iGEM 2025 — Team toulouse-insa-ut


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* ### Engineering
* [Overview of our engineering strategy](#bloc2)
* [Building an *E. coli* strain capable of orthogonal replication](#bloc3)
* [Development of a screening and selection platform for DeHa2 evolution](#bloc4)
* [Enhancing *P. putida*’s fluoride resistance](#bloc5)
* [References](#references)

![Engineering-Project](https://static.igem.wiki/teams/5755/images/header/headerfinaux/engineering1.webp)

## Overview of our engineering strategy

This page presents the Design-Build-Test-Learn (DBTL) cycles of our project PFAway. Here, we describe the rationale behind our design choices, the strategies we used to build and test our system, and the lessons we learned along the way. You will also find how we managed our experiments, overcame challenges, and improved our approach step by step. Throughout PFAway, we have completed three engineering cycles, focusing on three modules: the construction of a strain capable of orthogonal replication, the development of a screening technology for DeHa2, and the enhancement of *P. putida*’s fluoride resistance.

For each module, we performed a series of tests, analyzed the results, and adapted our strategy to improve outcomes. To visualize this process:

 • DBTL cycles (left boxes) are supported by experimental data (right boxes).

 • Down arrows indicate the iterations, highlighting the thoughts and new strategies that emerged from previous results.

 • Colors highlight outcomes of the cycles: red for unsuccessful attempts, green for successful results.

## Building an *E. coli* strain capable of orthogonal replication

Our primary objective was to introduce the replication operon (see part [BBa\_25HKSDJN](https://registry.igem.org/parts/bba-25hksdjn)) into *E. coli* DH10B in order to enable accelerated mutagenesis of the linear replicon (see part [BBa\_25794BPU](https://registry.igem.org/parts/bba-25794bpu)) for the directed evolution of DeHa2. Inspired by [Tian *et al*., 2024](https://www.science.org/doi/10.1126/science.adk1281), we first attempted chromosomal integration of the operon, before shifting to a strategy whereby the operon is carried on a low-copy plasmid.

| ![](https://static.igem.wiki/teams/5755/images/dbtl/head1.webp) | | |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/design-1.webp) We used the pTarget plasmid to deliver guide RNAs targeting the SS9 genomic region for Cas9, and the pCas plasmid to supply the molecular components necessary for both λ-Red recombination and Cas9 machinery. Together, these plasmids enable the insertion of the replication operon at the SS9 site within the bacterial chromosome (see [Design page](https://2025.igem.wiki/toulouse-insa-ut/design#bloc3-sub3)). |  |  |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/build.webp) Transformation of *E. coli* DH10B with pCas, pTarget, and, the replication operon. |  | ![](https://static.igem.wiki/teams/5755/images/dbtl/expres.webp)  ![Colony PCR of our strains after chromosomal insertion trials](https://static.igem.wiki/teams/5755/images/dbtl/figure2-3.webp)   **Figure 1: Colony PCR of our strains after chromosomal insertion trials.**  Well Mq shows the 1 kb Plus ladder from invitrogen, while well 1 to 16 corresponds to PCR on different colonies using primers flanking the chromosomal insertion. Bands at 1302 bp represent the SS9 site without insertion, whereas bands at 4982 bp correspond to the SS9 site with the chromosomal insertion.   WGS (data not shown) and colony PCR (Figure 1) indicate that the replication operon was not integrated at the SS9 site. Expected bands for successful insertion (4982 bp) were not observed; only bands corresponding to the empty SS9 site (1302 bp) are present. |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/arrowbis1.webp) |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/test.webp) Colony PCR screening of transformants and Whole Genome Sequencing (WGS).  ![](https://static.igem.wiki/teams/5755/images/dbtl/learn.webp) Despite several attempts, no successful integration was detected, neither from the chromosome sequencing data nor in colony PCR using primers flanking the SS9 site. These results suggest that the chromosomal insertion at SS9 was not efficient in our hands. |  |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/iteration1.webp) |  |  |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/head2bis.webp) | | |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/design-1.webp) We chose a one-copy plasmid (OCP). Using Gibson assembly, we attempted to insert the replication operon into the plasmid (see [Design page](https://2025.igem.wiki/toulouse-insa-ut/design#bloc3-sub3)). |  |  |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/build.webp) Gibson assembly of gBlocks A–B–C into the pUD1387 OCP backbone to construct pUD1387-operon. |  | ![](https://static.igem.wiki/teams/5755/images/dbtl/expres.webp)  ![Alignment of the expected one-copy plasmid and the plasmid extracted from transformants.](https://static.igem.wiki/teams/5755/images/dbtl/figure3.webp)   **Figure 2: Alignment of the expected one-copy plasmid and the plasmid extracted from transformants.**  The upper band corresponds to the expected one-copy plasmid containing the full synthetic operon, while the lower band represents the sequence of the plasmid extracted from the transformants. A mismatch or missing region is shown in red.   In Figure 2, alignment of the expected one-copy plasmid with the plasmid extracted from transformants shows that only part of the synthetic operon (gBlocks B and C) was successfully integrated. These results indicate that the current assembly approach needs optimization. Further experiments are ongoing, and results are pending. |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/arrowbis1.webp) |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/test.webp) Transformation of pUD1387-operon and sequencing.  ![](https://static.igem.wiki/teams/5755/images/dbtl/learn.webp) Transformation was partially successful: only gBlocks B and C were integrated into the plasmid, whereas gBlock A was missing. This indicates incomplete assembly or recombination in *E. coli*. |  |
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## Development of a screening and selection platform for DeHa2 evolution

Our objective was to implement a fluorescence-based biosensor for the detection of fluoride ions produced during the defluorination of short-chain PFAS by DeHa. The fluorescence intensity would report on the activity of the DeHa2 variants generated by orthogonal replication. The most active enzyme variants would then be selected. We first attempted to use a riboswitch-sfGFP construct that was described by the iGEM team [SDU Danemark 2023](https://2023.igem.wiki/sdu-denmark/index.html). Unfortunately, its sensitivity was insufficient for the low fluoride concentrations expected during the first evolution cycles. Inspired by recent work on RNA aptamers, we then turned to the FluorMango aptamer reported by [Husser *et al*., 2023](https://onlinelibrary.wiley.com/doi/abs/10.1002/smll.202205232). This RNA aptamer fluoresces upon fluoride binding and is expected to provide a specific signal. However, its extracellular nature made the aptamer difficult to use for bulk screening. To overcome this limitation and enable high-throughput applications, we developed a microfluidic-compatible screening protocol, see details in the [Design page](https://2025.igem.wiki/toulouse-insa-ut/design#bloc3-sub4).

| ![](https://static.igem.wiki/teams/5755/images/dbtl/head1.webp) | | |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/design-1.webp) Inspired by [SDU Danemark 2023](https://2023.igem.wiki/sdu-denmark/index.html), we constructed a pUC19 plasmid carrying a constitutive promoter that produces an RNA containing a riboswitch. In the presence of fluoride, this riboswitch enables the translation of sfGFP, resulting in fluorescence (see [Design page](https://2025.igem.wiki/toulouse-insa-ut/design#bloc3-sub4)). |  |  |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/build.webp) Construction of pUC19-sfGFP plasmid with fluoride riboswitch upstream of sfGFP. |  | ![](https://static.igem.wiki/teams/5755/images/dbtl/expres.webp)  ![Endpoint OD600 measurement of E. coli carrying pUC19-sfGFP under increasing NaF concentrations.](https://static.igem.wiki/teams/5755/images/dbtl/figure4-3.webp)   **Figure 3: Endpoint OD600 measurement of *E. coli* carrying pUC19-sfGFP under increasing NaF concentrations.**  Measurements were taken after overnight culture in a 96-well microplate. pUC19 bearing strain was used as a negative control, and p15A-PihfbsfGFP bearing strain was used as a positive control expressing sfGFP under the control of a constitutive promoter.   ![Normalized fluorescence (fluorescence/OD600 nm) of E. coli carrying pUC19-PJ23119-FRSsfGFP under increasing NaF concentrations.](https://static.igem.wiki/teams/5755/images/dbtl/figure5-3.webp)   **Figure 4: Normalized fluorescence (fluorescence/OD600 nm) of *E. coli* carrying pUC19-sfGFP under increasing NaF concentrations.**  Fluorescence measured at 𝜆excitation = 485 nm and 𝜆emission = 516 nm.   In Figure 3, growth curves (OD600) of *E. coli* carrying pUC19-sfGFP show reduced viability as NaF concentration increases. Normalized fluorescence (fluorescence/OD600) across NaF concentrations confirms that the riboswitch-sfGFP construct lacks sufficient sensitivity to respond to the low fluoride levels expected during initial mutagenesis cycles. Even at 25 mM NaF, no significant fluorescence was observed, casting some doubts on the utility of this biosensor under our conditions. |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/test.webp) Fluorescence was measured in 96-well microplates with increasing NaF concentrations. | ![](https://static.igem.wiki/teams/5755/images/dbtl/arrowbis1.webp) |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/learn.webp) The initial riboswitch design was not sensitive enough, making it incompatible with the small amounts of fluoride expected to be released by DeHa2 during early evolution cycles. |  |  |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/iteration2.webp) |  |  |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/head3.webp) | | |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/design-1.webp) A double-stranded DNA coding the FluorMango aptamer served as a template for *in vitro* transcription with the T7 RNA polymerase. This allowed for the expression of the aptamer, which was subsequently tested in microplates (see [Design page](https://2025.igem.wiki/toulouse-insa-ut/design#bloc3-sub4)). |

