Patent Application
20220235384
The invention relates to a nano-biohybrid (e.g., a nanoorg) organism (e.g. a bacteria) comprising different core-shell quantum dots (QDs) or gold nanoparticle clusters, with excitation ranges having ultraviolet to near-infrared energies, coupled with targeted enzyme sites. When illuminated by light, these nanorgs drive the renewable production of biofuel molecules and chemicals using carbon-dioxide (CO2), water, and nitrogen (from air) as substrates. The disclosed nanorgs catalyze light-induced air-water-CO2 reduction with a high turnover number (TON) of approximately 106-108 (mols of product per mol of cells) to biofuels including, but not limited to, isopropanol (IPA), butane diol, gasoline additives, gasoline substitutes, 2,3-butanediol (BDO), C11-C15 methyl ketones (MKs), and hydrogen (H2); and chemicals such as formic acid (FA), ammonia (NH3), ethylene (C2H4), and degradable bioplastics, e.g. polyhydroxybutyrate (PHB). These nanoorg cells function as nano-microbial factories powered by light.
Attempts were made to combine the desired functionality of direct light-activation in cell free extracts/purified enzymes (Reference No. 3) and whole non-photosynthetic bacteria (Reference No. 4), but these strategies have limited applicability due to either enzyme de-activation in the air, or specific tolerance of the bacteria to inorganic elements.
There has been intensive searches for new methods to combine multiple functionalities (e.g., light, voltage, or magnetic field stimulation) of inorganic nanomaterials with the versatility of designed synthetic metabolic networks in living cells, to simply “grow” such hybrid catalysts by the addition of inorganic nanomaterials to the media/buffered water, thereby combining specificity of biocatalysts with high-throughput of inorganic nanomaterials.
However, living cells do not interface naturally with nanoscale materials.
Because such artificial organisms are contemplated to have unprecedented multifunctional properties, such as wireless activation of enzyme function using electromagnetic stimuli, there is a need for new artificial organisms.
The invention relates to a nano-biohybrid (e.g., a nanoorg) organism (e.g. a bacteria) comprising different core-shell quantum dots (QDs) or gold nanoparticle clusters, with excitation ranges having ultraviolet to near-infrared energies, coupled with targeted enzyme sites. When illuminated by light, these nanorgs drive the renewable production of biofuel molecules and chemicals using carbon-dioxide (CO2), water, and nitrogen (from air) as substrates. The disclosed nanorgs catalyze light-induced air-water-CO2 reduction with a high turnover number (TON) of approximately 106-108 (mols of product per mol of cells) to biofuels including, but not limited to, isopropanol (IPA), butane diol, gasoline additives, gasoline substitutes, 2,3-butanediol (BDO), C11-C15 methyl ketones (MKs), and hydrogen (H2); and chemicals such as formic acid (FA), ammonia (NH3), ethylene (C2H4), and degradable bioplastics, e.g. polyhydroxybutyrate (PHB). These nanorg cells function as nano-microbial factories powered by light.
In one embodiment, the present invention provides a method of producing a biofuel, comprising, a) providing, i) a live engineered Cupriavidus necator bacteria comprising an exogenous protein enzyme that produces a biofuel molecule and a quantum dot that transmits electrons having energies in the range of the reduction potential of said enzyme upon exposure to radiation for boosting production of said biofuel molecule, ii) an illumination source for emitting radiation, and iii) compounds comprising CO2, H2O, O2 and N2, b) incubating said live engineered bacteria in the presence of said compounds in the dark, and c) illuminating said live engineered bacteria with said source for producing a biofuel molecule. In one embodiment, said biofuel is a biodiesel. In one embodiment, said biofuel is a methyl ketone (MK). In one embodiment, said biodiesel is a methyl ketone (MK). In one embodiment, said methyl ketone (MK) ranges from C11-C15. In one embodiment, said methyl ketone (MK) comprises C11-C15. In one embodiment, said biofuel is a biodiesel methyl ketone ranging from C11-C15. In one embodiment, said biofuel is a gasoline 2,3-butanediol (BDO) molecule. In one embodiment, said biofuel molecule is ethylene. In one embodiment, said biofuel molecule is isopropanol. In one embodiment, said biofuel molecule is an additive to fuel. In one embodiment, said biofuel molecule is butane diol. In one embodiment, said biofuel molecule is a gasoline additives. In one embodiment, said biofuel molecule is a gasoline substitute. In one embodiment, said illumination source is selected from the group consisting of a light bulb, a fluorescent bulb, a light-emitting diode (LED) and sunlight (e.g. solar radiation). In one embodiment, said boosting production is increasing production over said engineered bacteria that is not comprising said quantum dot.
In one embodiment, the present invention provides a core-shell quantum dot (QD), wherein said core is selected from the group consisting of cadmium sulfide (CdS), cadmium selenide (CdSe), indium phosphide (InP), and copper zinc tin sulfide (Cu2ZnSnS4) (CZTS), wherein said shell is a zinc sulfide (ZnS) shell. In one embodiment, said ZnS shell is a two monolayer shell. In one embodiment, said QD is selected from the group consisting of CZS1, CZS2, CZSe1, CZSe2, CZSe3, IPZS and CZTS. In one embodiment, said QD core has a zwitterion L-cysteine (CYS) ligand cap. In one embodiment, said QD core has a ligand cap selected from the group consisting of a 3-mercaptopropionic acid (MPA) capping ligand and a cysteamine (CA) ligand cap. In one embodiment, said QD has an excitation energy from wavelengths ranging from ultraviolet to near-infrared energies. In one embodiment, said QD has a range of emission energies after exposure to said excitation energies. In one embodiment, said QD further comprises a bacterium, wherein said bacterium expresses an enzyme which has a reduction potential energy range matching said QD emission energies.
In one embodiment, the present invention provides a method of using a core-shell quantum dot (QD), comprising, a) providing, i) a bacteria strain expressing an enzyme for producing a compound, said enzyme having a reduction potential in a range of energies, ii) a plurality of quantum dots (QDs) comprising core-shell quantum dot (QD), wherein said QD in turn comprises a core is selected from the group consisting of cadmium sulfide (CdS), cadmium selenide (CdSe), indium phosphide (InP), and copper zinc tin sulfide (Cu2ZnSnS4) (CZTS), wherein said a two monolayer zinc sulfide (ZnS) shell, and a zwitterion L-cysteine (CYS) ligand cap, wherein said QD emits energies within said range of reduction potential energies after illumination, ii) an illumination source capable of emitting radiation, and b) contacting said bacteria with said QDs such that said QDs are internalized by said bacteria after said ligand binding, and c) irradiating said QD contacted bacteria for providing said emission energies for boosting production amounts of said compound over amounts produced by said QD contacted bacteria prior to illumination. In one embodiment, said energy matched QDs increase enzyme activity of the energy-matched enzyme. In one embodiment, said illumination source emits radiation in wavelengths selected from the group consisting of near-UV, visible, NIR radiation, Light Emitting Diode (LED) and solar radiation. In one embodiment, said QD is selected from the group consisting of CZS1, CZS2, CZSe1, CZSe2, CZSe3, IPZS and CZTS. In one embodiment, said method further provides a solution, wherein said bacteria are added to said solution. In one embodiment, said solution is selected from the group consisting of buffer, growth media, and lysis solution. In one embodiment, said QD contacted bacteria are in said solution during step c), further comprising step d) precipitating said compound out of solution. In one embodiment, said lysis solution comprises a detergent. In one embodiment, said detergent is sodium dodecyl sulfate (SDS). In one embodiment, said method further provides a membrane for filtering said QD contacted bacteria out of solution, then step d), filtering said QD contacted bacteria out of solution. In one embodiment, said filtered QD contacted bacteria are suspended in said solution then repeating step c). In one embodiment, said filtered QD contacted bacteria are suspended in said buffer for harvesting said compounds from said illuminated QD contacted bacteria. In one embodiment, said method further provides a charged filtration membrane, wherein said filtered CZS-QD contacted bacteria are suspended in said lysis solution before or after step c), further comprising a step of filtering said lysed bacteria for recovering said QDs. In one embodiment, said method further comprises a step of using said recovered QDs as said plurality of QDs used in step b).
In one embodiment, the present invention provides a nanorg bacteria strain comprising a core-shell quantum dot (QD), wherein said QD in turn comprises a core is selected from the group consisting of cadmium sulfide (CdS), cadmium selenide (CdSe), indium phosphide (InP), and copper zinc tin sulfide (Cu2ZnSnS4) (CZTS), wherein said a two monolayer zinc sulfide (ZnS) shell, and a zwitterion L-cysteine (CYS) ligand cap.
In one embodiment, the present invention provides a composition comprising an Azobacteria vinelandii bacteria strain comprising a Cadmium sulfide (CdS) core and a zinc sulfide (ZnS) shell (CdS@ZnS) core-shell quantum dot (CZS-QD), and a molybdenum-iron nitrogenase (MFN) enzyme. In one embodiment, said bacteria contains ammonia (NH3) molecules above natural levels. In one embodiment, said bacteria contains hydrogen (H2) molecules above natural levels. In one embodiment, said amount is greater than 105 moles of NH3 per mole of bacteria cells. In one embodiment, said MFN enzyme is heterologous. In one embodiment, said bacteria comprises a cell lysate. In one embodiment, said bacteria are live bacteria. In one embodiment, said live bacteria are replicating. In one embodiment, said shell has two monolayers of zinc sulfide (ZnS). In one embodiment, said CZS-QD has a zwitterion L-cysteine (CYS) cap. In one embodiment, said strain is DJ995. In one embodiment, said composition further comprises CO2/O2 at a ratio of 4:1. In one embodiment, said composition further comprises a growth medium. In one embodiment, said growth medium comprises an amount of said NH3 above levels for said bacteria cells without a CZS-QD. In one embodiment, said method further
In one embodiment, the present invention provides a composition comprising a Cupriavidus necator bacteria strain comprising a Cadmium sulfide core (CdS) and a zinc sulfide (ZnS) shell (CdS@ZnS) core-shell quantum dot and a molybdenum-iron nitrogenase (MFN) enzyme. In one embodiment, said bacteria contains ammonia (NH3) above natural levels. In one embodiment, said bacteria contains hydrogen (H2) molecules above natural levels. In one embodiment, said Cupriavidus necator strain is an engineered strain comprising a pBBRl-efe plasmid. In one embodiment, said MFN enzyme is heterologous to said bacteria strain.
In one embodiment, the present invention provides a method of producing ammonia (NH3), comprising, a) providing, i) a bacteria strain comprising a molybdenum-iron nitrogenase (MFN) enzyme, wherein expression of said MFN enzyme results in ammonia (NH3) formation, a plurality of quantum dots (QDs), wherein said QD is a Cadmium sulfide core and a zinc sulfide (ZnS) shell (CdS@ZnS) (CZS) core-shell quantum dot, wherein said QD transmits electrons having energies in the range of the reduction potential of said MFN enzyme upon exposure to radiation that increases the activity of said MFN enzyme, ii) an illumination source capable of emitting radiation, iii) at least one compound selected from the group consisting of CO2, H2O, O2 and N2, and b) incubating said engineered bacteria in the presence of said at least one compound in the dark, and c) irradiating said engineered bacteria with said illumination source under conditions that produce ammonia (NH3). In one embodiment, said d production of NH3 molecules is above natural levels. In one embodiment, said production of NH3 is an amount greater than in said bacteria strain without said QD. In one embodiment, said production of NH3 is an amount greater than in said bacteria strain without said irradiating. In one embodiment, said bacteria strain is an Azobacteria vinelandii bacteria strain. In one embodiment, said Azobacteria vinelandii strain is DJ995. In one embodiment, said bacteria strain is a Cupriavidus necator bacteria strain. In one embodiment, said Cupriavidus necator strain is an engineered strain comprising a pBBRl-efe plasmid. In one embodiment, said MFN enzyme is heterologous. In one embodiment, said method further provides a bacterial growth medium. In one embodiment, said bacteria are irradiated in said growth medium. In one embodiment, said NH3 formation is in said growth medium. In one embodiment, said irradiation results in a yield amount greater than 105 moles of NH3 per mole of bacteria cells. In one embodiment, said bacteria comprise a cell lysate. In one embodiment, said bacteria are live bacteria. In one embodiment, said live bacteria are replicating. In one embodiment, said CZS-QD has two ZnS monolayers. In one embodiment, said CZS-QD has a zwitterion L-cysteine (CYS) ligand cap. In one embodiment, said radiation source emits wavelengths selected from the group consisting of near-UV, visible, NIR radiation, Light Emitting Diode (LED) and solar radiation. In one embodiment, said illumination source emits 400 nm radiation. In one embodiment, said illumination source emits yellow (near-infrared (NIR)) radiation. In one embodiment, said growth medium further comprises L-ascorbic acid. In one embodiment, said method further provides a growth medium further comprising 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer.
In one embodiment, the present invention provides a composition comprising a Cupriavidus necator bacteria strain comprising a Cadmium sulfide (CdS) core and a zinc sulfide (ZnS) shell (CdS@ZnS) (CZS) core-shell quantum dot (QD), and a pBBRl-yfp expression plasmid. In one embodiment, said QD is CZS2.
In one embodiment, the present invention provides a composition comprising a Cupriavidus necator bacteria strain comprising a cadmium selenide (CdSe) core and a zinc sulfide (ZnS) shell (CdSe@ZnS) (CZSe) core-shell quantum dot (QD), a pBBRl-yfp expression plasmid. In one embodiment, said QD is CZSe3. In one embodiment, said bacteria contains Polyhydroxybutyrate (PHB) molecules in amounts greater than in said bacteria strain without said QD. In one embodiment, said bacteria contains Polyhydroxybutyrate (PHB) molecules above natural levels. In one embodiment, said PHB contains an amount greater than 103 moles of PHB per mole of bacteria cells. In one embodiment, said Cupriavidus necator further comprises a H16+pBBR1-PphaC-YFP plasmid expressing an enzyme. In one embodiment, said Cupriavidus necator further comprises a pBBRl-yfp plasmid expressing a PHB enzyme sequence. In one embodiment, said enzyme sequence is codon optimized for expression in Cupriavidus necator. In one embodiment, said QD has a zwitterion L-cysteine (CYS) cap. In one embodiment, said QD has two ZnS monolayers. In one embodiment, said bacteria comprises a cell lysate. In one embodiment, said bacteria are live bacteria. In one embodiment, said live bacteria are replicating. In one embodiment, said illumination comprises 400 nm radiation. In one embodiment, said illumination comprises orange radiation (near-infrared (NIR)). In one embodiment, said composition further comprises CO2/O2 at a 4:1 ratio.
In one embodiment, the present invention provides a composition comprising a Azobacteria vinelandii bacteria strain comprising a core-shell quantum dot (QD) and Polyhydroxybutyrate (PHB) molecules. In one embodiment, said Polyhydroxybutyrate (PHB) molecules are in amounts greater than in said bacteria strain without said QD. In one embodiment, said Polyhydroxybutyrate (PHB) molecules are in amounts above natural levels.
In one embodiment, the present invention provides a method of producing Polyhydroxybutyrate (PHB), comprising, a) providing, i) a bacteria strain comprising a molybdenum-iron nitrogenase (MFN) enzyme, wherein expression of said MFN enzyme results in Polyhydroxybutyrate (PHB) formation, a plurality of core-shell quantum dots (QDs), wherein said QD transmits electrons having energies in the range of the reduction potential of said MFN enzyme upon exposure to radiation that increases the activity of said MFN enzyme, ii) an illumination source capable of emitting radiation, and iii) at least one compound selected from the group consisting of CO2, H2O, O2 and N2, b) incubating said engineered bacteria in the presence of said at least one compound in the dark, and c) irradiating said engineered bacteria with said illumination source under conditions that produce Polyhydroxybutyrate (PHB). In one embodiment, said QD is a cadmium sulfide (CdS) core zinc sulfide (ZnS) shell QD. In one embodiment, said QD is CZS2. In one embodiment, said QD is a cadmium selenide (CdSe core zinc sulfide (ZnS) shell QD. In one embodiment, said QD is CZSe3. In one embodiment, said QD has two ZnS monolayers. In one embodiment, said QD has a zwitterion L-cysteine (CYS) ligand cap. In one embodiment, said production of PHB molecules is above natural levels. In one embodiment, said production of PHB molecules is an amount greater than in said bacteria strain without said QD. In one embodiment, said production of PHB molecules is an amount greater than in said bacteria strain without said irradiation. In one embodiment, the method further provides bacteria growth medium, wherein said bacteria contact said growth medium. In one embodiment, said bacteria are illuminated when in contact with said growth medium. In one embodiment, said PHB molecules are secreted into said growth medium. In one embodiment, said greater amount is up to 150% greater. In one embodiment, said greater amount is up to 100 mg of said PHB per gram of bacteria cell dry weight (CDW). In one embodiment, said greater yield is obtained within a twenty-four hour illumination time. In one embodiment, said bacteria strain is a Cupriavidus necator bacteria strain. In one embodiment, said Cupriavidus necator strain is DJ995. In one embodiment, said bacteria strain is an Azobacteria vinelandii bacteria strain. In one embodiment, the method further comprises an extraction of PHB molecules with sodium hypochlorite/chloroform mixture. In one embodiment, the method further comprises the step of precipitating said PHB molecules with methanol/water from said chloroform extract. In one embodiment, the method further comprises the step of coagulating said PHB molecules by drying. In one embodiment, the method further comprises rehydrating PHB molecules in glacial acetic acid solution. In one embodiment, the method further comprises casting a PHB thin film from said rehydrated PHB molecules. In one embodiment, said radiation source emits wavelengths selected from the group consisting of near-UV, visible, NIR radiation, Light Emitting Diode (LED) and solar radiation. In one embodiment, said radiation source emits 400 nm radiation. In one embodiment, said radiation source emits orange (near-infrared (NIR)) radiation.
