#C-O2 - Color labs

[vincent-ix]

Ok, I think Orange deserves a little more attention than that so-

Welcome to: The Color Labs!

This is Orange!

This is our Protagonist!

It uses he/they/it pronouns

He is an experiment, it's scientific name is C-O2

The name Orange was given to them by Smiles

They are just a silly little guy, ok?

Is classified as C2 but is actually the only one who functions properly minus Smiles and C-G5, also known as Green!

They may not have a mouth but he does eat! It pulls a Wally darling and eats with his eyes, which as caused incidents

MORE DRAWINGS AND REFERENCES COMING SOON!

[abbkineeu]

New Post has been published on Biotech Advisers

New Post has been published on http://www.bioadvisers.com/basic-techniques-mammalian-cell-tissue-culture/

Basic Techniques for Mammalian Cell Tissue Culture

Tissue culture technology has found wide application in the field of cell biology. Cell cultures are utilized in cytogenetic, biochemical, and molecular laboratories for diagnostic as well as research studies. In most cases, cells or tissues must be grown in culture for days or weeks to obtain sufficient numbers of cells for analysis. Maintenance of cells in long-term culture requires strict adherence to aseptic technique to avoid contamination and potential loss of valuable cell lines.

An important factor influencing the growth of cells in culture is the choice of tissue culture medium. Many different recipes for tissue culture media are available and each laboratory must determine which medium best suits their needs. Individual laboratories may elect to use commercially prepared medium or prepare their own. Commercially available medium can be obtained as a sterile and ready-to-use liquid, in a concentrated liquid form, or in a powdered form. Besides providing nutrients for growing cells, medium is generally supplemented with antibiotics, fungicides, or both to inhibit contamination.

As cells reach confluency, they must be subcultured or passaged. Failure to subculture confluent cells results in reduced mitotic index and eventually cell death. The first step in subculturing monolayers is to detach cells from the surface of the primary culture vessel by trypsinization or mechanical means. The resultant cell suspension is then subdivided, or reseeded, into fresh cultures. Secondary cultures are checked for growth, fed periodically, and may be subsequently subcultured to produce tertiary cultures, etc. The time between passaging cells depends on the growth rate and varies with the cell line. The Basic Protocol describes subculturing of a monolayer culture grown in petri plates or flasks; the Alternate Protocol 1 describes passaging of suspension cultures.

CAUTION: When working with human blood, cells, or infectious agents, appropriate biosafety practices must be followed. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly.

Trypsinzing And Subculturing Cells From A Monolayer

A primary culture is grown to confluency in a 60mm petri plate or 25 cm2 tissue culture flask containing 5 ml tissue culture medium. Cells are dispersed by trypsin treatment and then reseeded into secondary cultures. The process of removing cells from the primary culture and transferring them to secondary cultures constitutes a passage, or subculture.

Materials Primary cultures of cells

PBS/HBSS without Ca2+ and Mg2+, 37°C

Trypsin/EDTA solution , 37°C

Complete medium with serum: e.g., supplemented DMEM with 10% to 15% (v/v) FBS, 37°C

Sterile Pasteur pipets

37°C warming tray or incubator

Tissue culture plasticware or glassware including pipets and 25-cm2 flasks or 60-mm petri plates, sterile

NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4.

1. Remove all medium from primary culture with a sterile Pasteur pipet. Wash adhering cell monolayer once or twice with a small volume of 37°C PBS/HBSS without Ca2+ and Mg2+ to remove any residual FBS that may inhibit the action of trypsin.

Use a buffered salt solution that is Ca2+ and Mg2+ free to wash cells. Ca2+ and Mg2+ in the salt solution can cause cells to stick together.

If this is the first medium change, rather than discarding medium that is removed from primary culture, put it into a fresh dish or flask. The medium contains unattached cells that may attach and grow, thereby providing a backup culture.

2. Add enough 37°C trypsin/EDTA solution to culture to cover adhering cell layer.

3. Place plate on a 37°C warming tray 1 to 2 min. Tap bottom of plate on the countertop to dislodge cells. Check culture with an inverted microscope to be sure that cells are rounded up and detached from the surface.

