LAB 5 Mammalian Cell Culture Tim Warmke and Catherine Sweeny, former SLCC-FV Biotechnology students, helped in the writing and initial teaching of this lab. STUDENT GUIDE GOAL The purpose of this lab is to provide the biotech student with experience in counting and seeding mammalian cells for culturing and to practice the fine art of aseptic technique. NOTE: You must take the Use and Care of the BSC quiz on the class Blackboard Website, scoring at least 80%, before you will be allowed to perform this lab. OBJECTIVES After completion, the student should be able to 1. Demonstrate excellent aseptic technique 2. Set up and use a Biological Safety Cabinet (BSC) for performing cell culture techniques 3. Operate an inverted light microscope 4. Estimate percent confluence of a cell culture 5. Determine the density of a cell culture using a hemocytometer 6. Dilute a cell culture for seeding 7. Maintain a cell culture for several weeks by passing the cells regularly TIMELINE: This lab will be continued through the first half of the semester BACKGROUND Tissue culture was first performed in the early 1900 s for studying animal cells free of the variations caused by homeostasis in the whole animal. For the first 50 years of tissue culturing, the technique was performed with fragments of explanted tissue, so the term tissue culture has stuck in spite of the fact that since the 1950 s dispersed cells in culture, or cell culture, has been practiced. A cell culture, therefore, refers to cultures derived from dispersed cells taken from the original tissue, from a primary culture, or from a cell line or cell strain. 1 In the 1950 s, it was discovered that human tumor cells could give rise to continuous cell lines (e.g., HeLa cells) 2 and since the 1960 s studies have been performed using normal cells with finite lifespans. 3 Further background information about cell culture is supplied in the Appendix of this manual. This lab is intended to apply knowledge of aseptic technique and cell culturing. They are extremely vital skills in biotech research and industrial labs. If aseptic technique is followed, there will be minimal chance of contamination from fungi, bacteria, or other hazardous organisms. All cell culture activities, except for the actual counting will be performed in the BSC, which must be thoroughly decontaminated using 70% ethyl alcohol (EtOH) and UV light. The CO 2 incubator where the cells will be grown must also be cleaned with 70% EtOH and the shelves should be sprayed with beta-iodine. If the incubator has a water bath, it must be washed and Fisher Clear Bath added when it is Page 5-1
refilled. Media bottles must be autoclaved within two days of use, even if they have been previously autoclaved. The neck and top of the bottle should not be touched after autoclaving and they should be stored away from any bacterial plates or other incubators used in microbiology classes. As you can see, you must be meticulously clean when performing tissue culture, especially if the lab serves a dual purpose. Before coming to lab, read the Corning handout General Procedures for the Cell Culture Laboratory in the Appendix section of this manual. It contains information on how to prepare a lab for tissue or cell culture. You should also read through the lecture notes Preparation and Use of the Biological Safety Cabinet. In this exercise you will prepare the media needed for the cells to survive and grow to an optimal density that could be used in an experiment. Although we will not use these cells for any experiment, the objective is to practice techniques involved with mammalian cell culture and to maintain the culture over a period of weeks. Cells will survive and continue to proliferate if they are diluted and used to inoculate new media on a regular basis. Even with this careful attention, however, cells can only be passed in this way about 20 times and then they will lose the ability to divide. The exception to this is cancer cells, which proliferate indefinitely, as long as they are passed correctly. In order to grow cells at the correct density for experimentation, they must be counted using a hemocytometer and then the concentration of the cells must be calculated in cells/ml of solution. Once the concentration is established, the dilution factor required can be calculated so that fresh media can be seeded for continued growth. See the sample calculations after Table 1. Table 1-Useful Numbers for Cell Culture gives the required seeding density required for different volumes of fresh media. If cells were confluent, a 1/4 dilution should be sufficient, however. If you need the culture to grow more slowly, dilute even more, for example, 1/8. Table 1. Useful Numbers for Cell Culture Page 5-2
