Part:BBa_K3151027
Hydrogenase c-di-GMP Phosphodiesterase Biosensor
Hydrogenase c-di-GMP Phosphodiesterase Biosensor
Sequence and Features
- 10COMPATIBLE WITH RFC[10]
- 12INCOMPATIBLE WITH RFC[12]Illegal NheI site found at 7
Illegal NheI site found at 30
Illegal NotI site found at 1166
Illegal NotI site found at 3782 - 21INCOMPATIBLE WITH RFC[21]Illegal BglII site found at 4210
- 23COMPATIBLE WITH RFC[23]
- 25INCOMPATIBLE WITH RFC[25]Illegal NgoMIV site found at 1025
Illegal NgoMIV site found at 4010
Illegal NgoMIV site found at 4354 - 1000INCOMPATIBLE WITH RFC[1000]Illegal BsaI.rc site found at 347
Illegal BsaI.rc site found at 2041
Overview
The sensory Magnetospirillum magneticum [NiFe] hydrogenase operon consists of four parts: 1 kb hurS encoding a small subunit, 1.6 kb hurL encoding a large subunit, 0.5 kb hupD encoding maturation protease, and 2 kb hurR encoding cyclic-di-GMP phosphodiesterase. Our hydrogenase c-di-GMP phosphodiesterase biosensor is comprised of large subunit [BBa_K3151012], small subunit [BBa_K3151013], maturation protease [BBa_K3151014] and cyclic-di-GMP phosphodiesterase [BBa_K3151015].
Literature
[NiFe] hydrogenases show higher affinity for molecular hydrogen compared to [FeFe] hydrogenases [1].
One of the many advantages of [NiFe] hydrogenases is their fast response to the presence of molecular hydrogen. They usually have high turnover numbers (Kcat, s-1), which means they are capable of oxidising a large number of hydrogen molecules per second. In a previous study, Pershad et al. investigated the redox activity of [NiFe] hydrogenase from Chromatium vinosum using protein film voltammetry [2]. They found that the Kcat value was between 1500 – 1900 s-1 at 10% partial hydrogen pressure, 30°C, and pH 5 – 8, although there was a strong force driving the reverse reaction, i.e. reduction of H+ to H2 [2]. A number of enzyme kinetic studies also revealed high Kcat of [NiFe] hydrogenases under various reaction conditions [3, 4]. The high catalytic activity means that these hydrogenases can rapidly generate large amounts of H+, which in turn activate a series of downstream signalling events and cellular response.
The future of hydrogen gas biosensing lies in whole cell-based biosensors which use genetically modified living organisms to detect the presence of target substances. A few designs were introduced in the past for monitoring gaseous ammonia and carbon monoxide [5, 6]. It is expected that whole cell-based biosensors for hydrogen detection will be made available in the near future.
Furthermore, [NiFe] hydrogenases maintain high-affinity oxidative activity in a broad range of tropospheric hydrogen concentration [7], which cannot be achieved by physicochemical hydrogen sensors. This feature is crucial for Team Macquarie Australia 2019 project in building a hydrogen biosensor to meet the requirement of overcoming the cross-sensitivity problem faced by current hydrogen gas sensors.
Assembly and design
We added EcoRI (GAATTC), XbaI (TCTAGA), SalI (GTCGAC), SpeI (ACTAGT), and PstI (CTGCAG) restriction enzyme cleavage sites within the sequence.
All genes within this plasmid are sequences obtained from Magnetospirillum magneticum and codon optimised to be expressed in Escherichia coli.
Usage
Hydrogenase (Large subunit hydrogenase + small subunit hydrogenase + Maturation protease)
The maturation protease will first cleave the C-terminus of [NiFe] hydrogenase large subunit after the Nickel insertion [8]. The large and small subunit will then come together to form the hydrogenase part of the biosensor.
The hydrogenase with all three components was used in the hydrogenase consumption assay.
Phosphodiesterase
The mature hydrogenase (large and small subunits together) will come together with the phosphodiesterase to form the sensory component of the hydrogen biosensor for Team Macquarie Australia 2019. Cyclic-di-GMP phosphodiesterase breaks down cyclic-di-GMP [8]. The change in cyclic-di-GMP levels in the cell will affect the amount of biofilm produced.