In one embodiment, the present invention provides a Cupriavidus necator bacteria comprising a CdS@ZnS (CZS) core-shell quantum dot (QD) and alcohol molecules. In one embodiment, said alcohol is selected from the group consisting of isopropanol, butane diol, gasoline additives and gasoline substitutes. In one embodiment, said bacteria contain said alcohol molecules. In one embodiment, said alcohol molecules are at a level greater than for said bacteria without a QD. In one embodiment, said composition further comprises a bacteria growth medium. In one embodiment, said alcohol molecules. In one embodiment, said growth medium contains said alcohol molecules at a level greater than for said bacteria without a QD. In one embodiment, said Cupriavidus necator bacteria comprises a H16_IPAl-10_int sequence. In one embodiment, said accumulated amounts ranging from 106-108 moles of isopropanol per mole of cells. In one embodiment, said bacteria comprise a cell lysate. In one embodiment, said bacteria are live bacteria. In one embodiment, said live bacteria are replicating. In one embodiment, said CZS-QD has two ZnS monolayers. In one embodiment, said CZS-QD has a zwitterion cysteine (CYS) cap. In one embodiment, the method further comprises 400 nm radiation. In one embodiment, the method further comprises red (near-infrared (NIR)) irradiation. In one embodiment, the method further comprises an irradiation source emitting wavelengths selected from the group consisting of near-UV, visible, and NIR irradiation. In one embodiment, said strain is DJ995. In one embodiment, the method further comprises a CO2/O2 4:1 mixture.
In one embodiment, the present invention provides a composition comprising a live engineered Azobacteria vinelandii bacteria strain comprising a CdS@ZnS (CZS) core-shell quantum dot (QD) and alcohol molecules.
In one embodiment, the present invention provides a method of producing an alcohol molecule, comprising, a) providing, i) a Cupriavidus necator strain comprising an enzyme that produces an alcohol molecule and a quantum dot (QD), wherein said QD is a CdS@ZnS (CZS) core-shell quantum dot, that transmits electrons having energies in the range of the reduction potential of said enzyme upon exposure to radiation for boosting production of said molecule, ii) an illumination source capable of emitting radiation, and iii) compounds comprising CO2, H2O, O2 and N2, b) incubating said bacteria in the presence of said compounds in the dark, and c) illuminating said live engineered bacteria for illuminating said bacteria for producing alcohol molecules in a yield amount greater than in said bacteria strain without said QD. In one embodiment, said alcohol is selected from the group consisting of isopropanol, butane diol, gasoline additives and gasoline substitutes. In one embodiment, the method further provides a bacteria growth medium. In one embodiment, said bacteria contact said growth medium. In one embodiment, said bacteria are illuminated in contact with said growth medium. In one embodiment, said alcohol molecules are in said growth medium. In one embodiment, said illumination emits energies selected from the group consisting of near-UV, visible, NIR radiation, Light Emitting Diode (LED) and solar radiation. In one embodiment, said Cupriavidus necator bacteria comprises a H16_IPAl-10_int sequence as a chromosome integration.
In one embodiment, the present invention provides a composition comprising a live Cupriavidus necator bacteria strain comprising a CdS@ZnS (CZS) core-shell quantum dot, wherein said bacteria contains carbon dioxide molecules. In one embodiment, said bacteria contain said carbon dioxide molecules at a level greater than for said bacteria without a QD.
In one embodiment, the present invention provides a composition comprising a live Azobacteria vinelandii bacteria strain comprising a CdS@ZnS (CZS) core-shell quantum dot, wherein said bacteria contains carbon dioxide molecules.
In one embodiment, the present invention provides a method of carbon dioxide sequestration in a live bacteria strain, comprising, a) providing, i) a live bacteria comprising an enzyme that produces carbon dioxide (CO2) and a quantum dot that transmits electrons having energies in the range of the reduction potential of said enzyme upon exposure to radiation for boosting production of said biofuel molecule, ii) an illumination source capable of emitting radiation, iii) a CO2/O2 4:1 air mixture, iv) a fermentation solution, v) flushing said solution with said air mixture, and b) suspending said QD containing bacteria in said fermentation solution, c) flushing said fermentation solution containing QD containing bacteria with said air mixture, and d) irradiating said bacteria in said flushed fermentation solution under conditions such that said carbon dioxide is sequestered in said QD containing bacteria. In one embodiment, said flushing is at a rate of 0.5 standard liter per minute. In one embodiment, said flushing is for 15 minutes in duration before step d).
In one embodiment, the present invention provides a nanobiohybrid composition comprising a core-shell quantum dot (QD), wherein said core is selected from the group consisting of cadmium sulfide (CdS), cadmium selenide (CdSe), indium phosphide (InP), and copper zinc tin sulfide (Cu2ZnSnS4) (CZTS), wherein said shell is a two monolayer zinc sulfide (ZnS) shell, further comprising an enzyme. In one embodiment, said QD is attached to said enzyme. The composition further comprising a microorganism. In one embodiment, said microorganism is a bacterium. In one embodiment, said QD is inside of a microorganism, wherein said QD containing microorganism is a nanorg. In one embodiment, said enzyme is selected from the group consisting of a molybdenum-iron (MoFe) nitrogenase (MFN), MoFe-MFN subunits, Methyl Ketone pathway enzymes, 2,3-butanediol (BDO) pathway enzymes, isopropanol pathway enzymes, ethylene-forming enzyme (EFE), and PHB pathway enzymes. In one embodiment, said enzyme is molybdenum-iron (MoFe) nitrogenase (MFN). In one embodiment, said MFN is endogenous. The composition, wherein said MFN is heterologous. In one embodiment, said enzyme is MFN and said microorganism contains ammonia (NH3) molecules. In one embodiment, said enzyme is MFN and said microorganism contains hydrogen (H2) molecules. In one embodiment, said molecules are at levels higher than for said microorganisms that does not contain a QD. In one embodiment, said enzyme is one or more enzyme in the Methyl Ketone (MK) pathway selected from the group consisting of enzymes encoded by genes: ‘tesA, fadB, Mlut_11700, and fadM. In one embodiment, said enzyme is one or more enzyme encoded by pJM20. In one embodiment, said Methyl Ketone (MK) pathway enzymes are endogenous. In one embodiment, said Methyl Ketone (MK) pathway enzymes are heterologous. In one embodiment, said microorganism is a bacterium. In one embodiment, said bacterium is C. necator strain H16. In one embodiment, said bacterium comprises pJM20. In one embodiment, said enzyme is encoded by pJM20 and said microorganism is a C. necator strain H16. In one embodiment, said enzymes form a complete MK pathway in said microorganism, wherein said microorganism contains C11-C15 methyl ketone compounds. In one embodiment, said C11-C15 methyl ketone compounds are at levels higher than for said microorganisms that does not contain a QD. In one embodiment, said enzyme is one or more enzyme in the isopropyl alcohol (IPA) forming pathway is selected from the group of enzyme encoding genes consisting of bktB(β-ketothiolase, H16_A1445) derived from C. necator H16; ctfAB(Succinyl-CoA transferase, AJ000086) from H. pylori, adc(acetoacetate decarboxylase, CA_P0165) from C. acetobutylicum, and sadh(secondary alcohol dehydrogenase, AAA23199.2) from C. beijerinckii. In one embodiment, said enzyme is one or more enzyme encoded in the H16_IPA1-10_int operon. In one embodiment, said d microorganism further comprises a H16_IPA1-10_int operon. In one embodiment, said microorganism has a chromosome, and said chromosome contains said H16_IPA1-10_int operon. In one embodiment, said microorganism contains IPA molecules. In one embodiment, said IPA molecules are at levels higher than for said microorganisms that does not contain a QD. In one embodiment, said enzyme is one or more enzyme in the 2,3-butanediol (BDO) forming pathway selected from the group of enzyme encoding genes consisting of alsS(acetolactate synsthase, BSU36010) and alsD(acetolactate decarboxylase, BSU36000) from B. subtilis and sadh(secondary alcohol dehydrogenase, AAA23199.2) from C. beijerinckii. In one embodiment, said enzyme is one or more enzyme encoded in the H16_BO2-20_int operon. In one embodiment, said microorganism further comprises a H16_BO2-20_int operon. In one embodiment, said microorganism has a chromosome, and said chromosome contains said H16_BO2-20_int operon. In one embodiment, said microorganism is a C. necator H16 strain. In one embodiment, said enzymes form a complete 2,3-butanediol pathway in said microorganism, wherein said microorganism is a bacterium contains BDO molecules. In one embodiment, said BDO molecules are at levels higher than for said microorganisms that does not contain a QD. In one embodiment, said enzyme is an ethylene-forming enzyme (EFE). In one embodiment, said EFE is derived from a Pseudomonas syringae bacterium. In one embodiment, said enzyme is an ethylene-forming enzyme (EFE). In one embodiment, said microorganism is a bacterium that contains ethylene molecules. In one embodiment, said ethylene molecules are at levels higher than for said microorganisms that does not contain a QD. In one embodiment, said enzyme is a Polyhydroxybutyrate (PHB) pathway enzyme. In one embodiment, said enzyme is a Polyhydroxybutyrate (PHB) pathway enzyme selected from the group consisting of PhaA, PhaB, and the heterodimeric PHB synthase PhaEC. The composition, wherein said enzyme is encoded by a PHB operon (phaCAB). In one embodiment, said enzyme is encoded by pBBR1-yfp. In one embodiment, said enzyme is a Polyhydroxybutyrate (PHB) pathway enzyme, wherein said microorganism is a bacterium contains PHB molecules. In one embodiment, said enzyme is a Polyhydroxybutyrate (PHB) pathway enzyme, wherein said microorganism is a bacterium contains CO2 molecules. In one embodiment, said PHB molecules are at levels higher than for said microorganisms that does not contain a QD. In one embodiment, said CO2 molecules are at levels higher than for said microorganisms that does not contain a QD. In one embodiment, said enzyme is a formic acid (FA) pathway enzyme. In one embodiment, said enzyme is a formate dehydrogenase. The composition, wherein said enzyme is a formate dehydrogenase derived from Clostridium carboxidivorans. In one embodiment, said bacterium contains formic acid molecules. In one embodiment, said enzyme is a methanol pathway enzyme. In one embodiment, said bacterium is lacking a functional formic acid (FA) pathway enzyme. In one embodiment, said bacterium contains hydrogen (H2) molecules. In one embodiment, said bacterium contains methanol molecules. In one embodiment, said bacterium contains accumulated CO2. In one embodiment, said formic acid molecules are at levels higher than for said microorganisms that does not contain a QD. In one embodiment, said hydrogen (H2) molecules are at levels higher than for said microorganisms that does not contain a QD. In one embodiment, said methanol molecules are at levels higher than for said microorganisms that does not contain a QD. In one embodiment, said QD core-shell has a zwitterion L-cysteine (CYS) ligand cap. In one embodiment, said QD core has a ligand cap selected from the group consisting of a 3-mercaptopropionic acid (MPA) capping ligand and a cysteamine (CA) ligand cap. In one embodiment, said QD is selected from the group consisting of CZS1, CZS2, CZSe1, CZSe2, CZSe3, IPZS and CZTS. In one embodiment, said QD has an excitation energy from wavelengths ranging from ultraviolet to near-infrared energies.
In one embodiment, the present invention provides a method of using a nanobiohybrid comprising, providing, i) a core-shell quantum dot (QD), wherein said core is selected from the group consisting of cadmium sulfide (CdS), cadmium selenide (CdSe), indium phosphide (InP), and copper zinc tin sulfide (Cu2ZnSnS4) (CZTS), wherein said shell is a two monolayer zinc sulfide (ZnS) shell, ii) an enzyme for producing a product compound from a substrate, having a level of catalytic activity for said substrate, iii) a substrate for said enzyme, and iv) an illumination source capable of emitting radiation, and b) contacting said QD with said enzyme forming a nanobiohybrid, c) irradiating said nanobiohybrids in the presence of said substrate for emitting radiation which increases said catalytic activity of said enzyme for increasing production of a product compound. In one embodiment, said production of a product is greater than for said nanobiohybrid that is not irradiated. In one embodiment, said production of a product is greater than for said enzyme that is not contacting a QD. In one embodiment, said production ranges from 105-108 moles per mole of bacteria cells. In one embodiment, said nanohybrid is contained within a microorganism forming a nanorg. In one embodiment, the method further provides a fermentation media, and a step of contacting said nanorg with said fermentation media. In one embodiment, the method further comprises compounds selected from the group comprising CO2, H2O, O2 and N2. In one embodiment, said CdS QD absorbs radiation in the ultraviolet range. In one embodiment, said CdSe QDs absorbs radiation in the visible light range. In one embodiment, said InP and CZTS QDs absorbs radiation in the near-infrared (NIR) range. In one embodiment, said product compound is selected from the group consisting of ammonia (NH3) molecules; hydrogen (H2) molecules; Methyl Ketones (MK) compounds; C11-C15 methyl ketone compounds; isopropyl alcohol (IPA); 2,3-butanediol (BDO); ethylene molecules; Polyhydroxybutyrate (PHB) molecules; CO2 molecules; formic acid (FA) molecules; and methanol molecules. In one embodiment, said PHB is degradable.
In one embodiment, the present invention provides a composition comprising a metal nanoparticle and a molybdenum-iron nitrogenase (MFN/Mo—Fe) enzyme. In one embodiment, said metal nanoparticle is capped with a plurality of glutathione (GSH) capping ligands. In one embodiment, said MFN enzyme comprises a plurality of histidine tags. In one embodiment, said metal nanoparticle contacts said MFN enzyme. In one embodiment, said metal nanoparticle is attached to said histidine tags. In one embodiment, said metal nanoparticle is a gold (Au) nanoparticle. In one embodiment, said metal nanoparticle is a nanocluster (NC). In one embodiment, said metal nanocluster comprises a plurality of gold (Au) nanoparticles. In one embodiment, said gold nanocluster is selected from the group consisting of gold nanoparticles having 10-12 atoms (Au10-12), 15 atoms (Au15), 18 atoms (Au18), 22 atoms (Au22) and 25 atoms (Au25). In one embodiment, said metal nanoparticle has an excitation energy from wavelengths ranging from ultraviolet to near-infrared energies. In one embodiment, said metal nanoparticle has a range of emission energies after exposure to said excitation energies. In one embodiment, said bacterium expresses said molybdenum-iron nitrogenase (Mo—Fe) enzyme having a reduction potential energy range matching said metal nanoparticle emission energies. In one embodiment, said metal nanoparticle further comprises a bacterium.
In one embodiment, the present invention provides a composition comprising an Azobacteria vinelandii bacteria strain comprising a gold nanoparticle, and a molybdenum-iron nitrogenase (Mo—Fe) enzyme. In one embodiment, said gold nanoparticle is a nanocluster (NC). In one embodiment, said method further comprising ammonia (NH3) molecules. In one embodiment, said bacteria contains ammonia (NH3) molecules above natural levels. In one embodiment, said bacteria contains hydrogen (H2) molecules above natural levels. In one embodiment, said Mo—Fe enzyme is heterologous. In one embodiment, said Mo—Fe enzyme comprises a 7× histidine tag. In one embodiment, said bacteria comprise a cell lysate. In one embodiment, said bacteria are live bacteria. In one embodiment, said live bacteria are replicating. In one embodiment, said strain is DJ995. In one embodiment, said method further comprising CO2/O2 at a ratio of 4:1. In one embodiment, said composition further comprises a growth medium. In one embodiment, said growth medium comprises an accumulation of said NH3 above levels for said bacteria cells without a metal nanoparticle.
In one embodiment, the present invention provides a method of producing ammonia (NH3), comprising, a) providing, i) an A. vinelandii bacteria strain comprising a molybdenum-iron nitrogenase (MFN) enzyme, wherein expression of said MFN enzyme results in ammonia (NH3) formation, a plurality of gold nanoparticles, ii) an illumination source capable of emitting radiation, iii) at least one compound selected from the group consisting of CO2, H2O, O2 and N2, and b) incubating said engineered bacteria in the presence of said at least one compound in the dark, and c) irradiating said engineered bacteria with said illumination source under conditions that produce ammonia (NH3). In one embodiment, said gold nanoparticle is a nanocluster (NC). In one embodiment, said gold nanocluster is selected from the group consisting of gold nanoparticles having 22 atoms (Au22) and 25 atoms (Au25). In one embodiment, said production of NH3 molecules is above natural levels. In one embodiment, said production of NH3 is an amount greater than in said bacteria strain without said plurality of gold nanoparticles. In one embodiment, said production of NH3 is an amount greater than in said bacteria strain without said irradiating. In one embodiment, said ammonia (NH3) molecules are greater than 105 moles of NH3 per mole of bacteria cells. In one embodiment, said ammonia (NH3) molecules are greater than 105 up; to 108 moles of NH3 per mole of bacteria cells. In one embodiment, said hydrogen (H2) molecules are greater than 105 moles of NH3 per mole of bacteria cells. In one embodiment, said Azobacteria vinelandii strain is DJ995. In one embodiment, said MFN enzyme is heterologous. In one embodiment, said method further providing a bacterial growth medium. In one embodiment, said bacteria are irradiated in said growth medium. In one embodiment, said NH3 formation is in said growth medium. In one embodiment, said irradiation results in a yield amount greater than 105 moles of NH3 per mole of bacteria cells. In one embodiment, said bacteria comprise a cell lysate. In one embodiment, said bacteria are live bacteria. In one embodiment, said live bacteria are replicating. In one embodiment, said radiation source emits wavelengths selected from the group consisting of near-UV, visible, NIR radiation, Light Emitting Diode (LED) and solar radiation. In one embodiment, said illumination source emits 405 nm radiation. In one embodiment, said illumination source emits yellow (near-infrared (NIR)) radiation. In one embodiment, said growth medium further comprises L-ascorbic acid. In one embodiment, said growth medium further comprises 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer.
To facilitate the understanding of this invention, a number of terms are defined below. Terms defined herein have meanings as commonly understood by a person of ordinary skill in the areas relevant to the present invention. Terms such as “a”, “an” and “the” are not intended to refer to a singular entity but also plural entities and also includes the general class of which a specific example may be used for illustration. The terminology herein is used to describe specific embodiments of the invention, but their usage does not delimit the invention, except as outlined in the claims.
The term “about” or “approximately” as used herein, in the context of any of any assay measurements refers to +/−5% of a given measurement.
The term, “enzyme” refers to a protein that acts as a catalyst to bring about a specific biochemical reaction, e.g. converting a specific set of reactants (called substrates) into specific products. An enzyme may also regulate the rate at which chemical reactions proceed in living organisms.
The term, “reduction potential” or “redox potential” or “oxidation reduction potential” refers to a tendency to gain electrons, i.e. become “reduced.
