iGEM 2022 — Team uniofbath

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{% block title %}Notebook{% endblock %}
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## What should this page contain?

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* Chronological notes of what your team is doing.
* Brief descriptions of daily important events.
* Pictures of your progress.
* Mention who participated in what task.

## Inspirations

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* [2018 Munich](http://2018.igem.org/Team%3AMunich/Notebook)
* [2019 Georgia State](https://2019.igem.org/Team%3AGeorgia_State/Notebook)
* [2019 Newcastle](https://2019.igem.org/Team%3ANewcastle/Notebook)
* [2020 IISER Pune India](https://2020.igem.org/Team%3AIISER-Pune-India/Notebook)
* [2020 Lund](https://2020.igem.org/Team%3ALund/Notebook)
* [2020 NOVA LxPortugal](https://2020.igem.org/Team%3ANOVA_LxPortugal/Notebook)
* [2020 RDFZ China](https://2020.igem.org/Team%3ARDFZ-China/NoteBook)

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[Phosphate Uptake](#slide1-chapter1)

[Functionality of the heterologous NOT gate in Bacillus subtilis](#slide1-chapter2)

[Phosphate Release](#slide1-chapter3)

[Testing the inducible promoter](#slide1-chapter4)

[Testing the Release Circuit](#slide1-chapter5)

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## Phosphate Uptake

In native Bacillus subtilis, phosphate inducible promoter PpstS [BBa\_K4418004](http://parts.igem.org/Part%3ABBa_K4418004), is repressed in increasing phosphate concentrations. 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 PpstS 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, we **DESIGNED** a NOT gate to ensure the expression of phosphate uptake systems in the presence of high phosphate. A schematic of this is shown in Fig. 1A-B. As shown in Fig.1A, in the presence of high phosphate, the PpstS 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 PpstS 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.

![](https://static.igem.wiki/teams/4418/wiki/images/engineeringfig1.png)

**Figure 1.**  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 PpstS. 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 PpstS 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 Bacillus subtilis

For the characterisation of the NOT gate, we initially wished to use the PxylA promoter ([BBa\_K1351039](http://parts.igem.org/Part%3ABBa_K1351039)) to drive the expression of the heterologous repressor SarA ([BBa\_K4418005](http://parts.igem.org/Part%3ABBa_K4418005)) of Staphylococcus aureus, the target promoter of which is the PsprC ([BBa\_K4418006](http://parts.igem.org/Part%3ABBa_K4418006))[[1]](#ref1). 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 PpstS promoter. A full schematic of the circuit is shown in Fig. 2. 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](http://parts.igem.org/Part%3ABBa_K090505)). To prevent read-through from PxylA, a strong transcriptional terminator [BBa\_K4418007](http://parts.igem.org/Part%3ABBa_K4418007) is used. The resulting circuit should show a reduction in luxABCDE in the presence of xylose through the composite part [BBa\_K4418008](http://parts.igem.org/Part%3ABBa_K4418008).

![](https://static.igem.wiki/teams/4418/wiki/images/engineeringfig2.png)

**Figure. 2**  Schematic of the PxylA inducible NOT gate ( [BBa\_K4418008](http://parts.igem.org/Part%3ABBa_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 PsprC promoter and luxABCDE activity. Boxed in blue is composite part comprising the xylose-inducible NOT gate, part [BBa\_K4418008](http://parts.igem.org/Part%3ABBa_K4418008).

We **BUILT** the circuit shown in figure 2 using Golden Gate cloning, assembling the NOT gate parts together and then assembling it into the plasmid pBSANDlux which contained our luciferase reporter (luxABCDE). Subsequently we transformed the assembled plasmid into Escherichia coli DH5α to test for successful integration by visible white colonies, indicating replacement of the RFP cassette in pBSANDlux by our plasmid. We then isolated transformed colonies, extracted our plasmid and transformed it into Bacillus subtilis, our chassis, for validation.

We **TESTED** our circuit by exposing our transformed Bacillus subtilis to increased xylose concentrations, measuring the relative luminescence units (RLU) with a plate reader, normalised by the optical density (OD 600) of the cultures. This measurement inform us of the relative promoter activity for a given concentration of cells.

Following xylose induction, we **LEARNT** that SarA is able to repress its cognate promoter PsprC 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. 3, 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.

![](https://static.igem.wiki/teams/4418/wiki/images/engineeringfig3.png)

**Figure 3.**  Activity of the xylose inducible NOT gate. Promoter activity of PsprC 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.

