Results

Background

Phosphate is an essential nutrient required to support plant growth, however excess levels due to the overuse of phosphate fertilisers results in agricultural run-off which can lead to damaging eutrophication which can devastate aquatic life. As a result, monitoring phosphate levels in soil is critical not only for monitoring plant health, but also providing direct measurements of environmental phosphate levels to implement appropriate clean up strategies in eutrophic rivers. To address this, we sought to engineer a phosphate biosensor using B. subtilis. Given the ability of this model Gram-positive to form spores, it makes an ideal chassis to enable long term storage and transport of a phosphate biosensor to provide in-situ monitoring of phosphate levels.

The P_pstS promoter – our design for a phosphate biosensor

The promoter P_pstS, native to B. subtilis, is regulated by the PhoPR Two-Component System which drives the expression of the pstS-pstC-pstA-pstBA-pstBB operon required for the expression of high affinity phosphate binding proteins and transporters to import phosphate upon phosphate limitation. In phosphate replete conditions, PhoR is unable to phosphorylate PhoP due to inhibition of PhoR by intermediates of Wall Teichoic Acid biosynthesis. The resulting unphosphorylated PhoP can no longer activate transcription of the P_pstS promoter, and thus phosphate uptake is inhibited. In phosphate deplete conditions, PhoR phosphorylates its cognate response regulator PhoP (PhoP-P). Once phosphorylated, PhoP-P is able to bind and activate the P_pstS promoter to initiate transcription of phosphate importers and binding proteins [1–3]. A simplified schematic of this is shown in Fig. 1 which outlines the design of the proposed phosphate biosensor. Here, the promoter P_pstS (BBa_K4418004) drives expression of a luciferase reporter (luxABCDE) in the plasmid pBSGGlux (donated by Prof. Susanne Gebhard) according to phosphate availability. In this system, under high phosphate conditions, the promoter is repressed (due to the presence of the unphosphorylated form of PhoP), whilst in low phosphate conditions, the promoter is activated (due to phosphorylation of PhoP by PhoR).

Figure 1. Circuit schematic for the P_pstS phosphate biosensor. A) Under phosphate replete conditions the promoter P_pstS is repressed as PhoP remains in a dephosphorylated state and is unable to activate the promoter. B) In phosphate deplete conditions, PhoP is phosphorylated (PhoP-P) and is able to activate the promoter P_pstS to drive luxABCDE (luciferase) activity.

PCR amplification of the P_pstS promoter

To construct the phosphate biosensor, we first amplified the P_pstS promoter from B. subtilis gDNA which resulted in a 161 bp fragment (Fig. 2A). This was subsequently used in a Golden Gate ligation using plasmid pBSGGlux which was subsequently transformed into DH5α, with white colonies screened for successful ligation of P_pstS. A single colony was identified which was screened for the insertion of the P_pstS promoter using colony PCR. As shown in Fig. 2B, a 234 bp fragment corresponding to the insertion of P_pstS into pBSGGlux was identified.

Figure 2. Cloning of the P_pstS promoter to generate a P_pstS-luxABCDE transcriptional fusion. A) P_pstS was amplified from B. subtilis gDNA the resulting product (161 bp) visualised using agarose gel electrophoresis. “L” corresponds to ladder, “P_pstS” is the promoter, “-“ corresponds to the negative control B) Ligation of P_pstS into pBSANDlux. “L” corresponds to ladder, “-“ corresponds to the negative control, “P” corresponds to positive control, “1” corresponds to the single identified colony. 1% gels with 1X TAE were used with the DNA stained using SYBR Safe. The gel was run for 100V for 30 minutes.