If cells are not sufficiently detached, return plate to warming tray for an additional minute or two.

4. Add 2 ml 37°C complete medium. Draw cell suspension into a Pasteur pipet and rinse cell layer two or three times to dissociate cells and to dislodge any remaining adherent cells. As soon as cells are detached, add serum or medium containing serum to inhibit further trypsin activity that might damage cells.

If cultures are to be split 1/3 or 1/4 rather than 1/2, add sufficient medium such that 1 ml of cell suspension can be transferred into each fresh culture vessel.

5. Add an equal volume of cell suspension to fresh plates or flasks that have been appropriately labeled.

Alternatively, cells can be counted using a hemacytometer or Coulter counter and diluted to the desired density so a specific number of cells can be added to each culture vessel. A final concentration of ∼5 × 104 cells/ml is appropriate for most subcultures.

For primary cultures and early subcultures, 60-mm petri plates or 25cm2 flasks are generally used; larger vessels (e.g., 150mm plates or 75cm2 flasks) may be used for later subcultures.

Cultures should be labeled with date of subculture and passage number.

6. Add 4 ml fresh medium to each new culture. Incubate in a humidified 37°C, 5% CO2 incubator.

If using 75cm2 culture flasks, add 9 ml medium per flask.

Some labs now use incubators with 5% CO2 and 4% O2. The low oxygen concentration is thought to simulate the in vivo environment of cells and to enhance cell growth.

For some media it is necessary to adjust the CO2 to a higher or lower level to maintain the pH at 7.4.

7. If necessary, feed subconfluent cultures after 3 or 4 days by removing old medium and adding fresh 37°C medium.

8. Passage secondary culture when it becomes confluent by repeating steps 1 to 7, and continue to passage as necessary.

Passaging Cells In Suspension Culture

A suspension culture is grown in culture flasks in a humidified 37°C, 5% CO2 incubator. Passaging of suspension cultures is somewhat less complicated than passaging of monolayer cultures. Because the cells are suspended in medium rather than attached to a surface, it is not necessary to disperse them enzymatically before passaging. However, before passaging, cells must be maintained in culture by feeding every 2 to 3 days until they reach confluency (i.e., until the cells clump together in the suspension and the medium appears turbid when the flask is swirled).

NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4.

1. Feed cells as follows every 2 to 3 days until the cultures are confluent:

a. Remove flask of suspension cells from incubator, taking care not to disturb those that have settled to the flask bottom.

b. Aseptically remove and discard about one-third of the medium from flask and replace with an equal volume of prewarmed (37°C) medium. If the cells are growing rapidly, add an additional 10% medium by volume in order to maintain optimum concentration of 1 × 106 cells/ml. Gently swirl flask to resuspend cells.

c. Return flask to incubator. If there is <15 ml of medium in the flask, incubate flask in horizontal position to enhance cell/medium contact.

At higher volumes of medium the flask can be incubated in the vertical position.

If using a 25-cm2 flask, there should be 20 to 30 ml of medium in the flask at confluency.

2. On the days cultures are not being fed, check them by swirling flask to resuspend cells and observing color changes in the medium that indicate good metabolic growth.

3. When cultures are confluent (∼2.5 × 106 cells/ml), passage culture as follows:

a. Remove flask from incubator and swirl flask so that cells are evenly distributed in the medium.

b. Aseptically remove half of the volume of cell suspension and place into a fresh flask.

c. Feed each flask with 7 to 10 ml prewarmed medium and return flask to incubator.

Some labs prefer to split the cells 1:3 or 1:4, although increasing the split ratio will result in a longer interval before subcultures reach confluency.

[bioadvisers]

Bioadvisers shared on Biotech Advisers

Basic Techniques for Mammalian Cell Tissue Culture

Tissue culture technology has found wide application in the field of cell biology. Cell cultures are utilized in cytogenetic, biochemical, and molecular laboratories for diagnostic as well as research studies. In most cases, cells or tissues must be grown in culture for days or weeks to obtain sufficient numbers of cells for analysis. Maintenance of cells in long-term culture requires strict adherence to aseptic technique to avoid contamination and potential loss of valuable cell lines.

