Notebook CIRI

August 4th :

Growth monitoring of Labrys Portucalensis to identify conditions and time points for RNA-seq

The objective is to generate growth curves in different media in order to identify the time points at which the culture enters the exponential phase and the stationary phase. The purpose of testing various media is to determine the least rich medium in which the bacterium can still grow normally. Using a medium that is not too rich helps avoid potential interferences between PFAS and nutrient-rich media during cultivation with PFAS, thereby allowing us to capture a transcriptomic response of the bacterium grown in the presence of PFAS. Inoculation of 4 mL TSB with different dilutions (1/1, 1/2, 1/5, 1/10, 1/20, 1/50, 1/100) using colonies of Larbys portuclanesis F11 spread on TSB agar plates by our PI. Cultures incubated at 30 °C with shaking (since the optimal growth temperature of L.Portucalensis is around 30°C). OD600 measurements were taken manually over several days, according to our availability. (Below are the results after 172 hours of monitoring)

Figure 1 : Growth curve Labrys portucalensis F11 in different TSB dilutions

The objective was to clone PFOA-responsive promoters identified in an RNA-seq study on E. coli exposed to PFOA, together with the lux operon genes split into two parts: luxA and luxB under promoter 1, and luxC, luxD, and luxE under promoter 2. The separation of the operon would enable a better specificity of reaction since both of them should be expressed for bacteria to emit luminescence. Before testing with PFAS-responsive promoters, we aimed to use inducible promoters as positive controls: Plac-LacI and TetR-Ptet. This would allow us to verify the function of the operon and assure us to see the signal. Three DNA fragments were ordered for insertion into pSEVA261 via Gibson Assembly. Fragments were designed with appropriate homologous sequences to enable assembly. Below are the three ordered fragments, the linearized plasmid, and the theoretical Gibson Assembly product. Illustration of the inserts and linearized pSEVA261 plasmid :

Plasmid cloning with PFOA-sensitive promoters or inducible promoters

The objective was to clone PFOA-responsive promoters identified in an RNA-seq study on E. coli exposed to PFOA, together with the lux operon genes split into two parts: luxA and luxB under promoter 1, and luxC, luxD, and luxE under promoter 2. The separation of the operon would enable a better specificity of reaction since both of them should be expressed for bacteria to emit luminescence. Before testing with PFAS-responsive promoters, we aimed to use inducible promoters as positive controls: Plac-LacI and TetR-Ptet. This would allow us to verify the function of the operon and assure us to see the signal. Three DNA fragments were ordered for insertion into pSEVA261 via Gibson Assembly. Fragments were designed with appropriate homologous sequences to enable assembly. Below are the three ordered fragments, the linearized plasmid, and the theoretical Gibson Assembly product. Illustration of the inserts and linearized pSEVA261 plasmid :

Figure 2 : Components of the construction for the inducible promoter plasmid.

A: First insert containing homology regions with the linearized plasmid and the second insert.

B: Second insert containing homology regions with the first and third inserts.

C: Third insert containing homology regions with the linearized plasmid and the second insert.

D: Linearized plasmid obtained after PCR linearization.

Figure 3 : Illustration of the theoretical Gibson assembly result

The planned strategy was to open this plasmid by digestion between the regions indicated as “restriction sites” in yellow on the plasmid map, and to clone in the Gibson fragments containing PFAS-sensitive promoters at 6h and 24h after PFOA exposure identified in the literature. These promoters were positioned in the opposite orientation so that they would drive the expression of the two constitutive operons of the biosensor :

Figure 4 : Inserts of the PFOA sensitive promoters in reverse orientation

A: First insert containing 2 promoter sensitive of PFOA concentration at 6h after exposure

B: Second insert containing 2 promoter sensitive of PFOA concentration at 24h after

Figure 5 : Illustration of digestion linearization to include PFOA sensitves promoters

Gibson cloning with the inserts containing the PFAS-sensitive promoters (in gray and purple) to generate the base plasmid for our biosensor, which is expected to express luciferase and emit light in the presence of PFOA.

Figure 6 : Illustration of the theoretical Gibson assembly result with PFOA sensitive promoters (grey and purple)

Linearization of pSEVA261 plasmid by high-fidelity PCR

PCR mix (final volume: 20 µL):

• 8 µL diluted pSEVA (including water to reach final volume)
• 1 µL primer P1 (10 µM)
• 1 µL primer P2 (10 µM)
• 10 µL Q5 2X Master Mix

PCR cycle :

• Expected fragment size: 2251 bp
• Extension rate: 20–30 s per kb → elongation time set to 1 min (sufficient)
• Tm calculated with NEB Tm calculator (link): ~63 °C with Q5 buffer
• P1: 62 °C
• P2: 72 °C
• Despite the difference, PCR performed with these values.

Gel preparation (1% agarose in 0.5X TAE):

• Stock solution prepared: 300 mL of 0.5X TAE with 1% agarose (stored, to be remelted in microwave when needed, BET room 2nd floor).
• BET added on gel rack before pouring molten agarose to reach final concentration 0.5 µg/mL.

Gel electrophoresis of PCR amplification

• Gel placed in tank with 0.5X TAE.
• Loaded 4 µL of plasmid amplification product.
• Migration: 25 min at 135 V.
• Visualization under UV.

Figure 7 : gel migration electrophoresis of the PCR linearized plasmid

August 5th :

Growth monitoring of Labrys portucalensis to identify sampling points for RNA-seq

Cultures monitored over time (OD600 measurements) to determine phases of interest for RNA extraction.

Plasmid cloning with PFOA-sensitive promoters or inducible promoters

DpnI digestion of the plasmid amplification

• Digestion with DpnI to degrade methylated (bacterial template) DNA and avoid transformation by residual initial circular plasmid.
• PCR product: ~17 µL (20 µL – 3 µL used for gel).
• Added 2 µL 10X Smart Buffer + 0.5 µL DpnI.
• Incubation: 30 min at 37 °C.

Clean-up (cf. Experiment : Monarch PCR & DNA Clean-up Kit).

