Team:Macquarie Australia/Results
RESULTS
Riboswitch
Our biosensor utilises a riboswitch to control the expression of enhanced green fluorescent protein (eGFP) through the modulation of intracellular cyclic-di-GMP concentration. This regulation occurs at the transcription level. As the riboswitch/eGFP is being transcribed by the RNA polymerase, cyclic-di-GMP can bind to the aptamer region of the riboswitch (Fig. 1), which will cause the formation of a hairpin terminator loop structure, preventing the transcription of the downstream eGFP gene.
Figure 1: BioBrick structure of the riboswitch/eGFP operon. The cyclic-di-GMP binds at the riboswitch aptamer, forming the terminator structure and preventing the transcription of the eGFP region.
In order to test the function of our riboswitch, we developed two riboswitch/eGFP operon constructs; one containing the terminator loop region within the riboswitch and one without. We hypothesised that the construct containing the terminator stem structure would constitutively express eGFP, regardless of cyclic-di-GMP concentration. Because of this constant activity, we refer to this construct as “R ON” (for riboswitch ON) [BBa_K3151002]. Alternatively, the construct with the terminator stem would have the expression of eGFP be repressed by native cellular concentrations of cyclic-di-GMP, with this construct being referred to as “R OFF” (for riboswitch OFF) [BBa_K3151011].
Figure 2: The hypothesised activity of the two riboswitch constructs. While the RNA polymerase (red) moves along the operon (blue), cyclic-di-GMP (green) will bind to the aptamer region of the riboswitch transcript (black). When bound to cyclic-di-GMP, the R OFF construct (Fig. 2.1a) will produce the terminator loop structure (Fig. 2.1b) and cease transcription, preventing the expression of eGFP. When cyclic-di-GMP binds to the R ON transcript (Fig. 2.2a) it does not form the terminator loop structure (Fig 2.2b), thereby having no effect on the expression of eGFP.
We ordered these sequences from Twist Bioscience and assembled them via BioBrick standard assembly. Assembly confirmation was performed using restriction digestion with EcoRI and PstI (Fig. 3) and sequencing (Macrogen). These constructs were transformed into E. coli DH5α.
Figure 3: Agarose gel of single and double digests of sequence confirmed parts.
The promoter selected for this construct is a stationary phase promoter [BBa_J45992], which was selected because cyclic-di-GMP concentration varies with cell growth cycles, with its highest level of expression being as it enters stationary phase[1]. We also tested the function of the constructs with two inducible promoters; Lac [BBa_R0010] and Tac [BBa_K180000] to characterise the efficacy of our promoter.
We collaborated with Team NTU-Singapore (Nanyang Technological University) to perform Quantitative Polymerase Chain Reaction (qPCR) to measure the rate of transcription of eGFP between the R ON and R OFF constructs under the control of the Lac promoter (Fig. 4). These results demonstrate that between 4 and 24 hours, when the cells are entering stationary phase, the increased concentration of cyclic-di-GMP results in a significant decrease in the transcription of the eGFP sequence in the R OFF transformants.
Figure 4: qPCR of R ON and R OFF constructs expressed in E. coli DH5ɑ under the Lac promoter. Samples were induced with 0.5 mM IPTG. The forward and reverse primers used for the qPCR both lay within the eGFP sequence.
In order to characterise the mode of action of the riboswitch/promoter combinations, we collaborated with Team Sydney Australia to perform a GFP fluorescence assay[2]. Samples were measured with the BMG Pherastar plate reader to measure eGFP (Ex485 nm Em520 nm) and OD (600 nm). These results demonstrate that the Lac and Tac promoters have significantly higher background expression of the R OFF construct.
This is likely due to the strength of the promoters, and the length of the transcripts produced by either construct. This is demonstrated clearly by the results of the Tac promoter (Fig. 6 & 9), in which by the end of the assay the R OFF samples were producing higher levels of eGFP. We believe that this is due to the fact that the binding of cyclic-di-GMP to the transcript results in shorter transcripts, allowing the RNA polymerase to produce more of the transcripts quickly and saturating the cyclic-di-GMP, resulting in a loss of inhibition of eGFP transcription. This effect can also be observed in the samples under control of the Lac promoter (Fig. 5 & 8). By contrast, the background level of transcription under the stationary phase promoter remains constant throughout the entire assay (Fig. 7), making it a more suitable candidate for our biosensor.