The term, “fuel” refers to a substance, such as a material or chemical, that is used to provide heat or power, usually by being burned. A fuel may be used as a single material or chemical and also as part of a fuel mixture comprising more than one material and/or chemical.
The term, “biofuel” refers to a fuel derived directly from living matter.
The term, “biodiesel” refers to a fuel comprising long-chain alkyl (methyl, ethyl, or propyl) esters, for example.
The term, “alcohol” refers to a compound in which one or more hydrogen atom positions in an alkane has a hydroxyl (—OH) group instead, including but not limited to primary (1°) alcohols where the C—OH is attached to one other carbon (on the end), e.g. Methanol CH3OH (Methyl alcohol), ethanol CH3CH2OH (Ethyl alcohol), 1-Propanol CH3CH2CH2OH (Propyl alcohol), 1-Butanol CH3CH2CH2CH2OH (Butyl alcohol), etc.; secondary (2°) alcohols where the C—OH is attached to two other carbons, e.g. 2-Propanol (CH3)2CHOH (IPA, Isopropanol, Isopropyl alcohol); and tertiary (3°) alcohols where the C—OH is attached to three other carbons (saturated carbon atom), e.g. 2-methylpropan-2-ol C7H16O2, 2-methylbutan-2-ol CH3CH2C(CH3)2OH (tert-Amyl alcohol), butane diol, gasoline additives, gasoline substitutes, etc.
The term, “sequestration” refers to storage. As one example, sequestration of carbon refers to storage of carbon dioxide or other forms of carbon atoms within a microbe.
The term, “light” refers to electromagnetic radiation in a wavelength range including but not limited to infrared, visible, ultraviolet, etc.
The term, “illumination” refers to an action of supplying or brightening with light.
The term, “source” in reference to light and illumination refers to a natural light, e.g. sunlight; an artificial light source, such as fluorescent light, Light Emitting Diodes (LEDs), Surface Mounted Device LEDs, Chip on Board LEDs, etc.
An “illumination source” may also be referred to as a measurement of the total quantity of light emitted by the source expressed in lumens per unit of area.
The term, “purified” or “isolated”, as used herein, may refer to a peptide composition that has been subjected to treatment (i.e., for example, fractionation) to remove various other components, and which composition substantially retains its expressed biological activity. Where the term “substantially purified” is used, this designation will refer to a composition in which the protein or peptide forms the major component of the composition, such as constituting about 50%, about 60%, about 70%, about 80%, about 90%, about 95% or more of the composition (i.e., for example, weight/weight and/or weight/volume). The term “purified to homogeneity” is used to include compositions that have been purified to “apparent homogeneity” such that there is single protein species (i.e., for example, based upon SDS-PAGE or HPLC analysis). A purified composition is not intended to mean that all trace impurities have been removed.
As used herein, the term “substantially purified” refers to molecules, either nucleic or amino acid sequences, that are removed from their natural environment, isolated or separated, and are at least 60% free, preferably 75% free, and more preferably 90% free from other components with which they are naturally associated. An “isolated polynucleotide” is therefore a substantially purified polynucleotide.
A “variant” of an oligonucleotide or protein is defined as a nucleotide or amino acid sequence that differs from a wild type oligonucleotide or protein by having deletions, insertions and substitutions. These may be detected using a variety of methods (e.g., sequencing, hybridization assays etc.).
A “deletion” is defined as a change in either nucleotide or amino acid sequence in which one or more nucleotides or amino acid residues, respectively, are absent.
An “insertion” or “addition” is that change in a nucleotide or amino acid sequence which has resulted in the addition of one or more nucleotides or amino acid residues, respectively, as compared to, for example, a naturally occurring C. necator.
A “substitution” results from the replacement of one or more nucleotides or amino acids by different nucleotides or amino acids, respectively.
The term “derivative” as used herein, refers to any chemical modification of a nucleic acid or an amino acid. Illustrative of such modifications would be replacement of hydrogen by an alkyl, acyl, or amino group. For example, a nucleic acid derivative would encode a polypeptide which retains desired biological characteristics.
The term “transfection” or “transfected” refers to the introduction of foreign DNA into a cell.
As used herein, the terms “nucleic acid molecule encoding”, “DNA sequence encoding,” and “DNA encoding” refer to the order or sequence of deoxyribonucleotides along a strand of deoxyribonucleic acid. The order of these deoxyribonucleotides determines the order of amino acids along the polypeptide (protein) chain. The DNA sequence thus codes for the amino acid sequence.
As used herein, the term “structural gene” refers to a DNA sequence coding for RNA or a protein. In contrast, “regulatory genes” are structural genes that encode products which control the expression of other genes (e.g., transcription factors).
As used herein, the term “gene” means the deoxyribonucleotide sequences comprising the coding region of a structural gene and including sequences located adjacent to the coding region on both the 5′ and 3′ ends for a distance of about 1 kb on either end such that the gene corresponds to the length of the full-length mRNA. The sequences which are located 5′ of the coding region and which are present on the mRNA are referred to as 5′ non-translated sequences. The sequences which are located 3′ or downstream of the coding region and which are present on the mRNA are referred to as 3′ non-translated sequences. The term “gene” encompasses both cDNA and genomic forms of a gene. A genomic form or clone of a gene contains the coding region interrupted with non-coding sequences termed “introns” or “intervening regions” or “intervening sequences.” Introns are segments of a gene which are transcribed into heterogeneous nuclear RNA (hnRNA); introns may contain regulatory elements such as enhancers. Introns are removed or “spliced out” from the nuclear or primary transcript; introns therefore are absent in the messenger RNA (mRNA) transcript. The mRNA functions during translation to specify the sequence or order of amino acids in a nascent polypeptide.
In addition to containing introns, genomic forms of a gene may also include sequences located on both the 5′ and 3′ end of the sequences that are present on the RNA transcript. These sequences are referred to as “flanking” sequences or regions (these flanking sequences are located 5′ or 3′ to the non-translated sequences present on the mRNA transcript). The 5′ flanking region may contain regulatory sequences such as promoters and enhancers which control or influence the transcription of the gene. The 3′ flanking region may contain sequences which direct the termination of transcription, posttranscriptional cleavage and polyadenylation.
The patent or application file contains at least one drawing executed in color. Copies of this patent or patent application publication with color drawing(s) will be provided by the Office upon request and payment of the necessary fee.
The invention relates to a nano-biohybrid (e.g., a nanoorg) organism (e.g. a bacteria) comprising different core-shell quantum dots (QDs) or gold nanoparticle clusters, with excitation ranges having ultraviolet to near-infrared energies, coupled with targeted enzyme sites. When illuminated by light, these nanorgs drive the renewable production of biofuel molecules and chemicals using carbon-dioxide (CO2), water, and nitrogen (from air) as substrates. The disclosed nanorgs catalyze light-induced air-water-CO2 reduction with a high turnover number (TON) of approximately 106-108 (mols of product per mol of cells) to biofuels including, but not limited to, isopropanol (IPA), butane diol, gasoline additives, gasoline substitutes, 2,3-butanediol (BDO), C11-C15 methyl ketones (MKs), and hydrogen (H2); and chemicals such as formic acid (FA), ammonia (NH3), ethylene (C2H4), and degradable bioplastics, e.g. polyhydroxybutyrate (PHB). These nanorg cells function as nano-microbial factories powered by light.
In one embodiment, the present invention contemplates a nano-biohybrid organism (or nanorg) including, but not limited to: (1) chemical coupling via affinity binding and self-assembly, (2) the energetic coupling between optoelectronic states of artificial materials with a cellular process, and (3) the design of appropriate interfaces ensuring biocompatibility. Here it is shown that at least seven different core-shell quantum dots (QDs), with excitations ranging from ultraviolet to near-infrared energies, coupled with targeted enzyme sites in bacteria. When illuminated by the appropriate light energy, these QDs drive the renewable production of biofuels and chemicals using carbon-dioxide (CO2), water, and nitrogen (e.g. from air) as substrates. These disclosed QDs, as described herein, use zinc-rich shell facets for affinity attachment to cellular proteins. Cysteine zwitterion ligands enable uptake through the cell wall, facilitating cell survival. Together, such compositions provide nanorgs catalyzing light-induced air-water-CO2 reduction with a high turnover number (TON) of approximately 106-108 (mols of product per mol of cells) to biofuels, such as isopropanol (IPA), 2,3-butanediol (BDO), C11-C15 methyl ketones (MKs), and hydrogen (H2); and chemicals such as formic acid (FA), ammonia (NH3), ethylene (C2H4), and degradable bioplastics such as polyhydroxybutyrate (PHB). Therefore, these resting cells function as nano-microbial factories powered by light.
External control (e.g. wireless) over specific cellular function has been a long-standing objective in biology (Ref. 1). While such control, e.g. external regulation, can provide unprecedented insights into molecular biology, it can also form the basis for several new biotechnological techniques ranging from diagnosis and therapeutics to the generation of biofuels and bioproducts.
In one embodiment, the present invention contemplates a platform technology comprising nano-biohybrid organisms (or nanorgs) using semiconductor nanoparticles which can be designed for affinity binding to desired proteins by facile transport, uptake, and self-assembly, and matched to the electrochemical potential of the enzyme to trigger them externally using electromagnetic radiation, e.g. light, etc. As a specific application and to demonstrate broader applicability of methods described herein, the formation of such living nano-biohybrid organisms (or nanorgs) using nonlimiting strains of Azotobacter vinelandii and Cupriavidus necator bacteria strains is shown with a desired enzyme activation for targeted chemical generation using light in these non-photosynthetic microbes.
In one embodiment, the presently contemplated engineered strains of naturally occurring and synthetic bacteria can accomplish industrial reactions using chemical energy to generate electrons and reduce renewable chemical feedstocks like, for e.g., CO2, H2O, and air, and can be labeled as living factories (
Unsuccessful attempts have previously been made to combine the desired functionality of direct light-activation in cell-free extracts or purified enzymes for in vitro biocatalysis or bioelectrocatalysis.3-6 But these strategies have some limitations due to enzyme de-activation in the air or during chemical conversion, without an ability to regenerate the enzyme using the living cell. Other in vivo efforts have been targeted in specific strains of whole non-photosynthetic bacteria,7,8 but can limit their applicability due to specific tolerance of the bacteria to inorganic elements and a smaller range of chemicals that can be made. Further, both these processes lack the desired specificity of enzyme activation in living cells, and there is a need to develop a platform technology for such desired living nano-biohybrids for applications beyond solar energy conversion and catalysis, to new avenues in diagnosis and therapeutics. There has been an intensive search for a new method to combine multiple functionalities (e.g., light, voltage, or magnetic field stimulation) of inorganic nanomaterials with the versatility of metabolic networks in living cells, to simply “grow” such hybrid catalysts or convert existing living cells into nanorgs, by the simple addition of inorganic nanomaterials to the cellular medium/water.
The present invention demonstrates the potential of multifunctional living nanorgs by suspending normally non-photosynthetic bacteria in buffered water (in the absence of any sugar) and converting renewable feedstocks like, for e.g., air and CO2 directly into biofuels and specialty chemicals using these solar-powered factories. Using different core semiconductor nanocrystals or quantum dots (QDs) with tunable bandgap energies (such as CdS, CdSe, InP, and Cu2ZnSnS4), and an optimized two monolayer ZnS shell, the chemical binding affinity of zinc is utilized with either a histidine-tagged MoFe nitrogenase in A. vinelandii or Fe—S clusters in hydrogenases and quinones in C. necator, demonstrating the facile formation of nanorgs by self-assembly and simple addition of QDs in the media/buffered water.
These biocatalysts demonstrate high conversion yields to target products (10-100 mg of product/g of dry weight of cells/day) without the utilization of sugar as a source of energy, comparable to or even exceeding (up to −150%) native production levels. The potentially high quantum yields from such light-driven chemical generation (up to 10-20%) depends on the optimization between biocatalyst turnover, incident light-flux, and the amount of light absorbed. Together, these results demonstrate an unprecedented opportunity for development of these nanorgs as renewable sugar-free microbial factories for the production of biofuels and chemicals using sunlight in a scalable process, but also as a means of externally regulating the cellular function of living cells using electromagnetic-stimuli such as light, sound, or magnetic field. In some embodiments, the present invention contemplates nanorg microbial factories including, but not limited to, light-driven renewable biochemical synthesis using quantum dot-bacteria nano-biohybrids.
In one embodiment, the present invention contemplates multifunctional living nanorgs by suspending a range of different normally non-photosynthetic bacteria in buffered water (in the absence of any sugar) and converting renewable feedstocks like, for e.g., air and CO2 directly into biofuels and specialty chemicals using these solar-powered factories (
A. Core-Shell Quantum Dots.
The core-shell quantum dots (QDs) as contemplated herein were empirically designed and tested. In one embodiment, the disclosed QDs comprise zinc-rich shells for attachment to intracellular proteins of living bacteria. Advantages of the present QDs overcome previous beliefs in the art that incorporation of QDs in living bacteria may cause host bacteria to die. In some embodiments, the present invention contemplates coating the QD with a cysteine zwitterion ligand coating that facilitates uptake of the QD through the cell and improves cell survival.
At least seven different core-shell quantum dots (QDs), with excitations ranging from ultraviolet to near-infrared irradiation were chosen during the development of the present invention based upon the electron emission energy released by the irradiated/illuminated QD for matching a particular energy targeted to the energy level of a particular enzyme site in bacteria. When illuminated by light, these QDs were contemplated to act as a catalyst for increasing enzyme activity of the energy-matched enzyme.
In some preferred embodiments, the production of biofuels comprises using carbon-dioxide (CO2), water, and with nitrogen oxygen provided by air, as substrates. In some embodiments, the production of biofuels is renewable, from the aspect that the same engineered bacteria strains can be grown in large batches and used to produce the same biofuel.
C. necator
Azobacteria (A.) vinelandii DJ995, wild-type and genetically modified C. necator strains used for creating nanorgs for providing compounds including but not limited to: H2, NH3, FA, C2H4, IPA, BDO, MKs, and PHB production.
The generation (accumulation) of the corresponding products are shown in
A. Biodiesel: C11-C15 methyl ketones, e.g. C2H4, and C11.
Wild-type and genetically-engineered Cupriavidus necator (C. necator) strains for methyl-ketone (MK) production were produced by inserting plasmid pJM20 into C. necator by electroporation as detailed in Muller et. al., Plasmid pJM20 contains the entire MK pathway (‘tesA, fadB, Mlut_11700, and fadM) under the control of BAD promoter. PBAD (araBp; arabinose promoter) is regulated by the addition and absence of arabinose.
Bacteria were grown at 30° C., with shaking at 200 rpm, in the minimal salt media (MSM) supplied with 0.2% fructose and 0.2% glycerol, and the production of MK was realized via induction with 0.2% L-arabinose after 24 h inoculation (count as t=0).
For MK analysis (Muller), 5 ml cell suspension was mixed with 2 ml hexane followed by vigorous shaking for 30 min. After centrifugation (5,000 rpm, 5 min), the upper hexane layer was collected and concentrated (using N2 flow) to 100 1 pd of the hexane layer was injected for GC analysis (180° C. constant oven temperature).
For methyl (MK) production, after induction with L-arabinose (0.2% w/V), 1 ml culture was sampled for assay.
Cupriavidus necator (C. necator) using IPZS. C. necator strains, showing accumulation of C11 products over time, up to 0.015 mM per 80 hours.
B. Gasoline: 2,3-butanediol (BDO).
Wild-type and genetically-engineered C. necator strains were utilized using 2,3-butanediol (H16_BO2-20_int) chromosome integration. The strains were grown at 30° C., with shaking at 200 rpm, in the minimal salt media (MSM) supplied with 0.2% fructose and 0.2% glycerol.
C. Alcohols: Isopropanol (IPA).
Wild-type and genetically-engineered C. necator strains were utilized for IPA using (H16_IPAl-10_int) chromosome integration. The strains were grown at 30° C., with shaking at 200 rpm, in the minimal salt media (MSM) supplied with 0.2% fructose and 0.2% glycerol. Additionally, alcohols such as butane diol, were produced for use as gasoline additives and gasoline substitutes.
For IPA and BDO analysis, 1 ml cell suspension was lysed with an ultrasonic probe (3 cycles, 1 min each) followed by centrifuging at 15,000 rpm for 10 min. The supernatant was directly injected (1 fil) into the GC. For IPA, a constant oven temperature (140° C.) was used, whereas programmed temperature ramping (140° C. for 2 min, and 10° C./min ramping to 200° C.) was used for BDO analysis.
D. Ethylene (C2H4).
Wild-type and genetically-engineered C. necator strains were utilized for Ethylene (C2H4) (pBBRl-efe) plasmid. The strains were grown at 30° C., with shaking at 200 rpm, in the minimal salt media (MSM) supplied with 0.2% fructose and 0.2% glycerol.
E. Degradable bioplastics: PHB.
Wild-type and genetically-engineered C. necator strains were utilized for PHB (pBBRl-yfp) plasmid. The strains were grown at 30° C., with shaking at 200 rpm, in the minimal salt media (MSM) supplied with 0.2% fructose and 0.2% glycerol.
To demonstrate the efficiency of our sugar-free nanorg system in different light-excited redox reactions, we compared the chemical production yield in our nanorgs (with wild-type and C. necator strains expressing heterologous genes for the production of C2H4, IPA, BDO, or MKs) test with the yield in their natural growth (organolithotropic with fructose/glycerol or formate) conditions (
Even for the un-optimized C2H4, IPA, BDO, and MK producing strains, we observed a 10-50% biofuel production, compared to their native conditions. Most notably, when comparing PHB production (a native metabolite produced by C. necator), nanorgs exhibit up to 150% the PHB yield of wild-type cells. Further, the production was easily scaled up from several milliliter tests to liters by simply using a conventional bioreactor with LED panels (
F. Fertilizer: Ammonia (NH3) and Hydrogen.
Wild-type and genetically-engineered C. necator strains were utilized for ammonia (NH3) and hydrogen.
