Cell and Cell Line Characterization

Cell and Cell Line Characterization A. Doyle, Wellcome Trust, London, England G. Stacey, National Institute of Biological Standards and Control, South Mimms, England Encyclopedia of Cell Technology Copyright © 2000 by John Wiley & Sons, Inc. All rights reserved. DOI: 10.1002/0471250570.spi031 Article Online Posting Date: January 15, 2003 Keywords: cell culture; cell storage; cell types Abstract Introduction Chromosome Analysis Slide Preparation—Karyology—Materials Monolayer Cultures Giemsa Staining Isoenzyme Analysis Preparation of Reagents Materials Evaluation of Results DNA Fingerprinting and DNA Profiling Preparation and Restriction Enzyme Digestion of High-Molecular-Weight DNA from Cell Lines Materials Southern Blot Materials Visualization of DNA Fingerprints by Chemiluminescence with the Multilocus Probe 33.15 Materials Interpretation of DNA Fingerprint Patterns Antibody Staining for Species Verification Preparation of Antiserum Materials Procedure for Test Cell Line Viability Testing using the MTT Assay Reagents Materials Procedure Neutral Red Assay Reagents Materials Conclusions Bibliography 1.​ Introduction The characterization of a new cell line is essential and should be carried out at both the earliest passage of cultures once established and at frequent intervals thereafter (R.J. Hay, 1988). The occurrence of cross-contamination is not merely anecdotal; documented cases have been widely reported (1, 2). Some earlier reports indicated that levels of cross-contamination may exceed 30% of cultures tested (3). The preferential inclusion of suspect cultures in these reports does not detract from the fact that cross-contamination is a serious problem. The classic example is that of HeLa contamination (1), for which conventional cytogenetic analysis in association with isoenzyme studies was used to verify the species and, with human samples, the race of origin. This is particularly easy in the case of the HeLa cell line, since it has characteristic cytogenetic markers, and in isoenzyme analysis, the type B rather than the more usual type A glucose-6-phosphate dehydrogenase isoform is present. A more traditional technique is that of immunological characterization—in essence a known sample of cells or tissue is used to raise an antibody in the rabbit. The antibody is then used in a fluorescence study against the test cells and an anti-rabbit globulin conjugated with FITC (fluorescein isothiocyanate) is added and then can be visualized on a fluorescence microscope. In recent times molecular biology has had a significant influence on cell characterization and DNA fingerprinting provides a particularly useful tool for identity testing. 2. Chromosome Analysis The determination of the chromosomal complement of a cell line provides a direct method of confirming the species of origin. It also allows the detection of gross aberrations in chromosome number and/or morphology. Cytogenetic analysis is very useful for specific identification of cell lines with unique chromosome markers. In one study of 47 cell lines reported by O'Brien et al. (4), two cell lines could not be differentiated by eight separate enzyme tests, but were readily distinguished by karyology. However, it should be borne in mind that very careful interpretation is required to differentiate cell lines of normal karyotype beyond the level of species. To follow is an outline of the standard technique for cytogenetic analysis for a monolayer cell culture. 2.1. Slide Preparation—Karyology—Materials Hypotonic trypsin/versene solution (Hypo-TV) at 37 °C for harvesting adherent cells Hypotonic KCl solution (Hypo-KCl) at room temperature Heparin, 10 units per mL Glacial acetic acid at room temperature (it is important to use this fresh) Working colcemid solution (100 or 50 µg/mL) at room temperature Slides, precleaned, wet, and chilled 3. Monolayer Cultures 3.1. Day 1 Passage the cells 1 day before use. One 80–90% confluent 75-cm2 culture is generally sufficient for a chromosome preparation. 3.2. Day 2 1. Add colcemid solution to the cells at a final concentration of 0.01–0.03 µg/mL and incubate at 37 °C for 1 h. The incubation time for colchicine treatment is 45 min for a fast-growing culture, 1–1.5 h for an “average” culture, 2–3 h for diploid cell cultures, and over 3 h for slow-growing cultures. 2. Label each centrifuge tube (15-mL size) and add 0.6 mL fetal bovine serum. 3. After incubation with colcemid, collect the medium in a centrifuge tube and centrifuge at 150 g for 5 min. After decanting supernatant, resuspend cells in Hypo-KCl and add to the harvest from the Hypo-TV treatment (Step 7). 4. Add 6 mL Hypo-TV per 75-cm2 flask and incubate at 37 °C for 10 min. 5. Aspirate to suspend cells. If a large number of cells are still attached, treat the cultures with Hypo-TV again. 