Molecular biology

PCR

We performed PCRs to obtain and amplify fragments corresponding to mutated mucA from genomic DNA of P. putida as the template.

Materials:
Template: Genomic DNA from P. putida or P. putida on LB agar plate
Forward and Reverse Primers
Q5 High-Fidelity 2X Master Mix (New England Biolabs)
MiliQ Water

Protocol:

• Dilute a colony ofP. putida from LB agar plate in 70µL of water (genomic DNA).
• In a PCR tube placed on ice (-4°C), add:
• 25 μL Q5 master mix (containing buffers and components required for PCR)
• 2.5 μL Reverse primer
• 2.5 μL Forward primer
• 1µL of genomic DNA (=template)
• Complete with MiliQ water to reach a final volume of 50µL

PCR program with Q5 polymerase:

Cycling Step	Temperature	Time
Cell lysis and initial denaturation	98°C	30s
25-30 cycles Denaturation	98°C	10s
25-30 cycles Hybridation	65°C	30s/kb
25-30 cycles Elongation	72°C	50s
Final elongation	72°C	2min
Hold	4°C	∞

Vector Linearization

We digested our vectors to enable the cloning of mutated mucA and mucA wt. All vector linearizations were performed by digestion with two restriction enzymes to prevent vector re-circularization.

Materials:
Enzyme buffer
Restriction Enzyme 1
Restriction Enzyme 2
Vector for digestion
MiliQ Water

Protocol

• In a 1.5mL tube, add:
• 5 µL enzyme buffer
• 1 µL Restriction Enzyme 1
• 1 µL Restriction Enzyme 2
• 22 µL of the vector to be linearized.
• 18 µL of MiliQ water
• Incubate for one hour at 37°C.

Cloning

We carried out the cloning of mutated mucA into the pKNG101 vector and cloning of mucA wt into the pJN105 vector using the NEBuilder kit from New England Biolabs.

Materials:
Linearized vector
Fragment(s) to be assembled
NEBuilder MasterMix
MiliQ Water

Protocol:

• In a 1.5mL tube, add:
• 2µL of the linearized vector
• 2µL of the fragment(s)
• 10µL of NEBuilder MasterMix
• Complete with MiliQ water to reach a final volume of 20µL
• Incubate for 15 minutes at 50°C.

Competent cell preparation

We transformed E. coli cc118λpir strains with our cloning products to verify the correct insertion of inserts into vectors and amplify cloning products. Prior to transformation, the cells need to be made competent.

Materials:
Cells to be made competent
LB
CaCl2 50mM
Glycerol 15%
Water
Ice

Protocol:

• The day before:
• Inoculate 10mL of LB with 100µL of cc118λpir cells.
• Incubate overnight at 37°C.
• On the day of the procedure:
• Prepare the following solutions:
• 50mL of 50mM CaCl₂ solution (Buffer 1)
• 50mL of 50mM CaCl₂ solution + 15% Glycerol (Buffer 2)
• Filter the solutions and store them at 4°C. They must remain sterile and be used while kept on ice.
• Inoculate 100mL of LB with 1mL of the overnight preculture.
• Incubate at 37°C until the OD600nm reaches between 0.3 to 0.5 (approximately 2 to 3 hours).
• Once the OD600nm is reached, divide the culture into two 50mL tubes, to be kept on ice (-4°C). All the following steps should be performed on ice.
• Centrifuge the culture at 4000 rpm at 4°C for 10 minutes.
• Remove the supernatant and resuspend the pellet in 25mL of Buffer 1.
• Incubate for 20 minutes on ice.
• Centrifuge the culture at 4000 rpm at 4°C for 10 minutes.
• Remove the supernatant and resuspend the pellet in 4mL of Buffer 2.
• Incubate for 20 minutes on ice.
• Prepare 350µL aliquots in 1.5mL Eppendorf tubes and store them at -80°C.

Transformation

We transformed competent E. coli cc118ypir cells with our cloning products (mutated mucA, mucA, alg8). The introduction of the recombinant plasmid is achieved through a heat shock.

