Media And Plates

Pouring Plates

Prepare and autoclave media containing agar.
- If cooling on a plate, add stir bar before autoclaving.
Immediately after autoclaving, cool for pouring.
- move molten agar to 55°C water bath.
- Cool to stir on plate.
Once cooled to 55°C, add any heat-labile components (vitamins, antibiotics) aseptically, then mix
- Avoid heavy stirring, which can cause bubbles to form
Under flame, pour agar from container into sterile petri dishes
- Only use enough liquid agar to cover the bottom of the dish
- Remove bubbles by briefly flaming them (rapidly pass flame of Bunsen burner over the top of still-liquid agar with bubbles; take care not to melt plastic or burn yourself)
Allow to solidify at room temperature, then invert and store in a labeled plastic sleeve at 4°C

Last modified: 5 August 2021; TBJ

LB Broth/LB Plates

Notes:

1 L of LB Agar can make ≥ 50 plates.

Chemical	Amount
Tryptone	10 g
Yeast Extract	5 g
NaCl	5 g*
Agar (optional)	15 g
dH₂O	Add to 1 L

Autoclave

*Some labs and published recipes use 10g/L NaCl

Last modified: 14 July 2021; TBJ

Last modified: 08/03/2021: DK

M9 Stock Solutions – version 1 from the lab website

M9 10X Base

Chemical	Amount
Na₂HPO₄anhydrous	60 g
KH₂PO₄anhydrous	30 g
NaCl	5 g
NH₄Cl	10 g
dH₂O	Add to 1 L

Dissolve in the order indicated. Dispense into bottles. Autoclave.

0.1 M MgSO₄

Chemical	Amount
MgSO₄anhydrous or MgSO₄· 7 H₂O	12.04 g / 24.65 g
dH₂O	Add to 1 L

Dissolve. Dispense into bottles. Autoclave.

0.01 M CaCl₂

Chemical	Amount
CaCl₂ · 2 H₂O	1.47 g
dH₂O	Add to 1 L

Dissolve. Dispense into bottles. Autoclave.

0.72 mM FeSO₄

Chemical	Amount
FeSO₄ · 7 H₂O	0.2 g
dH₂O	Add to 1 L

Dissolve. Dispense into bottles. Acidify with a drop or two of HCl. Titrate to pH 3.0 to 4.0. Autoclave. A precipitate WILL form upon autoclaving. This is normal.**

20% Glucose

Chemical	Amount
Glucose	200 g
dH₂O	Add to 1 L

Dissolve. Dispense into bottles. Autoclave. Do not OVER-autoclave!

M9 Media (using version 1 M9 stock solutions)

Notes:

Shake/swirl/stir well after metal additions to re-dissolve any precipitate.

Chemical	Amount
M9 10X Base	100 mL
20% Glucose	20 mL (final 0.4%)
0.1M MgSO₄	10 mL
0.01M CaCl₂	10 mL
0.72mM FeSO₄	10 mL
dH₂O	850 L

Mix all components aseptically into sterile vessel(s) or combine and filter sterilize.

M9 Stock Solutions – version 2 used by Zizi, Tyler, & Melanie

M9 10X Base (normal phosphate*, 0.643M PO₄)

Na₂HPO₄	60g
KH₂PO₄	30g
NaCl	5g
NH₄Cl	10g
ddH₂O to 1L

Dissolve in order indicated. Autoclave.

*Insufficient buffering capacity for WT anaerobic E. coli growth – media acidifies and inhibits growth

M9 10X Base (low phosphate**, 0.129M PO₄)

Na₂HPO₄	12g
KH₂PO₄	6g
NaCl	5g
NH₄Cl	10g
ddH₂O to 1L

Dissolve in order indicated. Autoclave.

**Does not support anaerobic E. coli growth

1M MgSO₄ solution

MgSO₄•7H₂O	24.65g
ddH₂O to 100mL

Dissolve. Autoclave.

1M CaCl₂ solution

CaCl₂•2H₂O	14.7g
ddH₂O to 100mL

Dissolve. Autoclave.