## Phosphate Release

Bacillus subtilis contains the gene encoding for thephosphodiesterase glpQ and the phosphatase encoding gene phoD ( [BBa\_K4418001](http://parts.igem.org/Part%3ABBa_K4418001)) both of which are activated under phosphate starvation are expressed and active in the degradation of teichoic acid in the cell wall. This leads to the release of phosphate as this is where it is natively stored in Bacillus subtilis. The phosphate released from this mechanism is then utilised by the bacteria for its metabolic processes.

For being used in a phosphate release circuit that induced by a plant signal released under phosphate stress, we **DESIGNED** a circuit in which an inducible promoter PmaeN [BBa\_K4418000](http://parts.igem.org/Part%3ABBa_K4418000) was inserted upstream of both genes. As shown in Fig. 4 PmaeN is induced by malate, an exudate released by plant roots when lacking in phosphate. As a result, phoD and glpQ will mediate the release of phosphate induced by malate i.e. when plants find themselves in need of phosphate. In this manner, plants will have a controlled supply of phosphate, one that is activated upon demand and that will not lead to excess phosphate levels in soil.

![](https://static.igem.wiki/teams/4418/wiki/images/engineeringfig4.png)

**Figure. 4**  Circuit schematic of the phosphate release circuit [BBa\_K4418003](http://parts.igem.org/Part%3ABBa_K4418003). Induction with malate allows transcriptional activation from the PmaeN 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.

## Testing the inducible promoter

The characterization the PmaeN promoter was carried out through the insertion of the promoter into the luciferase reporter plasmid pBSGGlux. A schematic of the circuit ( [BBa\_K4418003](http://parts.igem.org/Part%3ABBa_K4418003)) is show in Fig. 5 in which malate induction activates the MalK protein kinase that phosphorylates the PmaeN response regulator MalR. This would lead to luxABCDE expression which would then be measured through the emitted bioluminescence. Therefore, the circuit would emit a measurable signal upon malate induction allowing us to verify PmaeN functionality.

![](https://static.igem.wiki/teams/4418/wiki/images/engineeringfig5.png)

**Figure. 5**  Schematic of the PmaeN – luxABCDE biosensor in B. subtilis. The malate responsive promoter PmaeN 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 PmaeN promoter, resulting in measurable luxABCDE expression.

We **BUILT** the circuit shown in Fig. 5 using Golden Gate cloning, assembling PmaeN into the plasmid pBSGGlux containing luxABCDE. Subsequently we transformed the assembled plasmid into Escherichia coli DH5α to test for successful integration by visible white colonies, indicating replacement of the RFP cassette in pBSANDlux by our plasmid. We then isolated transformed colonies, extracted our plasmid and transformed it into B. subtilis, our chassis, for validation.

We **TESTED** our circuit by exposing our transformed B. subtilis to increased malate concentrations, measuring the relative luminescence units (RLU) with a plate reader, normalised by the optical density (OD 600) of the cultures. This measurement informed us of the relative promoter activity for a given concentration of cells.

We then **LEARNT** that the construct the luminescence emission occurred in a dose dependant manner upon malate induction (Fig. 6). This allowed us to verify the suitable working range of this promoter and, furthermore, to utilize it in our designed release circuit

![](https://static.igem.wiki/teams/4418/wiki/images/engineeringfig6.png)

**Figure. 6**  Dose response curve of the PmaeN-luxABCDE reporter. Cells harbouring the integrated PmaeN-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/OD600 of 35-45 minutes post induction. Note, due to the log-scale on the x-axis, values at 0.001 mM represent basal (uninduced) activity. Values presented show mean and ± standard deviation of triplicate measurements performed on three different days (n = 9).

## Testing the Release Circuit

Upon characterizing the inducible promoter PmaeN. We then proceeded to BUILD the release circuit containing glpQ and phoD (Fig. 4). Once again, we assembled the parts through Golden Gate Cloning. We assembled the pBSGGIC plasmid with glpQ, phoD and PmaeN. Before testing the circuit, we built two CRISPR plasmids for the deletion of the native glpQ and phoD genes, creating the ΔphoD/ΔglpQ strain, . In this way, our strain should not release phosphate under low phosphate conditions as the system naturally would do. This means that we could then test the strain under low phosphate conditions without phosphate being released, which would interfere with the analysis of our results. After deletion of both genes, the circuit was transformed into B. subtilis.

