Experiments

Our wet lab comprised a total of six in-person sessions at Institute for Research on Innovative Science (IRIS) Lab, headquartered in Seoul, South Korea. The biosafety provisions concerning our project can be found in the Safety page. Below, we list the experimental principle, methods, and materials for each experimentation we conducted over the course of our project:

Session Agenda #1: Micropipette Practice

Experimental Principle: Accurate pipetting is essential for precise DNA and bacterial handling. Practicing ensures consistency and avoids contamination.

Materials Needed:

• Micropipette
• Pipette tips
• Test tubes
• Distilled water

Experimental Procedure:

1. Check that the volume dial is set to the correct volume (10 µL).
2. Firmly press the pipette into a sterile tip until it clicks in place.
3. Tip slightly below the surface of the water and press the button to the first stop.
4. Make sure the pipette is perpendicular to the liquid.
5. Slowly release the button to draw water into the tip.
6. Make sure there are no air bubbles.
7. Press the button to the first stop and expel all liquid.
8. Pour the liquid close to the container in contact.
9. Pour the liquid onto the bottom middle of the container.
10. Repeat five times per person to practice.

Session Agenda #2: DNA Introduction into E. coli

Experimental Principle: Introducing the plasmid DNA into E. coli cells before heat shock allows transformation during the next step.

Materials Needed:

• 4 test tubes with E. coli solution on ice
• Alcohol burner
• Fume hood
• Plasmid DNA (1 µL per tube)
• LB media (100 µL)
• Micropipette & sterile tips

Experimental Procedure:

1. Place the test tubes containing E. coli on ice in a rack.
2. The test tubes are labelled with each group name.
3. Micropipette 1 µL of the plasmid DNA into the E.coli solution for each 4 test tubes.
4. We worked at the fume hood. An alcohol burner was lighted up in the fume hood to minimize airborne contamination.
5. Make sure to tap the side of each tube with your thumb after micropipetting in order to mix the solution evenly without creating bubbles.

Session Agenda #3: Heat Shock Transformation

Experimental Principle: Rapid heat shock creates temporary pores in bacterial membranes, allowing plasmid entry. LB media helps recovery and provides nutrition.

Materials Needed:

• Water bath (46 °C)
• 4 E. coli + DNA solution test tubes
• Ice container
• LB media (100 µL per tube)
• Micropipette & tips

Procedure

1. Transfer 4 tubes (E. coli + DNA) from ice into a 46 °C water bath for 1 min 30 sec.
2. Immediately return tubes to ice for 5 min recovery.
3. Pipette 100 µL of the LB (Luria-Bertani) media solution into each test tube.
4. Make sure to tap the test tube with your thumb to mix evenly.
5. Place tubes in a 37 °C incubator for 30 min to recover.

Session Agenda #4: Agar Plate Preparation

Experimental Principle: LB agar provides nutrients and a surface for colonies.

Materials Needed:

• Petri dishes
• LB broth high salt (5 g)
• Bacto agar (5 g)
• Distilled water (200 mL)
• Erlenmeyer flask
• Scale
• Wrap
• Microwave
• Erythromycin (80 µL)

Experimental Procedure:

1. Label petri dishes with date, team number, and names.
2. Weigh 5 g LB high salt, 3 g agar, and 200 mL distilled water and pour into the flask.
3. Microwave for 2 min 50 sec, until the solution turns transparent yellow with no solids.
4. Cover the flask opening with wrap and poke a small hole.
5. Pipette 80 µL erythromycin into the solution and swirl gently.
6. Pour ~20 mL into each petri dish (half height).
7. Avoid bubbles since it is hard to distinguish between bacteria colonies and solidified bubbles.
8. Allow agar to solidify at room temperature.

Session Agenda #5: Plating Bacteria

Experimental Principle: Spreading bacteria evenly on agar ensures individual cells from colonies. Incubation at optimal temperature promotes growth.

Materials Needed:

• Solidified agar plates with erythromycin
• Test tubes with recovered E. coli
• Micropipette + sterile tips
• Sterilized spreader
• Shaking Incubator (37 °C)

Experimental Procedure:

1. Pipette 100 µL bacterial culture onto the center of the agar plate on the petri dish.
2. Place a sterilized spreader on the plate; spread bacteria evenly across the surface.
3. Place the plates inside the 37 °C shaking incubator for 24 hours.