The decrease in quantum efficiency (or the saturation of NH3/H2 generation) with higher photon numbers was mainly due to accumulation of the products (functioning as inhibitors), and a replenishment of the reaction media (
G. Formic acid (FA).
Generation of formate from CO2 provides one embodiment of a method for sequestration of carbon from this greenhouse gas. In some embodiments, nanorgs of the present inventions are provided for producing formic acid and derivative chemicals from CO2. Enzymes include formate oxidases and formate dehydrogenases, e.g. derived from Clostridium carboxidivorans, for catalyzing the reduction of CO2. Wild-type and genetically-engineered C. necator strains were utilized for Formic acid (FA). In some embodiments, carbon sequestration refers to a process involved in carbon atom capture and storage, e.g. short-term or long-term, of atmospheric carbon, e.g. carbon dioxide (CO2). In some embodiments, stored carbon may be used for generating compounds as described herein.
With the replacement of LED lights to sunlight, the solar-powered, green product reactor for renewable generation of these targeted biofuels and bioproducts can also be realized commercially.
Cupriavidus
Cupriavidus
Cupriavidus
C. necator-
C. necator-
C. necator-
B. subtilis and
C. beijerinckii)
necator-pBBR1-
necator strain
necator strains, e.g. C.
necator H16
Cupriavidus (C.) necator
C. necator-chromosome
C. necator-Transformation
pylori, adc(acetoacetate
Choice of QD Core with Desirable Redox Potential.
One initial step towards the development of such living organisms is in vivo site-specific self-assembly, for chemically and energetically coupling the QDs and specific proteins within the synthetic bacteria. To accomplish the desired reaction driven by QD excitation, the reaction center and attachment site for the QD were identified. We designed these nanorgs by appropriately choosing the QD size and material ((Ref. 5, 6) (core-shells, since different materials were required for energetic alignment and chemical coupling/biocompatibility), QD surface charge and ligands (Reference Nos. 7, 8), and the desired site-specific attachment (9). The core QDs were selected using their size- and material-tunable conduction/valence bandedge positions (
Different sizes of CdS QDs (
The first step towards the development of such living organisms is in vivo site-specific self-assembly, for chemically and energetically coupling the QDs and specific proteins within the synthetic bacteria. While chemical conversion in biohybrid systems can also be achieved using the formation of intermediates using inorganic catalysis or electrochemistry, followed by utilization of these intermediates by bacteria,9-12 direct chemical conversion of inexpensive substrates using light in a bacteria as a platform is still challenging. Such a platform method, envisioned as non-genetically encoded (enzyme regeneration would lead to loss of photosensitization), self-assembled nano-biohybrid enzymes in nanorgs, could even allow selectively triggering cell function for diagnostic evaluation or therapy. To accomplish the desired reaction driven by QD excitation, the reaction center and attachment site for the QD were identified. We designed these nanorgs by appropriately choosing the QD size and material (Ref. 13, 14) (core-shells, since different materials were required for energetic alignment and chemical coupling/biocompatibility), QD surface charge and ligands, (Ref. 15, 16) and the desired site-specific attachment. (Ref. 17). The core QDs were selected using their size- and material-tunable conduction/valence bandedge positions (
Ensuring affinity attachment for chemical coupling and self-assembly between the QDs and the targeted enzymes inside the bacteria (
To maintain the desired energetic coupling of photoelectron production in CdS and the site-selective MFN binding with ZnS, CdS@ZnS core-shell quantum dots were designed to selectively trap the His-tagged MFN from the cell lysate (prepared from the A. vinelandii DJ995 bacteria) and conduct light-induced in situ redox reactions.
Optimizing the injection of photogenerated electrons from different core QDs to the enzyme active site, the core-shell QDs were synthesized using a layer-by-layer deposition technique (614), with precise control of the ZnS shell thickness (
The chemical attachment of QDs (capped with 3-mercaptopropionic acid or cysteine) to the proteins was demonstrated by Fourier-transformed infrared (FTIR) spectra of coupled QDs-CL after washing (to remove any unbound/weakly bound cellular component in CL,
Another aspect of the QD surface related to nanorg development was the ligand and the overall charge on QDs (9, 20-2317,28-31), which affected their biocompatibility, viability, and the uptake of designed QDs for intracellular self-assembly. Several attempts to combine the desired functionality of QDs with the synthetic versatility of the designed bacteria have relied on cell-free extraction of the enzymes and their coupling with QDs. These approaches showed limitations such as loss of enzyme activity or even deactivation (3, 10, 243-6,18,32) low enzyme concentrations, issues with scale-up, and low-activity and TON for chemical conversion. Using QDs capped with similar-sized ligands as well as different surface charge (negatively-charged MPA, positively-charged cysteamine (CA), and zwitterion cysteine (CYS)), we tested the viability of bacteria with QDs using three different methods. First, when monitoring cell growth by optical density, we observed that bacteria with CYS-capped QDs exhibited growth similar to no treatment, MPA-capped QDs impaired growth moderately, and CA-capped QDs strongly inhibited growth, especially at high concentrations (
Following the design and self-assembly of appropriate QD-bacteria biohybrid nanorgs (QDs with two monolayers thick ZnS shell, capped with cysteine ligand), we tested their ability to fix the energy of incident light-photons into specific chemical bonds using renewable and inexpensive substrates/feedstocks, like, for e.g., air, water, and CO2. Control experiments with either no QDs, light irradiation, or the cells, show no NH3 or C2H4 production in A. vinelandii and C. necator strains (
Further experiments also demonstrated the fixing of incident photon energy, as chemical fuels provide detailed measurements of light-intensity dependent chemical generation (
Following the design and self-assembly of appropriate QD-bacteria biohybrid nanorgs (QDs with two monolayers thick ZnS shell, capped with cysteine ligand), we tested their ability to fix the energy of incident light-photons into specific chemical bonds using renewable and inexpensive substrates/feedstocks, like, for e.g., air, water, and CO2. Control experiments with either no QDs, light irradiation, or the cells, show no NH3 or C2H4 production in A. vinelandii and C. necator strains (
After confirming the light-induced redox reaction by direct electron injection in nanorgs, we optimized the conditions further for chemical generation via enzyme activation, utilizing factors including bacterial cell density, irradiation intensity, sacrificial donor concentration (to improve) TOF of enzymes), and QD capping ligands and concentration (
These QDs form nanorgs with a high TON for NH3 (
The lower TON for the other nanorgs is mainly due to an unfavorable redox potential match or low absorptivity (especially for CZTS). We also demonstrated the flexibility of the nanorg platform by utilizing direct electron transfer to the living bacteria for different desired fuel generation. By coupling the QDs with a variety of bacterial strains (A. vinelandii DJ995, wild-type and genetically modified C. necator strains), we were able to create nanorgs for H2, NH3, FA, C2H4, IPA, BDO, MKs, and PHB production (
Optimizing the turnover number of light-driven chemical generation using nanorgs, we investigated the turnover frequency of the enzymes by comparing the photon flux for activation of the self-assembled QD-enzyme biohybrids, activated using light for converting photons to chemical bonds. First, we estimated the photon flux per biohybrid enzyme, by determining the light flux (approximately 1.6 mW/cm2) and using photon energy (approximately 2 eV) to obtain photon flux (5×1015 photons/sec/cm2). Using the optical density of the cells and the resulting nanorgs per unit area (approximately 108 nanorgs/cm2) and the estimated number of biohybrid enzymes (approximately 10,000 biohybrid enzymes/nanorg), we obtained the photon flux per biohybrid enzymes (approximately 5000 photons/biohybrid enzyme/sec). Comparing this to enzyme turnover (for MFN enzyme, 3000 nmol/mg/sec for 250 kDa enzyme≈(approximately equal to) 750/sec), we estimated an approximately 6-fold incident photons/enzyme turnover, thereby highlighting the mismatch between the number of biohybrid enzymes available for chemical generation in the nanorgs and the high incident flux of light. This would limit the maximum possible quantum efficiency to approximately 16-20%, further highlighting the high efficiency of enzyme activation using light (13% in our experiments), and the resulting biofuel and chemical generation using nanorgs. This could be optimized by utilizing synthetic biology tools to upregulate the enzymes, thereby coupling the photon flux to enzyme turnover, or reducing the photon flux. To study photon-energy related fuel production, we chose the nanorgs made from IPZS QDs (broader absorption spectrum) with the C2H4 producing C. necator strain, using the same test under different light sources (AM 1.5 (with 400 nm long pass filter), white, purple, blue, and green LED). All these light sources are able to excite the nanorgs for C2H4 production (
Most notably, when comparing PHB production (a native metabolite produced by C. necator), nanorgs exhibit up to 150% the PHB yield of wild-type cells. Further, the production was easily scaled up from several milliliter tests to liters by simply using a conventional bioreactor with LED panels (
Biochemical conversion can also be realized by using electricity. In electro-biochemical synthesis,34 the bacteria are immobilized on a modified electrode, which can inject electrons for downstream enzymatic reactions. The two processes: electro-biochemical synthesis and photo-biochemical synthesis, can be compared as photovoltaic driven-electrochemical synthesis and photocatalysis. The electro-biochemical approach circumvents several requirements of QDs-bacteria interactions, including QD uptake and low QD toxicity, making the design easier. However, electro-biochemical synthesis is limited to some special microbes that can exchange electrons with electrodes, and the requirements of immobilization and using redox mediators. Furthermore, there are no mechanisms in specifically targeting a desired enzyme or metabolic pathway for selective biochemical conversions. Therefore, photo-biochemical synthesis by using a designed QD-microbe biohybrid can offer additional advantages over other methods.
In conclusion, we have demonstrated a method for the formation of a living QD-bacterial nano-biohybrid nanorgs via the design of appropriate QDs and facile mixing, self-assembly, and affinity binding to the desired enzymes. Using a range of different light-absorbing QDs and targeted enzymes in different bacterial strains, we demonstrate the broad applicability of the proposed direct activation of the enzyme and the generation of biofuels and chemicals from non-photosynthetic microbes by simply suspending them in buffered water and bubbling air and/or CO2. Large turnover numbers and frequencies along with the high quantum efficiency for the direct conversion of light into chemicals were obtained for biofuel precursors and specialty chemicals including MKs, BDO, H2, IPA, NH3, FA, and PHB, demonstrating the potential and a possible application of the proposed method.
The biochemical conversion yields of the proposed method to simply utilize formed nanorgs, CO2 and water, in absence of sugar, even exceeded the natural yields in growing media (>150%), limited by the enzyme turnover. While maximum achievable quantum efficiency of light-to-chemical conversion can be 16-20% in the nanorgs, due to slow enzymatic conversion (˜1.3 msec) compared to absorbed light flux in the spontaneously self-assembled nano-biohybrids, high conversion efficiencies of light-activated chemical conversion (13%) highlights the potential of such simple platform for making self-assembled nanorgs and ability of electromagnetic enzyme activation. Further, such catalytic conversion can be optimized by upregulating the desired enzymes using tools in synthetic biology, and better matching the incident photon flux with the achievable turnover of the enzymes, for improved energy conversion.
This technique can easily be scaled up; be extended by improved screening for affinity binding to the proteins, expanding the scope of making nanorgs with other prokaryotes and eukaryotes; testing theories in molecular biology; and developing new diagnostic and therapeutic methods using other external stimuli, e.g. sound waves; magnetic field, etc.
A. A. vinelandii DJ995 Bacteria Growth and Cell Lysate Preparation.
In one embodiment, A. vinelandii DJ995 bacteria (wild type, with a histidine tagged MoFe nitrogenase on the C-terminus of the a-subunit), producing MoFe nitrogenase with 7× histidine tag on the C-terminal of the a-subunit was used. The cells were grown (31 s) in a nitrogen-free modified Burk media (500 ml for each batch, in a 2 L Erlenmeyer flask) with air bubbling (approximately 1 LPM) and shaking (approximately 300 rpm) for 24 hours (to an optical density of −1.5 at 600 nm). The resulting cells (dark brown color as shown in
1. Cell Lysate Activity Determination.
The cell lysate activity (Table S5, in the unit of nmol H2/NH3 per milliliter cell lysate per minute) in both enzymatic proton reduction (H2 generation) and the dinitrogen-water reduction was determined by a modified method reported by Dean et al, (Refs. 32S7 47).
2. Proton Reduction.
Enzymatic proton reduction was measured in a 25 ml septum sealed vial under a UPC argon atmosphere. One (1) ml from the reaction phase containing 25 mM TES (pH 7.4), 2.5 mM ATP, 5.0 mM MgCb, 30 mM creatine phosphate and 0.125 mg creatine phosphokinase (CPK) was fully degassed, sodium dithionite (solid) was then added to a final concentration of 20 mM. The headspace was charged with argon, and the vial was kept in a 30° C. water bath. The reaction was initiated by injection of 50 μl freshly thawed cell lysate (pellets stored in LN2) and terminated at 15 min by injecting 0.25 ml 2.5 M H2SO4. The headspace gas was analyzed by gas chromatography (SRI 8610C) with a molecular sieves 5 A column and a thermal conductivity detector (TCD) (sampling volume: 0.1 ml). The coefficient between peak area and amount of H2 (nmol) was determined by pure H2.
3. N2—H2O Reduction.
Enzymatic dinitrogen-water reduction was conducted using a method similar to the one used for proton reduction, with the replacement of the headspace gas by UHP N2 and the buffer by 35 mM HEPES (pH 7.4, to avoid interference in the NH3 assay). 0.25 ml 0.4 M EDTA (pH 8.0) was injected after 15 min to terminate the reaction. The amount of NH3 was determined using the o-phthalaldehyde fluorescence method (Refs. 3, 48). 25 μl from the reaction phase was added to 0.5 ml of assay reagent (pH=7.3) containing 20 mM o-phthalaldehyde, 0.2 M sodium phosphate, 5% ethanol and 3.4 mM mercaptoethanol. The mixture was maintained in the dark for at least 30 min and the emission was measured at 472 nm with a 410 nm excitation. NH3 calibration curves were obtained by using NH4CI in the same assay. Different concentration of NH4CI was used to obtain the calibration curve (
B. MoFe Nitrogenase (MFN) Purification.
The MFN protein from A. vinelandii DJ995 bacteria has a 7× histidine tag on the C-terminus of the a-subunit, allowing it to be purified using immobilized metal affinity chromatography (IMAC) (10, 25). The zinc ion was selected as the binding metal due to the use of sodium dithionite (DTT) reducing agent in the buffer. The column (Hitrap IMAC FF 1 ml, GE Healthcare) was charged with zinc (0.1 M ZnSO4/O and equilibrated with fully degassed equivalent buffer (with 2 mM DTT, pH 7.9). The cell lysate was loaded on to the column and washed with equivalent buffer and washing buffer (25 mM Tris-HCl, 0.5 M NaCl, 20 or 0.2? mM imidazole (PMSF) and 2 mM DTT, pH 7.9) to remove non-specific proteins. The His-tagged MFN protein was eluted using the elution buffer (25 mM Tris-HCl, 0.5 M NaCl, 250 mM imidazole and 2 mM DTT, pH 7.9). The elution (dark brown color) was stored as small pellets in LN2, for future use. Protein purity and concentration were determined using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and Bradford assay, respectively.
C. C. necator Culture.
Wild-type and genetically-engineered C. necator strainsS1,S8 were utilized for PHB (e.g. H16+pBBR1-PphaC-YFP) plasmid, (pBBRl-yfp) plasmid, etc., ethylene (pBBRl-efe) plasmid, isopropanol (H16_IPAl-10_int) chromosome integration, and 2,3-butanediol (H16_BO2-20_int) chromosome integration. The strains were grown at 30° C., with shaking at 200 rpm, in the minimal salt media (MSM) supplied with 0.2% fructose and 0.2% glycerol.
Strains for methyl-ketone (MK) production was grown in MSM supplied with 2% fructose, and the production of MK was realized via induction with 0.2% L-arabinose after 24 h inoculation (count as t=0).
D. Construction of C. necator Plasmids and Strains.
Plasmids were constructed in the pBBR1-MCS2 backbone. E. coli strains were propagated at 37° C. in lysogeny broth (LB). Where necessary, medium was solidified with 1.5% (wt/vol) agar and supplemented with 50 μg/ml kanamycin. The promoter from the phaC gene in C. necator was used to drive transcription of YFP (control), Ethylene-forming enzyme (efe, from Pseudomonas syringae). Ref S9. Conjugation was performed by transformation of plasmids into the mobilizing strain S-17 E. coli followed by incubation of the transformed cells overnight with C. necator on LB, followed by selection on LB+15 μl gentamicin (to select against S-17 cells) and 300 μl kanamycin. Ref. S10. Transformation of plasmids was verified by colony PCR and/or plasmid extraction and restriction digest validation.
E. Analysis of the Products Generated from C. necator.
For PHB, isopropanol (IPA), and 2,3-butanediol (BDO) production, the culture was sampled (1 ml) at a specific time point for analysis. For methyl (MK) production, after induction with L-arabinose (0.2% w/V), 1 ml culture was sampled for assay. C2H4 production was conducted in a septum-sealed vial, with 5 ml liquid phase containing OD600=1.0 cells in MSM media (with 40 mM formic acid). The headspace (25 ml) was sampled at a specific time point.
For PHB analysis (Ref. 33), 1 ml culture was centrifuged, and the pellet (washed once with MSM media) was mixed with 0.4 ml 3:7 HCkMeOH and 0.6 ml 1,2-dichloroethane (DCE) and incubated at 100° C. for two hours with gentle shaking (every 15 min). After cooling to room temperature, 0.3 ml DI water was added with vigorous shaking and the bottom layer (DCE layer) was injected into the GC (1 μl, oven temperature: 140° C.) for analysis.
For IPA and BDO analysis, 1 ml cell suspension was lysed with an ultrasonic probe (3 cycles, 1 min each) followed by centrifuging at 15,000 rpm for 10 min. The supernatant was directly injected (1 μl) into the GC. For IPA, a constant oven temperature (140° C.) was used, whereas programmed temperature ramping (140° C. for 2 min, and 10° C./min ramping to 200° C.) was used for BDO analysis.
For MK analysis (Ref 33), 5 ml cell suspension was mixed with 2 ml hexane followed by vigorous shaking for 30 min. After centrifugation (5,000 rpm, 5 min), the upper hexane layer was collected and concentrated (using N2 flow) to 100 μl. One (1) μl of the hexane layer was injected for GC analysis (180° C. constant oven temperature).
The generation (accumulation) of the corresponding products, together with the bacteria cell optical density (at 600 nm) are shown in
A. UV-VIS Determination of QDs-Protein Binding.
Further evidence supporting QD-attachment is provided by the UV-VIS measurements of solutions containing CL and a 500 nM concentration of CZSe3. Samples were incubated at room temperature for 30 min in 35 mM HEPES buffer prior to performing the following experiment.
