Jung Assay SOP — v2 (Serial Transfer Design), v1.0 Kathryn E. Caruso · 0009-0003-2436-1791 Foreman Lab · Center for Biofilm Engineering, Montana State University Updated March 2026

Standard Operating Procedure for 28-Day Ureolytic Activity Assay with Serial Transfers

Table of Contents

1 Overview 2 Safety 3 Equipment and materials Part I — Media prep Part II — Experiment setup, serial transfers, and sampling Part III — Reading the Jung assay References

1 Overview

Protocol for a 28-day ureolytic activity assay using the colorimetric method for quantifying urea concentration developed by jungNewColorimetricReaction1975. Uses a two-reagent system consisting of o-phthalaldehyde (OPA) and N-(1-naphthylethylenediamine) dihydrochloride (NED) that reacts with urea to produce a stable pink chromophore measurable at 505nm.

1.1 Isolate panel

2 Safety

⚠️ WARNING: Protocol involves handling concentrated acids and toxic reagents.

PPE:

• Lab coat
• Nitrile gloves
• Safety glasses or face shield
• Work in a fume hood when handling concentrated acids

Chemical hazards:

• Concentrated sulfuric acid (H₂SO₄): Corrosive, causes severe burns
• Concentrated nitric acid (HNO₃): Oxidizer, corrosive, causes severe burns
• o-Phthalaldehyde: Toxic, skin and respiratory sensitizer
• NED: May be carcinogenic; avoid inhalation and skin contact

Waste:

• Chemicals – liquid waste container in fume hood
• Solid waste – allow to evaporate in fume hood before disposing in autoclave trash

3 Equipment and materials

3.1 Equipment

• Single-channel pipettes (2–1000 µL range)
• Multi-channel pipettes (125 µL, 10 µL)
• Vortex
• Volumetric flasks (100 mL, 1 L)
• Graduated cylinders
• Analytical balance
• Magnetic stir bar and stir plate
• 1.5 mL microcentrifuge tubes
• 50 mL Falcon tubes
• Glass culture tubes: Durex™ Borosilicate, disposable, 16 × 150 mm (Cat. No. 47729-580) — for experimental cultures
• Tube caps/closures (autoclavable)
• 50 mL Falcon tubes (for R2A starter cultures)
• 0.2 µm syringe filters
• Sterile syringes
• 96-well plates (for Jung assay only)
• Reagent reservoirs for Reagents 2 and 3
• Microplate shaker
• Combustion oven (for glassware preparation)
• Incubator at 37°C (for Jung assay color development)
• Rotating shaker at 15°C (for experimental cultures; in fridge)
• Shaking incubator at 15°C (for starter cultures)
• Plate reader (need to reserve ahead of time)
• Spectrophotometer for OD₆₀₀ (direct reads in glass culture tubes)
• pH meter or pH strips
• Inoculating loops (sterile)

3.2 Chemicals and reagents

• Sodium succinate dibasic hexahydrate (MW 270.14 g/mol; Sigma-Aldrich Cat. No. S2378)
• Yeast extract
• K₂HPO₄ (potassium phosphate dibasic)
• Urea
• R2A broth (for starter cultures)
• Brain Heart Infusion (BHI) broth (for S. pasteurii) — see BHI Broth SOP
• Brain Heart Infusion (BHI) agar (for S. pasteurii plating) — see BHI Agar SOP
• o-Phthalaldehyde (OPA)
• N-1-naphthylethylenediamine dihydrochloride (NED)
• Brij-L23 surfactant
• Concentrated sulfuric acid (H₂SO₄, 95–98%)
• Concentrated nitric acid (HNO₃, 65–70%)
• Boric acid
• 1 M HCl (for pH adjustment)
• 1 M NaOH (for pH adjustment)
• Deionized (DI) / Milli-Q water

3.3 Total materials estimate

Part I — Media prep

Complete before Day 0

1 Glassware and equipment preparation

1.1 Combust glass culture tubes

→ See Combustion of Glassware SOP

• Combust ×66 minimum glass culture tubes per transfer (+ spares)

1.2 Autoclave tube closures and other equipment

→ See Autoclave Sterilization SOP

• Autoclave tube caps/closures for each transfer

2 Growth medium

→ See Succinate-Urea Growth Medium SOP for full formulation, preparation, urea supplementation, and volume planning.