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| ![](https://static.igem.wiki/teams/5755/images/dbtl/build.webp) The aptamer was produced by *in vitro* transcription of the DNA template. |  | ![](https://static.igem.wiki/teams/5755/images/dbtl/expres.webp)  ![Specificity tests of the FluorMango aptamer after 1 hour of incubation.](https://static.igem.wiki/teams/5755/images/dbtl/figure7-1.webp)   **Figure 5: Specificity tests of the FluorMango aptamer after 1 hour of incubation.**  Fluorescence is shown as a function of the tested molecules: TFA, NaF, NaCl, and microfluidics oil (Fluo-oil 135) with or without surfactant (FluoroSurf O at 2%). n = 3 biological replicates with two technical replicates each. For TFA and oil: n = 1 biological replicate. Fluorescence was measured at 𝜆excitation = 510 nm and 𝜆emission = 550 nm.   ![Calibration of the FluorMango aptamer after 1 hour of incubation.](https://static.igem.wiki/teams/5755/images/dbtl/figure8.webp)   **Figure 6: Calibration of the FluorMango aptamer after 1 hour of incubation**  Relative fluorescence is shown as a function of increasing NaF concentrations, measured at 𝜆excitation = 510 nm and 𝜆emission = 550 nm. Each biological replicate (n = 3) included two technical replicates. Grey points represent individual measurements (n = 3), and black symbols indicate the mean values. The shaded area represents the confidence interval at 95%.   In Figure 5, specificity tests of FluorMango aptamer in the presence of various salts and small molecules confirm the findings of [Husser *et al*., 2023](https://onlinelibrary.wiley.com/doi/10.1002/smll.202205232) and demonstrate strong aptamer specificity. In Figure 6, calibration experiments across different fluoride concentrations show that the biosensor produces a clear, proportional, and highly fluoride-specific fluorescence response. |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/test.webp) Fluorescence measurement in 384-well microplates in the presence of NaF or other compounds (negative controls). |

 ![](https://static.igem.wiki/teams/5755/images/dbtl/arrowbis1.webp) ||  |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/learn.webp) The aptamer functions as a fluoride-specific biosensor. However, its extracellular nature (it does not permeate the cell wall) requires a compatible strategy to screen DeHa2 variants produced in the orthogonal replication strain. |

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| ![](https://static.igem.wiki/teams/5755/images/dbtl/iteration3.webp) |  |  |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/head4bis.webp) | | |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/design-1.webp) Develop a droplet microfluidic screening platform to compartmentalize the FluorMango aptamer and a single cell of the orthogonal replication strain to quantify fluoride release by DeHa2 variants. |

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| ![](https://static.igem.wiki/teams/5755/images/dbtl/build.webp) Water-in-oil emulsion droplets containing the FluorMango aptamer and its ligand TO1-biotin, with or without fluoride ions, were generated in a microfluidic device. Droplets were screened one by one for their fluorescence and were subjected to sorting based on intensity thresholding. |

  | ![](https://static.igem.wiki/teams/5755/images/dbtl/expres.webp)  ![Confocal fluorescence microscopy images of emulsion droplets containing the FluorMango aptamer and TO1-biotin after 1 h of incubation.](https://static.igem.wiki/teams/5755/images/dbtl/figure9.webp)   **Figure 7: Confocal fluorescence microscopy images of emulsion droplets containing the FluorMango aptamer and TO1-biotin after 1 h of incubation.**  Droplets without fluoride (NaF –) show no detectable fluorescence, whereas droplets with fluoride (NaF +) display strong fluorescence measured at 𝜆excitation = 480/40 nm and 𝜆emission = 527/30 nm.   ![Microfluidic sorting of a mixed droplet population (20% NaF +, 80% NaF –).](https://static.igem.wiki/teams/5755/images/dbtl/figure10.webp)   **Figure 8: Microfluidic sorting of a mixed droplet population (20% NaF +, 80% NaF –).**   Fluorescence measured at 𝜆excitation = 480/40 nm and 𝜆emission = 527/30 nm. Sorting was realized at 𝜆excitation = 488 nm and 𝜆emission= 525 nm. Post-sorting analysis shows strong enrichment of fluorescent droplets, confirming the selection efficiency of the FluorMango–microfluidics platform.   Figure 7: shows confocal fluorescence microscopy images of emulsion droplets containing the FluorMango aptamer and its ligand TO1-biotin after 1 hour of incubation with (NaF +) or without (NaF –) fluoride. Droplets without fluoride exhibited no detectable fluorescence, while droplets with fluoride displayed strong fluorescence, confirming the system’s functionality. Figure 8 presents a mixture of 20% fluoride-containing droplets and 80% fluoride-free droplets subjected to microfluidic sorting. After sorting, fluorescence microscopy revealed a strong enrichment of fluorescent droplets, demonstrating the efficiency of the selection process. Together, these results validate the FluorMango–microfluidics strategy as a robust platform for high-throughput screening. Once coupled with DeHa2-mediated defluorination, this system could be applied to select the most efficient variants after successive mutagenesis rounds. |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/test.webp) A mock library consisting of a mixed population of droplets either containing (fluorescent) or not containing (nonfluorescent) NaF was prepared at 1:5 ratio, i.e., 20% of active droplets. Sorting efficiency was determined by sorting the fluorescent droplets that were subsequently recovered and analyzed under fluorescence microscopy. | ![](https://static.igem.wiki/teams/5755/images/dbtl/arrowbis1.webp) |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/learn.webp) The sorted fraction of droplets contained 80% of fluorescent droplets, demonstrating successful enrichment. We conclude that FluorMango coupled with droplet-microfluidic is a viable strategy for high-throughput screening and selection of orthogonal replication–generated variants. |

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## Enhancing *P. putida*’s fluoride resistance

Our objective was to evaluate the fluoride tolerance of *Pseudomonas putida* KT2440 as it must withstand the rise of fluoride ions released during the degradation of short-chain PFAS by the evolved DeHa enzyme. As a strategy to enhance fluoride resistance, we focused on overexpressing FluC, a fluoride-specific transporter capable of expulsing intracellular ions out of the cells. Initial experiments without FluC overexpression provided data to feed our computational model, which indicated that overexpression would be necessary for *P. putida*’s viability. By overexpressing FluC, we aimed to improve fluoride tolerance, ensuring sustained growth and efficient PFAS degradation under high intracellular fluoride stress.

| ![](https://static.igem.wiki/teams/5755/images/dbtl/head5bis.webp) | | |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/design-1.webp) Wild-type *P. putida* KT2440 was grown in different media to generate diverse data for feeding our computational model (see [Model page](https://2025.igem.wiki/toulouse-insa-ut/model#bloc5)). |  |  |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/build.webp) *P. putida* KT2440 was grown in 96-well microplates in M9 minimal medium supplemented with acetate under increasing fluoride concentrations supplied in the form of NaF to assess its natural tolerance. |

  | ![](https://static.igem.wiki/teams/5755/images/dbtl/expres.webp)  ![Growth curves of P. putida KT2440 exposed to increasing NaF concentrations.](https://static.igem.wiki/teams/5755/images/dbtl/figure11-4.webp)   **Figure 9: Growth curves of *P. putida* KT2440 exposed to increasing NaF concentrations.**  Growth was monitored over time in M9 minimal medium supplemented with 45 mM acetate, using a 96-well microplate at 30 °C under different NaF concentrations. Solid lines represent mean values, while dashed lines correspond to individual replicates (n = 3).   In Figure 9, growth curves of *P. putida* exposed to different NaF concentrations over time are shown. Cell growth decreases as fluoride concentration increases, reflecting stress at levels relevant for PFAS degradation. These data provide a baseline measurement of natural fluoride tolerance in the strain, from which a MIC of 75 mM was estimated. |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/test.webp) OD600 growth curves were monitored in liquid culture supplemented with NaF, enabling inhibition constant calculation to feed the computational model for simulations of PFOS degradation with WT *P. putida.* |

 ![](https://static.igem.wiki/teams/5755/images/dbtl/arrowbis1.webp) |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/learn.webp) Native fluoride tolerance may be insufficient to support degradation of polyfluorinated compounds, suggesting that enhancing fluoride resistance could be required. |  |  |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/iteration4.webp) |  |  |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/head6.webp) | | |
| ![](https://static.igem.wiki/teams/5755/images/dbtl/design-1.webp) A pSEVA438 plasmid carrying a constitutive promoter was constructed to drive FluC transcription (see [Design page](https://2025.igem.wiki/toulouse-insa-ut/design#bloc4)). |  |  |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/build.webp) The pSEVA438-*fluC* plasmid was assembled to overexpress FluC in *P. putida*. |  | ![](https://static.igem.wiki/teams/5755/images/dbtl/expres.webp)  ![Growth curves of P. putida KT2440-pSEVA438-fluC overexpressing FluC under increasing NaF concentrations.](https://static.igem.wiki/teams/5755/images/dbtl/figure12-1.webp)   **Figure 10: Growth curves of *P. putida* KT2440-pSEVA438-fluC overexpressing FluC under increasing NaF concentrations.**  Growth was monitored in M9 minimal medium supplemented with 45 mM acetate, using a 96-well microplate at 30 °C under different NaF concentrations. Solid lines represent mean values, while dashed lines correspond to individual replicates (n=3).   In Figure 10, growth curves of *P. putida* overexpressing FluC are shown under varying NaF concentrations. Compared to the baseline, cells exhibit improved growth at high fluoride concentrations, with MIC increasing from 75 mM to 100 mM. This demonstrates that constitutive FluC overexpression significantly enhances fluoride resistance. The data also confirm that the engineered strain maintains improved growth at fluoride concentrations relevant to PFAS degradation, highlighting FluC overexpression as a promising strategy to mitigate fluoride stress. |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/test.webp) *P. putida* KT2440 was transformed with pSEVA438-fluC and growth was monitored under increasing NaF concentrations. | ![](https://static.igem.wiki/teams/5755/images/dbtl/arrowbis1.webp) |
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| ![](https://static.igem.wiki/teams/5755/images/dbtl/learn.webp) Constitutive FluC expression improves fluoride resistance, enabling robust growth under high fluoride stress that may be caused by efficient PFAS degradation. |  |  |

+**References**

Husser, C., Vuilleumier, S., and Ryckelynck, M. (2023) [FluorMango, an RNA‐Based Fluorogenic Biosensor for the Direct and Specific Detection of Fluoride.](https://onlinelibrary.wiley.com/doi/10.1002/smll.202205232) Small 19: 2205232.

