Carbon Source Growth Assay, v1.0 Kathryn E. Caruso · 0009-0003-2436-1791 Foreman Lab · Center for Biofilm Engineering, Montana State University Updated March 2026

Isolates:

Original base media

Experimental conditions:

Experimental design summary:

• 4 isolates × 3 conditions × 3 replicates = 36 inoculated tubes
• 3 uninoculated media blanks (1 per condition) = 3 blank tubes
• Total: 39 tubes

Materials

Media/reagents:

• Yeast extract
• Sodium DL-malate, disodium salt (MW 178.05 g/mol; CAS 676-46-0) — racemic mixture; only the L-form is metabolically active as a TCA cycle intermediate
• Sodium succinate dibasic hexahydrate (disodium succinate hexahydrate; MW 270.14 g/mol; Sigma-Aldrich Cat. No. S2378; CAS 6106-21-4) — note: hexahydrate form; account for water of crystallization in mass calculations
• Sodium acetate, anhydrous, 99% (MW 82.03 g/mol, CAS 127-09-3)
• K₂HPO₄ (potassium phosphate)
• CH₄N₂O (urea)
• 1M HCl
• 1M NaOH
• Sterile deionized water

Equipment:

• Analytical balance
• Graduated cylinders or volumetric flasks
• Magnetic stir bar and stir plate
• 0.22 μm syringe filter
• Sterile syringe
• Sterile collection bottles/flasks
• Glass culture tubes: DurexTM Borosilicate, disposable, 16 × 150 mm (Cat. No. 47729-580) — ×39 minimum
• Color-coded stickers for tube labeling
• Inoculating loop (sterile)
• Rotating shaker at 15°C (in fridge)
• Spectrophotometer (OD600)
• Sterile pipettes and tips
• 70% ethanol
• pH strips
• Lab notebook

1 Pre-experiment preparation

1.1 Combust glassware

Combustion (dry heat sterilization) removes all organic residues and ensures tubes are free of contaminants that could interfere with growth or OD readings.

1. Wrap glass culture tubes (×39 minimum + spares) loosely in aluminum foil
2. Place in combustion oven
3. Combust at 450°C for 5 hours
4. Allow to cool completely inside the oven before removing — do not open door while hot (thermal shock can crack tubes)
5. Keep tubes wrapped/covered until use

1.2 Autoclave tube closures and other equipment

Autoclave all non-glass items that will contact sterile media or cultures.

Items to autoclave:

• Tube caps/closures

Procedure:

1. Wrap or bag each item loosely in aluminum foil — ensure steam can penetrate
2. Autoclave at 121°C, 15 psi, 15–20 min dry cycle
3. Allow to dry and cool completely inside autoclave
4. Store sealed/wrapped until use
5. Label autoclave tape with date and initials

2 Preparing media

Prepare 3 base media (one per carbon source condition) following the same procedure as the original urea-based growth medium protocol, substituting the carbon source as indicated.

2.1 Media formulations

Table 1. Base medium formulation per condition (per liter). Amounts calculated to match 16.67 mmol C/L (equivalent to 0.5 g/L glucose).

2.2 Volume planning

2.3 Prepare each base medium

For each of the 3 conditions, follow the base medium preparation steps from the original urea-based growth medium protocol (Sections 1.1–1.5):

1. Weigh components
2. Dissolve in ~80% of target Milli-Q water volume
3. Adjust pH to 6.8 ± 0.2
4. Bring to final volume to 200mL
5. Autoclave at 121°C, 15 psi, 20 min/liter (liquid cycle)
6. Cool to room temperature

2.5 Prepare urea solution

Follow the urea solution preparation from the original protocol (Section 2):

1. Weigh 20 g urea per 100 mL DI water (20% w/v stock)
2. Dissolve at room temperature (do not heat)
3. Filter sterilize through 0.22 μm syringe filter in biosafety cabinet
4. Store at 4°C; use within 1 week

2.6 Combine base medium and urea solution

For each condition, aseptically combine base medium and urea stock to achieve 2% (w/v) final urea concentration:

• Work in biosafety cabinet
• C₁V₁ = C₂V₂: (20%)(V₁) = (2%)(final volume)
• V₁ = 10% of final volume as urea stock
• Remaining 90% = base medium
• Mix gently by swirling

3 Inoculum preparation

3.1 Prepare starter cultures

For each of the 4 isolates:

1. Select an individual colony from the R2A agar plate
2. Using a sterile loop, transfer the colony into a 50 mL Falcon tube containing 10 mL sterile R2A broth
3. Incubate on a shaking incubator at 15°C until visibly turbid

3.2 Standardize inoculum

Target starting OD: 0.025

Measuring starter culture OD

Many starter cultures will be too dense to read directly — use a cuvette dilution series:

1. Add a small volume of starter culture to 1000 µL Milli-Q water in a cuvette
2. Measure OD600; if reading is >0.3, try a smaller volume or re-dilute
3. Back-calculate the true OD of the undiluted culture:
   • True OD = measured OD × (total cuvette volume ÷ sample volume added)
   • Example: 10 µL culture in 1000 µL total → True OD = measured OD × 100
4. Repeat with additional volumes if needed to confirm; average confirmatory reads
5. Record all dilution steps and measured values in lab notebook

Calculating inoculum volume

Use C₁V₁ with the confirmed true OD:

• C₁ = true OD of starter culture
• V₁ = inoculum volume to add (solve for this)
• C₂ = 0.025 (target starting OD)
• V₂ = 3000 µL (final tube volume)
• V₁ = (0.025 × 3000) ÷ C₁
• Round to nearest 5 µL for pipetting accuracy

Inoculum volumes (inoculation date: 2026-03-04)

3.3 Inoculate experimental tubes

1. Add the appropriate volume of starter culture to each tube, then bring to 3 mL final volume with the corresponding condition medium
2. Leave blank tubes uninoculated
3. Cap all tubes and record time of inoculation
4. Measure and record actual Day 0 OD600 for each tube to confirm starting density

4 Tube labeling

4.1 Color-coded sticker scheme

Assign one sticker color per isolate for quick visual identification:

4.2 Full tube list

5 Incubation

• Temperature: 15°C (fridge)
• Agitation: Rotating shaker, 150RPM
• Duration: ~5 days

6 OD600 measurements

6.1 Measurement schedule

Measurements are taken more frequently during early growth and spaced out as the experiment progresses. Days 1 and 2 each include three timepoints to capture early growth kinetics.

6.2 Measurement procedure

For each measurement session (single or one of multiple timepoints in a day):

1. Remove all tubes from the shaker
2. Blank the spectrophotometer with the corresponding uninoculated blank for each condition — blank once per condition, before reading all tubes in that condition:
   • Blank with BLK-Mal → read all 13 malate tubes
   • Blank with BLK-Suc → read all 13 succinate tubes
   • Blank with BLK-Ace → read all 13 acetate tubes
3. Read OD600 directly in the glass culture tubes
4. Record the time of measurement (HH:MM) for each reading
5. Return tubes to shaker promptly

7 Endpoint measurements

At the conclusion of the experiment (~Day 5):

7.1 Final OD600

• Take final OD600 reading as in Section 6.2

7.2 Endpoint pH

• Measure pH of each tube using a pH meter (preferred) or pH strips
• Record pH alongside the final OD600 reading
• Compare to initial pH (6.8) — an increase toward alkaline suggests urease activity