Abstract
Purpose of Review
We provide an overview of the current knowledge on cytochrome P450-mediated metabolism organized as metabolons and factors that facilitate their stabilization. Essential parameters will be discussed including those that are commonly disregarded using the dhurrin metabolon from Sorghum bicolor as a case study.
Recent Findings
Sessile plants control their metabolism to prioritize their resources between growth and development, or defense. This requires fine-tuned complex dynamic regulation of the metabolic networks involved. Within the recent years, numerous studies point to the formation of dynamic metabolons playing a major role in controlling the metabolic fluxes within such networks.
Summary
We propose that P450s and their partners interact and associate dynamically with POR, which acts as a charging station possibly in concert with Cytb5. Solvent environment, lipid composition, and non-catalytic proteins guide metabolon formation and thereby activity, which have important implications for synthetic biology approaches aiming to produce high-value specialized metabolites in heterologous hosts.
Similar content being viewed by others
Introduction
As sessile organisms, plants produce an impressive diversity of toxic chemical compounds to defend themselves from the continuum of biotic and abiotic challenges they are exposed to. The specialized metabolites produced depend on the specific plant species and type of challenges encountered, and their biosynthesis demands a high degree of functional organization to offer swift metabolic plasticity. Delicate mechanisms regulate the complex metabolic grids organizing biosynthesis and storage [1••]. Organellar compartmentation allows cells to bring enzymes and their substrates in close proximity, which serves to increase local concentration for biological process optimization. In plants, many pathways involved in both general and specialized metabolism have been proposed to organize in protein complexes, metabolons [2•]. This extends the idea of compartmentalization at the molecular level. Rapid metabolic reconfigurations, as required for biosynthetic diversity, may be controlled by formation of dynamic metabolons [3] facilitating channeling of labile and toxic intermediates out of the bulk phase, increasing local substrate concentrations and preventing undesired metabolic cross talk [4]. Dynamic assembly and disassembly permit swift adaptation of the metabolite profile to environmental challenges [5, 6]. Beyond the fundamental understanding of cellular metabolism, it has become increasingly important to understand how enzyme systems catalyzing complex reactions are spatially organized and their possible enrolment as part of dynamic metabolons. Intensive efforts to maximize product yield from genetically engineered pathways were undertaken without much attention to this aspect [7,8,9]. Current synthetic biology approaches based on the function of biological modules and the possibility to combine these in new ways are challenged up front by constraints arising from lack of understanding the impact of proper structural organization [10, 11•, 12]. The concept of metabolon formation in plants and animals was first suggested in 1974 [13] and predicted to be of relevance to both general and specialized metabolism. Pathways known to involve the formation of metabolons include the dhurrin pathway [14, 15••], the core phenylpropanoid pathway [5, 16], the flavonoid pathways [17, 18], the isoflavonoid pathways [19•, 20], the biosynthesis of sporopollenin subunits [21], the TCA cycle [22•], the Krebs cycle [23•], the Calvin-Benson cycle [24], the biosynthesis of bitter acids in hop [25], the biosynthesis of lipids [26], the fatty acid biosynthetic network [27], the spermine/spermidine synthesis [28], the photosynthetic complex [29], the auxin synthesis [30,31,, 31••, 32], the cholesterol synthesis [33], the de novo purine synthesis [34•], the urea cycle of rat hepatocytes [35], the citrate cycle of porcine liver [36], the glycolytic pathway in rabbit skeletal muscle and mouse fibroblasts [37, 38] and the glycolytic pathway in Saccharomyces cerevisiae [39]. Interested readers can read some of the excellent reviews written over the years on this topic [1••, 2•, 13, 14, 40,41,42]. Despite this rather exhaustive accumulation of data, there is still skepticism about the necessity of metabolons for organizing cellular metabolism mainly due to the technical challenges encountered in efforts directed towards isolating the active complexes. However, scientific papers on this subject have accumulated and the number of papers mentioning metabolons has rapidly increased within the last 5 years (Fig. 1).