• Final elution volume: 20 µL.
• Nanodrop: 25.4 ng/µL.

Note:

clean-up removes free nucleotides, confirming presence of linearized plasmid. Concentration higher than expected from residual template → PCR likely successful.

Gibson Assembly

• Equimolar calculation for 100 ng final plasmid:
• Insert 1: (2150 / 11,771) × 100 = 18.3 ng
• Insert 2: (3739 / 11,771) × 100 = 31.8 ng
• Insert 3: (3755 / 11,771) × 100 = 31.7 ng
• Linearized pSEVA261 vector: (2251 / 11,771) × 100 = 19.1 ng
• Target resuspension concentrations:
• Inserts: 40 ng/µL
• PCR plasmid pSEVA261: 25.4 ng/µL (from clean-up)
• Resuspension volumes from Twist insert delivery:
• Insert 1: 803 ng → 20 µL
• Insert 2: 601 ng → 15 µL
• Insert 3: 969 ng → 24.2 µL
• Final: inserts at 40 ng/µL, plasmid at 25.4 ng/µL. Volumes for Gibson assembly calculated to achieve target masses above.
• Gibson Assembly Mix:
• Insert 1: 0.46 µL
• Insert 2: 0.80 µL
• Insert 3: 0.79 µL
• pSEVA261 plasmid: 0.75 µL
• NEBuilder HiFi Assembly Mix: 2.8 µL (same total volume as inserts + vector)
• Incubation: 30 min at 50 °C

E.coli DHAα Transformation

• Thaw competent Heat Shock E.coli DHAα on ice.
• Add 3 µL Gibson assembly product, incubate 10 min on ice.
• Heat shock: 42 °C, 1 min, then 10 min on ice.
• Recovery: 1 mL LB, 30–60 min, allows expression of resistance cassette.

Plating

• Spin: 4,000 g, 2.5 min.
• Remove supernatant, leave ~100 µL, resuspend pellet.
• Plate on kanamycin agar: spread ~150 µL on half the plate with a spreader, then use same spreader for other half.
• Incubate overnight at 37 °C.

August 6th :

Colony observation

~200 colonies visible.

Figure 8 : Observation of the petri dish containing colonies of transformed DH5α

Need to verify plasmid identity and functionality.

Verification options (complementary):

1. PCR + gel electrophoresis to check fragment size → not suitable here (at that point we did not have appropriate primers to test our construct yet).
2. Colorimetric test and fluorescence test / TECAN measurement with IPTG and ATC induction → chosen method.
3. Miniprep + sequencing → to be done on promising colonies.

Culture setup

• 200 µL LB + 1X kanamycin in 12 wells of a 96-well plate.
• Inoculate 12 selected colonies using sterile pipette tip, under Bunsen flame.

Incubation

• 1 h at 37 °C with shaking.

Induction

• Split cultures: 2 × 100 µL.
• Add 4 µL of 50X IPTG + ATC solution.

Note:

stock 2× too concentrated → final IPTG 1 mM, ATC 2 µM; intended: 500 µM IPTG, 1 µM ATC

Induction incubation

• 1 h at 37 °C.

TECAN measurement

• No fluorescence nor luminescence detected.
• Likely Gibson cloning failure.
• Check again 24 h later → still negative.

Next steps

1. Start pre-culture of colony n°1, 2, 3, 4 for miniprep and sequencing to analyze the cause.
2. Potential problem: previous PCR quality low.
3. Try again PCR in order to perform Gibson assembly and E.coli DHAα transformation

PCR retry setup

• Dilute primers to 10 µM.
• Dilute pSEVA261 plasmid to 0.3 ng/µL (10× dilution from 3 ng/µL).
• PCR mix (20 µL):
• 1 µL each primer
• 8 µL plasmid (0.3 ng/µL → 2.4 ng total)
• 10 µL Q5 2X Master Mix
• Cycle parameters: same as previous PCR.

August 7th :

Re do Cloning and transformation Gel electrophoresis of PCR amplification of 6th of August

• ~20 mL agarose 1% gel, add 4 µL BET (for visualization of PCR migration).
• Gel placed in tank with 0.5X TAE.
• Loaded 4 µL of linearized plasmid amplification product.
• Migration: 25 min at 135 V.
• Visualization under UV.

Figure 9 : Observation of the petri dish containing colonies of transformed DH5α

Gibson Assembly

• Mix:
• Insert 1: 0.46 µL
• Insert 2: 0.80 µL
• Insert 3: 0.79 µL
• pSEVA261 plasmid: 0.75 µL
• NEBuilder HiFi Assembly Mix: 2.8 µL (same total volume as inserts + vector since it is a 2X solution)
• Incubation: 30 min at 50 °C

E.coli DHAα Transformation (Transformation n°2)

• Thaw competent Heat Shock E.coli DHAα on ice.
• Add 3 µL Gibson assembly product, incubate 10 min on ice.
• Heat shock: 42 °C, 1 min, then 10 min on ice.
• Recovery: 1 mL LB, 30–60 min, allows expression of resistance cassette.

Plating

• Spin: 4,000 g, 2.5 min.
• Remove supernatant, leave ~100 µL, resuspend pellet.
• Plate on kanamycin agar: spread ~150 µL on half the plate with a spreader, then use same spreader for other half.
• Incubate overnight at 37 °C.

Plasmid purification and sequencing

1. Plasmid from transformation was purified by miniprep for sequencing.
2. Pellet culture: Spin down overnight culture from the colony n° 1, 2, 3, 4: 5 min at 13 rpm speed. Pellet to be used for miniprep of a transformant colony.
3. Miniprep: Follow NEB Miniprep kit protocol (cf : Experiment : Plasmid miniprep). Final elution: 30 µL.
4. Nanodrop measurement: 33.8 ng/µL (1 µL used for measurement). Total DNA obtained: ~980 ng.
5. Plasmid purified by miniprep was sent to Microsynth for sequencing.

August 8th :

Cloning and transformation n°2

Colony observation

• 14 colonies visible.