Integration of P_pstS-pBSANDlux into the B. subtilis genome

Following the cloning of the promoter P_pstS into plasmid pBSGGlux, the resulting plasmid was purified from an O/N culture of DH5α and transformed into B. subtilis. The resulting transformants were patched onto an LB agar plate supplemented with chloramphenicol and screened for integration of the plasmid at the sacA locus via double cross-over. As shown in Fig. 3, all four screened transformants showed double cross-over at the sacA locus, revealing stable integration (up and down) into the genome. Up-integration and down-integration resulted in expected band sizes of 946 and 930 bp, respectively.

Figure 3. Verification of the integration of P_pstS-pBSANDlux at the sacA locus. B. subtilis transformants were screened for the integration of the P_pstS-luxABCDE reporter plasmid at up and down regions of the sacA locus. “L” corresponds to the DNA ladder, “-ve” corresponds to the negative control (water), “+ve” corresponds to the positive control, and screened colonies are indicates (1-4).

The PhoPR regulated P_pstS-luxABCDE biosensor displays the behaviour of a NOT gate.

Following the confirmation of stable integration of the plasmid P_pstS-pBSGGlux into the B. subtilis genome, we next sought to assess the functionality of the phosphate biosensor. Cells with the integrated circuit were grown in MCSE medium containing 1% casamino acids and in which the phosphate from the MOPS solution mixture was removed to generate phosphate deplete conditions. Once the cells reached an OD₆₀₀ of 0.1 (~0.4 in 1 cm cuvette), they were subsequently induced with various increasing phosphate concentrations. As shown in Fig. 4, the dose-response of this circuit is consistent with the regulation of the P_pstS promoter, where increasing the concentration of phosphate (indicative of phosphate replete conditions) begins to repress the activity of the promoter by preventing PhoP phosphorylation, and inhibiting subsequent activation of the P_pstS promoter. This results in a drop in RLU/OD₆₀₀ activity of 14-fold where promoter repression is saturated at 0.3 mM of phosphate. The result logic of this circuit, when the presence of phosphate represses promoter activity, is NOT logic.

Figure 4. Activity of the P_pstS-luxABCDE phosphate biosensor. Measurement of P_pstS promoter activity in pBSGGlux upon induction with various phosphate concentrations. Promoter activity of the strain is luminescence (RLU) normalised to OD₆₀₀ (RLU/OD₆₀₀) of cells grown in phosphate lacking MCSE medium supplemented with 1% casamino acids. Data are mean values and ± SD of two biological replicates.

Conclusion & future directions

In this section of our project, we sought to design and test the functionality of a phosphate biosensor based on the native P_pstS promoter in B. subtilis, regulated by the PhoPR two-component system. Our work presents successful cloning of the P_pstS promoter and demonstrates the P_pstS promoter is repressible in the presence of increasing phosphate concentrations, functioning as a NOT logic gate circuit. The promoter P_pstS (BBa_K4418004) will likely see application in a variety of other related projects where phosphate is the topic of interest and B. subtilis is the chassis organism of choice.

Future directions using promoter P_pstS in B. subtilis will aim to explore the effect repeated rounds of sporulation and germination on the behaviour of the circuit, the incorporation of this sensor into microfluidics devices to enable portable monitoring of phosphate, and to establish methodology to extract soil phosphate to ascertain as to whether this sensor would be able to provide a means of in situ monitoring of soil phosphate.

Background

As we have shown using out phosphate inducible promoter BBa_K4418004, increasing phosphate concentrations result in repression of the promoter P_pstS. For application in phosphate remediation where phosphate concentrations in solution are elevated, such as in eutrophic rivers or wastewater, this would mean the expression of any phosphate importers used to uptake phosphate would be inhibited as phosphate levels are in excess. As a result, the regulation of the P_pstS promoter obeys the logic of a NOT gate, where the presence of an input (high phosphate) causes an output of 0 (luciferase), and the absence of an input (low phosphate), causes the output to be 1.

