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Review

Engineering Electron Transfer Pathway of Cytochrome P450s

by
Jingting He
1,
Xin Liu
2,3,* and
Chun Li
2,3,4,*
1
Key Laboratory for Green Processing of Chemical Engineering of Xinjiang Bingtuan, School of Chemistry and Chemical Engineering, Shihezi 832003, China
2
Department of Chemical Engineering, Tsinghua University, Beijing 100084, China
3
Key Lab for Industrial Biocatalysis, Ministry of Education, Tsinghua University, Beijing 100084, China
4
Center for Synthetic and Systems Biology, Tsinghua University, Beijing 100084, China
*
Authors to whom correspondence should be addressed.
Molecules 2024, 29(11), 2480; https://doi.org/10.3390/molecules29112480
Submission received: 15 April 2024 / Revised: 20 May 2024 / Accepted: 21 May 2024 / Published: 24 May 2024
(This article belongs to the Special Issue Cytochrome P450)
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Abstract

:
Cytochrome P450s (P450s), a superfamily of heme-containing enzymes, existed in animals, plants, and microorganisms. P450s can catalyze various regional and stereoselective oxidation reactions, which are widely used in natural product biosynthesis, drug metabolism, and biotechnology. In a typical catalytic cycle, P450s use redox proteins or domains to mediate electron transfer from NAD(P)H to heme iron. Therefore, the main factors determining the catalytic efficiency of P450s include not only the P450s themselves but also their redox-partners and electron transfer pathways. In this review, the electron transfer pathway engineering strategies of the P450s catalytic system are reviewed from four aspects: cofactor regeneration, selection of redox-partners, P450s and redox-partner engineering, and electrochemically or photochemically driven electron transfer.

1. Introduction

Cytochrome P450s (P450s) are a superfamily of heme-containing enzymes named for their absorption band at 450 nm in the form of a carbon monoxide-bound complex and widely distributed in kingdoms of life. As the most versatile biocatalyst, P450s can catalyze C-H hydroxylation, C=C double bond epoxidation, N-, and S-oxidation, O-, N- and S-dealkylation, aromatic coupling, C-C bond breaking, and many other common and uncommon reactions [1,2,3,4,5,6]. P450s have highly regio-selective and stereospecific oxidation activity against a variety of substrates and are widely involved in the metabolism of various exogenous substances such as environmental compounds, antibiotics, and drugs [7], as well as in the biosynthesis pathways of endogenous molecules such as sterols and fatty acids [8,9,10,11]. Therefore, P450s play an important role in drug development [12], natural product biosynthesis, and biotechnology [13,14,15,16].
The typical catalytic system of P450 is an electron transfer system composed of the P450 enzyme, substrate, electron shuttle carrier (cytochrome P450 redox-partners, CRPs), electron donor (NAD(P)H), and oxidizer (O2). However, the large-scale application of P450s is limited by the poor stability of enzymes and the requirement for redox-partners or expensive cofactors. P450s and their redox-partners need to be organized in the right stoichiometry and orientation to ensure efficient electron transfer and support the P450s catalytic process. With more and more research on the structure and catalytic system of the P450 enzyme, P450s have been heterologously expressed in animal, plant, and microbial host cells, such as in the baculovirus-insect cell expression system [17], the tobacco osmotic transient expression system [18,19], and the prokaryotic/eukaryotic microbial expression system. Due to the unknown and complex nature, of the whole cell P450 catalytic system, many researchers have realized the catalysis of P450 by designing electron transfer pathways in vitro.
Both in vivo and in vitro catalytic systems have been limited by several technical bottlenecks, including low heterologous expression enzyme activity, the need for electron transfer partners, and the expensive cofactor NAD(P)H [20]. To solve these problems, strategies such as P450s mining and screening, host cell screening, redox-partner engineering, and NAD(P)H regeneration have been proposed. Herein, we reviewed the application of electron transfer engineering strategies in recent years from four aspects: cofactor regeneration, selection of redox-partners, P450 and CRP engineering, and electrochemically/photochemically driven electron transfer engineering.

2. Cytochrome P450s

The highly diverse P450s superfamily (with an amino acid sequence identity as low as 16% [21]) maintains a highly conserved 3D structural fold in which cysteine is fully conserved as the fifth (axial) ligand of heme iron and a catalytic cycle that is used together for O2 isomerization cleavage and the formation of highly active substances [22,23]. In most instances, the catalytic cycle of the P450s is based on the cleavage of dioxygen, resulting in the hydroxylation of the substrate and the formation of water [24]. Electrons are transferred from NAD(P)H to iron atoms in the heme groups of P450s by redox-partners [5].

2.1. Classification of P450s Systems

The P450s can be divided into ten categories [25,26], as shown in Figure 1, according to the redox-partner protein required by P450s in the catalytic reaction process. Classes I commonly found in bacteria and mitochondria consist of a P450 enzyme, a ferredoxin reductase redox-partner (FdR; cofactor flavin adenine dinucleotide (FAD)), and a small iron-sulfur protein-containing ferredoxin (Fdx). Classes II, present in eukaryotes, consist of a P450 enzyme and a membrane-bound cytochrome P450 reductase (CPR) as redox-partner, containing both FAD and flavin mononucleotide (FMN). Classes III: found in bacteria, the Fdx is replaced by a flavodoxin (Fld) containing FMN. Classes IV were first discovered in Sulfolobus solfataricus, where the FdR is replaced by 2-oxo-acid-ferredoxin oxidoreductase (OFOR). Classes V, first discovered in Methylococcus capsulatus, are fused with their redox-partners, Fdx. Classes VI, first discovered in Rhodococcus rhodochrous, are fused with their redox-partners, Fld. Class VII, first discovered in Rhodococcus, consists of an N-terminal P450 heme domain and a C-terminal reductase domain, consisting of an FMN-binding domain, an NADH-binding domain, and a [2Fe-2S] ferredoxin domain. Class VIII enzymes, first discovered in Bacillus megaterium, are self-sufficient enzymes, that contain binding sites for heme fused with CRP through a linker. Class IX is the only water-soluble eukaryotic P450 enzyme and can directly use NAD(P)H as an electron donor. Class X is independent of oxygen, NAD(P)H, and CRP however, uses acyl hydroperoxides as both substrate and oxygen donor.

2.2. P450 Catalytic Mechanism

In order to achieve the reductive activation of the inert O2 to the substrate mono-oxygenation, the vast majority of P450s interact with CRPs to obtain a reduction equivalent from NAD(P)H. In most instances, the NAD(P)H→CRP(s)→P450→Substrate electron transfer (ET) system is formed [27].
First, the substrate enters the reaction pocket to replace the bound water molecule at the active site, FeIII-OH2, interacting with FeIII and binding the C-H bond to form the high-spin FeIII-RH. The CRP then transfers one electron from NAD(P)H to FeIII-RH, forming FeII-RH. After that, it combines with molecular oxygen to form FeII-O2, receives a second electron transferred from CRP, and produces a FeIII-OOH complex by protonation of protons obtained from the solution. The O-O bond of FeIII-OOH is then broken by a second proton from the solvent, releasing an H2O molecule to form the high-priced FeⅣ=O, a cation that uses a radical mechanism to insert oxygen atoms into the substrate R-H bond to form FeⅣ-OH. The substrate free radical dissociates from iron to produce the product, and a molecule of water returns to the FeIII coordination, as shown in Figure 2.
Most P450s share a common monooxygenation mechanism, as mentioned above, which depends on NAD(P)H and CRPs. However, a few P450s (such as CYP152 peroxygenases) have evolved to directly utilize H2O2 as the sole oxygen and electron donor, forming FeIII-OOH complexes that catalyze fatty acids to hydroxylation products via the peroxide shunt pathway (dashed arrow in Figure 2) [28,29]. However, most P450s have a low H2O2 tolerance, which greatly limits their H2O2 shunt pathway [30]. Therefore, many researchers have made an effort to modify P450 monooxygenases into their peroxizyme by directed evolution or site-directed mutagenesis to simplify the common electron transfer chain and expand P450’s practical application [31,32]. Fan et al. developed a dual-functional small molecule (DFSM) strategy in which one end of DFSM binds to the P450BM3 protein with the anchoring group and the other end binds to the imidazolyl group. DFSM promoted the O-O cleavage of the adduct FeIII-OOH, which is beneficial to the formation of FeⅣ=O of the active species, thus converting P450BM3 monooxygenase into peroxygenase [33]. Podgorski et al. modified selected residues within the I-helix of CYP199A4 to more closely resemble those of a natural peroxygenase and obtained a better mutant that functioned better at lower H2O2 concentrations and kept regio- and stereoselective hydroxylation ability [34].

3. Electron Transfer Pathway Engineering

In general, a functional P450 system requires a redox-partner to transfer electrons from NAD(P)H to heme [35]. Therefore, in order to improve the catalytic efficiency of P450s, optimizing or reshaping the electron transfer pathway is an important approach, including the supply of electrons, the selection of CRPs, and enzyme engineering (Figure 3).

3.1. Cofactor Regeneration

In natural biocatalytic processes, cytochrome P450 enzymes catalyze a variety of oxidative transformations using the pyridine nucleotide NADH or NADPH as a cofactor [36]. However, due to the relatively low intracellular NAD(P)H concentration and the relatively expensive exogenous addition, the catalytic rate and application of P450s are greatly limited. Regeneration of NAD(P)H requires the transfer of one proton and two electrons (from the sacrificed hydride donor, such as glucose, formate, phosphite, or triethanolamine (TEOA)) to NAD(P)+. In general, the regeneration of cofactor NAD(P)H can be carried out by enzymatic, chemical, photocatalytic, and electrochemical methods [37,38]. The method of using enzymes to regenerate NAD(P)H is considered a favorable system due to its compatible reaction conditions, high selectivity, and high efficiency, and is widely used in practical industrial production [39].
The most widely used enzymes for cofactor regeneration in commercial processes are formate dehydrogenase (FDH) and glucose dehydrogenase (GDH), whereas phosphite (PDH), alcohol (ADH), and glucose 6-phosphate dehydrogenases (G6PDH) have been tested at laboratory scale [40,41]. Ma et al. [42] mutated formate dehydrogenase from Candida dubliniensis, and the catalytic efficiency of the mutant CdFDH-M4 was improved 75-fold, with total cofactor turnover numbers (TTN) ranging from 135 to 986. Xu et al. [43] mutated the alcohol dehydrogenase from Clostridium beijerinckii to promote the NADPH coenzyme cycle, so that the conversion rate of the LkADH and CbADH reactions reached 100% within 6 h. Xiong et al. [44] obtained a self-contained NADPH cofactor regeneration system when ethanol dehydrogenase (ADH) and cyclohexanone mono-oxygenase (CHMO) were co-expressed in E. coli, and the maximum yield of ε-caprolactone was 0.80 mol/mol when the ADH/CHMO ratio was 0.34. Zhang et al. [45] conjured thioredoxin1 (Trx1) and thioredoxin reductase (TR) from Thermus thermophilus to NADPH-dependent oxidoreductases (alcohol dehydrogenase and cyclohexanone monooxygenase) to achieve NADPH regeneration.
Because the natural cofactor NAD(P)H is widely involved in various metabolic reactions in the cell, a change in its concentration in the cell will cause disturbances to cell life activities [46]. Therefore, unnatural cofactors such as nicotinamide cytidine dinucleotides (NCD) [47] and nicotinamide mononucleotides (NMN) were designed to be coupled to oxidoreductases [48].

