Results FitD

Overview

FitD is a toxin naturally produced by Pseudomonas protegens. The fitD gene is part of a virulence operon which is regulated by three regulatory proteins, including an activator encoded by the fitG gene and a type I secretion system. The FitD toxin has been shown to have insecticidal activity by inducing the midgut epithelium's apoptosis and paralyzing and destroying insect phagocytes (Péchy-Tarr et al., 2013). Interestingly for our project, FitD is also known for its potential as a molluscicidal agent (Molloy et al., 2013).
Hence, we tried to overexpress the FitD protein in P. protegens using plasmids bearing either an extra copy of the fitG or fitD genes. By overexpressing fitD or its activator fitG in P. protegens, we expected that it would impact the survivability of quagga mussels.

explanation of FitD
Figure 1: Plasmid maps of the constructed plasmids for the overexpression of the FitD toxin in P. protegens.

Construction of the overexpression fitD and fitG plasmids

To improve FitD production in P. protegens, we thought of overexpressing the toxin by either (a) introducing simply an extra copy of its gene (fitD) on a plasmid, or (b) introducing an extra copy of its activator fitG. Considering that expressing extra copies of these genes might be toxic or represent a significant metabolic burden on our cells, we designed vectors bearing both genes under the regulation of a constitutive or inducible promoter (Figure 1). We chose pSEVA plasmids as backbones for our cloning as they are designed to be compatible with Pseudomonas bacteria. Gibson assembly was used to clone either the fitD or fitG gene in our vector of choice. The inducible plasmids are based on the promoter lacIq-Ptrc, which IPTG can induce. The constitutive plasmids are based on the promoter EM7. Ultimately, the following vectors were designed:

  • pSEVA234 + fitD
  • pSEVA234 + fitG
  • pSEVA2313 + fitD
  • pSEVA2313 + fitG

pSEVA234 refers to an IPTG inducible plasmid and pSEVA2313 refers to a constitutive plasmid.

The fitD and fitG genes were PCR amplified using the genomic DNA of P. protegens as template. The obtained gene fragments were cloned into their respective backbone through Gibson assembly and were transformed into E. coli DH5α. After plasmid extraction, the constructs were transformed into P. protegens.By performing colony PCR and subsequent sequencing on the resulting transformants, we also were able to confirm the successful cloning of all of our plasmids (Figure 2).

explanation of FitD
Figure 2: Electrophoresis gel confirming the correct assembly of our fitD and fitG constructs in P. protegens by colony PCR scale is given by 1kb plus ladder, A) P. protegens pSEVA2313 fitG shows a band at the expected size of 1968 bp, B) P. protegens pSEVA234 fitG shows a band at the expected size of 3356 bp, C) P. protegens pSEVA2313 fitD shows a band at the expected size of 3480 bp, D) P. protegens pSEVA234 fitD shows a band at the expected size of 3480 bp.

Validation of the killing activity of our engineered P. protegens against quagga mussels

After a trial and error process (see Engineering) we successfully transformed P. protegens CHA0 with our newly-built constructs. We then set out to test the efficacy of the obtained four engineered P. protegens strains in killing quagga mussels. After testing various conditions and troubleshooting measurements, we initially assess the killing activity of our strains on 10 mussels using a final lysed bacterial concentration of 8·108 cells/mL of water (Figure 3).

explanation of FitD
Figure 3: Lysed and engineered P. protegens kill quagga mussels. Kaplan-Meier graph shows probability of mussel survival over time when subjected to different lysed bacterial treatment applied to a concentration of 8·108 cells per mL. Gehan-Breslow-Wilcoxon test, * significant for p-value < 0.05, n = 10 mussels.

The data obtained showed that our engineered P. protegens strains were efficiently killing quagga mussel (Figure 3). While the rate of killing appeared extremely similar between the wild–type P. protegens and our strain P. protegens pSEVA2313 fitD, we observed a trend suggesting our strain P. protegens pSEVA2313 fitG has better activity than the wild-type. However, the bacterial concentration used was relatively high (8·108 cells per mL) and 10 mussels was too little to have enough statistical power to detect significant differences.
We therefore decided to increase the number of mussels tested per condition to 45 as well as to test the efficacy of a lower concentration of cells. Since the workload was exponentially increasing with the number of mussels we were planning to use, we also chose to focus our experiments on the most promising strains only for further testing. As one of the strains is the inducible fitD, we wanted to use a strain without an inducer as another control. We ended up testing the following strains:

  • P. protegens pSEVA234 fitD induced with IPTG
  • P. protegens pSEVA234 fitD without IPTG
  • P. protegens pSEVA2313 fitG (constitutive)
  • Wild type (i.e containing fitD in the genome)

We also tested a lower concentration of cells applied to the mussel in order to see if we could limit the amount of product needed and still significantly improve the killing of mussels (Figure 4).

explanation of FitD
Figure 4: Lysed P. protegens cells engineered to overexpress the fitG gene show improved killing of quagga mussels compared to wild-type P. protegens. Kaplan-Meier graph shows probability of mussel survival over time when subjected to different lysed bacterial treatment applied at a concentration of 2·108 cells per mL. Gehan-Breslow-Wilcoxon test, * p-value < 0.05, n = 45 mussels.