Tian, R., Zhao, R., Guo, H., Yan, K., Wang, C., Lu, C., et al. (2023) [Engineered bacterial orthogonal DNA replication system for continuous evolution.](https://www.nature.com/articles/s41589-023-01387-2) Nat Chem Biol 19: 1504–1512.

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| Next, find out about our results here: |
| Results |

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* ### RESULTS
* [Evaluation of encapsulation beads for dual-strain water treatment](#bloc1)
  + [The bacterial strains continue to grow within the beads](#bloc1-sub1)
  + [Activated-carbon beads adsorb PFOA from aqueous solutions](#bloc1-sub2)
* [PFOS degradation by *Labrys portucalensis*](#bloc2)
  + [*L. portucalensis* and PFOS degradatio](#bloc2-sub1)
  + [Quantification of toxicity effects by NaF, TFA and PFPrA](#bloc2-sub2)
* [Engineering an orthogonal replication system in *E. coli*](#bloc3)
  + [The replication operon was successfully assembled and verified](#bloc3-sub1)
  + [Construction of a one-copy plasmid carrying the replication operon](#bloc3-sub2)
  + [The linear replicon was successfully assembled in pUC19 vector](#bloc3-sub3)
  + [Production of linear replicon with ITRs from pUC19-replicon](#bloc3-sub4)
  + [Creation of a dehalogenase-free linear replicon](#bloc3-sub5)
* [Screening ORep strains with FluorMango and microfluidics](#bloc4)
  + [FluorMango fluorescence is sensitive to NaF concentration variations](#bloc4-sub1)
  + [FluorMango responds specifically to fluoride ions](#bloc4-sub2)
  + [FluorMango detects fluoride released from enzymatic defluorination](#bloc4-sub3)
  + [FluorMango enables fluorescence detection of fluoride in microfluidic droplets](#bloc4-sub4)
  + [FluorMango allows for fluorescence-based sorting in microfluidics](#bloc4-sub5)
* [Enhancing the resistance of *P. putida* to fluoride ions with FluC](#bloc5)
  + [Successful construction of the pSEVA438-*fluC* plasmid](#bloc5-sub1)
  + [Overexpression of *fluC* accelerates cellular adaptation to fluoride stress](#bloc5-sub2)
  + [Process limits were assessed by PFPrA and TFA toxicity on *P. putida*](#bloc5-sub3)
* [Discussion and perspectives](#bloc6)

![Results-project](https://static.igem.wiki/teams/5755/images/header/headerfinaux/resultsf.webp)

This page is dedicated to the achievements of our experiments in the lab. We will focus on key results:

![Card introduction](https://static.igem.wiki/teams/5755/images/results/figonebis.webp)

## Evaluation of encapsulation beads for dual-strain water treatment

**Context:** Our bioremediation approach relies on two bacterial strains that cannot be co-cultured due to their different growth rates. To overcome this, each strain was encapsulated separately in activated carbon beads. This strategy allows us to combine the effects of both strains without co-culture, prevents the release of GMOs into the treated water, and leverages the PFAS adsorption capacity of activated carbon. These pilot experiments aim to evaluate the long-term viability of the bacteria within the beads, the potential release of encapsulated strains into water, and the adsorption of PFOA onto the beads.

### Bacterial viability is ensured within the beads

**Objective:** We aimed at testing the viability of both strains over time within the beads to estimate the lifespan of our degradation process.

**Method:** The release and stability of bacteria were evaluated by incubating dried beads in sterile water (Figure 1), encapsulating either *Pseudomonas putida* with 10% or 20% activated carbon, or *Labrys portucalensis* with 20% activated carbon. Control beads without bacteria were also prepared using sterile water instead of bacterial culture.

![Figure 1](https://static.igem.wiki/teams/5755/images/results/fig1.webp)

**Figure 1: Pictures of A) dried activated-carbon beads and B) beads in solution.**

The production of this support matrix for our strains was carried out at our partner’s headquarter YpHen, in Grenoble (FR).

Supernatant samples were collected at different time points to measure bacterial release, and beads from dedicated tubes were dissolved with 1% tricitrate solution to assess bacterial viability within the beads. Bacterial counts were determined by serial dilutions, plating, and colony forming unit (CFU) enumeration.

![Figure 2](https://static.igem.wiki/teams/5755/images/results/fig2bis.webp)

**Figure 2: Stability and release of encapsulated bacteria over time.**

Time course of A) bacterial viability in encapsulated beads and B) bacterial release from encapsulated beads. Samples were incubated at room temperature (25°C) under shaking (200 rpm): *Pseudomonas putida* (10% and 20% activated carbon, AC), *Labrys portucalensis* (20% AC), and control beads without bacteria over 30 days.

**Results:** All strains remained viable and even showed growth for at least 10 days (Figure 2A). *Pseudomonas putida* in 10% activated carbon beads continued to grow for up to 30 days. Although the other conditions were not tested for the full 30-day period, similar trends are likely. No significant differences in bacterial viability were observed between beads containing 10% or 20% activated carbon, indicating that carbon content does not compromise survival.

Regarding bacterial release, Figure 2B shows that the number of bacteria released into water increases over time, at levels comparable to the bacteria remaining in the beads. This indicates that the current bead formulation is not optimized to retain bacteria. Major adjustments to the formulation will be discussed with YpHen to ensure bacterial confinement and comply with biosafety regulations.

### Activated-carbon beads adsorb PFOA from aqueous solutions

**Objective:** We aimed at testing the efficiency of PFAS removal from aqueous solutions of activated-carbon beads.

**Method:** The adsorption of PFOA on beads (10% and 20% activated carbon, 10% and 20% talc, and a negative control without beads) was assessed by incubating 1.0 g of beads in a 50 µM PFOA solution.

![Figure 3](https://static.igem.wiki/teams/5755/images/results/fig3.webp)

**Figure 3: Evolution of PFOA concentration in presence of beads.**

Bacteria-free beads containing activated carbon or talc (● or ▲) at 10% or 20% loading (grey or dark) were incubated at room temperature (25°C) under shaking (200 rpm) in 20 mL of 50 µM PFOA solution. The no-bead negative control is shown in red.

**Results:** Figure 3 clearly shows a distinct difference in PFOA concentration over time between beads with or without activated carbon, as well as the negative control without beads. The results indicate that beads containing 20% activated carbon adsorbed 90.2% of PFOA within 120 minutes, while those with 10% activated carbon achieved 96.9% adsorption. In contrast, beads containing talc showed much lower adsorption efficiencies, 36.7% for 20% talc and 33.5% for 10% talc. The control exhibited a 26% decrease in PFOA concentration, likely due to adsorption onto the walls of the tubes used in the experiment.

These findings confirm that the presence of activated carbon in the beads is primarily responsible for PFOA adsorption and highlight the potential synergistic effect between adsorption and bacterial degradation. Interestingly, the 10% activated-carbon beads demonstrated slightly higher adsorption efficiency than the 20% beads. This suggests that similar or even better removal performance can be achieved with a lower activated carbon content, thereby reducing production costs while maintaining high adsorption capacity.

We then attempted to characterize PFOS degradation by *L. portucalensis*.

## PFOS degradation by *Labrys portucalensis*

**Context:** As the first axis of PFAway, *L. portucalensis* is used to break down long-chain PFAS such as PFOS. This newly described strain needs further characterization before implementation in the PFAway process.

### *L. portucalensis* and PFOS degradation

**Objective:** We aimed at following PFOS degradation by the strain as performed in [Wijayahena et al., 2025](https://www.sciencedirect.com/science/article/pii/S0048969724085061), but this time by increasing 10 fold the starting concentration of bacteria to measure the impact on degradation efficiency.

**Method:** Degradation of 10 mg/L of PFOS as the sole carbon source was monitored in M9 minimal medium for 30 days by LC-MS/MS. *L. portucalensis* was inoculated at two different starting concentrations, OD600=0.1 and 1.

**Results:** Changes in optical density over time suggested different bacterial growth responses depending on the initial inoculum (Figure 4).

