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Media

General information

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.

Rich media

LBS

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

SWT

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.

LB

Lysogeny Broth

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

BHI

Brain Heart Infusion

Amount Reagent
37 g Brain Heart Infusion powder
to 1000 ml total with dI H2O

TBS

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

Minimal Media

Tris minimal medium

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%)

General media recipes

DSW 2X

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.

Glucose (20% v/v)

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.

Glycerol (50% v/v)

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.

Instant Ocean (70% v/v)

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.

IPTG (100 mM)

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.

Xgal (20 mg/ml)

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.

DAP (300 mM, 1000x)

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.

Thymidine (100 mM, 333x)

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.

Making media

General information

  • 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
  • Clean the media preparation area when finished making media
    • Dust off the scale, make sure its doors are closed

Liquid media

  1. Make media and measure out in 100 mL aliquots into milk dilution bottles
  2. Put lids on bottles loosely (don't close all the way)
  3. Place autoclave tape on seam between lid and bottle
  4. Put bottles in square metal cage
  5. Autoclave
  6. 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
  7. Put media away on designated shelf

Plates

  1. Make media in appropriate sized flask with stir bar
  2. Add 15 g agar per 1 L of media (makes 1.5% agar plates)
  3. Cover top of flask with aluminum foil and place autoclave tape on top
  4. Turn on 55°C water bath
  5. Autoclave
  6. After autoclaving, immediately place media in 55°C water bath and wait until cool enough to touch with bare hands (~30 min)
  7. 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
  8. 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
  9. Light Bunsen burner
  10. 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
  11. 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
  12. 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)
  13. Leave plates out at room temperature for 1-2 days to solidify and dry out along with a piece of tape labeled with the media type, date, and your initials
  14. After the plates are solidified, place them in a labeled plastic sleeve and place in the cold room on the appropriate shelf

Antibiotics

  • 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 ug/mL 25 mg/uL 1 mL
Chloramphenicol (Cam) 5 ug/mL 25 mg/uL 200 uL
Erythromycin (Erm) 150 ug/mL N/A 0.150 g of powder
Erythromycin (Erm) 5 ug/mL 10 mg/uL 500 ul
Kanamycin (Kan) 50 ug/mL 50 mg/uL 1 mL
Kanamycin (Kan) 100 ug/mL 50 mg/uL 2 mL
Spectinomycin (Spc) 50 ug/mL 50 mg/uL 1 mL
Tetracycline (Tet) 15 ug/mL 10 mg/uL 1.5 mL
Tetracycline (Tet) 5 ug/mL 10 mg/uL 500 uL
Carbenicillin (Carb) 100 ug/mL 100 mg/uL 1 mL