. 7. Transfer the embryos to another sterile dish and use a pair of sterile fine forceps to pinch off the head and remove the liver from each embryo. 8. Transfer the remainder of the carcass into a fresh culture dish and gently mince the tissue using the fine sterile scissors into pieces small enough to be drawn into a 10-mL disposable pipet. 9. Add 0.5 mL of MEF culture medium per embryo to the minced tissue and draw the slurry up into a syringe of the appropriate volume through a sterile 18-gage 01/Helgason/1-12 7 8/26/04, 9:09 AM 8 Helgason blunt needle. Expel and draw up the minced tissue through the needle four to five times to generate small clumps of cells. 10. Add 10 mL of MEF culture medium per two embryos and culture in a 100-mm tissue-culture-treated (not Petri style) cell culture dishes. This is considered passage 1 (P1). 11. Incubate overnight at 37°C, 5% CO2 in a humidified cell culture incubator. Clusters of adherent cells should be visible, attached to the surface of the dish. Aspirate the medium containing floating cell debris and add an equal volume of fresh MEF culture medium. 12. Cultures should become confluent in 2–3 d. The expected yield is 1 × 107 cells per confluent 100-mm dish. 3.2.2. Subculture of MEF Mouse embryonic fibroblasts should be subcultured when they reach 80–90% confluence. If the MEF are allowed to reach 100% confluence, growth arrest can result with a decrease in the subsequent proliferative potential of the cells. 1. Aspirate the MEF medium from the dishes that have achieved the desired level of confluence and wash the monolayer of cells with 2–3 mL o...
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  • Fall '19
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