Facing climate change and scarcity of resources, the investigation of more sustainable pathways for the production of fine and platform chemicals is one of the most urgent challenges for science. Processes employing atmospheric oxygen and carbon dioxide as well as water substrates seem to be especially promising since they are readily available and per se environmentally friendly. Furthermore, light is the most abundant and reliable energy source of our planet. We were inspired by a publication from Gutekunst and co-workers about the fermentation of dihydrogen with an engineered strain of the cyanobacteria Synechocystis.[1] The hydrogenase HoxYH was fused to the PsaD subunit of photosystem I (PSI), allowing direct electron transfer from the PSI to the hydrogenase. This electron transfer, which is in competition with the reduction of the electron carrier ferredoxin (Fd) and the utilization of the reduction equivalents within other metabolic processes, enables the photosynthetic production of dihydrogen from only light and water.[1] This conversion is regarding its sustainability superior over the production of hydrogen from natural gas in the steam reforming process and the electrolysis of water since no non-renewable resources are consumed and no conversion of light into electricity is necessary. Our ideas on how this PSI-HoxYH construct could be used included various in vivo and in vitro applications, such as the production of hydrogen in isolated thylakoids to avoid complications related to GMOs.[2]
Fig. 1: thylakoid electron transport chain in the Synechocystis mutant with HoxYH-PSI fusion. Both photosystems are highlighted in green, the hydrogenase is highlighted in light blue. The electron transport chain is marked in dark blue. (PSII: photosystem II; PQ: plastoquinone; Cyt-b6f complex; PC: plastocyanin; PSI: photosystem I; Fd: ferredoxin).
However, we quickly noticed that the system described by Gutekunst and co-workers suffers from several drawbacks: The in vivo production of hydrogen is at a very initial stage. The achieved concentrations are very low and cannot compete in any way with commercial methods for hydrogen production. Furthermore, the presence of oxygen leads to the inactivation of the hydrogenase and the formation of reactive oxygen species. This is why the oxygen level needs to be kept low, using glucose as a reductant in combination with glucose oxidase and catalase.[1] In this way, at least one molecule of glucose is needed to synthesize one molecule of hydrogen. Furthermore, self-designed gas sensors or electrochemical measurements are often utilized in literature for hydrogen measurements, which requires appropriate know-how.[1,3] Therefore, the measurement of the nonetheless low hydrogen concentration could prove to be difficult. Finally, working with hydrogen gas always leads to safety risks, since hydrogen is a highly flammable gas. Despite the inspiring work of Gutekunst and co-workers, we excluded working with hydrogenases from our iGEM project, but took this as an opportunity to think about fusion constructs between redox-active enzymes.
As we continued with the speculations about the utilization of fusion proteins in metabolic processes, it was important to consider the advantages and disadvantages of fermentation in order to find suitable targets. The main advantages are the independence from the petrochemical industry and the avoidance of toxic reagents and solvents, which are often required in synthetic processes. No expensive metal catalysts are required, which are sometimes produced in filthy mining operations, and the arising waste from the fermentation broth is per se biodegradable. Furthermore, the construction of stereocenters can be performed in a gentle and efficient manner in contrast to most chemical processes. In contrast to the production of high-value natural products through cultivation and extraction in plants, there is no need for large areas of acreage, the harvest is not subject to annual and seasonal fluctuations and microorganisms grow much faster compared to crops.
Table 1: advantages and drawbacks of fermentation.
Advantages Drawbacks
  • Sustainable
    • Independant from petrochemical industry
    • No toxic reactants or solvents
    • No expensive metals required
    • Biodegradable waste
  • Easy Stereocontrol
  • In contrast to extraction from plants no annual/seasonal variations, less area required, corresponding plants often rare and slow growing
  • Depending on the product, often low titers
  • All required enzymes and genes need to be well known
  • Separation of complex mixtures
  • Poor space-time yields
  • Scale-up issues
  • Long time until organisms are optimized for production of the desired product
The major drawbacks of fermentation are that in many cases only low titers of the desired products are achieved, all of the required enzymes have to be described and the corresponding sequences of the enzymes have to be known as well as that the fermentation products needs to be seperated from a highly complex mixture. The low titers often lead to poor space-time yields compared to chemical processes, and the scale-up of the fermentation processes can be challenging. Most of these drawbacks can be tackled by delicate metabolic engineering until the employed organisms are highly optimized to the requirements needed, but these optimizations can prove to be very time consuming and last for years until satisfactory results are obtained.