To allow expression of phosphate importers to in the presence of high phosphate for environmental remediation, a NOT gate would be required to ensure the expression of phosphate uptake systems in the presence of high phosphate. A schematic of this is shown in Fig. 5A-B. As shown in Fig.5A, in the presence of high phosphate, the P_pstS promoter is repressed, such that the expression of a downstream repressor can no longer take place. As a result, the cognate promoter of the repressor (Prepressor) becomes active and can allow for the expression of phosphate importers in the presence of high amounts of phosphate. When the levels of phosphate deplete, the P_pstS promoter becomes active, allowing for the expression of the repressor which subsequent represses transcription of a downstream phosphate uptake system. This proposed design ensures phosphate uptake only occurs when phosphate levels in solution are elevated.

Figure 5. Design of a phosphate regulated NOT gate. A) Regulation of the phosphate regulated NOT gate in phosphate replete. High phosphate conditions allow for the repression of a repressor driven by the promoter P_pstS. With the repressor no longer made, the cognate promoter of this repressor can allow for transcription of phosphate uptake genes to import external phosphate. Note, the NOT gate section of the circuit is boxed in light blue. B) Regulation of the phosphate circuit in the phosphate deplete conditions. The presence of phosphate causes activation of transcription from P_pstS and subsequent expression of a repressor which is able to repress its’ cognate target promoter and expression of phosphate uptake genes.

Functionality of the heterologous NOT gate in B. subtilis

For the characterisation of the NOT gate, we initially wished to use the PxylA promoter (BBa_K1351039) to drive the expression of the heterologous repressor SarA (BBa_K4418005) of Staphylococcus aureus, the target promoter of which is the P_sprC (BBa_K4418006)[4]. The use of the PxylA promoter was favoured to first assess as to whether the circuit functions prior to testing the final design using our characterised P_pstS promoter. A full schematic of the circuit is shown in Fig. 6. In this circuit design, the expression of the regulator SarA is induced by the promoter PxylA in the presence of xylose, and subsequently translated by the consensus B. subtilis RBS (BBa_K090505). To prevent read-through from PxylA, a strong transcriptional terminator BBa_K4418007 is used. The resulting circuit should show a reduction in luxABCDE in the presence of xylose through the composite part BBa_K4418008.

Figure 6. Schematic of the PxylA inducible NOT gate ( BBa_K4418008). In the proposed xylose inducible SarA based NOT, the presence of xylose allows for the expression of the repressor SarA from PxylA which leads to subsequent repression of the P_sprC promoter and luxABCDE activity. Boxed in blue is composite part comprising the xylose-inducible NOT gate, part BBa_K4418008.

Following xylose induction, SarA is able to repress its’ cognate promoter P_sprC which drives the expression of a luciferase reporter (luxABCDE). As a result, luciferase output shown decrease as a function of increasing xylose concentrations. As shown in Fig. 7, consistent with our theorised design, the repression of luciferase activity occurs in a xylose dependent manner with basal activity of 293025 RLU/OD₆₀₀ units with output reduced to 40703 RLU/OD₆₀₀ (7.2 -fold repression) in the presence of 0.2 % xylose.

Figure 7. Activity of the xylose inducible NOT gate. Promoter activity of P_sprC in response to xylose inducible regulation of the repressor SarA, expressed as luminescence in RLU/OD₆₀₀. Data show mean values and ± SD of a single biological replicate carried out using duplicate technical measurements.

Conclusion & future directions

Our data confirms that the heterologous SarA-P_sprC regulated circuit generates a functional NOT gate heterologous which can repress the luciferase output reporter. Due to time limitations, we were unable to test the circuit designed using the phosphate regulated promoter P_pstS and to see as to whether increasing phosphate concentration would correspond to an increase in luciferase output, and as to whether decreasing phosphate concentration would cause a reduction in luciferase output. Additionally, we were unable to (due to time limitation) test our design with a phosphate uptake system, or in application to phosphate bioremediation using phosphate importers. Nevertheless, we demonstrate the functionality of this circuit which provides useful parts for the implementation of NOT gates in B. subtilis. Future directions in this work will require RBS fine-tuning strategies to increase P_sprC by SarA to provide greater repression of the output module used with this NOT gate circuit.

