GFP Fluorescence Distribution of Cells Containing the Original Dehalogenases

It was necessary to access the effectivity of the original dehalogenases and therefore an experiment was set up using FACS and violin plots (Fig. 3). This experiment determined the distribution of fluorescence from sfGFP in samples with the different DeHAs, a sample containing RFP at the insertion site. It can be seen from the figure below that the riboswitch is inducible with fluoride in all the plasmids harboring DeHas (Fig. 4-7). Notably, it can be observed that there are often fluorescent and non-fluorescent populations within the same sample. No fluorescence induction could be observed in wild type (NEB #C2566I) strain (Fig. 8). Notably a big fluorescence population is observable at 0 mM NaF. This indicates a leakage of the riboswitch.

Selecting for efficient error-prone DeHas via FACS

To obtain a variety of different DeHas (DeHa1, 2, 4, and 5 respectively) an error-prone PCR was performed. Golden Gate was performed to insert the error-prone DeHas into the RGFr carrying plasmid, resulting in a diverse mix of plasmids with slightly different versions of the respective DeHa. Subsequently this DeHa mutant libraries were transformed into NEB T7 (NEB #C2566I) and plated out on LB-media plates with an antibiotic for selection and a PFOA layer (88 µL/cm² of evaporated 1.2 mM PFOA solution). Therefore, colonies with more efficient DeHa versions would result in higher fluorescent cells.

The plates were incubated overnight at 37°C for optimal growth of the bacteria and were then moved to 25°C, since it was estimated that the enzymes have a higher efficiency at that temperature.

The selection of our highly efficient mutated DeHas was performed using FACS. The samples were prepared by resuspending and pooling all colonies of a transformation plate, mixing them thoroughly. This was performed for each DeHa (1, 2, 4, and 5) respectively. The single top 0.1% fluorescent cells were sorted at 2000 events/second into 96-well plates. 40 variants from all DeHas that showed subsequently the highest fluorescence on the 96-well plate in the plate reader were sent for sequencing.

Furthermore, the fluorescence profile of the original DeHa populations (also grown on PFOA plates) were measured and compared to the fluorescent profile of the mutant populations (Fig. 9) The recorded data were used to compare the fluorescence distribution of each original DeHas compared to the corresponding mutated dehalogenase pool (Fig. 9).

When looking at Figure 9, a clear difference can be observed between the original DeHa and the mutated DeHa pool of DeHa 2 and DeHa 5. Based on the plots of DeHa 2 and DeHa 5 it indicates that the original DeHa 2 and DeHa 5 are not very efficient at degrading PFOA compared to the mutated counterparts, under the same conditions. However, DeHa 1 and DeHa 4 show approximately the same level of sfGFP as their mutated counterparts. Considering that there were thousands of variants generated per DeHa to begin with and only a small fraction of the pooled variants were analyzed, there is a certain likelihood that no beneficail mutations were present in this pool of mutants.

Evaluating performance of selected DeHa versions

The selected 40 variants were tested in a cell system, where each mutation variant, and the original DeHas, were plated out on agar plates with a concentrated PFOA layer in the same manner as for the plates for the FACS experiment. Fluorescence of the colonies was monitored with exposure to blue light in the GelDoc (BIO-RAD 10000125685) to determine which of the plates had the most fluorescent colonies. For further details on how this experiment was conducted, please visit our Experimental Design page. A side-by-side comparison in fluorescence of the mutated variants and original DeHas can be seen in figure 10.

The pattern that can be seen in Figure 10 matches the previously obtained FACS data seen in Figure 9. Both the mutated DeHa 2 and DeHa 5 variants show an overall increase in fluorescence compared to the original DeHa 2 and DeHa 5, whereas the fluorescence of the mutated DeHa 1 and DeHa 4 variants does not show an overall increase in fluorescence. Based on Figure 10 it can be concluded that most of the DeHa 2 and DeHa 5 mutant variants express an increased amount of sfGFP compared to the original DeHa 2 and DeHa 5. Since the expression of sfGFP comes from an increased amount of fluoride, which activates the riboswitch, it can be assumed that the increase in fluorescence is due to higher efficiency of the mutated DeHas. Whether the enzymes show the same increase in efficiency in vitro making them applicable still needs further investigation.

