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Wet lab results

Rhamnolipids synthesis :

  • Conjugation :

After receiving our four strains, a donor E. coli strain with the plasmid carrying the Mini-Tn7 transposon, two helper E. coli strains, and the P. putida KT2440 recipient strains, we carried out conjugation for three days to obtain a KT2440 strain that had integrated the transposon carrying the gentamicin resistance gene and the two genes encoding the RhlA and RhlB proteins, enabling rhamnolipid synthesis by Pseudomonas putida. It should be noted that the transposon integrates specifically on the att sites of the bacterial genome, in an operon that carries genes involved in cell membrane biosynthesis and therefore constitutively expressed. Once conjugation is complete, gentamicin-resistant strains producing rhamnolipids are expected.

Figure 1: Photo of our isolation medium with our different bacteria. On the left, these are the cetrimide plate and on the right these are the cetrimide + gentamicin plate. In the first row, these are the contaminated tube of E.coli DH5α (Pb) (left of the plate) and the stored ones (right of the plate). In the second row, these are P.putida WT (left of the plate) and the conjugates (right of the plate). In the last row, these are the two E.coli helper strain.

Cetrimide is a nitrogenous antiseptic molecule acting as an antibacterial. However, Pseudomonas bacteria are known to be resistant to this type of nitrogenous antiseptic, so cetrimide plates can be used for species-specific selection. We grew all the strains we had in the laboratory on cetrimide plates, with or without gentamicin, to check that helper and donor E. coli were not growing, and to isolate correctly transformed clones. As expected, the wild-type (KT2440) grew on cetrimide without antibiotics and did not grow with gentamicin, and our modified strains (RL) grew in both cases (middle row, right and left boxes). Unsurprisingly, the helper strains (PTRII and HB101) did not grow on cetrimide plates (bottom line, right and left boxes). On the other hand, one of our DH5a cultures was contaminated by a transformed KT2440 strain, since the plates grown with DH5a contained bacteria that were supposed to be unable to grow on this medium. After re-cultivating this strain on new cetrimide plates (top row, right and left plates), we observed that the strains from our contaminated culture (Pb) could grow, but the original strains as received could not (WT), except on the right-hand dish, where E. coli DH5a grew slightly, but not in the presence of gentamicin. In addition, we noticed that the P. putida KT2440 bacterial pellets obtained after centrifugation showed a reddish pellet, unlike the E. coli with a whitish pellet (not shown here). This test served as a further argument that the contamination was due to P. putida, since the pellets of the (Pb) strains were red, but not the DH5a from the original tube. This setback nevertheless taught us to handle the bacteria with much greater care, given its mobility, and the problem never recurred thereafter. From these cultures, we isolated 4 gentamicin-resistant transformed KT2440 strains for subsequent rhamnolipid production tests.

Achieved to have a mini-TN7 in the genome are not the goal of our technique and then, we had to test the rhamnolipid production.

Choi, KH., Schweizer, H. mini-Tn7 insertion in bacteria with single attTn7 sites: example Pseudomonas aeruginosa. Nat Protoc 1, 153–161 (2006). https://doi.org/10.1038/nprot.2006.24.

  • Hemolytic test:

In the course of our research, we discovered that biosurfactants such as rhamnolipids can have a hemolytic action. We tried to detect the production of rhamnolipids by highlighting this hemolytic capacity. To do this, we inoculated our transformed Pseudomonas putida on sheep blood agar plates, hoping that the rhamnolipids would degrade the blood through their hemolytic action and form a halo around the colonies. However, when we tried this technique, no halo was visible, indicating that rhamnolipid production was not effective or that this technique is not suitable for rhamnolipid detection. We repeated these experiments three times and no halo was ever observed, even after several days.

Figure 2: Culture of Pseudomonas putida KT2440 strains on sheep blood agar (7.5%) after 5 days at 30°C: from left to right, WT strain, strain modified to produce rhamnolipids RL1-RL4 and strain RL1 but in culture without glucose.