Initially, strong PL and absorbance of the QDs in
B. Fourier-Transformed Infrared Spectroscopy (FTIR).
Thermo Nicolet 6700 FTIR instrument was used for infrared spectroscopy measurements, using a germanium attenuated total reflectance (ATR) accessory. QDs-cell lysate, QDs, and cell lysate samples were prepared for the test. Cell lysate (prepared from A. vinelandii DJ995 or C. necator pBBRl-efe) in Tris buffer was desalted (using 10 kDa centrifugal device) and washed with DI water twice to remove interfering Tris components. QDs-cell lysate complex was prepared by mixing the cell lysate with MPA-coated CZSe3 and incubated at room temperature for 30 min, followed by centrifugation at 15,000 rpm, the resulting pellets were washed twice with DI (deionized) water. No precipitate was observed when cell lysate or MPA-coated QDs suspension (in pH 11 water) was centrifuged at 15,000 rpm up to 30 min. The pellets formed in the mixture are due to the formation of QDs-protein complex. Samples were drop-casted on clean glass slides followed by air-drying.
FTIR was used to show proteins coupled to the QDs. The QDs are able to trap the proteins from the cell lysate, as shown by the similarity of the QDs-CL and CL spectra (
C. SDS-PAGE Protein Electrophoresis: Determination of Protein Bound to CdS or ZnS.
To evaluate the selectivity of CdS and ZnS in protein binding, cellular protein trapped by their corresponding particles were analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE).
1. Selective Protein Binding to CdS or ZnS.
CdS and ZnS particles were synthesized using the reaction of 0.1 M Cd2+ (CdCl2) or Zn2+ (ZnSO4) with 0.1 M S2+ (Na2S). The particles were washed with water (twice) and equivalent buffer (twice) and suspended in equivalent buffer. About 10 mg CdS or ZnS were charged with 1 ml cell lysate and incubated at 4° C. for 1 hour. The mixture was then centrifuged and washed with equivalent buffer (0.3 ml, twice) and wash buffer (0.3 ml) and re-suspended in 0.1 ml equivalent buffer.
2. Determination of Protein Bound to CdS or ZnS.
The proteins bound to CdS or ZnS were determined using SDS-PAGE. The CdS and ZnS particles with trapped (bound) proteins were boiled with SDS sample buffer and centrifuged at 10,000 rpm to remove the particles. The samples were loaded on the 12% SDS-PAGE gel, and the electrophoresis was run using constant voltage (200 V), followed by staining with Coomassie G250 to show the protein bands (
Selective His-tagged protein binding can be clearly seen with ZnS particles, showing an almost single protein band (
The A. vinelandii MoFe protein has two alpha (MoFe co-factor) and two beta (P-cluster) subunits, with molecular weights of 55.3 (plus approximately 1 for 7× histidine tag) and 59.5 kDa, respectively.
D. Agarose Gel Electrophoresis (AGE).
Agarose Gel Electrophoresis (AGE) was performed to prove the selective coupling between the ZnS-coated QDs and His-tagged MFN. Different proteins in the cell lysate are separated based on their molecular weight and charge, and the coupling of proteins to the QDs could show a change in the electrophoretic migration pattern. To use visible QDs, we used MPA-capped CdSe@ZnS QDs (CZSe1 and CZSe2 with peak emission at 540 and 570 nm, two monolayer ZnS coating) instead of non-fluorescent CdS@ZnS QDs for fluorescent detection of the QDs. The fresh-thawed cell lysate was mixed with QDs and incubated at room temperature for half an hour, followed by loading on 1% agarose gel. The electrophoresis was performed under constant voltage (200 V) mode for 1 hour. The gel was then gently rinsed with DI water and imaged using Gel Doc EZ imager on a UV tray (Bio-Rad), with excitation at 470 nm and emission at 570 nm, (
Fluorescence images clearly shows the existence and the position of QDs in the gel (
E. Inductively-Coupled Plasma Mass Spectroscopy (ICP-MS).
ICP-MS was used to determine the level of cadmium to quantify the efficiency and selectivity of QDs-protein coupling. The CZSe2-cell lysate lane (
F. QDs-MFN Biohybrid for Light-Induced Proton Reduction: CdX:Nitrogenase Biohybrid Photocatalytic Proton Reduction.
MPA-capped CdS or CdSe QDs were anaerobically mixed with the purified nitrogenase. The mixture was incubated at room temperature for about 10 min and diluted with fully degassed 100 mM L-ascorbic acid (pH=7.4). The mixture (with 200 nM QDs and 66 nM nitrogenase) was anaerobically transferred to several argon-purged 2 ml GC vials (with a small magnetic stirrer, 0.3 ml liquid volume). Light-induced proton reduction was performed by irradiating the system with a 400 nm LED panel at 1.6 mW/cm2. Headspace gas was sampled after 30 min irradiation. The net H2 turnover number (TON) are shown in
G. Light-Induced Redox Reaction with QDs-CL Mixture.
Here, the QDs refer to CZSI with nominal 0 to approximately 3 monolayer ZnS shells, synthesized from 3.55 nm CdS QDs, and capped with MPA ligands.
Light-induced proton reduction reaction was taken in a 2 ml vial under stirring, with a total reaction volume of 0.3 ml. The reaction phase contained 200 nM QDs, 100 mM ascorbic acid (pH 7.4) was vacuum-degassed and charged with argon (approximately 1.7 ml headspace). Anaerobically thawed cell lysate (15 μl) was swiftly injected into the vial with an air-tight syringe. The mixture was incubated at 30° C. for 10 min, followed by irradiating using a 400 nm LED panel (with −1.6 mW/cm2 at reaction site) for 30 min. The headspace gas was analyzed by gas chromatography (0.1 ml sampling) using the above-mentioned method, and as described herein.
The light-induced N2—H2O reduction was done in a similar condition, with the replacement of ascorbic acid by 300 mM HEPES and the headspace gas by UHP grade N2. H2 and NH3 were analyzed using the methods mentioned above, and as described herein, (
To evaluate how the selective binding of the His-tagged MFN to the ZnS-coated QDs would effect H2 production, the reaction was performed in media with the addition of 250 mM imidazole and with a higher acidity (pH 5.9). In both cases, the selective binding was interrupted.
The total amount of H2 produced from the QDs-cell lysate systems (xQDs-CL) was compared to the QDs (xQDs) (x=0-3 indicating the number of ZnS shells) systems. The OQDs-CL shows (
For the light-induced dinitrogen-water reduction with QDs:cell lysate biohybrids, both H2 and NH3 were generated (
In this section, and as described herein, cellular uptake (elemental analysis and confocal microscopy) and a series of viability tests (cell growth assay, resazurin dye assay, CFU assay) were performed to study the interactions between the QDs (CZSI with nominal 2ML ZnS shell, if not specified) and the bacteria cells (A. vinelandii and C. necator). Nitrogen-free Burk media or photocatalytic media (35 mM HEPES and 5 to approximately 25 mM L-ascorbic acid, pH=7.4) were used for A. vinelandii test, and minimal salt media (MSM) was used for C. necator test.
A. Cellular QDs Uptake Assay.
A. vinelandii DJ995 cells were collected at mid-log phase (OD600 approximately 1.0) and washed twice with ASC5 media (35 mM HEPES, 5 mM L-ascorbic acid, pH=7.4). Mixture with 200 nM MPA, CYS, or CA-coated QDs and OD6oo approximately 1.0 bacteria cells in ASC5 media (total volume 200 μl) were incubated at 30° C. for 30 min. The mixture was then centrifuged at 6,000 rpm and the cell pellets were quickly washed with ASC5 media (0.5 ml, twice) and finally re-suspended in 0.5 ml ASC5 media. The amount of Cd (in ppb) is measured by ICP-MS. The Cd level in the QDs suspension (1, 10, 100, 1000 nM) was also measured. A calibration curve was used to correlate the QDs concentration (in nM) to the Cd level (in ppb).
The QDs uptake (
QDs uptake (%)=ICP−MS measured QDs concentration×500×100%
B. Laser-Scanning Confocal Microscopy.
A. vinelandii cells were taken from growth media and centrifuged at 4,000 rpm for 2 min, washed twice with 25 mM HEPES buffer at pH 7.4, and resuspended in the same HEPES buffer to an OD of 0.1. MPA-capped CZSe were added to cells in HEPES buffer for a final concentration of 250 nM. The QDs-cell mixture was vortexed briefly and placed in a shaker at 37° C. and 225 rpm for one hour. The mixture was then centrifuged, washed with HEPES buffer three times, resuspended to an OD of 0.1, and deposited into a well-plate treated with poly-L-lysine. Samples were incubated at room temperature for 30 min, after which the wells were rinsed twice with DI water. 0.1 mL 4% paraformaldehyde solution was added to each well and allowed to incubate for 10 min at room temperature. After fixation, wells were rinsed with DI water twice and evacuated of liquid prior to imaging. Samples were imaged using Nikon AIR laser scanning confocal microscope with immersion oil and a 100× oil objective. Z-scan images were taken using a step-size of 0.25 m with an x-y resolution of 200 nm.
Confocal images were recorded (
C. Photocatalytic Reaction for Nanoparticle-Cell Lysate (NP-CL) Biohybrid.
Photocatalytic proton reduction reaction was taken in a 2 ml vial under stirring, with a total reaction volume of 0.3 ml. The reaction phase contained 200 nM nanoparticles, 100 mM ascorbic acid (pH 7.4) was vacuum-degassed and charged with argon (−1.7 ml headspace). Anaerobically thawed cell lysate (15 ul) was swiftly injected into the vial with an air-tight syringe. The mixture was incubated at 30° C. for about 5 min, followed by irradiating using a 400 nm LED panel (with about 1.6 mW/cm2 at reaction site) for 30 min. The headspace gas was analyzed by gas chromatography (0.1 ml sampling) using the method mentioned above. Reaction media with the addition of 250 mM imidazole and with higher acidity (pH=5.9) were also used.
Photocatalytic N2 reduction was taken in a similar condition, with the replacement of ascorbic acid by 300 mM HEPES and the headspace gas by UHP grade N2. H2 and NH3 were analyzed using the methods mentioned herein.
The total amount of hydrogen produced from nanoparticle-cell lysate systems (xNP-CL) was compared with the nanoparticle- (xNP) systems (x=0˜3 indicating the number of ZnS shells. The ONP-CL shows (
In the case of dinitrogen reduction with MoFe nitrogenase, both H2 and NH3 were generated (
D. Cell Growth Curve Measurement.
For the following test using living cells, if not specified, the nanoparticles refer to CdS@ZnS nanoparticles with nominal two-monolayer ZnS shell (CZS). The photocatalytic media refers to 35 mM HEPES buffer (pH=7.4) with 5, 10 or 25 mM L-ascorbic acid (ASC5, ASC10, and ASC25, respectively) as sacrificial hole quencher.
Cell growth measurement was performed in either Burk media or photocatalytic media (PCM), with a variation of QDs (nanoparticle) capping ligands (MPA, CYS, and CA) and concentration (50, 100, 200, 500, 750, and 1000 nM). A. vinelandii DJ995 bacteria grown in nitrogen-free Burk media were harvested at OD600 approximately 1.0 (mid-log phase) and washed twice and re-suspended in the Burk media or photocatalytic media. The cell growth curve (by measuring the optical density at 590 nm) was taken in the 96 well microplate (30° C., vigorous shaking) and monitored using a microplate reader (TECAN GENios) controlled by Megellan 7.2 software. Cells treated with QDs in photocatalytic media followed by growing in nitrogen-free Burk media were also tested to evaluate their viability after QDs treatment. The cells were first incubated in photocatalytic media (with QDs) for two hours (either in the dark or under 1.6 mW/cm2, 400 nm irradiation), followed by washing and re-suspending in Burk media for the growth. For all these cell growth measurements, the initial cell OD600 is 0.1.
The inhibition of cell growth indicates the QDs toxicity, which is strongly ligand-dependent (
No cell growth was observed (
Cell treatment in the dark revealed the capping ligand-dependent cell viability. While no remarkable cell viability loss was seen with MPA or CYS-capped QDs, a decrease of cell viability was observed with CA-coated QDs at higher concentration (small decrease at 500 nM and a complete cease of cell growth at 750 or 1000 nM). This indicates that dark cytotoxicity is very low for MPA or CYS-capped QDs, where the nanoparticle surface is negatively charged or has zwitterion characters, respectively. On the other hand, CA-capped QDs with positive surface non-selectively bind to all cell components, showing high cell toxicity.
The photo-toxicity of these QDs is similar to their dark toxicity. While CYS-capped QDs show no loss of cell viability up to 1000 nM, some cell viability loss was observed with a high concentration (750 and 1000 nM) MPA-capped QDs treatment. And almost complete ceasing of cell growth in higher concentration CA-capped QDs demonstrates their high toxicity.
D. Resazurin Dye Cell Viability Assay.
Cell viability assay was performed in a 96 well microplate using resazurin dye as an indicator. The bacteria cells (OD600 approximately 1.0) were for two hours treated QDs in the dark or under light (1.6 mW/cm2, 400 nm irradiation) irradiation, followed by washing (twice) and re-suspended in the nitrogen-free Burk media (OD600=1.0), as mentioned in the cell growth curve measurements. Resazurin was added to a final concentration of 0.1 mg/ml, and the fluorescence (excited at 485 nm) was measured at 620 nm using the microplate reader.
As shown in
For CA-capped QDs, cell toxicity was initially seen at 500 nM, and complete loss of cell viability was observed with higher concentrations (750 and 1000 nM). Compared with cell treatment in different media charged with or without CYS-coated nanoparticles (500 nM), a very small decrease of cell viability is shown. Furthermore, cell treatment (without nanoparticles) in different media shows no statistical loss of cell viability with ASC5 and ASC10 compared to cells without treatment (directly growth in Burk media). With L-ascorbic acid at higher concentration (25 mM), a small inhibition effect was observed.
Conclusions obtained from cells treated with QDs (nanoparticles) under light irradiation (
E. Colony forming unit (CFU) assay.
CFU assay was also used for evaluating the cell viability. Cell cultures were collected at OD600=1.0 from nitrogen-free Burk media and washed twice with ASC5 media. Mixtures with OD600=1.0 bacteria cell, 500 nM MPA, CYS or CA-capped QDs in ASC5 media were incubated at 30° C. for 2 hours in the dark, followed by washing (twice) and re-suspending in the same amount of nitrogen-free Burk media. 10 ul of the suspension with different cell OD600s (1, 10−2, 10−4, 10−6, 10−8) was inoculated on the B-plate (nitrogen-free Burk media with agar, in a squared petri dish) and incubated under 30° C. The CFU was counted by naked eyes (
Similar to the cell growth and resazurin cell viability test mentioned herein (
F. In-Vivo Photocatalytic Ammonia and Hydrogen Generation Test.
In vivo photocatalytic reactions were conducted in either 96 well microplates or small test tubes and tested either in the air or pure dinitrogen atmosphere. The mixtures basically contain Azotobacter vinelandii DJ995 cells, nanoparticles, and photocatalytic media and were incubated at 30° C. for 30 min and 150 mixture was added to the wells and a LED panel with 400 nm emission was used to irradiate the system through the cover, in a top-down mode. Ammonia production was determined using fluorescence assay described above, with the same mixture without irradiation (dark) as a baseline. To optimize the condition for ammonia yield, variations of cell optical density (ODeoo), capping ligands of the nanoparticles, nanoparticle concentration, and irradiation intensity were used.
First, 200 nM MPA-coated nanoparticles were used in ASC5 (5 mM L-ascorbic acid, 35 mM HEPES, pH 7.4) and irradiated with 400 nm light at 1.6 mW/cm2 for 1 hour. The Azotobacter vinelandii DJ995 culture from the Burk media was centrifuged at 6000 rpm and washed twice with ASC5. The cells were added to the above suspension with final OD600 from 0.1 to 1.0. The net ammonia production is shown in
The ammonia production increases with cell optical density, but not linearly. As from Azotobacter vinelandii DJ995, OD6oo=1.0 is at the mid-log phase of its growth and cells will start lysing at higher density. Therefore, we will use OD6oo=1.0 for our following optimization.
With fixed cell optical density (OD6oo=1.0), the nanoparticles with different capping ligands (MPA, CYS, CA) and concentrations were used. The ammonia generation is presented in
Control experiments with the removal of some components (cells or nanoparticles) from the mixture were also taken. As shown in
Irradiation intensity-dependent ammonia yield is also measured, with 500 nM CYS-coated nanoparticles and ODeoo=1-0 bacteria cells. Light intensity at reaction site from 0.16 to 2.42 mW/cm2 was used in this assay. With low irradiation intensity, the ammonia yield is low (
With fixed bacteria, cell optical density (OD600=1.0) and nanoparticle concentration (200 and 500 nM), photocatalytic dinitrogen reduction was taken in the air or pure dinitrogen atmosphere. A small test tube with 150 1 mixture was sealed with a septum and for replacing air with pure dinitrogen gas, the headspace air was vacuumed and recharged with UHP grade N2 using a syringe needle connected to the Schlenk line. The vacuum degassing and N2 recharging were repeated for three cycles to ensure low 02 level in the reaction system. As shown in
With the above optimization, time-dependent ammonia production was measured using photocatalytic ASC5, ASC10, ASC25 (35 mM HEPES with 5, 10, 25 mM L-ascorbic acid, pH=7.4) and Burk media. The reaction mixture was scaled up from 150 1 to 1 ml to allow multiple sampling. The photocatalytic test was taken in a small test tube covered with aluminum foil and magnetically stirred to ensure enough air supply. 25 microliter reaction phase was sampled at certain time point for ammonia assay. As shown in
The saturation of ammonia production could be due to depletion of the reducing agent (L-ascorbic acid or sucrose in photocatalytic or Burk media) or an increase of ammonia (inhibitor for MoFe nitrogenase) level in the reaction phase. To prove this assumption, ammonia was removed by separation the cells with centrifugation and replace the reaction phase with new media with nanoparticles every 1.5 hours. As shown from
We tested the cell-nanoparticle system in photocatalytic hydrogen production in the air atmosphere. The photocatalytic reaction (1 ml total volume) was performed in a small test tube as described above, with a rubber septum to retain the gas phase used for hydrogen quantification with gas chromatography. Headspace gas (total volume: 7 ml) was sampled at the certain time point and 0.1 ml gas was injected for hydrogen detection. Reaction phase contains 500 nM CYS-coated nanoparticles and OD6oo=LO bacteria cells in photocatalytic media (ASC5, ASC10, and ASC25). The result is presented in
The turnover number of the ASC5-CYS500 system for ammonia and hydrogen production was calculated, taking the number of cells (4.5×108/ml×1 ml=4.5×108 at ODgoo=1.0) into account. The result is presented in
A. Experiment and Control Groups.
The QDs-living bacteria nano-biohybrids were formed by mixing the bacteria cells with QDs in the suitable media followed by incubating at room temperature for about half an hour. Either open-air or septum-sealed reactors were used for photocatalytic air-water or CO2-water reduction tests. Typically, air-water reduction (NH3 production) using A. vinelandii was performed in either a 96 well microplate (150 ul total reaction volume, for end-point assay) or a small test tube (1 ml total reaction volume, for kinetics study). CO2-water reduction using A. vinelandii or C. necator was performed in either a 2 ml GC vial (0.3 ml total reaction volume, for end-point assay) or 30 ml septum-sealed vial (5 ml total reaction volume, for kinetics study). Experiment group for light-induced air-water reduction includes bacteria cells (OD600=1.0), 500 nM CZS QDs (CYS-capped), photocatalytic media (HEPES+L-ascorbic acid), air atmosphere, 1.6 mW/cm2 400 nm LED irradiation. For CO2-water reduction, the headspace was purged with CO2 for 15 min. Gentle agitation was applied to facilitate gas-liquid phase contact. Control groups with the removal of one or more components were used to prove the product generation from light-activated nanorgs instead of from media or contamination.