Insertion and sequencing of the mutant DeHa variants

Sequencing of the selected mutant variants confirmed the insertion of 7 DeHa 1 variants, 9 DeHa 2 variants, 7 DeHa 4 variants, and 7 DeHa 5 variants. When taking a closer look at the translations of the mutant variants it could be seen that a lot of the mutant variants had mutations that led to one or several stop-codons in the amino acid sequence. The amino acid sequences of the amino acids were aligned to see if there were any locations where several variants have mutations to determine sites of the enzymes that most likely are essential for the activity of the enzyme. Figure 11 shows an alignment of the DeHa 1 variants, where the stop codon in DeHa1_var1 and the regions with mutations are highlighted.

The same alignment was done for the DeHa 2 variants (Figure 12). There were however no DeHa 2 variants that didn’t have a stop codon. Furthermore, DeHa2_var3 and DeHa2_var4 had frameshift mutations, which is why they were excluded from the alignment. The frameshift mutations might however still have had a gain-of-function impact on the enzyme structure, since the fluorescence experiment showed high fluorescence in DeHa2_var4. All of the stop codons and positions where multiple variants have mutations are highlighted in Fig. 12

In the alignment of DeHa 4 and its variants (Figure 13), multiple variants had frameshift mutations and those were excluded from the alignment. Since there also was not seen an increase in fluorescence of the mutated DeHa 4 variants compared to the original DeHa 4 the other DeHas were prioritized for further analysis.

Two of the DeHa 5 mutant variants had frameshift mutations which is why they were excluded from the alignment (figure 14). Furthermore, the overall mutation rate was lower compared to the other DeHas. The DeHa 5 mutant variants did however show the biggest increase in fluorescence compared to the original DeHa 5.

Though the sequencing results show many variants containing one or more stop codons it does not necessarily mean that the mutated variants DeHas will not be fully transcribed or functional as enzymes. For example, it has been reported that bacteria (such as E. coli) have varying UGA readthrough, which means that bacteria can in certain situations transcribe past a stop codon⁽²⁾.

Other studies supported varying efficiencies for E. coli to terminate transcription. Furthermore, another study suggested that the base pair following the stop codon dictates whether transcription halts or proceeds³. Other studies suggest that the stop codon is nothing more than an indication of where to terminate the signal and there are many other factors needed for transcription termination⁴.

There is also a possibility that the enzymes could be transcribed until the stop codon, and folded, while maintaining their enzymatic pocket and thereby still be functional enzymes. Therefore, the stop codons are not to be seen as determining factors for the possible further investigations of the mutated variants enzymes. Additionally, the sequencing results revealed the cells showing the most fluorescent when running them in FACS, had in fact mutations forming stop codons in their sequence.

Additionally, docking was used to simulate the binding of PFOA to the mutated DeHa variants. See more on this link.

Protein purification

For testing the DeHas in a cell-free system they had to be purified. First, there had to be proven DeHa expression. This was done by purifying them using a His-tag purification kit (Cube Biotech 80101) and then running the purified proteins on an SDS page (Fig. 15). DeHa 1 and DeHa 5 were run on a gel together and DeHa 2 and 4 on another gel, as seen on figures 15. As expected, the bands for Deha 4 and Deha 5 are visible at ~34 kDa and ~25 kDa respectively. Conversely, for Deha 1 and DeHa 2, there were no clear bands at the expected size of 26 kDa. The expected protein sizes can be seen in table 1. Likely the enzymes were nonetheless expressed, but the purification protocol remains to be optimized for these enzymes, as they neigher could be found in the flow-through or wash fractions of their respective purification (data not published).

If there was more time at disposal, it would have been possible to optimize the purification process using different temperatures and different elution solutions for instance.