We **TESTED** the strain by growing the cells in LB medium to OD600 = 0.8. Cells were spun down and washed to remove residual phosphate. Cells were then resuspended in malate supplemented solution for 1 hour, after which the supernatant was collected, and phosphate levels were measured using the commercial phosphate measuring kit (SpectroQuant®) (Fig. 7).

![](https://static.igem.wiki/teams/4418/wiki/images/engineeringfig7.png)

**Figure. 7**  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.

We **LEARNT** that B. subtilis ΔphoD/ΔglpQ circuit containing strain had a significantly higher release of phosphate upon malate induction. This allowed us to conclude that the circuit worked in the intended manner. This shows that we successfully went through the DBTL cycle, showing the success of our parts step by step leading to the understanding of how our final circuits worked.

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[Plasmid Prep](#slide1-chapter1)

[Golden Gate Assembly](#slide1-chapter2)

[Transformation of the assembled plasmid into E. coli DH5α](#slide1-chapter3)

[Making a Patch Plate/a>](#slide1-chapter4)

[Colony PCR](#slide1-chapter5)

[Transformation of Bacillus subtilis](#slide1-chapter6)

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The following chapters detail the course of a single set of cloning and transformation experiments that made up the core of our project.

## Plasmid Prep

1. Find the cryovial with the plasmid (in our case pBSANDlux, light pink in colour), KEEP THIS ON DRY ICE AT ALL TIMES. Using a sterile loop, scrape some of the frozen cells from the cryovial so that the loop becomes inoculated and streak this onto an LB agar plate supplemented with an appropriate antibiotic for selection (in our case, ampicillin) to obtain single colonies. Do this in a flow hood and keep the tube on dry ice.
2. Once streaked, incubate the plate at 37oC O/N in a static incubator
3. Set up a 2 mL O/N culture of LB medium supplemented with Ampicillin, inoculate this with a colony from the streak plate with the plasmid, allow this to grow O/N at 37oC in a shaking incubator at 200 rpm. – Allow it to grow for 12-18 hours ideally.
4. Follow the plasmid prep kit instructions on how to prep the plasmid
5. Quantify the amount of plasmid using a nanodrop (in µg/µL) WRITE THE CONCENTRATION AND DATE ON THE SIZE OF THE EPPENDORF AS WELL AS THE DATE.

## Golden Gate assembly

###### Reagents

* 10X T4 Ligase Buffer
* 10X CutSmart Buffer
* 400,000 U/ml T4 DNA ligase
* 20,000 U/ml BsaI-HFvs2
* X ng of plasmid
* X ng of insert (this might be an amplified PCR fragment or a purchased gBlock

1. Use the NEB ligation calculator to determine the mass of plasmid you need. NEB recommends 27 fmol of plasmid to ensure optimal ligation – the mass in ng calculated form the fmol concentration will depend on the size of the plasmid. Using the [online calculator](https://nebiocalculator.neb.com/#!/dsdnaamt), and click on the tab mass -> moles, then click moles to mass. plasmid pBSANDlux (which is 10927 bp) at 27 fmol gives a mass of 182.3 ng (this is the final concentration required).
2. To get to a final concentration 182.3 ng. Divide this value by the concentration of pBSANDlux determined by the nanodrop. E.g if you have a plasmid concentration of 100 ng/uL based on the nanodrop, then you need to add a volume of 1.96 µL of purified pBSANDlux to the GoldenGate mix below.
3. For your insert, which is the NOT gate gBlock, use the [NEB ligation calculator](https://nebiocalculator.neb.com/#!/ligation). Into this, entire the size of the plasmid (10927 bp) the size of the insert NOT gate gBlock (814 bp) and then the calculated mass required for this plasmid (182.3 ng). This will calculate the concentration of the NOT gate gBlock that needs to be added to the GoldenGate mix. When I did this, the calculator gave me a concentration of 13.58 ng of the NOT gate gBlock (for a 1:1 ratio of insert to plasmid). Note: The gBlocks have been resuspended to a concentration of 10 ng/uL, so if you need a final concentration of 13.58 ng, you need 1.35 uL of the NOT gate gBlock in your Golden Gate mix.
4. For the GoldenGate mix, add reagents to the following amounts as required. MAKE SURE YOU KEEP ALL THE REAGENTS ON ICE. See the table below, and make up the mix in a PCR microfuge tube.