1. Plasmid Construction.
Primers were designed to amplify the efe gene with flanking Ndel and BamHI sites at the 5′ and 3′ ends respectively. The efe gene from Pseudomonas syringae pv. phaseolicola was synthesized by Eurofins (e.g. Eurofins Genomics LLC, Louisville Ky. 40299, USA) and was used as a template for PCR amplification utilizing exemplary primers EFEPF ′5 TTT CCC CAT ATG ATG ACC AAC CTA CAG ACT TT 3′ and EFEPR 5′ GGG AAA GGA TCC TCA TGA GCC TGT CGC GCG GG 3′. Both the efeP and YFP plasmids were constructed in the pBBR1-MCS2 backbone. The promoter from the phaC gene in C. necator was used to drive transcription of YFP (control), and the ethylene-forming enzyme (efe, from Pseudomonas syringae). The pBBRl vector was digested with Ndel and BamHI and the efe gene was ligated in to the vector downstream of the Pphac promoter, resulting in the construction of pBBRl-efe. Transformant colonies were selected on Kanamycin (50 mg/ml). Growing transformants were screened via colony PCR utilizing EFEPF and EFEPR Transformants were screened via colony PCR utilizing EFEPF and EFEPR. Sequences, plasmids, transformants, etc., were verified by Sanger sequencing (Source Bioscience). Expression of EFE in C. necator was confirmed via Western blots with EFE antibody (Thermofisher).
The IPA and 2,3-BDO strains were constructed as shown in
2. Integrated Operons of IPA and 2,3-BDO.
In one embodiment, genes were codon optimized for expression in C. necator H16 (e.g. strain). In one embodiment, bktB(β-ketothiolase, H16_A1445) is native; ctfAB(Succinyl-CoA transferase, AJ000086) from H. pylori, adc(acetoacetate decarboxylase, CA_P0165) from C. acetobutylicum, sadh(secondary alcohol dehydrogenase, AAA23199.2) from C. beijerinckii, alsS(acetolactate synsthase, BSU36010) and alsD(acetolactate decarboxylase, BSU36000) from B. subtilis. pBAD promoter with araC gene was obtained from pCM291rfp plasmid (1) The operons were integrated by a two step homologous recombination using sacB as the selection marker as described previously (2, S11) replacing the native PHB operon (phaCAB).
Primers used for plasmid construction are shown in Table S6. Genes were assembled using the USER assembly method, with the USER cloning kit (New England Biolabs, NEB).
Briefly, a suicide vector pL03(J) (Ref 3) was used for gene deletion and integration. 700 bp upstream of phaC and downstream sequence of phaBl genes were used as homologous regions. The IPA1 and BD2 operons were cloned in between the homologous arms. Alternatively, operons starting from the rrnB T2 terminator were cloned in between the homologous arms.
Suicide vectors, including those described; designed and constructed herein, were transformed via conjugation using E. coli S-17 cells carrying the desired vector and C. necator strains using the protocol described herein and in (Reference (Ref.) No. 2).
Conjugation was performed by transformation of the IPA1 and BD2 suicide plasmids into the mobilizing strain S-17 E. coli followed by incubation of the transformed cells overnight with C. necator on LB, followed by selection on LB+15 μl gentamicin (to select against S-17 cells) and 12.5 μg/ml tetracycline. In some embodiments, transconjugants were selected on minimal media plates with 0.4% fructose and appropriate antibiotics.
A second recombination step was carried out by inoculating single colonies from the first cross-over into low salt LB medium with 15% sucrose without antibiotics overnight. Sucrose resistant colonies were plated onto low salt LB agar plates with 15% sucrose and then single colonies were selected for further screening. Colonies which did not exhibit antibiotic resistance were selected and successful integration of the IPA1 and BD2 operons was confirmed via PCR, utilizing primers flanking upstream and downstream of the homologous sequences. In some embodiments, colonies which did not exhibit antibiotic resistance were selected and successful integration or gene deletions were analyzed using PCR with primers flanking upstream and downstream of the chosen homologous sequences.
Integration of both the IPA1 and BD2 operons and subsequent deletion of the PHB1 operon was confirmed using Sanger sequencing (Eurofins genomics GmbH). Integration or gene deletions were also confirmed using Sanger sequencing (Eurofins genomics GmbH).
For production of the methyl ketones, the plasmid pJM20 was conjugated into C. necator as detailed in Muller et. al., (Ref 33, and as described herein).
3. Transformation of C. necator.
In preferred embodiments, plasmids were transformed into C. necator by electroporation. However, it is not meant to limit the method of transformation. As one example, both the pBBR1 efep and pBBR1 YFP plasmids were transformed into C. necator by electroporation. Cells were made competent by growing a 10 ml overnight culture in SOB (Hannahan's broth) media in a Falcon™ tube at 30° C. at 200 rpm with 10 μg/ml gentamycin. The overnight culture was then used to inoculate a 50 ml culture to an OD600 of 0.05 with 10 μg/ml gentamycin in SOB media and incubated at 30° C. at 200 rpm until they reached an OD600 of 0.3-0.4. Cells were then washed three times with 10 ml ice cold 1 mM HEPES at 4° C. Cells were pelleted by centrifugation at 8000 RPM for 5 minutes at 4° C. The cells were resuspended in 200 ul of 1 mM HEPES. 100-500 ng of pBBRl-efe was added to 100 μl of competent cells in a pre-chilled electroporation cuvette and left on ice for 5 minutes. Electroporation was performed at 2.5 kV, 200Q and 25 uF by a Bio-Rad gene pulser. After electroporation, 0.9 ml SOC media was added to the cells and the cells were transferred to an eppendorf tube and incubated at 30° C. at 200 rpm for 2 hours for outgrowth. After 2 hours of outgrowth dilutions of cells were plated onto LB agar plates with the appropriate antibiotics (300 μg/ml kanamycin in the case of pBBRl-efe) and incubated at 30° C. Transformants appear after 48 hours and were confirmed by amplification of the efe gene by colony PCR.
4. Ethylene Measurements to Confirm Productivity in pBBR1 Efe.
The pBBR1 efe strain was tested for ethylene production using gas chromatography (GC). A single colony was used to inoculate liquid FGN medium, supplemented with 300 μg/mL kanamycin and the cultures was grown for 24 hrs at 30° C. Cultures were then diluted to an OD600 nm of 0.08 in 10 mL fresh FGN medium supplemented with kanamycin (300 ug/ml). 3 mL aliquots were grown overnight in triplicate in 10 mL rubber-capped GC serum bottles at 30° C., 200 rpm. 2 mL of the headspace was collected with a gas syringe after 4, 8, 12, 24, 48 and 72 hrs and analyzed using a Trace™ 1300 gas chromatograph (Thermo Scientific™) under the following conditions: column size: 0.53 mm×40 mm; solid phase: Porapak N column; column temperature: 60° C.; carrier gas: helium and detector: TCD. OD values were determined using a spectrophotometer set at the wavelength λ=600 nm. Ethylene production peaked at 24 hours at 300 nmol/OD600/mL ethylene. No ethylene was detected in the control strain pBBR1YFP.
5. Analytical Methods to Confirm Productivity in C. necator IPA1 and BD2.
The C. necator IPA-1 and BD2 strains were both cultivated in minimal media with a C:N (carbon to nitrogen) of 60 (mol C/mol N), utilising fructose as the sole carbon source to induce the stringent response in 50 ml shake flasks at 200 rpm, 30° C. L-Arabinose was added at a final concentration of 1 g/L (0.1% w/v) to the cultures for IPA-1 and BD2 gene induction. Supernatants from the cultivations were obtained by centrifuging the culture samples for 5 min at 13000 rpm. R,R-2,3-BDO and isopropanol were analysed using HPLC with Aminex HPX-87H column (Bio-Rad, Hercules, Calif.), 5 mM H2SO4 as mobile phase, equipped with UV and RI detectors. The flow rate of the mobile phase was 0.5 mL/min with a column temperature of 50° C. Quantifications were performed from the standard curves obtained using standards purchased from Sigma Aldrich. Biomass growth was quantified based on optical density measurement at 600 nm using a spectrophotometer. R, R-2, 3-BDO was produced at 0.045 g/L in the BD2 strain and isopropanol was produced at 0.11 g/L in the IPA-1 strain. No R, R-2, 3-BDO and isopropanol could be detected in the control strains.
6. Dark Vs. Light Test.
Control tests were performed with the removal of light irradiation. The lack of any detectable levels of NH3, H2, HCOOH, PHB, C2H4, IPA, or BDO demonstrates that these products were not generated from the natural metabolism of the bacteria. It also rules out the possibility of any organic compounds (HEPES, L-ascorbic acid, etc.) acting as a source for product generation.
7. Photocatalytic Test with Bacteria Cells or QDs.
Control tests carried out with either no QDs or no bacteria cells showed negligible (
8. Photocatalytic Test in an Argon Atmosphere.
To further clarify the origin of the nitrogen and carbon in the NH3 and C2H4 products, the same tests were performed in an argon atmosphere instead of air or CO2. The cell cultures (A. vinelandii or C. necator) were bubbled with argon for 1-2 hours to remove any remaining intracellular nitrogen or CO2 gas. The collected cells were washed twice and suspended in argon-purged reaction media. The photocatalytic reactions were carried out in the same condition in a septum-sealed vial with argon headspace. No NH3 or C2H4 was detected up to 3 hours and 48 hours, respectively (
9. Photocatalytic Test with the Microparticles.
To prove our assumption that the photocatalytic air-water and CO2-water reduction reaction was performed intracellularly, we compared the parallel tests using the sub-5 nm QDs (q-CZS and q-CZSe, as used in the experiment groups) and their bulk counterpart (b-CZS and b-CZSe, acting as the control groups) for both NH3 and C2H4 generation. Bulk CdS was synthesized using chemical precipitation method with 0.1 M CdCl2 and 0.1 M Na2S solution. Similarly, bulk CdSe was synthesized with CdCl2 and Na2SSeO3 (by dissolving Se powder in hot Na2S2O3 solution). The b-CZS and b-CZSe were synthesized by coating two monolayer ZnS on the bulk CdS or CdSe with SILAR. Photocatalytic tests were done with replacement of 500 nM QDs with 0.2 mg/ml b-CZS or b-CZSe. No or negligible production of NH3 or C2H4 indicates the necessity of intracellular coupling of QDs with related enzymes for catalytic fuel generation.
10. Photocatalytic Test with the Addition of 2,4-dinitrophenol (DNP).
To prove the direct electron transfer from the QDs to MFN (in A. vinelandii) or hydrogenase (in C. necator) to initiate the air-water or CO2-water reduction reaction, instead of an indirect, ATP-dependent NADPH-mediated process, control experiments with the addition of ionosphere (DNP) at different concentration were conducted (S12, 34). Ionophores can disrupt the cell membrane potential and inhibit the formation of ATP, thus blocking the NH3 or C2H4 generation under natural production conditions (with consumption of sugar). We have observed (
11. Photocatalytic Tests with Different Bacteria Strains.
To further prove photocatalytic product generation, we compared different strains of A. vinelandii and C. necator, where the control groups have different metabolic pathways resulting in much lower or no desired product generation. For NH3 production, A. vinelandii DJ1003 strain (same as DJ995 but also contains an insertion and deletion mutation within nifB) was used as a control, the strain produces apo-nitrogenase (lack of MoFe cofactor) with a low or no nitrogen fixation capability (urea as a nitrogen source is hence required in the culture). The same test was conducted within 2-hours and resulted in trace amounts of NH3 generation (
To prove the effects of zinc-histidine binding in in vivo ammonia production, nanorgs made from A. vinelandii DJ995 and A. vinelandii Wards42 (produces a nitrogenase without histidine tag) with CdS (without ZnS shell) and CZS (with two-monolayer ZnS shell) were also tested. As shown in
12. 13C Isotope Labeling Tests.
13C isotope labeling tests were used to further prove biofuel production from inorganic 13CO2. The tests were performed similar to the other nanorg tests, with the replacement of non-labeled CO2 with 13C-bicarbonate. Typically, 20 mM NaH13CO3 were supplied, with headspace charged with argon. Headspace gas was monitored using a mass spectrometer (Balzers Prisma QME-200) with QUADSTAR software. The fragments (m/q=30 and 47 for 13C2H4+ and 13C2H5O+, respectively,
B. Experimental Parameter Optimization for Improving Product Yield.
The efficiency for in vivo photocatalytic air-water or CO2-water reduction using the QDs-living bacteria biohybrid (nanorgs) depends on many factors. These factors can be briefly classified as the intrinsic QDs part (light absorption, electronic transport, etc.), intrinsic living bacteria part (enzymatic activity, etc.), and the interactions between them (electronic coupling, cellular toxicity, etc.). Experimentally, factors including QDs surface functioning, QDs concentration, bacteria cell optical density, use of electron donor, irradiation intensity, etc. can be varied to obtain optimal product generation. The optimization was performed on NH3 producing strains with nanorgs prepared from CZS1 QDs and A. vinelandii DJ995 bacterial cells. The results can guide the optimization of other QDs and bacteria strains for other fuel production.
1. Bacteria Cell Optical Density.
Firstly, in one embodiment, mixtures with variations in cell optical density (OD600=0.1 to approximately 1.0) were tested, with 200 nM MPA-capped QDs in ASC5 media. Photocatalytic reactions were performed in air, with 400 nm light irradiation at 1.6 mW/cm2 for 1 hour. As shown in
2. QDs Capping Ligands and QDs Concentration.
With fixed cell optical density (OD600=1.0), QDs with different capping ligands (MPA, CYS, CA) and concentrations were tested. As presented in
3. Irradiation Intensity.
NH3 production with varying light intensity (from 0.16 to 2.42 mW/cm2 at reaction site) was also tested (500 nM CYS-capped QDs and OD600=1.0 bacteria cells, 1 hour irradiation time).
Low NH3 yield with low irradiation intensity is (
4. Effect of the Electron Donor.
To investigate the function of the sacrificial quencher (L-ascorbic acid, and sulfide here), we performed the same photocatalytic test in the different medias or buffers, (e.g. HEPES buffer ((e.g. 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid buffer)), MOPS (e.g. 3-(N-morpholino)propanesulfonic acid) buffer, PIPES (e.g. 1,4-Piperazinediethanesulfonic acid) buffer, PBS) with the removal of L-ascorbic acid, using the optimized CYS-capped QDs at 500 nM for 2 hours. As shown in
To further understand the function of the sacrificial donor in photocatalytic NH3 production, the time evolution of NH3 was monitored in the media (HEPES) with or without the ascorbic acid. Interestingly, the total amount of NH3 in the two tests reached the same level (
Similar tests were also performed in Burk media and photocatalytic media supplied with 5, 10, and 25 mM L-ascorbic acid (ASC5, ASC10, and ASC25). As shown in
C. Recovery Tests.
The saturation of NH3 production could be due to the depletion of the sacrificial hole quencher (L-ascorbic acid) or the accumulation of NH3 (an inhibitor for MoFe nitrogenase) in the reaction phase. To prove this hypothesis, the bacteria cells were separated by centrifugation and recharged with new QDs-containing media for photocatalytic reaction every 1.5 hours. Taking the cell loss due to centrifugation, the cell OD were re-measured before each cycle, and the NH3 TON and recovery were normalized to the corrected cell OD (measured OD for the three cycles were 1.00, 0.728, and 0.565, respectively). NH3 production was resumed after each cycle, and no obvious yield loss was seen (almost 100% recovery as shown in
D. Extension to Other QDs and Bacteria Strains for Other Solar Fuel Production.
The knowledge obtained from the optimization of NH3 production can be utilized to conduct other nanorgs tests with different combinations of QDs and bacteria strains, with the benefits of extending the light absorption spectra from near-UV, to visible, to near-IR using different QDs or different excitation sources, and obtain different fuels using different genetically modified strains. Some of the results are presented in
E. Estimation of the Internal Quantum Efficiency (IQE).
The internal quantum efficiency (IQE), defined as the ratio of electron production to the total amount of photon absorbed, was estimated based on some of the following parameters:
Based on the Lambert-Beer's law, the light absorbed A=ε(400)bc=0.313, and the transmittance and can be calculated by A=2−log(% T), and T=48.6% and the absorbed part is 1−T=51.4%.
The incident photon number can be calculated from the irradiation intensity:
And the incident photon flux (Finc=N/t) is Finc=ISλ/hc=1.676×1014 s−1.
The absorbed photon by the nanorgs is Fabs=Finc×(1−T)×uptake=1.2×1014 s−1.