Culture setup to induce promoter and measure reporter gene activity

• 200 µL LB + 1X kanamycin in 12 wells of a 96-well plate.
• Inoculate 12 selected colonies using sterile pipette tip, under Biological Safety Cabinet.

Incubation

• 1 h at 37 °C with shaking.

Induction

• Split cultures: 2 × 100 µL.
• Add 2 µL of 50X IPTG + ATC solution (1µM)

Induction incubation

• 1 h at 37 °C.

TECAN measurement

• No fluorescence nor luminescence detected.
• Likely Gibson cloning failure.
• Check again 24 h later → still negative.

August 11th :

Growth monitoring of Labrys portucalensis to identify sampling points for RNA-seq

The Labrys portucalensis culture reached the stationary phase :

Figure 1 : Growth curve Labrys portucalensis F11 in different TSB dilutions

We identified that the 1:5 TSB dilution is the highest dilution allowing proper growth of Labrys portucalensis F11. The selected time points for RNA-seq sampling are 12 h and 24 h. We will start the growth of Labrys portucalensis with different concentrations of PFOA and TFA the next day, and perform sampling at 12 h and 24 h after the culture is initiated. 2 measures at 168h and 172h were performed to ensure the stationary phase was truly reached.

Plasmid cloning with PFOA-sensitive promoters or inducible promoters

Plasmid sequencing result and alignment with the desired plasmid :

Figure 10 : Illustration of the plasmid alignment on the expected sequence

Colonies 1, 2, 3, and 4 from the first transformation only integrated the template plasmid, not the cloned construct. Several hypotheses could explain this outcome: insufficient Gibson reaction time, incomplete DpnI digestion, plasmid size being too large for efficient transformation, or a generally low transformation efficiency.

We ordered primers to perform colony PCRs in order to quickly test the transformants and screen for those that have integrated the correct plasmid. The amplification overlaps the 3 inserts:

Figure 11 : Illustration of the plasmid PCR tested to validate the nexttransformation

Primer forward (5'-3'): GAGAAAATTGGGGAGGTTGG (50% GC, Tm : 57°C, length : 20pb)

Primer reverse (5'-3') : AAGATTTCAACCTGGCCGG (53% GC, Tm : 58°C, length : 19pb)

Inoculation and miniprep

Inoculation of colonies no. 1, 2, and 4 (from the 14 colonies obtained after transformation n°2) into 4 mL of LB supplemented with 4 µL of kanamycin. These cultures will be used to perform a miniprep and subsequently sent for sequencing.

August 12th :

Culture of Labrys portucalensis under PFAS and control conditions

Objective

To grow Labrys portucalensis cultures under different PFAS concentrations for transcriptomic analysis.

Culture setup

• Starting culture: preculture at OD600 = 1.9
• Inoculated 3 mL cultures under the following 5 conditions, each in 4 replicates:
• 100 µM PFOA
• 0.24 nM PFOA
• 100 µM TFA
• 0.87 nM TFA
• Water (control)
• Cultures grown in 15 mL screw-cap Falcon tubes (safety measure due to high toxicity of PFOA and TFA).
• Incubation: 30 °C, shaking.

August 13th :

Culture of Labrys portucalensis under PFAS and control conditions

Second sampling

• 12 h after inoculation, the second sampling was performed.
• A volume corresponding to 7.5 × 10⁸ bacterial cells was collected.
• Add 2 volumes of RNAprotect® Bacteria Reagent (Qiagen) to the sample.
• Incubate for 5 minutes at room temperature.
• Centrifuge for 10 minutes at 5,000 g.

Supernatant removal

• Carefully decant the supernatant.
• Gently dab the inverted tube once onto a paper towel to remove residual liquid.
• Important: Do not attempt to remove the remaining supernatant by pipetting, as this may result in pellet loss. The residual volume should not exceed ~80 µL.

Storage

• Store the samples at –80 °C until refrigerated transport to the sequencing facility.
• The transporter picked up the sample a few hours after the 2nd sampling.

Plasmid cloning with PFOA-sensitive promoters or inducible promoters

Colony PCR

Procedure

• Collected colonies no. 1–7 using the tip of a pipette cone.
• Suspended each colony in 50 µL of ultrapure water in a PCR tube (1 tube per colony).

Colony PCR premix (for 9 reactions)

• 1.125 µL primer FW (Gibsoncheckfw, 100 µM)
• 1.125 µL primer RV (Gibsoncheckrv, 100 µM)
• 56.25 µL DreamTaq mix

PCR setup

• In new PCR tubes, added 6.5 µL of premix + 6 µL of colony suspension → final volume of 12.5 µL per tube.
• Total: 7 PCR tubes
• Tubes 1–7: colonies 1–7

Next steps

• Quick spin of the PCR tubes before starting the PCR program.

PCR program

• Launched PCR with 35 cycles.
• Expected amplicon size: 3847 bp.
• DreamTaq elongation rate: ~1 kb/min.

Gel electrophoresis of PCR amplification

• Pour ~20 mL agarose 1% gel, add 4 µL BET (for visualization of PCR migration).
• Gel placed in tank with 0.5X TAE.
• Loaded 4 µL of each colony PCR amplification product.
• Migration: 20 min at 135 V.
• Visualization under UV.

Figure 12 : UV-visualisation of the gel after electrophoresis in order to verify the transformation

We decided to adjust the protocol in a few aspects: doubling the DpnI digestion time, doubling the Gibson incubation time, and performing the plasmid linearization by PCR using only 1:100 of the initially used amount.

Plasmid cloning with PFOA-sensitive promoters or inducible promoters : third attempt

PCR to linearize and amplify pSEVA261 plasmid

• Prepared 2 identical tubes to ensure enough material for subsequent cloning.