Background

Malate is one of the organic acid exudates which plants release upon encountering phosphate deficiency which allows them to increase local soil pH and readily solubilise more phosphate within soil [5]. In our long-term goal of genetically modifying B. subtilis to release phosphate according to plant phosphate deprivation signals, such as malate, we first sought to characterise a promoter responsive to malate.

Design of a malate responsive biosensor

To design a malate sensing circuit, we utilised the P_maeN promoter ( BBa_K4418000) native to B. subtilis which is regulated by the MalKR Two-Component System. In this system, the sensory kinase MalK phosphorylates its cognate response regulator MalR in response to malate in order to activate transcription of the P_maeN promoter, allowing for expression of enzymes to enable to usage of malate as a carbon source [6]. A simplified schematic of this is shown in Fig. 8. To build this circuit, we cloned the P_maeN promoter into the luciferase reporter plasmid pBSGGlux.

Figure 8. Schematic of the P_maeN – luxABCDE biosensor in B. subtilis. The malate responsive promoter P_maeN is regulated by the MalKR Two-Component System. In this circuit design, malate induction causes activation of the sensor kinase MalK, leading to subsequent phosphorylation of the response regulator MalR and activation of the P_maeN promoter, resulting in measurable luxABCDE expression.

Activity of the BBa_K4418000  biosensor in B. subtilis

To test the activity of the sensor, we grew strains in MCSE medium and induced with various concentrations of malate to the point of promoter saturation. The resulting dose-response is shown in Fig. 9. As shown, RLU/OD₆₀₀ activity increases as a function of malate concentration covering a range within two orders of magnitude, with promoter activity saturation at 0.5 mM of malate. The resulting data demonstrate the successful construction of a malate sensor and validates its use in the design of a malate dependent phosphate release circuit, as we have done using the composite part BBa_K4418003.

Figure 9. Dose response curve of the P_maeN-luxABCDE reporter. Cells harbouring the integrated P_maeN-luxABCDE circuit were grown to mid-exponential phase (OD₆₀₀ = 0.1) in MCSE medium and challenged with various concentrations of malate shown. Promoter activity following malate induction are expressed as luminescence in RLU/OD_600¬ of 35-45 minutes post induction. Values presented show mean and ± standard deviation of three biological replicates.

Conclusions & Future Directions

In this section of our project, we sought to characterise the part BBa_K4418000, a malate responsive promoter native to B. subtilis. We demonstrate successful engineering of this promoter into B. subtilis as a luxABCDE reporter fusion and demonstrate the operational range of this sensor. We subsequently used this part to generate the composite part BBa_K4418003 which we utilised to trigger the degradation of Wall Teichoic Acid (WTA) into shorter phosphate rich subunits which can be used as a phosphate source to support plant growth. Future directions will aim to improve the dynamic range and sensitivity of this promoter in order to increase transcriptional output and provide sensitive detection of malate from plant exudates grown hydroponically.

Background 

Phosphate is an essential nutrient required to support plant growth, yet overuse of commercial phosphate fertiliser not only depletes available stores, but also contributes to agricultural run-off which can lead to damaging eutrophication of rivers. To resolve this issue, we sought to engineer B. subtilis as a smart biofertiliser capable of releasing phosphate to plants in a manner accordingly to the plants physiological needs. To do this, we initially characterised the part BBa_K4418000, a malate inducible promoter. Malate is a plant exudate released during phosphate limitation which is used to solubilise soil phosphate. To generate a phosphate release circuit, we took advantage enzymes involved in the phosphate starvation response in B. subtilis. Upon entering phosphate limited environments, B. subtilis induces the expression of PhoD and GlpQ – two secreted phosphodiesterase enzymes which catalyse the degradation of Wall Teichoic Acid (WTA), a phosphate rich polymer (polyglycerol-phosphate) attached to peptidoglycan which serves as a phosphate store [7]. We speculated, that by coupling the induction of the P_maeN ( BBa_K4418000) to the release of both enzymes GlpQ ( BBa_K4418001) and PhoD  ( BBa_K4418002), we could trigger malate inducible phosphate release, enabling plants to obtain phosphate from depolymerised WTA to subsequently support growth.  

