Pouring Plates

1. Prepare and autoclave media containing agar.
   • If cooling on a plate, add stir bar before autoclaving.
2. Immediately after autoclaving, cool for pouring.
   • move molten agar to 55°C water bath.
   • Cool to stir on plate.
3. Once cooled to 55°C, add any heat-labile components (vitamins, antibiotics) aseptically, then mix
   • Avoid heavy stirring, which can cause bubbles to form
4. 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)
5. 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
Na2HPO4 anhydrous	60 g
KH2PO4 anhydrous	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
MgSO4 anhydrous 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
KH2PO4 anhydrous	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

1. Autoclave water.
2. Mix all components aseptically into sterile vessel(s).
3. 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

1. Mix all components with ~half the final volume of water
2. Once all components are dissolved, adjust to appropriate final volume
3. Filter sterilize using screw cap filter
4. 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
K2HPO4 anhydrous	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 · 7 H2O	1 mL
25g/L Na2MoO4 · 2 H2O	1 mL
2.5g/L FeSO4 · 7 H2O	1 mL
20g/L CaCl2 · 2 H2O	1 mL
5mg/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 • 7H2O	2g
Na2MoO4 · 2 H2O	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

Zymomonas minimal media for Hungate (ZMH) media Adaptive Evolution Experiment 

This makes 100mL of minimal media with the desired concentration of isobutanol, the 100mL are then separated into empty, anaerobic, autoclaved Hungate tubes, 10mL each. 

First add each components below into single 100mL Hungate bottle, make anaerobic and then autoclave. (Except for the Calcium Pantothenate, this is made and filtered sterilized in the anaerobic chamber into an empty, anaerobic, autoclaved 100mL Hungate bottle.) 

Components 

• 75mL ddH2O 

• 2.00g/100mL MgSO4 · 7 H2O 

• 2.50g/L Na2MoO4 ·  H2O 

• 0.25g/L FeSO4 · 7 H2O 

• 2.0g/L CaCl2 · 2 H2O 

• 20g/100mL (20%) Glucose Solution 

• Minimal media 10X Base 

• 10 g of KH2PO4  

• 10 g of K2HPO4  

• 5 g of NaCl  

• 10 g of (NH4)2SO4  

• Fill to 1 L with dH2O 

• Do not use this unless you need nitrogen-free minimal media 10X Base 

• 10 g of KH2PO4  

• 10 g of K2HPO4  

• 5 g of NaCl  

• 10 g of K2SO4 

• Fill to 1 L with dH2O 

• 50.00mg/100.00mL Calcium Pantothenate (After making, store in the fridge) 

• First make an empty 100mL Hungate bottle anaerobic and then autoclave. 

• Make the 50.00mg/100.00mL Calcium Pantothenate with autoclaved ddH2O 

• In the anaerobic chamber, with a 50mL syringe, filter and needle inject it into the empty anaerobic autoclaved 100mL Hungate bottle. 

Components for 100mLs of Minimal Media
ddH20	75 mL
Minimal Media 10X Base	10 mL
20g/100mL (20%) glucose solution	10 mL
2.00g/100mL MgSO4 · 7 H2O	1 mL
2.50g/L Na2MoO4 ·  H2O	1 mL
0.25g/L FeSO4 · 7 H2O	1 mL
2.0g/L CaCl2 · 2 H2O	1 mL
50.00mg/100.00mL Calcium Pantothenate	1 mL

For isobutanol media, make a 4.616% isobutanol solution with ddH2O in a 100mL Hungate bottle, make anaerobic and then autoclave. 

Add the following components, pay attention at the isobutanol molarity. 

Minimal Media for 100mL Hungate Bottle
Minimal Media 10X Base	10 mL
20g/100mL (20%) glucose solution	10 mL
2.00g/100mL MgSO4 · 7 H2O	1 mL
2.50g/L Na2MoO4 ·  H2O	1 mL
0.25g/L FeSO4 · 7 H2O	1 mL
2.0g/L CaCl2 · 2 H2O	1 mL
50.00mg/100.00mL Calcium Pantothenate	1 mL
DDH2O needed for:
0.00M Isobutanol	75 mL
0.05 M Isobutanol	65 mL
0.10 M Isobutanol	55 mL
0.13 M Isobutanol	49 mL
0.15 M Isobutanol	45 mL
0.16M Isobutanol	43 mL
0.17M Isobutanol	41 mL
0.18M Isobutanol	39 mL
0.19M Isobutanol	37 mL
0.20M Isobutanol	35 mL
4.616% Isobutanol needed for:
Control	0 mL
0.05 M Isobutanol	10 mL
0.10 M Isobutanol	20 mL
0.13 M Isobutanol	26 mL
0.15 M Isobutanol	30 mL
0.16 M Isobutanol	32 mL
0.17 M Isobutanol	34 mL
0.18M Isobutanol	36 mL
0.19M Isobutanol	38 mL
0.20M Isobutanol	40 mL

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

1. 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)
2. Attach a new, sterile syringe to a new, sterile needle aseptically
3. Make syringe anaerobic by drawing up headspace gas from sterile, anaerobic bottle of 2.5% Cys-HCl
4. Use syringe to draw up 0.2mL SolB per 5mL final volume MTC, then add solB (to tube containing solA)
5. Repeat steps 2 and 3 with a new syringe
6. Use syringe to draw up 0.1mL SolC per 5mL final volume MTC, then add solC
7. Repeat steps 2 and 3 with a new syringe
8. Use syringe to draw up 0.1mL SolD per 5mL final volume MTC, then add solD
9. Repeat steps 2 and 3 with a new syringe
10. 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

1. Dissolve above in 900mL of ddH₂O
2. Adjust pH to 7 for thermocellum or 6.8 for T. saccharolyticum
3. 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
4. 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 MgCl₂ are 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

1. Move serum bottles and pressure tubes to be made anaerobic to vacuum manifold
2. Turn on vacuum pump, switch line to vacuum, open nitrogen tank and regulator to provide ~10psi
3. Attach each tube/bottle to manifold by puncturing stopper with attached needles
4. 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)
5. Switch line to N₂ to flush media with nitrogen, let sit 30 seconds
6. Repeat steps 4 and 5 three times total
7. Close main valve on nitrogen tank, turn off vacuum pump, then vent N₂ by opening manifold valve
8. Close manifold valve, then vent N₂ in 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)
9. Close valves above each tube/bottle, remove them from needles
10. 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 N₂ pressure 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