Reaction mix

• 8 µL diluted pSEVA261 diluted 1/100 (~8 ng)
• 1 µL primer P1 (10 µM)
• 1 µL primer P2 (10 µM)
• 10 µL Q5 2X Master Mix

PCR program

• Annealing temperature: 60 °C
• 35 cycles
• Elongation: 1 min 30 s (Q5 polymerase elongates ~1 kb per 30 s)

Gel electrophoresis of PCR amplification

• Pour ~20 mL agarose 1% gel, add 4 µL BET (for visualization of PCR migration).
• Gel placed in tank with 0.5X TAE.
• Loaded 4 µL of each PCR amplification product.
• Migration: 20 min at 135 V.
• Visualization under UV.

Figure 13 : UV-visualisation of the gel after electrophoresis in order to verify the PCR

DpnI digestion to remove DNA template

• Pooled PCR tubes → total volume 35 µL.
• Took 3 µL and transferred into a separate tube → this sample will not undergo DpnI treatment (tube labeled NON-DPN1).
• To the remaining 32 µL of PCR product, added:
• 3.5 µL DpnI buffer (10X)
• 1 µL DpnI enzyme
• Tube labeled DIGESTION DPN1.
• Incubated DIGESTION DPN1 in thermocycler at 37 °C for 60 min.

Gel electrophoresis of DpnI digestion to test if DpnI digests the amplicons

• Pour ~20 mL agarose 1% gel, add 4 µL BET (for visualization of PCR migration).
• Gel placed in tank with 0.5X TAE.
• Loaded 4 µL of each DpnI digestion product.
• Migration: 22 min at 135 V.
• Visualization under UV.

Figure 14 : UV-visualisation of the gel after electrophoresis in order to verify the digestion with DPN1

DpnI digestion does not digest our amplification. We can not assess if it actually digests the DNA template as we did not have enough DNA template amount.

Clean-up of the sample digested with DpnI

• Protocol: Monarch PCR & DNA Clean-up Kit.
• Final elution volume: 20 µL.
• Nanodrop: 266 ng/µL.
• 1:10 Dilution for final sample (26.6 ng/µL).

Gibson Assembly preparation

Gibson assembly is performed in a final volume of 5 µL, so all volumes suggested on NEBuilder Calculator are divided by 4.

Gibson Assembly reaction

• 0.675 µL of vector at 26 ng/µL
• 0.425 µL of insert 1 at 40 ng/µL
• 0.725 µL of insert 2 at 40 ng/µL
• 0.725 µL of insert 3 at 40 ng/µL
• 2.5 µL of NEBuilder HiFi DNA Assembly Master Mix

Incubated the sample at 50 °C for 60 min in a thermocycler, then stored at -20°C before transformation.

Transformation of DH5α with Gibson reaction and linearized plasmid

• Retrieved 2 tubes of 100 µL competent DH5α cells from -80°C, transported and kept on ice.
• Once thawed, under BSL-2 conditions:
• Added 4 µL of Gibson reaction mix to the first tube of bacteria.
• Added 0.540 µL of linearized plasmid digested by DpnI (26 ng/µL) to the second tube (equivalent to the amount of plasmid in 4 µL of the Gibson mix, for a control of the quantity of the circular plasmid able to transform the bacteria in the mix).
• Incubated on ice for 45 min.
• Heat shock: 30 - 40 sec at 42°C in thermomixer.
• Placed samples on ice for 10 min.
• Added 1 mL LB to each tube under BSL2.
• Incubated Eppendorf tubes in incubator for 1 h.
• After 1 h, centrifuged 5 min at 4000 g, discarded 900 µL of supernatant, and resuspended pellet in remaining 200 µL.

Plating on LB+Kan plates

• Deposited 200 µL on one side of the plate.
• Spread with a sterile loop across half the plate, then streaked from this half to the other half.
• Repeated for both samples: bacteria transformed with linearized plasmid (background control) and bacteria transformed with Gibson reaction mix.

August 14th :

Figure 15 : Observation of the petri dish containing colonies of transformed DH5α

We obtained a large number of colonies. However, there are also colonies on the plates transformed with the linearized plasmid. This is surprising because it was digested with DpnI (if it hadn’t been digested, it would be normal to see so many transformants). This might be due to the original plasmid, but the number still seems quite high.

Colony PCR for colonies n° 1–11 from bacteria transformed by the Gibson product

Procedure

• Collected colonies no. 1–11 using the tip of a pipette cone.
• Suspended each colony in 50 µL of ultrapure water in a PCR tube (1 tube per colony).

Colony PCR premix (for 12 reactions)

• 1.5 µL primer FW (Gibsoncheckfw, 100 µM)
• 1.5 µL primer RV (Gibsoncheckrv, 100 µM)
• 75 µL DreamTaq mix

PCR setup

• In new PCR tubes, added 6.5 µL of premix + 6 µL of colony suspension → final volume of 12.5 µL per tube.
• Total: 11 PCR tubes
• Tubes 1–11: colonies 1–11

Next steps

• Quick spin of the PCR tubes before starting the PCR program.

PCR program

• Launched PCR with 35 cycles.
• Expected amplicon size: 3847 bp.
• DreamTaq elongation rate: ~1 kb/min.

Gel electrophoresis of PCR amplification

• For ~20 mL agarose 1% gel, add 4 µL BET (for visualization of PCR migration).
• Gel placed in tank with 0.5X TAE.
• Loaded 4 µL of each colony PCR amplification product.
• Migration: 20 min at 135 V.
• Visualization under UV.

Figure 16 : UV-visualisation of the gel after electrophoresis in order to verify the PCR

On this gel, most colonies show faint bands of various sizes, which appear to be non-specific bands of unclear origin. Colony 11 displays a band at the expected size, between 3 and 4 kb.

To test whether some of these colonies — particularly colony 11 — successfully integrated the plasmid, we decided to start a culture in a 96-well plate until the bacteria reach the exponential growth phase, followed by induction with IPTG and anhydrotetracycline.

Inoculation of 96-well plate (LB + Kan) with bacteria transformed either with the Gibson product or with the plasmid linearized

Procedure:

• Inoculated 200µL LB + Kan medium in a 96-well plate (black-bordered).
• Green wells: colonies from the plate labeled “Gibson” (= colonies transformed by Gibson cloning).
• Red wells: colonies from the plate labeled “Plasmid” (= colonies transformed with the linearized empty plasmid, which likely re-ligated, or possibly carried over template DNA).
• Placed the plate in a shaking incubator at 37 °C.
• Covered the wells with a sealing film to prevent cross-contamination (photo).