We were pleased to observe that our engineered and lysed P. protegens cells overexpressing the fitG gene were able to significantly reduce the probability of survival of the tested mussels (p-value 0.0117; Figure 4). Indeed, our data show that the probability of survival of the quagga mussels decreases remarkably to 86 % after 80 hours when exposed to our engineered strain, compared to the wild-type P. protegens did not impact the mussel’s fitness. Although we could not measure a statistically significant difference in the probability of survival of the mussels between the wild-type P. protegens treatment and our engineered strain overexpressing the fitD gene, we noted that the latter, however, seems to reduce the mussel survival as well modestly. We would therefore argue that both our modified P. protegens strains, expressing fitD or fitG, are able to kill the invasive quagga mussels.
Importantly, at the lower final bacterial concentration of 2·108 cells per mL tested here, the wild-type P. protegens cells do not seem to affect quagga mussels at all, whereas our engineered cells demonstrate the killing of mussels. These results show that our constructs significantly improve the quagga mussel-killing capacity of P. protegens and would allow its use at low concentrations which could prove to provide a cost-effective alternative to the currently marketed Zequanox product (see proof of concept).

Results Zosteric Acid

Overview

Zosteric Acid (ZA) has been proven to prevent foot attachment of mussels to surfaces (Ram et Al., 2012). We sought to produce heterologously ZA in Escherichia coli by introducing a two-step enzymatic pathway such that tyrosine is first converted into coumaric acid (pHCA) and then pHCA is converted to ZA by the addition of a sulfate group (Figure 1). Therefore, another set of genes, cysDNC, cysQ, cysPUWA and cysP, must additionally be added to the pathway as they are responsible for the entry and the transport of sulphate into the cell (cfr design)(Jendresen & Nielsen, 2019).

explanation of FitD
Figure 5: Schematic representation of the ZA production pathway. The TAL enzyme converts the tyrosine, naturally present in the bacteria, into pHCA. CysP or CysPUWA are responsible for the uptake of sulphate into the cell. The CysDNC enzymes replace two phosphate groups of ATP by the sulphate. The sulphate carrier is used by SULT1A1 to incorporate sulphate into coumaric acid creating zosteric acid. CysQ molecule is essential to recycle ATP and allow for a second cycle to start.

Construction of the zosteric acid biosynthesis plasmids

a. Gene amplification:

explanation of FitD
Figure 6: Plasmid maps showing the constructed plasmids used for ZA production. A) and B) represent the 2 plasmids containing the catalytic enzymes tal-fjo for the conversion of tyrosine into pHCA and SULT1A1 for the conversion of pHCA into ZA. C) and D) represent the 2 plasmids containing sulphate transporter genes cysDNCQ for both, responsible for the transport of sulphate inside the cell, and two different genes (cysP and cysPUWA) for the entry of sulphate inside the cell.

Ultimately, we designed our pathway to be split onto two separate vectors with one ‘transport’ plasmid, bearing the genes responsible for the intake of sulphate, and one ‘catalytic’ plasmid, bearing the genes responsible for the synthesis of ZA (Figure 2). We obtained the genes for our transport vector, cysDNC, cysQ, cysPUWA and cysP, by PCR-amplifying them all from the genome of E. coli but for the latter, which we got from the genome of Bacillus subtilis. The tal-fjo and SULT1A1 genes necessary for the catalysis of the two conversion reactions (tyrosine to pHCA, and pHCA to ZA, respectively) were ordered from Twist Bioscience. The ‘transport’ genes were to be cloned into the pCola_Duet vector, while the ‘catalytic’ genes were to be inserted into the pET17b plasmid (Experiments).

explanation of FitD
Figure 7: A) agarose gel electrophoresis of amplified genes. Expected sizes: 1=cysP (1017bp), 2=cysQ (741bp), 3=cysPUWA (3812bp), 4=cysDNC (2943bp), 5=SULT1A1 (876bp), 6=tal-fjo (1520bp). B) agarose gel electrophoresis of amplified inter-regions: 1= Inter region of pCOLA (135bp), 2= Inter region of pET17b-S-T (157bp), 3=Inter region of pET17b-T-S (150bp). C) agarose gel electrophoresis of the amplified sequences: 1=pCola_Duet inter-region (135bp), 2= pCola_Duet backbone (3.3Kb). D) agarose gel electrophoresis of the amplified sequences: 1= pET17ST BB (Backbone for the insertion of SULT1A1 and tal-fjo), 2=pET17TS BB (Backbone for the insertion of tal-fjo and SULT1A1), 3 = LacI gene for SULT1A1 and tal-fjo plasmid, 4= LacI gene for tal-fjo and SULT1A1 plasmid.