![Figure 4](https://static.igem.wiki/teams/5755/images/results/fig4.webp)

**Figure 4: Growth curves of *L. portucalensis* during the PFOS degradation assays.**

Growth in M9 minimal medium supplemented with 10 mg/L of PFOS as the sole carbon source was monitored in tubes. Replicates are shown as dashed lines, and the mean as a solid line (n=2).

At starting OD600=1.0, we observed a gradual decrease from 1.00 to 0.50 OD600 over time, indicating a decline in the bacterial population.

In contrast, at an initial OD600 of 0.1, a slight increase was observed, suggesting that a smaller starting population could better utilize PFOS as a carbon source, likely due to reduced competition.

Further experiments with intermediate inoculum densities may help identify the maximum threshold below OD600=1 that still allows growth, thereby clarifying the relationship between initial biomass, resource availability, and bacterial activity under PFOS exposure.

PFOS degradation assays turned out to be inconclusive. During sampling, the filtering step used to remove bacteria before LC-MS/MS analysis led to adsorption of PFOS to the filter, precluding quantification of PFOS concentration. For future experiments, we recommend substituting the filtration step with cell pelleting by centrifugation and removal of the supernatant containing PFOS for mass spectrometry quantitation.

### Quantification of toxicity effects by NaF, TFA and PFPrA

**Objective:** Degradation of PFOS generates PFPrA, TFA, and fluoride. These products are toxic to cells and might determine the maximal PFOS concentration that our process will be able to treat. We therefore assayed the minimal inhibitory concentration (MIC) of PFPrA, TFA, and NaF on *L. portucalensis*.

**Method:** Growth curves were monitored in the presence of NaF, TFA, and PFPrA by monitoring OD600 kinetics in a 96-well microplate reader on glucose as the carbon source in M9 minimal medium.

**Results:** All the collected data are available via our [Kinetic Viewer](https://devlinlucas.shinyapps.io/PFAway-KineticViewer/). This allows a complete and easy visual presentation of the kinetic data, and the corresponding R code is available on [Contribution page](https://2025.igem.wiki/toulouse-insa-ut/contribution#bloc6). The most relevant data are presented here. *L. portucalensis* was able to grow until 40 mM of both TFA and PFPrA, (Figure 5A). However, MIC for NaF was not reached, and lies in a range over 40 mM (Figure 5B). Concentrations above 50 mM NaF were tested but the resulting growth curves could not be analyzed because the M9 medium crystallized at such high concentrations. Interestingly, this effect was not observed in toxicity assays with *Pseudomonas putida* (as shown later on this [page](https://2025.igem.wiki/toulouse-insa-ut/results#bloc5)).

![Figure 5](https://static.igem.wiki/teams/5755/images/results/fig5.webp)

**Figure 5: Toxicity assays of NaF, PFPrA and TFA on *Labrys portucalensis*.**

A) Growth curves of *L. portucalensis* for different NaF (top), PFPrA (middle) and TFA (bottom) concentrations. Technical and biological replicates (n=6) are shown as transparent lines and the mean as bold lines. Growth was monitored in M9 minimal medium supplemented with 15 mM glucose in a 96-well microplate at 30°C across different concentrations of fluorinated compounds. B) Minimal inhibitory concentrations (MIC) of NaF, PFPrA and TFA on *L. portucalensis*.

As shown in Figure 5A, we observed important variations between biological replicates. This variability is primarily due to the difficulty of synchronizing *Labrys portucalensis* cultures, as the strain exhibits a long lag phase in pre-cultures (2 days). This issue is evident for all molecules shown in the figure and across all experiments with *L. portucalensis* (see on the [Kinetic Viewer](https://devlinlucas.shinyapps.io/PFAway-KineticViewer/)). While this variability does not prevent the determination of MIC values, it makes it challenging to accurately define characteristic growth parameters such as growth rates (μ) and lag time. To address this, we decided to normalize the biological replicates against their control condition (0 mM) in order to highlight trends in the evolution of μ and lag time as a function of molecule concentration (Figure 6).

![Figure 6](https://static.igem.wiki/teams/5755/images/results/fig6.webp)

**Figure 6: Kinetic parameters of *Labrys portucalensis* growth in presence of NaF, PFPrA, and TFA.**

Kinetic parameters were normalized to the 0 mM control within each replicate to account for variability across replicates. Normalized A) growth rate and B) lag time of *L. portucalensis* in the presence of NaF, PFPrA, or TFA at the indicated concentrations. Data show the mean of three normalized replicates (n=3). Means are shown as bold points, replicates are shown as transparent points.

The maximum growth rate decreases as the concentration increases for the different fluorinated compounds. No clear trend could be detected for the lag time.

These findings indicate that, although growth inhibition by PFOS degradation products can be quantified, variability in the physiological state of the replicates limits the precise determination of kinetic growth parameters.

We next turned to the second axis of the project: directed evolution of DeHa2 for degradation of short-chain PFAS.

## Engineering an orthogonal replication system in *E. coli*

**Context:** We designed an orthogonal replication system (ORep) in *E. coli*, to generate a library of genetic variants of DeHa2 (see our [Design page](https://2025.igem.wiki/toulouse-insa-ut/design#bloc3-sub3)). The system requires two components: the linear replicon containing the dehalogenase gene (*dehH2*) flanked by its ITRs, and the replication operon encoding the error-prone polymerase and associated proteins that replicate the linear replicon in continuous cycles.

### The replication operon was successfully assembled and verified

**Objective:** We aimed to assemble the replication operon, which encodes a double-mutant error-prone DNA polymerase (ODNAP) and associated proteins (TP, DSB, and SSB) required for continuous replication of the linear replicon, with the goal of either integrating it into the chromosome of *E. coli* or cloning it into a single-copy plasmid.

**Method:** The replication operon was initially synthesized into three gBlocks (A, B and C) by IDT. Each fragment contained 60 bp overlapping sequences to allow for Gibson assembly as well as homology regions for chromosomal integration in *E. coli*. The assembled operon was then amplified by PCR using KOD polymerase. To validate the good assembly of the operon, we performed agarose gel electrophoresis on the PCR performed on the assembled product and confirmed these results by linear amplicon sequencing using nanopore technology (Eurofins Genomics).

**Results:** Gel electrophoresis analysis showed a PCR product of the expected size (Figure 7). In addition, sequencing of the purified fragment confirmed the correct assembly of the replication operon (see part [BBa\_25HKSDJN](https://registry.igem.org/parts/bba-25hksdjn)). This validates the construct for transformation into a bacterial host, with the goal of generating a strain carrying the ORep system to enable continuous mutagenesis.

![Figure 7](https://static.igem.wiki/teams/5755/images/results/fig7.webp)

**Figure 7: Agarose gel electrophoresis** confirming the successful assembly of the replication operon by PCR amplification from the Gibson-assembled construct.

The main objective was to integrate the operon into *E. coli* chromosome to implement the ORep mutagenesis system, a modification made possible using two helper plasmids (pCas and pTarget) enabling CRISPR-assisted recombineering. However, this approach did not yield the expected results (see our [Engineering page](https://2025.igem.wiki/toulouse-insa-ut/engineering#bloc3)). We therefore sought for an alternative using a one-copy plasmid.

### Construction of a one-copy plasmid carrying the replication operon

**Objective:** We aimed at inserting the replication operon into *E. coli* to generate the ORep strain, prior transformation with the linear replicon. The first strategy was chromosomal integration at the SS9 locus using CRISPR-assisted λ-Red recombination (see our [Engineering page](https://2025.igem.wiki/toulouse-insa-ut/engineering#bloc3)). As this was not achieved, an alternative strategy was designed based on assembling the operon into the one-copy plasmid pUD1387, which carries chloramphenicol resistance.

**Method:** The replication operon was amplified by PCR while excluding pre-existing SS9 sites within its sequence. The pUD1387 plasmid backbone was amplified by PCR to generate a linear fragment. Following Gibson assembly with both fragments, the product was transformed into competent *E. coli* cells and plated on LB agar supplemented with chloramphenicol for selection. Colonies were screened by colony PCR, and the presence of the operon was verified by PCR on purified plasmid and subsequent sequencing.

**Results:** PCR amplifications produced bands at the expected sizes: 3 804 bp for the operon and 6 945 bp for the linearized pUD1387 backbone (Figure 8).

![Figure 8](https://static.igem.wiki/teams/5755/images/results/fig8.webp)

**Figure 8: Agarose gel electrophoresis showing the linear operon and backbone fragments generated by PCR.**

Lane ABC: amplification of the operon at the expected size of 3 804 bp.
Lane OCP: amplification of the one-copy plasmid pUD1387 at the expected size of 6 945 bp.

Colonies grew on LB-chloramphenicol plates after transformation, confirming uptake of plasmid DNA. However, diagnostic PCR on miniprepped clones did not yield the expected amplicon size for the assembled plasmid; instead, smaller fragments were obtained (Figure 9).

![Figure 9](https://static.igem.wiki/teams/5755/images/results/fig9.webp)

**Figure 9: Agarose gel electrophoresis of colony PCR from four independent clones.**

Left side of the ladder: amplification product was expected at 5 572 bp, smaller bands were observed instead. Right side of the ladder: amplification product was expected at 5 243 bp, with bands matching the expected size. The discrepancy on the left suggests that a portion of gBlock A may be missing.

Sequencing confirmed incomplete assembly: only part of gBlock A was integrated, while gBlocks B and C were recovered intact in the plasmid (Figure 10).