Session Agenda #6: Transferring Grown Bacteria to Test Tube Solutions

Experimental Principle: To extract the bacteria that grew on the agar plates, colonies were transferred into fresh liquid media so the bacteria can multiply, producing more plasmid copies.

Session Agenda #1: Harvesting Bacterial Cells

Experimental Principle: Centrifugation separates bacterial cells from liquid media. The dense cells result at the bottom while the LB medium remains above.

Materials Needed:

• 4 bacterial culture tubes
• Refrigerated centrifuge
• Ice bucket

Experimental Procedure:

1. Centrifuge the 4 bacterial culture tubes at 3220 rpm, 4 °C, for 3 min.
2. Make sure they are balanced across the rotor.
3. 2. Pour out the LB media into a waste container, leaving only the pelleted bacteria.

Session Agenda #2: Plasmid Extraction

Experimental Principle: The bacterial cells are lysed to release DNA. Potassium acetate neutralizes the solution, precipitating proteins and chromosomal DNA, leaving plasmid only.

Materials Needed:

• EX Pure Plasmid Kit
• S1 Resuspension Buffer (NaOH +RNase)
• S2 Lysis Buffer (alkaline solution, NaOH +SDS)
• S3 Neutralization Buffer (potassium acetate)
• Vortex mixer
• Microcentrifuge
• Microtubes

Experimental Procedure:

1. Add 250 µL S1 Resuspension Buffer to the bacterial pellet.
2. Vortex each tube until the suspension is uniform with no clumps.
3. Label fresh microtubes and transfer 300 µL of the suspension to each.
4. Add 250 µL S2 Lysis Buffer to each tube and mix gently.
5. Add 350 µL S3 Neutralization Buffer to each tube.
6. Mix it well until the mixture turns white (precipitate forms).
7. Centrifuge the tubes at 13,000 rpm, 21 °C, for 10 min.
8. After centrifugation, a white pellet forms (cell debris, proteins) with a clear supernatant (plasmid DNA).
9. Pipette 750 µL supernatant plasmid DNA into the DNA binding column.
10. Centrifuge at 13,000 rpm, 25 °C, for 1 min.

Session Agenda #3: Washing and Eluting Plasmid DNA

Experimental Principle: Washing buffers remove proteins, salts, and contaminants, leaving plasmid DNA bound to the membrane. Elution releases purified plasmid DNA into a clean tube.

Materials Needed:

• DNA binding column
• AW Washing Buffer (ethanol-based)
• ED Elution Buffer
• Microcentrifuge
• Collection tubes

Experimental Procedure:

1. Add 700 µL AW Washing Buffer into the DNA binding column.
2. Centrifuge at 13,000 rpm, 25 °C, for 1 min.
3. Remove all the non-plasmid substances.
4. Centrifuge again at 13,000 rpm, 25 °C, for 1 min to remove the Washing Buffer.
5. Remove the “insert” residues to a new test tube.
6. Add 50 µL ED Elution Buffer directly to the membrane
7. Let stand for 1 min at room temperature.
8. Centrifuge at 13,000 rpm, 25 °C, for 1 min.
9. Remove column insert so that the bottom tube now contains purified plasmid DNA.

Session Agenda #4: DNA Quantification

Experimental Principle: Nanodrop uses UV absorbance to measure DNA concentration & purity.

Materials Needed:

• Nanodrop spectrophotometer
• ED buffer
• Micropipette + tips
• Alcohol wipes

Experimental Procedure:

1. Open Nanodrop software and select dsDNA mode.
2. Test ED buffer for the control.
3. Clean pedestal with alcohol wipes.
4. Load 2 µL of each DNA sample, close the lid, and measure the concentration.

Results for DNA Concentration:

Results for Final DNA Concentration and Purity Ratio:

Session Agenda #1: SDS-PAGE Sample Preparation

Experimental Principle: Samples of different conditions in the presence of the plasmid and whether Lactobacillus reuteri underwent lysis to assess the production of the pgxC protein.