Summary: Succinate-based medium + 2% urea (20 g/L), pH 6.8 ± 0.2. Prepare ~230 mL per transfer window (~1 L total for 4 transfers).

2.1 5% Nitric acid (for sample preservation)

→ See 5% Nitric Acid SOP for full preparation procedure.

2.2 R2A broth (for starter cultures)

→ See R2A Broth SOP for preparation.

Volume needed for this experiment: ~250 mL (20 isolates × 10 mL + 15% overage). Dispense 10 mL per 50 mL Falcon tube aseptically (×20 tubes + spares).

2.3 BHI media (for S. pasteurii positive control)

→ See BHI Agar SOP for agar plate preparation (used for plating and freezer stocks). → See BHI Broth SOP for broth preparation (used for S. pasteurii starter cultures).

BHI broth: 10 mL per starter culture in a 50 mL Falcon tube.

3 Starter culture preparation (pre-experiment)

3.1 Positive control (Sporosarcina pasteurii)

→ See S. pasteurii Starter Culture SOP for full retrieval and revival procedure.

For this experiment: Use Section 4.1 (15°C protocol) — grow in 10 mL BHI at 15°C, 150 RPM for 4 days to match test isolate conditions.

3.2 Isolates

Start isolates on R2A four days before Day 0 (i.e., Day −4).

For each of the 20 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, 150 RPM for 4 days (until visibly turbid)

Part II — Experiment setup, serial transfers, and sampling

Days 0–28

1 Standardize inocula

**Target starting OD₆₀₀: 0.02–0.04 For each starter culture:

1. Measure OD₆₀₀ of the starter culture (dilute if necessary to get an accurate reading)
2. Back-calculate the true OD if diluted:
   • True OD = measured OD × (total volume ÷ sample volume)
   • Example: 10 µL culture in 1000 µL total → True OD = measured OD × 100
3. Calculate inoculum volume using C₁V₁ = C₂V₂:
   • V₁ = (0.025 × 3,000 µL) ÷ True OD
4. Round to nearest 5 µL for pipetting accuracy
5. Record all values in lab notebook

2 Transfer 1 — Inoculation and assay initiation (Day 0)

1. For each isolate, inoculate 3 replicate tubes of succinate + urea medium (3 mL per tube)
2. Inoculate 3 replicate tubes of S. pasteurii positive control (1:100 from starter)
3. Prepare 3 replicate tubes of uninoculated medium (negative control)
4. Cap all tubes and record time of inoculation
5. Collect T0 samples immediately (see Part II, §4)
6. Measure and record Day 0 OD₆₀₀ to confirm starting density
7. Measure and record pH
8. Place all tubes in rotating shaker at 15°C, 150 RPM

Total tubes: 66

3 Serial transfers (Days 7, 14, 21)

3.1 Transfer day workflow

3.2 Transfer procedure

For each culture tube:

1. Vortex or gently mix the outgoing culture to ensure homogeneity
2. Transfer 30 µL of culture into the corresponding fresh tube containing 3 mL succinate + urea medium (1:100 dilution)

3.3 Negative control

• Prepare new uninoculated medium tubes at each transfer

3.4 Positive control

• Transfer S. pasteurii 1:100 in the same manner as test isolates

3.5 Notes

• Work in Biosafety Cabinet for all inoculation and transfer steps
• Use sterile pipettes and tubes
• Record all observations at each transfer (turbidity, color, odor, pH)
• Label tubes clearly with isolate ID, transfer number, replicate number, and date

4 Sampling

4.1 Sampling timeline

4.2 Measurements at each timepoint

At each of the 12 timepoints, collect from every tube (66 total):

1. Urea sample — preserved for Jung assay (see §4.3)
2. OD₆₀₀ — for growth tracking (see §4.4)
3. pH — for tracking alkalinization

4.3 Sample collection for urea measurement

Procedure:

1. Immediate preservation
   • Option A (Acid): Mix 30 µL culture + 570 µL of 5% HNO₃ (1:20 dilution)
2. Label and store
   • Label tubes with: Isolate ID, Transfer #, Timepoint (T0/Mid/Final), Date

Method A: Acid preservation

Advantages: Instantaneous enzyme inactivation, stable samples, no filtration required Disadvantages: Requires acid handling, standards must also be prepared in acid

• Pre-prep: Pipette 570 µL of 5% HNO₃ into labeled microcentrifuge tubes before collecting samples
  • Prepare enough tubes for all timepoints and samples
  • Label each tube with sample ID and timepoint
• Sampling: Collect exactly 30 µL of sample and add directly into the preloaded 570 µL of 5% HNO₃
  • This yields a 1:20 dilution (final volume = 600 µL)
  • Acidification instantly stops urease activity
• Mixing: Immediately vortex each tube for 5–10 seconds
• Storage: Preserved samples are stable at room temperature for several days, or refrigerate at 4°C for longer storage

4.4 OD₆₀₀ measurements

Purpose: Optical density at 600 nm provides a proxy for cell density, used to normalize urease activity to biomass and track growth kinetics throughout the assay.

Procedure:

1. Remove all tubes from the rotating shaker
2. Blank the spectrophotometer with the corresponding uninoculated negative control tube
3. Read OD₆₀₀ directly in the glass culture tubes
4. Record the time of measurement (HH:MM) for each reading
5. Return tubes to rotating shaker promptly

Part III — Reading the Jung assay

Run after collecting preserved samples. Can be batched — does not need to happen on sampling day if using acid preservation.

1 Jung assay reagent preparation

Note: Always add acid to water (never water to acid) when diluting concentrated acids. Work in a fume hood.

1.1 Reagent 1: Brij-L23 Solution (33% v/v)

Purpose: Surfactant to enhance color development and stabilize the chromophore Preparation:

1. Add 29 mL of deionized water to a clean graduated cylinder or beaker
2. Add 1 mL of Brij-L23 surfactant
3. Mix thoroughly by gentle swirling
4. Transfer to a labeled bottle and store at room temperature

Storage: Room temperature, stable for several months. Final concentration: 33% v/v.

1.2 Reagent 2: o-Phthalaldehyde (OPA) Solution

Purpose: Primary reagent that reacts with urea in the presence of sulfuric acid Volume: 1 L (sufficient for ~4,000 wells) Preparation (in fume hood):

1. Add approximately 800 mL of deionized water to a 1 L volumetric flask
2. Carefully add 74 mL of concentrated H₂SO₄ (SLOWLY, always acid to water)
   • Pour the acid slowly down the side of the flask while swirling gently
   • The solution will heat up significantly; this is normal
3. Allow the solution to cool to room temperature (20–30 minutes)
4. Weigh 200 mg of o-phthalaldehyde powder
5. Add the o-phthalaldehyde to the cooled acidic solution
6. Add 1 mL of Reagent 1 (Brij solution)
7. Swirl gently until all o-phthalaldehyde is completely dissolved
8. Add deionized water to bring the final volume to exactly 1 L

Storage: Store in an amber bottle at room temperature. Stable for several weeks.

1.3 Reagent 3: NED Solution

Purpose: Secondary reagent that completes the chromophore formation, producing the pink color Volume: 1 L (sufficient for ~4,000 wells)

Preparation (in fume hood):

1. Add approximately 600 mL of deionized water to a 1 L volumetric flask
2. Weigh 5.0 g of boric acid
3. Add the boric acid to the water and swirl until completely dissolved
4. Carefully add 222 mL of concentrated H₂SO₄ (SLOWLY, always acid to water)
   • Pour slowly down the side of the flask while swirling
   • Allow adequate cooling time due to heat generation
5. Allow the solution to cool to room temperature (30–45 minutes)
6. Weigh 600 mg of N-1-naphthylethylenediamine dihydrochloride (NED)
7. Add the NED to the cooled solution
8. Add 1 mL of Reagent 1 (Brij solution)
9. Swirl gently until all NED is completely dissolved
10. Add deionized water to bring the final volume to exactly 1 L

Storage: Store in an amber bottle at room temperature. Stable for several weeks. Protect from light. Discard if solution turns dark brown.

