Mammalian Cell Culture - STLCC

BIOTECHNOLOGY I – MAMMALIAN CELL CULTURE

EBoedeker revised 8/24/2011 Page 5-1

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 CO2 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



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



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.

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



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

CO2 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

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

sterile 50 ml tubes

* 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


EBoedeker revised 8/24/2011

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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.)



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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



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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



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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% CO2 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.

95% confluent 98% confluent



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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

Clumps of cells due to poor

triteration at last passing.

75-80% confluent

80% confluent Cells after trituration –

still too many clumps



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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



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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.



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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 CO2 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



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