Materials Needed:

• 2 Lactobacillus reuteri-containing solutions, one treated with plasmid and the other not.
• 4 small test tubes
• Centrifuge
• Lysis buffer
• Vortex machine
• Sonicator

Experimental Procedure:

1. Centrifuge two 50 mL test tubes, each containing the solution of no-plasmid L. reuteri and plasmid-inserted L. reuteri , for 2 min at 3000rpm and 4 °C.
2. Pipette 5mL of the centrifuged solution, ideally only pulling out the liquid from each centrifuged test tube, and transfer them into small test tubes. Label the small test tubes as “+ plasmid solution,” “- plasmid solution.”
3. Discard the rest of the solution, leaving only the L. reuteri cells in each test tube.
4. Add 10mL of lysis buffer to each of the test tubes containing Lactobacillus reuteri, which functions to burst the cells and to stabilize the pgxC protein.
5. Vortex both test tubes that contain the lysis buffer and Lactobacillus reuteri for approximately 45 seconds to attain a homogeneous solution without clumped cells.
6. Transfer the solution into smaller tubes to fit them into the sonicator.
7. Insert both test tubes into the Sonicator (“Ultrasonic Processor”) to burst the cells.
8. Run the sonicator for 30 seconds, set at pulse 1 sec/1(1 sec on, 1 sec off), amp 1/20%.
9. Repeat for 30 seconds, set at pulse 1 sec/1(1 sec on, 1 sec off), amp 1/20%.
10. Run the sonicator again for 1 minute, set at pulse 1 sec/1, amp 1/20%.
11. Run the sonicator again for 1 minute, set at pulse 1 sec/1, amp 1/40%.
12. Run the sonicator again for 2 minutes, set at pulse 1 sec/1, amp 1/40%.
13. Centrifuge both sonicated test tubes at 3000rpm and 19 °C for 2 minutes to segregate the cell leftovers, which will be submerged after centrifuging, and the pgxC protein.
14. Transfer each solution to test tubes, labelling each as “+ plasmid cell,” “- plasmid cell.”
15. As a result, there are 4 small test tubes labelled as: +plasmid cell, +plasmid solution, -plasmid cell, -plasmid solution.

Session Agenda #2: Gel Electrophoresis

Experimental Principle: For the verification of the size of the protein produced, and to confirm that it has the size of the protein desired of production, pgxC.

Materials Needed:

• SDS 1x solution
• 4 pgxC samples from previous agenda
• Electrophoresis device
• SDS PAGE Running Buffers
• Protein marker

Experimental Procedure:

1. Insert 50 µL SDS 1x solution into all 4 small test tubes and mix the solution in each test tube with the pipette with repeated insert-eject motion.
2. Fill the electrophoresis device (“Mini-PROTEAN Tetra System”) with SDS PAGE Running Buffers loaded on both upper and lower components
3. Load the protein marker 3µL into the first slot of the electrophoresis well.
4. Load the 4 samples, each 10µL, into the electrophoresis well, in the order of: -plasmid solution, +plasmid solution, -plasmid cell, +plasmid cell.
5. The previous step is repeated– now the 4 different samples are each loaded in two wells, resulting in a total of 8 samples loaded.
6. Initiate electrophoresis at 160V, 0.03 Amperes for 40 min.

Session Agenda #3: Gel Staining

Experimental Principle: SDG-PAGE gel staining was performed to make the strips of protein that have undergone electrophoresis easily observable.

Materials Needed:

• Electrophoresis gel that has undergone SDS-PAGE
• Blue pigmentation solution: Coomassie Brilliant Blue R-250 Staining Solution
• Nutating mixer

Experimental Procedure:

1. Remove the Gel after electrophoresis carefully, as it tends to rip easily.
2. Cut off the unnecessary well walls with a green scraper.
3. Submerge the gel into a blue pigmentation solution (Coomassie Brilliant Blue R-250 Staining Solution) for around 1 min, on the nutating mixer.

Session Agenda #1: Bradford Assay Sample Preparation

Experimental Principle: To conduct the Bradford Assay, through which the concentration of pgxC can be verified, the following four samples need to be prepared: +plasmid cell, +plasmid solution, -plasmid cell, -plasmid solution.