2 Urea standards

⚠️ CRITICAL: Standards must be matrix-matched to your samples. If samples are in 5% HNO₃, prepare standards in 5% HNO₃. If samples are in DI water, prepare standards in DI water.

2.1 Urea stock (2.0 g/L)

Preparation:

1. Accurately weigh 0.200 g (±0.001 g) of urea using an analytical balance
2. Transfer to a 100 mL volumetric flask
3. Add approximately 80 mL of either:
   • Deionized water (for DI-based standards), OR
   • 5% HNO₃ solution (for acid-preserved samples)
4. Swirl gently until urea is completely dissolved
5. Add solvent to bring final volume to exactly 100 mL
6. Mix thoroughly by inverting 10 times
7. Label with concentration, date, and initials

Storage: Refrigerate at 4°C in a tightly sealed container. Stable for one week. Discard and prepare fresh weekly.

2.2 Working standards

Prepare working standards fresh on the day of analysis by diluting the 2.0 g/L stock solution.

Table. Dilution instructions for preparing 10 mL of each standard concentration.

Storage: Working standards should be used fresh (same day). If necessary, refrigerate at 4°C for up to 24 hours. Bring to room temperature before use.

3 96-Well plate assay

3.1 Scale per timepoint

• 66 samples × 3 technical replicates = 198 sample wells
• 6 standards × 3 replicates = 18 standard wells
• Total: 216 wells per timepoint → 3 plates
• Include one set of standards per plate for internal consistency

3.2 Plate layout (suggested)

Plate 1 of 3 (per timepoint) — Isolates 1–8 + Standards

Note: This is a suggested layout. Adjust as needed for your pipetting workflow.

Plate 2 of 3 — Isolates 9–16 + Standards Plate 3 of 3 — Isolates 17–20 + Controls (S. pasteurii, uninoculated) + Standards

3.3 Assay volumes per well

3.4 Color development guide

3.5 Step-by-step procedure

Step 1: Sample and standard addition

• Add 10 µL of each standard to 3 wells (triplicates)
  • Include all six standards: 2.0, 1.5, 1.0, 0.5, 0.25, and 0.0 g/L
  • Mix each standard thoroughly before pipetting
• Add 10 µL of each prepared sample to 3 wells (triplicates)

Important: Ensure samples are well-mixed before pipetting. If samples have settled, vortex briefly before adding to the plate.

Step 2: Reagent 2 addition

• Pour adequate volume of Reagent 2 (OPA solution) into a reagent reservoir
• Using a multi-channel pipette, add 125 µL of Reagent 2 to each well
  • Pipette slowly to avoid bubble formation
  • Ensure reagent is added to the bottom of each well

Step 3: Reagent 3 addition

• Pour adequate volume of Reagent 3 (NED solution) into a clean reagent reservoir
• Using a multi-channel pipette, add 125 µL of Reagent 3 to each well
  • Total volume per well is now 260 µL

Step 4: Mixing

• Place the microplate on a microplate shaker
• Mix at 450 RPM for 1 minute
  • Check for bubbles; if present, gently tap the plate to remove them

Step 5: Incubation

• Cover the plate with a plate lid
• Incubate at 37°C for 30 minutes
  • The pink chromophore develops during this incubation period

Step 6: Absorbance reading

• After exactly 30 minutes, read absorbance on a microplate reader
• Set wavelength to 505 nm
• Record absorbance values for all wells

Expected color: Wells should display a pink color. Higher urea concentrations will show darker pink, while low concentrations will be lighter pink or nearly colorless.

Step 7: Waste disposal ⚠️ CRITICAL: All liquid in the plate and excess reagents must be disposed of in a properly labeled hazardous waste container. Do not pour down the drain. Follow institutional chemical waste protocols.