The total electrons produced from the nanorgs can be calculated from the TOF of NH3 (3 electrons per NH3 molecule) and H2 (2 electrons per H2 molecule), and for one cell, the electron flux is Fe/cell=3×TOF(NH3)+2×TOF(H2)=34890 s−1.
With OD600=1.0 cell in 1 ml total volume, the total electron flux is Fe=1.5×1013 s−1.
Therefore, IQE=Fe/Fabs×100%=13.1%.
F. Scaled-Up PHB Production in a Photobioreactor.
To investigate the scaling up of solar-biofuel production, the nanorg test scale was amplified from 5 ml to several liters. The scaled-up production was conducted in a benchtop bioreactor (BioFlo/CelliGen 115). The fermentation (approximately 4 L) was conducted in the MSM media with 200 nM Cys-capped QDs (CZS2 in this case). Wild-type C. necator pBBRl-yfp strain obtained from FGN media (OD600 between 2 and 3) was washed twice with MSM media and re-suspended in the fermentation media with an initial OD600 of 1.695. The media was flushed with CO2/O2 mixture (4:1, 0.5 SLPM (standard litre per minute)) for 15 min before irradiation. The temperature was maintained at 30° C., and the agitation was kept at 200 rpm. The pH was monitored during the whole process (approximately 6.0), and CO2/O2 was recharged every 12 hours. Five 400 nm LED panels (2 on the side wall and 3 at the bottom) were used as irradiation source. The fermentation was stopped after 68 hours, and the nanorgs were collected using centrifugation (7,000 rpm). After washing twice with D.I. water, PHB from the nanorgs was extracted with sodium hypochlorite/chloroform mixture (35) and the chloroform layer obtained from centrifugation was pulled to 70% methanol for precipitating the PHB, which were collected and dried in 70° C. overnight. The whole process was shown in
For scaling up the process for commercial production, our process in lab scale (several liters) is amenable to a pilot scale plant (approximately 1000 L), and eventually even to a commercial level (>40,000 L) with little change of the configuration. The cells with biofuel or bioproduct can simply be filtered using commercial membranes, reuse the buffered water, and simply lyse the cells using detergent (sodium dodecyl sulfate) solution, followed by a charged filtration membrane to capture/recycle all the QDs (due to their surface charge). Once QDs are filtered, the remaining solution with biodegradable plastic, biofuel, or other specialty chemicals as bioproducts can be precipitated (as depicted in
Described herein are compositions and methods used during the development of the present inventions, including but not limited to embodiments of: Quantum dots (QDs) Synthesis; CdS and CdSe QDs; InP@ZnS QDs (IPZS); Cu2ZnSnS4 QDs (CZTS); ZnS shell growth; Ligand exchange; Characterization of QDs, including but not limited to: Optical spectroscopy; Elemental analysis, Electrochemical analysis, and Zeta potential measurements; Cellular enzyme preparation and characterization; Azobacter vinelandii DJ995 bacteria growth and cell lysate preparation; Cell lysate (CL) activity determination; MoFe nitrogenase (MFN) purification; Cupriavidus necator (C. necator) culture; Analysis of product generated from C. necator culture; QDs-enzyme complex preparation and characterization; UV-VIS determination of QDs-protein binding; Fourier-Transformed Infrared Spectroscopy (FTIR); SDS-PAGE protein electrophoresis; Agarose Gel Electrophoresis (AGE); Inductively-coupled plasma mass spectroscopy (ICP-MS); QDs-MFN biohybrid for light-induced proton reduction; Light-induced redox reaction with QDs-CL mixture; Interactions between QDs and the living bacteria; Cellular QDs uptake assay; Laser-scanning confocal microscopy; Cell growth curve measurement; Resazurin dye cell viability assay; Colony forming unit (CFU) assay; Formation of QDs-living bacteria nano-biohybrid (nanorgs) for light-induced air-CO2-water to fuel production; Experiment and Control Groups; Experimental parameter optimization for improving product yield; Recovery tests; Extension to other QDs and bacteria strains for other solar fuel production; Estimation of the internal quantum efficiency (IQE); and Scaled-up PHB production in a photobioreactor.
Results described herein include but are not limited to embodiments of: UV-VIS spectra and photoluminescence (PL) spectra of the QDs; Electrochemical characterization of the QDs; Zeta potential CYS-capped CZS; Natural growth and production with different C. necator strains; Proof of QDs and cellular protein binding; Light-induced H2 or NH3 production with QDs-MFN or QDs-CL biohybrids; Laser scanning confocal images; Cell growth curve assay; Cell viability tests with resazurin; Control experiments with A. vinelandii and C. necator; Optimization of experimental parameters for improved NH3 production; Recovery test for NH3 production; TON of different fuels; and Scaled-up production of PHB, etc.
VIII. Quantum Dot-Azotobacter vinelandii Living Nano-Biohybrid Organisms Cause Light-Driven Air-Water Reduction: Solar-Powered Living Factories.
Many naturally occurring and synthetic bacteria can accomplish industrially relevant reactions, like, for e.g., conversion of nitrogen to ammonia in ambient conditions, using chemical energy to generate electrons and reduce readily available chemical feedstocks, and can be labeled as living factories. Refs. 1-6 However, they derive the chemical energy needed sometimes from valuable food stocks, thereby reducing their attraction for energy conversion to useful solar or biofuels. Inorganic photocatalysts directly derive energy from sunlight to generate photoelectrons for reduction of inexpensive and abundant chemical feedstocks like, for e.g., air, water, and carbon-dioxide, but their lack of selectivity, low efficiency, and sometimes use of conditions such as high-temperature and pressure limit their widespread application. Refs. 7-18.
Combining these desired functionalities of direct stimuli-activations via light, voltage, or magnetic field, with the versatility of designing desired synthetic metabolic networks in living cells can provide an unprecedented platform for designing and creating multifunctional living nano-biohybrid organisms (or nanorg's), and for specific applications as living solar-powered factories for direct energy conversion to solar fuels. Ref19.
One initial step towards development of such living organisms is chemical coupling, site-specific self-assembly20-24 from dispersion, and energetic coupling between QDs and synthetic bacteria by appropriately choosing QD size and material (core-shells, if different materials required for energetic alignment and chemical coupling/biocompatibility), QD surface charge and ligands, and desired site-specific attachment. To ensure good energetic alignment and efficient electron injection from the conduction band of photoexcited QDs to molybdenum-iron nitrogenase (MFN) enzyme in Azotobacter vinelandii25 for multielectron reduction of water to hydrogen, we conducted in-situ experiments with MFN from cell-lysate with different cadmium chalcogenide QDs. Since the choice of chalcogen virtually fixes the valence band state and any change in size and therefore quantum confinement tunes the conduction band position,26 we identified different sizes of cadmium sulfide (CdS,
To identify suitable chemical coupling (and ensure biocompatibility, discussed later) and design site-specific attachment and self-assembly in chosen QDs, we tested large CdS and ZnS nanoparticles. These particles were suspended with cell lysate prepared from Azotobacter vinelandii DJ995 bacteria followed by separation and the resulting protein-bound particles were analyzed using gel electrophoresis (SDS-PAGE) to identify the type of enzymes attached to the nanoparticle surface (
Another requirement for making living nano-biohybrid nanorg's was cell uptake30-33 and viability34-39 of designed QDs. An aspect of this, besides biocompatibility of ZnS coating, is the ligand and charge on QD surface. Using three-different similar-sized QD ligands with different surface charge: mercaptopropionic acid (MPA, negative charge), cysteamine (CA, positive charge) and cysteine (CYS, zwitterion), we tested cell viability of CdS@ZnS QDs using three-different methods.40-42 First, using cell growth (monitored using optical density) in the growing media (Burk media) with nanoparticles, we have observed high growth inhibition for MPA- and CA-coated nanoparticles (
Following design and self-assembly of appropriate living QD-Azotobacter vinelandii biohybrid nanorg's (CdS@ZnS with 2 monolayer shells, with cysteine ligand coating and site-specific attachment with histidine-tagged MFN enzyme), we tested their ability to fix light-energy into specific bonds using inexpensive chemical feedstocks like, for e.g., air and water. Optimized bacteria cell optical density (
While different capping ligands with CdS@ZnS QDs lead to different optimal QD concentrations for improved ammonia production (
In conclusion, we have demonstrated the formation of a living QD-Azotobacter vinelandii DJ995 nano-biohybrid nanorgs via the design of appropriate QDs and facile mixing, self-assembly, and site-specific attachment of desired nanorg's. Based on the success of in-vitro testing, photocatalytic living cell ammonia and hydrogen production are realized in-vivo through air-water and water reduction using light irradiation in non-growing cells. We have shown the importance of optimal QD material and size design due to alignment and charge injection of a photogenerated electron to MFN-enzyme, the function of biocompatible ZnS shell in site-specific Histidine-tagged MFN enzyme binding, charge transport tuning, CdS cytotoxicity reduction. The cysteine-coated CdS@ZnS (2 monolayers thick) QDs showed sufficient cell uptake and cell viability of nano-biohybrids, to facilitate high-efficiency and high-selectivity in-vivo photocatalytic production of ammonia and hydrogen with an optimized turnover frequency (TOF) of 8.73×103 s″1 and 4.35×103 s″1, respectively. This could pave the way of designing highly efficient solar-powered living factories for solar fuel and solar fertilizer generator, using readily available chemical feedstocks. Furthermore, this idea could be extended as a platform technology to design, synthesize, and test other engineered living nano-biohybrid systems, with different combinations of semiconductor nanomaterials and synthetic microorganisms, to harness the power of desired biological processes with multifunctional properties of designer materials like, for e.g., external stimuli-activation with light, electrical pulses, or magnetic field, to wireless communication with living cells.
Living nano-biohybrid organisms or nanorgs combine the specificity and well-designed surface chemistry of an enzyme catalyst site, with the strong light absorption and efficient charge injection (for biocatalytic reaction) from inorganic materials. Previous efforts in harvesting sunlight for renewable and sustainable photochemical conversion of inexpensive feedstocks to biochemicals using nanorgs focused on the design of semiconductor nanoparticles or quantum dots (QDs). However, metal nanoparticles and nanoclusters (NCs), such as gold (Au), offer strong light absorption properties and biocompatibility for potential application in living nanorgs. Here we show that optimized, sub-1 nanometer Au NCs-nanorgs can carry out selective biochemical catalysis with high turnover number (108 mol/mol of cells) and turnover frequency (>2×107 h−1). While the differences of size, light absorption, and electrochemical properties between these NCs (with 18, 22, and 25 atoms) are small, large differences in their light-activated properties dictate that 22 atom Au NCs are best suited for forming living nanorgs to drive photocatalytic ammonia production from air. Further, by comparing the light-driven ammonia production yield between strains producing Mo—Fe nitrogenase with and without histidine tags, we demonstrate that preferential coupling of Au NCs to the nitrogenase through Au-histidine interactions is crucial for effective electron transfer and subsequent product generation. Together, these results provide the design rules for forming Au NCs-nanorgs, and may have implications for carrying out light-driven biochemical catalysis for renewable solar fuel generation.
Biochemical conversion of inexpensive feedstocks like, for e.g., air, water, and carbon dioxide (CO2) into desired chemicals and fuels offers specificity and low cost, but typically requires energetic substrates such as sugar to supply the energy required for conversion. Inorganic catalysts can directly utilize sunlight for photocatalytic conversion at high efficiencies, but suffers from lack of specificity.1-3 Recently, nano-biohybrid catalysts have been suggested an alternative, to combine the best properties of high turnover, efficiency, and selectivity, in a single biocatalyst both using in vitro4-7 and in vivo8-10 studies.
Moreover, living or whole-cell biohybrids offer additional advantage of self-replication or growth, avoiding enzyme deactivation, and enzyme regeneration and repair. Living nano-biohybrid organism, or nanorgs, combine these functionalities as a platform where targeted interfacial chemistry and specific enzyme attachment can be used to ensure facile uptake, self-assembly, and light-driven catalysis to specific chemicals or solar fuels.10
Further, these nanorgs do not require any energetic substrates like, for e.g., sugars, glucose, or NADH, and can directly trigger an energetically uphill biocatalytic transformation of inexpensive substrates (air, CO2) using wireless transfer of energy from light (or sunlight), for a range of selective biochemical or fuel generation as living microbial factories. To further advance nanorgs beyond semiconductor nanoparticles, here we demonstrate nanorgs made from small, sub-1 nanometer Au NCs.11-13 Besides the biocompatibility of Au and their large extinction coefficient,14-16 this work will also serve to advance expanding the nanorg platform to a wide range of materials and microbial platforms.
As described herein, nanorgs were engineered by screening different Au NCs17-20 for biocompatibility with exemplary bacterial strains described herein. For providing good biocompatibility, facile uptake, low hydrodynamic radius, and efficient removal/clearance of unattached Au NC, we focused on Au NCs capped with the zwitterionic ligand, glutathione (GSH).21-23 These atomically-precise Au NCs were synthesized using simple wet-chemistry, with tunable optoelectronic properties (
To allow direct electron transfer from light-activated Au NCs to the intracellular nitrogenase enzyme, cellular uptake and biocompatibility of these nanoclusters are required. Due to their extremely small sizes and favorable surface charge (zwitterionic), intracellular incorporation of these nanoclusters was as high as 90% (
To further probe the effect of cell viability and coupling chemistry between Au NCs and nanorg enzymes, we carried out light-driven ammonia production using these Au NCs-A. vinelandii nanorgs in sugar-free media. This reaction relies on the injection of light-induced electrons from Au NCs to the nitrogenase in the bacteria cells. Since nitrogenase is extremely sensitive to oxygen and is normally protected inside living cells, loss of cell viability could render the complete or partial loss of enzyme activity if rendered in air.4,10,24,25 In order to verify this and correlate the cell viability with product yield, we compared the photocatalytic ammonia turnover number carried out in air (contains dinitrogen and dioxygen) and pure dinitrogen (oxygen-free). Using respective ammonia yield performed in pure dinitrogen as a reference, no change of ammonia production was observed in air with nanorgs made from Au22 NCs, while the ammonia turnover number with Au18 NCs decreased to 30% of the yield in dinitrogen (
As mentioned previously, we selected the DJ995 strain that produces a nitrogenase with a 7× histidine tag in order to facilitate its coupling to the Au NCs (using Au-histidine interactions). As a comparison, a wild-type strain (A. vinelandii Wards) with no such affinity was also used to form the nanorgs. Under the same photocatalytic tests, their ammonia yield is half compared to our optimized systems, which could be explained by the poor electron transfer from the Au22 to the nitrogenase due to unfavorable coupling between them. This again highlights the importance of (specific) coupling between the Au NCs and the enzyme for effective fuel production, as reported previously in the CdS/ZnS QDs-His-tagged nitrogenase biohybrids.10 Similar tests were also conducted on nanorgs made from A. vinelandii DJ1003 strain, which produces an apo-nitrogenase (also with histidine-tag) that lacks the essential Mo—Fe cofactors for nitrogen fixation. As expected, negligible amount of ammonia was produced.
To optimize the turnover number, turnover frequency, and the photon-to-chemical conversion yield (quantum yield, QY), we conducted concentration-dependent light-driven air-water reduction reaction with different Au NCs-A. vinelandii nanorgs. Kinetics of ammonia production indicates no loss of nanorg viability or nitrogenase activity for more than 4 hours in Au22 NC nanorgs (
In conclusion, we have demonstrated light-driven biochemical catalysis using Au NC-A. vinelandii living nano-biohybrids (nanorgs), with a high turnover number (108 mol/mol of cells) and turnover frequency (>2×107 h−1) for ammonia production in ambient air. By screening different atomically-precise Au NCs capped with zwitterionic ligand glutathione, we have selected the best candidate, Au22 NCs, combining desired properties of high cellular uptake, effective light capture, suitable redox potential for electron transfer, and non-toxicity/biocompatibility, for building the nanorgs. In addition, the coupling of Au NCs to the His-tagged nitrogenase produced by A. vinelandii DJ995 strain demonstrates the importance of strong histidine-Au binding for high-efficiency biocatalytic applications. The optimized system can yield a high turnover frequency >2×107 h−1, with photon-to-chemical conversion efficiency (QY yield of 14%) approaching to the theoretical limit (16-20%). Since such high turnover number and frequency is difficult to achieve using conventional synthetic tools and such precise control over individual enzyme activity is not possible with most energetic substrates/conventional substrate-driven enzymatic catalysis, it could provide a new pathway for potentially better design of or high-throughput optimization of such biocatalytic reaction in optimized microbial systems.
These results can pave the way for expanding the choice of available benign and biocompatible materials, some with smaller sized light capturing/emitting particles than QDs, with high absorption cross-sections for use with optimized microbes, e.g. engineered, for building living nanorgs that can carry out light-driven biochemical catalysis for renewable solar fuel generation.
The use of such QDs and gold nanoclusters may also provide benefits over other systems for producing chemicals including reducing costs and lowering negative environmental impact of production.
Although various embodiments are specifically illustrated and described herein, it will be appreciated that modifications and variations of the present disclosure are covered by the above teachings, and as described herein, and are within the purview of the appended claims without departing from the spirit and intended scope of the disclosure. For example, while the engineered organisms are described herein for providing biofuel, such engineered organisms may also be useful for other applications such as plastics.
All patents, patent applications, and publications identified are expressly incorporated herein by reference for the purpose of describing and disclosing, for example, the methodologies described in such publications that might be used in connection with the present invention. These publications are provided solely for their disclosure prior to the filing date of the present application. Nothing in this regard should be construed as an admission that the inventors are not entitled to antedate such disclosure by virtue of prior invention or for any other reason. All statements as to the date or representation as to the contents of these documents is based on the information available to the applicants and does not constitute any admission as to the correctness of the dates or contents of these documents.
Atomically-precise gold NCs capped with glutathione (GSH) capping ligands were synthesized based on the methods reported by Ghosh et al., Zhang et al., and Kamat et. al.11-20
Au18 NCs were synthesized by dissolving 150 mg HAuCl43H2O in 1.2 mL methanol and adding 1.8 mL DI Water. 300 mg of glutathione was added to this solution and sonicated to dissolve. Once the glutathione dissolved and the color changed from yellow to almost colorless. 96 mL methanol was then added and stirred for 10 min. 4.5 mL of a 220 mM NaBH3CN solution was added under vigorous stirring for 30 min. After 30 min the precipitate was removed by centrifugation and washed with methanol repeatedly to remove any remaining precursor. Finally, the precipitate was dissolved in water and freeze-dried to obtain a pale red powder identified as Au18SG14.