Examples for Calculating Cell Density 1. Starting Concentration = 8.4 x 10^5 cells per milliliter Seeding density required for T-25 flask = 0.7 x 10^6 cells (See Table 1.) Final volume in T-25 flask = 5ml Use a ratio and solve for X: 8.4 x 10^5 cells = 0.7 x 10^6 cells 1 ml X X = 0.83 ml = 830 μl Therefore, for proper seeding density, add 830 μl of the original culture to 4.17 ml of new culture in your T-25 culture flask. However, what is usually done is to add 1 ml of cells into 4 ml of new culture, which is close enough when you are just passing cells. 2. Starting Concentration = 5.2 x 10^6 cells per milliliter Seeding density required for each well of a 6 well plate = 0.3 x 10^6 cells (See Table 1.) Final volume of each well of a 6 well plate = 12.5 ml You therefore need a total volume of 6 x 12.5 ml or 75 ml. To be on the safe side, you would make sure you have more volume, so 80 ml of diluted cells should be sufficient. (5.2 x 10^6) = 0.3 x 10^6 1 ml X X = 0.06 ml = 60 μl Since you need 12.05ml media + 450ul of original cell suspension SAFETY GUIDELINES Check with the instructor on the type of cells you will use. We have used NIH 3T3 CRL- 1658, Mus musculus (mouse) embryonic fibroblast cells. Go to the American Type Culture Collection website to view more information about these cells, which are Biosafety Level I. (http://www.atcc.org/common/catalog/cellbiology/cellbiologyindex.cfm). All pipettes, tubes, and flasks must be disposed in the biohazard bag for autoclave sterilization. Correct sterile technique must be practiced so that aerosols do not leave the BSC. Ethyl alcohol is highly flammable. Use appropriate caution (e.g., no open flames) when cleaning the BSC. Safety glasses, lab coats and gloves are required. Page 5-3
MATERIALS To be ordered Sigma D5796 media Sigma N4637 serum (Fetal Bovin Serum FBS) Sigma T3924 trypsin Biosafety Level I cells (*NIH 3T3 CRL-1658, *RAW 264.7, *MM3MG, or *BHK-21 cells could be used check with the instructor) Phenol red (ph 6.8-yellow 8.2-red indicator) Trypan Blue aliquot 50 µl per 1.5 ml microcentrifuge tube per team CO 2 gas Sterile Phosphate Buffered Saline (PBS), 1M or Hank s balanced salt solution For Bringing up Cells T-25 Flasks Media: 100 ml of 10% FBS D5796 (+ 1% Antibiotics if instructor chooses) Serological Pipettes 15 ml Conical Centrifuge Tubes (Sterile for aliquoting Serum and Antibiotics) Per Class (Amounts depend on # of groups): D5796 Media FBS Cultured cells or ATCC cell stock 100X penicillin/streptomycin stock, [final] = 100 µg/ml check with instructor 1 M sterile PBS or Hank s (except for RAW cells) 0.05% Trypsin-EDTA (except for RAW) Trypan Blue 50 µl aliquot/team Autoclaved or Sterile 100 ml Media bottles I per team (Glass is better) 5 ml, 10 ml, 25 ml serological pipettes Cell scrapers (only for RAW cells) Flasks - T-25 sterile 50 ml tubes Spray bottle for 70% EtOH Parafilm 15 ml tubes and holders (racks) Hemocytometers Cell counters Light microscopes Inverted light microscope Water bath (cleaned w/ fresh water) Labeling tape To be left in the BSC: 10% bleach solution container for used pipettes two electric pipette pumps beaker for old media sterilization * NIH 3T3 CRL-1658, Mus musculus (mouse) embryonic fibroblast cells RAW 264.7, Abelson murine leukemia virus-induced tumor cells MM3MG, Mouse mammary gland epithelial cells BHK-21, Baby Hamster Kidney cells Page 5-4