Design of a phosphate release circuit  

To generate a Bacillus subtilis strain that would function as a smart phosphate releasing biofertiliser, we sought to assemble a composite part comprising the promoter P_maeN BBa_K4418000, and both WTA degrading enzymes GlpQ ( BBa_K4418001) and PhoD  ( BBa_K4418002) - both of which are translated by the consensus B. subtilis RBS sequence ( BBa_K090505). A schematic of the resulting composite part BBa_K4418003 is shown in Fig. 10. In this circuit, malate (released in plant exudates during phosphate limitation) triggers the expression of both GlpQ and PhoD to allow for subsequent WTA degradation and release of phosphate into the culture medium. 

Figure 10. Circuit schematic of the phosphate release circuit BBa_K4418003. Induction with malate allows transcriptional activation from the P_maeN promoter, allowing for expression of the genes glpQ and phoD which encode Wall Teichoic Acid (WTA) degrading enzymes. Expression of both enzymes allows for phosphate release from WTA degradation.

Amplification of GlpQ flanking regions   

Since GlpQ and PhoD are both expressed under phosphate limiting conditions natively in B. subtilis, we sought to knock out GlpQ (as well as PhoD) to ensure any phosphate release was resulting from our engineered phosphate release circuit, not the native mechanism [7]. This required a genetic deletion of both enzymes, for which we utilised the CRISPR-Cas9 plasmid pJOE8999. We first had to amplify the flanking regions of GlpQ in B. subtilis to clone into the plasmid. These flanking regions provide a template for the bacteria to repair the double stranded break induced by the Cas9 endonuclease and remove the native copy of GlpQ. The gel showing the amplified flanking regions is shown in Fig. 11. 

Figure 11. Amplification of 5’ and 3’ GlpQ flanking regions. A 1% agarose gel stained with SYBR safe ran at 100V for 30 minutes. + indicates reactions with the addition of B. subtilis W168 DNA, - indicates control reactions with the addition of PCR grade H2O instead.

Successful amplification of both 5’ and 3’ flanking regions of GlpQ was shown by the presence of bands at 1317bp and 1001bp respectively. We then digested the flanking regions and pJOE8999 with Sfil and then ligated these flanking regions into the plasmid. From here the plasmid with integrated flanking regions was transformed into DH5a E. coli and then colonies were screened via a colony PCR to check if the flanking regions successfully integrated into the plasmid (Fig. 12). Colonies with the plasmid were taken forward, grown overnight and miniprepped using a commercial kit.  

Figure 12. Colony PCR screening of pJOE8999 for insertion of glpQ flanking regions. E. coli colonies were screened for the insertion of 5’ and 3’ flanking regions for glpQ. A 1% agarose gel using 1X TAE was used with SYBR safe DNA stain. Gels were run at 100V for 30 minutes. “L” represents ladder, “-“ represents the negative control, “+” represents positive control (pJOE8999), with the colonies screened numbered accordingly.   

Insertion of gRNA into CRISPR-Cas9 plasmid    

Insertion of the gRNA to direct the Cas9 machinery was achieved by ligation of the gRNA into the plasmid containing the flanking regions through a Goldengate reaction. This plasmid was then transformed again into DH5a E. coli which were streaked onto kanamycin and X-gal selection plates (See Fig. 13). Colonies showing a positive result (white) indicate that the gRNA had been implemented and disrupted the LacZ cassette. These were grown overnight and then miniprepped. 