Au22 NCs were prepared by mixing 12.5 mL 20 mM HAuCl4 and 7.5 mL 50 mM glutathione solution in a 500 mL flask containing 180 mL of DI water. After vigorously stirring for 2 min, the pH was raised to 12.0 with 1M NaOH, after which 0.24 mg NaBH4 in 0.1 mL DI water was added to the reaction with sitting at 500 rpm. After 30 min, the pH was lowered to 2.5 with 0.33 M HCl. The reaction solution was then sealed airtight with stirring at 200 rpm and allowed to react for 8 hours forming a red-emitting Au22 solution. The NCs were cleaned by using isopropyl alcohol and centrifugation and finally resuspended in water and kept in the fridge away from light for future use.
Au10-12, Au15, and Au25 NCs were synthesized by carbon monoxide (CO) reducing techniques. Briefly, in a 125 ml flask with 20 ml distilled water, 100 mM HAuCl4 and 200 mM reduced glutathione were added to a final concentration of 1 mM and 2 mM, respectively. The pH of the mixture was adjusted to 7, 9, and 11 for Au10-12, Au15, and Au25 NCs synthesis, respectively. The flask was sealed with a rubber septum and flush with pure CO gas through a syringe needle for 2 min. The mixture was violently stirred for 24 hours and the resulting Au NCs was precipitated with an excess amount of isopropanol, followed by separating with centrifugation at 5,000 rpm. The precipitates were dried with clean air and re-suspend in distilled water. The Au NCs were stored in 4° C. for future tests.
Ultraviolet-visible (UV-VIS) spectra were measured using the UV1600PC UV-VIS spectrometer (VWR). Dynamic light scattering (DLS) was performed on Litesizer 500 (Anton-Paar) to quantify the hydrodynamic size and zeta potential of the Au NCs.
The conduction band (CB) of the NCs was characterized using differential pulse voltammetry (DPV), with a Bio-logic SP200 potentiostat. A three-electrode configuration with a 3 mm glassy carbon working electrode, a platinum wire counter electrode, and a Ag/AgCl reference electrode was used. Au NCs suspension (in 0.1 M Na2SO4 electrolyte) was bubbled with argon for 10 min before the measurement. DPV was taken with the following parameters: 50 ms pulse width, 50 mV pulse height, 200 ms step width, and 4 mV step height (˜20 mV/s scan rate).
A. vinelandii DJ995 and DJ1003 strain were kindly provided by the Dennis group (Virginia Tech). Typically, bacteria were grown in nitrogen-limited Burk media (with 3 mM urea) at 30° C., with 200 rpm shaking. Cultures at an optical density (OD) ˜1.5 (overnight culture) were collected for future tests.
Cell growth curves and resazurin assays were performed in Burk media and photocatalytic media, respectively, with a variation of Au NC size. A. vinelandii DJ995 was first grown in nitrogen-free Burk media and harvested near OD 1.0, washed twice and resuspended in photocatalytic media. The cells and their respective NCs were then incubated and subject to light-exposure to produce NH3. After production was complete (˜4 hrs) the nanorg mixtures were subject to both growth and viability measurements. The cell growth curves were taken in a 96-well microplate at 30 C with vigorous shaking and monitored using a microplate reader (TECAN GENios) controlled by Megellan 7.2 software. An initial OD of 0.1 in Burk media was used for the growth curves. The resazurin assay was performed separately in a 96-well plate with the bacteria maintained at an OD of 1.0 after the photocatalytic tests. Resazurin was added to a final concentration of 0.1 mg/mL and the fluorescence was measured at 620 nm (485 excitation) over the course of 2 hours.
Uptake of Au NCs by A. vinelandii was determined by a UV-vis spectrophotometric method. NCs first had their UV-vis spectra recorded in photocatalytic media at the desired concentration (20 μM). The initial absorption value at the first excitonic peak was taken to be the 0% uptake mark (Absi). Then the NCs at the desired concentration (20 μM) were incubated with the bacteria at OD 1.0 in photocatalytic media for 15 minutes on a shaker at 250 rpm. The mixtures were then centrifuged at 5,000 rpm and aliquots of the supernatant were taken for UV-vis measurement. The absorption values of the supernatant at the corresponding excitonic peak position (Absf) were used to calculate the uptake % using the following equation:
Uptake %=Abs2Absf/Abs1×100%
Ammonia production with the nanorgs was achieved by first growing A. vinelandii to an OD of ˜1.0 in Burk media. The cells were then washed twice with sugar-free photocatalytic media (25 mM ascorbic acid 35 mM HEPES) to remove any residual sugars. Cells were then resuspended in photocatalytic media and diluted to a final OD of 1.0. The Au NCs were then added to the bacteria and allowed to incubate for 15 min before subjecting to production. For production, the nanorgs were subjected to light irradiation (405 nm LED, 1.6 mW/cm2) for 5 hours on a shaker at 250 rpm. 25 μL aliquots were taken at various time points throughout the reaction to monitor the kinetics.
Ammonia was quantified using a fluorescent assay described previously. Typically, 25 μl sample was added to 0.5 ml o-phthalaldehyde assay reagent and incubated in dark for 30 min, followed by measuring the fluorescence at 472 nm (excitation at 410 nm).
CdS and CdSe nanoparticles for QDs were synthesized using a modified method developed by Peng et. al, (12/20). Their size can be controlled by varying the amount of oleic acid (OA) capping ligand. A total 12 g mixture containing 38.4 mg CdO, 316, 1904, or 5712 μl OA and 1-octadecene (ODE) was vacuum-degassed at 80° C. and refilled with argon for three cycles. The mixture was then heated up to 300° C. and injected with a sulfur precursor (4.8 mg sulfur powder dispersed in ODE). The resulting reaction phase was cooled down to 250° C. and the CdS QDs were grown for 1 hour. The removal of unconsumed cadmium and oleic acid was performed by extraction with warm CH3OH (50° C.) in a separation funnel. This process was repeated three times and the resulting ODE layer was obtained, the removal of residual CH3OH was carried out under vacuum at 80° C. To transfer the CdS nanoparticles to chloroform (CHCI3), the ODE layer was precipitated with chloroform and acetone followed by washing for at least three times. The CdS nanoparticles for QDs were re-dispersed in chloroform and stored in the dark. CdSe nanoparticles for QDs with varying sizes were synthesized using the same method, with the sulfur being replaced with a selenium precursor (12 mg selenium powder dispersed in ODE with 41 μl tributylphosphine (TBP)). Ultraviolet-visible (UV-VIS) spectra (
IPZS were synthesized using a modified method developed by Fichter et. al, (26). A mixture of 9 ml oleylamine (OLA), 119.4 mg InCl3, and 73.5 mg ZnCb were subjected to a vacuum, using a Schlenk system, at 110° C. for 1 h, followed by recharging with argon and ramping the temperature to 220° C. After 15 min, 0.24 ml tris(trimethylsilyl)phosphine ((TMS)3P) was swiftly injected into the mixture, and the InP cores were allowed to continue growing for 10 min. The reaction phase was then cooled down to 80° C., and 1.27 ml of 1-dodecanethiol (DDT) was added drop-wise. The system was vacuum-degassed and recharged with argon, followed by raising the temperature to 200° C. for 1 hour. After cooling down to 70° C., 5 ml hexane was added, and the mixture was centrifuged at 4,000 rpm for 10 min to remove the large aggregates. The supernatant was precipitated with excess acetone, and the particles were obtained by centrifugation at 5,000 rpm for 20 min. After drying with N2, the precipitate was suspended in 10 min OLA with 142.2 mg zinc stearate. The mixture was degassed and recharged with argon, and the temperature was raised to 180° C. After 3 hours, the system was cooled down, and the IPZS were obtained by centrifugation as mentioned above, and as described herein, and re-suspended in hexane.
CZTS QDs were synthesized following the procedure outlined by Yang et. al,S2 (27). Briefly, 0.13 g copper acetylacetonate, 0.05 g of zinc acetate dihydrate, 0.048 g of tin chloride, and 0.033 g of sulfur were added to 10 mL of oleylamine in a 40 mL round-bottom flask. The mixture was stirred under vacuum for 2 hours, then heated to 110° C. while purging with Ar and held for 30 min. The temperature was then raised to 280° C. and held for 1 hour, then cooled to room temperature. Precipitation of the nanoparticles was achieved by adding ethanol and centrifuging at 5,000 rpm for 20 min. Redispersion in chloroform and centrifugation at 5,000 rpm for 5 min was used to isolate and discard aggregates in the resulting pellet. ZnS shell growth follows the same protocols for the CdS or CdSe (see section D below, ZnS Shell Growth on CdS nanoparticle cores).
The synthesis (e.g., growing) of the CdS@ZnS core-shell nanoparticles for QDs (CZS) was adapted from the method reported by Peng et. al, (4428). The size and concentration of the CdS cores (i.e. nanoparticles) were determined using the following formula:
D=(−6.6521×10−8)λ3+(1.9557×10−4)λ2−(9.2352×10−2)λ+13.29
ε5500×E×D2.5
c=A/εl
where λ, E, A are the wavelength, photon energy, and extinction at the first exciton peak, respectively. l is the optical path of the cuvette. D, ε, and c are the diameter, extinction coefficient and concentration of the CdS nanoparticles for QDs. The CdS stock solution (dispersed in ODE) was determined to have a concentration of 0.0377 mM.
The zinc precursor (0.1 M Zn2+) was prepared by a heating a degassed mixture containing 82 mg ZnO, 2.82 ml (2.51 g) OA and 7.2 ml ODE to 250° C. The resulting clear solution was cooled down and stored in a septum sealed vial. The precursor was gently heated up to 60° C. before use.
The sulfur precursor (0.1 M) was prepared by dispersing 32 mg sulfur powder in 10 ml ODE with sonication. The resulting clear solution was bubbled with argon for 30 min and stored in a septum sealed vial.
ZnS shells were grown using a layer-by-layer deposition. A mixture containing 120 nmol CdS cores (3.55 nm; 3.2 ml CdS stock solution) and 2 ml oleylamine (OLA) was vacuum degassed and recharged with argon for three cycles under 120° C. A defined amount of zinc and sulfur precursors were injected simultaneously, and the reaction phase was kept at 120° C. for 5 min. The reaction was then raised up to 220° C. for the growth (20 min) of the first ZnS layer. The reaction was then cooled down to 120° C., and the UV-VIS spectrum was taken to determine the extinction coefficient of the CdS@ZnS core-shell nanoparticles (CZS). The resulting solution could be either washed (similar to CdS nanoparticles for QDs) or used for 2nd or 3rd ZnS layer growth. To grow nominal 1 approximately 3 monolayer ZnS shells, 0.44, 0.60 and 0.77 ml of zinc and sulfur precursors (each) in solvent were used, respectively, (determined by simple geometrical calculation, as reported by Peng et al.) zinc and sulfur precursors (each) were used, respectively.
The real thickness of CdS shell and extinction coefficient of CdS@ZnS core-shell nanoparticles (summarized in Table S1, Table S1-2) were determined by UV-VIS spectrum (
The real thickness of the ZnS coating can be estimated from the cadmium and zinc ratio (determined by ICP-MS) by using simple geometrical calculations shown in
Similar to the CZS, CdSe@ZnS core-shell QDs (CZSe) can be synthesized using the same layer-by-layer deposition technique. Starting with CdSe QDs with the first exciton peak position at 500, 525 and 580 nm, CZSe1, CZSe2, and CZSe3 with green, yellow, and orange emission (
QDs (nanoparticles) suspended in CHCl3 were phase transferred into aqueous solution by ligand exchange with 3-mercaptopropionic acid (MPA), L-cysteine (CYS) or cysteamine (CA). For ligand exchange with MPA, 0.1 ml MPA was added to 0.3 ml QD suspension (approximately 10 mM). 0.3 ml ethanol (EtOH) was added, and the mixture was vigorously stirred with gentle heating. 1 ml 1 M NaOH solution was then added, and stirred for a further 5 min. The upper part (aqueous phase) was collected and centrifuged at 15,000 rpm. The obtained QD precipitates were re-suspended in water pH 11. The QD suspension was further concentrated with a 3,000 Da centrifugal filter and washed twice with water pH 11. The concentration of the QD suspension (in water) was determined using the UV-VIS spectra. Ligand exchange with CYS is similar. For the ligand exchange with CA, cysteamine hydrochloride was used, with the replacement of 1 M NaOH solution by DI water and finally re-suspended in water pH 4. The ligand-exchanged QDs (nanoparticle suspension) were stable for up to 1 week in the 4° C. fridge.
A. Optical Spectroscopy.
Ultraviolet-visible spectra of the QDs were measured using the UV1600PC UV-VIS spectrometer (VWR), and the Photoluminescence (PL) spectra were taken using a QM-6 steady-state fluorimeter (PTI). The QDs optical spectra are shown in
B. Elemental Analysis
Real thickness of CdS shell and extinction coefficient of CdS@ZnS core-shell nanoparticles.
The real thickness of CdS shell and extinction coefficient of CdS@ZnS core-shell nanoparticles (summarized in Table S1, Table S1-2) were determined by UV-VIS spectrum (
The real thickness of the ZnS coating can be estimated from the cadmium and zinc ratio (determined by ICP-MS) by using simple geometrical calculations shown in
Taking the 1 ML sample as an example: The volume and moles of the CdS core (diameter D1=3.55 nm, the density of CdS p(CdS)=4.82 g/cm3, MW is an approximate MW=molecular weight):
V(CdS)=π/6×D13=2.34×10−20 cm3
n(CdS)=p(CdS)×V(CdS)/MW(CdS)=7.83×10−22 mol
The total amount of Cd and Zn (in ppb) was determined by ICP-MS. Due to Zn impurity in Cd precursor used in the synthesis, the value for the shell Zn was corrected by the amount of Zn in the CdS core (assume in a single nanoparticle, the amount of Cd is the same):
The molar ratio of Zn to Cd (AW=approximate atomic weight):
The moles and volume of the ZnS shell (the density of ZnS p(ZnS)=4.10 g/cm3):
n(ZnS)=r(Zn:Cd)×n(CdS)=3.62×10−22 mol
V(ZnS)=n(ZnS)×MW(ZnS)/p(ZnS)=0.860×10−20 cm3
The total volume and diameter of the CdS @ZnS QDs:
V(CdS@ZnS)=V(CdS)+V(ZnS)=3.20×10−20 cm3
D(CdS@ZnS)=(6×V(CdS@ZnS)/π)1/3=3.94 nm
The real thickness (in ML unit, ZnS monolayer thickness d(ZnS)=0.312 nm):
Thickness (ZnS)=(D(CdS@ZnS)−D(CdS))/(2×d(ZnS))=0.63ML
The extinction coefficient (at a specific wavelength) of the QDs (core-shell, with nominal 2 monolayer ZnS shell) can be determined using the amount of QDs cores and the UV-VIS of the core-shell QDs. These values are summarized in Table S2.
C. Differential Pulse Voltammetry (DPV) Electrochemical Analysis
Differential pulse voltammetry (DPV) with QDs (in) suspension was used to determine the conduction and valence band positions of the CdX (X=S, Se) QDs. Refs. 27, 29, 45. This was done using a three-electrode configuration with a 2 mm platinum plate electrode, platinum wire, and silver wire as the working, counter, and (quasi-) reference electrode. Ferrocene was used as an internal reference. The CdX QDs (nanoparticles) were suspended in CH2Cl2 with 100 mM n-Bu4NPF6 as the electrolyte. The whole system was purged with argon, and the DPV was measured using a Bio-logic SP200 potentiostat with the following parameters: 50 ms pulse width, 50 mV pulse height, 200 ms step width and 4 mV step height (which corresponds to a 20 mV/s scan rate). The results are presented in
D. CdS, CdSe, CdS(S>ZnS Nanoparticle Thin Film Electrochemistry.
QD thin film electrochemistry was conducted to evaluate the charge carrier dynamics of the QDs. The CdS, CdSe and CdS@ZnS nanoparticles (in ODE) were transferred into CHCI3, as described herein. 50 ul of about 5 uM suspension of nanoparticles were drop-casted on clean fluorinated-tin oxide (FTO) coated glass (about 0.5 cm2) and fully dried in a vacuum desiccator. The CdS, CdSe, and CdS@ZnS QD electrodes were fabricated by drop-casting 50 μl of 5 μM QDs suspension (in chloroform) on the clean fluorinated-tin oxide (FTO) coated glass slides (approximately 0.5 cm2).
Electrochemical measurements were taken using a three-electrode configuration, with the QD electrodes/nanoparticle coated FTO glass, platinum wire and Ag/AgCl electrode as a working, counter, and reference electrodes. 0.5 M sodium sulfate (pH=6.4) solution was used as an electrolyte. The whole system was purged with argon for 20 min before the measurements were taken.
Electrochemical impedance spectroscopy (EIS) was used to evaluate the charge trapping and charge transfer in these QDs under light irradiation. Measurements were taken under 365 nm UV irradiation (˜5 mW/cm2) at open circuit potential (OCP), with a frequency range from 100 kHz to 100 MHz. These spectra are shown with the Nyquist plot (
Open circuit potential (OCP) was used to evaluate the charge transfer and charge recombination on the QD surface. The OCP of the QD electrode was first measured in the dark, followed by irradiation with 365 nm UV light (˜5 mW/cm2) until the OCP reached constant values (
E. Zeta Potential Measurements.
The zeta potential of the CYS-capped CZS (suspended in water with different pH values) was quantified using a Litesizer 500 (Anton-Paar). As expected (
Each of which are herein incorporated by reference in its entirety:
A. Non-Parenthetical Numbers
B. Parenthetical Numbers
Each of which are herein incorporated by reference in it's entirety.
Each of which are herein incorporated by reference in it's entirety:
Each of which are herein incorporated by reference in it's entirety:
This invention was made with government support under grant number CBET1351281 awarded by the National Science Foundation. The government has certain rights in the invention.
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/US20/32899 | 5/14/2020 | WO | 00 |
| Number | Date | Country | |
|---|---|---|---|
| 62927452 | Oct 2019 | US | |
| 62847653 | May 2019 | US |