Table 1) Overview of molecular weight of RFP and dehalogenases 1, 2, 4, and 5, both with and without a 6x His-tag.

	Mol. weight (kDa)	Mol. weight w/ 6x His-tag (kDa)
RFP	25.30	-
DeHa 1	25.69	26.53
DeHa 2	25.51	26.35
DeHa 4	33.6	34.44
DeHa 5	24.14	24.98

F-NMR

Fluorine-19 nuclear magnetic resonance spectroscopy (19F-NMR) was used to prove the enzymes did break down PFOA. For this purpose, one sample was taken after 80 hours, 68 hours, and 44 hours. 19F-NMR-spectra were obtained. The spectra can be seen below, in figure 16:

Unfortunately, there were no differences in the spectra when they were compared to the samples with PFOA. This indicated that the enzymes did not work on PFOA and therefore no C-F bonds were broken. This could have been due to the low concentration of enzyme (5 µg/mL), or the enzyme unfolded before they were able to interact with PFOA. One explanation could be that the enzymes have a degradation rate bigger than 80 hours. Additionally, temperature optimization could be investigated. The protocol for the used kit for protein purification could likewise be adjusted.

Supplementary results

Adaptive Laboratory Evolution

If you are interested in the Whole Genome Sequencing results for the ALE1 and ALE2 strains, please contact: sofielmoebes@gmail.com

Prior to the plasmid assembly it was hypothesized that it would benefit the project if the bacteria could tolerate an increased amount of fluoride, as fluoride ions are released during the enzymatic degradation of PFOA. So, the first idea of the project was that the bacteria could be integrated into a closed system that could work as a live PFAS degradation filter. The aim was to enable the bacteria to tolerate a NaF tolerance of 200 mM.

Therefore, a side project was initiated aimed to create an E. coli with increased fluoride tolerance. However, the project and the corresponding results did not meet the criteria, and the side project was discontinued.

We aimed to increase this tolerance to 200 mM NaF using adaptive laboratory evolution (ALE). In the first ALE experiment, the bacteria were cultivated in a 96-well plate containing LB media with different NaF concentrations. Over a 4-week period, we inoculated a new 96-well plate containing a NaF dilution row with an aliquot of the well from the day before. We will refer from here on to the strain obtained through this ALE as ALE1. We attempted another ALE-experiment in which we grew the bacteria on LB-agar plates containing NaF. Each day, we streak-plated colonies onto fresh plates a higher NaF concentration than the day before. We refer to the mutant strain obtained from this experiment as ALE2 from here onwards. For an overview of the experimental setup, please refer to figure 9S.

To conclude the experiment, the growth of both ALE1 and ALE2 on different concentrations of NaF, ranging from 0 mM to 400 mM, was compared to that of the original Rosetta (Fig. 10S). The three different strains were plated out in a 1:10 serial dilution from a factor 1 to a factor of 10^-7, on LB agar plates containing 0 mM, 50 mM, 100 mM, 150 mM, 200 mM, 250 mM, 300 mM, 350 mM, and 400 mM NaF. Overall, there are no remarkable differences in bacterial growth between the strains on plates with low (0 mM and 50 mM) NaF. On plates which contain 200 mM NaF a distinct difference in growth could be observed between the two strains. ALE1 does not show any clear difference in growth to the original Rosetta strain, with both only showing growth down to a dilution of 10^-2 at a concentration of 200 mM. ALE2, conversely consistently grew a clear lawn down to a dilution of 10^-3 at a concentration of 200 mM.

At concentrations above 350mM NaF, none of the cells grew.

At conclusion the ALE strains were not suitable for the purpose of this specific study. They, however, exhibit significant potential in studies that rely on the tolerance of E. coli to fluoride.

From the results above, it can be concluded that ALE2 does have increased fluoride tolerance, compared to the original Rosetta strain. Unfortunately, fluorescence microscopy images show that ALE1 and ALE2 are unsuitable for selection of optimized dehalogenases. Please see figure 11S for comparison of fluorescence of ALE1, NEB and original Rosetta.