An important factor influencing the growth of cells in culture is the choice of tissue culture medium. Many different recipes for tissue culture media are available and each laboratory must determine which medium best suits their needs. Individual laboratories may elect to use commercially prepared medium or prepare their own. Commercially available medium can be obtained as a sterile and ready-to-use liquid, in a concentrated liquid form, or in a powdered form. Besides providing nutrients for growing cells, medium is generally supplemented with antibiotics, fungicides, or both to inhibit contamination.

As cells reach confluency, they must be subcultured or passaged. Failure to subculture confluent cells results in reduced mitotic index and eventually cell death. The first step in subculturing monolayers is to detach cells from the surface of the primary culture vessel by trypsinization or mechanical means. The resultant cell suspension is then subdivided, or reseeded, into fresh cultures. Secondary cultures are checked for growth, fed periodically, and may be subsequently subcultured to produce tertiary cultures, etc. The time between passaging cells depends on the growth rate and varies with the cell line. The Basic Protocol describes subculturing of a monolayer culture grown in petri plates or flasks; the Alternate Protocol 1 describes passaging of suspension cultures.

CAUTION: When working with human blood, cells, or infectious agents, appropriate biosafety practices must be followed. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly.

Trypsinzing And Subculturing Cells From A Monolayer

A primary culture is grown to confluency in a 60mm petri plate or 25 cm2 tissue culture flask containing 5 ml tissue culture medium. Cells are dispersed by trypsin treatment and then reseeded into secondary cultures. The process of removing cells from the primary culture and transferring them to secondary cultures constitutes a passage, or subculture.

Materials Primary cultures of cells

PBS/HBSS without Ca2+ and Mg2+, 37°C

Trypsin/EDTA solution , 37°C

Complete medium with serum: e.g., supplemented DMEM with 10% to 15% (v/v) FBS, 37°C

Sterile Pasteur pipets

37°C warming tray or incubator

Tissue culture plasticware or glassware including pipets and 25-cm2 flasks or 60-mm petri plates, sterile

NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4.

1. Remove all medium from primary culture with a sterile Pasteur pipet. Wash adhering cell monolayer once or twice with a small volume of 37°C PBS/HBSS without Ca2+ and Mg2+ to remove any residual FBS that may inhibit the action of trypsin.

Use a buffered salt solution that is Ca2+ and Mg2+ free to wash cells. Ca2+ and Mg2+ in the salt solution can cause cells to stick together.

If this is the first medium change, rather than discarding medium that is removed from primary culture, put it into a fresh dish or flask. The medium contains unattached cells that may attach and grow, thereby providing a backup culture.

2. Add enough 37°C trypsin/EDTA solution to culture to cover adhering cell layer.

3. Place plate on a 37°C warming tray 1 to 2 min. Tap bottom of plate on the countertop to dislodge cells. Check culture with an inverted microscope to be sure that cells are rounded up and detached from the surface.

If cells are not sufficiently detached, return plate to warming tray for an additional minute or two.

4. Add 2 ml 37°C complete medium. Draw cell suspension into a Pasteur pipet and rinse cell layer two or three times to dissociate cells and to dislodge any remaining adherent cells. As soon as cells are detached, add serum or medium containing serum to inhibit further trypsin activity that might damage cells.

If cultures are to be split 1/3 or 1/4 rather than 1/2, add sufficient medium such that 1 ml of cell suspension can be transferred into each fresh culture vessel.

5. Add an equal volume of cell suspension to fresh plates or flasks that have been appropriately labeled.

Alternatively, cells can be counted using a hemacytometer or Coulter counter and diluted to the desired density so a specific number of cells can be added to each culture vessel. A final concentration of ∼5 × 104 cells/ml is appropriate for most subcultures.

For primary cultures and early subcultures, 60-mm petri plates or 25cm2 flasks are generally used; larger vessels (e.g., 150mm plates or 75cm2 flasks) may be used for later subcultures.

Cultures should be labeled with date of subculture and passage number.

6. Add 4 ml fresh medium to each new culture. Incubate in a humidified 37°C, 5% CO2 incubator.

If using 75cm2 culture flasks, add 9 ml medium per flask.