| Reagent | Volume | Final Amount |
| --- | --- | --- |
| Insert(s) | 1.35µL | 13.58ng |
| Plasmid | X µL | 196.1 ng |
| 10x T4 Ligase Buffer | 1 µL | 1x |
| 10x CutSmart Buffer | 1 µL | 1x |
| 400,000 U/mL T4 DNA ligase | 0.25 µL | 10,000 U/mL |
| 20,000 U/mL BsaI-HFvs2 | 0.5 µL | 1000 U/mL |
| Water | X µL | (Make up to 10 µL) |

Using the example concentrations on the previous page, the volume of plasmid would be 1.96 uL and the mass in ng would be 196.1 ng. For the fragment, the volume of purified PmaeN would be 2.85 uL and the mass would be 2.85 ng.

###### Golden Gate Cycles

| Temperature (oC) | Time (mins) |  |
| --- | --- | --- |
| 37 | 10 | x 10 Cycles |
| 16 | 10 |
| 16 | 30 |
| 50 | 5 |
| 80 | 10 |
| 4 | Hold |

After the thermocycler step, the resulting mix (10 µL) can be transformed directly into competent Escherichia coli DH5α cells using standard heat-shock and plated onto the appropriate selective medium. – If you don’t want to do this the same day as the golden gate, then keep the mixture in the -20oC freezer.

### Transformation of the assembled plasmid into E. coli DH5α

Once you have assembled the plasmid pBSANDlux with your NOT gate gBlock, you need to transform it into E. coli so the plasmid with your circuit can be replicated and lots of copies made so that we can extract out the plasmid, and put it into B. subtilis to see if it works.

1. Thaw a tube of DH5-alpha Competent E. coli cells on ice for 10 minutes.
2. Add 10 µl of the Golden Gate mix directly to the cells. Carefully flick the tube 4-5 times to mix cells and DNA. Do not vortex.
3. Place the mixture on ice for 15 minutes. Do not mix.
4. Heat shock at exactly 42°C for exactly 90 seconds. Do not mix.
5. Place on ice for 5 minutes. Do not mix.
6. Pipette 950 µl of room temperature SOC medium into the mixture.
7. Place at 37°C for 60 minutes. Shake vigorously (250 rpm) or rotate.
8. Warm selection plates to 37°C. – These are LB agar plates supplemented with ampicillin (100 µg/mL)
9. Mix the cells thoroughly by flicking the tube and inverting, then perform three10-fold serial dilutions in SOC medium
10. Spread 100 µl of each dilution onto a selection plate and incubate overnight at 37°C. Alternatively, incubate at 30°C for 24-36 hours or 25°C for 48 hours.

Once you’ve seen that you have colonies, the colonies which should have the insert will be white, anything that is red means there was plasmid that wasn’t cut by the enzyme and the RFP cassette wasn’t removed.

### Making a Patch Plate

Once you have inspected your transformation plate, you need to make a patch plate. A patch plate is just an LB agar plate with antibiotic (for the plasmid screening in E. coli, this needs to be an LB plate with ampicillin) that you use to pick colonies from your transformation plate and streak them onto a new fresh plate so the cells can grow to a bigger cell density.

1. To do this, pick a white colony (using a pipette tip) and carefully streak a small line onto the fresh LB agar plate with ampicillin which will be your patch plate. I STRONGLY recommend that on your patch plate, you draw a grid of about 10 squares which are carefully labelled numerically. Into each grid you will make a small streak of a single colony from your transformation plate. I.e pick a colony and streak it into square one, pick another colony and streak it into square two, etc.
2. Incubate the plate for a 3-4 hours at 37’C and check to see if you can see any cells beginning to grow. Now, you can do the colony PCR using the GoTaq Green MasterMix. See the reaction mixture below – If not, incubate overnight if you have a busy day.

The patch plate will be used to do a colony PCR.