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| Figure 5: eGFP produced by cells transformed with the Lac riboswitch constructs over 60 hours. | Figure 6: eGFP produced by cells transformed with the Tac riboswitch constructs over 60 hours. | Figure 7: eGFP produced by cells transformed with the Stationary phase riboswitch constructs over 60 hours. |
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| Figure 8: eGFP produced by cells transformed with the Lac riboswitch constructs over 30 hours. Data provided by Usyd iGEM. | Figure 9: eGFP produced by cells transformed with the Tac riboswitch constructs over 30 hours. Data provided by Usyd iGEM. |
Phosphodiesterase
Cyclic-di-GMP acts as the secondary messenger within our detect-response system. It prevents the response (eGFP production) by binding to the terminator stem of the riboswitch. The hydrogenase we are utilising as our sensor contains a phosphodiesterase domain, which is modulated by the presence of molecular hydrogen. This allows it to digest cyclic-di-GMP, reducing the cellular concentration and allowing for the production of eGFP.
Figure 10: Hypothetical mode of action of the sense/response system of our biosensor. The hydrogenase will have its phosphodiesterase domain activated in the presence of molecular hydrogen. This will lead to a decrease in intracellular cyclic-di-GMP, preventing the riboswitch from forming the terminator loop structure and allowing the transcription of eGFP.
In order to test this functionality, we assembled a plasmid containing a well characterised cyclic-di-GMP phosphodiesterase, yhjH [BBa_K861090], with an inducible Lac promoter [BBa_R0010] to create a new composite part [BBa_K3151008]. We also assembled a plasmid with just the yhjH gene with no promoter and terminator as a control. We ordered these sequences from Integrated DNA Technologies and assembled into BioBricks using standard assembly. Assembly was confirmed through restriction digestion (Fig. 3) and sequencing. These constructs were then transformed into the E. coli DH5α and Nissle 1917.
As intracellular cyclic-di-GMP concentration is directly correlated with biofilm formation[2], we can utilise the biofilm forming capability of the transformants as a proxy for intracellular cyclic-di-GMP concentration and phosphodiesterase activity. Nissle 1917 E. coli cells transformed with these constructs were spotted onto LB agar plates supplemented with Congo Red and Coomassie Blue[3], and the cells transformed with the construct containing the promoter and terminator sequences produced significantly less cellulose than cells transformed with the construct without the promoter and terminator, as well as parental Nissle 1917 cells (Fig. 11), indicating significant phosphodiesterase activity within the cell.
Figure 11: E. coli Nissle 1917 cells transformed with the yhjH phosphodiesterase with (A) and without (B) promoter and terminator, as well as negative control (C). Cells were spotted onto LB agar plates supplemented with Congo Red and Coomassie Blue, and visualised with Blue Light LEDs. Nissle 1917 was selected as it produces significantly higher levels of cellulose in its biofilm. The red colour observed in (B) and (C) is the Congo Red staining the cellulose.
E. coli DH5α cells transformed with these same constructs were used to perform a microtitre plate adherence capacity assay[X], another measure of the cells’ capacity for forming biofilms. This assay was particularly prone to error as the Crystal Violet used to stain the biofilm was particularly difficult to remove entirely, and extraneous stain increased the absorbance reading (Fig. 12).
Figure 12: Results from adherence capacity assay with E. coli DH5α transformed with the yhjH phosphodiesterase w/ and w/o promoter and terminator, as well as negative control parental cells.
This yhjH phosphodiesterase has been used in a previous composite part [BBa_K2471001], which utilised the T7 promoter for its expression. However, as we were not planning on assembling our constructs into cells with T7 RNA polymerase, as well as wishing to have inducible expression of our parts, we decided to improve upon the design of the previous BioBrick to better suit out purposes. We transformed both our Lac yhjH BioBrick as well at the T7 yhjH BioBrick into the E. coli strains DH5a, Nissle 1917 and BL21(DE3). These transformants were then spotted onto LB plates supplemented with Congo Red and Coommassie Blue in order to assay for their biofilm production capability (Fig. 13).