Materials Needed:

• 2 Lactobacillus reuteri-containing solutions, one treated with plasmid and the other not.
• 4 small test tubes
• Centrifuge
• Lysis buffer
• Vortex machine
• Sonicator

Experimental Procedure:

1. Centrifuge two 50 mL test tubes, each containing the solution of no-plasmid Lactobacillus reuteri and plasmid-inserted L. reuteri, for 3 min at 3000 rpm and 4 °C.
2. Pipette 1000 μl of the centrifuged 50 mL test tubes into small sample test tubes, tagging each as plasmid (+) solution and plasmid (-) solution.
3. Discard the centrifuged solution in the 50 mL test tubes, leaving only the clumped cells.
4. Pipette 8.5 mL of lysis buffer into each 50 mL test tube, using the pipette mixing technique for a homogeneous mixture of cells and lysis buffer.
5. Transfer the solutions into smaller test tubes for sonication.
6. Sonication for 15 min at 1/40% amplitude, pulse 3 sec/3.

Session Agenda #2: Bradford Assay

Experimental Principle: Assessing the concentration of the pgxC protein.

Materials Needed:

• 6 small test tubes
• 50 mL test tube
• Distilled water
• Bradford Dye Solution
• 94 well
• Incubator
• Spectrophotometer

Experimental Procedure:

1. Label 6 Test tubes from S0 to S5.
2. Pipette as follows:

3. Label a test tube as “Bradford Dye.” Pipette 1440 μl of distilled water into the test tube and 360 μl of Bradford Dye solution.
4. In the 94-well, pipette 195 μl of Bradford Dye solution into 10 different wells.
5. In each of the 10 Bradford Dye solutions, pipette 5 μl each of the 6 solutions of S0 to S5 and 4 solutions of plasmid (+) solution and plasmid (-) solution, and plasmid (+) cell and plasmid (-) cell from the sonicated test tubes from the previous agenda.
6. Incubate the 94-well with lid on for 3 minutes at 37 °C.
7. Determine observance of the 94 well using the Spectramax iD3 spectrophotometer, wavelength at 595nm, set at no shake.

Session Agenda #1: DNS Reagent Preparation

Experimental Principle: To determine the ability of the pgxC protein to degrade pectin, the DNS assay is utilized, which determines the concentration of sugar upon sugar degradation.

Materials Needed:

• 3,5-Dinitrosalicylic acid (DNS)
• Sodium potassium tartrate
• Sodium hydroxide
• Distilled water
• Weighing boats
• Test Tube

Experimental Procedure:

1. Label 3 weighing boats each as “DNS,” “SPT,” and “NaOH,” and weigh each component with an electronic scale. Add 0.5g DNS, 0.8g NaOH to the test tube
2. Add 30 mL of distilled water to the same tube, then shake the tube for homogeneity.
3. Add 15 g of sodium potassium tartrate and continue stirring until fully dissolved.
4. Adjust volume to 50 mL with distilled water.

Session Agenda #2: Preparation of Evaluation Points for the DNS Assay

Experimental Principle: To run the DNS assay, there needs to be benchmarking points to draw a standard curve to evaluate the concentration of sugar that occurs when pgxC degrades pectin.

Materials Needed:

• D-galacturonic acid
• Distilled water
• 6 Test tubes

Experimental Procedure:

1. Dissolve 100 mg of D-galactronic acid in 100 mL of distilled water as a stock.
2. Label 6 test tubes based on concentration:

Session Agenda #3: Enzymatic Reaction with Pectin Preparation

Experimental Principle: We prepared conditions for pgxC to react with pectin to find the optimal time of incubation for pgxC to most efficiently degrade pectin molecules.

