Week 9.Cell.Culture.FA23

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Morgan State University *

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Feb 20, 2024

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Week 9 Introduction to Cell Culture Learning Objectives Identify cell culture medium components, buffer, and supplements Describe NIH 3T3 cell growth properties, and attachment to surfaces. Use of PBS washing and trypsin/EDTA digestion in passaging NIH-3T3 cells Diagram a laminar flow cell culture hood with HEPA filtered air. Describe the use of 70% alcohol to surface-sterilize the work surface of the hood and the materials to be brought into the hood. Demonstrate sterile pipetting technique within the hood. Describe correct disposal of solid waste and waste media from hood Determine cell concentration using a hemocytometer Ethanol fixation of NIH 3T3 cells to prepare for PI staining Background Mammalian cell cultures are an essential tool in biology because they allow rapid growth and proliferation of different cell types for experimental analysis. In order to successfully work with mammalian cell lines, they must be grown under controlled conditions and require their own specific growth medium. We are using NIH 3T3 cells. This is a mammalian cell line derived from the fibroblasts of a mouse. They are cultured in Dulbecco’s Modified Eagles Medium (DMEM+). The “+” signifies that the DMEM contains 10% (v/v) bovine calf serum and 1% (v/v) antibiotic/antimycotic. Bovine calf serum (BCS) is a growth supplement for cell culture media because of it contains growth promoting factors. It supplies many defined and undefined components that have been shown to satisfy specific metabolic requirements for the culture of cells. Antibiotic-antimycotic is used to prevent the growth of bacteria and fungi in culture. The 100X solution has 10,000 units/ml of penicillin, 10,000 ug/ml of streptomycin and 25 ug/ml of amphotericin B. Mouse 3T3 cells are adherent, which means they attach to the bottom of the flask and grow in a single monolayer. To guarantee consistency, their growth must be monitored at regular intervals. When a cell line reaches about 80% confluence, the cells must be subcultured (passaged) to ensure proper growth and health of the cells. A confluency of 80% means that 80% of the surface of a culture vessel is covered with cells. If needed, you can passage cells before 80% confluency. When we passage cells, we must detach the cells from the flask and then dilute the cells with media in a new flask. Then the cells must be allowed to reattach a new surface. The cells need to be incubated at 37°C and at 5% CO 2 . The pH of the medium is dependent on the delicate balance of dissolved carbon dioxide (CO 2 ) and bicarbonate (HCO 3 – ), and changes in the atmospheric CO 2 can alter the pH of the medium. Therefore, it is necessary to use exogenous CO2 when using media buffered with a CO 2 -bicarbonate based buffer. Passaging NIH 3T3 cells involves releasing cells from the culture flask and from each other to form a suspension of single cells. This requires trypsin and EDTA. The 3T3 cells secrete collagen, fibronectin and other extracellular matrix (ECM) proteins to attach themselves to the surface of the culture flask. Also, cells adhere to other cells through cadherins, ca lcium-d ependent adherin g proteins. To remove all these attachments, trypsin with EDTA is used. Trypsin is a protease and cleaves the ECM proteins, releasing the cells from the dish. EDTA is a chelator which removes calcium

ions from solution and allows cadherin-mediated cell-cell contacts to come apart. Once the cadherins lose their calcium, they become sensitive to trypsin digestion and are removed from the cell surface. The first step in passaging cells is removing the DMEM+ growth medium and washing cells with phosphate buffered saline (PBS) which lacks calcium. The DMEM+ growth medium has 10% bovine calf serum (BCS) and must be removed before adding trypsin because serum contains trypsin inhibitor. So we will remove the old medium and wash the cells with 4 mLs of PBS, removing both serum and calcium. After removing the PBS wash, we will add 1 mL of trypsin/EDTA and allow it to work. When the cells have detached, we will add 4 mLs of PBS plus 0.500 mL of BCS to dilute and inhibit the added trypsin. After centrifugation, the supernatant will be discarded in the hood, and the cell pellet will be resuspended in 1 mL DMEM + 10% BCS + 1% AA and kept on ice. After counting, a small portion of the cell suspension will be used to seed another culture flask in new DMEM + 10% BCS + 1% AA, and the cell passaging will be complete. Even though our media includes antibiotics and antifungals, it is very easy to contaminate cell cultures because the media is full of nutrients and growth factors. Contamination of your cells could lead to contamination throughout the entire incubator because the air circulator of the incubator will spread the contaminants into other cultures. To keep our cultures free of bacteria and fungi, we will be using cell culture hoods and aseptic techniques. The two videos below will better explain these techniques. Watch this video to learn how the Cell Culture hood stays sterile https://www.jove.com/science-education/5036/an-introduction-to-working-in-the- hood Watch this video to learn how to culture your cells using Aseptic technique https://www.youtube.com/watch?v=cyJ2zUbLMMo Your TA will walk through more specifics about your hood and aseptic techniques specific for your NIH-3T3 cells.  Our Cell Culture hoods use vertical laminar flow of air passed through HEPA filters.  Always wear gloves in the cell culture room.  Never open your cell culture dishes outside of the hood including when imaging on the microscope.  You must place your materials only on the solid surface of the hood.  Spray 70% ethanol on everything that goes in the hood including your gloved hands.  Make sure you have all your materials inside the hood before you begin any protocols.  Do not open any media or cells until you are completely ready. You do not want to take your hands out of the hood unless absolutely necessary.  You and your partner will receive your own sterile tubes of DMEM+, PBS, Trypsin/EDTA and BCS for your use over the weeks we will be growing cells. Please mark the tubes and caps with your names and hood number using the Fisher Brand makers that are resistant to 70% ethanol. Use only your own solutions. While you are