The above discussed advantages and drawbacks suggest the fermentation of scarce, chiral natural products with a high market value within our iGEM project, since in contrast to the production of platform chemicals, lower space-time yields can be justified and thus even less efficient processes are still in proximity to economical applications. Terpenes and terpenoids appeared to be a promising subgroup of natural products for our requirements. Many terpenes and terpenoids are high value products and employed as the active ingredients in medical applications as well as fragrances and flavours in cosmetic products, perfumes, food, beverages and cleaning agents. Many different aspects of the fermentation of terpenoids are discussed in scientific literature. There is a large market and great commercial interest in these substances, and other iGEM Teams used terpene biosynthesis before,[4,5] so there were many starting points and a lot of inspiration for us. Moreover, once established in an organism, terpene biosynthesis offers a variable platform, since the synthesis of all terpenes and terpenoids diverges from two monomer-like building blocks that are always identical. The functionalization of the backbones takes place via conversions with cytochromes, so that the idea of using redox-active fusion proteins can be taken up here for the engineering of terpene biosynthesis.[6]
Terpenes are a group of natural products which represent oligomers of isoprene. Terpenes are a group of hydrocarbons, whereas terpenoids are higher functionalized molecules with functional groups such as alcohols, ethers or carbonyls like esters and ketones. With more than 40,000 described substances, terpenes and terpenoids are one of the largest subgroups of natural products. Terpenes and terpenoids are classified based on the number of connected isoprene units. Linear, cyclic as well as bridged structures of the carbon backbone are common.[6]
Table 2: classification of terpenes and terpenoids based on the number of connected isoprene units.
subclass number of isoprene-units number of carbon atoms
hemiterpene 1 5
monoterpene 2 10
sesquiterpene 3 15
diterpene 4 20
sesterterpene 5 25
triterpene 6 30
tetraterpene 8 40
polyterpene >>8 >>40
Typically found in plants, terpenes and terpenoids often exhibit a large spectrum of biological activities and serve as a defense against herbivores, attract pollinators or and play a central role in plant disease resistance. However, some terpenes also occur in other organisms such as microorganisms. The fragrance linalool is a typical example for a linear monoterpene alcohol. It has widespread uses in cleaning agents and hygiene products such as shampoos. An example for a cyclic terpene is limonene, which can be found in the peels of citrus fruits. Beneath its application as a fragrance it is also used in the food industry. The bicyclic monoterpene α-pinene is isolated from conifers and one of the most widespread terpenes. It is repellent to insects and used as a feedstock for the production of other terpenes. The diterpene alcohol phytol is a constituent of chlorophylls.[6]
Figure 2: Structures of isoprene, selected monoterpenes and the diterpene alcohol phytol.
The central metabolites in terpene biosynthesis are isopentenyl pyrophosphate (IPP) and dimethylallyl pyrophosphate (DMAPP) as isoprene equivalents. The biosynthesis of these intermediates can take place over two pathways: the mevalonate pathway (MVA pathway) and the non-mevalonate pathway (also known as DXP-pathway). In the following, only the MVA pathway will be discussed.
The MVA pathway starts from the widespread metabolite acetyl coenzyme A (Ac-CoA). Three molecules are connected in two steps with acetoacetyl coenzyme A as an intermediate to form 3-hydroxy-3-methylglutaryl coenzyme A (HMG-CoA). The following step is rate determining in this pathway: the reduction of HMG-CoA to mevalonate, catalyzed by the HMG-CoA reductase. The following two phosphorylation steps yield mevalonate pyrophosphate, which is then decarboxylated into IPP. IPP is isomerized into DMAPP by the IPP isomerase; both compounds are in equilibrium with each other.[7,8] The heterologous expression of the MVA pathway enzymes in various organisms is well established in literature.[9] Plasmids with the corresponding genes are available from the organization AddGene for Escherichia coli.[10]
Figure 3: Intermediates and enzymes of the MVA pathway.