3.2. Selection of Redox-Partner

Due to the difference in structure and catalytic system of P450s from different sources, their heterologous expression effect in microbial cells is completely different. In most prokaryotic microorganisms, the P450s catalytic system mediates the conformational change of ferredoxin (Fdx) through electrostatic interactions between Fdx, P450s heme, and reductase domains, thereby transferring electrons to P450s [49,50]. In eukaryotes, p450 is usually localized to the membrane structure via a terminal transmembrane domain, and electrons are transferred to the P450 heme domain via CPR [26,51,52]. Many P450s recombinants have been used in E. coli to synthesize a variety of chemicals. At the same time, yeast cells are generally considered to be the preferred system for heterologous expression of membrane-binding proteins due to their endoplasmic reticulum. The redox-partner, an essential component of most P450s systems, transfers two electrons from NAD(P)H sequentially to the heme-iron reaction center, activating oxygen to insert the C-H bond, a rate-limiting step in the catalytic cycle. However, little information is available about the homologous redox-partner of P450s [53], which has greatly limited the functional exploration and practical application of P450s [54]. Therefore, exploring and trying different redox-partners to construct various hybrid P450 electron transfer systems is the main strategy to improve the catalytic efficiency of P450. Prokaryotic microbial systems do not encode CRPs, so exogenous introduction of CRP is required to re-establish the function of P450s. Giang et al. [55] reconstructed the CYP108N12 catalytic system using cymredosin, a [2Fe-2S] redox chaperone of P450, to increase electron transfer rate (from 13 ± 2 to 70 ± 1 μM NADH/min/μM CYP108N12) and NADH utilization efficiency (so-called coupling efficiency from 13% to 90%).
Although eukaryotic microorganisms have endogenous CRPs, exogenous CRPs are often introduced to maximize the catalytic effect of P450s. For example, pairing CFS with plant-derived ATR1 increased yeast plant-derived nicotine production by more than 50 times compared to yeast endogenous CRP [56]. Therefore, exogenous CRPs are often introduced to improve the catalytic efficiency of P450s. Liao et al. [57] found that heterogeneously expressed recombinant protein MaCPR2 from mulberry leaves in yeast can use NADPH as an electron donor to reduce cytochrome c and ferricyanide and catalyze the formation of chlorogenic acid. Wang et al. [58] showed that after the integration of Glycyrrhiza uralensis Cb5 in the S. cerevisiae strain, which integrated CYP88D6, CYP72A154, β-amyrin synthetase, and the NADPH-cytochrome P450 reductase POR1 gene of Arabidopsis thaliana, the triterpenoid acid titer was increased by eight times. Pairing with different CRPs often also makes a substantial difference in P450s activity [56,59,60]. The Fe2S2 Fdxs have been found to be more likely to provide high electron transfer efficiency for bacterial Class I P450s [61]. Liu et al. [54] compared three commonly used alternative redox-partners, SelFdx1499/SelFdR0978, Adx/AdR, and Pdx/PdR. SelFdx1499/SelFdR0978 has a higher electron transfer efficiency when interacting with P450 monooxygenase PikC, and their catalytic efficiency for YC-17 was the highest, reaching 99.1%. Istiandari et al. [62] found that mutation of the CRP gene LjCPR1 in Lotus japonicus had no effect on product yield, while deletion of the CRP gene LjCPR2-1 significantly reduced soybean saponin precursor yield. When constructing the P450s catalytic system, it is necessary to consider not only their species in addition to, their expression quantity. Excess CRP activity is toxic to microbial cells because it causes the overproduction of reactive oxygen species. Some studies have shown that the optimal ratio of P450s to CRP is about 12:1 [63]. Therefore, adjusting the expression levels of P450s and CRP appears to be critical for achieving higher P450s catalytic efficiency [64,65,66].
Of course, in the microbial production process, in addition to the P450s catalytic system that requires heterologous expression of rAedox-partners, there are also some self-sufficient P450s (Class VIII in Figure 1) that do not require additional redox-partners [67], in which the P450 heme domain is covalently bound to the reduction domain [68]. Such as the CYP116B subfamily [69], P450 BM3 [70], and P450 AZC1 [71]. The P450-CRP fusion arrangement promotes rapid electron transfer from NADPH to P450 heme. Due to the proximity and specific localization of their domains and the need for no additional redox-partners, they have excellent electron transfer systems [53]. Cha et al. [72] explored the optimal synergistic ratio of (+)-nootkatone production by considering the oxidation of membrane-anchored cytochrome P450/P450 reductase system (HPO/AtCPR) by establishing high-throughput screening technology and metabolic engineering, and the adjustment of the ratio of HPO/AtCPR increased the yield of (+)-nootkatone by two times in Saccharomyces cerevisiae.

3.3. P450s and CRP Engineering

In the catalytic process of P450s, electrons from cofactors need to be transferred to the P450s heme domain through CRP. In order to improve the electron transfer rate of this step, the researchers engineered P450s and CRP from the protein structure (Figure 4).

3.3.1. Key Amino Acid Mutation

P450s-mediated metabolism through proteins depends on the electron transfer chain—the interaction of a protein with its primary CRP. Traditionally, the P450s heme domain and CRP surface amino acid residues have been considered the main determinants of proper alignment in this interaction, in which hydrophobic interactions are also involved [73]. At present, many programs that can predict protein structure and calculate the protein-protein interaction have been designed for protein modification, such as AlphaFold2 (AF2), molecular dynamics (MD) simulation, QM/MM calculations, and so on. Cytochrome P450BM3 (CYP102A1) is a type VIII self-contained flavin hemoglobin with a heme and flavin domain in a polypeptide chain [74] that catalyzes subterminal oxidation of saturated and unsaturated fatty acids. Ivanov et al. [75] used AlphaFold2 and AlphaFold polymer (AFMultimer) programs to predict the three-dimensional structure of CYP102A1 (wild-type) protein monomer and homologous dimer and found that the homologous dimer F262 aromatic group is closer to heme. It increases the speed of electron transfer. Meng et al. [76] used molecular dynamics simulation and electron jump analysis of P450 BM3 mutants from Bacillus megaterium and showed that substituting the amino acid residues connecting FMN and heme iron with aromatic amino acids could increase the electron transfer rate and the catalytic effect by 13.9 times. Mutations of cytochrome P450 BM3, such as P450BM3 M7 mutants and P450BM3 M9 mutants, have improved electron transfer [77]. Velazquez et al. [78] expressed POR (OMIM:*124015, HNGC:9208) wild type and P228L variant in bacteria, and based on computer and in vitro studies, predicted that the change of proline to leucine might change the rigidity of protein, change the conformation of POR, and lead to a decrease in the rate of electron transfer to cytochrome c and thiazole blue tetrazole (MTT) significantly. Yu et al. [79] used a colorimetric high-throughput screening (HTS) system with DDA as the true substrate to conduct directed evolution and obtained a P450 mutant (R14R/D629G) with higher activity. Molecular docking analysis, kinetic parameter determination, and protein electrophoresis showed that the decrease in electron transfer distance between FMN and FAD caused by the D629G mutation is the main reason for the increased activity.

3.3.2. Protein Fusion

CRPs, which are often fused to the C-terminal of P450s [80] to enhance their interactions and shorten the electron transfer chain, have been successfully applied in many model strains [81,82,83,84]. When constructing P450s-redox-partner fusion protein, it is important to consider the length of the linker, the amino acid content, and the position relative to P450s and CRP protein. Ligand length can affect the orientation, folding, correct conformation, and flexibility of enzymes [82]. Proline-rich rigid ligands [85,86] and glycine-rich flexible ligands [84,87] have been used more frequently to better control the spatial arrangement of enzymes and the structural isolation of fusion domains from surrounding amino acids and redox-partners, improving electron transfer [16,88]. Bakkes et al. [35] reported that the fusion efficiency of P450s and CRP with glycine-rich linkers was the highest (81.2%). Li et al. [89] used the GSTSSG linker to fuse CYP716A12 with ATR1 with N-terminal truncation, which increased the triterpene oleanolic acid (OA) yield to 129.9 mg/L.

3.3.3. Enzyme Immobilization

Alternative techniques such as immobilization help to support the redox activity of P450 [81]. Park et al. [90] reported an electron channel strategy based on the application of CipB stent-protein, which allows efficient electron transfer between P450 and reductase by bringing these enzymes close together. The final lutein strain produced 218.0 mg/L of lutein in fed-batch fermentation. Wang et al. [91] constructed a protein scaffold multi-enzyme assembly to improve the coupling efficiency of the P450 system for efficient biosynthesis of daidzein from (2S)-naringenin.

3.4. Electro/Photo-Chemically Driven Electron Transfer Engineering

Despite the great potential of P450s, their dependence on NAD(P)H and redox-partners has limited their application in industry. In industrial processes, the production of NAD(P)H in cells is limited, exogenous addition is very expensive, and suitable redox-partners are difficult to find [92]. In order to overcome the dependence of cofactor NAD(P)H and redox-partner chaperones in the catalytic pathway of P450s, other approaches have been tried, such as electrochemical and photochemical methods on the P450s catalytic system [93,94]. In principle, the photochemical/electrochemical drive allows the conversion of luminous or electrical energy into chemical energy in the form of electron carriers, such as reduction cofactors, free radicals, or redox-partners, which are then used to drive the P450 catalytic reactions [95,96,97].