Figure 13. Blue-white screening in E. coli DH5α to identify the insertion of the glpQ targeting gRNA.  A LB agar plate with 10 µg/mL kanamycin and 50 µg/mL -gal showing the blue and white screening for E. coli DH5α.  If the gRNA has been integrated, the LacZ cassette would be disrupted, leading to white colonies. If the gRNA is not integrated, LacZ produces β-galactosidase which hydrolyses X-gal to produce an insoluble blue pigment, indicating a negative result. 

Deletion of GlpQ in B. subtilis     

Following successful construction of the CRISPR plasmid in E. coli, the purified plasmid was subsequently transformed in B. subtilis. Once colonies were obtained, they were streaked onto LB agar without antibiotics and underwent temperature shift steps in order to cure the cells of the CRISPR plasmid, and to render them sensitive to kanamycin. After the temperature shift steps, colonies were then screened to identify those that had been cured of the plasmid. These colonies were then checked to see if glpQ had been successfully knocked out (Fig. 14). The reduction in band size in relation to the wild-type control indicates that glpQ has been successfully deleted, with all four screened colonies showing successful deletion. The final B. subtilis colony with the glpQ deletion was taken forward and grown, to then knock out phoD. 

Figure 14. Colony PCR screening of B. subtilis colonies for the deletion of GlpQ using CRISPR-Cas9. A 1% agarose gel using 1X TAE buffer was stained with SYBR safe and ran at 100V for 30 minutes. L represents NEB 1kb plus DNA ladder, -ve the negative control with the addition of H2O, WT is wildtype B. subtilis w168 DNA, these were ran alongside the 4 tested colonies.   

Deletion of PhoD from B. subtilis      

To delete the gene phoD in the genetic background in which we had removed the gene glpQ and cured the cells of the CRISPR-Cas9 plasmid which conferred kanamycin resistance, we used a B. subtilis ΔphoD strain from the Bacillus subtilis BKK collection (kindly provided by Prof. Susanne Gebhard). Isolated genomic DNA from this B. subtilis ΔphoD strain was used to transform the prior constructed B. subtilis ΔglpQ strain in order to remove the gene phoD, generating a double mutant (B. subtilis ΔglpQ/ΔphoD), as shown in Fig. 15A-B. 

Figure 15. PhoD deletion in a B. subtilis ΔglpQ genetic background. Isolated genomic DNA (gDNA) from a B. subtilis ΔphoD strain was used to transform a B. subtilis ΔglpQ strain. A) Negtaive control without the addition of B. subtilis ΔphoD gDNA. B) B. subtilis ΔglpQ strain transformed with gDNA from B. subtilis ΔphoD. Transformants were plated onto LB agar supplemented with 10 µg/mL kanamycin.   

Building the phosphate release circuit       

To build the phosphate release circuit, we utilised the plasmid pBSGG1C (donated by Prof. Susanne Gebhard) into which we cloned amplified fragments for the malate promoter P_maeN (BBa_K4418000), and the enzymes GlpQ (BBa_K4418001) and PhoD (BBa_K4418002) – the later two which are translated by the B. subtilis consensus RBS sequence (BBa_K090505) . The resulting gel following amplification of these fragments is shown in Fig. 16A. These parts were assembled into pBSGG1C using Golden Gate, transformed into DH5α and then the resulting colonies screen to identify colonies with the correctly assembly circuit, as shown in Fig. 16B. Out of 16 screened colonies, 10 showed a band size which corresponding to a fully assembled phosphate release circuit (2828 bp). Colonies with the correct size band were grown overnight with the plasmids mini-prepped for transformation in B. subtilis.  