Ultimately, after thorough primer design for all the parts we wished to amplify, we successfully PCR-amplified all the genes and linear backbone fragments required to perform our cloning (Figure 3).

b. Gene assembly:

The obtained gene fragments were cloned into their respective backbone through Gibson assembly and were transformed into E. coli DH5α. By performing colony PCR and subsequent sequencing on the resulting transformants, we also were able to confirm the successful cloning of all of our plasmids (Figure 4).

explanation of FitD
Figure 8: Gel electrophoresis of colony PCR performed on E.coli DH5a transformed with our plasmids. A) agarose electrophoresis of colony PCR performed on the second MCS of both transporter plasmids containing the genes cysDNCQ. Expected band size= 3.7Kb. Observed band size=~3.7 Kb. B) agarose electrophoresis of colony PCR performed on the first MCS of one of the two transporter plasmids containing the gene cysP. Expected band size= 1Kb. Observed band size=~1Kb. C) agarose electrophoresis of colony PCR performed on the first MCS of one of the two transporter plasmids containing the gene cysPUWA. Expected band size= 3.8Kb. Observed band size=~3.8Kb .

The newly-built plasmids were then co-transformed into E. coli BL21 (DE3), a strain more adapted for protein production. Indeed, we wished to generate four distinct strains, each containing a different combination of two plasmids, one bearing sulfate uptake genes and one bearing the catalytic enzymes. Unfortunately, we were not able to co-transform E. coli BL21 (DE3) containing pCola_Duet_P_DNCQ and pET_17b_Tal_SULT1A1. We, however, successfully obtained the three following co-transformant that we analyzed further for ZA production (Fig. 5).

  • pCola_Duet_PUWA_DNCQ + pET_17b_Tal_SULT1A1 (PUWA-TS)
  • pCola_Duet_P_DNCQ + pET_17b_SULT1A1_ Tal (P-ST)
  • pCola_Duet_PUWA_DNCQ + pET_17b_SULT1A1_ Tal (PUWA-ST)

Zosteric acid production from our engineered strains

To test the ability of our engineered strains to produce ZA, we used HPLC to analyze the supernatant of our cells grown in liquid cultures to detect ZA and pHCA.

a. Expected results from HPLC:

We first purchased the pure compounds ZA and pHCA and ran them into the HPLC as standards (Fig 5). This allowed us to confirm that we could detect ZA and pHCA with our method, with a retention time (RT) of 5 and 9 min, respectively.

explanation of FitD
Figure 9: HPLC chromatograms of standards for Za and pHCA. A) Zosteric acid and B) pHCA.

Next, we analyzed the supernatants of our three mutant strains to assess ZA production. To do so, we cultured overnight the strains in M9 medium supplemented with 4 mM of pHCA, 10 mM of K2SO4-, and 0.1 mM IPTG induction. When supplied with all molecules, we expected our engineered cells to utilize pHCA and produce ZA, resulting in an absorbance peak at 5 min RT corresponding to ZA and the reduction of the peak at 9 min RT for pHCA, for a non-induced control (Fig.6).
Indeed, as a reference control, we additionally analyzed the supernatant of cells grown with the substrates pHCA and K2SO4 (Figure 6) but without IPTG induction. In these conditions, we expected to observe no production of ZA but a significant amount of pHCA corresponding to the added initial concentration that cannot be utilized by the cells.
Lastly, we prepared a second control for each mutant strain where we added IPTG but no supplementation of pHCA and K2SO4. The rationale of this control was to assess the efficacy of the tal-fjo gene-encoded protein to convert tyrosine into pHCA effectively. Without added sulfate and pHCA, we reasoned that any detected pHCA would be the product of the tal-fjo protein activity. pHCA would most likely accumulate as no added sulfate would limit its further conversion to ZA. Here, we expected to observe the production of pHCA and virtually no ZA. In the case where the basal level of sulfate present in the M9 media used for our experiment would be sufficient for conversion of ZA, we would then expect minimal production of ZA and a reduced amount of pHCA (Figure 6).