3.4.1. Electrochemically Driven Electron Transfer Engineering

Since the catalytic cycle of P450s involves electron transfer processes [74,98], electrochemically driven P450s catalytic systems are considered to be a simple, clean, and pollution-free alternative to complex biological systems.
There are two main ways of electrochemically catalyzing P450s. One is to fix the enzyme on the surface of the electrode by surface modification. This way increases the electron coupling by increasing the contact area between the enzyme and the electrode surface, so that the electrons are transferred directly to the active center of the P450s [99,100,101,102,103,104], where redox-partners and the cofactor NAD(P)H are not necessary. Or transfer to the P450s active center via redox chaperones in fusion proteins. Castrignano et al. combined the heme domains of bacterial CYP3A4 and CYP116B5 with the reductase domain of BM3 to construct an efficient electron transfer chain [105]. Some studies have shown that the electrode surface characteristics, the porous structure of nanomaterials, and environmental conditions may affect the electron transfer efficiency [106,107,108]. These factors can control the orientation of P450s on the electrode, the electron loss of adverse reactions, and the amount of ROS, thus changing the efficiency of the electron coupling between heme and the electrode [109,110,111]. The most commonly used electrode surface modifications for the electrochemical catalysis of P450s include lipid film (didodecyldimethylammonium bromide (DDAB), polydienyldimethylammonium chloride, polydienyldimethylammonium bromide, polysodium p-styrene sulfonate), sol-gel methyltriethoxysilane of alkylmercaptan self-assembled monolayer (SAM), and composite materials based on various nanomaterials (gold nanoparticles, carbon nanomaterials), conductive polymers (polypyrrole, polythiophene, polyaniline, mercaptocarboxylic acid), etc. Electrochemical methods have high potential to investigate the different functions of P450s, including the study of the complex catalytic mechanisms of cytochrome P450s [103], as well as the design and development of new substrates, inhibitors, and modulators [112,113]. It has a promising application prospect in drug sensing devices, new drug search tools, and bioreactors [101]. Different P450s electrochemical catalytic systems will produce different catalytic effects, as shown in Table 1.
The other is to construct a P450 bioelectrocatalytic system (BES) [130], which can synergistically utilize electrons from electrodes and carbon source oxidation to achieve intracellular NAD(P)H regeneration and glucose oxidation through an electron shuttle carrier-mediated extracellular electron transfer (EET) pathway to promote P450 enzymatically catalyzed reactions. Common electron shuttles include riboflavin [131], methylviologen (MV) [132], and neutral red (NR) [130]. In BES with NR as electron shuttle carrier, the 7α-OH-DHEA yield catalyzed by CYP7B1 in Saccharomyces cerevisiae reached 288.6 ± 7.8 mg L−1, 2.4 times higher than that without EET (122.1 ± 3.7 mg L−1) [133].

3.4.2. Photochemically Driven Electron Transfer Engineering

In the process of photochemically driven P450s catalysis, the photosensitizer and protein form a complex through covalent or non-covalent bonding using photogenerated electrons or electrons produced by the electron donor to drive the P450s-catalyzed reaction. Photosensitizers induce the direct or indirect activation of redox enzymes by photoinduced electron transfer, and light promotes electrons into higher energy states, while the holes vacated by excited electrons are filled at the expense of oxidation of the electron donor, such as water, triethanolamine (TEOA), or ethylenediaminetetraaceticacid (EDTA). Conduction band or triplet/singlet photoexcited electrons transfer to the redox centers of enzymes, such as P450 heme, flavin, and metal clusters. There are three main methods of light-driven electron transfer: (1) Active oxygen is generated in situ, and then P450 is catalyzed by the peroxide shingle pathway; (2) Redox-partners transfer electrons to the heme domain; and (3) Electrons are transferred directly to the heme domain [134]. Photosensitizers catalyze the redox conversion of the substrate by exchanging electrons at the active site of the enzyme [135,136]. The commonly used photosensitizers are protein photosensitizers (PS-I, PS-II, PSP), organic photosensitizers (porphyrin and Ru-pyridine) [137], and semiconductors (TiO2, carbon dot, CdS, g-C3N4, etc.) [138].
Girhard et al. [139] photoexcited flavin (riboflavin, flavin mononucleotide, or flavin adenine dinucleotide) with EDTA as an electron donor, and a photochemically driven CYP152A1 system to produce hydrogen peroxide in situ. Kim et al. [140] firstly used photosynthesis-derived electrons to immobilize P450-containing microsomes in spinach chloroplasts for light-driven bioreactors, and the chloroplasts of spinach were used for photosynthesis to produce NADPH for the oxidation of 7-ethyl coumarin (7-EC) to 7-hydroxycoumarin (7-HC) catalyzed by the CYP1A1-CRP fusion enzyme. Wang et al. [141] can effectively degrade amaranth by directly transferring photogenerated electrons generated by CdS to P450 through the electron transfer chain of Shewanella under the light-driven anaerobic condition. Lee et al. [95] used eosin Y as a photosensitive dye, TEOA as an electron donor, and [Cp*Rh (bpy) H2O] as an electron medium to drive NADPH regeneration and achieve O-dealkylation catalyzed by P450 BM3 variants (Y51F/F87A, BM3M2). Liu et al. [142] regenerated NADPH by combining engineered Bacillus auriculare with InP nanoparticles, which increased NADPH levels by 84 times.
Until Jensen recombinant PSI and purified CYP79A1 performed in vitro studies, the system relied on Fdx to transfer electrons from PSI to CYP79A1 without the involvement of NADPH and CRP, thus simplifying the previously proposed electron transfer pathway [96]. Natural flavin, as a photosensitizer, is often used to construct vision-driven whole cell platforms for P450s photocatalysis. The general applicability of light-driven flavin-mediated P450s photocatalysis systems was demonstrated in whole cells containing CYP1A1 [143], CYP1A2, CYP1B1, CYP2B4, and CYP3A4 [144,145,146]. In CdTe quantum dots (QDs) prepared by a covalent combination of CYP2D6 and QDs, electrons are transferred from the conduction band (CB) of CdTe quantum dots to CYP2D6 as a photocatalyst for drug metabolism [147]. When CYP119 is coupled with CeO2-3TiO2 semiconductor composite material, the coupling system shows higher photocatalytic reactivity than the uncoupled system under light [148]. In the hybrid P450 BM3 enzyme, the covalent linking of Ru (II)-diimide photosensitizer non-natural single cysteine residues enables rapid electron injection into the heme domain of several P450s for P450 oxidative functionalization on the trifluoromethylation substrate under visible light activation [149].

4. Conclusions and Future Perspectives

So far, many strategies have been developed to optimize the electron transfer process in P450s catalytic systems to improve the catalytic efficiency of P450s, and calculations such as molecular dynamics (MD) and quantum mechanics/molecular mechanics (QM/MM) are used to elucidate the electron transfer mechanism of P450s [150]. These include an adequate supply of electrons for the catalytic system, the selection and modification of P450s and CRP through rational/semi-rational design and directed evolution, the construction of photocatalysis and electrocatalysis systems, etc. Light as a free, green, and pollution-free energy, coupling P450s with the photosynthetic machinery of a plant, can use photogenerated electrons to drive the catalytic reaction [151], thus reducing the catalytic cost and improving the catalytic efficiency. With the pursuit of green, sustainable processes, the application of light or electricity to drive P450 catalysis may become mainstream. Combined with the advantages of biocatalysis and photoelectrocatalysis, the use of green energy for selective conversion has many advantages, including novel reactivity, high enantioselectivity, green synthesis, and high yield, which provide the basis for the application and development of P450s as bioreactors and biosensors [152]. Photo/electrically induced electron or energy transfer enables the synthesis method to complement the traditional dual electron transfer process for P450s or provide an orthogonal pathway for the development of new reactions. Enzymes can be regulated by directed evolution to exert control over intermediates, thereby inhibiting adverse reactions and providing high chemical and stereoselectivity. These strategies are combined to achieve green and efficient substance synthesis. In the future, the stability of P450s in vitro systems, the compatibility of CRP, electron transfer, and utilization efficiency will be the focus of P450s catalysis research.

Author Contributions

J.H.: Conceptualization, Methodology, Investigation, Writing—original draft, Writing—review and editing. X.L.: Conceptualization, Methodology, Writing—review and editing. C.L.: Funding acquisition, Project administration, Supervision, Writing—original draft, Writing—review, and editing. All authors have read and agreed to the published version of the manuscript.

