microbiological media preparation

GENERAL MICROBIOLOGICAL TECHNIQUES
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.GENERAL MICROBIOLOGICAL TECHNIQUES
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.GENERAL MICROBIOLOGICAL TECHNIQUES
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.GENERAL MICROBIOLOGICAL TECHNIQUES
PREPARATION OF BACTERIAL MEDIA
Nearly all procedures in molecular biology that involve using bacteria to clone DNA are done
with the assistance of specific derivatives of Eschericia coli (E. Coli) strain K- 12. E. Coli is a
rod shaped gram-negative bacterium with a circular chromosome of ca. 3 million base pairs. It
also harbors plasmids, a characteristic of great importance to molecular biologists. It can readily
be grown on minimal media that contain a carbon source (glucose) and salts that provide
nitrogen, phosphorus and trace metals. However, its growth is much more rapid on enriched
media which provide cells with amino acids. In such rich medium, a culture during exponential
growth will double every 20-30 minutes at 37oC. This growth rate will continue until the culture
is "saturated" which occurs in most strains of E. coli at a cell density of 1-2 x 109 cells.
The most commonly used medium is known as LB medium (also referred to as Luria-Bertaini
broth). This medium contains tryptone, yeast extract and NaCl. For most applications during this
course this is the medium that will be utilized.
Equipment
1L Erlenmeyer flask
Balance
Weigh boats or paper
Autoclave
Sterile 100 mm Petri dishes
37oC incubator
55oC water bath
Magnetic stirrer and bars
pH paper
Reagents and Materials
Bacto-tryptone
Yeast extract
NaCl
Bacto-Agar
ddH2O
0.1M IPTG (filter sterilized)
2% X-gal in dimethylformamide
100 mg/ml Ampicillin (filter sterilized)
1 M NaOH
Preparation of LB liquid medium:
1. In a 1 L flask in which you have calibrated the 500 ml mark add 400 ml of ddH2O and the
following:
5 g Bacto tryptone
2.5g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0 (use pH
paper, not a pH electrode as the tryptone is not good for the electrode). Bring up the volume
to 500 ml using MQ water. Note: The medium is not very highly buffered and its pH drops
significantly as the culture nears saturation.. Remove the stir bar.
3. Make cotton/cheese cloth plugs to plug the flasks and cover the plugged tops with foil. Put a
label (indicating the contents, group's identity and date made) near the flask neck. Also put a
piece of autoclave tape on each flask.
4. Autoclave for 20 minutes. BE SURE TO PUT ALL ITEMS IN AN AUTOCLAVABLE
TRAY INSIDE THE AUTOCLAVE AND SET THE AUTOCLAVE FOR SLOW
"LIQUID" EXHAUST.
The sterile medium can be stored where designated and will be used during the course. As
more is needed, it will be up to your group to replenish the supply. For most procedures,
ampicillin will be added to the LB medium prior to use. As ampicillin is not stable when
stored at room temperature, a sterile stock solution (which will be stored frozen) will be
added to the sterile LB medium just before it is to be used.
Preparation of LB/amp-100 agar plates:
1. In a 1 L flask in which you have calibrated the 500 ml mark. add 400 ml ddH2O and the
following:
5 g Bacto tryptone
2.5 g yeast extract
5 g NaCl
2. Add a stir bar and stir until in solution. Adjust pH with 1 M NaOH to a pH near 7.0.
3. Add 7.5 g of agar (not agarose!) Bring up the volume to 500 ml using ddH2O.
4. Plug flask with cotton/cheesecloth plug and foil and label (indicating the contents, date made
and by which group). Put the flask in an autoclave pan and AUTOCLAVE ON SLOW
"LIQUID" EXHAUST for 20 minutes.
5. After autoclaving, cool the flask to ca. 50-55oC by letting the flask sit in a 55oC water bath.
Do not check the temperature with a thermometer, as you will contaminate the medium. A
good rule of thumb for estimating the temperature of your media is “if you can hold the flask
it is cool enough”.
Note: it is critical that you cool the agar medium before pouring the plates for two reasons:
(1) the ampicillin that you will add is heat labile and will be rapidly degraded if added to hot
medium; (2) if plates are poured from medium that is too hot tremendous condensation will
form in the plates as they cool. Such "wet-plates" are a no-no as it is impossible to culture
discrete colonies in a film of water.
When the medium reaches this temperature you must move quickly or it will solidify. If this
happens, do not open the flask. Just reheat in the microwave until the agar is melted, and
then recool.
6. For each 1 liter of medium, aseptically add 1 ml of a 100 mg/ml stock solution of sterile
ampicillin. This should result in a final concentration of 100 μg/ml ampicillin in the LB
plates.
Note: This concentration of ampicillin is at the high end of the range for use. Many protocols
call for 30-50 μg/ml final concentration. We have found that a 100 μg/ml concentration will
permits the growth of all strains of bacteria harboring ampicillin resistant plasmids. In
addition, the plates, if stored at 4oC, can be used over a 2-3 week period before the
ampicillin is degraded below restrictive levels.