6. Transfer the cell suspension into the tube prefilled with serum (Step 2) and centrifuge 150 g for 5 min. 7. Decant supernatant without disturbing the cell pellet; add Hypo-KCl little by little with continuous agitation to a final volume of 4 mL. After centrifugation and removal of supernatant, add KCl gradually by increasing the volume with intermittent agitation. Finally, add the contents of one (or two) Pasteur pipettes to each. Between each addition, agitate cells by gentle pipetting. 8. Leave at room temperature for 10 min, then centrifuge and decant most of the supernatant, leaving an equal volume to the cell pellet for resuspending cells. 9. Resuspend cells gently but thoroughly to make a uniform cell suspension. 10. Add freshly prepared ice cold 1:3 glacial acetic acid/methanol fixative slowly as indicated for Hypo-KCl (Step 8) to a total of 4 mL and leave for 15 min at room temperature. The fixative must be made fresh just before use. After mixing, it must always be kept on ice. 11. Decant as much supernatant as possible without disturbing the cell pellet. Repeat Step 10. 12. Repeat Step 10; however, leave at room temperature for 10 min and then centrifuge. 13. Remove most of the supernatant, leaving behind a volume, which is about ten times the cell mass, and then resuspend the cells. 14. Pick up a wet, clean, chilled slide with a pair of forceps. Hold the frosted edge with your fingers and shake off excess water using a fanning motion. (Be sure the frosted side faces up.) Hold the slide slightly downward (at about a 30 ° angle), place a few drops of cell suspension onto the upper edge of the slide just below the frosted edge to let the suspension run down slowly, and at the same time blow gently and evenly over the surface to spread cells over the entire surface. Wipe off excess liquid from the edges and back of the slide, and air dry by leaving the slide on a paper towel. 15. When dried, examine the quality of the chromosomes under phase microscopy to assess metaphase spreads and cell density. Cells should be evenly distributed and chromosomes from each cell should be close together but without frequent overlapping. If the quality is good, make at least 10 slides. Experience suggests that slides prepared the same day as the cell harvest give the best quality spreading and staining. However, it may be advisable to also store the fixed cell suspension for repeat analyses. To make slides from this stored material, cells are again treated in fresh fixative at least three times with a 10-min incubation between each centrifugation. 16. Use an indelible marker pen to record on the slide: cell line, number of slide, date of preparation. 17. Leave slides overnight at room temperature for further drying and then store in a slide box. Slides kept at room temperature may be used even after one month of storage and still give good staining. As slides get older, staining results become unpredictable. Keep slides in a cold and dry environment, or seal in a container filled with an inert gas (e.g., argon) and store at 4 °C. 3.3. Day 3 Stain slides. 4. Giemsa Staining The most commonly used procedure for chromosome banding is Giemsa staining (5, 6). The method allows for a simple chromosome count and provides an estimation of the rate of polyploidy, a key issue related to cell stability. 1. Immerse slides in 1% Giemsa solution in a Coplin jar, at room temperature, for 30 min. 2. Rinse thoroughly with distilled water. Use a squeezing bottle to stream water over the slide surface evenly. Examine under 63x and 100x water-emulsion objective lenses. 5. Isoenzyme Analysis Isoenzyme analysis is used for the speciation of cell lines and for the detection of contamination of one cell line with another, although a relatively high level of contamination is necessary (>10%). This method utilizes the property that isoenzymes have similar substrate specificity, but different molecular structures, which in turn affects their electrophoretic mobility. Each species therefore has a characteristic isoenzyme mobility pattern. While the species of origin of a cell line is usually indicated with only two isoenzyme tests (lactate dehydrogenase and glucose-6-phosphate dehydrogenase) (4), specific identification of a cell line requires a larger battery of tests (3). Generally the use of four isoenzymes can give adequate results (7). To follow is the standard technique for isoenzyme analysis utilizing the “Authentikit” system supplied by Innovative Chemistry, Inc., Marshfield, MA. 6. Preparation of Reagents 1. Cell extraction buffer Prepare a 50 mM solution of Tris (for final volume of 100 mL) in 80 ml water. Adjust to pH 7.5 (either 1 M HCl or 1 M NaOH). Add 1 mM EDTA and then 2% Triton X-100. Adjust the pH if necessary and make up the volume with deionized water. Store at 4 °C. 2. Enzyme substrates. The list of tabulated migration distances and ratios supplied by the kit manufacturer includes the following: Aspartate aminotransferase (AST) Glucose-6-phosphate dehydrogenase (G6PD) Lactate dehydrogenase (LDH) Malate dehydrogenase (MD) Mannose phosphate isomerase (MPI) Nucleoside phosphorylase (NP) Peptidase B (Pep B) 3. Visualization. Detection of the enzyme bands is provided by an insoluble purple formazan dye when 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) is reduced in the presence of phenazine methosulfate (PMS) and the appropriate substrate. 