50mM FeCl₃ solution

FeCl₃•4H₂O	0.99g
ddH₂O to 100mL

Dissolve. Filter sterilize. Store at 4°C. Solution should be very yellow and should not precipitate.

20% glucose solution

Glucose (dextrose)	20g
ddH₂O to 100mL

Dissolve. Autoclave. Remove from autoclave immediately upon completion.

M9 Media (using version 2 M9 stock solutions)

Notes:

Reliably stores on benchtop 1-2+ weeks; check before use as media will precipitate after a time.
Shake/swirl/stir well after metal additions to re-dissolve any precipitate.

Chemical	Amount
M9 10X Base	100 mL
20% Glucose	20 mL (final 0.4%)
1M MgSO₄	400 µL
1M CaCl₂	20 µL
50mM FeCl₃	300 µL
dH₂O	876.4 L

Mix all components aseptically into sterile vessel(s) or combine and filter sterilize.

MSgg Stock Solutions

100 mM KH₂PO₄

Chemical	Amount
KH₂PO₄anhydrous	13.61 g
dH₂O	Add to 1 L

Dissolve, pH to 7.0, and autoclave.

0.5 M MOPS

Chemical	Amount
MOPS	104.63 g
dH₂O	Add to 1 L

Dissolve, pH to 6.5, and autoclave.

1 M MgCl₂

Chemical	Amount
MgCl₂	95.21 g
dH₂O	Add to 1 L

Dissolve and autoclave.

0.1 M CaCl₂

Chemical	Amount
CaCl₂ · 2 H₂O	14.70 g
dH₂O	Add to 1 L

Dissolve and autoclave.

50 mM MnCl₂

Chemical	Amount
MnCl₂	6.29 g
dH₂O	Add to 1 L

Dissolve and autoclave. Store at 4⁰C.

50 mM FeCl₃

Chemical	Amount
FeCl₃	8.11 g
dH₂O	Add to 1 L

Acidify with HCl to solubilize, filter sterilize, wrap in foil, and store in 4⁰C.

50 mM ZnCl₂

Chemical	Amount
ZnCl₂	6.81 g
dH₂O	Add to 1 L

Dissolve and autoclave. Store at 4⁰C. Note: this will likely not dissolve without adding HCl. Simply continue adding HCL until the solution is mostly clear (doesn’t have to be perfect). Autoclaving should bring all precipitate into solution.

2 mM Thiamine-HCl

Chemical	Amount
Thiamine-HCl	0.675 g
dH₂O	Add to 1 L

Dissolve, filter sterilize, and store in 4⁰C.**

50% (v/v) Glycerol

Chemical	Amount
Glycerol (100% solution)	500 mL
dH₂O	500 mL

Mix and autoclave.

10% (w/v) Glutamate

Chemical	Amount
Glutamate (potassium glutamate)	100 g (137 g)
dH₂O	1 L

Dissolve and autoclave.

MSgg Media

Chemical	Amount
100mM KH₂PO₄	50 mL
0.5M MOPS	10 mL
1M MgCl₂	2 mL
0.1M CaCl₂	7 mL
50mM MnCl₂	1 mL
50mM FeCl₃	1 mL
50mM ZnCl₂	20 μL
2mM Thiamine-HCl	1 mL
50% (v/v) Glycerol	40 mL
10% (w/v) Glutamate	100 mL
dH₂O	788 mL

Autoclave water.
Mix all components aseptically into sterile vessel(s).
pH to 6.8.

Last modified by LNL 08/09/21

10X MC recipe (B. subtilis transformation medium)

Compound	For 100 ml MC	For 200 ml MC	For 500 ml MC
K₂HPO₄ (potassium phosphate dibasic)	10.7 g	21.4 g	53.6 g
KH₂PO₄ (potassium phosphate monobasic)	5.2 g	10.5 g	26.2 g
Glucose (dextrose)	20 g	40 g	100 g
Na₃C₆H₅O₇ ∘ 2H₂O (trisodium citrate dihydrate)	0.88 g	1.8 g	4.4 g
1000X Ferric Ammonium Citrate (2.2% stock)*	1 ml	2 ml	5 ml
Casein Hydrolysate (Oxoid)	1 g	2 g	5 g
Potassium Glutamate monohydrate	2.2 g	4.4 g	11 g
ddH₂O	100 ml	200 ml	500 ml

Mix all components with ~half the final volume of water
Once all components are dissolved, adjust to appropriate final volume
Filter sterilize using screw cap filter
Store 10X solution in freezer aliquots. Thaw before using in transformation.