working in the cell culture hoods, the Trypsin/EDTA and BCS tubes will be kept in a small insulated, ice-filled cup inside the hood. DMEM+ and PBS tubes will be kept at room temperature in the hood.  For long-tern storage, all DMEM+ and PBS tubes from your section will be stored in a Styrofoam rack at 4° with your TA’s name and lab day on it. Likewise, your Trypsin/EDTA and BCS tubes will all be stored in a single rack at -20°. If you require more media, PBS, trypsin or BCS, your TA will refill your tubes under sterile conditions. You will also do the first step for PI staining. You will be ethanol-fixing cells, which will allow the stain into the cells. More background will be given next week. Materials Microscope - Inverted microscope Cell Culture Hoods, 70% Ethanol, mini-ice bucket (an insulated cup to hold ice in the hood) 37°C, 5% CO 2 Incubator Pipet Aids, Pipets, pipet tips, solid and liquid waste disposal containers Media, Trypsin/EDTA, sterile PBS, BCS 25 cm 2 flasks, 15mL conical tubes Confluent NIH 3T3 cells in a 25 cm 2 flask Cold 70% Ethanol, non-sterile PBS, Centrifuges Procedure Do NOT use the Bunsen burners this week. People will be spraying ethanol around the room as well as in the cell culture room. We don't want anything in flames. Experiment: Passage cells While waiting for a Cell Culture hood, l ook at your cells using the inverted microscope. Do not take the cap off the culture flask outside of the cell culture hood. You can see your cells through the flask. Set up an inverted phase contrast scope with your smart phone attached. Focus on your cell culture, and take a photograph. Your TA will show you how to work in your hood.  You must place your materials only on the solid surface of the hood.  Spray 70% ethanol on everything that goes in the hood including your gloved hands.  Make sure you have all your materials inside the hood before you begin any protocols.  Do not open any media or cells until you are completely ready. You do not want to take your hands out of the hood unless absolutely necessary.

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You will need pipets, tips, pi-pump and micropipettors and two waste beakers, one for solids, one for liquids, in your hood, and a 70% alcohol spray outside your hood. You will need a 15 mL conical tube, a 25 cm 2 flask, your cells (in a 25 cm 2 flask), MEM+, Trypsin/EDTA, sterile PBS, and BCS. 1. Remove expired media from cells without disturbing the cells. Do this by tilting the flask and pouring the media out into the liquid waste beaker. 2. Pipette 4 mLs sterile PBS into the flask at the neck, and tilt the flask to rinse cells. Pour out the PBS without disturbing the cells. 3. Pipette 1mL trypsin/EDTA into the flask. Then tilt the flask side to side until the trypsin/EDTA has contacted the entire dish surface. Leave in the hood, and observe the cells every two minutes using the inverted microscope. Please make your observations quickly. Others may need to use the microscope. 4. Under the scope, when the cells round up and detach, tilt the flask until there are no attached cells. The cells should visibly be floating around in the trypsin at this point. 5. Add 4 mLs PBS and 0.500 mL BCS to the flask. Tilt the plate, and pipette the PBS-BCS-trypsin/EDTA solution over the flask to rinse any remaining adherent cells off the surface. This usually only takes 3-4 times. 6. Put the ~5 mLs of cell suspension into a 15 mL conical tube. 7. Centrifuge 1,200 rpm, 5 min. (1,000 x g in IEC centrifuge). Spin in a group. 8. In the hood, pour the entire supernatant into the waste beaker, and add 1 mL of PBS to the cell pellet, resuspending the cells by gently finger-vortexing until the pellet is fully dispersed (no clumps). Put your cell suspension on ice. 9. Determine the cell concentration using a hemocytometer. See next section. 10. Before pipetting your cells into the new flask, finger-vortex the cell suspension to bring all cells up into the PBS. Cells left undisturbed for several minutes will fall to the bottom of the 15 mL tube. Add 6.6 x 10 4 cells to the new flask with 4 mLs of fresh DMEM+ (“+” means it has 10% BCS and 1X antibiotics). Swirl the flask to distribute your cells across the dish evenly, and label with your name. Put the flask in your section’s basket. Your TA will place it in the incubator for next week. 11. Take the 15mL conical with your cell suspension back to your bench to fix with ethanol. 12. Please clean the hood, and store your solutions:

