For plates, add 15 g Agar and a stir bar. Cool in the 55 °C bath prior to pouring, until the media can be touched with bare hands. Add antibiotics if required. Stir for approximately 1 min, then pour.
Additional detail on making media is listed below.
LB Salt
| Amount | Reagent |
|---|---|
| 25 g | LB Broth powder |
| 10 g | NaCl |
| 50 ml | Tris (1 M, pH 7.5) |
| 950 ml | dI H2O |
Note: Above is equivalent to the following (i.e. Tris-buffered LB with double salt)
| Amount | Reagent |
|---|---|
| 10 g | Bacto-tryptone |
| 5 g | Yeast extract |
| 20 g | NaCl |
| 50 ml | Tris (1 M, pH 7.5) |
| to 1000 ml total | with dI H2O |
Seawater tryptone
| Amount | Reagent |
|---|---|
| 5 g | Bacto-Tryptone |
| 3 g | Yeast Extract |
| 6 ml | 50% Glycerol |
| 700 ml | Instant Ocean [1] |
| to 1000 ml total | with dI H2O |
[1] Instant Ocean at 35 ppt. Dispense from an "E" tripour into a laboratory graduated cylinder or beaker. Ensure that media glassware does not touch squid "E" areas or Instant Ocean source barrels.
| Amount | Reagent |
|---|---|
| 25 g | LB Broth powder |
| 1000 ml | dI H2O |
Note: Above is equivalent to:
| Amount | Reagent |
|---|---|
| 10 g | Bacto-tryptone |
| 5 g | Yeast extract |
| 10 g | NaCl |
| to 1000 ml total | with dI H2O |
Brain Heart Infusion
| Amount | Reagent |
|---|---|
| 37 g | Brain Heart Infusion powder |
| to 1000 ml total | with dI H2O |
Tryptone broth
| Amount | Reagent |
|---|---|
| 10 g | Bacto-tryptone |
| 20 g | NaCl |
| 50 ml | Tris (1 M, pH 7.5) |
| to 1000 ml total | with dI H2O |
Mix the following:
| Amount | Reagent |
|---|---|
| 50 ml | Tris base, pH 7.5 (1M) |
| 500 ml | DSW (2X) |
| 450 ml | dI H2O |
Filter, then add each filter-sterilized solutions:
| Amount | Reagent |
|---|---|
| 1 ml | K2HPO4 (5.8%) |
| 1 ml | FeSO4 (10 mM) |
| . | . |
| to 0.2% (w/v) final | N-acetyl-glucosamine as N+C-source |
| . | or |
| to 0.2% (w/v) final | Carbon Source |
| 5 ml | NH4Cl (20%) |
Defined seawater 2X
Note: This is defined sea water for minimal media and not for animal incubation (for the latter use Instant Ocean). Final concentrations:
- 100 mM MgSO4
- 20 mM CaCl2
- 600 mM NaCl
- 20 mM KCl
In a 2 L beaker add 1 L dI H2O and set aside.
In three smaller beakers dissolve each of the constituent solutions as described. Ensure they are completely dissolved, then add to the beaker above.
| Beaker | Reagent(s) | Dissolve completely in |
|---|---|---|
| 1 | 49.5 g MgSO4 • 7 H2O | 250 ml dI H2O |
| 2 | 5.75 g CaCl2 • 2 H2O | 250 ml dI H2O |
| 3 | 70.25 g NaCl + 3 g KCl | 250 ml dI H2O |
Adjust total volume to 2 L with dI H2O. Filter sterilize and store at room temperature. Lasts 1-2 months.
In a 1 liter graduated cylinder mix:
| Amount | Reagent |
|---|---|
| 200 g | glucose |
| 800 ml | dI H2O |
Cover with parafilm and invert multiple times to mix. Adjust final volume to 1 L.
Dispense 100 ml aliquots into milk dilution bottles.
Autoclave, store at room temperature.
In a 1 liter graduated cylinder mix:
| Amount | Reagent |
|---|---|
| 500 ml | glycerol |
| 500 ml | dI H2O |
Cover with parafilm and invert multiple times to mix. Glycerol is very viscous, so make sure it mixes completely and does not stick to the container. Dispense 100 ml aliquots into milk dilution bottles and autoclave.
In a 1 liter graduated cylinder mix:
| Amount | Reagent |
|---|---|
| 300 ml | dI H2O |
| 700 ml | Dissolved Instant Ocean at approximately 33-38 ‰ |
Cover with parafilm and invert multiple times to mix. Dispense 100 ml aliquots into milk dilution bottles and autoclave.