![Figure 10](https://static.igem.wiki/teams/5755/images/results/fig10bis.webp)

**Figure 10: Alignment of the expected one-copy plasmid carrying the replication operon (upper sequence) with the sequencing result (lower sequence).**

A large missing region is highlighted in red, corresponding to most of gBlock A at the position where the DNA polymerase gene was expected.

Repeated attempts with purified PCR products and optimized assembly conditions yielded the same outcome. The incomplete recovery of the operon suggests instability of the construct in *E. coli*, possibly linked to the presence of foreign DNA polymerase genes. Future work will involve testing alternative cloning strategies to achieve stable maintenance of the complete operon.

### The linear replicon was successfully assembled in pUC19 vector

**Objective:** We aimed to assemble the linear replicon designed to carry the dehalogenase gene (*dehH2*), the kanamycin-resistance gene (*KanR*) and flanked by ITRs, then to clone it into the pUC19 backbone, and validate the construction by sequencing.

**Method:** Linear replicon was first synthesized as two gBlocks (D and E) from IDT. Both fragments harboring *dehH2*, *KanR* and flanked by ITRs were cloned into the pUC19 vector by Gibson assembly to generate the replicon. The assembled construct was transformed into *E. coli* DH5α and plated on LB agar supplemented with kanamycin and ampicillin (*KanR* from gBlock D and *AmpR* from pUC19). Colonies were screened by restriction digestion. Plasmid DNA was extracted and subsequently sequenced by whole-plasmid sequencing using nanopore technology (Eurofins Genomics) to confirm the correct assembly and integrity of the linear replicon.

**Results:** Colonies were obtained on LB agar plates containing kanamycin and ampicillin, indicating successful transformation with the pUC19-containing linear replicon construct. Plasmid DNA from five selected clones was digested with BamHI, producing the expected band pattern (Figure 11). This result suggests correct assembly of the linear replicon within the pUC19 backbone.

![Figure 11](https://static.igem.wiki/teams/5755/images/results/fig11.webp)

**Figure 11: Agarose gel electrophoresis for validation of the pUC19-replicon biobrick by BamHI restriction digestion.**

A) Predicted digestion profile. B) Digestion patterns obtained from five independent colonies.

Sequencing of the miniprepped plasmids confirmed the correct assembly of the biobrick (see pUC19-replicon sequence on [Materials page](https://2025.igem.wiki/toulouse-insa-ut/materials#bloc2-sub1)).

The sequence-verified pUC19-replicon was subsequently employed for the expression of DeHa2 in the fluorescence assays using the FluorMango biosensor (see later on this [page](https://2025.igem.wiki/toulouse-insa-ut/results#bloc4-sub1)).

### Production of linear replicon with ITRs from pUC19-replicon

**Objective:** We aimed at amplifying and purifying the linear replicon from the pUC19-replicon biobrick. This linear fragment will later be introduced into the DH10B host carrying the replication operon to enable continuous orthogonal replication of the encoded *dehH2* gene.

**Method:** Two primers targeting the linear replicon were designed to amplify the ITR-flanked fragment from the pUC19-replicon biobrick by PCR using KOD One polymerase. The PCR product was verified by agarose gel electrophoresis, purified, and sent for sequencing by whole-plasmid sequencing using nanopore technology (Eurofins Genomics).

**Results:** Agarose gel electrophoresis of the PCR product showed a single band at the expected size (Figure 12), confirming successful amplification of the linear replicon from the biobrick.

![Figure 12](https://static.igem.wiki/teams/5755/images/results/fig12.webp)

**Figure 12: Agarose gel electrophoresis of the linear replicon amplified by PCR from the pUC19-replicon biobrick.**

The expected band at 2 214 bp, corresponding to the linear replicon, is observed.

Sequencing of the purified fragment confirmed the linear replicon had the correct sequence (see part [BBa\_25794BPU](https://registry.igem.org/parts/bba-25794bpu)).

We conclude that the linear replicon is ready for transformation into the bacterial host, where it provides the template carrying the DeHa2 enzyme to be subjected to continued mutagenesis by the ORep machinery.

### Creation of a dehalogenase-free linear replicon biobrick

**Objective:** We aimed at generating a version of the linear replicon biobrick lacking the *dehH2* coding the DeHa2 enzyme. This modified biobrick serves as a versatile kit, allowing users to insert their own target gene for continuous orthogonal replication and directed evolution.

**Method:** Two PCRs were performed from the biobrick: one amplifying the left half of the biobrick and the other the right half, both excluding the *dehH2* sequence. PCR products were purified and joined by Gibson assembly to reconstruct the biobrick lacking *dehH2*. The assembled construct was transformed into *E. coli* for amplification and extracted by miniprep from multiple clones. Colonies were screened by restriction digestion to verify the absence of the *dehH2* sequence. Plasmids from selected clones were sequenced by whole-plasmid sequencing using nanopore technology (Eurofins Genomics) to confirm the correct deletion and integrity of the biobrick.

**Results:** Agarose gel analysis of the two PCR products showed single bands (an upper band of lower intensity is also observed in one PCR product) of the expected size (~ 2 000 bp each), confirming successful amplification (Figure 13).

![Figure 13](https://static.igem.wiki/teams/5755/images/results/fig13.webp)

**Figure 13: Agarose gel electrophoresis showing PCR linearization of the pUC19-linear replicon construct into two fragments.**

Amplification was designed to exclude the dehalogenase gene. The expected bands were obtained: 1 932 bp (left) and 1 838 bp (right).

Gibson assembly products were amplified in *E. coli*. Restriction digestion of miniprepped plasmids showed that only clones 3 and 4 had the expected digestion pattern corresponding to the biobrick without *dehH2* (Figure 14). Comparison with the original biobrick containing *dehH2* corroborated the deletion.

![Figure 14](https://static.igem.wiki/teams/5755/images/results/fig14.webp)

**Figure 14: Removal of the dehalogenase was confirmed by restriction digestion.**

A) Expected profile: Lane 1 is the biobrick with dehalogenase gene and lane 2 is the biobrick without the dehalogenase gene for the kit. B) Experimentally obtained profile.

Sequencing results of plasmids from clones 3 and 4 confirmed the removal of the *dehH2* gene and the correct sequence of the remaining biobrick (see pReplicon\_KIT sequence on [Materials page](https://2025.igem.wiki/toulouse-insa-ut/materials#bloc2-sub1)).

We then aimed at developing a screening method to be combined with ORep: fluoride-sensitive biosensor integrated with droplet microfluidics.

## Screening ORep strains with FluorMango and microfluidics

#### Measurement in microplates

**Context:** We aimed to establish a FluorMango biosensor-based microfluidic screening approach that will later be applied to select active DeHa2 variants (for more details, see our [Design page](https://2025.igem.wiki/toulouse-insa-ut/design#bloc3-sub4)). We first performed microplate-based tests to evaluate FluorMango’s sensitivity, specificity, and proof-of-concept performance. For more information on the FluorMango microplate assays, see our [Best Measurement page](https://2025.igem.wiki/toulouse-insa-ut/measurement#bloc2).

### FluorMango fluorescence is sensitive to NaF concentration variations

**Objective:** We aimed at quantifying the responsiveness of FluorMango to varying amounts of fluoride ions.

**Method:** FluorMango aptamer was incubated for 1 hour with TO1-biotin, 1X refolding buffer and increasing NaF concentrations, and fluorescence was measured in 384-well plates.

**Results:** Figure 15 clearly demonstrates that the fluorescence intensity increases with the fluoride concentration up to 3 mM NaF. Moreover, the aptamer is sensitive to 0.1 mM NaF.

![Figure 15](https://static.igem.wiki/teams/5755/images/results/fig15.webp)

**Figure 15: FluorMango/TO1-biotin complex response to fluoride ions.**

FluorMango (400 nM) aptamer was incubated for 1 hour with TO1-biotin (700 nM), 1X refolding buffer and increasing NaF concentrations. Fluorescence of FluorMango/TO1-biotin complex was directly measured at 𝜆excitation=510 nm and 𝜆emission=550 nm. The x-axis represents NaF concentration and is displayed on a logarithmic scale. Grey points represent individual measurements (n=6), and black symbols indicate the mean values. The shaded area represents the confidence interval at 95%.

The next step was to show if this response is specific to fluoride ions.

### FluorMango responds specifically to fluoride ions

**Objective:** We assessed the specificity of FluorMango fluorescence. To this end, we exposed the aptamer to a variety of molecules relevant to our experiments and measured its fluorescence.

**Method:** FluorMango aptamer was incubated for 1 hour with TO1-biotin (400 nM), 1X refolding buffer and NaF, NaCl, TFA or fluorinated microfluidic oil. Fluorescence was measured in 384-well plates.

**Results:** A strong fluorescence signal, up to 14-fold, was observed exclusively in the presence of fluoride ions, whereas fluorescence remained low in the presence of NaCl, TFA, or the fluorinated oil used in microfluidics (Figure 16). This remarkable specificity, together with its quantitative responsiveness, validates FluorMango as a reliable biosensor for fluoride ions.

