Part a

Maintenance

This is Part A, Maintenance, under the module topic, Cell Culture Techniques. This topic part has one section: Content Tutorials

Content Tutorials

I. MAINTENANCE – Cell Culture Experimental Overview

Cultures should be examined daily, observing the morphology, the color of the medium and the density of the cells. A tissue culture log should be maintained that is separate from your regular laboratory notebook. The log should contain: the name of the cell line, the medium components and any alterations to the standard medium, the dates on which the cells were split and/or fed, a calculation of the doubling time of the culture (this should be done at least once during the semester), and any observations relative to the morphology, etc.

A. Growth Pattern

Cells will initially go through a quiescent or lag phase that depends on the cell type, the seeding density, the media components, and previous handling. The cells will then go into exponential growth where they have the highest metabolic activity. The cells will then enter into stationary phase where the number of cells is constant, this is characteristic of a confluent population (where all growth surfaces are covered).

B. Subculturing & Harvesting

Cells are harvested when the cells have reached a population density which suppresses growth. Ideally, cells are harvested when they are in a semi- confluent state and are still in log phase. Cells that are not passaged and are allowed to grow to a confluent state can sometimes lag for a long period of time and some may never recover. It is also essential to keep your cells as happy as possible to maximize the efficiency of transformation. Most cells are passaged (or at least fed) three times a week.

1. Suspension culture. Suspension cultures are fed by dilution into fresh medium.

2. Adherent cultures. Adherent cultures that do not need to be divided can simply be fed by removing the old medium and replacing it with fresh medium. When the cells become semi-confluent, several methods are used to remove the cells from the growing surface so that they can be diluted:

Mechanical – A rubber spatula can be used to physically remove the cells from the growth surface. This method is quick and easy but is also disruptive to the cells and may result in significant cell death. This method is best when harvesting many different samples of cells for preparing extracts, i.e., when viability is not important.

Proteolytic enzymes – Trypsin, collagenase, or pronase, usually in combination with EDTA, causes cells to detach from the growth surface. This method is fast and reliable but can damage the cell surface by digesting exposed cell surface proteins. The proteolysis reaction can be quickly terminated by the addition of complete medium containing serum

EDTA – EDTA alone can also be used to detach cells and seems to be gentler on the cells than trypsin. The standard procedure for detaching adherent cells is as follows:

a. Visually inspect daily. b. Release cells from monolayer surface: -Wash once with a buffer solution – Treat with dissociating agent. – Observe cells under the microscope and incubate until cells become rounded and loosen when flask is gently tapped with the side of the hand. – Transfer cells to a culture tube and dilute with medium containing serum. – Spin down cells, remove supernatant and replace with fresh medium. c. Count the cells in a hemacytometer, and dilute as appropriate into fresh medium.

C. Media and growth requirements

1. Physiological parameters – temperature – 37C for cells from homeotherms – pH – 7.2-7.5 and osmolality of medium must be maintained – humidity is required – gas phase – bicarbonate concentration and CO2 tension in equilibrium – visible light, can have an adverse effect on cells; light induced production of toxic compounds can occur in some media; cells should be cultured in the dark and exposed to room light as little as possible

2. Medium requirements: (often empirical) – Bulk ions – Na, K, Ca, Mg, Cl, P, Bicarb or CO2 – Trace elements – iron, zinc, selenium – sugars – glucose is the most common – amino acids – 13 essential – vitamins – B, etc. – choline, inositol – serum, contains a large number of growth promoting activities such as buffering toxic nutrients by binding them, neutralizes trypsin and other proteases, has undefined effects on the interaction between cells and substrate, and contains peptide hormones or hormone-like growth factors that promote healthy growth. – antibiotics, although not required for cell growth, antibiotics are often used to control the growth of bacterial and fungal contaminants.

3. Feeding – 2-3 times/week.

4. Measurement of growth and viability. The viability of cells can be observed visually using an inverted phase contrast microscope. Live cells are phase bright; suspension cells are typically rounded and somewhat symmetrical; adherent cells will form projections when they attach to the growth surface. Viability can also be assessed using the vital dye, trypan blue, which is excluded by live cells but accumulates in dead cells. Cell numbers are determined using a hemacytometer.

5. Observing cells. Everything that the cell plates or flasks come into contact with must be wiped with 70% ethanol. In complying with this, closely follow each step:

1. Wipe incubator door and microscope stage with 70% ethanol.
2. Wipe gloved hands with 70% ethanol.
3. Check cells under microscope.
4. Place cells back into incubator.

6. Feeding Cells

1. Do steps 1-4. Listed above.
2. Wipe laminar hood with 70% ethanol.
3. Take out media from refrigerator, wipe down with 70% ethanol and place in the hood.
4. Measure out the desired amount of media and pipette into a centrifuge tube.
5. Set the centrifuge tube on bench to warm up for at least 15 minutes.
6. Put hood UV light for at least 15 minutes.
7. Take cells out of the incubator and place inside the hood.
8. Wipe media tube with 70% ethanol and place inside the hood.
9. Aspirate off existing media from the flask or microplate.
10. Pipette fresh media into the flask.
11. Check cells under microscope.
12. Place cells back in the incubator.
13. Wipe surface of hood with 70% ethanol.
14. Clean aspirator hose with autoclaved SigmaClean water bath solution.
15. Leave UV light on for at least 5 minutes.

7. Subculturing UMR-106 cells (can be modified for other cell lines)

1. Take out 0.25% Trypsin-EDTA from -80°C freezer and let it thaw.
2. Do steps 1-4 in “Observing Cells” and steps 2-9 in “Feeding Cells”.
3. Pipette 6 ml of 0.25% Trypsin-EDTA into flask and incubate for two minutes.
4. Check flask under microscope to see evidence of cell detachment.
5. Tap bottom of flask lightly to detach more cells.
6. Pipette 6 ml of media into flask and pipette up and down against cell wall until cells have become suspended.
7. Pipette cell suspension into a 15 mL centrifuge tube.
8. Place tube into ultra centrifuge for 5 minutes at 600 rpm with a counter balance.
9. Wipe centrifuge tube with 70% ethanol and place back into the hood.
10. Aspirate the media, leaving a small layer of media on top of the cell pellet.
11. Slowly pipette 5 ml of media into the tube and re-suspend the cells.
12. Pipette out 100 ul of media into eppendorf tube.
13. Use this eppindorf for cell counting. Pipette out 20 ul of cell suspension and 80 ul of trypan blue into another eppendorf tube. Pipette enough to coat the surface of the hemocytometer.
14. Count cells and calculate the number of cells to seed into the flask. (Approximately 1,000,000 cells for a T-75 flask, 20,000 cells/well for a 24 well plate, 3200 cells/well for a 96 well plate.)
15. Re-suspend cells and pipette cell suspension into a flask. Add fresh media.
16. Check cells under microscope and place in incubator.
17. Wipe surface of hood with 70% ethanol.
18. Clean aspirator hose with autoclaved SigmaClean” water bath solution.
19. Turn on UV light for at least five minutes.