The spot assay shows a significant reduction in cellulose synthesis in Nissle 1917 cells transformed with the Lac+yhjH compared with negative control, and little to no reduction in cells transformed with the T7+yhjH (Fig. 13). No significant change was seen in any of the DH5a cells (Fig. 15). No significant difference in colony morphology was observed between negative control BL21(DE3) and BL21(DE3) cells transformed with T7+yhjH, though some reduction in Congo-Red staining was observed in BL21(DE3) cells transformed with Lac-yhjH (Fig. 14), which indicates a reduction in cellulose biosynthesis and therefore an increase in phosphodiesterase activity.
Figure 13: Spot Assay of the negative control Nissle 1917 (A), Nissle 1917 with T7+yhjH (B), and Nissle 1917 with Lac+yhjH (induced with 1 mM IPTG) (C). Plate observed under blue light to enhance visualisation of Congo Red-stained cellulose.
Figure 14: Spot Assay of the negative control BL21(DE3) (A), BL21(DE3) with T7+yhjH (B), and BL21(DE3) with Lac+yhjH (induced with 1 mM IPTG) (C). Plate observed under white light to enhance visualisation of colony morphology.
Figure 15: Spot Assay of the negative control DH5a (A), DH5a with T7+yhjH (B), and DH5a with Lac+yhjH (induced with 1 mM IPTG) (C). Plate observed under white light to enhance visualisation of colony morphology.
Phosphodiesterase and Riboswitch
In order to characterise the interaction between the function of our riboswitch and phosphodiesterases, we constructed two plasmids containing the coding sequences both the riboswitch/eGFP operon (either R ON and R OFF) under the stationary phase promoter and the yhjH gene under the Lac promoter, referred to as yhjH + RON [BBa_K3151029] and yhjH + ROFF [BBa_K3151030] respectively. We confirmed the assembly through restriction digestion as well as sequence confirmation. These plasmids were then transformed into the E. coli strains DH5α and Nissle 1917.
E. coli DH5α cells transformed with these plasmids we used to perform fluorescence assays using the BMG Pherastar plate reader to measure eGFP (Ex485 nm Em520 nm) and OD (600 nm). These results (Fig. 16) confirm our hypothesis that the ROFF construct will produce significantly lower levels of eGFP due to inhibition from cellular cyclic-di-GMP.
Figure 16: The constitutive riboswitch [BBa_K3151011] was expressed in DH5ɑ cells. The black lines represent R ON samples, the blue lines represent R OFF samples. The increased phosphodiesterase activity in the ROFF samples allowed the eGFP to be expressed at higher than baseline levels, though still significantly lower than RON samples.
Nissle 1917 E. coli cells transformed these plasmids were spotted onto LB agar plates supplemented with Congo Red and Coomassie Blue. We compared the effect of induction of the yhjH phosphodiesterase between the yhjH+R ON and yhjH+R OFF transformants. Induction of yhjH appeared to have no effect on the fluorescence of yhjH+R ON transformants (Fig. 17A, Fig. 17C), which corroborates our hypothesis that its expression of eGFP is not influenced by the intracellular concentration of cyclic-di-GMP. Induction of yhjH appeared to significantly increase the fluorescence of yhjH+R OFF transformants (Fig. 17B, Fig. 17D), again corroborating our hypothesis that a decrease in intracellular cyclic-di-GMP will allow for an increase in expression of eGFP.
Figure 17: LB agar plate supplemented with Congo Red and Coomassie blue, spotted with yhjH+R ON induced (A), yhjH+R OFF induced (B), yhjH+R ON uninduced (C), and yhjH+R OFF uninduced (D).
Hydrogenase
We ordered hydrogenase constructs pUC57Kan-HydA (Hydrogenase large and small subunits, maturation protease) and pUC57Amp-HydB (cyclic-di-GMP phosphodiesterase) from GeneWiz. They were transformed into E. coli DH5α competent cells and amplified. To confirm the presence of the two parts, we performed single and double restriction enzyme digestions, followed by agarose gel electrophoresis and SDS-PAGE. The results were consistent with our expectations (Fig. 18–20).
For pUC57Kan-HydA (Fig. 18), we observed a single band at about 6000 bp from single digestion, representing the linearised plasmid DNA; for double digestion, a band at 3300 bp represented HydA, and the two bands at 900 bp and 1500 bp corresponded to the cleaved pUC57Kan.
Figure 18: Agarose gel electrophoresis profiles of Magnetospirillum magneticum [NiFe] hydrogenase operon upstream part HydA. Four biological replicates were analysed.