Materials Needed:

• 8 Test Tubes
• 1% Pectin Solution in Sodium Acetate Buffer (pH 6.0)
• +/- plasmid solutions
• Sterilization Cabinet
• Alcohol Lamp
• Incubator
• Spectrophotometer
• DNS reagent solution from Agenda 1
• 96-well

Experimental Procedure:

1. Label 8 test tubes: 4 plasmid and 4 no plasmid. Each of the four “plasmid” / “no plasmid” is further labelled with “0hr,” “12hr,” “24hr,” and “48hr.”
2. Pipette 4.5mL of 1% pectin solution in sodium acetate buffer (pH 6.0) into all test tubes.
3. Pipette 500 microliters + plasmid solution or - plasmid solution (bacterial supernatant) into the 8 tubes accordingly, referring to the label on each test tube. Perform the pipetting inside the sterilization cabinet with the alcohol lamp.
4. Incubate the test tubes that contain the + plasmid solution and - plasmid solution at 37°C for 0 hours, 12 hours, 24 hours, and 48 hours, referring to the label on test tube.
5. Immediately after each incubation, centrifuge the test tube if needed. Pipette 1 mL of the solution from the test tube into the 96-well, then mix with 500 µL of the DNS reagent solution for the DNS assay. Conduct the DNS assay with a Spectrophotometer.

Session Agenda #1: Fruit Peel Sample Preparation

Experimental Principle: Fruit peels contain pectin (a polysaccharide). Preparing peel extracts creates substrate samples to test enzymatic degradation. Centrifugation separates soluble components from large solid debris.

Materials Needed:

• Fruits: Korean pear, watermelon, banana, kiwi, tomato (x4)
• Balance
• Distilled water
• Blender/grinder
• 50 mL conical tubes
• Centrifuge

Experimental Procedure:

1. Collect peel from each fruit:

• Watermelon: 78.9 g
• Banana: 53.09 g + 50 mL water
• Kiwi: 29.61 g
• Tomato: 42.63 g
• Pear: 52.62 g

2. Add 50 mL of distilled water to each sample.
3. Grind each fruit peel + water until homogenized into sludge.
4. Transfer each slurry into labeled 50 mL conical tubes.
5. Centrifuge tubes at 2000 rpm, 22 °C, for 2 min.
6. After spinning, collect the liquid phase and avoid pellet debris.

Session Agenda #2: Enzyme Reaction Preparation

Experimental Principle: The goal is to test enzyme activity on fruit peel extracts. Adding plasmid solution (+ vs -) allows comparison of enzyme-expressing vs non-expressing samples. DNS assay detects reducing sugars released from polysaccharide breakdown.

Materials Needed:

• Fruit peel supernatants
• 10 small test tubes
• Plasmid solution (enzyme-expressing)
• Plasmid solution (control)
• Micropipette + sterile tips
• Vortex mixer
• DNS assay reagent
• Heat block / Dry bath heat
• Additional 10 small test tubes

Experimental Procedure:

Step A: Reaction Setup

1. Label 10 small test tubes: 2 for each fruit (one “+”, one “-”)
2. Pipette 300 µL fruit peel extract into each tube, avoiding chunks.
3. To “+” tubes, add 200 µL + plasmid solution.
4. To “-” tubes, add 200 µL - plasmid solution.
5. Vortex all tubes for 5 sec.

Set B: DNS Assay Prep

1. Label another 10 small test tubes: 2 for each fruit (one “+”, one “-”)
2. Pipette 50 µL DNS solution into each tube.
3. Add 50 µL of each enzyme reaction sample into the corresponding DNS tube.
4. Vortex all DNS tubes to mix.
5. Dry bath heat all DNS tubes at 100 °C for 5 min.
6. DNS reacts with reducing sugars to produce a colored product.

Session Agenda #3: Absorbance Measurement

Experimental Principle: DNS assay turns orange-red when reducing sugars are present. Measuring absorbance at 540 nm quantifies enzymatic breakdown of fruit peel polysaccharides.

Material Needed:

• DNS assay reaction tubes (from Agenda 2)
• 96-well microplate
• Micropipette + tips
• Plate reader (set to 540 nm)

Experimental Procedure:

1. Label wells on the 96-well plate clearly (fruit name, +/-).
2. Pipette 80 µL of each DNS reaction into its assigned well.
3. Add one well with DNS reagent only (blank/control).
4. Place plate into spectrophotometer/plate reader.
5. Measure absorbance at 540 nm.
6. Record and compare absorbance values (+ vs - plasmid).