Procedure Turn on the water bath to 37 C. Part I. Sterilization of the Biological Safety Cabinet (BSC) 1. With gloved hands, sterilize the BSC with 70% EtOH. 2. Gather equipment and supplies needed. 3. Spray your gloved hands and anything else you are placing into the BSC with 70% EtOH before you move it in. Place supplies and equipment back at least 4 inches from the front airflow. Remember to keep contaminated items towards the back of the BSC. Place the bin for trash and the used pipette container in the back of the BSC. Equipment and supplies you will need in the hood: Autoclaved or Sterile 100 ml Media bottles (Glass is better) 5 ml, 10 ml, 25 ml serological pipettes Cell scrapers or 0.05% Trypsin-EDTA @ 37 C T-25 Flasks Spray bottle of 70% EtOH Parafilm Sterile 15 ml tubes and rack Sterile 50 ml tubes and rack 10% bleach solution in capped bottle Container for used pipettes Two electric pipette pumps Beaker for contaminated media disposal label P-1000 automatic micropipette and tips 4. Turn on the UV light and blower for 15 minutes before working. Part II. Preparation and Use of the BSC 1. Turn off the UV light but leave the blower running. 2. Once your hands are in the BSC, DO NOT MOVE THEM OUT until you are finished. Move your hands slowly while working. 3. No one should be walking in the vicinity of the BSC and the doors of the laboratory should stay closed while anyone is working. 4. Do not block the front vents or rear exhaust. 5. Clean up immediately any spilled cells. (Note: If BSLII cells or higher, you must wait 10 minutes before resuming work. We will be using only BSLI cells.) 6. Do not work over open cultures; this allows contaminants to fall into the sterile cells or media. 7. If you are the last one to use the BSC, add 10% bleach solution to the contaminated media, which should turn from pink to clear. Spray all surfaces with 70% EtOH before exiting and wipe dry with Kim Wipes. (Note: Bleach evaporates quickly from solutions, even in capped bottles, so the 10% bleach solution needs to be remade often. If the media does not lose its color quickly, the bleach solution is not strong enough.) 5-5
8. When finished, spray gloved hands with 70% EtOH right after exiting the BSC. Remove gloves, inside out, and place in the biological/hazardous waste. Immediately wash hands with germicidal soap. 9. Leave the fan running. Remove your lab coat before exiting the lab. Part III. Preparation of the Media What you must take into the BSC: DMEM media Antibiotic stock FBS NOTE: You must verify your calculations with your lab partner and the instructor before proceeding. 1. Using a sterile container in the safety cabinet, add a 10% volume of FBS. 2. Add a 1% volume of antibiotic stock [100 μg.ml]. 3. Bring to volume (BTV) of 100ml with DMEM. 4. Swirl gently. 5. Place in a 37 C water bath for 20 minutes. While waiting, the instructor will demonstrate how to use the hemocytometer; explain how to determine cell density, and how to calculate the correct seeding density for T-25 flasks. Review Table 1 and the sample calculations given in the Background information at the beginning of this lab. Part IV. Cell Counting What you must take into the BSC: 1.5 ml microcentrifuge tube with 50 µl Trypan Blue prepared media (see Part III) @37 C 0.05% Trypsin-EDTA @37 C or cell scrapers sterile Phosphate Buffered Saline (PBS), 1 M (except for RAW cells) Flask of confluent cell culture started previously 5 and 10 ml serological pipettes and pump P-1000 automatic micropipetter and tips 1. Obtain a T-25 flask of confluent cells and observe with the inverted light microscope. Note the percent confluence (see examples in section VI of this lab). Document this (draw or photograph) in your lab notebook. 2. Remove old media with serological pipette and discard. 3. Gently add 3 ml of sterile 1 M PBS to the side of the flask. 4. Swirl the PBS gently and briefly across bottom of flask. 5. Remove PBS solution. 6. Gently add 0.5 to 1 ml of 0.05% Trypsin-EDTA to the side of the flask. 7. Swirl the Trypsin-EDTA gently and briefly across bottom of flask and return the solution to the side of the flask. Cells may be placed at 37ºC for 2-5 minutes to facilitate the dissociation from the surface of the flask. Cells must be checked frequently (every 1-2 minutes) if placed back in incubator at 37ºC. Over 5-6