In order to show how IPP and DMAPP are connected to form terpenes and how these are functionalized to yield terpenoids in the following metabolic processes, the biosynthesis of the sesquiterpene zerumbone will serve as an example. First, farnesyl pyrophosphate (FPP), the common precursor of all sesquiterpenes, is synthesized by the FPP synthase. This is done stepwise by two condensation reactions. At first, one molecule of IPP is condensed with one molecule of DMAPP. The resulting geranyl pyrophosphate (GPP) then undergoes a further condensation reaction with DMAPP, resulting in the synthesis of FPP.[6,8] The biosynthesis of cyclic sesquiterpenes then proceeds with an FPP cyclase as the catalyst. In the case of humulene biosynthesis, this cyclase is α-humulene synthase; the pyrophosphate is first used as a leaving group to generate a carbenium ion. After nucleophilic attack of the corresponding double bond, the intermediately formed carbocation reacts by deprotonation and formation of a new endocyclic double bond to form α-humulene.[11-13]
Figure 4: Biosynthesis of α-humulene, starting from IPP and DMAPP, the primary products of the MVA pathway.
The primary synthesized terpenes are oxidized and thus functionalized by cytochrome P450 (CYP P450) monooxygenases. In the biosynthesis of zerumbone, the involved enzyme is α-humulene-8-hydroxylase. The resulting 8-hydroxy-α-humulene is converted to zerumbone by a dehydrogenase in the final step of the biosynthesis.[12]
Figure 5: Oxidation of α-humulene to zerumbone.
IPP and DMAPP are not only the starting material for the biosynthesis of terpenes, but terpene-derived structural elements are also involved in the construction of other classes of natural products. For example, terpenes are precursors for steroids and monoterpene units can be found in the structures of cannabinoids such as THC and CBD. Monoterpene indole alkaloids such as the well-known strychnine are also examples of structures whose synthetic precursors are derived from the MVA pathway. Our approach using the MVA pathway in combination with redox-active fusion proteins is thus not only limited to the production of terpenoids, but can be understood as a more general approach towards the biotechnological production of various complex natural products.[8,14,15]
Figure 6: Structures of THC and strychnine. The terpene derived moieties are highlighted in red.
In general, monooxygenases are enzymes that can introduce an oxygen atom into a substrate, using molecular oxygen as an oxidant. The second oxygen atom of the oxidant is reduced to water, often with NADPH or another electron carrier as Ferredoxin or a flavoprotein as the reductant. The common feature of all CYP P450 enzymes is that they are heme proteins and thus exhibit hemes on their active site. The prosthetic heme group is bound to the backbone of the enzyme via a cysteine residue. Plants generally exhibit a large variety of CYP P450 enzymes, which are for example involved in the biosynthesis of secondary metabolites. The electron transfer between the CYP P450 and the final reductant often involves a cytochrome P450 reductase (CPR) as an electron mediator. Natural forms of plant CYP P450 and the corresponding CPRs are generally localized in the membrane of the endoplasmic reticulum.[16,17,18]
Figure 7: Reaction mechanism of CYP P450 monooxygenases with peroxide shunt.
A great challenge for the expression of plant CYP P450 in E. coli as the most convenient platform for synthetic biology is the membrane anchor domain of both, the CYP P450 and the corresponding CPR, rendering the proteins insoluble. The membrane recognition signal of eukaryotic proteins can furthermore be incompatible for prokaryotic host organisms. With the above discussed redox active fusion proteins in mind, we envisioned the linkage of truncated versions of CYP P450s with appropriate CPRs without membrane domains to use them for fermentation purposes in prokaryotic organisms. Indeed, the value of this approach is already outlined by examples from literature. For example, such fusion proteins had already been employed for the biosynthesis of plant isoflavones in E. coli. Furthermore, various fusion proteins had been designed with CYP P450 from mammalian cells to investigate drug metabolism and the activation of carcinogens. Despite all of these previous efforts, we felt that the literature lacks of a generalization of this approach: The corresponding fusion constructs are usually designed for a specific application. For this reason, we envisioned a toolbox approach for the construction of CYP P450/CPR fusion proteins to enable easier access to these constructs. We suggest a standardization of the procedure which should involve a standardization for the removal of the membrane-binding domain and the introduction of only a few CPRs that are broadly applicable for a large variety of CYP P450 enzymes. In addition, bioinformatic methods should be investigated to design linkers between the CYP P450 subunit and the CPR: To ensure electron transport between the CYP P450 and the CPR, both subunits of the fusion protein must be able to interact correctly. An appropriate linker can significantly increase the performance of the fusion construct. However, in addition to designing soluble fusion constructs, membrane-bound fusion proteins can also be expressed in E. coli if the membrane-binding domain of CYP P450 in the fusion construct is exchanged for one that is functional in E. coli. In addition to the possibility of using plant CYP P450s in organisms such as E. coli, the use of fusion constructs has also other advantages. CYPBM-3 is a natural fusion of a CYP P450 enzyme and a CPR and exhibits a remarkably high turnover rate: the electrons can be directly channeled to the reactive site of the CYP P450. This is another opportunity offered by the use of such CYP P450-CPR fusion proteins: The reaction rates of CYP P450 enzymes can be increased, which could be used to optimize metabolic pathways and could be a way to increase product titers and make biotechnological production of chemicals more effective.[18,19, 20]
Figure 8: Construction of CYP P450/CPR fusion proteins with and without artificial membrane anchors.