## Colony PCR

Colony PCR is used to screen bacterial colonies to see if have a plasmid integrated, have a plasmid which successfully had a gBlock or PCR fragment ligated into it, if a genetic deletion has been successful, amongst many other applications. (you’re all scientists, so you know all the applications of screening cells)

| Reagent | Volume | Final Concentration |
| --- | --- | --- |
| GoTaq(R) Green Master Mix, 2x | 12.5 µL | 1x |
| Upstream Primer, 10µM | 0.25-2.5 µL | 0.1-1.0 µM |
| Downstream Primer, 10µM | 0.25-2.5 µL | 0.1-1.0 µM |
| DNA Template | 1-5 µL | <250ng |
| Nuclease-Free Water made up to | 25 µL | NA |

For plasmid pBSANDlux, the primers you need to add are IGB03/IGB04 – these are primers which anneal either side of the RFP sites in pBSANDlux so will show you if an insert has been placed in between the BsaI sites. (This will test if your Golden Gate was successful). This is better than using insert specific primers because if you assemble multiple inserts at the same, you can check for the incorporation of multiple inserts using these standard primers. – The annealing temperature for these primers is 60oC

###### Setting up the Colony PCR

1. The DNA template here is the actual bacterial colony. First set up your PCR tubes (don’t forget a positive control (pBSANDlux plasmid, 10 ng) and a negative control (no plasmid), and label then with the numbers from the grid (i.e tube 1 = colony 1 from grid 1 of the patch plate, tube 2 = colony 2 from grid 2 of the patch plate). I strongly recommend you set up a mastermix for this. Multiple the volumes above by the number of tubes you will have, and add everything into a single 1.5 mL Eppendorf and then aliquot 25 uL into each PCR tube. DO NOT ADD THE pBSANDlux PLASMID INTO THE MasterMix. Make the mastermix first, aliquot 25 uL into the PCR tubes, and then to your tube labelled negative control, you add nothing else, but to your positive control, you add 1 uL of 10 ng of pBSANDlux plasmid. Dilute the stock of plasmid accordingly to achieve this.
2. To add the bacterial colony to the other tubes, use a pipette tip (attached to a pipette) and just slightly touch the colony of the patch plate so that the pipette tip gets some bacterial cells onto the tip. Then, put this tip that now has the cells from, for example colony 1, into the PCR tube and pipette up and down and move the tip of the pipette which touched the cells around the inside walls of the PCR tube in the colony PCR mix. Do this for all the tubes. Be careful not to pick a large density of cells as it means you end up with a lot of debris at the bottom of the gel you will use to visualise if you have your gBlock inserted.

###### PCR Conditions

The annealing temperature for the Fwd and Rev primers the check for the insertion of genes into the Golden Gate site is 60oC.

| Temperature (oC) | Time |  |
| --- | --- | --- |
| 95 | 10 mins |
| 95 | 30 secs | x30 Cycles |
| 60 | 30 secs |
| 72 | 90 secs |
| 72 | 1 min |

1. Whilst the PCR is running, make a 1% agarose gel using 1X TAE buffer (50 mL gel) and be sure the gel has enough wells to accommodate the number of samples you want to run including your ladder, negative control and positive control.
2. Once the PCR is done, load the PCR mixtures directly into the wells of your gel and run for 20 mins at 140 V. The GoTaq has green dye in it so you don’t have to add loading dye separately.
3. The band size for the addition of the xylose inducible NOT gate is 883 nt.
4. Pick the colony which has the correct band size, and make an O/N culture of the one with the correct band size
5. Plasmid prep this plasmid and send it for sequencing using the eurofins kit.
6. Check the sequencing result against the reference plasmid (in our case, the xylose NOT gate-luciferase reporter).

### Transformation of Bacillus subtilis

1. Using the enzyme ScaI, cut 1-2 µg of your transformant (in our case, the pBSANDlux-NOT gate) for 1 hour as per the NEB enzyme digestion protocol (use a 50 uL reaction) – look this up online. After 1 hour, heat the enzyme mix on a heat block to 80°C for 10 mins to inactive the enzyme.
2. Inoculate 10 ml MNGE to OD600 = 0,1 (or simply 1/100) with overnight culture of *B. subtilis* W168. I usually do, 250 µL of *B. subtilis* cells in LB medium and 9.75 mL of MNGE medium (see below). Add this into an autoclaved conical flask, cover with foil, label with your name, the culture and date.
3. Let this grow to OD600 = 0.9-1.3 at 37°C with agitation (at least 200 rpm!). Use the 1 cm cuvette and as a blank use LB medium. Once OD600 has been reached, take 400 µL of cells. To grow to OD600 = 0.9-1.3 takes about 4 hours.
4. Once you reach the right OD, take 400 µL of your cells from the conical flask and add to a sterile 15 mL falcon tube. Make x2 version of this.
5. Add DNA (1-2 µg linearized plasmid) to one falcon tube, and to the other add nothing.
6. Let it grow for 1 h at 37’C (shaking incubator)
7. Let grow for 1 h
8. Plate on selective media – LB agar with chloramphenicol. – incubate O/N at 37’C in a static incubator.