Funding

This research was supported by National Key R&D Program of China (2023YFA0913600), China Postdoctoral Science Foundation (2022M711792, 2022TQ0172), and National Natural Science Foundation of China (22138006).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

No new data were created or analyzed in this study. Data sharing is not applicable to this article.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Nelson, D.R.; Kamataki, T.; Waxman, D.J.; Guengerich, F.P. The P450 Superfamily: Update on New Sequences, Gene Mapping, Accession Numbers, Early Trivial Names of Enzymes, and Nomenclature. DNA Cell Biol. 1993, 12, 1–51. [Google Scholar] [CrossRef] [PubMed]
  2. Guengerich, F.P. Common and uncommon cytochrome P450 reactions related to metabolism and chemical toxicity. Chem. Res. Toxicol. 2001, 14, 611–650. [Google Scholar] [CrossRef] [PubMed]
  3. Zhang, X.; Li, S. Expansion of chemical space for natural products by uncommon P450 reactions. Nat. Prod. Rep. 2017, 34, 1061–1089. [Google Scholar] [CrossRef] [PubMed]
  4. Schuler, M.A.; Werck-Reichhart, D. Functional genomics of P450s. Annu. Rev. Plant Biol. 2003, 54, 629–667. [Google Scholar] [CrossRef] [PubMed]
  5. Vermilion, J.L.; Ballou, D.P.; Massey, V.; Coon, M.J. Separate roles for FMN and FAD in catalysis by liver microsomal NADPH-cytochrome P-450 reductase. J. Biol. Chem. 1981, 256, 266–277. [Google Scholar] [CrossRef] [PubMed]
  6. Renault, H.; Bassard, J.E.; Hamberger, B.; Werck-Reichhart, D. Cytochrome P450-mediated metabolic engineering: Current progress and future challenges. Curr. Opin. Plant Biol. 2014, 19, 27–34. [Google Scholar] [CrossRef] [PubMed]
  7. Paddon, C.J.; Keasling, J.D. Semi-synthetic artemisinin: A model for the use of synthetic biology in pharmaceutical development. Nat. Rev. Microbiol. 2014, 12, 355–367. [Google Scholar] [CrossRef] [PubMed]
  8. Isin, E.M.; Guengerich, F.P. Substrate binding to cytochromes P450. Anal. Bioanal. Chem. 2008, 392, 1019–1030. [Google Scholar] [CrossRef] [PubMed]
  9. Srinivasan, P.; Smolke, C.D. Biosynthesis of medicinal tropane alkaloids in yeast. Nature 2020, 585, 614–619. [Google Scholar] [CrossRef]
  10. Brown, S.; Clastre, M.; Courdavault, V.; O’connor, S.E. De novo production of the plant-derived alkaloid strictosidine in yeast. Proc. Natl. Acad. Sci. USA 2015, 112, 3205–3210. [Google Scholar] [CrossRef]
  11. Ajikumar, P.K.; Xiao, W.H.; Tyo, K.E.J.; Wang, Y.; Simeon, F.; Leonard, E.; Mucha, O.; Phon, T.H.; Pfeifer, B.; Stephnopoulos, G. Isoprenoid Pathway Optimization for Taxol Precursor Overproduction in Escherichia coli. Science 2010, 330, 70–74. [Google Scholar] [CrossRef] [PubMed]
  12. Nakagawa, F. An epidemiological modelling study to estimate the composition of HIV-positive populations including migrants from endemic settings. AIDS 2017, 31, 417–425. [Google Scholar] [CrossRef] [PubMed]
  13. Cravens, A.; Payne, J.; Smolke, C.D. Synthetic biology strategies for microbial biosynthesis of plant natural products. Nat. Commun. 2019, 10, 2142. [Google Scholar] [CrossRef] [PubMed]
  14. Li, S.; Li, Y.; Smolke, C.D. Strategies for microbial synthesis of high-value phytochemicals. Nat. Chem. 2018, 10, 395–404. [Google Scholar] [CrossRef]
  15. Guengerich, F.P. Cytochrome P450 enzymes in the generation of commercial products. Nat. Rev. Drug Discov. 2002, 1, 359–366. [Google Scholar] [CrossRef] [PubMed]
  16. Bakkes, P.J.; Biemann, S.; Bokel, A.; Eickholt, M.; Girhard, M.; Urlacher, V.B. Design and improvement of artificial redox modules by molecular fusion of flavodoxin and flavodoxin reductase from Escherichia coli. Sci. Rep. 2015, 5, 12158. [Google Scholar] [CrossRef] [PubMed]
  17. Amarneh, B.; Simpson, E.R. Expression of a recombinant derivative of human aromatase P450 in insect cells utilizing the baculovirus vector system. Mol. Cell. Endocrinol. 1995, 109, R1–R5. [Google Scholar] [CrossRef] [PubMed]
  18. Reed, J.; Osbourn, A. Engineering terpenoid production through transient expression in Nicotiana benthamiana. Plant Cell Rep. 2018, 37, 1431–1441. [Google Scholar] [CrossRef] [PubMed]
  19. Reed, J.; Stephenson, M.J.; Miettinen, K.; Brouwer, B.; Leveau, A.; Brett, P.; Goss, R.J.M.; Goossens, A.; O’connell, M.A.; Osbourn, A. A translational synthetic biology platform for rapid access to gram-scale quantities of novel drug-like molecules. Metab. Eng. 2017, 42, 185–193. [Google Scholar] [CrossRef]
  20. Bernhardt, R.; Urlacher, V.B. Cytochromes P450 as promising catalysts for biotechnological application: Chances and limitations. Appl. Microbiol. Biotechnol. 2014, 98, 6185–6203. [Google Scholar] [CrossRef]
  21. Wang, H.; Napoli, K.L.; Strobel, H.W. Cytochrome P450 3A9 catalyzes the metabolism of progesterone and other steroid hormones. Mol. Cell. Biochem. 2000, 213, 127–135. [Google Scholar] [CrossRef] [PubMed]
  22. Sono, M.; Roach, M.P.; Coulter, E.D.; Dawson, J.H. Heme-Containing Oxygenases. Chem. Rev. 1996, 96, 2841–2888. [Google Scholar] [CrossRef]
  23. Denisov, I.G.; Makris, T.M.; Sligar, S.G. Structure and Chemistry of Cytochrome P450. ChemInform 2005, 36, 2253–2278. [Google Scholar] [CrossRef]
  24. Hannemann, F.; Bichet, A.; Ewen, K.M.; Bernhardt, R. Cytochrome P450 systems--biological variations of electron transport chains. Biochim. Biophys. Acta 2007, 1770, 330–344. [Google Scholar] [CrossRef] [PubMed]
  25. Li, S.; Du, L.; Bernhardt, R. Redox Partners: Function Modulators of Bacterial P450 Enzymes. Trends Microbiol. 2020, 28, 445–454. [Google Scholar] [CrossRef] [PubMed]
  26. Cook, D.J.; Finnigan, J.D.; Cook, K.; Black, G.W.; Charnock, S.J. Cytochromes P450: History, Classes, Catalytic Mechanism, and Industrial Application. Adv. Protein Chem. Struct. Biol. 2016, 105, 105–126. [Google Scholar] [PubMed]
  27. Guengerich, F.P. Rate-limiting steps in cytochrome P450 catalysis. Biol. Chem. 2002, 383, 1553–1564. [Google Scholar] [CrossRef] [PubMed]
  28. Jiang, Y.; Peng, W.; Li, Z.; You, C.; Zhao, Y.; Tang, D.; Wang, B.; Li, S. Unexpected Reactions of α,β-Unsaturated Fatty Acids Provide Insight into the Mechanisms of CYP152 Peroxygenases. Angew. Chem. Int. Ed. 2021, 60, 24694–24701. [Google Scholar] [CrossRef]
  29. Munro, A.W.; Mclean, K.J.; Grant, J.L.; Makris, T.M. Structure and function of the cytochrome P450 peroxygenase enzymes. Biochem. Soc. Trans. 2018, 46, 183–196. [Google Scholar] [CrossRef]
  30. Matthews, S.; Belcher, J.D.; Tee, K.L.; Girvan, H.M.; Mclean, K.J.; Rigby, S.E.; Levy, C.W.; Leys, D.; Parker, D.A.; Blankley, R.T.; et al. Catalytic Determinants of Alkene Production by the Cytochrome P450 Peroxygenase OleT(JE). J. Biol. Chem. 2017, 292, 5128–5143. [Google Scholar] [CrossRef]
  31. Wise, C.E.; Hsieh, C.H.; Poplin, N.L.; Makris, T.M. Dioxygen Activation by the Biofuel-Generating Cytochrome P450 OleT. ACS Catal. 2018, 8, 9342–9352. [Google Scholar] [CrossRef]
  32. Kumar, S.; Liu, H.; Halpert, J. Engineering of Cytochrome P 450 3 A 4 for Enhanced Peroxide-Mediated Substrate Oxidation Using Directed Evolution and Site-Directed Mutagenesis. Drug Metab. Dispos. 2006, 34, 1958–1965. [Google Scholar] [CrossRef] [PubMed]
  33. Fan, S.; Cong, Z. Emerging Strategies for Modifying Cytochrome P450 Monooxygenases into Peroxizymes. Acc. Chem. Res. 2024, 57, 613–624. [Google Scholar] [CrossRef]
  34. Podgorski, M.N.; Akter, J.; Churchman, L.R.; Bruning, J.B.; De Voss, J.J.; Bell, S.G. Engineering Peroxygenase Activity into Cytochrome P450 Monooxygenases through Modification of the Oxygen Binding Region. ACS Catal. 2024, 14, 7426–7443. [Google Scholar] [CrossRef]
  35. Bakkes, P.J.; Riehm, J.L.; Sagadin, T.; Ruhlmann, A.; Schubert, P.; Biemann, S.; Girhard, M.; Hutter, M.C.; Bernhardt, R.; Urlacher, V.B. Engineering of versatile redox partner fusions that support monooxygenase activity of functionally diverse cytochrome P450s. Sci. Rep. 2017, 7, 9570. [Google Scholar] [CrossRef] [PubMed]