Note: Solution is not checked for salinity. The purpose of the 70% solution is to provide a growth-unsupportive medium for dilution plating. Solution is inappropriate for animal work.
isopropyl-β-D-thiogalactopyranoside
| Amount | Reagent |
|---|---|
| 238 mg | IPTG |
| 10 ml | ddH2O |
Filter sterilize, then store 0.5 ml aliquots at -20 °C.
When adding to autoclaved media prior to pouring, add 1 μl IPTG solution per 1 ml agar for a final concentration of 0.1 mM.
If spreading on plate, consider that a typical plate agar volume is 25 ml. Dispense 75 μl sterile water on the plate containing 10 beads, add 25 μl IPTG to the plate, then shake to distribute evenly.
5-bromo-4-chloro-3-indolyl-β-D-galactoside
| Amount | Reagent |
|---|---|
| 200 mg | Xgal |
| 10 ml | N,N-dimethylformamide (Dispense in hood) |
Mix in Corning polypropylene 15 mL tube. Wrap tube in foil as Xgal is light-sensitive
Note that dimethylformamide dissolves some plastics (e.g., polystyrene conical tubes).
Mix well and store at -20°C.
When adding to autoclaved media prior to pouring, add 5 mL to 1 L of agar.
Add 100 μl to agar plates for β-galactosidase assays.
The most uniform distribution in an agar plate is accomplished by adding Xgal to the liquid agar after the media has been autoclaved and can be touched (~60 °C), then mix well prior to plating. Alternately, if Xgal is spread on a plate directly, pipet onto multiple foci and use 10 glass beads to distribute it rapidly before it sets into local areas of the plate.
Diaminopimelic acid
MW = 190.19
| Amount | Reagent |
|---|---|
| 1.14g | DAP |
| 20 ml | ddH2O |
At this concentration, the DAP will not go into solution, so aliquot quickly after mixing. Autoclave, then store at room temp.
Vortex before adding to media.
Deoxythymidine
MW = 242.231
| Amount | Reagent |
|---|---|
| 242 mg | Thymidine |
| 10 ml | ddH2O |
Filter sterilize, then store 1.0 ml aliquots at -20 °C.
When adding to autoclaved media prior to pouring, add 3 μl Thymidine solution per 1 ml agar for a final concentration of 0.3 mM.
- When pouring plates, ignite a Bunsen Burner near you to decontaminate the air around you.
- Make sure the flame IS NOT UNDER THE WOOD SHELF!
- Once ignited, wrap a paper towel under the neck of the flask and pour slowly to avoid adding bubbles.
- Make sure the paper towel does not touch the media being poured! This would contaminate all of the plates, as our paper towel is not sterile.
- If poured correctly, you should yield about two sleeves of plates per liter of media.
- A good rule of thumb to follow to ensure the proper amount of media is in each plate is to pour slowly until the bottom is completely filled. AS SOON as the bottom is filled, stop pouring.
- Pouring the correct amount of media takes time and practice. If your plates are too thin or too thick, ask a senior labmate for tips and tricks. Everyone has different methods that work for them, and they may have some good advice!
- Once done pouring, rinse the flasks with HOT tap water, and then DI water.
- Cold tap water can cause the agar to harden, sticking to the glass. This is especially bad if an antibiotic media was in this flask. Place on the drying rack above the sink when done.
- Flasks used with antibiotic media should be washed before reuse.
- Let plates sit on the benchtop for two days.
- After two days, place in plate sleeve and close with label tape with media type, date made, and initials.
- Place it in the cold room on appropriate shelf against the back wall.
- Wear gloves while making media.
- When making media to be autoclaved, make sure the volume of the container is twice that of the amount of media you are autoclaving (i.e. if you are making 1 L of media, make it in a 2 L flask to autoclave it).
- Too small of a container will result in boil-over.
- Use the largest stirbar that fits the vessel. Autoclave the stirbar with the media.
- Always use weigh boats/weigh paper, which is stored in the drawer under the scales.
- If you accidentally take too much powder, do not put the extra back into the source bottle as this can contaminate the source.
- Clean the media preparation area when finished making media.
- Dust off the scale, ensure any scale doors are closed.