Figure 17 : 96-well plate layout

Induction and 96-well plate layout

Measured OD of the cultures → values between 0.5 and 0.7. Decided to induce.

Prepared two Falcon tubes

• 6 mL LB + 6 µL Kan + 6 µL IPTG (500mM) + 2.6 µL ATC (2 mg/mL)
• 6 mL LB + 6 µL Kan (no inducers).

Inoculation in 96-well plate

• Added 195 µL of medium per well.
• Inoculated with 5 µL of each culture (final dilution 1/40 instead of 1/100).

Plate layout

• Green & Red wells: initial LB + Kan cultures that reached OD 0.5–0.7.
• Yellow wells: cultures with inducers.
• Purple wells: cultures without inducers.

Next step: Incubate for 2 h before taking measurements.

After several measurements, no fluorescence or luminescence values were detected.

Given the difficulties we are facing in cloning this plasmid, we decided to work with a simpler plasmid containing only a single promoter, and then gradually increase the complexity of the construct step by step.

August 15th :

We started from a low-intermediate copy (p15A ORI ~ 10-20 copies) plasmid that already contained LacI and pLac to express an RFP. Below is the plasmid map:

Figure 18 : Illustration of the plasmid used to test the RFP with an inductible promotor

At the same time, we ordered primers to amplify and linearize the plasmid while excluding LacI, pLac, and the Lac operator. We also ordered additional primers to specifically amplify the PFAS-sensitive promoters (those sensible at 24h PFOA exposure) that were present in the initially ordered inserts (see map from August 4), designed with homology regions to match the linearized plasmid.

Plasmid linearization :

Figure 19 : Illustration of the plasmid used to test the RFP with an inductible promotor

Figure 20 : Illustration of the linearised plasmid

Promoter b3021_24h amplification and addition of plasmid homology region (we did exactly the same thing with the b0002_24h promoter) :

Figure 21 : Illustration of the PCR made on the fragment previously used to isolate and amplify the promoter

Figure 22 : Illustration of the amplified promoter for Gibson

Figure 23 : Illustration of the plasmid PCR tested to validate the next transformation

Primers ordered for the construction

• primer “IGEM1” to linearize the plasmid
• primer “IGEM2” to linearize the plasmid
• primer “b0002RV” to amplify and add homology sequence in b0002_24h promoter
• primer “b0002FW” to amplify and add homology sequence in b0002_24h promoter
• primer “b3021RV” to amplify and add homology sequence in b3021_24h promoter
• primer “b3021FW” to amplify and add homology sequence in b3021_24h promoter

Our supervisor already had frozen bacteria transformed with this plasmid, and we cultured them in 3mL LB containing chloramphenicol 1X.

Culture and induction of MG1655 containing the plasmid

• Launched a culture of MG1655 bacteria carrying “François’ plasmid”.
• Inoculated 196 µL of LB + chloramphenicol (1X) with 4 µL of stationary phase culture in a 96-well plate (black-bordered).
• Agitated for approximately 30 min.
• Added IPTG to reach a final concentration of 0.5 mM in each well.

Plate layout

• Wells 1–4: induced
• Wells 5–7: non-induced
• Well 8: LB only (no untransformed bacteria available at that time)

August 18th :

We started from a plasmid that already contained LacI and pLac to express an RFP. Below is the plasmid map:

August 19th :

Cloning and transformation for biosensor

Performed PCR on the inserts to amplify the promoters, and colony PCR to amplify and linearize the plasmid contained within the bacteria.

PCR mix (final volume: 20 µL) for plasmid linearization:

• 8 µL of solution from the MG1655 strain carrying the inducible RFP plasmid (scraped bacterial pellet from frozen stock with a loop, then resuspended in 50 µL of water).
• 1 µL primer IGEM1 (10 µM)
• 1 µL primer IGEM2 (10 µM)
• 10 µL Q5 2X Master Mix

PCR mix (final volume: 20 µL) for promoter b0002_24h amplification:

• 1 µL of insert Insert_promoteur_24H
• 7 µL of water
• 1 µL primer b0002RV (10 µM)
• 1 µL primer b0002FW (10 µM)
• 10 µL Q5 2X Master Mix

PCR mix (final volume: 20 µL) for promoter b3021_24h amplification:

• 1 µL of insert Insert_promoteur_24H
• 7 µL of water
• 1 µL primer b3021RV (10 µM)
• 1 µL primer b3021FW (10 µM)
• 10 µL Q5 2X Master Mix

PCR cycle:

• Extension rate: 20–30 s per kb → elongation time set for each amplification

Gel preparation (1% agarose in 0.5X TAE):

• Stock solution prepared: 300 mL of 0.5X TAE with 1% agarose (stored, to be remelted in microwave when needed, BET room 2nd floor).
• BET added on gel rack before pouring molten agarose to reach final concentration 0.5 µg/mL.

Gel electrophoresis of PCR amplification

• Gel placed in tank with 0.5X TAE.
• Loaded 4 µL of each PCR product
• Migration: 25 min at 135 V.
• Visualization under UV.