4. Standards. Extracts of the mouse cell line NCTC Clone L929 (standard) and the human cell line HeLa (control) can be obtained from the kit manufacturer in lyophilized form. If required, they can be prepared from growing cultures, but these should be obtained from a documented and authenticated source (i.e., culture collection). 6.1. Materials Agarose film–agarose gel on transparent polystyrene sheet (Authentikit, Innovative Laboratories, Marshfield, MA) Barbital buffer 0.05 mM, pH 8.6 Hamilton syringe, 5–10 µL with tapered needle Electrophoresis cell (Innovative Laboratories) Power supply to provide 160 V DC, constant voltage (Innovative Laboratories) A minimum of 107 cells is necessary for each analysis. 1. Prepare a cell pellet (centrifugation at 150 g for 5 min). Attached cells must be removed with the appropriate enzyme. 2. Decant the culture medium, resuspend the cell pellet in 15 ml Earle's balanced salt solution at 4 °C. Centrifuge the cells at 150 g for 5 min. 3. Mark the volume of the pellet on the side of the tube, and then drain the buffer by inverting the tube on absorbent paper. Dry the inside of the tube, taking care not to touch the pellet. Add an equal volume of cell extraction buffer to the pellet, mix with a micropipette tip, and stand on ice for 15 min. 4. Mix again and examine a drop sandwiched between a glass microscope slide and a coverslip. The cells should now be lysed. If necessary, mix again and leave for another 15 min on ice. 5. Centrifuge at 150 g at 4 °C for 10 min. Remove the supernatant with a micropipette to a microtube and place on ice. 6. Into each chamber of the electrophoresis cell base place 95 mL 0.05 M barbital buffer. Ensure that the liquid levels are equal. Then fill the chamber cover with 500 mL ice-cold water. 7. Remove the agarose gel (Authentikit) from the rigid mould by peeling the film back. Lay the gel face up with the plastic film against the work surface. 8. Using a Hamilton syringe with a Teflon tip, place 1 µL sample into each of the pre-formed slots. 9. Position the gels into the chamber cover with the agarose facing upward. Ensure that the positive and negative indicators on the gel match those of the chamber. 10. Connect the power and run the gel at 160 V DC for 25 min. The time must be precise. 11. Switch off the power, lift the chamber cover, and stand on paper towels to drain for 30 sec. 12. Remove the agarose film and place gel side up on the work surface. Pour the contents of the enzyme substrate vial down the center of the gel, and using the edge of a pipette, spread over the gel until fully covered. 13. Drain off excess substrate by holding the corner of the film on a paper towel. Place gel side up onto an incubator tray with a sheet of damp filter paper. Cover the tray with the lid and incubate at 37 °C until the enzyme bands appear (i.e., up to 20 min). 14. Stop the reaction by immersing in tap water and leave until fully destained. 15. Dry the gels at 60 °C in an oven for 60 min. 7. Evaluation of Results To complete this it is necessary to obtain a copy of the tabulated results from Authentikit (Fig. 1). Figure 1. Migration distances for various species. Source: Cell and Tissue Culture: Laboratory procedures. A. Boyle and J.B. Griffiths (eds.) John Wiley & Sons, Chichester. [Full View] 1. Measure the distance the enzyme bands have migrated. 2. Check that the values for the standard and control are within ±2 mm of the listed values. 3. If they are, proceed to Step 5. If not, apply the following correction factor: (1) 4. Multiply the actual values for the control and test samples by the factor and compare these with the listed values. They should now be within 1–2 mm of their listed values. 5. Calculate the migration ratio as follows: (2) The values for the control and test samples are compared with those given in the tabulated results provided. Identify the species, giving similar ratios for each enzyme examined, and list them. By a process of elimination, the species should be identified (Fig. 1). Note: If a cell line of human origin is being checked, it is necessary to include G6PD in the examination, as this enzyme can detect HeLa contamination. This cell line has the Type B G6PD, which is very unusual in cell lines derived from Caucasians. The enzymes selected for analysis may initially depend on the supposed species of the cell lines being tested. However, it is recommended to use at least four enzymes to confirm such preliminary results. The routine use of glucose-6-phosphate dehydrogenase and lactate dehydrogenase will give an indication of species of origins, and nucleoside phosphorylase and malate dehydrogenase will provide confirmations of a particular species match. 