  36. Guengerich, F.P. Mechanisms of Cytochrome P450-Catalyzed Oxidations. ACS Catal. 2018, 8, 10964–10976. [Google Scholar] [CrossRef] [PubMed]
  37. Wu, H.; Tian, C.; Song, X.; Liu, C.; Jiang, Z. Methods for the regeneration of nicotinamide coenzymes. Green. Chem. 2013, 15, 1773–1789. [Google Scholar] [CrossRef]
  38. Wittmann, C. Biosystems Engineering I: Creating Superior Biocatalysts; Springer: Berlin/Heidelberg, Germany, 2010. [Google Scholar]
  39. Wang, X.; Saba, T.; Yiu, H.H.P.; Howe, R.F.; Shi, J. Cofactor NAD(P)H Regeneration Inspired by Heterogeneous Pathways. Chem 2017, 2, 621–654. [Google Scholar] [CrossRef]
  40. Hollmann, F.; Arends, I.W.C.E.; Holtmann, D. Enzymatic reductions for the chemist. Green. Chem. 2011, 13, 2285–2314. [Google Scholar] [CrossRef]
  41. Ganguli, P.; Khanna, R.; Prickril, B. Technology Transfer in Biotechnology; John Wiley & Sons: Hoboken, NJ, USA, 2005. [Google Scholar]
  42. Ma, W.; Geng, Q.; Chen, C.; Zheng, Y.C.; Yu, H.L.; Xu, J.H. Engineering a Formate Dehydrogenase for NADPH Regeneration. Chembiochem 2023, 24, e202300390. [Google Scholar] [CrossRef]
  43. Xu, J.; Zhou, H.; Yu, H.; Deng, T.; Wang, Z.; Zhang, H.; Wu, J.; Yang, L. Computational design of highly stable and soluble alcohol dehydrogenase for NADPH regeneration. Bioresour. Bioprocess. 2021, 8, 12. [Google Scholar] [CrossRef] [PubMed]
  44. Xiong, J.; Chen, H.; Liu, R.; Yu, H.; Zhuo, M.; Zhou, T.; Li, S. Tuning a bi-enzymatic cascade reaction in Escherichia coli to facilitate NADPH regeneration for ε-caprolactone production. Bioresour. Bioprocess. 2021, 8, 32. [Google Scholar] [CrossRef] [PubMed]
  45. Zhang, N.; Müller, B.; Kirkeby, T.R.; Kara, S.; Loderer, C. Development of a Thioredoxin-Based Cofactor Regeneration System for NADPH-Dependent Oxidoreductases. ChemCatChem 2022, 14, e202101625. [Google Scholar] [CrossRef]
  46. Matsuoka, Y.; Kurata, H. Computer-Aided Rational Design of Efficient NADPH Production System by Escherichia coli pgi Mutant Using a Mixture of Glucose and Xylose. Front. Bioeng. Biotechnol. 2020, 8, 277. [Google Scholar] [CrossRef] [PubMed]
  47. Guo, X.; Liu, Y.; Wang, Q.; Wang, X.; Zhao, Z.K. Non-natural Cofactor and Formate Driven Reductive Carboxylation of Pyruvate. Angew. Chem. Int. Ed. 2019, 59, 3143–3146. [Google Scholar] [CrossRef] [PubMed]
  48. Black, W.B.; Zhang, L.; Mak, W.S.; Maxel, S.; Cui, Y.; King, E.; Fong, B.; Sanchez Martinez, A.; Siegel, J.B.; Li, H. Engineering a nicotinamide mononucleotide redox cofactor system for biocatalysis. Nat. Chem. Biol. 2020, 16, 87–94. [Google Scholar] [CrossRef] [PubMed]
  49. Chiliza, Z.E.; Martinez-Oyanedel, J.; Syed, K. An overview of the factors playing a role in cytochrome P450 monooxygenase and ferredoxin interactions. Biophys. Rev. 2020, 12, 1217–1222. [Google Scholar] [CrossRef] [PubMed]
  50. Wang, Z.; Shaik, S.; Wang, B. Conformational Motion of Ferredoxin Enables Efficient Electron Transfer to Heme in the Full-Length P450(TT). J. Am. Chem. Soc. 2021, 143, 1005–1016. [Google Scholar] [CrossRef] [PubMed]
  51. Mellor, S.B.; Vinde, M.H.; Nielsen, A.Z.; Hanke, G.T.; Abdiaziz, K.; Roessler, M.M.; Burow, M.; Motawia, M.S.; Moller, B.L.; Jensen, P.E. Defining optimal electron transfer partners for light-driven cytochrome P450 reactions. Metab. Eng. 2019, 55, 33–43. [Google Scholar] [CrossRef]
  52. Jensen, K.; Moller, B.L. Plant NADPH-cytochrome P450 oxidoreductases. Phytochemistry 2010, 71, 132–141. [Google Scholar] [CrossRef]
  53. Sabbadin, F.; Hyde, R.; Robin, A.; Hilgarth, E.M.; Delenne, M.; Flitsch, S.; Turner, N.; Grogan, G.; Bruce, N.C. LICRED: A versatile drop-in vector for rapid generation of redox-self-sufficient cytochrome P450s. Chembiochem 2010, 11, 987–994. [Google Scholar] [CrossRef]
  54. Liu, X.; Li, F.; Sun, T.; Guo, J.; Zhang, X.; Zheng, X.; Du, L.; Zhang, W.; Ma, L.; Li, S. Three pairs of surrogate redox partners comparison for Class I cytochrome P450 enzyme activity reconstitution. Commun. Biol. 2022, 5, 791. [Google Scholar] [CrossRef]
  55. Giang, P.D.; Churchman, L.R.; Stok, J.E.; Bell, S.G.; De Voss, J.J. Cymredoxin, a [2Fe-2S] ferredoxin, supports catalytic activity of the p-cymene oxidising P450 enzyme CYP108N12. Arch. Biochem. Biophys. 2023, 737, 109549. [Google Scholar] [CrossRef] [PubMed]
  56. Trenchard, I.J.; Smolke, C.D. Engineering strategies for the fermentative production of plant alkaloids in yeast. Metab. Eng. 2015, 30, 96–104. [Google Scholar] [CrossRef] [PubMed]
  57. Liao, Y.; Du, W.; Wan, J.; Fan, J.; Pi, J.; Wu, M.; Wei, Y. Mining and functional characterization of NADPH-cytochrome P450 reductases of the DNJ biosynthetic pathway in mulberry leaves. BMC Plant Biol. 2024, 24, 133. [Google Scholar] [CrossRef] [PubMed]
  58. Wang, C.; Su, X.; Sun, M.; Zhang, M.; Wu, J.; Xing, J.; Wang, Y.; Xue, J.; Liu, X.; Sun, W.; et al. Efficient production of glycyrrhetinic acid in metabolically engineered Saccharomyces cerevisiae via an integrated strategy. Microb. Cell Fact. 2019, 18, 95. [Google Scholar] [CrossRef]
  59. Zhu, M.; Wang, C.; Sun, W.; Zhou, A.; Wang, Y.; Zhang, G.; Zhou, X.; Huo, Y.; Li, C. Boosting 11-oxo-beta-amyrin and glycyrrhetinic acid synthesis in Saccharomyces cerevisiae via pairing novel oxidation and reduction system from legume plants. Metab. Eng. 2018, 45, 43–50. [Google Scholar] [CrossRef]
  60. Dai, Z.; Liu, Y.; Sun, Z.; Wang, D.; Qu, G.; Ma, X.; Fan, F.; Zhang, L.; Li, S.; Zhang, X. Identification of a novel cytochrome P450 enzyme that catalyzes the C-2alpha hydroxylation of pentacyclic triterpenoids and its application in yeast cell factories. Metab. Eng. 2019, 51, 70–78. [Google Scholar] [CrossRef]
  61. Zhang, W.; Du, L.; Li, F.; Zhang, X.; Qu, Z.; Han, L.; Li, Z.; Sun, J.; Qi, F.; Yao, Q. Mechanistic Insights into Interactions between Bacterial Class I P450 Enzymes and Redox Partners. ACS Catal. 2018, 8, 992–10003. [Google Scholar] [CrossRef]
  62. Istiandari, P.; Yasumoto, S.; Seki, H.; Fukushima, E.O.; Muranaka, T. Class I and II NADPH-cytochrome P450 reductases exhibit different roles in triterpenoid biosynthesis in Lotus japonicus. Front. Plant Sci. 2023, 14, 1214602. [Google Scholar] [CrossRef]
  63. Biggs, B.W.; Lim, C.G.; Sagliani, K.; Shankar, S.; Stephanopoulos, G.; De Mey, M.; Ajikumar, P.K. Overcoming heterologous protein interdependency to optimize P450-mediated Taxol precursor synthesis in Escherichia coli. Proc. Natl. Acad. Sci. USA 2016, 113, 3209–3214. [Google Scholar] [CrossRef]
  64. Li, Z.; Jiang, Y.; Guengerich, F.P.; Ma, L.; Li, S.; Zhang, W. Engineering cytochrome P450 enzyme systems for biomedical and biotechnological applications. J. Biol. Chem. 2020, 295, 833–849. [Google Scholar] [CrossRef]
  65. Xu, L.H.; Du, Y.L. Rational and semi-rational engineering of cytochrome P450s for biotechnological applications. Synth. Syst. Biotechnol. 2018, 3, 283–290. [Google Scholar] [CrossRef]
  66. Giuriato, D.; Correddu, D.; Catucci, G.; Di Nardo, G.; Bolchi, C.; Pallavicini, M.; Gilardi, G. Design of a H2O2-generating P450(SPalpha) fusion protein for high yield fatty acid conversion. Protein Sci. 2022, 31, e4501. [Google Scholar] [CrossRef] [PubMed]
  67. Zhang, S.T.; Li, T.; Deng, S.K.; Spain, J.C.; Zhou, N.Y. A cytochrome P450 system initiates 4-nitroanisole degradation in Rhodococcus sp. strain JS3073. J. Hazard. Mater. 2023, 458, 131886. [Google Scholar] [CrossRef] [PubMed]
  68. Kim, B.S.; Kim, S.Y.; Park, J.; Park, W.; Hwang, K.Y.; Yoon, Y.J.; Oh, W.K.; Kim, B.Y.; Ahn, J.S. Sequence-based screening for self-sufficient P450 monooxygenase from a metagenome library. J. Appl. Microbiol. 2007, 102, 1392–1400. [Google Scholar] [CrossRef] [PubMed]
  69. Correddu, D.; Di Nardo, G.; Gilardi, G. Self-Sufficient Class VII Cytochromes P450: From Full-Length Structure to Synthetic Biology Applications. Trends Biotechnol. 2021, 39, 1184–1207. [Google Scholar] [CrossRef]