Some labs now use incubators with 5% CO2 and 4% O2. The low oxygen concentration is thought to simulate the in vivo environment of cells and to enhance cell growth.

For some media it is necessary to adjust the CO2 to a higher or lower level to maintain the pH at 7.4.

7. If necessary, feed subconfluent cultures after 3 or 4 days by removing old medium and adding fresh 37°C medium.

8. Passage secondary culture when it becomes confluent by repeating steps 1 to 7, and continue to passage as necessary.

Passaging Cells In Suspension Culture

A suspension culture is grown in culture flasks in a humidified 37°C, 5% CO2 incubator. Passaging of suspension cultures is somewhat less complicated than passaging of monolayer cultures. Because the cells are suspended in medium rather than attached to a surface, it is not necessary to disperse them enzymatically before passaging. However, before passaging, cells must be maintained in culture by feeding every 2 to 3 days until they reach confluency (i.e., until the cells clump together in the suspension and the medium appears turbid when the flask is swirled).

NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4.

1. Feed cells as follows every 2 to 3 days until the cultures are confluent:

a. Remove flask of suspension cells from incubator, taking care not to disturb those that have settled to the flask bottom.

b. Aseptically remove and discard about one-third of the medium from flask and replace with an equal volume of prewarmed (37°C) medium. If the cells are growing rapidly, add an additional 10% medium by volume in order to maintain optimum concentration of 1 × 106 cells/ml. Gently swirl flask to resuspend cells.

c. Return flask to incubator. If there is <15 ml of medium in the flask, incubate flask in horizontal position to enhance cell/medium contact.

At higher volumes of medium the flask can be incubated in the vertical position.

If using a 25-cm2 flask, there should be 20 to 30 ml of medium in the flask at confluency.

2. On the days cultures are not being fed, check them by swirling flask to resuspend cells and observing color changes in the medium that indicate good metabolic growth.

3. When cultures are confluent (∼2.5 × 106 cells/ml), passage culture as follows:

a. Remove flask from incubator and swirl flask so that cells are evenly distributed in the medium.

b. Aseptically remove half of the volume of cell suspension and place into a fresh flask.

c. Feed each flask with 7 to 10 ml prewarmed medium and return flask to incubator.

Some labs prefer to split the cells 1:3 or 1:4, although increasing the split ratio will result in a longer interval before subcultures reach confluency.

[abbkine]

BioAdvisers said on Biotech Advisers

Basic Techniques for Mammalian Cell Tissue Culture

Tissue culture technology has found wide application in the field of cell biology. Cell cultures are utilized in cytogenetic, biochemical, and molecular laboratories for diagnostic as well as research studies. In most cases, cells or tissues must be grown in culture for days or weeks to obtain sufficient numbers of cells for analysis. Maintenance of cells in long-term culture requires strict adherence to aseptic technique to avoid contamination and potential loss of valuable cell lines.

An important factor influencing the growth of cells in culture is the choice of tissue culture medium. Many different recipes for tissue culture media are available and each laboratory must determine which medium best suits their needs. Individual laboratories may elect to use commercially prepared medium or prepare their own. Commercially available medium can be obtained as a sterile and ready-to-use liquid, in a concentrated liquid form, or in a powdered form. Besides providing nutrients for growing cells, medium is generally supplemented with antibiotics, fungicides, or both to inhibit contamination.

As cells reach confluency, they must be subcultured or passaged. Failure to subculture confluent cells results in reduced mitotic index and eventually cell death. The first step in subculturing monolayers is to detach cells from the surface of the primary culture vessel by trypsinization or mechanical means. The resultant cell suspension is then subdivided, or reseeded, into fresh cultures. Secondary cultures are checked for growth, fed periodically, and may be subsequently subcultured to produce tertiary cultures, etc. The time between passaging cells depends on the growth rate and varies with the cell line. The Basic Protocol describes subculturing of a monolayer culture grown in petri plates or flasks; the Alternate Protocol 1 describes passaging of suspension cultures.

CAUTION: When working with human blood, cells, or infectious agents, appropriate biosafety practices must be followed. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly.