4 Analysis

4.1 Quality control

Before proceeding with calculations, verify:

• Coefficient of variation (CV) for triplicates: Should be <5%
• Blank absorbance: Should be <0.05 at 505 nm
• Standard curve linearity: R² ≥ 0.97 (preferably ≥ 0.99)

4.2 Standard curve construction

Step 1: Average triplicate absorbance values

For each standard, calculate the mean absorbance of the three replicate wells: A_mean = (A₁ + A₂ + A₃) / 3

Step 2: Subtract blank

Subtract the mean absorbance of the 0.0 g/L standard (blank) from all other values: A_corrected = A_mean − A_blank

Step 3: Plot standard curve

Create a scatter plot with:

• X-axis: Urea concentration (g/L)
• Y-axis: Corrected absorbance (A_corrected)

Step 4: Perform linear regression

Fit a linear regression line: y = mx + b

• y = corrected absorbance
• x = urea concentration (g/L)
• m = slope (sensitivity)
• b = y-intercept (should be near zero after blank correction)

Acceptance criteria: R² ≥ 0.97 (preferably ≥ 0.99). If R² is below 0.97, troubleshoot and repeat the assay.

4.3 Sample concentration calculation

Step 1: Average sample triplicates

A_sample = (A₁ + A₂ + A₃) / 3

Step 2: Subtract blank

A_corrected = A_sample − A_blank

Step 3: Calculate measured concentration

C_m = (A_corrected − b) / m

Step 4: Apply dilution factor

C_sample = C_m × DF

4.4 Reporting

• Concentration: Report in g/L or mg/L as appropriate
• Precision: Report mean ± standard deviation (SD) from triplicates
• Dilution factor: Note the dilution used
• Preservation method: Specify acid preservation or filtration
• Calibration equation and R²: Include for quality assurance

Example entry: “Sample GG7-T2-Mid: Urea concentration = 17.0 ± 0.4 g/L (mean ± SD, n=3). Sample diluted 1:20 in 5% HNO₃. Calibration curve: y = 0.498x + 0.005, R² = 0.998.”

References

Protocol adapted from:

Jung, D., Biggs, H., Erikson, J., & Ledyard, P. U. (1975). New Colorimetric Reaction for End-Point, Continuous-Flow, and Kinetic Measurement of Urea. Clinical Chemistry, 21(8), 1136–1140. https://doi.org/10.1093/clinchem/21.8.1136

Quick reference

Experimental design

• 28-day assay with 4 serial transfer windows (Days 0–7, 7–14, 14–21, 21–28)
• 20 isolates × 3 biological replicates = 60 culture tubes
• Controls: S. pasteurii (positive, ×3) + uninoculated medium (negative, ×3) = 6 tubes
• 66 total culture tubes per transfer window
• Culture vessel: Combusted glass culture tubes (16 × 150 mm) on rotating shaker
• Starter cultures: R2A broth (isolates) / BHI broth (S. pasteurii), 50 mL Falcon tubes, 15°C shaking 150 RPM, 4 days
• 1:100 transfer ratio into fresh medium every 7 days
• 3 timepoints per transfer: T0, Mid (~Day 3–4), Final (Day 7)
• 12 total timepoints across the experiment
• OD₆₀₀: Direct reads in glass tubes via spectrophotometer

Growth medium

• Sodium succinate dibasic hexahydrate: 1.126 g/L
• Yeast extract: 0.1 g/L
• K₂HPO₄: 0.3 g/L
• Urea: 20 g/L (2%; from 10% stock, filter-sterilized)
• pH 6.8 ± 0.2
• Incubation: 15°C, rotating shaker, 150 RPM

Sample preparation

• Acid method: 30 µL sample + 570 µL 5% HNO₃ (1:20 dilution)
• Filtration method: Filter through 0.2 µm, dilute with DI water

Jung assay volumes per well

• Sample/Standard: 10 µL
• Reagent 2 (OPA): 125 µL
• Reagent 3 (NED): 125 µL

Jung assay conditions

• Temperature: 37°C
• Time: 30 minutes
• Mixing: 450 RPM for 1 minute before incubation
• Detection: 505 nm
• Linear range: 0–2 g/L urea

Acceptance criteria

• Standard curve R² ≥ 0.97 (preferably ≥ 0.99)
• Triplicate CV < 5%
• Blank absorbance < 0.05