* For 100 ml of 1000X Ferric Ammonium Citrate (2.2% stock)

Ferric Ammonium Citrate 2.2 g
ddH₂O to 100 ml

Last modified: 5 August 2021; TBJ

Last modified: 9 August 2021; LNL

10x SPIZ (Spizizen’s minimal salts)

Reagent	For 1L	For 2L
(NH₄)₂SO₄	20g	40g
K₂HPO₄anhydrous	140g	280g
KH₂PO₄	60g	120g
Sodium citrate dihydrate) (Na₃Citrate ● 2H₂O)	10g	20g
MgSO₄ ● 7H₂O)	2g	4g
ddH₂O	Dilute to volume

10X S750

Reagent	For 1L	For 2L
MOPS (free acid)	104.7g	209.4g
(NH₄)₂SO₄	13.2g	26.4g
KH₂PO₄	6.8g	13.6g
ddH₂O	Dilute to volume

Adjust pH to 7.0 using KOH

Zymomonas mobilis Rich Media (ZRMG)/ZRMG Plates

Notes

1 L of ZRMG Agar can make ≥ 50 plates

Chemical	Amount
Yeast Extract	10 g
KH₂PO₄	2 g
Glucose	20 g
Agar (optional)	20 g
dH₂O	1 L

Autoclave. Place in 55⁰C water bath to cool before pouring plates.

Zymomonas mobilis Minimal Media (ZMM) 10X Base

Chemical	Amount
KH₂PO₄	10 g
K₂HPO₄	10 g
NaCl	5 g
(NH₄)₂SO₄	10 g
dH₂O	1 L

Autoclave

Zymomonas mobilis Minimal Media (ZMM) version 3

Notes

Make all stock solutions separately and autoclave all except the pantothenic acid; the pantothenic acid can be filter sterilized instead.
The pH of this media should be between 6-6.4.
Lower glucose concentrations lower growth rate, but do not significantly alter final OD.
This media does not need to be filter sterilized if made aseptically; however, filter sterilization can be performed ~1 hour after preparation to prevent formation of precipitants.
Version 2 of this media uses 25g/L FeSO₄ 7 H₂O and 10g/L CaCl₂ · 2 H₂O stock solutions

Chemical	Amount
ZMM 10X Base	100 mL
20g/100mL (20%) glucose solution	100 mL (final 2%)
20g/100mL MgSO₄ · 6 H₂O	1 mL
25g/L NaMoO₄ · 2 H₂O	1 mL
2.5g/L FeSO₄ · 7 H₂O	1 mL
20g/L CaCl₂ · 2 H₂O	1 mL
1mg/mL Calcium Pantothenate	1 mL
dH₂O	795 mL

Last modified: 20 July 2021; TBJ

Zymomonas minimal media for Hungate (ZMH) media v1

Tyler Jacobson

Solution A (zA) (Same as ZMMG base)

KH₂PO₄	10g
K₂HPO₄	10g
NaCl	5g
(NH₄)₂SO₄	10g
ddH₂O	1L

pH should be 6.0 to 6.4

Solution B (zB)

MgSO₄ • 6H₂O	2g
Na₂MoO₄• 2H₂O	0.25g
ddH₂O	200mL

Solution C (zC)

FeSO₄• 7H₂O	0.25g
ddH₂O	200mL

Solution D (zD)

CaCl₂• 2H₂O	0.2g
ddH₂O	200mL

Solution E (zE)

Calcium Pantothenate hydrate	10mg
ddH₂O	200mL

Easier to make by diluting Pantothenate solution from ZMMG media rather than directly making