###### 10x MN-Medium:

* 136g K2HPO4 (x3 H2O)
* 60g KH2PO4
* 10g Na-citrat (x2 H2O)

###### MNGE-Medium:

* 9.2 ml 1 x MN-Medium (920 µl 10x MN + 8.28 ml sterile water)
* 1 ml Glucose (20%)
* 50 µl K-Glutamat (40%)
* 50 µl Fe[III]- ammonium-citrate (2,2 mg/ml)
* 100 µl Tryptophan (5 mg/ml)
* 30 µl MgSO4 (1M)
* (100 µl threonine (5 mg/ml) for strains carrying an insertion in thrC)

###### Expression Mix:

* 500 µl yeast extract (5%)
* 250 µl casamino-acids (CAA) (10%)
* 250 µl H2O
* 50 µl Tryptophan (5 mg/ml)

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{% block title %}Results{% endblock %}
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[The PpstS promoter – our journey to make a phosphate biosensor.](#slide1-chapter1)

[Design of a heterologous NOT gate for B. subtilis](#slide1-chapter2)

[PmaeN – A malate inducible promoter from Bacillus subtilis](#slide1-chapter3)

[Generating a phosphate release circuit](#slide1-chapter4)

[Project Achievements](#slide1-chapter5)

[References](#slide1-chapter6)

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## The PpstS promoter – our journey to make a phosphate biosensor.

###### 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 PpstS promoter – our design for a phosphate biosensor

The promoter PpstS, 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 PpstS 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 PpstS promoter to initiate transcription of phosphate importers and binding proteins [[1–3]](#ref1). A simplified schematic of this is shown in Fig. 1 which outlines the design of the proposed phosphate biosensor. Here, the promoter PpstS ([BBa\_K4418004](http://parts.igem.org/Part%3ABBa_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).

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig1.png)

**Figure 1.**  Circuit schematic for the PpstS phosphate biosensor. A) Under phosphate replete conditions the promoter PpstS 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 PpstS to drive luxABCDE (luciferase) activity.

###### PCR amplification of the PpstS promoter

To construct the phosphate biosensor, we first amplified the PpstS 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 PpstS. A single colony was identified which was screened for the insertion of the PpstS promoter using colony PCR. As shown in Fig. 2B, a 234 bp fragment corresponding to the insertion of PpstS into pBSGGlux was identified.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig2.png)

**Figure 2.**  Cloning of the PpstS promoter to generate a PpstS-luxABCDE transcriptional fusion. A) PpstS was amplified from B. subtilis gDNA the resulting product (161 bp) visualised using agarose gel electrophoresis. “L” corresponds to ladder, “PpstS” is the promoter, “-“ corresponds to the negative control B) Ligation of PpstS 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 PpstS-pBSANDlux into the B. subtilis genome

Following the cloning of the promoter PpstS 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.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig3.png)

**Figure 3.**  Verification of the integration of PpstS-pBSANDlux at the sacA locus. B. subtilis transformants were screened for the integration of the PpstS-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 PpstS-luxABCDE biosensor displays the behaviour of a NOT gate.

Following the confirmation of stable integration of the plasmid PpstS-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 PpstS 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 PpstS 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.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig4.png)

**Figure 4.**  Activity of the PpstS-luxABCDE phosphate biosensor. Measurement of PpstS 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 PpstS promoter in B. subtilis, regulated by the PhoPR two-component system. Our work presents successful cloning of the PpstS promoter and demonstrates the PpstS promoter is repressible in the presence of increasing phosphate concentrations, functioning as a NOT logic gate circuit. The promoter PpstS ([BBa\_K4418004](http://parts.igem.org/Part%3ABBa_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 PpstS 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.

## Design of a heterologous NOT gate for B. subtilis

###### Background

As we have shown using out phosphate inducible promoter [BBa\_K4418004](http://parts.igem.org/Part%3ABBa_K4418004), increasing phosphate concentrations result in repression of the promoter PpstS. 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 PpstS 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 PpstS 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 PpstS 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.