PCR product clean-up (cf. Experiment : Monarch PCR & DNA Clean-up Kit)

• Final elution volume: 20 µL.
• Nanodrop linearized plasmid: 50 ng/µL
• Nanodrop b0002 promoter: 70 ng/µL
• Nanodrop b3021 promoter: 70 ng/µL

After calculating the optimal amount of plasmid and insert for each cloning on NEBuilder calculator:

Gibson assembly using NEBuilder HiFi DNA Assembly Master Mix

• Diluted inserts b3021 and b0002 1:4 because their concentration were too high compared to the concentration of the vector.
• New concentrations:
• b3021 → 17.5 ng/µL
• b0002 → 19.75 ng/µL

Gibson mix for b3021:

• 0.35 µL b3021 insert (1:4 dilution)
• 1.6 µL vector
• 3.05 µL DNase-free water
• 5 µL NEBuilder HiFi DNA Assembly Master Mix

Gibson mix for b0002:

• 0.4 µL b0002 insert (1:4 dilution)
• 1.6 µL vector
• 3 µL DNase-free water
• 5 µL NEBuilder HiFi DNA Assembly Master Mix

Incubation:

• 30 min at 50°C in thermocycler (15 min is suggested for <4 fragments by the NEB calculator; doubled for safety)
• Then placed on ice until transformation

Transformation:

• Thaw competent Heat Shock E.coli MG1655 ΔRM on ice.
• Add 3 µL Gibson assembly product, incubate 10 min on ice.
• Heat shock: 42 °C, 1 min, then 10 min on ice.
• Recovery: 1 mL LB, 30–60 min, allows expression of resistance cassette.

Plating

• Spin: 4,000 g, 2.5 min.
• Remove supernatant, leave ~100 µL, resuspend pellet.
• Plate on chloramphenicol agar: spread ~150 µL on half the plate with a spreader, then use same spreader for other half.
• Incubate overnight at 37 °C.

RNA-seq

We learned that the sequencing platform was unable to purify enough RNA to perform RNA-seq. The bacterium used is not a model organism; it is a capsulated strain, and we only had a rough estimate of the relationship between OD and cell concentration. Therefore, we need to consider alternative options for RNA-seq: a bacterium that is both well-characterized and reasonably resistant to PFAS.

In the meantime, we discussed with the 2024 iGEM team from Padua, who had tested E. coli growth and found that it grows well in the presence of several PFAS up to concentrations of 1 mM. Additionally, a 2025 study performed RNA-seq on E. coli MG1655 exposed to PFOA, which is how we identified genes that are upregulated in response to PFOA and traced back to their promoters.

Based on this, we decided to work with a sequencing platform closer to our lab (that we already reached out for technical advice, so that was quick to set everything up for this new RNA-seq), with stronger expertise in RNA sequencing. This will allow us to perform RNA-seq on E. coli MG1655 exposed to TFA, another PFAS for which we aim to design a biosensor, as it is increasingly present in the environment.

August 20th :

Cloning and transformation for biosensor

Figure 23: Illustration of the plasmid PCR tested to validate the next transformation

Colony PCR

Procedure:

• Collected colonies no. 1-5 from each of the plate using the tip of a pipette cone.
• Suspended each colony in 50 µL of ultrapure water in a PCR tube (1 tube per colony).

Colony PCR premix (for 12 reactions):

• 1.5 µL primer FW (FR15, 100 µM)
• 1.5 µL primer RV (FR14, 100 µM)
• 78 µL DreamTaq mix

Figure 26: Amplification Layout

PCR setup:

• In new PCR tubes, added 6.5 µL of premix + 6 µL of colony suspension → final volume of 12.5 µL per tube.
• Total: 10 PCR tubes

Next steps:

• Quick spin of the PCR tubes before starting the PCR program.

PCR program:

• Launched PCR with 35 cycles.
• Expected amplicon size: 1100 bp.
• DreamTaq elongation rate: ~1 kb/min.

Gel electrophoresis of PCR amplification

• Pour ~20 mL agarose 1% gel, add 4 µL BET (for visualization of PCR migration).
• Gel placed in tank with 0.5X TAE.
• Loaded 4 µL of each colony PCR amplification product.
• Migration: 20 min at 135 V.
• Visualization under UV.

The first wells (1–5) originate from colonies obtained with the transformation using b0002_24, and the second set of wells (1–5) originate from colonies obtained with the transformation using b3021_24. The second “1” well was loaded two times by mistake.

In parallel, we started liquid precultures in 3 mL LB + chloramphenicol for these 10 colonies (5 colonies with b0002 and 5 colonies with b3021) + 1 preculture of MG1655 with IPTG-inducible RFP plasmid + 1 preculture of MG1655 WT (to be monitored this evening)

Sanger sequencing

2 µL of amplification product from b0002_24h promoter colonies #1 and #2, and from b3021_24h promoter colonies #1 and #3, mixed with 1 µL of primer FR15, were sent for Sanger sequencing.

Overnight fluorescence time course on TECAN (96-well plate)

Several hours later, we used the precultures 4 of these 10 colonies to set up a 96-well plate (black-bordered) for overnight fluorescence monitoring on the TECAN.

Bacteria–medium premixes for overnight growth:

• 3 mL LB + 3 µL Cm + 30 µL MG1655 ΔRM with François’ plasmid + 3 µL IPTG → row H
• 3 mL LB + 3 µL Cm + 30 µL MG1655 ΔRM with François’ plasmid (no IPTG) → row G
• 3 mL LB + 30 µL MG1655 ΔRM without plasmid → row F
• 3 mL LB + 3 µL Cm + 30 µL b0002-1 → row B
• 3 mL LB + 3 µL Cm + 30 µL b0002-2 → row C
• 3 mL LB + 3 µL Cm + 30 µL b3021-1 → row D
• 3 mL LB + 3 µL Cm + 30 µL b3021-3 → row E

PFOA dilutions prepared (11 concentrations):

• 1 mM, 500 µM, 100 µM, 20 µM, 4 µM, 800 nM, 160 nM, 32 nM, 10.4 nM, 1.28 nM, 0.256 nM, 0

Plate setup:

• Added 190 µL of each premix to the wells.
• Added 10 µL of each PFOA dilution → final dilution 1/20.

Cultures were incubated overnight in the 96-well plate inside the TECAN reader.

Measurements (OD and fluorescence RFP) were recorded every 10 minutes throughout the night.