Filter sterilize rather than autoclave

Carbon source (zF)

200g/L glucose (or other carbon source)

Preparing Media (assuming 10mL final volume in hungate tube)

Dispense 1mL of zA into hungate tube and add 7mL of ddH₂O
- can also add things like isobutanol at this step if needed, leaving a final volume of 8mL in the tube
- if you need to add additional post-autoclave components to the media, leave out water to make space
Prepare all other solutions (except zE) separately in pressure vials
Seal hungate tubes and pressure vials and make anaerobic using vacuum manifold
Autoclave to sterilize sealed tubes
Aseptically add 1mL of zF (carbon source solution)
- Use 2.5% Cys-HCl solution to remove oxygen from syringe first
Aseptically add 200uL of zB, zC, zD, zE
- Use 2.5% Cys-HCl solution to remove oxygen from syringe first

Last modified: 14 July 2021; TBJ

Detection/Isolation Enrichment Media (DEM)/DEM Plates for Zymomonas mobilis

Notes:

After autoclaving, you can add 3% (v/v) ethanol and 20 µg/mL cycloheximide(actidione) as antifungals, which should be prepared in ethanol or DMSO as a 20 mg/mL (1000x) stock solution and stored at -20C.

Chemical	Amount
Malt Extract	3 g
Yeast Extract	3 g
Glucose	20 g
Peptone	5 g
Agar (optional)	17.5 g for plates; 7 g for soft agar; 2.5g for phage overlays
dH₂O	Add to 1 L

pH media to 4.0 with HCl. Autoclave. Place in 55⁰C water bath to cool before pouring plates.

Last modified: 14 July 2021; TBJ

MTC media (for Clostridium thermocellum and Thermoanaerobacterium saccharolyticum)

Assembling MTC media

Start with SolA in tube or bottle (SolA volume should be 90% of desired final media volume; ie 9mL solA for 10mL MTC media final)
Attach a new, sterile syringe to a new, sterile needle aseptically
Make syringe anaerobic by drawing up headspace gas from sterile, anaerobic bottle of 2.5% Cys-HCl
Use syringe to draw up 0.2mL SolB per 5mL final volume MTC, then add solB (to tube containing solA)
Repeat steps 2 and 3 with a new syringe
Use syringe to draw up 0.1mL SolC per 5mL final volume MTC, then add solC
Repeat steps 2 and 3 with a new syringe
Use syringe to draw up 0.1mL SolD per 5mL final volume MTC, then add solD
Repeat steps 2 and 3 with a new syringe
Use syringe to draw up 0.1mL SolE per 5mL final volume MTC, then add solE

Notes:

All media components (SolA-E) used to make media should be made anaerobic (see below) and sterilized before beginning this process.
Prepare the media aseptically and do not autoclave it – autoclaving will destroy the vitamins and cause precipitation.
Large amounts of media can be prepared in a single serum bottle, then separated into individual tubes or smaller bottles using syringes
- The recipient tubes should be made anaerobic and left with a vacuum, then autoclaved to sterilize (see Making tubes and serum bottles anaerobic)

If you accidentally make a stopper non-sterile, the top of the septa may be sterilized using 95-100% alcohol and flaming (dip the top of the vessel in alcohol, pass through a flame to ignite and let the alcohol burn off. Do not hold the vessel in the flame; just burn off the alcohol)

Preparing solutions needed for 1L MTC medium

Solution A (SolA; 1.1x MTC base)

5g carbon source (usually cellobiose for C. thermocellum or glucose for T. saccharolyticum)
- Can also leave out soluble sugars and include a strip of filter paper or small portion of crystalline cellulose for C. thermocellum
9.3g MOPS (3-[N-morphholinol]-2-hydroxypropanesulfonic acid) sodium salt (pH with HCl)
- Can substitute 8.4g MOPS free acid instead (pH with NaOH)
~Optional~ 1mL of 2% rezasurin solution (redox indicator – on Dave’s bench)
For plates: add either 1% Gel-Rite (gellan gum) with 1g MgCl₂•6H₂O OR 2-4% agar