  • Coomassie blue dye :

Figure 3: Right: control curve obtained from an overnight culture of a wild type strain which does not produce rhamnolipids. Left: superimposed curves resulting from the analysis of the supernatant of all overnight cultures of rhamno-producing strains (in black RL1, in red RL2 and in blue RL3).

The methylene blue assay method is a quick and simple method for analyzing rhamnolipids by studying the complexation of rhamnolipids and methylene blue. The method is based on measuring the absorbance (at 638 nm) of the rhamnolipid-methylene blue complex which is distributed in a chloroform phase. Following the extraction of extracellular lipids with the chloroform phase, we produced absorption spectra between 200-700 nm of this chloroform phase. We emphasize that in the article on which we based ourselves, they obtain the same spectra shapes, leading us to believe that the experiment was correctly carried out. The first thing we observe is the absorption peak at 638nm which therefore hypothetically corresponds to rhamnolipids. On the other hand, this absorption peak is present for the same absorbance (approximately less than 0.500) both in the control (left) and for the strains supposed to produce rhamnolipids (right), which leads us to believe that there is clearly no production of rhamnolipid, and that this absorbance is perhaps due to the imperfect sensitivity of the method which could detect other types of extracellular lipids. We specify that we repeated these experiments three times, in particular by varying the glucose concentration, and obtained similar results.

Pinzon, N.M., Ju, LK. Analysis of rhamnolipid biosurfactants by methylene blue complexation. Appl Microbiol Biotechnol 82, 975–981 (2009). https://doi.org/10.1007/s00253-009-1896-9.

Sophorolipids synthesis :

  • Transformation :

Gibson assembly is a molecular biology technique used to assemble several DNA fragments into a larger DNA molecule. It is a powerful and efficient method for creating recombinant DNA constructs, such as plasmids, synthetic genes or other DNA constructs for genetic engineering or molecular biology research. Gibson assembly offers several advantages, including its ability to join DNA fragments without the use of restriction enzymes or ligases, and its capacity to seamlessly combine multiple fragments with overlapping sequences. This is why we decided to use this technique to create our plasmid. We discovered that 4 enzymes were required for sophorolipid synthesis: Cytochrome P450 CYP52 M1, UDP-glucosyltransferase A1 (UGTA1), UDP-glucosyltransferase B1 (UGTB1) and Lactonase (or Lactone esterase) These four enzymes are naturally produced in eukaryotic cells such as Starmerella bombicola. Our aim was to engineer P. putida to induce the synthesis of these four proteins, leading to the production of sophorolipids. To this end, we turned to Pseudomonas expert Dr Victor Delorenzo. After several meetings with him, we set about designing a plasmid incorporating the genes for these various enzymes. To create this plasmid, we asked Dr De Lorenzo to supply us with plasmid pSEVA438, which would serve as our backbone. We also designed different biobricks containing each gene required for sophorolipid synthesis. First, we attempted to linearize pSEVA438 by PCR. After numerous attempts under different conditions, we were unable to linearize the plasmid using this technique (see photo of gels). We therefore decided to linearize the plasmid with restriction enzymes. Next, we used Gibson's assembly protocol to fuse our different fragments and create our plasmid for sophorolipid production. Finally and after many trials, we successfully transformed Pseudomonas putida by electroporation. We also tried heat shock, and found that this technique resulted in less bacterial death.

Figure 4 : Bacteria shown here are grown in LB in the presence of streptomycin 30mg/L: WT KT2440 (left tube), transformed bacteria from electroporation (middle tube and right Erlenmeyer flask).