![Figure 16](https://static.igem.wiki/teams/5755/images/results/fig16.webp)

**Figure 16: Fluorescence of FluorMango/TO1-biotin complex in presence of different compounds.**

FluorMango aptamer (400 nM) was incubated for 1 hour with TO1-biotin (700 nM), 1X refolding buffer and 1 mM of NaF, NaCl, TFA or fluorinated microfluidics oil (Fluo-oil 135) with or without surfactant (FluoroSurf O at 2%). Fluorescence of FluorMango/TO1-biotin complex was directly measured at 𝜆excitation=510 nm and 𝜆emission=550 nm. Values are the mean of three independent experiments (n=3), and errors bars correspond to standard deviation. The individual data points are also displayed in grey.

We then tested whether the biosensor could detect fluoride released during bacterial defluorination of a fluorinated substrate.

### FluorMango detects fluoride released from enzymatic defluorination

**Objective:** We investigated if FluorMango is compatible with the detection of fluoride in the presence of bacteria, i.e., under conditions close to the screening of DeHa2 variants with ORep *E. coli* cells.

**Method:** FluorMango was incubated with TO1-biotin, 2-fluoropropionate (2-FP) as the monofluorinated substrate for DeHa2, and *E. coli* cells expressing recombinant DeHa2 (from pUC19-replicon plasmid). Substrate degradation kinetics were monitored over 4 hours in 384-well plates by directly measuring the fluorescence of the FluorMango/TO1-biotin complex.

**Results:** We found that the defluorination of 2-FP and the accompanying production of F- by DeHa2 (with IPTG) could successfully be detected by FluorMango (upper curve in Figure 17), while the fluorescence signal remained low in the control samples (no IPTG induction, only bacteria, only 2-FP). Summarizing, we provided a detailed characterization of FluorMango responsiveness, establishing this biosensor as a quantitative tool for the directed evolution of DeHa2.

![Figure 17](https://static.igem.wiki/teams/5755/images/results/fig17.webp)

**Figure 17: Real-time monitoring of fluoride (F⁻) release during IPTG-induced DeHa2 expression in *E. coli* DH5α.**

Cultures were grown overnight in LB at 37 °C, washed in PBS, and incubated in a microplate reader at 25°C with IPTG (0.1 mM), 2-FP (50 mM), FluorMango (400 nM), TO1-biotin (700 nM) and 1X refolding buffer. Fluorescence was recorded with λexcitation=510 nm and λemission=550 nm. Values are the mean of two independent experiments (n=2), and errors bars correspond to the standard deviation.

Having validated FluorMango as a sensitive and specific biosensor in microplate assays and even in the presence of living bacteria, the next step was to test whether this technology could be translated into a droplet-based format. Microfluidic encapsulation not only provides the high-throughput capacity required for directed evolution but also allows linking enzymatic activity to individual genetic variants. We therefore explored whether FluorMango retains its functionality inside droplets and whether it can support the sorting of active versus inactive variants in a pooled screening setup.

#### Measurement in droplet-microfluidics

**Context:** Pooled genetic screens of DeHa2 variants require a method to efficiently link the gene sequence to enzymatic activity. During hypermutagenesis in the ORep *E. coli* strain, a large diversity of the *dehH2* gene is created within the bacterial population. Isolation of genetic variants and their phenotypic outputs is then essential for directed evolution.

Our strategy consisted in compartmentalizing individual ORep cells along with FluorMango inside water-in-oil emulsion droplets generated by microfluidics. To find out more about the microfluidic assays, see our [Best Measurement page](https://2025.igem.wiki/toulouse-insa-ut/measurement#bloc3).

### FluorMango enables fluorescence detection of fluoride in microfluidic droplets

**Objective:** We aimed at detecting fluoride ions in microfluidic droplets using FluorMango.

**Method:** FluorMango was incubated with TO1-biotin, 1X refolding buffer and either NaF (1 mM, for positive-fluorescent droplets) or RNAse-free water (for negative droplets). Droplets were generated using two approaches: in the first one, FluorMango activation occurred only after encapsulation, whereas in the second one, FluorMango was pre-activated by fluoride prior encapsulation. Fluorescence of FluorMango/ TO1-biotin in droplets was measured by fluorescence microscopy.

**Results:** We visualized the positive and negative droplets collected from the microfluidic chip by fluorescence microscopy. With both sample preparation and droplet generation methods, no fluorescence was detected in negative control droplets, whereas positive control droplets consistently exhibited strong fluorescence (Figure 18). These results demonstrate the successful detection of fluoride ions by FluorMango in microfluidic droplets.

![Figure 18](https://static.igem.wiki/teams/5755/images/results/fig18.webp)

**Figure 18: Fluorescence detection of fluoride (F⁻) in microfluidic droplets using the FluorMango biosensor**

Bright-field (top) and fluorescence (bottom) microscopy images of negative (+RNAse-free water) and positive (+NaF) droplets generated with A) method 1 where FluorMango activation occurred after encapsulation, B) method 2 where FluorMango activation occurred prior encapsulation. Fluorescence was recorded with 𝜆excitation=480/40 nm and 𝜆emission=527/30 nm.

We conclude that its implementation in microfluidics does not impair biosensor functionality.

### FluorMango allows for fluorescence-based sorting in microfluidics

**Objective:** We aimed at sorting active droplets based on fluorescence intensity thresholding from a mixed population of positive (with fluoride → fluorescent) and negative (no fluoride → nonfluorescent) droplets. This ‘mock library’ assay would validate the method for the prospective screening and selection of DeHa2 variants.

**Method:** Positive droplets and negative droplets generated using Method 2 (described above) were mixed at a 2:8 ratio (v/v) to create a mock library of active and inactive variants. Droplets were screened using the integrated fluorescence detector with 𝜆excitation=488 nm and 𝜆emission=525 nm and pre-sort and sort population analyzed using fluorescence microscopy.

Figure 19 shows a droplet sorting experiment which was performed under an oil pressure of 500 mbar and a sample pressure of 360 mbar to ensure efficient separation of droplets. When a droplet passing the detector exceeds the fluorescence threshold, the electrode is activated, deflecting the droplet toward the lower outlet for collection of the fluorescent droplets. Droplets with fluorescence below the threshold are directed to the upper outlet as waste.

![Figure 19](https://static.igem.wiki/teams/5755/images/results/fig19.webp)

**Figure 19: Bright-field microscopy images of the sorting area of R-Sort100.**

The flow direction and position of the fluorescence detector are indicated. Droplets are directed to one of the two outlets depending on their fluorescence intensity.

**Video 1: Droplets sorting**

**Results:** Fluorescence-based sorting of the mixed droplet population (‘mock library’) yielded time-resolved events of screened positive droplets (Figure 20). These detection hits activated the sorting of droplets that were collected in an Eppendorf tube. We assessed the sorting efficiency by imaging the collected droplets under a fluorescence microscope. The images (bright field and fluorescence) revealed an enrichment of the positive droplets from 20% (pre-sort) to 80% after sorting (Figure 21).

![Figure 20](https://static.igem.wiki/teams/5755/images/results/fig20.webp)

**Figure 20: Fluorescence detection of hits during droplet sorting.**

Events above threshold are represented as red lines on top of the graph.

![Figure 21](https://static.igem.wiki/teams/5755/images/results/fig21.webp)

**Figure 21: Microscopy images of pre-sort and sort populations showing enrichment of positive droplets.**

All the droplets are visualized with the bright-field mode (top), while active droplets (+NaF) are observed in the fluorescence channel (bottom). Fluorescence measured at 𝜆excitation=480/40 nm and 𝜆emission=527/30 nm.

We conclude that FluorMango, combined with fluorescence-based droplet microfluidic sorting, enables successful enrichment of positive droplets in a mock library containing an excess of inactive droplets. The ‘mock library’ validates this method for the prospective screening and selection of DeHa2 variants.

We then turned to enhancing the fluoride resistance of our chassis *Pseudomonas putida*, which will be bearing the expression of DeHa2 for short-chain PFAS defluorination.

## Enhancing the resistance of *P. putida* to fluoride ions with FluC

**Context:** In the PFAway process, *L. portucalensis* degrades long-chain PFAS, releasing PFPrA and TFA, whose subsequent defluorination by the evolved DeHa2 produces large amounts of fluoride ions. These ions, as well as short-chain PFAS, can inhibit essential metabolic enzymes, compromising the expression of the evolved DeHa2 by *Pseudomonas putida* KT2440 and more generally the cell’s viability. We therefore wanted to characterize and enhance the resistance to fluoride ions of our modified *P. putida* strain which will bear the expression of the evolved DeHa2.

### Successful construction of the pSEVA438-*fluC* plasmid

**Objective:** The fluoride-specific transmembrane transporter FluC endogenously expressed under the control of fluoride-responsive mechanisms was overexpressed in *P. putida* KT2440.

**Method:** The gene encoding for FluC was cloned into the expression plasmid pSEVA438 under the control of the strong *xylS*/*Pm* transcription promoter. *P. putida* was transformed with the resulting construct, pSEVA438-*fluC*. Following induction with *m*-toluic acid, the growth of the recombinant strain KT2440-pSEVA438-*fluC* was assessed in the presence or absence of 50 mM NaF by monitoring OD600 kinetics in 96-well microplates in LB medium. Whole-plasmid sequencing was performed using nanopore technology at Eurofins Genomics.

**Results:** After cloning and transformation of *P. putida*, plasmid extraction, and purification, pSEVA438-*fluC* was sent for sequencing. The alignment shows that *fluC* has successfully been cloned in pSEVA438. See the complete sequencing of pSEVA438-*fluC* in the [Materials page](https://2025.igem.wiki/toulouse-insa-ut/materials#bloc2-sub1).