Materials:
Competent cells
Cloning product
Ice bath
LB agar plates + antibiotic
Water bath capable of reaching 42°C
Shaking incubator at 37°C

Protocol:

• Prepare an ice bath and a water bath set to 42°C.
• Thaw a tube stored at -80°C containing competent cells in the ice bath.
• In a 2mL tube, add:
• 100µL of competent cc118 λpir cells
• 10µL of the cloning product
• Incubate for 30 minutes in the ice bath (-4°C).
• Incubate for 45 seconds at 42°C.
• Incubate for 10 minutes in the ice bath.
• Add 1mL of LB to the tube.
• Incubate for 1 hour at 37°C under agitation (300rpm).
• During this time, prepare LB agar plates with antibiotic.
• Spread 100µL of the culture onto LB agar plates with antibiotic using a spreader or a bent glass rod.
• Once the plates are dry, incubate the plates overnight at 37°C.

Colony PCR

We conducted colony PCR to verify the insertion of inserts into vectors after cloning mutated mucA and wild type mucA. Colonies containing the recombinant plasmid were selected for sequencing after this step. Once the sequencing is confirmed, the cells carrying the recombinant plasmid will be used to amplify the plasmid for extraction. We performed colony PCR using EconoTaq Plus 2X Green Master Mix by Biosearch technologies.

Materials:
EconoTaq Master Mix 2X
Forward and Reverse Primers
emplate = colony on a plate
LB agar plates + antibiotic containing the plasmid's resistance marker

Protocol:

• Prepare LB agar plates with the appropriate antibiotic containing the plasmid's resistance marker.
• In a PCR tube, add:
• 12.5µL EconoTaq Master Mix 2X
• 0.25µL Forward Primer
• 0.25µL Reverse Primer
• Complete with water to a final volume of 25µL.
• Pick a colony from the plate with transformed colonies using a toothpick and streak onto a new LB agar plate with the appropriate antibiotic.
• Place the toothpick into the PCR tube and scrape it in the tube to transfer cells (= PCR template).
• Thermocycler Program:
• Initial denaturation: 95°C for 2 minutes
• Denaturation: 95°C for 30 seconds
• Hybridation: 55°C for 30 seconds
• Elongation: 72°C for 1 minute
• Final elongation: 72°C for 5 minutes
• Hold: 4°C

Cycling Step	Temperature	Time	Number of cycles
Initial denaturation	94°C	2 min	1
Denaturation	94°C	30 sec	30
Annealing	65°C	30 sec	30
Extension	72°C	1kb/min	30
Final extension	72°C	6min	1
Hold	4°C	∞	1

Agarose gel electrophoresis

We conducted agarose gel electrophoresis to verify the linearization of plasmids and the presence of amplicons at the expected size after PCR/colony PCR.

Materials:
1% Agarose powder
1X TBE buffer
Water

Protocol:

⚠ Caution! TBE buffer is toxic and should be handled with gloves.

• Preparation of Agarose Gel:
• Prepare 100mL of 1X TBE buffer.
• Add 1g of 1% agarose.
• Transfer the solution to a flask and heat it until the agarose melts.
• Pour the solution into an agarose gel mold.
• Allow the gel to polymerize.
• Electrophoresis
• Add the migration buffer (1X TBE) to the electrophoresis.
• Place the gel, still within the mold, into the electrophoresis chamber.
• Using a micropipette, dispense between 5 to 10μL of the sample into the well, depending on the well size. Don't forget the size marker.
• Allow the samples to migrate for 30 minutes at 135 volts.
• Visualize the gels.

Conjugation in Pseudomonas

We introduced the recombinant pKNG101 plasmids through conjugation to integrate them into the chromosome via homologous recombination. We also used conjugation to introduce pNJ105 carrying mucA wt.