  70. Munro, A.W.; Leys, D.G.; Mclean, K.J.; Marshall, K.R.; Ost, T.W.; Daff, S.; Miles, C.S.; Chapman, S.K.; Lysek, D.A.; Moser, C.C.; et al. P450 BM3: The very model of a modern flavocytochrome. Trends Biochem. Sci. 2002, 27, 250–257. [Google Scholar] [CrossRef]
  71. Yildirim, D.; Ozic, C.; Ensari, Y. Expression and Characterization of a New Self-Sufficient P450 Monooxygenase (P450 AZC1) from Azorhizobium caulinodans. Chembiochem 2023, 24, e202300065. [Google Scholar] [CrossRef]
  72. Cha, Y.; Li, W.; Wu, T.; You, X.; Chen, H.; Zhu, C.; Zhuo, M.; Chen, B.; Li, S. Probing the Synergistic Ratio of P450/CPR To Improve (+)-Nootkatone Production in Saccharomyces cerevisiae. J. Agric. Food Chem. 2022, 70, 815–825. [Google Scholar] [CrossRef]
  73. Campelo, D.; Esteves, F.; Brito Palma, B.; Costa Gomes, B.; Rueff, J.; Lautier, T.; Urban, P.; Truan, G.; Kranendonk, M. Probing the Role of the Hinge Segment of Cytochrome P450 Oxidoreductase in the Interaction with Cytochrome P450. Int. J. Mol. Sci. 2018, 19, 3914. [Google Scholar] [CrossRef] [PubMed]
  74. Urlacher, V.B.; Girhard, M. Cytochrome P450 Monooxygenases in Biotechnology and Synthetic Biology. Trends Biotechnol. 2019, 37, 882–897. [Google Scholar] [CrossRef] [PubMed]
  75. Ivanov, Y.D.; Taldaev, A.; Lisitsa, A.V.; Ponomarenko, E.A.; Archakov, A.I. Prediction of Monomeric and Dimeric Structures of CYP102A1 Using AlphaFold2 and AlphaFold Multimer and Assessment of Point Mutation Effect on the Efficiency of Intra- and Interprotein Electron Transfer. Molecules 2022, 27, 1386. [Google Scholar] [CrossRef] [PubMed]
  76. Meng, S.; Li, Z.; Ji, Y.; Ruff, A.J.; Liu, L.; Davari, M.D.; Schwaneberg, U. Introduction of aromatic amino acids in electron transfer pathways yielded improved catalytic performance of cytochrome P450s. Chin. J. Catal. 2023, 49, 81–90. [Google Scholar] [CrossRef]
  77. Zhao, L.; Guven, G.; Li, Y.; Schwaneberg, U. First steps towards a Zn/Co(III)sep-driven P450 BM3 reactor. Appl. Microbiol. Biotechnol. 2011, 91, 989–999. [Google Scholar] [CrossRef]
  78. Velazquez, M.N.R.; Noebauer, M.; Pandey, A.V. Loss of Protein Stability and Function Caused by P228L Variation in NADPH-Cytochrome P450 Reductase Linked to Lower Testosterone Levels. Int. J. Mol. Sci. 2022, 23, 10141. [Google Scholar] [CrossRef] [PubMed]
  79. Yu, J.; Ge, J.; Yu, H.; Ye, L. Improved Bioproduction of the Nylon 12 Monomer by Combining the Directed Evolution of P450 and Enhancing Heme Synthesis. Molecules 2023, 28, 1758. [Google Scholar] [CrossRef] [PubMed]
  80. Liu, F.; Yang, H.; Wang, L.; Yu, B. Biosynthesis of the High-Value Plant Secondary Product Benzyl Isothiocyanate via Functional Expression of Multiple Heterologous Enzymes in Escherichia coli. ACS Synth. Biol. 2016, 5, 1557–1565. [Google Scholar] [CrossRef] [PubMed]
  81. Zhao, F.; Bai, P.; Liu, T.; Li, D.; Zhang, X.; Lu, W.; Yuan, Y. Optimization of a cytochrome P450 oxidation system for enhancing protopanaxadiol production in Saccharomyces cerevisiae. Biotechnol. Bioeng. 2016, 113, 1787–1795. [Google Scholar] [CrossRef]
  82. Guo, X.; Sun, J.; Li, D.; Lu, W. Heterologous biosynthesis of (+)-nootkatone in unconventional yeast Yarrowia lipolytica. Biochem. Eng. J. 2018, 137, 125–131. [Google Scholar] [CrossRef]
  83. Sun, J.; Zhang, C.; Nan, W.; Li, D.; Ke, D.; Lu, W. Glycerol improves heterologous biosynthesis of betulinic acid in engineered Yarrowia lipolytica. Chem. Eng. Sci. 2019, 196, 82–90. [Google Scholar] [CrossRef]
  84. Li, D.; Wu, Y.; Zhang, C.; Sun, J.; Zhou, Z.; Lu, W. Production of Triterpene Ginsenoside Compound K in the Non-conventional Yeast Yarrowia lipolytica. J. Agric. Food Chem. 2019, 67, 2581–2588. [Google Scholar] [CrossRef] [PubMed]
  85. Aalbers, F.S.; Fraaije, M.W. Enzyme Fusions in Biocatalysis: Coupling Reactions by Pairing Enzymes. Chembiochem 2019, 20, 20–28. [Google Scholar] [CrossRef] [PubMed]
  86. Papaleo, E.; Saladino, G.; Lambrughi, M.; Lindorff-Larsen, K.; Gervasio, F.L.; Nussinov, R. The Role of Protein Loops and Linkers in Conformational Dynamics and Allostery. Chem. Rev. 2016, 116, 6391–6423. [Google Scholar] [CrossRef] [PubMed]
  87. Moon, J.H.; Lee, K.; Lee, J.H.; Lee, P.C. Redesign and reconstruction of a steviol-biosynthetic pathway for enhanced production of steviol in Escherichia coli. Microb. Cell Fact. 2020, 19, 20. [Google Scholar] [CrossRef] [PubMed]
  88. Haga, T.; Hirakawa, H.; Nagamune, T. Fine tuning of spatial arrangement of enzymes in a PCNA-mediated multienzyme complex using a rigid poly-L-proline linker. PLoS ONE 2013, 8, e75114. [Google Scholar] [CrossRef]
  89. Li, D.; Wu, Y.; Wei, P.; Gao, X.; Li, M.; Zhang, C.; Zhou, Z.; Lu, W. Metabolic engineering of Yarrowia lipolytica for heterologous oleanolic acid production. Chem. Eng. Sci. 2020, 218, 115529. [Google Scholar] [CrossRef]
  90. Park, S.Y.; Eun, H.; Lee, M.H.; Lee, S.Y. Metabolic engineering of Escherichia coli with electron channelling for the production of natural products. Nat. Catal. 2022, 5, 726–737. [Google Scholar] [CrossRef]
  91. Wang, Z.; Dai, Y.; Azi, F.; Wang, Z.; Xu, W.; Wang, D.; Dong, M.; Xia, X. Constructing Protein-Scaffolded Multienzyme Assembly Enhances the Coupling Efficiency of the P450 System for Efficient Daidzein Biosynthesis from (2S)-Naringenin. J. Agric. Food Chem. 2024, 72, 5849–5859. [Google Scholar] [CrossRef]
  92. Cirino, P.C.; Arnold, F.H. A Self-Sufficient Peroxide-Driven Hydroxylation Biocatalyst. Angew. Chem. Int. Ed. 2003, 42, 3299–3301. [Google Scholar] [CrossRef]
  93. Zhao, H.; Donk, W.A. Regeneration of cofactors for use in biocatalysis. Curr. Opin. Biotechnol. 2003, 14, 583–589. [Google Scholar] [CrossRef] [PubMed]
  94. Hollmann, F.; Arends, I.W.C.E.; Buehler, K. Biocatalytic Redox Reactions for Organic Synthesis: Nonconventional Regeneration Methods. Chemcatchem 2010, 2, 762–782. [Google Scholar] [CrossRef]
  95. Lee, S.H.; Kwon, Y.C.; Kim, D.M.; Park, C.B. Cytochrome P450-catalyzed O-dealkylation coupled with photochemical NADPH regeneration. Biotechnol. Bioeng. 2013, 110, 383–390. [Google Scholar] [CrossRef]
  96. Jensen, K.; Jensen, P.E.; Moller, B.L. Light-driven cytochrome p450 hydroxylations. ACS Chem. Biol. 2011, 6, 533–539. [Google Scholar] [CrossRef] [PubMed]
  97. Lee, W.H.; Kim, M.; Jin, Y.S.; Seo, J.H. Engineering of NADPH regenerators in Escherichia coli for enhanced biotransformation. Appl. Microbiol. Biotechnol. 2013, 97, 2761–2772. [Google Scholar] [CrossRef] [PubMed]
  98. Bernhardt, R. Cytochromes P450 as versatile biocatalysts. J. Biotechnol. 2006, 124, 128–145. [Google Scholar] [CrossRef]
  99. Mi, L.; Wang, Z.; Yang, W.; Huang, C.; Zhou, B.; Hu, Y.; Liu, S. Cytochromes P450 in biosensing and biosynthesis applications: Recent progress and future perspectives. TrAC Trends Anal. Chem. 2023, 158, 116791. [Google Scholar] [CrossRef]
  100. Guengerich, F.P. Intersection of the Roles of Cytochrome P450 Enzymes with Xenobiotic and Endogenous Substrates: Relevance to Toxicity and Drug Interactions. Chem. Res. Toxicol. 2017, 30, 2–12. [Google Scholar] [CrossRef] [PubMed]
  101. Shumyantseva, V.V.; Koroleva, P.I.; Bulko, T.V.; Shkel, T.V.; Gilep, A.A.; Veselovsky, A.V. Approaches for increasing the electrocatalitic efficiency of cytochrome P450 3A4. Bioelectrochemistry 2023, 149, 108277. [Google Scholar] [CrossRef]
  102. Brett, C.; Brett, A.M.O. Electrochemistry. Principles, methods and applications: Christopher, M.A. Brett and Ana Maria Oliveira Brett Oxford Science Publications 1993, 427 pp., 25. Electrochim. Acta 1994, 39, 853. [Google Scholar]
  103. Shumyantseva, V.V.; Kuzikov, A.V.; Masamrekh, R.A.; Bulko, T.V.; Archakov, A.I. From electrochemistry to enzyme kinetics of cytochrome P450. Biosens. Bioelectron. 2018, 121, 192–204. [Google Scholar] [CrossRef] [PubMed]
  104. Masamrekh, R.A.; Kuzikov, A.V.; Haurychenka, Y.I.; Shcherbakov, K.A.; Veselovsky, A.V.; Filimonov, D.A.; Dmitriev, A.V.; Zavialova, M.G.; Gilep, A.A.; Shkel, T.V.; et al. In vitro interactions of abiraterone, erythromycin, and CYP3A4: Implications for drug-drug interactions. Fundam. Clin. Pharmacol. 2020, 34, 120–130. [Google Scholar] [CrossRef]