SDS-PAGE results (Fig. 19) showed distinct protein bands at approximately 17 kDa in two biological replicates, which corresponded to the maturation protease HupD (Sample 4 and 7, Fig. 18b). As the hydrogenase operon was organised in a sequential order of hurS, hurL, followed by hupD, the expression of HupD suggested that both hurS and hurL should have been transcribed and translated into proteins.
Figure 19: SDS-PAGE profiles of Magnetospirillum magneticum [NiFe] hydrogenase operon upstream part HydA. Four biological replicates were analysed.
For double digestion of pUC57Amp-HydB, the bands at 2500 bp and 3000 bp represented pUC57Amp and HydB, respectively. Single digestion of pUC57Amp-HydB gave an intense band at approximately 3500 bp, which was the uncut circular plasmid. (Fig.20).
Figure 20: Agarose gel electrophoresis profile of Magnetospirillum magneticum [NiFe] hydrogenase operon downstream part HydB. Four biological replicates were analysed.
To test the functionality of the hydrogenase part HydA of M. magneticum [NiFe] hydrogenase, we measured the hydrogen saturation and desaturation rates (mV/s) in water, HydA7 (DH5ɑ +hydrogenase) and DH5ɑ (-hydrogenase) using a Clark-type electrode sensor. HydA7 was used for the functionality test, because it has the highest level of protein over-expression. The results showed a significant difference between the DH5ɑ and HydA7 bacterial cells in their rates of hydrogen absorption.
The hydrogen saturation and desaturation rates (Fig. 21 and 22), are represented as mV per second. The saturation rate in water was the highest, followed by HydA7 and then DH5ɑ (Fig. 21). Similarly, the desaturation rate in water was the highest, followed by HydA7 and the lowest, DH5ɑ (Fig. 22). The three samples represented in each graph are water, DH5ɑ (-hydrogenase), and (DH5ɑ +hydrogenase).
Figure 21: Maximum hydrogen saturation rates of water (standard), DH5α (negative control), and DH5α HydA7 transformant. Error bars are +/- StDev, n=2.
Figure 22: Maximum hydrogen desaturation rates of water (standard), DH5α (negative control), and DH5α HydA7 transformant. Error bars are +/- StDev, n=2.
Water is the standard, representing the maximum hydrogen saturation rates. When comparing the DH5ɑ with and without hydrogenase, a clear difference is demonstrated in the maximum rate of H2 saturation (mV/s; Fig. 21). H2 rate of consumption is higher in our HydA7 sample compared to untransformed DH5ɑ cells. There is overexpression of hydrogenase in the DH5ɑ HydA7 sample, therefore, in the presence of hydrogen, the HydA7 cells appeared to bind hydrogen, thus allowing for more hydrogen to be absorbed into solution before its maximum saturation is reached. This may be specifically attributed to the hydrogen binding in the hydrogenase active site. Conversely, the DH5ɑ cells without the hydrogenase did not appear to absorb as much hydrogen, resulting in lower hydrogen saturation rate. This result demonstrates that there is significant hydrogen binding in hydrogenase binding site of our modified HydA7 cells
A clear difference when comparing the DH5ɑ with and without hydrogenase is demonstrated once more with rate of H2 desaturation. When comparing the data for DH5ɑ (-hydrogenase) and our DH5ɑ (+hydrogenase), the desaturation rate for the HydA7 is higher compared to that of the DH5ɑ cells without the hydrogenase, demonstrating the ability of the hydrogenase to bind hydrogen.
Hydrogenases are known to be reversible, where they can both produce H2 or Detect H2. In order to demonstrate that our hydrogenase ([BBa_K3151025]) does not function as a H2 producer, we performed a hydrogen electrode test (Fig. 23) comparing it to the hydrogen producing gene cluster (HPGC). This gene cluster was taken from the Macquarie_Australia iGEM 2017 team. In black, is represented by E. coli (+HPGC) and in blue, is represented by E. coli (+Hydrogen sensor). The clear indication of hydrogen gas production can be seen by the HPGC, as opposed to the HydA7 (Hydrogen Sensor), which shows no indication that any hydrogen gas is being produced. This result shows that our [NiFe] hydrogenase taken from M. magneticum operates only in the direction of oxidizing hydrogen, and that the oxidation is not reversible.
Figure 23: Hydrogen production in DH5α HydA7 transformant following addition on 20 mM glucose.
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