trypsinization and cell lysis can occur very quickly at this elevated temperature. Alternatively, you can just allow the cells to sit at room temperature for 2-5 minutes, but do monitor them regularly. When the cells are falling off the surface, you will be able to see a slurry of material moving across the flask when you pick it up and tip it one direction or another. 8. Add 5 ml of fresh media directly to the bottom of the flask. Cells should release into the media. You may see masses floating in the liquid. 9. Triturate 5 times with a serological pipette. 10. Verify that cells are mostly separated, by viewing the cells in the flask with the inverted light microscope. 11. Return to the BSC and transfer 250 l of cell suspension to the 1.5 ml microfuge tube containing 50 µl of Trypan Blue and close. 12. TURN OFF THE ELECTRIC PUMP! 13. Exit the BSC properly, taking this tube of cell culture to the bench. Place the flask of culture back in the incubator standing upright (this helps prevents reattachment of the cells to the bottom surface). You can leave your media in the hood. 14. At the bench, calculate the dilution factor of the removed cells. Set up a light microscope and a hemocytometer. You will also need your P20 automatic micropipetter and tips. 15. Remove 10 µl of the cell culture and apply to the hemocytometer. Place the hemocytometer onto the stage of your microscope and view the counting grid with cells on 40x total magnification. Use the guide below to determine both viable and dead cell numbers. Dead cells are distinguished from viable cells by their uptake of Trypan Blue stain, which diffuses into dead cells but not live cells. Your instructor will demonstrate this in class. Determine the cell density and the percent viability of the original culture. 16. Disinfect the bench, microscope and hemocytometer with 70% EtOH. This cell counting guide is included with cell counting equipment such as hemocytometers 5-7
Part V. Seeding (Subculturing) Cells 1. Determine the amount of original cell suspension to add to a T-25 Flask (See Table 1 and the examples given in the background information.) All calculations must be included in your lab notebook. 2. In the BSC, add the original cell suspension to the correct amount of fresh media in a new T-25 flask. 3. Incubate the cells @ 37 C with 5% CO 2 until at least 70% confluent. Part VI. Passing Cells Every 2-4 days, your cell culture must be passed. You must make arrangements to come into the lab and check your cultures for growth. When the cells are at least 70% confluent they can be passed. 1. Obtain your team s T-25 flask of cells and observe under the inverted light microscope. Note their percent confluence. Document this in your lab notebook (draw or photograph). Examples are shown, below. 98% confluent 95% confluent 5-8
80% confluent Cells after trituration still too many clumps Clumps of cells due to poor triteration at last passing. 75-80% confluent 2. In the BSC, remove old media with serological pipettes and discard. 3. Gently add 3 ml of sterile 1 M PBS to the side of the flask. (Except for RAW) 4. Swirl the PBS gently across bottom of flask briefly. (Except for RAW) 5. Remove PBS solution. (Except for RAW) 6. Gently add 0.5-1 ml of 0.05% Trypsin-EDTA to the side of the flask. 7. Swirl the Trypsin-EDTA gently across bottom of flask briefly and return the solution to the side of the flask. 8. Once cells are moving freely, add 5 ml of media directly to the bottom of the flask. Triturate 5 times with a serological pipette. 9. Cap the flask and view the culture using the inverted light microscope. If the cells are still in clumps, triturate again and check again. 10. When cells are well separated, use Table 1 to determine the correct volume to transfer to a new T-25 flask with fresh media labeled with your team number or your initials, the type of cell, the passage number and the date. If cells were confluent, a 1/4 dilution should be sufficient. If you need the 5-9
culture to grow more slowly, dilute even more, for example, 1/8. For the NIH 3t3 cells, for Tuesdays classes, a ¼ dilution should be sufficient. For Thursdays, a harsher slip should allow the cells to make it until the following Tuesday (so a 1/8 split). 11. Place the passed cells into the incubator. Make sure that the flask is placed in the correct position, with the neck pointing upward (see the photo). Correct Incorrect 5-10