Pubmed occurrences about metabolon papers. Pubmed (https://www.ncbi.nlm.nih.gov/pubmed) queried towards the end of November 2016. With the queries: “protein-protein interaction,” “protein complex,” metabolon, and plant and metabolon
In this review, we will focus mostly on recent papers and use the dhurrin metabolon as a case study. Dhurrin is a cyanogenic glucoside present in the cereal crop plant Sorghum bicolor. Dhurrin is produced from the amino acid l-tyrosine and synthesized by two membrane-anchored cytochrome P450 enzymes (CYP79A1 and CYP71E1) and by a soluble UDP-glucosyltransferase (UGT85B1). Although multiple studies have focused on the formation of metabolons involving P450 enzymes, and even if metabolon formation seems essential to organize and direct flux across biosynthetic networks [6], it is still unknown how the dynamic assembly of such complexes is regulated and organized. Here, we will illustrate different possible models of organization, which are best in agreement with the published data. In the first parts of this review, we will discuss briefly factors that are classically considered when studying metabolon formation, i.e., channeling of intermediates and protein-protein interactions. Then, we will review more extensively factors that are likely to regulate the dynamic assembly process, i.e., membrane environment, solvent environment, non-catalytic proteins and higher-order clustering, that are still largely disregarded.
Definition and Challenges of P450-Based Metabolons
The expansion of the plant-specialized - metabolites during evolution notably results from the massive recruitment of cytochromes P450 (P450s) that catalyze multiple types of catalytic reactions [43]. Eukaryotic P450s are type-II membrane-anchored enzymes and proposed as nucleation platforms anchoring metabolons to the endoplasmic reticulum (ER), i.e., in the lignin [5], the isoflavonoid [19•], and the dhurrin biosynthetic pathways [15••]. The plant ER is a dynamic multitasking organelle connected to several compartments of the cell and involved in cell processes like protein synthesis, signaling, and primary and secondary metabolism [44]. The ER membrane is highly crowded with proteins [45], which points to a tight organization of the ER proteins to avoid hazardous crosstalk and to prevent inadvertent competition for common substrates between overlapping P450-mediated metabolic pathways.
Channeling, Protein-Protein Interactions and Allosteric Regulation
Metabolons are defined as organization of sequential biosynthetic enzymes into complexes and involve substrate channeling and protein-protein interactions. Furthermore, metabolons involved in specialized metabolism are commonly considered highly dynamic, responding to environmental conditions. Initially, substrate channeling studies pioneered the investigation of metabolons [46], providing evidence for intermediate channeling between two consecutive enzymes of the phenylpropanoid pathway, the phenylalanine ammonia lyase (PAL) and the cinnamate 4-hydroxylase (C4H), a P450. Although the results of succeeding studies on channeling between these enzymes have been contradictory [47, 48], this phenomenon has been observed in several pathways and remains a key feature of metabolons [2•, 14, 41]. Another key parameter is the occurrence of multiple weak protein-protein interactions leading to stabilization of larger complexes. More and more studies demonstrate protein interactions in virtually all parts of plant metabolism, including the phenylpropanoid pathway [5, 19•, 21, 49], the dhurrin pathway [15••, 50], and lipid biosynthesis [26]. Complex formation also involves P450-P450 complexes [51••] with reported oligomers ranging from dimers up to 40-mers [52]. In eukaryotes, P450s are nearly always present as membrane proteins, with N-terminal hydrophobic amino acids forming an α-helix anchor to the ER membrane and their catalytic domain facing the cytosol. In this manner, P450s constitute a nucleation platform facilitating localization of metabolons to the ER. Evidence has shown that metabolism of substrates by a given P450 may be influenced by the specific interaction with other P450s [51••], pointing to competition or allosteric effects between P450s. Thus, the last parameter typically considered is the allosteric effect between enzymes involved in metabolon formation. For example, we observed that the mere physical presence of UGT85B1, even in the absence of its UDP-glucose cofactor, stimulated the catalytic properties of CYP79A1, a non-direct partner in the dhurrin pathway [15••], supporting the assembly of a metabolon in planta.