explanation of FitD
Figure 10: Schematic representation of the testing conditions. The upper part of the image shows the E. coli strains that we constructed and tested. The table in the figure represents the specificity for each test and control condition. As we can see, in every test condition, the M9 media has been implemented with 4mM of pHCA, 10mM of K2SO4, and induced with 0.1mM. Control 1 media has been implemented with 4mM of pHCA, and 10 mM of K2SO4, but no IPTG was added; the plasmids are therefore not induced. Control 2 has no implementation of chemicals. Instead, the plasmids have been induced with 0.1mM of IPTG. The two last columns represent the expected results for each condition: ZA should be produced in Test and Control 2, and pHCA should be detected in Control 1.

b.Observed HPLC results:

After a few preliminary analyses to determine the best conditions under which to perform the HPLC, the following results were obtained:

explanation of FitD
Figure 11: HPLC analyses of our modified E. coli strains’ supernatant after overnight culture in M9. A) Decrease of pHCA compared to the negative control. Analysis of each transformed BL21(DE3) strain’s (PUWA-TS, P-ST, PUWA-ST) supernatant in test condition : with addition of 4mM pHCA, 10mM SO4- and induction with 0.1mM IPTG. B) Negative control showing the amount of pHCA that was added at the beginning of the incubation. Analysis of each transformed BL21(DE3) strain’s (PUWA-TS, P-ST, PUWA-ST) supernatant with addition of 4mM pHCA, 10mM K2SO4- but no IPTG induction (negative CTRL). C) Production of coumaric acid. Analysis of each transformed BL21(DE3) strain’s (PUWA-TS, P-ST, PUWA-ST) supernatant without implementation of chemicals (pHCA, K2SO4-) but with 0.1mM IPTG induction.

Indeed, we analyzed the supernatant of our three different strains, each bearing a different combination of the transport and catalytic plasmids. We, however, observed the strongest signal with the P-ST and PUWA-ST co-transformants. For clarity, we decided to report only the results obtained for these strains (Fig. 7). The absorption profile of the supernatant isolated from the PUWA-ST strain showed that in control 2 growth conditions, cells could convert tyrosine into pHCA as a great peak with the correct retention time can be observed while the molecule was not added here (Fig. 7C, PUWA-ST-CHEM). However, comparing the test and control 1 condition for this strain, no decrease of pHCA and detection of ZA can be seen, which suggests that no pHCA is being utilized. Altogether, these observations indicate that the tal-fjo is functional in producing pHCA but the PUWA does not appear to allow significant sulfate uptake to permit conversion of pHCA to ZA. On the other hand, the P-ST strain showed no detectable pHCA in control 2 conditions, while we observed a clear decrease in pHCA between the test and control 1 condition when it was added in the culture. Although we do not detect a peak for ZA itself, we argue that this data suggests that ZA is being produced, albeit below the detection limit of the method used, as pHCA decreases specifically when the cells are induced. We thus considered a notable decrease in pHCA as a proxy for ZA production. Comparing our two strains, we can also conclude that the CysP protein seems more functional than the CysPUWA protein in sulfate uptake, which is effectively used to convert pHCA to ZA, as the decrease of pHCA could only be detected with the cysP gene.
In conclusion, we are pleased to report that our data so far suggest our P-ST strain can convert pHCA to ZA. Further work will, however, be carried out to improve our growing conditions and the detection of ZA to confirm this.

References

  1. Péchy-Tarr, M., Borel, N., Kupferschmied, P., Turner, V., Binggeli, O., Radovanovic, D., … Keel, C. (2013). Control and host-dependent activation of insect toxin expression in a root-associated biocontrol pseudomonad. Environmental Microbiology, 15(3), 736–750. https://doi.org/10.1111/1462-2920.12050
  2. Molloy, D. P., Mayer, D. A., Gaylo, M. J., Morse, J. T., Presti, K. T., Sawyko, P. M., … Griffin, B. H. (2013). Pseudomonas fluorescens strain CL145A - a biopesticide for the control of zebra and quagga mussels (Bivalvia: Dreissenidae). Journal of Invertebrate Pathology, 113(1), 104–114. https://doi.org/10.1016/j.jip.2012.12.012
  3. Ram, J. L., Purohit, S., Newby, B. Z., & Cutright, T. J. (2012). Evaluation of the natural product antifoulant, zosteric acid, for preventing the attachment of quagga mussels – a preliminary study. Natural Product Research, 26(6), 580–584.https://doi.org/10.1080/14786419.2010.541873
  4. Jendresen, C. B., & Nielsen, A. T. (2019). Production of zosteric acid and other sulfated phenolic biochemicals in microbial cell factories. Nature Communications, 10(1). https://doi.org/10.1038/s41467-019-12022-x‌https://doi.org/10.1038/s41467-019-12022-x‌