Dissolve above in 900mL of ddH₂O
Adjust pH to 7 for thermocellum or 6.8 for T. saccharolyticum
Aliquot 90% of desired culture volume (ie 9mL SolA for 10mL final media) into appropriate container (pressure tube or serum bottle – leave minimum 1/3 of container volume as headspace), make anaerobic, and autoclave
Unused SolA can be stored after autoclaving in a normal media bottle (without making anaerobic)

Solution B (SolB; 25x stock) – add 0.2mL per 5mL final media

2g potassium citrate monohydrate
1.3g citric acid monohydrate
- Can substitute 1.19g citric acid, anhydrous
1g Na₂SO₄(sodium sulfate)
1g KH₂PO₄(Potassium phosphate, monobasic)
2.5g NaHCO₃ (Sodium bicarbonate)

Dissolve in 40mL ddH₂O, make anaerobic, and autoclave

Solution C for C. thermocellum (SolC; 50x stock) – add 0.1mL per 5mL final media

2g Urea

Dissolve in 20mL ddH₂O, make anaerobic, and autoclave

Solution C for T. saccharolyticum (SolC; 50x stock) – add 0.1mL per 5mL final media

1.5g NH₄Cl (ammonium chloride)
- Can substititute 1.5 NH₄SO₄ (ammonium sulfate)

Dissolve in 20mL ddH₂O, make anaerobic, and autoclave

Solution D (SolD; 50x stock) – add 0.1mL per 5mL final media

1g MgCl₂•6H₂O (magnesium chloride hexahydrate)
0.20g CaCl₂•2H₂O (calcium chloride dihydrate)
0.10g FeCl₂•4H₂O (ferrous chloride tetrahydrate)
1g L-cysteine HCl monohydrate
1mL solution F

Dissolve in 20mL ddH₂O, make anaerobic, and autoclave

Solution E (SolE; 50x stock) – add 0.1mL per 5mL final media

0.02g pyridoxamine HCl (MW = ~241.1)
0.004g para-aminobenzoic acid (MW = ~137.1)
0.002g biotin (MW = ~244.3)
0.002g vitamin B12 (cobalamin; MW = ~1355.4)
0.004 thiamine (MW = ~337.3)

Dissolve in 20mL ddH₂O, filter sterilize into degassed serum bottle

Solution F (SolF) – does not need to be sterilized; add to solution D before sterilizing solD

0.0005g Mn₂Cl₂•4H₂O (manganese chloride tetrahydrate)
0.0005g CoCl₂•6H₂O (cobalt chloride hexahydrate)
0.0002g ZnCl₂(Zinc Chloride)
0.0001g Cu₂Cl₂•2H₂O (cupric chloride dihydrate)
0.0001g H₃BO₃ (boric acid)
0.0001g Na₂MoO₄•2H₂O (sodium molybdenate dihydrate)
0.0001g NiCl₂•6H₂O (nickel(II) chloride hexahydrate)

Dissolve in 1L ddH₂O, store in media bottle on bench, no need to sterilize

Notes:

Solutions A-D can be stored at room temperature on your bench, but solE and solF may last longer if stored at 4°C and protected from light
Dave has a large stock of rezasurin solution and SolF and Tyler has a stock of SolE that they might share with you if you ask.

Last modified: 5 August 2021; TBJ

SOB/SOC Media

Notes

It’s recommended to make SOC media fresh for each transformation.

SOB Media

Chemical	Amount
Tryptone	20 g
Yeast Extract	5 g
5M NaCl	2 mL
2M KCl	1.25 mL
dH₂O	990 mL

Autoclave.