Growth monitoring of E.coli MG1655 to identify sampling points for RNA-seq after TFA exposure

M9 medium preparation (cf. Experiment Growth curve):

Preparation of 200 mL M9 medium

• 176 mL sterile pure water
• 20 mL M9 salts (stock of 100 mL containing: 6 g Na₂HPO₄, 3 g KH₂PO₄, 1 g NH₄Cl, 0.5 g NaCl)
• 1.2 mL MgSO₄ (0.33 M)
• 20 µL CaCl₂ (1 M)
• 200 µL thiamine hydrochloride (1 mg/mL)
• 2 mL glucose (40%)

We performed growth tests in both minimal medium (M9) and TSB medium diluted with sterile water. The M9 medium composition was as follows: 6 g/L Na2HPO4, 3 g/L KH2PO4, 1 g/L NH4Cl, 0.5 g/L NaCl, 2 mM MgSO4, 0.1 mM CaCl2, 4 g/L glucose, and 1 mg/L thiamine.

Once again, however, we were unable to perform growth assays with TFA due to laboratory biosafety regulations.

Figure 28 : Growth curve of E.coli on different medium

We chose to perform RNA-seq in TSB diluted at 1:2 medium

We identified that the 1:2 TSB dilution is the highest dilution allowing proper growth of E.coli MG1655. We decided not to use the M9 because the growth was very different as the ones reported in the literature. We suspected a problem in the M9 design. The selected time points for RNA-seq sampling are 4h and 8h. We will start the growth of E.coli MG1655 with different concentrations TFA only the next day, and perform sampling at 4h and 8h after the culture is initiated.

August 21st :

RNA-seq Sampling Protocol

The protocol for harvesting bacteria at selected time points was established in collaboration with the IGFL laboratory, which performed the sequencing. For each sample, a culture volume corresponding to approximately 10⁹ bacteria was collected.

We started E.coli MG1655 culture with either 100 µM of TFA, 0.87nM of TFA (european limit allowed) or water.

Experiment layout

14:00 (t = +4 h)

• OD₆₀₀ = 0.727 in tube TFA 100 µM #4.
• → 1.8 mL of culture was transferred into a 2 mL Eppendorf (Lock).
• Centrifugation: 10 min, 4000 g, 4°C.
• Supernatant removed by inversion.
• Pellets flash-frozen in liquid nitrogen and stored overnight at –80 °C.

18:00 (t = +8 h)

• OD₆₀₀ (after 2× dilution) = 0.776 → actual OD₆₀₀ = 1.552.
• → 886 µL of each culture was transferred into 2 mL Eppendorf tubes.
• Centrifugation: 10 min, 4000 g, 4°C.
• Supernatant removed by inversion.
• Pellets flash-frozen in liquid nitrogen and stored overnight at –80 °C.

Samples were transported to IGFL on dry ice by certified courier.

RNA purification was performed at the sequencing facility using the Illumina Stranded Total RNA Prep, Ligation with Ribo-Zero Plus kit, enabling rRNA depletion and library preparation.

Cloning and transformation for biosensor

After one night time course fluorescence we obtained these results :

In these graphs, fluorescence intensity was normalized to OD, ensuring that differences in fluorescence are not simply due to different growth rates.

For bacteria transformed with the construct containing the PFOA-sensitive promoter driving RFP expression, a common pattern can be observed: higher PFOA concentrations lead to stronger RFP expression, starting at around 5.5 h. However, the time course ends just as the curves begin to diverge more clearly (from ~8 h onward). It would therefore have been preferable to extend the time course, or at least avoid starting measurements at the very beginning of the culture when cell density was still very low.

In addition, replicates would be necessary to allow statistical analysis and confirm whether there is indeed a significant difference in fluorescence intensity between conditions. That’s what will be done tomorrow.

As expected, in the WT condition, E. coli MG1655 shows no difference in fluorescence/OD ratios across the different PFOA exposure conditions.

The initial parts of the curves may look unusual because the fluorescence/OD ratio decreases over time. This can be explained by the fact that the fluorescent response occurs later in bacterial growth: OD increases first, followed by the onset of fluorescence expression and signal intensity. This phenomenon is illustrated in the two graphs below.

Sanger sequencing result :

This result is consistent with our promoters: colonies #1 and #2 transformed with the plasmid containing the b0002_24h promoter, and colonies #1 and #3 transformed with the plasmid containing the b3021_24h promoter, were confirmed to carry their respective plasmids.

August 22nd/23rd/24th :

We performed a time-course (between t = 3h and t = 11h, then we measure it at t = 24h, t = 27h and a final time course between t = 27h and t = 35h) measurement of the transformed bacteria carrying PFOA-sensitive promoters with the reporter gene (RFP), in quadruplicate, and at different concentrations of PFOA.

96 wells layout

The goal is to identify the time point where fluorescence response is differential regarding the PFOA concentration. To identify this, we focused on the difference in the fluorescence/OD ratio of the transformed bacteria in the presence of PFOA compared to the same ratio without PFOA, in order to determine at which time points the biosensor was most effective and at which time points a proportional relationship might exist between PFOA concentration and fluorescence expression.

Two time points seem to meet our requirements for b0002: at t = 41,400 s (at t = 162,000 s E.coli growth is decreasing making the measure not reliable)

Two time points seem to meet our requirements for b0002: at t = 86,400s (=24h) s and t = 110,000 s (=30h). We plotted the PFOA-dependent response at the time point where it is most significant: for bacteria transformed with b3021, this occurs around t = 24 h and t = 30h, while for bacteria transformed with b0002, it occurs around t = 12 h.

We plotted the PFOA-dependent response at the time point where it is most significant: for bacteria transformed with b3021, this occurs around t = 24 h and t = 30h, while for bacteria transformed with b0002, it occurs around t = 12 h.

E. coli MG1655 ΔRM transformed with the reporter construct containing promoter b0002 is capable of displaying a PFOA-dependent response at t = 12 h. This response is sensitive to PFOA concentrations in the initial solution (in the 10 µL added to 190 µL of culture medium) of 800 µM (40 µM final concentration in the well), 4 mM (200 µM final concentration), and 20 mM (1 mM final concentration). Lower concentrations do not lead to a significantly different response. Significance testing was performed using a t-test after verifying the assumption of homoscedasticity (n = 4 for each condition).