  105. Castrignano, S.; Di Nardo, G.; Sadeghi, S.J.; Gilardi, G. Influence of inter-domain dynamics and surrounding environment flexibility on the direct electrochemistry and electrocatalysis of self-sufficient cytochrome P450 3A4-BMR chimeras. J. Inorg. Biochem. 2018, 188, 9–17. [Google Scholar] [CrossRef] [PubMed]
  106. Algov, I.; Grushka, J.; Zarivach, R.; Alfonta, L. Highly efficient flavin-adenine dinucleotide glucose dehydrogenase fused to a minimal cytochrome c domain. J. Am. Chem. Soc. 2017, 139, 17217–17220. [Google Scholar] [CrossRef] [PubMed]
  107. Udit, A.K.; Hindoyan, N.; Hill, M.G.; Arnold, F.H.; Gray, H.B. Protein-surfactant film voltammetry of wild-type and mutant cytochrome P450 BM3. Inorg. Chem. 2005, 44, 4109–4111. [Google Scholar] [CrossRef] [PubMed]
  108. Cui, D.; Mi, L.; Xu, X.; Lu, J.; Qian, J.; Liu, S. Nanocomposites of Graphene and Cytochrome P450 2D6 Isozyme for Electrochemical-Driven Tramadol Metabolism. Langmuir 2014, 30, 11833–11840. [Google Scholar] [CrossRef] [PubMed]
  109. Yoshioka, K.; Kato, D.; Kamata, T.; Niwa, O. Cytochrome P450 Modified Polycrystalline Indium Tin Oxide Film as a Drug Metabolizing Electrochemical Biosensor with a Simple Configuration. Anal. Chem. 2013, 85, 9996–9999. [Google Scholar] [CrossRef]
  110. Zernia, S.; Ott, F.; Bellmann-Sickert, K.; Frank, R.; Klenner, M.; Jahnke, H.G.; Prager, A.; Abel, B.; Robitzki, A.A.; Beck-Sickinger, A.G. Peptide-Mediated Specific Immobilization of Catalytically Active Cytochrome P450 BM3 Variant. Bioconjug Chem. 2016, 27, 1090–1097. [Google Scholar] [CrossRef] [PubMed]
  111. Rusling, J.F. ChemInform Abstract: Enzyme Bioelectrochemistry in Cast Biomembrane-Like Films. ChemInform 1998, 31, 363–369. [Google Scholar] [CrossRef]
  112. Shumyantseva, V.V.; Bulko, T.V.; Kuzikov, A.V.; Masamrekh, R.A.; Konyakhina, A.Y.; Romanenko, I.; Max, J.B.; Köhler, M.; Gilep, A.A.; Usanov, S.A.; et al. All-electrochemical nanocomposite two-electrode setup for quantification of drugs and study of their electrocatalytical conversion by cytochromes P450. Electrochim. Acta 2020, 336, 135579. [Google Scholar] [CrossRef]
  113. Panicco, P.; Castrignano, S.; Sadeghi, S.J.; Nardo, G.D.; Gilardi, G. Engineered human CYP2C9 and its main polymorphic variants for bioelectrochemical measurements of catalytic response. Bioelectrochemistry 2021, 138, 107729. [Google Scholar] [CrossRef] [PubMed]
  114. Shumyantseva, V.; Bulko, T.; Koroleva, P.; Shich, E.; Makhova, A.; Kisel, M.; Haidukevich, I.; Gilep, A. Human Cytochrome P450 2C9 and its Polymorphic Modifications: Electroanalysis, Catalytic Properties and Approaches to the Regulation of Enzymatic Activity; Social Science Electronic Publishing: New York NY, USA, 2022. [Google Scholar]
  115. Kuzikov, A.V.; Filippova, T.A.; Masamrekh, R.A.; Shumyantseva, V.V. Electrochemical determination of (S)-7-hydroxywarfarin for analysis of CYP2C9 catalytic activity. J. Electroanal. Chem. 2022, 904, 115937. [Google Scholar] [CrossRef]
  116. Yang, M.; Kabulski, J.L.; Wollenberg, L.; Chen, X.; Subramanian, M.; Tracy, T.S.; Lederman, D.; Gannett, P.M.; Wu, N. Electrocatalytic drug metabolism by CYP2C9 bonded to a self-assembled monolayer-modified electrode. Drug Metab. Dispos. 2009, 37, 892–899. [Google Scholar] [CrossRef] [PubMed]
  117. Kuzikov, A.V.; Filippova, T.A.; Masamrekh, R.A.; Shumyantseva, V.V. Biotransformation of phenytoin in the electrochemically-driven CYP2C19 system. Biophys. Chem. 2022, 291, 106894. [Google Scholar] [CrossRef]
  118. Vargas, A.F.L.; Buitrago, W.G.Q.; Silva, D.C.C. Voltammetric Responses of a CYP2D6-Based Biosensor to 3,4-methylenedioxymethamphetamine (MDMA) and the Synthetic Cathinone α-pyrrolidinopentiophenone (α-PVP). Chem. Sel. 2022, 42, e202202748. [Google Scholar] [CrossRef]
  119. Shu, T.; Wang, J.; Li, X.; Wang, X.; Wang, S. Cytochrome P450 2D6 biosensor for perphenazine based on multi-walled carbon nanotube/ionic liquid and tetrathiafulvalene-tetracyanoquinodimethane salt/ionic liquid gels. Electrochim. Acta 2023, 439, 141656. [Google Scholar] [CrossRef]
  120. Kuzikov, A.V.; Masamrekh, R.A.; Filippova, T.A.; Tumilovich, A.M.; Strushkevich, N.V.; Gilep, A.A.; Khudoklinova, Y.Y.; Shumyantseva, V.V. Bielectrode Strategy for Determination of CYP2E1 Catalytic Activity: Electrodes with Bactosomes and Voltammetric Determination of 6-Hydroxychlorzoxazone. Biomedicines 2024, 12, 152. [Google Scholar] [CrossRef]
  121. Koroleva, P.I.; Gilep, A.A.; Kraevsky, S.V.; Tsybruk, T.V.; Shumyantseva, V.V. Improving the Efficiency of Electrocatalysis of Cytochrome P450 3A4 by Modifying the Electrode with Membrane Protein Streptolysin O for Studying the Metabolic Transformations of Drugs. Biosensors 2023, 13, 457. [Google Scholar] [CrossRef]
  122. Shumyantseva, V.V.; Koroleva, P.I.; Gilep, A.A.; Napolskii, K.S.; Ivanov, Y.D.; Kanashenko, S.L.; Archakov, A.I. Increasing the Efficiency of Cytochrome P450 3A4 Electrocatalysis Using Electrode Modification with Spatially Ordered Anodic Aluminum Oxide-Based Nanostructures for Investigation of Metabolic Transformations of Drugs. Dokl. Biochem. Biophys. 2022, 506, 215–219. [Google Scholar] [CrossRef]
  123. Cheropkina, H.; Catucci, G.; Cesano, F.; Marucco, A.; Gilardi, G.; Sadeghi, S.J. Bioelectrochemical platform with human monooxygenases: FMO1 and CYP3A4 tandem reactions with phorate. Bioelectrochemistry 2023, 150, 108327. [Google Scholar] [CrossRef]
  124. Kuzikov, A.; Masamrekh, R.; Shkel, T.; Strushkevich, N.; Gilep, A.; Usanov, S.; Archakov, A.; Shumyantseva, V. Assessment of electrocatalytic hydroxylase activity of cytochrome P450 3A4 (CYP3A4) by means of derivatization of 6beta-hydroxycortisol by sulfuric acid for fluorimetric assay. Talanta 2019, 196, 231–236. [Google Scholar] [CrossRef]
  125. Kuzikov, A.V.; Masamrekh, R.A.; Filippova, T.A.; Haurychenka, Y.I.; Gilep, A.A.; Shkel, T.V.; Strushkevich, N.V.; Usanov, S.A.; Shumyantseva, V.V. Electrochemical oxidation of estrogens as a method for CYP19A1 (aromatase) electrocatalytic activity determination. Electrochim. Acta 2020, 333, 135539. [Google Scholar] [CrossRef]
  126. Wu, Y.; Jiang, N.; He, Z.; Yang, Y.; Li, Y. Direct electrochemical detection of extracellular nitric oxide in Arabidopsis protoplast based on cytochrome P450 55B1 biosensor. Nitric Oxide 2023, 132, 8–14. [Google Scholar] [CrossRef]
  127. Ray, M.; Mhaske, S.D.; Haram, S.K.; Mazumdar, S. Covalent Conjugation of Single-Walled Carbon Nanotube with CYP101 mutant for Direct Electrocatalysis. Anal. Biochem. 2021, 626, 114204. [Google Scholar] [CrossRef]
  128. Shumyantseva, V.V.; Kuzikov, A.V.; Masamrekh, R.A.; Khatri, Y.; Zavialova, M.G.; Bernhardt, R.; Archakov, A.I. Direct electrochemistry of CYP109C1, CYP109C2 fraom Sorangium cellulosum So ce56. Electrochim. Acta 2016, 192, 72–79. [Google Scholar] [CrossRef]
  129. Dai, Q.; Yang, L.; Wang, Y.; Cao, X.; Xu, X. Surface charge-controlled electron transfer and catalytic behavior of immobilized cytochrome P450 BM3 inside dendritic mesoporous silica nanoparticles. Anal. Bioanal. Chem. 2020, 412, 4703–4712. [Google Scholar] [CrossRef]
  130. Chen, X.; Cao, Y.; Li, F.; Tian, Y.; Song, H. Enzyme-Assisted Microbial Electrosynthesis of Poly(3-hydroxybutyrate) via CO2 Bioreduction by Engineered Ralstonia eutropha. ACS Catal. 2018, 8, 4429–4437. [Google Scholar] [CrossRef]
  131. Yang, Y.; Ding, Y.; Hu, Y.; Cao, B.; Rice, S.A.; Kjelleberg, S.; Song, H. Enhancing Bidirectional Electron Transfer of Shewanella oneidensis by a Synthetic Flavin Pathway. ACS Synth. Biol. 2015, 4, 815–823. [Google Scholar] [CrossRef]
  132. Rowe, S.F.; Le Gall, G.; Ainsworth, E.V.; Davies, J.A.; Lockwood, C.W.J.; Shi, L.; Elliston, A.; Roberts, I.N.; Waldron, K.W.; Richardson, D.J.; et al. Light-Driven H-2 Evolution and C=C or C=O Bond Hydrogenation by Shewanella oneidensis: A Versatile Strategy for Photocatalysis by Nonphotosynthetic Microorganisms. ACS Catal. 2017, 7, 7558–7566. [Google Scholar] [CrossRef]