![](https://static.igem.wiki/teams/4418/wiki/images/engineeringfig1.png)

**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 PpstS. 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 PpstS 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](http://parts.igem.org/Part%3ABBa_K1351039)) to drive the expression of the heterologous repressor SarA ([BBa\_K4418005](http://parts.igem.org/Part%3ABBa_K4418005)) of Staphylococcus aureus, the target promoter of which is the PsprC ([BBa\_K4418006](http://parts.igem.org/Part%3ABBa_K4418006))[[4]](#ref4). 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 PpstS 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](http://parts.igem.org/Part%3ABBa_K090505)). To prevent read-through from PxylA, a strong transcriptional terminator [BBa\_K4418007](http://parts.igem.org/Part%3ABBa_K4418007) is used. The resulting circuit should show a reduction in luxABCDE in the presence of xylose through the composite part [BBa\_K4418008](http://parts.igem.org/Part%3ABBa_K4418008).

![](https://static.igem.wiki/teams/4418/wiki/images/engineeringfig2.png)

**Figure 6.**  Schematic of the PxylA inducible NOT gate ( [BBa\_K4418008](http://parts.igem.org/Part%3ABBa_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 PsprC promoter and luxABCDE activity. Boxed in blue is composite part comprising the xylose-inducible NOT gate, part [BBa\_K4418008](http://parts.igem.org/Part%3ABBa_K4418008).

Following xylose induction, SarA is able to repress its’ cognate promoter PsprC 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.

![](https://static.igem.wiki/teams/4418/wiki/images/engineeringfig3.png)

**Figure 7.**  Activity of the xylose inducible NOT gate. Promoter activity of PsprC 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-PsprC 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 PpstS 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 PsprC by SarA to provide greater repression of the output module used with this NOT gate circuit.

## PmaeN – A malate inducible promoter from Bacillus subtilis

###### 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]](#ref5). 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 PmaeN promoter ( [BBa\_K4418000](http://parts.igem.org/Part%3ABBa_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 PmaeN promoter, allowing for expression of enzymes to enable to usage of malate as a carbon source [[6]](#ref6). A simplified schematic of this is shown in Fig. 8. To build this circuit, we cloned the PmaeN promoter into the luciferase reporter plasmid pBSGGlux.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig8.png)

**Figure 8.** Schematic of the PmaeN – luxABCDE biosensor in B. subtilis. The malate responsive promoter PmaeN 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 PmaeN promoter, resulting in measurable *luxABCDE* expression.

###### Activity of the [BBa\_K4418000](http://parts.igem.org/Part%3ABBa_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](http://parts.igem.org/Part%3ABBa_K4418003).

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig9.png)

**Figure 9.** Dose response curve of the PmaeN-luxABCDE reporter. Cells harbouring the integrated PmaeN-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/OD600¬ 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](http://parts.igem.org/Part%3ABBa_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](http://parts.igem.org/Part%3ABBa_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.

## Generating a phosphate release circuit

###### 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](http://parts.igem.org/Part%3ABBa_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]](#ref7). We speculated, that by coupling the induction of the PmaeN ( [BBa\_K4418000](http://parts.igem.org/Part%3ABBa_K4418000)) to the release of both enzymes GlpQ ( [BBa\_K4418001](http://parts.igem.org/Part%3ABBa_K4418001)) and PhoD  ( [BBa\_K4418002](http://parts.igem.org/Part%3ABBa_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 PmaeN [BBa\_K4418000](http://parts.igem.org/Part%3ABBa_K4418000), and both WTA degrading enzymes GlpQ ( [BBa\_K4418001](http://parts.igem.org/Part%3ABBa_K4418001)) and PhoD  ( [BBa\_K4418002](http://parts.igem.org/Part%3ABBa_K4418002)) - both of which are translated by the consensus B. subtilis RBS sequence ( [BBa\_K090505](http://parts.igem.org/Part%3ABBa_K090505)). A schematic of the resulting composite part [BBa\_K4418003](http://parts.igem.org/Part%3ABBa_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.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig10.png)

**Figure 10.** Circuit schematic of the phosphate release circuit [BBa\_K4418003](http://parts.igem.org/Part%3ABBa_K4418003). Induction with malate allows transcriptional activation from the PmaeN 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]](#ref7). 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.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig11.png)

**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.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig12.png)

**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.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig13.png)

**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*.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig14.png)

**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 Δ*phoD* strain from the Bacillus subtilis BKK collection (kindly provided by Prof. Susanne Gebhard). Isolated genomic DNA from this B. subtilis Δ*phoD* strain was used to transform the prior constructed B. subtilis Δ*glpQ* strain in order to remove the gene *phoD*, generating a double mutant (B. subtilis Δ*glpQ*/Δ*phoD*), as shown in Fig. 15A-B.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig15.png)