ALTERNATIVE PROTOCOL for RAW cells only Releasing Cells by Scraping An alternative method for removing cells from the flask is to use a sterile cell scraper. Cells are in larger clumps and it will require more trituration to separate them enough for passing. 1. In the laminar hood, remove the old media with serological pipette and discard. 2. Prepare a new T-25 flask by adding 5 ml of sterile media. 3. Gently apply 5 ml of fresh media to the side of the flask with the cells, but not directly on top of the cells. 4. Using a cell scraper, gently scrape the bottom of the flask side to side for several minutes until the entire surface has been scraped free of cells. 5. With an electric pipette pump and a 5 ml serological pipette, slowly triturate 20-30 times without adding bubbles of air so as not to make the media foam. 6. Cap the flask and view the culture using the inverted light microscope. If cells are still in clumps, triturate again and check again. 7. When cells are well separated, pass one-fourth of the culture to the new T-25 flask with fresh media. If it will be longer than 3 days until you can pass the cells again, transfer only one-eighth of the triterated cell culture to the 5 ml of new media. 8. Label the newly inoculated T25 flask and place into the incubator. Make sure that the flask is placed in the correct position, with the neck pointing upward (see the photos, above). DATA ANALYSIS Drawings and/or photographs of the magnified cell cultures would be appropriate for your laboratory notebook. See the last page of this lab for an example of how to maintain your laboratory notebook over the next several weeks while passing the cells. In this case, it is permissible to record observations from different days on the same page. In addition to written descriptions, as seen in the example, you are encouraged to include drawings or photos. How to use the image capture setup on the inverted light microscope MAKE SURE THE CAMERA IS ON (See the MTI power router on the top of the computer) 1. Turn on the microscope and position your culture flask over the objective. The 4X objective should be in place for determination of confluence. Use the 10X or 20X only to view individual cells. 2. Use the focus adjustment to bring the image into sharp focus. Adjust light with the light diaphragm lever and/or with the condenser adjustment knob at the back right of the microscope. 3. Switch the light path to go through the camera and the USB video capture device rather than through the ocular lenses by rotating the small knob just behind the right focus adjustment. 5-11
4. On the Desktop, click Start Programs Accessories Imaging 5. The imaging software will open; Click on File and select Acquire Image. 6. A pop-up menu will appear showing the image. If the menu is black, then the image path knob has not been rotated to send it through the camera to the video capture device. (See step 3.) NOTE: To view the image on the screen using the ceiling projector, just turn on the projector. 7. Use the focus adjustment knob on the microscope to get a sharp image on the pop-up menu on the computer. 8. Click on Capture Still Image and a larger image will appear. 9. Click print to send the image to the printer. It will be actual size, which is about 6 cm high x 8 cm long. 10. If you want to save the image, do so by clicking Save. Save it to the desktop and when you are finished capturing your images, you will need to transfer the saved image to your own floppy or portable hard drive (mini-cruiser, key, etc.). The file will be removed from the hard drive when the computer is shut down. 11. Please TURN OFF the microscope and replace the cover before you leave. QUESTIONS Refer to the Lecture notes, information in this lab and the handout from Culture of Animal Cells by R. Ian Freshney in the lab manual Appendix. 1. Are you doing tissue culture or cell culture in this lab? 2. Why is a CO 2 incubator used for mammalian cell culture? Explain the chemistry involved. 3. Why is serum-free selective media used to propagate cells of a specific phenotype? 4. Explain why the strict aseptic conditions required for mammalian cell culture are not required with bacterial cell cultures used in microbiology. 5. What is the purpose of using Trypsin-EDTA? Trypan Blue? Sources: 1. Freshney, R. Ian. (1994) Culture of Animal Cells, Third Edition. Wiley Publishing, New York. 2. Gey, G.O., Coffman, W.D., Kubicek, M.T. (1952) Tissue culture studies of the proliferative capacity of cervical carcinoma and normal epithelium. Cancer Research, 12:364-365. 3. Hayflick, L., Moorhead, P.S. (1961) The serial cultivation of human diploid cell strains. Exp. Cell Res. 25:585-621. 4. Sweeney, Catherine. BS Biology, SLCC-CS Biotechnology 5. Warmke, Timothy. SLCC-AAS Biotechnology 5-12
Title: Project No. Book No. From Page No. To Page No. 5-13