8. DNA Fingerprinting and DNA Profiling The term DNA fingerprinting has been applied to a variety of methods involving PCR or Southern blotting and a range of primer sequences or nucleic acid probes for different targets in the genome. All these methods are based on a determination of the size of DNA sequences, which are hypervariable between different individuals. However, the specific genetic identity of an individual can only be demonstrated by analyzing a number of loci. This can be achieved by analyzing a number of specific loci using a panel of DNA probes (8) or multiplex PCR. In addition, there are techniques that identify multiple genetic loci simultaneously by the use of random primers (9) or DNA probes that cross-hybridize with a range of related hypervariable DNA sequences. The latter technique is commonly called multilocus DNA fingerprinting and was the first DNA fingerprinting technique to be reported (10). This technique has been used widely for human paternity and forensic studies and in analysis of population genetics for a wide range of animals and plants. The original probes 33.15 and 33.6 reported by Jeffreys (11) have proven extremely valuable for authentication and quality control of animal cell cultures (12, 13), and DNA fingerprinting is now regarded as a useful component of routine identity testing for cell lines used in the production of biological reagents (14). The probe 33.15 has proved to be extremely useful for analyzing cell lines from a wide range of species and has been validated for routine use at the European Collection of Cell Cultures (ECACC, Salisbury, UK) (13, 15). The current methodology uses an oligonucleotide for the consensus core sequence of the original 33.15 probe (11) and is described in the following. 9. Preparation and Restriction Enzyme Digestion of High-Molecular-Weight DNA from Cell Lines 9.1. Materials Cell suspension solution: 0.2 M sodium acetate (pH 7.5), RNAse (0.34 mg/mL) Chloroform/isoamyl alcohol (24:1) Phenol/chloroform: Phenol (saturated solution of pH >7.6 in tris buffer) mixed 1:1 with chloroform/isoamylalcohol (24:1) TE buffer: 10 mM Tris, 1 mM disodium EDTA, pH 7.5 Electrophoresis buffer: 10 fold dilution in distilled water of 10 × TBE (162 g/L Tris, 46.3 g/L ortho-boric acid, 9.5 g/L disodium EDTA, pH 7.5) with 0.5 mg/L ethidium bromide Depurination solution: 0.25 M hydrochloric acid in distilled water Denaturation solution: 1.5 M sodium chloride, 0.5 M sodium hydroxide Neutralization solution: 3.0 M sodium chloride, 0.5 M Tris-HCl pH 7.5, 1 mM disodium EDTA X20 SSC: 175 g/L sodium chloride, 88.2 g/L trisodium citrate at pH 7.4 Stop mix: 1 mL loading buffer (e.g., Sigma G2526), 0.25 mL ethidium bromide (10 mg/mL), 0.25 mL 10XTBE. Sterile microtubes Microfuge 10% (w/v) SDS pH 7.2 RNAse type 1A (10 mg/mL) Proteinase K (10 mg/mL) Micropipetter (1 mL) and phenol-resistant disposable tips Incubator or infrared lamp uv spectrophotometer Restriction enzymes (HinfI, HaeIII) and digestion buffer (from enzyme manufacturer) Sterile distilled water (0.22 µm filtered) Submarine horizontal electrophoresis equipment Agarose (low endo-osmosis) Whatman 3 MM chromatography paper (or equivalent) Nylon membrane (20 cm × 20 cm) (e.g., Hybond-N, Amersham) Wash trays (approximately 28 cm × 22 cm × 4 cm) 1. Resuspend a pellet of approximately 5 × 106 cells in a microtube with 200-µL cell resuspension. 2. Add 25 µL 10% SDS and mix by inversion. 3. Incubate at 50 °C for 5 h. 4. Microfuge (10,000 g for 5 min) and transfer the upper aqueous phase to a fresh labeled microtube. 5. Mix by inversion with 200 µL phenol/chloroform. 6. Microfuge (10,000 g for 5 min) and transfer the aqueous phase to a fresh microtube. 7. Repeat steps 4, 5 and 6 twice. 8. Mix by inversion with 200 µL chloroform and repeat step 6. 9. Precipitate the DNA with 2 volumes of cold absolute ethanol. 10. The DNA is pelleted by microfuging (12,000 g for 5 min), the alcohol is aspirated, and the pellet partially dried in air (37 °C or under an infrared lamp). 11. The DNA pellet is resuspended and dissolved at 37 °C overnight with mixing) in 30–40 µL TE and quantified by uv spectrophotometry at 260 and 280 nm. An optical density (OD) value of 1 corresponds to a double-stranded DNA concentration of 50 µg/mL. The ratio A260/A280 gives an indication of nucleic acid purity, and this ratio should be 1.8 for pure DNA. 12. A 5 µg sample of the DNA is digested at 37 °C overnight after mixing with Hinf1 or HaeIII enzyme (5–10 units/µg DNA), 4 µL 10 × enzyme reaction buffer and sterile water up to a total volume of 40 µL. 13. The enzyme digest is stopped by addition of 8 µL 6 × stop mix. 10. Southern Blot 10.1. Materials Lambda/HinDIII molecular weight marker Electrophoresis buffer (as in the preceding) Plastic wash trays (1 L capacity) 20xSSC (see above) uv transilluminator (315 nm) 1. Quantify the DNA in each digest and run an analytical 0.8% agarose gel of at least 20 cm in length and electrophorese 5 µg di