Materials:
Donor strain (E. coli cc118ypir)
Recipient strain (P. putida)
Mobilizing strain (E. coli prK2013)
LB agar plates
LB medium
PIA medium

Protocol:

• The Day Before the Procedure:
  • Prepare overnight precultures of the recipient, donor, and mobilizing strains in LB medium.
• On the Day of the Procedure:
  • Spot 25µL of the donor and mobilizing strain precultures on LB agar plate.
  • Incubate for 2 hours at 37°C.
  • During this time, incubate the recipient strain preculture at 42°C.
  • Combine with the recipient strain by spotting 25µL of its preculture onto a plate containing the mobilizing and donor strains.
  • Incubate for 6 hours at 37°C.
  • Resuspend the patch in 1mL of LB and incubate for 1 hour at 37°C with agitation.
  • Select clones on PIA with Streptomycin 2000 μg/ml (3 plates with 75μl/150µL/300µL).
  • Incubate overnight at 37°C.

Recombination in Pseudomonas

We used homologous recombination to replace the native mucA gene in the genome with a mutated mucA gene encoding a truncated MucA. The first homologous recombination event occurs after conjugation. This protocol describes how to proceed with the second homologous recombination event.

Protocol:

• The first day:
• Isolate 6 colonies on 6 plates with LB agar containing 6% sucrose and no NaCl. (It is crucial to perform true isolation to increase the chances of obtaining more isolated clones.)
• Leave the plates at room temperature (RT) for 2-3 days until isolated clones, approximately 50-100, become SucR (sucrose-resistant).
• The next day:
• Re-patch the isolated clones on LB plates, LB plates with 2000 μg/ml Sm (Sm2000), and LB plates with 6% sucrose and no NaCl (LB ssNaCl Suc 6%).
• Incubate at 37°C overnight.
• Perform colony PCR to screen for the recombination event that occurred.

Production and testing of alginate beads

Making alginate beads

In order to optimize the characteristics (component concentration, size, etc.) of our alginate beads to achieve the best water retention possible for the intended application, we created alginate beads using commercially available sodium alginate (ThermoFisher) for various testing purposes.

Materials:
Sodium alginate
CaCl2
Water
Pasteur pipette or bead mold provided by the team

Protocol:

Initially, we created the beads as follows:

• Dissolve sodium alginate in water with vigorous stirring to achieve a 1% concentration. Continue stirring until the solution is completely homogeneous.
• Prepare a 0.5 mM CaCl₂ solution in a beaker.
• Using a Pasteur pipette, draw up the sodium alginate solution and drop it into the beaker containing the CaCl₂ solution. The beads will form immediately.
• Remove the beads using forceps or a small scoop.

Subsequently, we devised a mold to obtain standardized bead sizes, with details provided in the wiki's contribution section. The protocol for using the mold is as follows:

• Prepare solutions of 1% sodium alginate and 1M CaCl₂.
• Secure the two parts of the mold using elastic bands or a clamp (preferable).
• Using a micropipette (p1000), draw up 300 µL of 0.5 mM CaCl₂ and dispense it into one of the mold's holes.
• Draw up 600 µL of 1% sodium alginate and dispense it into the second hole. Be sure to remove any air bubbles beforehand, as the presence of air may affect bead formation. ⚠ It is essential not to use the same holes for both solutions, as this will prevent bead formation.
• Separate the two mold pieces and retrieve the beads.

Water retention test by alginate beads

Initially, we pondered on how to demonstrate the water retention capability of alginate beads (alginate 1%; [CaCl₂] 1M). Therefore, we devised the following protocol, which we subsequently tested:

• Prepare alginate beads using the dedicated protocol.
• In a Petri dish, add 2 mL of water (condition 1).
• In another Petri dish, add 5 g of soil (condition 2).
• In a third Petri dish, add 5 g of soil + 2 mL of water (condition 3).
• Add 2 g of beads with the same alginate and CaCl₂ concentration to each of the Petri dishes.
• Incubate the Petri dishes without lids at 30°C.
• Weigh the Petri dishes every two hours.

Calculation mass of water in the balls: Allow the beads to dry for several days and record the weight to determine the dry weight of the beads. Subtract the original weight from the dry weight.