**Figure 15.**  PhoD deletion in a B. subtilis Δ*glpQ* genetic background. Isolated genomic DNA (gDNA) from a B. subtilis Δ*phoD* strain was used to transform a B. subtilis Δ*glpQ* strain. A) Negtaive control without the addition of B. subtilis Δ*phoD* gDNA. B) B. subtilis Δ*glpQ* strain transformed with gDNA from B. subtilis Δ*phoD*. 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 PmaeN ([BBa\_K4418000](http://parts.igem.org/Part%3ABBa_K4418000)), and the enzymes GlpQ ([BBa\_K4418001](http://parts.igem.org/Part%3ABBa_K4418001)) and PhoD ([BBa\_K4418002](http://parts.igem.org/Part%3ABBa_K4418002)) – the later two which are translated by the B. subtilis consensus RBS sequence ([BBa\_K090505](http://parts.igem.org/Part%3ABBa_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.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig16.png)

**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, PmaeN, 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 Δ*phoD*/Δ*glpQ* 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.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig17.png)

**Figure 17.**  Screening colonies for integration of the phosphate release circuit in the B. subtilis Δ*phoD*/Δ*glpQ*. 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 Δ*phoD*/Δ*glpQ* 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 Δ*phoD*/Δ*glpQ* 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 Δ*phoD*/Δ*glpQ* 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.

![](https://static.igem.wiki/teams/4418/wiki/images/resultsfig18.png)

**Figure 18.** Testing phosphate release in the presence of malate. B. subtilis cells, including the B. subtilis Δ*phoD*/Δ*glpQ* 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.

## Project Achievements

###### Successes

* Engineering PhoBac to sense phosphate and upregulate genes in response using our NOT gate circuit
* Engineering PhoBac to sense malate and
* Feeding the experimental data into an ODE model to better characterise our biosensor.
* Successfully developing a prototype lab-on-chip portable phosphate detection device.

###### Failures

* The Xylose promoter improved to have less leaky expression failed to be induced.
* The asRNA repression circuit designed, built and tested when SarA repressor NOT gate did not function.
* The survey we developed for the public and farmers and published using Facebook advertisement campaigns did not attract respondents.

###### Next Steps

* Integrate both the phosphate uptake and release circuits into a single strain.
* Improve the phosphate uptake ability of PhoBac.
* Conduct studies of PhoBac in soil, measuring survivability and ecological interactions

## References

1. Qi, Y., Kobayashi, Y. & Hulett, F. M. The pst operon of Bacillus subtilis has a phosphate-regulated promoter and is involved in phosphate transport but not in regulation of the Pho regulon. J. Bacteriol. 179, 2534–2539 (1997).
2. Liu, W., Qi, Y. & Hulett, F. M. Sites internal to the coding regions of phoA and pstS bind PhoP and are required for full promoter activity. Mol. Microbiol. 28, 119–130 (1998).
3. Devine, K. M. Activation of the PhoPR-mediated response to phosphate limitation is regulated by wall teichoic acid metabolism in Bacillus subtilis. Front. Microbiol. 9, 1–7 (2018).
4. Mauro, T., Rouillon, A. & Felden, B. Insights into the regulation of small RNA expression: SarA represses the expression of two sRNAs inStaphylococcus aureus. Nucleic Acids Res. 44, 10186–10200 (2016).
5. Canarini, A., Kaiser, C., Merchant, A., Richter, A. & Wanek, W. Root exudation of primary metabolites: Mechanisms and their roles in plant responses to environmental stimuli. Front. Plant Sci. 10, (2019).
6. Tanaka, K., Kobayashi, K. & Ogasawara, N. The Bacillus subtilis YufLM two-component system regulates the expression of the malate transporters MaeN (YufR) and YfIS, and is essential for utilization of malate in minimal medium. Microbiology 149, 2317–2329 (2003).
7. Myers, C.L., Li, F.K.K., Koo, B.-M., El-Halfawy, O.M., French, S., Gross, C.A., Strynadka, N.C.J. and Brown, E.D., 2016. Identification of Two Phosphate Starvation-induced Wall Teichoic Acid Hydrolases Provides First Insights into the Degradative Pathway of a Key Bacterial Cell Wall Component. Journal of Biological Chemistry [Online], 291(50), pp.26066–26082. Available from: https://doi.org/10.1074/jbc.m116.760447 [Accessed 12 October 2022].

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