Trypsinzing And Subculturing Cells From A Monolayer

A primary culture is grown to confluency in a 60mm petri plate or 25 cm2 tissue culture flask containing 5 ml tissue culture medium. Cells are dispersed by trypsin treatment and then reseeded into secondary cultures. The process of removing cells from the primary culture and transferring them to secondary cultures constitutes a passage, or subculture.

Materials Primary cultures of cells

PBS/HBSS without Ca2+ and Mg2+, 37°C

Trypsin/EDTA solution , 37°C

Complete medium with serum: e.g., supplemented DMEM with 10% to 15% (v/v) FBS, 37°C

Sterile Pasteur pipets

37°C warming tray or incubator

Tissue culture plasticware or glassware including pipets and 25-cm2 flasks or 60-mm petri plates, sterile

NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4.

1. Remove all medium from primary culture with a sterile Pasteur pipet. Wash adhering cell monolayer once or twice with a small volume of 37°C PBS/HBSS without Ca2+ and Mg2+ to remove any residual FBS that may inhibit the action of trypsin.

Use a buffered salt solution that is Ca2+ and Mg2+ free to wash cells. Ca2+ and Mg2+ in the salt solution can cause cells to stick together.

If this is the first medium change, rather than discarding medium that is removed from primary culture, put it into a fresh dish or flask. The medium contains unattached cells that may attach and grow, thereby providing a backup culture.

2. Add enough 37°C trypsin/EDTA solution to culture to cover adhering cell layer.

3. Place plate on a 37°C warming tray 1 to 2 min. Tap bottom of plate on the countertop to dislodge cells. Check culture with an inverted microscope to be sure that cells are rounded up and detached from the surface.

If cells are not sufficiently detached, return plate to warming tray for an additional minute or two.

4. Add 2 ml 37°C complete medium. Draw cell suspension into a Pasteur pipet and rinse cell layer two or three times to dissociate cells and to dislodge any remaining adherent cells. As soon as cells are detached, add serum or medium containing serum to inhibit further trypsin activity that might damage cells.

If cultures are to be split 1/3 or 1/4 rather than 1/2, add sufficient medium such that 1 ml of cell suspension can be transferred into each fresh culture vessel.

5. Add an equal volume of cell suspension to fresh plates or flasks that have been appropriately labeled.

Alternatively, cells can be counted using a hemacytometer or Coulter counter and diluted to the desired density so a specific number of cells can be added to each culture vessel. A final concentration of ∼5 × 104 cells/ml is appropriate for most subcultures.

For primary cultures and early subcultures, 60-mm petri plates or 25cm2 flasks are generally used; larger vessels (e.g., 150mm plates or 75cm2 flasks) may be used for later subcultures.

Cultures should be labeled with date of subculture and passage number.

6. Add 4 ml fresh medium to each new culture. Incubate in a humidified 37°C, 5% CO2 incubator.

If using 75cm2 culture flasks, add 9 ml medium per flask.

Some labs now use incubators with 5% CO2 and 4% O2. The low oxygen concentration is thought to simulate the in vivo environment of cells and to enhance cell growth.

For some media it is necessary to adjust the CO2 to a higher or lower level to maintain the pH at 7.4.

7. If necessary, feed subconfluent cultures after 3 or 4 days by removing old medium and adding fresh 37°C medium.

8. Passage secondary culture when it becomes confluent by repeating steps 1 to 7, and continue to passage as necessary.

Passaging Cells In Suspension Culture

A suspension culture is grown in culture flasks in a humidified 37°C, 5% CO2 incubator. Passaging of suspension cultures is somewhat less complicated than passaging of monolayer cultures. Because the cells are suspended in medium rather than attached to a surface, it is not necessary to disperse them enzymatically before passaging. However, before passaging, cells must be maintained in culture by feeding every 2 to 3 days until they reach confluency (i.e., until the cells clump together in the suspension and the medium appears turbid when the flask is swirled).

NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4.