Optimal Concentration Testing of Alginate and CaCl₂

We sought to determine the optimal concentrations of alginate and CaCl₂ for achieving the best water retention in our beads. To assess water retention, we conducted experiments using beads with varying concentrations of alginate and CaCl₂, following this protocol:

• Prepare beads with different concentrations of alginate and CaCl₂. We prepared beads at either 0.25% 0.5%, 0.75% or 1% alginate concentration and 0.1M, 0.2M, 0.4M, 0.5M , 0.8M, 1M, 2M, 4M CaCl₂ concentration.
• In a Petri dish, add 2 mL of water (condition 1).
• In another Petri dish, add 5 g of soil (condition 2).
• In a third Petri dish, add 5 g of soil + 2 mL of water (condition 3).
• Add 2 g of beads with the same alginate and CaCl₂ concentration to each of the Petri dishes.
• Incubate the Petri dishes without lids at 30°C.
• Weigh the Petri dishes every two hours.

The best performing beads will be those that have lost water the slowest.

Calculation mass of water in the balls: Allow the beads to dry for several days and record the weight to determine the dry weight of the beads. Subtract the original weight from the dry weight.

Water Retention Tests Based on the Quantity of Beads in Relation to the Amount of Soil

Subsequently, we explored the amount of beads (alginate 1%; [CaCl₂] 1M) required in relation to the quantity of soil to achieve optimal water retention. To address this issue, we implemented the following protocol:

• Prepare alginate beads using the dedicated protocol.
• In a Petri dish, add 2 mL of water (condition 1).
• In another Petri dish, add 5 g of soil (condition 2).
• For each of the conditions, conduct tests in multiple Petri dishes with varying amounts of beads. We tested the following quantities: 10%, 20%, 30%, 40%, and 50% (Example: 10% = 5 g of soil + 0.5 g of beads).
• Weigh the Petri dishes every hour for a duration of 4 hours.

From the result in condition 3, substract what you got in condition 1 and 2 to control for the different parameters

Testing Alginate Production in P. putida:

Alginate Assay

Materials:
LB medium
Water
Alginate 0.5 mg/mL
96% Ethanol
Sulfuric acid
Ice
P. putida cells

Sample Preparation for Quantification:

• Start with a bacterial culture in 2 mL of LB (in a Greiner tube) that was inoculated with a single colony and incubated overnight at 30°C.
• Centrifuge the culture at 5,000 rpm for 10 minutes.
• Carefully collect 1 mL of the supernatant and transfer it to a 2 mL Eppendorf tube. Keep this tube on ice.
• Add 1 mL of cold (-20°C) 100% isopropanol to the Eppendorf tube.
• Centrifuge the Eppendorf tube at 14,000 rpm for 30 minutes.
• Discard the supernatant and retain the pellet.
• Resuspend the pellet in 1 mL of distilled water. Use vortexing and sonication if necessary to ensure complete resuspension.

Preparation of the Reference Range:

• Prepare a stock solution of alginate at 0.5 mg/mL in water. Use vortexing and sonication if necessary to ensure complete dissolution.
• Prepare dilutions as follows:

	1	2	3	4	5	6	7	8	9
[Alginate] (µg/mL)	500	400	350	300	250	200	150	100	50
Alginate (µg)	15	12	10.5	9	7.5	6	4.5	3	1.5
Vi (µL)	200	160.0	140.0	120.0	100.0	80.0	60.0	40.0	20.0
VH20 (µL)	0	40.0	60.0	80.0	100.0	120.0	140.0	160.0	180.0

Preparation of Solutions:

⚠ To be done in a fume hood:

• Stock Solution:
• Sulfuric Acid (H2SO4) – Pure
• Ethanol (96%)
• Borate Solution: Dissolve 24.74g of H3BO3 in a 4M KOH solution.
• Solutions to Prepare Before Assay:
• Borate-Sulfuric Acid 0.1M: Dilute the borate solution with pure H2SO4 at a 1/40 ratio (40% for 10 mL, which equals 250 µL).
• Carbazole 0.1%: Weigh and dissolve the carbazole in ethanol. For 44 wells, prepare 10 mL, which requires 10 mg of carbazole.