Figure 16. Construction of the phosphate release circuit. A) Amplification of parts in the release circuit. Lanes 1-4 correspond to NEB 1kb plus ladder, P_maeN, GlpQ and PhoD, respectively. B) Screening of DH5α colonies for complete construction of the phosphate release circuit. Lanes 1; NEB 1kb plus ladder, Lane 2; negative control, lane 3; positive control (pBSGG1C), lanes 4-19 correspond to the screening of colonies 1-16. The fully assembled circuit has a band size of 2828 bp. A 1% agarose gel (1X TAE) was used with SYBR safe DNA binding dye.      

Integration of the release circuit into B. subtilis    

The resulting assembled phosphate release circuit was transformed into the B. subtilis ΔphoD/ΔglpQ genetic background. As the plasmid pBSGG1C integrates at the amyE locus in B. subtilis, transformants were patched onto starch agar plates supplemented with chloramphenicol (in parallel with transformants being patched onto LB agar plates supplemented with chloramphenicol), to which 2% iodine solution was added. Integration of the plasmid at this locus disrupts α-amylase production and prevents starch degradation, as shown in Fig. 17. 

Figure 17. Screening colonies for integration of the phosphate release circuit in the B. subtilis ΔphoD/ΔglpQ. Integration of the phosphate release circuit in the plasmid pBSGG1C disrupts α-amylase production. If the plasmid has not integrated at the amyE locus via double cross-over, colonies appear with a bright halo following the addition of 2% iodine.  If a plasmid has integrated by double cross-over at this locus, the inability of cells to produce α-amylase prevents starch degradation, thus no bright halo appears. Colonies 1, 3, 5, 9, 12, 13 and 16 show correct integration.      

Testing phosphate release in the presence of malate   

To test for the release of phosphate in response to malate induction, B. subtilis ΔphoD/ΔglpQ with the integrated release circuit were grown in LB medium to OD₆₀₀ = 0.8, spun-down, the supernatant removed and the pellet washed with 0.85% saline solution. This was repeated three times to remove residual phosphate after which the cells were resuspended in 8 mL of saline with or without 1 mM malate supplementation and grown for 1 hour at 37 °C. After 1 hour of growth, cells were pelleted, the supernatant removed and subsequently tested with a commercial phosphate detection kit (SpectroQuant®). The resulting release is shown in Fig. 18. Whilst some variation is observed, the general trend observed shows consistently higher phosphate release in cells induced with malate, relative to cells in which do not have an integrated release circuit and only a double ΔphoD/ΔglpQ deletion. Likewise, cells induced with malate also generally show higher release compared to those in the absence of malate. The reason why phosphate signal is still detectable in the supernatants from the B. subtilis ΔphoD/ΔglpQ genetic background without an integrated circuit remains unclear. However, we speculate this may be due to other metabolites released during cell growth which likely have passed through the filtration step following growth in saline and cross-reacted with reagents in the assay. Note, no cross-reactivity with the phosphate detection kit was observed using 0.85% saline alone.

Figure 18. Testing phosphate release in the presence of malate. B. subtilis cells, including the B. subtilis ΔphoD/ΔglpQ strain (no circuit) and cells with the integrated release circuit were grown (1 hr, 37 °C) without (circuit -ve) and with (circuit +ve) malate induction (1 mM) following resuspension of cells in 0.85% saline. Saline was included as a control. The supernatants from these cells were filter sterilised and tested using a commercial kit (SpectroQuant® Phosphate Test Cell) for phosphate release. Data are presented are the values from three independent biological replicates.  

Conclusions & Future directions  

In this work, we sought to design, construct and test a phosphate release circuit which can allow for malate inducible phosphate release following the degradation of B. subtilis WTA. Our results show successful cloning of the circuit and functionality in B. subtilis. The resulting supernatants from malate induced cells were collected and used as part of our proof-of-concept experiment to assess as to whether the bacterial derived phosphate, release following malate induction, can support plant growth. 

Future work should explore the use of fine-tuning RBS strength to improve WTA depolymerisation and increase phosphate release, as well as testing lower malate concentrations to assess as to whether the circuit will function at environmentally relevant concentrations of malate release in plant exudates.