Artemisinin is a sesquiterpene lactone from the plant Artemisia annua. It has a complex polycyclic structure and exhibits an endocyclic peroxide as a characteristic structural feature. Since its first isolation and description at the beginning of the 1970s, artemisinin and its derivatives has proven to be a very efficient drug against Plasmodium falciparum, a unicellular parasite which is the deadliest Plasmodium species causing malaria. Malaria affects mostly people in the tropical and subtropical regions and caused 409,000 deaths and estimated 229 million cases in 2019.[8,21,22,23] In 2015, Tu Youyou was awarded with the nobel prize in medicine or physiology for her discovery and the initial investigations of artemisinin.[24] However, due to fluctuations in the harvests of Artemisia annua, the artemisinin market is not stable.[8]
Figure 9: Structure of artemisinin.
To follow in the footsteps of iGEM team Hamburg 2018, which was working on a different approach to fight malaria by tackling the spread of the parasite via mosquitoes,[25] we selected artemisinic acid (AA) as one of our target products. AA is the precursor to semisynthetic artemisinin and its biotechnological production as an alternative for the extraction from Artemisia annua is a long story: In the early 2000s, Keasling and co-workers were working on genetically modified E. coli strains for the production of the terpene amorphadiene, which exhibits the same carbon backbone as AA.[26] After identifying CYP71AV1 as the key enzyme for the conversion of amorphadiene into AA, the fermentative production of AA in yeast was first reported in 2006.[8]
Figure 10: Conversion of FPP to amorphadiene and subsequent oxidation to artemisinic acid.
After promotion by the Bill and Melinda Gates Foundation and further improvements in the biotechnological production of AA by Keasling and the company Amyris,[21] the company Sanofi managed a scale-up of the chemical conversion of AA into artemisinin in good yields of 55%. Artemisinin was synthesized in 370 kg isolated material per batch with a total amount of 35 tons in 2013. The reaction sequence starts with a stereoselective hydrogenation and subsequent conversion of the carboxylic acid moiety into a mixed anhydride by ethyl chloroformate. The activated form of dihydro AA is then exposed to singlet oxygen, leading to the formation of an allylic hydroperoxide, which is rearranged in a Hock cleavage, induced by trifluoroacetic acid. After a subsequent second oxidation step with triplet oxygen, diverse acetalization steps occur, leading to the formation of artemisinin.[27] We wanted to pick up this impressive demonstration of sophisticated synthetic biology merged with clever downstream conversions of the fermentation product to synthesize a valuable drug. Our approach would use plasmids, designed by Keasling and co-workers, which encode for the required enzymes to establish the MVA pathway in E. coli[9,10] in combination with the FPP and amorphadiene synthase as well as CYP71AV1 to catalyze the crucial functionalization of amorphadiene to AA. We envisioned further increases in the product titers of AA to enable new perspectives for the production of semisynthetic artemisinin.
Figure 11: Synthesis of artemisinin from artemisinic acid.