1. Feed cells as follows every 2 to 3 days until the cultures are confluent:

a. Remove flask of suspension cells from incubator, taking care not to disturb those that have settled to the flask bottom.

b. Aseptically remove and discard about one-third of the medium from flask and replace with an equal volume of prewarmed (37°C) medium. If the cells are growing rapidly, add an additional 10% medium by volume in order to maintain optimum concentration of 1 × 106 cells/ml. Gently swirl flask to resuspend cells.

c. Return flask to incubator. If there is <15 ml of medium in the flask, incubate flask in horizontal position to enhance cell/medium contact.

At higher volumes of medium the flask can be incubated in the vertical position.

If using a 25-cm2 flask, there should be 20 to 30 ml of medium in the flask at confluency.

2. On the days cultures are not being fed, check them by swirling flask to resuspend cells and observing color changes in the medium that indicate good metabolic growth.

3. When cultures are confluent (∼2.5 × 106 cells/ml), passage culture as follows:

a. Remove flask from incubator and swirl flask so that cells are evenly distributed in the medium.

b. Aseptically remove half of the volume of cell suspension and place into a fresh flask.

c. Feed each flask with 7 to 10 ml prewarmed medium and return flask to incubator.

Some labs prefer to split the cells 1:3 or 1:4, although increasing the split ratio will result in a longer interval before subcultures reach confluency.

[bioadvisers]

Bioadvisers shared on Biotech Advisers

Basic Techniques for Mammalian Cell Tissue Culture

Tissue culture technology has found wide application in the field of cell biology. Cell cultures are utilized in cytogenetic, biochemical, and molecular laboratories for diagnostic as well as research studies. In most cases, cells or tissues must be grown in culture for days or weeks to obtain sufficient numbers of cells for analysis. Maintenance of cells in long-term culture requires strict adherence to aseptic technique to avoid contamination and potential loss of valuable cell lines.

An important factor influencing the growth of cells in culture is the choice of tissue culture medium. Many different recipes for tissue culture media are available and each laboratory must determine which medium best suits their needs. Individual laboratories may elect to use commercially prepared medium or prepare their own. Commercially available medium can be obtained as a sterile and ready-to-use liquid, in a concentrated liquid form, or in a powdered form. Besides providing nutrients for growing cells, medium is generally supplemented with antibiotics, fungicides, or both to inhibit contamination.

As cells reach confluency, they must be subcultured or passaged. Failure to subculture confluent cells results in reduced mitotic index and eventually cell death. The first step in subculturing monolayers is to detach cells from the surface of the primary culture vessel by trypsinization or mechanical means. The resultant cell suspension is then subdivided, or reseeded, into fresh cultures. Secondary cultures are checked for growth, fed periodically, and may be subsequently subcultured to produce tertiary cultures, etc. The time between passaging cells depends on the growth rate and varies with the cell line. The Basic Protocol describes subculturing of a monolayer culture grown in petri plates or flasks; the Alternate Protocol 1 describes passaging of suspension cultures.

CAUTION: When working with human blood, cells, or infectious agents, appropriate biosafety practices must be followed. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly.

Trypsinzing And Subculturing Cells From A Monolayer

A primary culture is grown to confluency in a 60mm petri plate or 25 cm2 tissue culture flask containing 5 ml tissue culture medium. Cells are dispersed by trypsin treatment and then reseeded into secondary cultures. The process of removing cells from the primary culture and transferring them to secondary cultures constitutes a passage, or subculture.

Materials Primary cultures of cells

PBS/HBSS without Ca2+ and Mg2+, 37°C

Trypsin/EDTA solution , 37°C

Complete medium with serum: e.g., supplemented DMEM with 10% to 15% (v/v) FBS, 37°C

Sterile Pasteur pipets

37°C warming tray or incubator

Tissue culture plasticware or glassware including pipets and 25-cm2 flasks or 60-mm petri plates, sterile

NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4.

1. Remove all medium from primary culture with a sterile Pasteur pipet. Wash adhering cell monolayer once or twice with a small volume of 37°C PBS/HBSS without Ca2+ and Mg2+ to remove any residual FBS that may inhibit the action of trypsin.

Use a buffered salt solution that is Ca2+ and Mg2+ free to wash cells. Ca2+ and Mg2+ in the salt solution can cause cells to stick together.