We then aimed to verify that FluC was successfully overexpressed by comparing the growth curve of the recombinant strain to that of wild-type *P. putida* in the presence of NaF, both with or without induction with *m*-toluic acid (Figure 22).

![Figure 22](https://static.igem.wiki/teams/5755/images/results/fig22.webp)

**Figure 22: Growth curves and kinetics of *Pseudomonas putida* KT2440 (WT) and *P. putida* KT2440-pSEVA438-*fluC*.**

Strains were cultivated in LB medium at 30°C with 50 mM of NaF. Technical replicates (transparent circles or dashed lines) and means (solid circles or lines) are represented (n=3). A) Growth curves in the presence or absence of 500 µM *m*-toluic acid (induced or not induced). B) Specific growth rates and C) lag time for both strains, in the presence or absence of 500 µM *m*-toluic acid (induced or not induced).

Figure 22A shows that for the WT strain, there is no significant difference in fluoride resistance with or without the inducer. This suggests that the presence of *m*-toluic does not affect bacterial growth. For *P. putida* KT2440-pSEVA438-*fluC*, the lag time is reduced by 2-fold compared to the WT strain in the presence of the inducer, while the growth rate is unchanged (Figure 22 B and C). These results suggest a positive effect on fluoride stress adaptation. Further investigation to characterize these effects were pursued.

### Overexpression of *fluC* accelerates cellular adaptation to fluoride stress

**Objective:** We further investigated the effect of *fluC* overexpression on *P. putida*’s response to fluoride stress. Additional experiments were conducted in M9 minimal medium to better mimic the conditions encountered during water treatment.

**Method:** Growth of the recombinant strain *P. putida* KT2440-pSEVA438-*fluC* in M9 minimal medium was monitored in the presence of *m*-toluic acid inducer, across different NaF concentrations. Two individual carbon sources were used, acetate and propionate, as they are the expected degradation products of TFA and PFPrA. Lag time, specific growth rate, and Minimal Inhibitory Concentrations (MIC) were evaluated.

**Results:** The complete dataset and results are provided on the [Kinetic Viewer](https://devlinlucas.shinyapps.io/PFAway-KineticViewer/). The growth kinetics of the recombinant strain on both carbon sources were compared, as shown in Figure 23.

![Figure 23](https://static.igem.wiki/teams/5755/images/results/fig23.webp)

**Figure 23: Growth curves and kinetics of *P. putida* KT2440-pSEVA438-*fluC* on propionate or acetate as carbon source, in the presence of fluoride ions.**

Biological replicates (transparent circles and dashed lines) and means (solid circles and lines) are represented (n=3). A) Growth curves on M9 minimal medium supplemented with 30 mM and 45 mM of propionate or acetate respectively, in the presence of different NaF concentrations. OD600 was monitored in 96-well microplates at 30°C. B) Specific growth rates and C) lag time on both carbon sources, for different NaF concentrations. D) Comparative table of MIC of NaF on the recombinant strain.

In the absence of fluoride ions, *P. putida* pSEVA438-*fluC* grows similarly on propionate or acetate (Figure 23A). In fact, specific growth rates are equivalent with values of 0.58 h-1 on acetate and 0.63 h-1 on propionate (Figure 23B). When increasing the NaF concentration, the specific growth rates follow the same trend, and carbon source does show any influence. However, lag times for acetate are clearly reduced compared to that on propionate (Figure 23C). Overall, the MIC for NaF on acetate is clearly higher (over 100 mM) than the MIC on propionate (75 mM) (Figure 23D).

To further characterize the effect of the overexpression of *fluC* on growth, we then compared growth of the wild-type strain and the recombinant strain on acetate in the presence of different NaF concentrations (Figure 24).

![Figure 24](https://static.igem.wiki/teams/5755/images/results/fig24.webp)

**Figure 24: Growth curves and kinetics of the WT and recombinant *P. putida* KT2440 strain in minimal M9 acetate medium in the presence of NaF.**

Biological replicates (transparent circles and dashed lines) and means (solid circles and lines) are represented (n=3). A) Growth curves of *Pseudomonas putida* KT2440 (WT) and *P. putida* KT2440-pSEVA438-*fluC* in M9 minimal medium supplemented with 45 mM acetate, recorded in a 96-well microplate at 30°C under different NaF concentrations. B) Growth rate and C) lag time of *P. putida* KT2440 (WT) and *P. putida* KT2440-pSEVA438-*fluC* as a function of NaF concentration under the same growth conditions. D) Comparative table of MIC of NaF for the recombinant and WT strain.

We found that higher concentrations of NaF led to reduced growth rates (µ) and increased lag times for both strains (Figure 24). This confirms that the toxic effect of fluoride ions on bacterial growth is not confined to a threshold concentration leading to cell death, but encompasses a progressive impairment of metabolic activity ([Qin et al., 2006](https://pubs.acs.org/doi/10.1021/bi051558s)).

Interestingly, the recombinant strain was less affected than the WT strain at a given NaF concentration. *P. putida* KT2440-pSEVA438-*fluC* was able to grow at NaF concentrations up to 100 mM, whereas the WT strain only grew up to 60 mM (Figure 24A). The resulting MIC for *P. putida* WT is congruent with the MIC found in literature in M9 minimal medium supplemented with glucose, which lies in the range of 75 mM ([Calero et al., 2022](https://onlinelibrary.wiley.com/doi/abs/10.1111/1462-2920.16110)) (Figure 24D). A difference of 25 mM is then observed in the MIC analysis, showing that the overexpression of the FluC transporter enables growth in the presence of higher fluoride concentrations.

The overexpression of *fluC* also accelerated the adaptation of *P. putida* to high fluoride concentrations. Specific growth rates were less affected by NaF for the recombinant strain, and lag times were significantly reduced compared to that of the WT strain (Figure 24B and C). For example at 40 mM NaF, the lag time of KT2440-pSEVA438-*fluC* was 12 h compared to 25 h for the WT, and the growth rate was 0.32 h-1 versus 0.25 h-1, respectively. Similar trends were observed in M9 medium supplemented with propionate: data are provided in the supplementary Figure S1. The complete dataset and results are provided on the [Kinetic Viewer](https://devlinlucas.shinyapps.io/PFAway-KineticViewer/).

+ Figure S1

![Figure S1](https://static.igem.wiki/teams/5755/images/results/figs1.webp)

**Figure S1: Growth curves and kinetics of the WT and recombinant *P. putida* KT2440 strain in minimal M9 propionate medium in the presence of NaF.**

Biological replicates (transparent circles and dashed lines) and means (solid circles and lines) are represented (n=3). A) Growth curves of *P. putida* KT2440 (WT) and *P. putida* KT2440-pSEVA438-*fluC* in M9 minimal medium supplemented with 30 mM propionate, recorded in a 96-well microplate at 30°C under different NaF concentrations. B) Growth rate and C) lag time of *P. putida* KT2440 (WT) and *P. putida* KT2440-pSEVA438-*fluC* as a function of NaF concentration under the same growth conditions. D) Comparative table of MIC of NaF for the recombinant and WT strain.

Inhibition constants (*Ki*) were also calculated in order to estimate the inhibition law of *P. putida* by fluoride ions in our Model, as represented in Figure 25. See our [Model page](https://2025.igem.wiki/toulouse-insa-ut/model#bloc5-sub1) for more details on calculations and how these data were exploited.

![Figure 25](https://static.igem.wiki/teams/5755/images/results/fig25.webp)

**Figure 25: Estimation of the inhibition constants *Ki* for NaF for *P. putida* WT and *P. putida* pSEVA438-*fluC*.**

Error bar represents ± standard deviation. \* *P* value < 0.05 (=0.0183 with Graphpad two-tailed unpaired *t* test).

*Ki* were estimated for the WT and recombinant strains at 29.9 ± 6.8 mM and 57.4 ± 10.3 mM, respectively (Figure 25), indicating that the recombinant strain is less impacted by the presence of fluoride. These results were used to evaluate the inhibition law of the strains in our Model.

In our envisioned PFAway process, the recombinant strain *P. putida* KT2440-pSEVA438-*fluC* will be embedded within the beads to metabolize the PFAS degradation products generated by *L. portucalensis*. Therefore, the toxicity of the metabolites TFA and PFPrA was assessed on the recombinant strain overexpressing the FluC transporter.

### Process limits were assessed by PFPrA and TFA toxicity on *P. putida*

**Objective:** Since PFAS are toxic to cells and might determine the maximal PFOS concentration that our process will be able to treat, we assayed the minimal inhibitory concentration (MIC) of PFPrA and TFA on the fluoride resistant strain *P. putida* KT2440-pSEVA438-*fluC*. Two individual carbon sources were used, acetate and propionate, as they are the expected degradation products of TFA and PFPrA.

**Method:** The growth of the recombinant strain was assessed in the presence of TFA and PFPrA by monitoring OD600 kinetics in 96-well microplate reader in M9 minimal medium supplemented with acetate and propionate as carbon sources. Lag time, specific growth rate, and Minimal Inhibitory Concentrations (MIC) were evaluated.

**Results:** As shown in Figure 26A, *Pseudomonas putida* KT2440-pSEVA438-*fluC* was able to grow at TFA and PFPrA concentrations up to 30 mM.