Sterile Magnesium Solution

Chemical	Amount
1M MgSO₄ · 7 H₂O	2.46 g
1M MgCl₂ · 6 H₂O	2.03 g
dH₂O	Add to 10 mL

Filter sterilize solution with 0.22 µM filter. Add 10 mL to SOB media

SOC Media

Add glucose solution to a final concentration of 2% to SOB media (e.g., 10 mL of a 20% glucose solution to 90 mL of SOB)

WM6026 and Mating Plates

Notes

These plates are used in the “Conjugation of ZM4 using WM6026 as donor strain” protocol

WM6026 Plates

Chemical	Amount
Tryptone	10 g
Yeast Extract	5 g
NaCl	5 g
Agar	15 g
dH₂O	Add to 1 L

Autoclave. Place in 55⁰C water bath to cool. Same as LB, but add 250 μL of 100 μg/μl spectinomycin and 10 mL of 10 mM 2,3-Diaminopropionic Acid (DAP).

Mating Plates

Chemical	Amount
Tryptone	10 g
Yeast Extract	10 g
KH₂PO₄	2 g
Glucose	20 g
Agar	17.5 g
dH₂O	Add to 1 L

Autoclave. Place in 55⁰C water bath to cool. Add 10 mL of 10 mM DAP.

Antibiotic Concentrations for Medias and Plates

Notes

These concentrations are suggestions and can change depending on your experiment.

Antibiotic	E. coli/Zymomonas (µg/mL)	B. subtilis (µg/mL)
Kanamycin	30-50	30
Spectinomycin	40-50	40
Erythromycin	0.5	0.5
Chloramphenicol	30	30
Gentamycin	12
Tetracycline	12.5
Ampicillin	60 (E. coli)	100

Phosphate Buffered Saline (PBS) 10X Solution

Notes

This protocol is from Cold Springs Harbor and can be found at: http://cshprotocols.cshlp.org/content/2006/1/pdb.rec8247
CaCl₂ and MgCl₂are necessary if working with cell culture.

Chemical	Amount (g)
NaCl	80
KCl	2
Na₂HPO₄	14.4
KH₂PO₄	2.4
CaCl₂ · 2 H₂O (optional)	1.33
MgCl₂ · 6 H₂O (optional)	1.0
dH₂O	Add to 1 L

pH to 7.2-7.4. Filter sterilize or autoclave. Store at RT.

Making tubes and serum bottles anaerobic

See this video Melanie made for a step by step visual guide

Move serum bottles and pressure tubes to be made anaerobic to vacuum manifold
Turn on vacuum pump, switch line to vacuum, open nitrogen tank and regulator to provide ~10psi
Attach each tube/bottle to manifold by puncturing stopper with attached needles
Open valves above in-use needles to allow vacuum to de-gas tubes (2min under vacuum for low volumes or for cysteine solutions; 5min for larger volumes)
Switch line to N₂ to flush media with nitrogen, let sit 30 seconds
Repeat steps 4 and 5 three times total
Close main valve on nitrogen tank, turn off vacuum pump, then vent N₂by opening manifold valve
Close manifold valve, then vent N₂in bottles by opening valve to an unused needle (if you are using every needle, you may need to pull a tube/bottle off to free up a needle and “take turns” venting excess N₂)
Close valves above each tube/bottle, remove them from needles
Tubes/bottles can be covered (optional) and autoclaved

Notes:

An empty bottle or tube with a drop of water (to make autoclaving effective) can be degassed using the above protocol. Instead of stopping after adding N₂ the final time, a final round of vacuuming can be performed, then the bottle removed from the manifold and autoclaved. These “vacuum bottles” work well as sterile bottles to filter sterilize solutions into so that you aren’t fighting the pressure buildup from adding liquid to a sealed bottle
It is important to release the N₂pressure after the last time media is flushed. Failing to do so will leave the bottles pressurized, which can cause them to blow the back out of a syringe when you try to use them.
Serum bottles can have their stoppers covered with 2 layers of foil for autoclaving. Tubes have green caps that can be used to cover their stoppers.
Instead of covering, serum bottles and tubes can be used as is. Each time contents are accessed with needles the top of the septa may be sterilized using 95-100% alcohol and flaming (dip the top of the vessel in alcohol, pass through a flame to ignite and let the alcohol burn off. Do not hold the vessel in the flame; just burn off the alcohol)

Last modified: 5 August 2021; TBJ