Procedure for Sample Preparation and Measurement:

• Place the 96-well plate on an ice box.
• Add 200 µL of Borate-sulfuric acid 0.1M to each well. Allow it to cool for a few minutes.
• Add 30 µL of the sample to be quantified to each well. Let it cool for a few minutes.
• Add 20 µL of 0.1% carbazole solution to each well. Vortex the carbazole if necessary and perform two pipetting cycles in the wells.
• Place the plate in an incubator at 55°C for 30 minutes. The samples will turn violet during this step.
• After incubation, measure the absorbance of the samples at 530nm using a Tecan plate reader. The reading should be done in two rounds with the Alginate CK program.
• The coloration will remain stable for approximately 2 hours after measurement.

Culture under Water Stress

Since P. putida produces alginate as a component of its biofilm during membrane stress, resulting in a mucoid phenotype on agar plates, we aimed to test the appearance of this phenotype under water stress. We cultivated the bacterium in LB medium supplemented with different concentrations of NaCl before quantifying the alginate produced under these conditions.

Materials:
LB medium
Diluted NaCl solution
LB agar medium
P. putida cells

Protocol:

• Inoculate 5 to 6 colonies of P. putida in 5 mL of LB.
• Incubate at 30°C with shaking at 180 rpm.
• Measure the OD_600nm, at the stationary phase, take 20 µL of the culture and place the drop on an LBa dish plate supplemented or not with NaCl:
• LB agar
• LB agar + NaCl 0,1M
• LB agar + NaCl 0,3M
• LB agar + NaCl 0,5M
• Streak the liquid drop on the plates and incubate at 30° overnight.
• The next morning, take pictures of the plates.

Monitoring the growth of P. putida in a glycerol/LB media

Materials:
LB medium
Glycerol
P. putida cells

Protocol:

• The day before:
• Make a pre-culture of P. putida: one colony in 10 mL of LB.
• The next day:
• Inoculate 100 µL of the preculture in 10 mL of the different media: Gly 40%, 20%, 10%, 5% and LB.
• Place erlens at 30° with agitation.
• Take the OD every hour until as late as possible in the day. ⚠ Make the blank with the corresponding media
• Repeat OD the next day and continue measurements if necessary.
• Make curves with results and calculate generation time.

NB : the medium must be sterile to be sure to detect only the P. putida’s growth.

Building alginate plates

Here is how we built our alginate plates for our collaboration with Jungle.bio.

Alginate plate creation protocol:

Material

• Preferably a box for casting PCR gels (easier to handle), otherwise a rectangular box.
• a 20ml syringe
• 100ml 1% alginate
• 100ml 1M CaCl2

Note: The PCR gel casting box has detachable parts, making it quick and easy to make alginate plates. Prefer this or similar systems.

Protocol

• Prepare the dish for casting PCR gels.
• Using a syringe, draw up some alginate and place it in a very thin layer on all edges of the dish.
• Then deposit the alginate in the rest of the dish in a regular back-and-forth motion. Don't hesitate to use the syringe (without pouring alginate) to distribute the alginate in any holes.

Note: the layer should be as thin as possible, so don't apply too much alginate. For a 6cm by 13 plate, alginate should be removed 3 times.

Leave to stand for 30 minutes to 1 hour, to allow the alginate to spread into as smooth a surface as possible. During this time, clean the syringe with water. If alginate remains in the syringe, use 96% ethanol.

Draw up CaCl₂ and drop into the center of the dish. Once a circle has formed, spread it over the entire dish. Once this is done, add more CaCl₂. The edges of the alginate should shrink and the whole plate will begin to float.

Recover the plate and place in a suitable container with water to prevent drying out. Excess CaCl₂ in the dish can be used to make other plates.

P.S: after a week in the open air, the plates start to contain contamination. It's best to store them in a fridge.