E. coli MG1655 ΔRM transformed with the reporter construct containing promoter b3021 is partially capable of displaying a PFOA-dependent response at t = 24 h. This response is sensitive to PFOA concentrations in the initial solution (in the 10 µL added to 190 µL of culture medium) of 4 mM (200 µM final concentration), and 20 mM (1 mM final concentration). However, we observed that the fluorescent output measured under control conditions was higher than the output measured at concentrations of 1.28 µM, 6.4 µM, and 32 µM, which is very surprising. We currently have difficulty explaining this phenomenon, but it could be possible that the water used for the control was contaminated with PFAS in a greater concentration that 32µM, or by mistake when preparing the premix. Removing this point displays a very nice Fluorescence response of PFOA concentration. We did not have the time to redo the experience. Significance testing was performed using a t-test after verifying the assumption of homoscedasticity (n = 4 for each condition).

E. coli MG1655 ΔRM transformed with the reporter construct containing promoter b3021 is capable of displaying a PFOA-dependent response at t = 30 h. This response is sensitive to PFOA concentrations in the initial solution (in the 10 µL added to 190 µL of culture medium) of 800 µM (40 µM final concentration in the well) and 4 mM (200 µM final concentration), at 20mM the growth of E.coli started to decrease earlier, altering the ratio Fluorescence/OD. Lower concentrations do not lead to a significantly different response. Significance testing was performed using a t-test after verifying the assumption of homoscedasticity (n = 4 for each condition)

Histogram :

We also measured fluorescence in the same bacteria exposed to different concentrations of PFOA diluted in environmental water to assess whether promoter activation was specific to PFOA or interfered by compounds present in environment water. Fluorescence was recorded at four time points: t = 0, t = 20 h, t = 24 h, and t = 40 h. A response was detectable from t = 20 h onward; however, for both promoters, the responses at t = 20 h and t = 24 h showed a U-shaped pattern, making it impossible to link proportionally to the PFOA concentration. While the b3021 promoter exhibited a strong response even in the presence of water alone, the b0002 promoter displayed a significant increase in fluorescence (p < 0.05) when the PFOA concentration in the well was equal to or greater than 1 nM. Although the response was not proportional to PFOA concentration, it may still serve as a useful ON/OFF indicator for PFOA. At t=40 h, the response is likely unreliable, as E. coli had already entered the decline phase.

September 15th :

We obtained the data from the RNAseq facility :

A single-end, reverse-stranded RNA-seq was performed with an average sequencing depth of 28,518,648 reads per sample.

Data processing from raw reads to the generation of the count table was carried out on Galaxy Europe. The first step consisted of assessing data quality using the tool Falco (Galaxy Version 1.2.4+galaxy0).

Given the high quality of the raw data and the issues encountered when generating the count matrix after trimming, we decided, in consultation with a supervising instructor, to omit the trimming step. Reads were mapped using the “Map with BWA” tool (Galaxy Version 0.7.19), and read counting was performed with HTSeq-count (Galaxy Version 2.0.9+galaxy0). This analysis workflow was validated by our instructors based on the good quality of the raw data.

The resulting count matrix was subsequently analyzed using DESeq2 in R, yielding in particular the log2 fold change (L2FC) between two experimental conditions, as well as the adjusted p-value, which provide an estimate of both the magnitude of the transcriptional changes undergone by each gene and their statistical significance.

Starting from a DESeq2 output matrix (log2 fold change, adjusted p-value) and the raw count matrix, we selected genes with a log2 fold change (L2FC) > 1.5 and an adjusted p-value < 0.05. These genes were then ranked from highest to lowest L2FC.

As we did with the dataset of Wintenberg & al, 2025, we wanted to identify genes upregulated during TFA exposure, by looking at the L2FC value.

Among these candidates, we focused on genes with very low read counts under control conditions (H2O) but high read counts under TFA treatment, in order to minimize background noise. Importantly, we avoided selecting genes with strong L2FC values when their expression in the untreated condition was already high, since these would reduce biosensor sensitivity.

For each TFA treatment condition, we selected two genes meeting these criteria and belonging to distinct signaling or metabolic pathways, to reduce the likelihood of co-regulation.

At 4 h, the data did not reveal any particularly strong candidates, with the exception of ER3413_28, corresponding to carB (carbamoyl-phosphate synthetase large subunit). The average read count for this gene increased from 1,501 under control conditions (H₂O) to 4,629 under TFA exposure. However, a baseline read count as high as 1,501 would generate excessive background noise, making this gene unsuitable for use in a biosensor due to insufficient signal specificity.

At 8 h, several more upregulated genes were identified. Notably, ER3413_2576, corresponding to purM (phosphoribosylformylglycinamide cyclo-ligase), increased from an average of 429 reads (control) to 3,079 reads (TFA), and ER3413_3765, corresponding to xanP (xanthine:H⁺ symporter), increased from 184 reads (control) to 3,116 reads (TFA). Although purM and xanP are not part of the same operon, both are repressed by purR and are therefore co-regulated. To avoid potential co-regulation, we replaced purM with ER3413_1665, corresponding to gtrS (CPS-53 (KpLE1) prophage; serotype-specific glucosyl transferase YfdI), which belongs to a distinct metabolic pathway and is unlikely to be co-regulated with xanP.

We couldn’t analyse data from condition where E.coli MG1655 was exposed to 0,87 nM of TFA because when performing statistical analyses involving multiple hypothesis testing, such as with DESeq2, raw p-values are adjusted to control the false discovery rate (FDR). Methods like the Benjamini–Hochberg procedure rescale the p-values according to the number of tests and their rank. As a result, adjusted p-values can sometimes mathematically exceed 1. Since p-values are by definition bounded between 0 and 1, the software caps these values at 1.

In practice, this means that adjusted p-values close to or equal to 1 indicate no statistical evidence against the null hypothesis after correction. In consequence the statistical comparison between both conditions didn’t greater FDR < 0.9. It suggests that 0,87 nM of TFA does not lead to differential gene expression compared to water exposure.