  133. Zhang, Z.; Li, F.; Cao, Y.; Tian, Y.; Song, H. Electricity-driven 7α-hydroxylation of a steroid catalyzed by a cytochrome P450 monooxygenase in engineered yeast. Catal. Sci. Technol. 2019, 9, 4877–4887. [Google Scholar] [CrossRef]
  134. Shalan, H.; Kato, M.; Cheruzel, L. Keeping the spotlight on cytochrome P450. Biochim. Biophys. Acta Proteins Proteom. 2018, 1866, 80–87. [Google Scholar] [CrossRef]
  135. Chen, S.; Wang, L.W. Thermodynamic Oxidation and Reduction Potentials of Photocatalytic Semiconductors in Aqueous Solution. Chem. Mater. 2012, 24, 1658–1671. [Google Scholar] [CrossRef]
  136. Park, J.H.; Lee, S.H.; Cha, G.S.; Da, S.C.; Chan, B.P. Cofactor-Free Light-Driven Whole-Cell Cytochrome P450 Catalysis. Angew. Chem. 2015, 54, 969–973. [Google Scholar] [CrossRef]
  137. Kato, M.; Lam, Q.; Bhandarkar, M.; Banh, T.; Heredia, J.; Andrew, U.; Cheruzel, L. Selective C–H bond functionalization with light-driven P450 biocatalysts. Comptes Rendus Chim. 2016, 20, 237–242. [Google Scholar] [CrossRef]
  138. Lee, S.H.; Choi, D.S.; Kuk, S.K.; Park, C.B. Photobiocatalysis: Activating Redox Enzymes by Direct or Indirect Transfer of Photoinduced Electrons. Angew. Chem. 2017, 57, 7958–7985. [Google Scholar] [CrossRef]
  139. Girhard, M.; Kunigk, E.; Tihovsky, S.; Shumyantseva, V.V.; Urlacher, V.B. Light-driven biocatalysis with cytochrome P450 peroxygenases. Biotechnol. Appl. Biochem. 2013, 60, 111–118. [Google Scholar] [CrossRef]
  140. Kim, Y.S.; Hara, M.; Ikebukuro, K.; Miyake, J.; Karube, I. Photo-induced activation of cytochrome P450/reductase fusion enzyme coupled with spinach chloroplasts. Biotechnol. Technol. 1996, 10, 717–720. [Google Scholar] [CrossRef]
  141. Wang, Y.; Jin, M.; Wang, J.; Bai, L.; Yang, Y.; Dai, H.; Cui, D.; Zhao, M. Light-driven biodegradation of azo dyes by Shewanella decolorationis-CdS biohybrid in wastewater lacking electron donors. Appl. Microbiol. Biotechnol. 2023, 107, 447–457. [Google Scholar] [CrossRef]
  142. Liu, K.; Wang, F.-Q.; Liu, K.; Zhao, Y.; Gao, B.; Tao, X.; Wei, D. Light-driven progesterone production by InP–(M. neoaurum) biohybrid system. Bioresour. Bioprocess. 2022, 9, 93. [Google Scholar] [CrossRef]
  143. Torrado, A.; Connabeer, H.M.; Rottig, A.; Pratt, N.; Baylay, A.J.; Terry, M.J.; Moore, C.M.; Bibby, T.S. Directing cyanobacterial photosynthesis in a cytochrome c oxidase mutant using a heterologous electron sink. Plant Physiol. 2022, 189, 2554–2566. [Google Scholar] [CrossRef]
  144. Le, T.K.; Kim, J.; Anh Nguyen, N.; Huong Ha Nguyen, T.; Sun, E.G.; Yee, S.M.; Kang, H.S.; Yeom, S.J.; Beum Park, C.; Yun, C.H. Solar-Powered Whole-Cell P450 Catalytic Platform for C-Hydroxylation Reactions. ChemSusChem 2021, 14, 3054–3058. [Google Scholar] [CrossRef]
  145. Le, T.-K.; Park, J.H.; Choi, D.S.; Lee, G.-Y.; Choi, W.S.; Jeong, K.J.; Park, C.B.; Yun, C.-H. Solar-driven biocatalytic C-hydroxylation through direct transfer of photoinduced electrons. Green. Chem. 2019, 21, 515–525. [Google Scholar] [CrossRef]
  146. Archakov, V. Fluorescence Spectroscopy Study of the Interaction of Cytochrome P450 2B4 with Riboflavin. Biophysics 2004, 49, 13–18. [Google Scholar]
  147. Qian, J.; Zhu, W.; Mi, L.; Xu, X.; Yu, J.; Cui, D.; Xue, Y.; Liu, S. Nanohybrids of quantum dots and cytochrome P450 for light-driven drug metabolism. J. Electroanal. Chem. 2014, 733, 27–32. [Google Scholar] [CrossRef]
  148. Jiang, H.; Li, X.; Li, M.; Niu, P.; Wang, T.; Chen, D.; Chen, P.; Zou, J.-P. A new strategy for triggering photocatalytic activity of Cytrochrome P450 by coupling of semiconductors. Chem. Eng. J. 2019, 358, 58–66. [Google Scholar] [CrossRef]
  149. Sosa, V.; Melkie, M.; Sulca, C.; Li, J.; Tang, L.; Li, J.; Faris, J.; Foley, B.; Banh, T.; Kato, M.; et al. Selective Light-Driven Chemoenzymatic Trifluoromethylation/Hydroxylation of Substituted Arenes. ACS Catal. 2018, 8, 2225–2229. [Google Scholar] [CrossRef]
  150. Li, N.; Yan, S.; Wu, P.; Li, J.; Wang, B. Local Electric Fields Drives the Proton-Coupled Electron Transfer within Cytochrome P450 Reductase. ACS Catal. 2024, 14, 7893–7900. [Google Scholar] [CrossRef]
  151. Agustinus, B.; Gillam, E.M.J. Solar-powered P450 catalysis: Engineering electron transfer pathways from photosynthesis to P450s. J. Inorg. Biochem. An. Interdiscip. J. 2023, 245, 112242. [Google Scholar] [CrossRef]
  152. Shumyantseva, V.V.; Koroleva, P.I.; Bulko, T.V.; Agafonova, L.E. Alternative Electron Sources for Cytochrome P450s Catalytic Cycle: Biosensing and Biosynthetic Application. Processes 2023, 11, 1801. [Google Scholar] [CrossRef]
Molecules 29 02480 g001
Figure 1. Schematic organization of different P450s systems. Abbreviations: FdR, ferredoxin reductase; FAD, flavin adenine dinucleotide; Fdx, ferredoxin; CPR, cytochrome P450 reductase; ER, endoplasmic reticulum; Fld, flavodoxin; FMN, flavin mononucleotide; OFOR, 2-oxo-acid-ferredoxin oxidoreductase; PFOR, phthalate family oxygenase reductase.
Figure 1. Schematic organization of different P450s systems. Abbreviations: FdR, ferredoxin reductase; FAD, flavin adenine dinucleotide; Fdx, ferredoxin; CPR, cytochrome P450 reductase; ER, endoplasmic reticulum; Fld, flavodoxin; FMN, flavin mononucleotide; OFOR, 2-oxo-acid-ferredoxin oxidoreductase; PFOR, phthalate family oxygenase reductase.
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Figure 2. Schematic diagram of the P450 catalytic mechanism.
Figure 2. Schematic diagram of the P450 catalytic mechanism.
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Figure 3. Strategies for engineering the electron transfer pathway of cytochrome P450s.
Figure 3. Strategies for engineering the electron transfer pathway of cytochrome P450s.
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Figure 4. Schematic diagram of P450s and CRP engineering.
Figure 4. Schematic diagram of P450s and CRP engineering.
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Table 1. Catalysis of P450s in different electrochemical systems.
Table 1. Catalysis of P450s in different electrochemical systems.
SpeciesElectrodesKm/μMSubstrateReference
CYP2C9SPE 1/DDAB 245 ± 5diclofenac[114]
CYP2C9CE 3/DDAB3.03 ± 0.38(S)-7-hydroxywarfarin[115]
CYP2C9GOLD/SAM3warfarin[116]
CYP2C19CE/DDAB25.8 ± 2.04-hydroxyphenytoin[117]
CYP2D6CE/SPE-MDMA and α-PVP[118]
CYP2D6MWNT/[BMIM][PF6]-[TTF–TCNQ]/[BMIM][PF6]5.52Perphenazine[119]
CYP2E1SPE/DDAB78 ± 9chlorzoxazone[120]
CYP3A4SPE/DDAB/SLO 4207 ± 2.5erythromycin[121]
CYP3A4DDAB70erythromycin[101]
CYP3A4DDAB--[122]
CYP3A4GCE/GO/DDAB29.6 ± 4.1phorate[123]
CYP3A4SPE/(PB-b-PDMAEMA/MWCNTs)48 ± 8diclofenac[112]
CYP3A4SPE/DDAB10 ± 26β-hydroxycortisol[124]
CYP19A1GE/DDAB4.2 ± 1.5estrone[125]
CYP55B1PGE/GA/BSA11.64 × 10−3Nitric oxide reductase[126]
CYP101GC/SWCNT--[127]
CYP109D1SPE/DDAB-myristic acid[128]
P450 BM3GCE/NH2-DMSN244.82testosterone[129]
1 screen-printed electrodes. 2 didodecyldimethylammonium bromide. 3 graphite electrodes. 4 streptolysin O.
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He, J.; Liu, X.; Li, C. Engineering Electron Transfer Pathway of Cytochrome P450s. Molecules 2024, 29, 2480. https://doi.org/10.3390/molecules29112480

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He J, Liu X, Li C. Engineering Electron Transfer Pathway of Cytochrome P450s. Molecules. 2024; 29(11):2480. https://doi.org/10.3390/molecules29112480

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He, Jingting, Xin Liu, and Chun Li. 2024. "Engineering Electron Transfer Pathway of Cytochrome P450s" Molecules 29, no. 11: 2480. https://doi.org/10.3390/molecules29112480

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He, J., Liu, X., & Li, C. (2024). Engineering Electron Transfer Pathway of Cytochrome P450s. Molecules, 29(11), 2480. https://doi.org/10.3390/molecules29112480

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