If this is the first medium change, rather than discarding medium that is removed from primary culture, put it into a fresh dish or flask. The medium contains unattached cells that may attach and grow, thereby providing a backup culture.

2. Add enough 37°C trypsin/EDTA solution to culture to cover adhering cell layer.

3. Place plate on a 37°C warming tray 1 to 2 min. Tap bottom of plate on the countertop to dislodge cells. Check culture with an inverted microscope to be sure that cells are rounded up and detached from the surface.

If cells are not sufficiently detached, return plate to warming tray for an additional minute or two.

4. Add 2 ml 37°C complete medium. Draw cell suspension into a Pasteur pipet and rinse cell layer two or three times to dissociate cells and to dislodge any remaining adherent cells. As soon as cells are detached, add serum or medium containing serum to inhibit further trypsin activity that might damage cells.

If cultures are to be split 1/3 or 1/4 rather than 1/2, add sufficient medium such that 1 ml of cell suspension can be transferred into each fresh culture vessel.

5. Add an equal volume of cell suspension to fresh plates or flasks that have been appropriately labeled.

Alternatively, cells can be counted using a hemacytometer or Coulter counter and diluted to the desired density so a specific number of cells can be added to each culture vessel. A final concentration of ∼5 × 104 cells/ml is appropriate for most subcultures.

For primary cultures and early subcultures, 60-mm petri plates or 25cm2 flasks are generally used; larger vessels (e.g., 150mm plates or 75cm2 flasks) may be used for later subcultures.

Cultures should be labeled with date of subculture and passage number.

6. Add 4 ml fresh medium to each new culture. Incubate in a humidified 37°C, 5% CO2 incubator.

If using 75cm2 culture flasks, add 9 ml medium per flask.

Some labs now use incubators with 5% CO2 and 4% O2. The low oxygen concentration is thought to simulate the in vivo environment of cells and to enhance cell growth.

For some media it is necessary to adjust the CO2 to a higher or lower level to maintain the pH at 7.4.

7. If necessary, feed subconfluent cultures after 3 or 4 days by removing old medium and adding fresh 37°C medium.

8. Passage secondary culture when it becomes confluent by repeating steps 1 to 7, and continue to passage as necessary.

Passaging Cells In Suspension Culture

A suspension culture is grown in culture flasks in a humidified 37°C, 5% CO2 incubator. Passaging of suspension cultures is somewhat less complicated than passaging of monolayer cultures. Because the cells are suspended in medium rather than attached to a surface, it is not necessary to disperse them enzymatically before passaging. However, before passaging, cells must be maintained in culture by feeding every 2 to 3 days until they reach confluency (i.e., until the cells clump together in the suspension and the medium appears turbid when the flask is swirled).

NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4.

1. Feed cells as follows every 2 to 3 days until the cultures are confluent:

a. Remove flask of suspension cells from incubator, taking care not to disturb those that have settled to the flask bottom.

b. Aseptically remove and discard about one-third of the medium from flask and replace with an equal volume of prewarmed (37°C) medium. If the cells are growing rapidly, add an additional 10% medium by volume in order to maintain optimum concentration of 1 × 106 cells/ml. Gently swirl flask to resuspend cells.

c. Return flask to incubator. If there is <15 ml of medium in the flask, incubate flask in horizontal position to enhance cell/medium contact.

At higher volumes of medium the flask can be incubated in the vertical position.

If using a 25-cm2 flask, there should be 20 to 30 ml of medium in the flask at confluency.

2. On the days cultures are not being fed, check them by swirling flask to resuspend cells and observing color changes in the medium that indicate good metabolic growth.

3. When cultures are confluent (∼2.5 × 106 cells/ml), passage culture as follows:

a. Remove flask from incubator and swirl flask so that cells are evenly distributed in the medium.

b. Aseptically remove half of the volume of cell suspension and place into a fresh flask.

c. Feed each flask with 7 to 10 ml prewarmed medium and return flask to incubator.

Some labs prefer to split the cells 1:3 or 1:4, although increasing the split ratio will result in a longer interval before subcultures reach confluency.