![Figure 26](https://static.igem.wiki/teams/5755/images/results/fig26.webp)

**Figure 26: Growth curves and kinetics of the recombinant *P. putida* strain in minimal M9 acetate medium in the presence of TFA and PFPrA.**

Biological replicates (transparent circles and lines) and means (solid circles and lines) are represented (n=3). A) Growth curves of *P. putida* KT2440-pSEVA438-*fluC* cultivated in M9 minimal medium supplemented with 45 mM acetate and measured in a 96-well microplate at 30°C across different TFA and PFPrA concentrations. B) Growth rate and C) lag time of *P. putida* KT2440-pSEVA438-*fluC* as a function of TFA and PFPrA concentration. D) Determined MIC for TFA and PFPrA for *P. putida* KT2440-pSEVA438-*fluC*.

Increasing TFA and PFPrA concentrations caused a drop of the specific growth rate at 30 mM, while the lag time increased. For instance, lag time ranged from 5 h without TFA to 12 h at 30 mM TFA. (Figure 26C). The MIC of TFA and PFPrA for *P. putida* KT2440-pSEVA438-*fluC* were determined to be 40 mM (Figure 26D). Similar behaviors were observed under other conditions, such as growth in M9 medium supplemented with propionate for which the MIC also lies in the range of 40 mM (see supplementary Figure S2). A link to the complete dataset and results is provided on the [Kinetic Viewer](https://devlinlucas.shinyapps.io/PFAway-KineticViewer/).

+ Figure S2

![Figure S2](https://static.igem.wiki/teams/5755/images/results/figs2.webp)

**Figure S2: Growth curves and kinetics of the recombinant *P. putida* strain in minimal M9 propionate medium in the presence of TFA and PFPrA.**

Biological replicates (transparent circles and dashed lines) and means (solid circles and lines) are represented (n=3). A) Growth curves of *P. putida* KT2440 (WT) and *P. putida* KT2440-pSEVA438-*fluC* in M9 minimal medium supplemented with 30 mM propionate, recorded in a 96-well microplate at 30°C under different TFA and PFPrA concentrations. B) Growth rate and C) lag time of *P. putida* KT2440 (WT) and *P. putida* KT2440-pSEVA438-*fluC* as a function of TFA and PFPrA concentration under the same growth conditions. D) Determined MIC for TFA and PFPrA on for *P. putida* KT2440-pSEVA438-*fluC*.

## Discussion and perspectives

Our team aimed at implementing a degradation process of PFAS using two complementary bacterial strains embedded in activated carbon beads.

We first determined key parameters for the full process development as detailed in [Entrepreneurship page](https://2025.igem.wiki/toulouse-insa-ut/entrepreneurship#bloc2-sub1):

• The viability of *L. portucalensis* and *P. putida* was ensured for at least 30 days in the encapsulation beads. This indicates the lifespan of the beads before recycling, and thus the lifespan of the degradation process.

• Activated carbon showed efficient adsorption of 96.9% of PFOA when used at 10% concentration in the beads. Its presence did not compromise bacterial viability.

We also determined the maximum concentration of degradation products of PFOS (PFPrA, TFA and fluoride ions) that could tolerate the recombinant *P. putida* strain and *L. portucalensis*. Minimal inhibitory concentrations (MICs) were deduced:

**Table 1: Experimental MIC determined for *L. portucalensis*, *P. putida* KT2440 Wild Type and *P. putida* KT2440-pSEVA438-*fluC*.** These were estimated in M9 medium supplemented with acetate or glucose as carbon source for *P. putida* or *L. portucalensis* respectively.

![Table 1](https://static.igem.wiki/teams/5755/images/results/table1bisbis.avif)

In addition, we found that overexpression of the fluoride transporter FluC in *P. putida* KT2440 significantly increased the strain’s tolerance to fluoride ions, raising the MIC from 75 mM in the wild-type to >100 mM in the recombinant strain. Knowing that PFOS complete defluorination releases 17 fluoride ions, the maximum PFOS concentration that can be fully defluorinated before growth inhibition rises from 4.4 mM for the WT to 5.9 mM for the recombinant strain, representing a 34% improvement. This enhanced resistance directly extends the operational range for complete PFOS degradation, demonstrating the importance of engineering fluoride tolerance to optimize bioremediation of PFAS.

Moreover, at a 1:1 stoichiometric ratio, PFPrA and TFA are formed at concentrations up to 5.9 mM, which remain far below the MIC values for *P. putida* KT2440-pSEVA438-*fluC* (40 mM) and *L. portucalensis* (50 mM). Consequently, the accumulation of these intermediates should not hinder PFOS biodegradation under the tested conditions.

Although such high PFOS concentrations are unlikely to occur in natural environments, the increased fluoride tolerance conferred by FluC is still highly relevant. In real applications, it provides an additional safety margin against growth inhibition, ensuring robust strain performance even under conditions of accumulation of PFOS in the activated carbon beads, transient concentration peaks, or scale-up processes where fluorinated compounds may accumulate.

Further perspectives are to be explored:

+Performing PFOS toxicity assays

We attempted to test PFOS toxicity on cells, but we were limited by its low solubility in M9 minimal medium. Consequently, growth in the presence of PFOS could not accurately be measured. A possible cause was the crystallization of PFOS in the microplate wells, which interfered with the optical density measurements. Additional tests at around 5.9 mM of PFOS should be performed to ascertain its effect on both strains.

+Countering bacterial release

Minimizing bacterial release from the beads remains a critical challenge. Collaborative efforts with YpHen have focused on optimizing bead formulation to enhance encapsulation efficiency and ensure adherence to biosafety regulations.

+Engineering *Labrys portucalensis*

Engineering efforts initially targeted *Pseudomonas putida* for *fluC* overexpression and eventual expression of the evolved dehalogenase. *Labrys portucalensis* could potentially serve as a single host for both functions, simplifying the process. However, as a recently identified strain, its genetic tractability remains uncharacterized. Yet, this represents promising prospects for future development.

These considerations helped us design our large-scale implementation plan, and the collected data served for the construction of our mathematical Model. For more details, please visit our Entrepreneurship and Model pages:

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| ![Logo](https://static.igem.wiki/teams/5755/images/boutons/model.webp) Model | ![Logo](https://static.igem.wiki/teams/5755/images/boutons/entrepreneurship2.webp) Entrepreneurship |

Moreover, we built the key components for implementing our directed evolution approach to improve DeHa2. For enzyme mutagenesis via orthogonal replication, we successfully synthesized the two essential elements of the system: the replication operon and the linear replicon carrying the DeHa2 gene. Unfortunately, we did not manage to establish the functional system in *E. coli* DH10B within the project duration. Convinced that this technique could become a valuable tool for future iGEM teams, we prepared a user’s guide to facilitate its wide adoption in the community. The guide includes a schematic overview of the workflow for establishing the orthogonal replication strain, along with the materials we synthesized and are ready to share with future iGEMers. Find out about the kit [here](https://static.igem.wiki/teams/5755/images/hp/orthogonal-replication-system-user-guide.pdf).

We recommend the following experiments to prepare the ORep strain and deploy it for the evolution of DeHa2:

+Stable implementation of the replication operon in *E. coli*

We attempted to achieve integration of the replication operon through chromosomal insertion at the SS9 site, as well as expression via a single-copy plasmid. These approaches, however, did not yield the expected results, suggesting that alternative chromosomal loci or different expression strategies may need to be explored to achieve stable operon function, with the help of our guide and materials.

+Stable implementation of the replication operon in *E. coli*

We attempted to achieve integration of the replication operon through chromosomal insertion at the SS9 site, as well as expression via a single-copy plasmid. These approaches, however, did not yield the expected results, suggesting that alternative chromosomal loci or different expression strategies may need to be explored to achieve stable operon function, with the help of our guide and materials.

+Transforming the ORep strain with the linear replicon

This step is also outlined in the guide. We provided the pReplicon-KIT plasmid, which allows insertion of any gene of interest for targeted mutagenesis. This tool forms the basis for future experiments aimed at evolving specific genes within the orthogonal replication system.

+Mutation rates measurement

Once both the replication operon and the linear replicon are successfully implemented, the functionality of the system can be evaluated by quantifying mutation rates. Specifically, mutation frequencies of the bacterial chromosome, assumed to remain stable, can be compared with those of the linear replicon, which is replicated by the orthogonal DNA polymerase. This comparison provides a direct measure of the system’s ability to introduce targeted genetic variability while preserving the host genome.

Finally, we provided evidence that FluorMango could be established as a sensitive, specific, and robust fluorescent biosensor for the detection of fluoride ions. When combined with droplet-based microfluidics, this integrated pipeline will enable high-throughput sorting of active DeHa2 enzymes from pooled variants within an ORep cell population.

We suggest future experiments for the optimization of the FluorMango-based screening system and its integration in directed evolution workflows:

+Optimization of droplet generation

So far, microfluidic experiments were performed without bacteria. When moving to bacterial encapsulation, reducing droplet size would be preferable to improve the coupling between substrate degradation and fluorescence detection by FluorMango.

+Integration with ORep-based mutagenesis

Coupling the FluorMango sensor with the orthogonal replication system will enable real-time selection of improved enzymatic variants directly linked to their efficiency to degrade PFAS, thereby allowing directed evolution of DeHa2.

Find out about the complete technique on our Best Measurement page:

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| ![Logo](https://static.igem.wiki/teams/5755/images/boutons/measurement.webp) Best Measurement |

![Logo](https://static.igem.wiki/teams/5755/images/homepage/up.webp)

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