7. If you are making plates for a blue/white screen using the expression of the LacZ gene, you
will need to add 2 ml of a 20 mg/ml X-gal solution and 2 ml of a 0.1M IPTG solution to the
cooled media.
8. Swirl the flask to thoroughly mix the ampicillin, X-gal and IPTG and pour into ca. 24-28
sterile, plastic Petri dishes. Pour each dish to a level of 1/3-1/2 full (ca. 20-25 ml) and before
solidified, flame the top of each surface with a Bunsen flame to eliminate any bubbles.
Note: Generally rather thin plates (i.e., 1/4 - 1/3 full) give better color reactions (but more
satellite colonies) when used to select transformed bacteria. We maintain stocks of colonies
on somewhat thicker (i.e.. 1/2 full) plates.
Cover the dishes and let stand at room temperature for several hours (or overnight) before
putting back in the plastic sleeve that had contained the Petri dishes. Label the sleeve of agar
dishes with the contents, date and group name and put in designated area of cold room. Do
not store the plates on their sides or right side up. Rather, store the plates upside-down.
OBTAINING SINGLE COLONIES of BACTERIA
The object of streaking of a bacterial culture is to obtain a single colony on a plate that represents
a clone of a single bacterium. This is important as it ensures that each cell in the population is
descended from a single cell, and thus, all have the same genetic make up. During cultured
growth, transformed bacterial cells may lose their plasmids or parts of plasmids. It is, therefore,
critical that you know that all cells in your culture all descended from one ancestral cell and that
all harbor the correct plasmid.
To obtain single colonies, a bacterial inoculum is streaked across one region of an agar plate (see
following diagrams). The loop is then resterilized (or a new sterile toothpick is obtained) and is
then passed once through the first streak and streaked across a fresh region of the plate. This
procedure is repeated one or more times after which the plate is incubated, upside down, at 37oC,
overnight.
Alternatively, a log culture can be diluted (serial dilution) and an aliquot of the diluted bacteria is
then spread with a sterile glass rod onto the agar.
Equipment
Wire bacterial loop
Bunsen burner
37oC incubator
Reagents and materials
LB agar plates
Bacterial colony or culture
Streaking plates to obtain single colonies:
1. Take a sterile bacterial transfer loop and streak each of the plates with the stock cultures
(undiluted log-phase cultures). Be sure to do this as demonstrated in the following diagram,
as the objective in this is to obtain single cultures in the final pass of the loop across the
plates.
2. Label these plates on the side and place them UPSIDE-DOWN in the 37oC incubator
overnight (ca. 12-16 hr). Always be sure to check that the incubator is at 37oC and minimize
the time the door is opened.
3. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
Spreading diluted stock cultures to obtain single colonies:
Equipment
Spreader (“hockey stick”)
Turntable
Bunsen Burner
Sterile culture tubes
Reagents and materials
Bacterial culture
Sterile LB liquid media or ddH2O
1. You will need to dilute each of your bacterial suspensions before plating out on a plate. If
your suspensions have grown into post-log phase, you should have ca. 109 cells per ml. If you
spread 100 μl per plate, you could expect to have ca. 108 colonies developing on your plate.
What you would see if you did this would be a "lawn" of cells on your plate. You would not
be able to delineate single colonies. For each of your cultures, dilute a subaliquot of your
culture so that you have no more than 103 cells per ml. Be very careful to keep your labeled
samples straight and do not mix up your cultures.
You should be able to calculate this. Your calculations should reveal that you should do a
million - fold dilution (10-6) dilution. To do this, take 1 μl of your agitated cell suspension
and dilute it in 1000 μl (1 ml) LB medium. This should give you a dilution of 10-3. Take 1 μl
of your 10-3 dilution and add it to another 1 ml of LB medium. This should effectively give
you a cell concentration of 103 cells/ml.
2. For each of your cultures you will spread 100 μl. To spread use a bent glass or metal rod
(“hockey stick”) and a turntable as demonstrated by the instructors.
1. Incubate the plates upside down at 37oC overnight (ca. 16- 20h).
Note: Incubating ampicillin-containing plates for periods longer than 16 h (or incubating on
very thin ampicillin plates) results in the appearance of "satellite" colonies. These colonies
will appear as much smaller, white colonies forming a halo around the larger plasmidcontaining
colonies. The "satellites" do not contain plasmids but are able to grow because as
the culture ages, the ampicillin in the medium decreases. In addition, colonies with plasmids
produce compounds that actively degrade ampicillin thus conferring their "resistance” to
this drug. In the environs directly surrounding ampicillin-degrading colonies, colonies
lacking plasmids may grow.
2. Remove your plates from the 37oC incubator. Wrap the plate edges in Parafilm and store at
4oC. Count and record the number of colonies you get on each of your plates. Compare this
with the number of colonies you expect to get from your dilution calculations. Record all
your results carefully in your notebook.
6
Figure 2. Plating Bacterial Cultures onto LB Agar Plates.