  • - SDS-Page and Western-Blot:

After bacterial transformation, we wanted to check whether all four proteins were expressed. To do this, we analyzed protein expression in SDS-page electrophoresis and transformed pseudomonas after inoculation with 0mM, 1mM, and 2mM salicylate for 4 hours. We stained our gel with Coomassie stain and observed different profiles between SDS-page and transformed pseudomonas. The appearance of bands between 50 and 75 kDa for transformed Pseudomonas could demonstrate the presence of new transformation-related proteins. Indeed, the WT strains, which were grown separately for 4H, all three (Figure ?, the three left-hand columns) show the same migration profile, indicating that the results are robust and reproducible. Similarly, the transformed strains have exactly the same migration profile. On the other hand, the profiles vary between strains, with some bands being highly concentrated in the wild-type but not in the transformed strains, and vice versa. Most interesting are the four bands located between 75 and 50 kDa in the profile of the transformed bacteria, as they correspond approximately to the molecular weight of the proteins contained in the sophorolipid plasmid (CYP52 61.8 kDa, UGTA1 50.5 kDa, UGTB1 46.1 kDa and lactone esterase 45.6 kDa). It should also be noted that our bacteria carry TAGs of a few kDa (less than 5kDa), which could vary migration slightly. Nevertheless, for the profile of transformed bacteria, there was no difference in the concentration of these bands in relation to the concentration of salicylate inducer. To explain this lack of induction, we may hypothesize that our promoter has a fairly strong basal activity (the XylS/Pm promoter being activatable with many different molecules (Morgenroth et al.) and that salicylate did not activate the promoter in any measurable way, or else that the plasmid is not functioning. These results are encouraging, but we need to go further to determine whether the bands correspond to our four proteins. We had to design gene sequences with tags to easily target our neo-proteins. We didn't have time to carry out the Western blot for the wikifreeze but will do so before the Grand Jambore.

Figure 5: Result of Coomassie blue staining of SDS-Page carried out on six conditions, three control conditions with a gradual induction of "no induction," "1mM salicylate," and "2mM salicylate" and three conditions transformed with the plasmid sophorolipid with the same gradual induction. Left well: BioRad dual color ladder.

The future experiments :

  • Sophorolipds dosage :

In our upcoming experiment, we will be utilizing High-Performance Liquid Chromatography (HPLC) to quantify the levels of sophorolipides in our samples. HPLC is a powerful analytical technique that is frequently used in the separation and quantification of complex mixtures. It will allow us to determine the concentration of sophorolipides produced by our modified Pseudomonas putida strains, giving us a precise measurement of our biosurfactant yields. This data will be crucial in assessing the effectiveness of our genetic modifications and in refining our strategies for enhancing sophorolipide production.

  • Rhamnolipds dosage :

To determine the production of rhamnolipids, we will perform a rhamnolipid assay. This will involve culturing our genetically modified Pseudomonas putida strains and extracting the rhamnolipids from the culture medium. We will then employ a rhamnolipid assay kit to quantify the rhamnolipid concentration in the samples. This will offer us a quantitative measurement of rhamnolipid production, aiding us in assessing the performance of our engineered strains. Additionally, we will investigate whether modifying various culture conditions can further enhance rhamnolipid production.

  • Combined Biosurfactant Production :

Our overarching goal is to create strains of Pseudomonas putida that are capable of producing both sophorolipides and rhamnolipides. This dual biosurfactant production has the potential to offer a broader range of applications and benefits, such as enhanced oil recovery and improved bioremediation. We will explore the feasibility of engineering strains that can simultaneously produce both biosurfactants by integrating the necessary genetic components for both pathways. This will be a challenging but rewarding endeavor, as it has the potential to open up new opportunities in the field of biotechnology.

  • Advanced Characterization :

In our future experiments, we will dive deeper into the characterization of the sophorolipides and rhamnolipides produced by our modified strains. This includes analyzing their surface-active properties, emulsification capabilities, and stability under various environmental conditions. Such in-depth characterization is essential for understanding the potential applications and performance of these biosurfactants in different industrial and environmental settings.

  • Application Development :

Our ultimate aim is to harness the biosurfactants produced by our engineered Pseudomonas putida strains for real-world applications. We will explore the use of sophorolipides and rhamnolipides in various fields, including bioremediation, enhanced oil recovery, and pharmaceuticals. Through collaborative efforts with industry partners and research organizations, we will work towards developing practical applications that can benefit society and the environment.