As described above, the terpene α-humulene is the precursor for the sesquiterpene ketone zerumbone. It was for the first time described after the isolation from the essential oils of the rhizomes of Zingiber zerumbet in 1956 and can be found in various ginger species and other subtropical plants. Zerumbone exhibits manifold biological activities, beneath anti-inflammatory, antioxidant and immunoregulatory activities. However, most attention comes to zerumbone because of the discussed anticancer properties. It is reported that zerumbone can prevent cancer as well as inhibit the proliferation of various cancer cell lines by lowering the glutathione concentration through Michael reactions, leading to an altered redox potential within the cells. Normal cell lines are not affected.[28,29,30] Due to this interesting medical properties and since we can employ the same plasmids for the MVA pathway and the synthesis of FPP as for the biosynthesis of artemisinic acid, we chose zerumbone as another target molecule for our cytochrome/reductase fusion approach.
Figure 12: Structures of α-humulene and zerumbone.
Another sesquiterpene lactone which attracted our attention is parthenolide. Within the sesquiterpene lactones, this belongs to the subgroup of germacranolides and can be isolated from the aerial parts of the fewerfew plant (Tanacetum parthenium). Like zerumbone, it exhibits a broad spectrum of biological activities. For example, applications as a migraine prophylaxis agent and for treatment of cancer were discussed in the literature.[31] From a biosynthetic point of view, parthenolide is derived from the terpene germacrene A, which itself is a cyclization product of FPP by the corresponding terpene synthase. The biosynthesis of parthenolide occurs by several CYP P450 enzymes, which makes this biosynthesis particularly interesting for our approach. First, germacrene-12-ol is synthesized by germacrene A hydroxylase from germacrene A. Subsequently, germacrene-12-ol is converted into the corresponding carboxylic acid by dehydrogenases. After further functionalization and esterification by costunolide synthase, another cytochrome, the corresponding double bond is epoxidized by parthenolide synthesase, the third cytochrome in this biosynthetic sequence. Since costunolide, the biosynthetic precursor of the germacranolide parthenolide, is also the precursor for various guainolides and eudesmanolides, this biosynthetic pathway could prove useful for the synthesis of a variety of biologically active molecules in later studies.[32,33,34]
Figure 13: Biosynthesis of parthenolide from FPP with germacrene A and costunolide as key intermediates. Costunolide is a biosynthetic intermediate for various sesquiterpene lactones such as germacranolides, guaianolides and eudesmanolides.
Still our initial ideas regarding the sustainable production of hydrogen in mind, which enables the green production of platform chemicals and which can also be used as a biofuel, we were interested to read about the CYP P450 OleT: This enzyme can decarboxylate a broad scope of carboxylic acids to the corresponding terminal alkenes, which presents important building-blocks for the petrochemical industry, for example to adjust the crystallinity of polymers such as polyethylene.[35,36] Terminal olefins are usually produced from petrochemical feedstocks and the substitution of those by fatty acids to synthesize terminal olefins seems to be promising. Although the fusion of OleT with a CPR is already described in literature,[37] we wanted to include OleT in our toolbox for designing CYP P450/CPR fusion proteins to demonstrate the general applicability of this approach also outside of terpene biosynthesis and to further push the possible applications of OleT in synthetic and industrial chemistry.
Figure 14: Decarboxylation of carboxylic acids by OleT.
At the end of our thoughts and inspirations about redox active fusion proteins, we came back to the idea to employ the PSI of cyanobacteria as an electron source: When CYP P450 enzymes are merged with the electron carrier Fd, it should be possible to use the Fd as some kind of linker between the cytochrome and PSI to harness electrons directly from the thylakoid membranes.[13] This could enable a more precise control of the flow of reduction equivalents so that the desired reaction does not directly compete with other metabolic processes which consume NADH, to increase the effectiveness of fermentation processes in cyanobacteria. The use of phototrophic organisms for chemical production applications appears to be particularly promising and sustainable, as they can produce the desired products like terpenes directly from light and atmospheric carbon dioxide. If Fd is used as a mediator between a CYP P450 and the PSI, it may be possible to circumvent the challenges which appear by creating mutants of cyanobacteria with a fusion construct of the PSI and the desired CYP. Since the construction of such PSI fusion proteins interferes with the primary metabolism of the cyanobacteria, this interference dramatically slow down or inhibits the growth of the cyanobacteria - a problem, with which Gutekunst and co-workers were faced to when they developed the hydrogenase/PSI fusion protein.[1] Based on the activity of the promoters used, the competition between the CYP P450/Fd fusion protein and the natural Fd, which provides reduction equivalents for the usual metabolic processes, could be accurately adjusted.
Figure 15: PSI with membrane and CYP P450/Fd fusion.
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