The specific steps you take will depend on whether you are preparing liquid media or agar plates:
- For liquid media bottles, aliquot the non-sterile liquid into
milk dilution bottlesand then autoclave the individual bottles. - For agar plates, autoclave the large flask(s), allow them to stir/cool, add antibiotics if needed, and then pour the plates.
- Make media and measure out in 100 mL aliquots into
milk dilution bottles - Put lids on bottles loosely (don't close all the way)
- Place autoclave tape on seam between lid and bottle
- Put bottles in square metal cage
- Autoclave
- After autoclaving, let bottles cool at room temperature until you can touch them with bare hands, then tighten the caps
- Important to tighten caps after cooled, or else they will get stuck on
- Put media away on designated shelf
- Make media in appropriate sized flask with stir bar
- Add 15 g agar per 1 L of media (makes 1.5% agar plates)
- Cover top of flask with aluminum foil and place autoclave tape on top
- Cover flasks/beakers to be autoclaved with aluminum foil, then with a small piece of autoclave tape.
- Label the foil with the specific media type and your initials.
- Please keep the tape on the foil, not on the flask unless necessary, as it makes the glass gooey.
- Turn on 55°C water bath
- Autoclave
- After autoclaving, immediately place media in 55°C water bath and wait until cool enough to touch with bare hands (~30 min)
- When ready to pour plates, place flask on stir plate and stir at low speed (~200 rpm) so bubbles aren't produced
- If making plates with antibiotics, add antibiotic to flask at this point and let stir for ~1 min
- Set up plates for pouring; keep the plastic sleeve the plates come in to store the poured plates
- Use scissors to open sleeve of plates closest to the top seam (to get the most length out of the sleeve for reuse)
- Stack plates in sets of 5 and line up on bench top
- Light Bunsen burner
- Take flask off of stir plate and pour media into plates
- Pour just enough to cover the bottom of the plate
- Pour gently so as not to create bubbles
- Make sure to put lid back plate on immediately after pouring
- If bubbles end up in plate, pass flame from Bunsen burner over surface of media
- Typically 1 L of media yields 2 sleeves of plates (i.e., 45-50 plates).
- When all the media has been poured, immediately rinse out flask with hot water (so the residual media doesn't solidify), turn off the water bath, turn gas off of Bunsen burner
- Label the sides of the plates with permanent marker according to the guide at the plate pouring bench (i.e. LB is one black line)
- Leave plates out at room temperature for 2 days to solidify and dry out along with a piece of tape labeled with the media type, date, and your initials
- On the day after the plates are poured, validate one plate from each stock with the test strains. Use a plate from the middle of the stack (i.e., not the wettest plate on the top of the stack).
- After the plates are solidified and passed validation, place them in a labeled plastic sleeve and place in the cold room on the appropriate shelf.
- Antibiotic stocks are kept in the -20°C freezer in a labeled box
- The numbers on the tubes of antibiotics refer to the concentration of antibiotic in mg/mL
- If you need a concentration of antibiotic that is not indicated on a tube, then you'll have to do some math (i.e. to make LBS-Kan100, add 2 tubes of Kan50 to 1 L of LBS); see table below
- Antibiotics should be added to media AFTER autoclaving and after the media has cooled to a usable temperature (or else the antibiotics could degrade)
- Fully thaw antibiotics and vortex just before adding them to media
- Allow the media to stir for ~1 min after adding antibiotics
| Antibiotic | Final concentration in 1 L media | Stock concentration | Volume of stock to add to 1L media |
|---|---|---|---|
| Chloramphenicol (Cam) | 25 μg/ml | 25 mg/μl | 1 ml |
| Chloramphenicol (Cam) | 5 μg/ml | 25 mg/μl | 200 ul |
| Erythromycin (Erm) | 150 μg/ml | N/A | 0.150 g of powder |
| Erythromycin (Erm) | 5 μg/ml | 10 mg/μl | 500 ul |
| Kanamycin (Kan) | 50 μg/ml | 50 mg/μl | 1 ml |
| Kanamycin (Kan) | 100 μg/ml | 50 mg/μl | 2 ml |
| Spectinomycin (Spc) | 50 μg/ml | 50 mg/μl | 1 ml |
| Tetracycline (Tet) | 15 μg/ml | 10 mg/μl | 1.5 ml |
| Tetracycline (Tet) | 5 μg/ml | 10 mg/μl | 500 μl |
| Carbenicillin (Carb) | 100 μg/ml | 100 mg/μl | 1 ml |