Part Characterisation
We have successfully integrated the hydrogenase part Hyd A of M. magneticum [NiFe] hydrogenase into E. coli DH5α cells. The hydrogen consumption assay has confirmed that our hydrogenase is functional.
In order to test the functionality of the hydrogenase part of M. magneticum [NiFe] hydrogenase, we measured hydrogen desaturation rates (mV/s) in water and in E. coli DH5α Hyd A transformant cells (which contain hydrogenase) using Clark-type electrode. This test is called the hydrogenase consumption assay.
Testing the functionality of the hydrogenase part requires a sample that has all three components (small subunit, large subunit, and maturation protease). As seen in Figure 1, HydA 7 which has all three components present was chosen for the assay.
Figure 1. SDS-PAGE showing the three parts of the hydrogenase biosensor. <i>HurL (large subunit), HurS (small subunit), and HupD (maturation protease). Four biological replicates of HydA 7, HydA 3, HydA 4, and HydA 2. HydA 7 was chosen for the assay.</i>
As seen in Figure 2 and 3, the results showed higher rates of hydrogen saturation and desaturation for HydA 7 transformant than DH5α cells without [NiFe] hydrogenase. In the presence of hydrogen, HydA 7 cells appeared to consume hydrogen, which resulted in more hydrogen being absorbed into the solution and decreased of hydrogen concentration in the electrode chamber. In contrast, non-transformant DH5α cells did not show significant intake of hydrogen from the environment, resulting in lower hydrogen saturation rate. The results from this experiment demonstrated that our [NiFe] hydrogenase was functional and was capable of oxidising molecular hydrogen into protons and electrons.
Figure 2. Maximum hydrogen saturation rates of water (negative control), DH5α (positive control), and DH5α HydA 7 transformant. Standard deviation n=2.
Figure 3. Maximum hydrogen desaturation rates of water (negative control), DH5α (positive control), and DH5α HydA 7 transformant. Standard deviation n=2.
References
[1] Frey M. Hydrogenases: Hydrogen-Activating Enzymes. ChemBioChem. 2002;3(2-3):153-160.
[2] Pershad H, Duff J, Heering H, Duin E, Albracht S, Armstrong F. Catalytic Electron Transport inChromatium vinosum[NiFe]-Hydrogenase: Application of Voltammetry in Detecting Redox-Active Centers and Establishing That Hydrogen Oxidation Is Very Fast Even at Potentials Close to the Reversible H+/H2Value. Biochemistry. 1999;38(28):8992-8999.
[3] Preissler J, Wahlefeld S, Lorent C, Teutloff C, Horch M, Lauterbach L et al. Enzymatic and spectroscopic properties of a thermostable [NiFe]‑hydrogenase performing H2-driven NAD+-reduction in the presence of O2. Biochimica et Biophysica Acta (BBA) - Bioenergetics. 2018;1859(1):8-18.
[4] van der Linden E, Burgdorf T, de Lacey A, Buhrke T, Scholte M, Fernandez V et al. An improved purification procedure for the soluble [NiFe]-hydrogenase of Ralstonia eutropha: new insights into its (in)stability and spectroscopic properties. JBIC Journal of Biological Inorganic Chemistry. 2006;11(2):247-260.
[5] Bohrn U, Stütz E, Fuchs K, Fleischer M, Schöning M, Wagner P. Monitoring of irritant gas using a whole-cell-based sensor system. Sensors and Actuators B: Chemical. 2012;175:208-217.
[6] Bohrn U, Stütz E, Fuchs K, Fleischer M, Schöning M, Wagner P. Air Quality Monitoring using a Whole-Cell based Sensor System. Procedia Engineering. 2011;25:1421-1424.
[7] Greening C, Berney M, Hards K, Cook G, Conrad R. A soil actinobacterium scavenges atmospheric H2 using two membrane-associated, oxygen-dependent [NiFe] hydrogenases. Proceedings of the National Academy of Sciences. 2014;111(11):4257-4261.
[8] Greening C, Biswas A, Carere C, Jackson C, Taylor M, Stott M et al. Genomic and metagenomic surveys of hydrogenase distribution indicate H2 is a widely utilised energy source for microbial growth and survival. The ISME Journal. 2015;10(3):761-777.
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