Store your DMEM+ and PBS in your section’s rack in the refrigerator. Store your Trypsin/EDTA and BCS in your section’s rack in the freezer. Solid waste to red bag trash. Pour the liquid waste in your waste beaker into the 4L brown glass culture waste jug near the window sink in 184. Squirt a small amount of 10% bleach into your waste beaker, swirl and dispose in the waste jug. (All used media must be treated with bleach in the jug for at least 1 hour before it can be disposed of by pouring down the drain.) Wipe down hood surfaces with 70% alcohol. Ethanol fix cells for PI staining When fixing cells, we are killing the cells. The fixed cells can be worked with outside of the culture hood because they will no longer be used for culturing and do not need to be kept sterile. Use non-sterile PBS for this procedure. Do not use your sterile PBS. 1. Spin cells at 1200 x g for 5 minutes. 2. Remove supernatant using the P1000 pipet at your bench. 3. Wash your cells in 1mL PBS (keep in 15 mL conical tube). 4. Spin cells at 1200 x g for 5 minutes. 5. Repeat the PBS wash one more time (steps 2-4). 6. Resuspend your cells in 500 uL of non-sterile PBS. 7. Add 5 ml ice-cold 70% ethanol drop-wise while vortexing. If cells are not vortexed while ethanol is being added, they will be fixed to each other in clumps. The tube of cold ethanol should remain in ice at all times while 1 ml volumes are being withdrawn. 8. After ethanol addition is complete, keep cells on ice. Reseal the tube firmly. Label the tube with your initials, TA, lab room and day, and give the tube to your TA. 9. Cells will be stored at -20°C until next week’s PI staining and DNA content analysis. You will continue the protocol next week

Measuring Cells per mL in a Hemocytometer Passaging cells requires a count of the number of cells per mL. We will be counting our cells in a hemocytometer. Load 10 -15 uL of the cell suspension into a hemocytometer, and count the cells by phase contrast at 100X. The chamber formed by the thick cover glass resting on the support strips and the platform with the etched grad is 1/10 th of a millimeter deep. The chamber is filled by placing the filled tip of a pipettor on the platform next to the edge of the cover glass. The solution is slowly released until the volume between the cover glass and the platform is filled. Then the pipet tip is withdrawn. After counting, the hemocytometer is cleaned by running dH 2 O over the cover glass and the platform. Gently blot with a Kimwipe, and return to the box. To prevent the platform from being scratched, the hemocytometer should be stored face-down on the folded Kimwipe in the box, and the cover glass should be placed inside the plastic sleeve. Below is the pattern you will see when you focus on the hemocytometer slide at 100X phase contrast in an Axiostar, Primostar-Lumin or an inverted microscope. Above the central square millimeter, the 0.1 millimeter space between cover glass and etched Hemocytometer raised support strips moats thick cover glass platform with etched grid patterns thick cover m

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platform encloses a volume 1 x 10 -4 cubic centimeters. The formula for calculating cells/mL is cells in 1 sq mm x 10 4 = cells/mL. 1 mm In counting, if a cell is bisected by a line, count the cell if it is on the top or left line but not if it is on the bottom or right line. Count at least 200 cells. You are asked to count a total of at least 200 cells by counting all the cells in the 1 mm 2 central area. If the central area has less than 200 cells, count a second 1 mm 2 area of the same grid.

Name:___Raghu Chinta____________________ Sec_______ TA_______Emily__________________ Week 9 Lab Report Introduction to Cell Culture 10 pts 1. Photograph of your cell culture before passaging with magnification, type of microscopy and the estimate of percentage of confluence. Post the photograph here. 2 pts Phase Contrast 100x with a 7-10% confluence

2. From the hemocytometer count, calculate the total cells in the final cell suspension. Count was 36 cells in 9 grid squares A. Begin with cells per 1 mm 2 in the hemocytometer. 1 pt 36 cells/9 = 4 cells per 1 mm 2 B. Calculate cells per ml. 1 pt 4 cells/1 mm 2 * 1*10 4 = 4*10^4 cells/mL. C. Multiply by the volume of the final cell suspension to yield total cells. 2 pts 4*10^4 cells/ml * 1 mL = 4*10^4 cells. 3. What volume of cell suspension was added to the new 25 cm 2 flask for the new culture? 1 mL for the 40000 cells/mL 2 pts 4. If you begin this culture flask with 6.6 x 10 4 cells, calculate how many cells will be present when you next passage this flask, assuming a 24 hour cell division time. 2 pts Final cell count = 6.6*10^4 cells *2^(7) = 8448000 cells. Cell divides once every day for 7 days till next lab. The completed lab report can be uploaded in the Student Assessment section of the Canvas website. This report is due at midnight 6 days after Lab 9 has ended.

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Week 9.Cell.Culture.FA23

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