Stoichiometric Imbalance Leads to Dynamic Assembly
The catalytic cycle of all microsomal P450s requires two single-electron transfer reactions catalyzed by the shared NADPH-dependent cytochrome P450 oxidoreductase (POR). This makes POR an essential component of P450-mediated metabolism, even if it is typically not considered in the understanding of metabolic networks and metabolons. POR, as P450s, is an ER-anchored protein, and the physical interactions between P450s and POR are facilitated and modulated by the membrane environment [53, 54]. It is still not clear if electron transfer from POR to a P450 enzyme requires formation of a 1:1 functional complex [55] or the involvement of higher-order multimers of the P450s [51••, 56]. In most organisms, the P450s are present in a large excess in comparison to POR with reported stoichiometric ratios from 5:1 to 40:1 [15••, 52, 57,58,59,60,61]. Nevertheless, POR must provide electrons to each of the different P450s present. This may be controlled by the multiple functional states [62] and conformational dynamics of POR [63, 64]. The large number of different P450s compared to the presence of sub-stoichiometric amounts of POR has prompted the hypothesis of transient interactions between P450s and POR. P450-mediated metabolism may therefore be limited by the POR availability. Thus, highly complex and efficient regulations must occur to ensure the coordinated operation of all P450s belonging to various biosynthetic pathways. P450-P450 interactions can alter P450-mediated metabolism by affecting their ability to interact with POR, to bind substrate, or by both types of mechanisms [51••]. In contrast to yeasts and mammals, the P450-POR couple is even more complex in plants. Plant POR isoforms usually group into two phylogenetically distinct classes [65, 66]. The number of POR isoforms present within a single plant species is between one and four, and they may be derived from a single or both phylogenetic classes [67]. The two POR classes have not been shown to display structural specificity towards individual P450. Direct in vitro interaction and reduction assays of P450s demonstrated that both POR classes performed equally well [66]. However, differential expression patterns of the two classes suggest functional distinct roles. The POR class I enzymes are constitutively expressed and suggested to mainly be involved in primary and basal specialized metabolism. The class II PORs have an adaptive role in specialized metabolism [65, 66] responding to environmental stimuli such as wounding, pathogen infection, or light exposure [67,68,69,70]. The hypothesis is that class II POR provides electrons for highly expressed P450s involved in “on-demand” specialized metabolism. This highlights the evolutionary strategy that ensures an efficient activation of P450s in planta.
Microenvironments Regulate Metabolon Assembly: an Unknown Terrain
Accumulating experimental data support the importance of individual protein-protein interactions in order to accomplish substrate channeling. However, the experimental approaches used typically involve targeted screening based on prior knowledge. Very little is known about the triggering mechanisms that stimulate assembly of metabolons and the biomembrane microenvironments required to stabilize these dynamic complexes. Here, we will highlight three areas that remain poorly understood: the neighboring proteins not essential for in vitro functional studies, the lipid bilayer environment, and the local solvent medium properties.
Neighbor Proteins in the Microenvironment
In animals and yeast, a small number of proteins have been found associated to P450s without being directly involved in their catalytic function. BiP proteins are co-detected with P450s and seem important for their ER retention [71]. CYP2C2 has been shown to interact with the ubiquitous BAP31 integral membrane protein [72] and reported to associate with the cytoskeletal components actin and myosin [73]. Erg28p has been reported to interact with CYP51 in yeast. The Erg28p protein was proposed to tether a large complex of yeast sterol biosynthetic enzymes [74]. In plants, several ER-resident non-catalytic protein partners have repeatedly been found associated together: P450s, POR, cytochrome b5 (Cytb5), reticulons, synaptotagmin (SYTA), band7 protein, sterol methyltransferase (SMT), vesicle-associated protein (VAP), membrane steroid-binding protein (MSBP) and others [5, 15••, 19•, 75]. These proteins are typically not required for activity in vitro and are generally ignored, but seem to be part of the microenvironment around metabolons and might have a role in their stabilization and activity.
Hildebrandt and Estabrook first reported the involvement of Cytb5 in P450-catalyzed reactions [76]. Cytb5 is a small (15 kDa) hemoprotein known to enhance the activity of several P450s, thus increasing the complexity of the electron donation chain of P450-mediated metabolism, but still largely overlooked. While Cytb5 is usually considered as non-essential for P450 activity in vitro, several studies have demonstrated the complex in vivo functional role of Cytb5 in P450 catalysis [77]. POR is required for delivering the primary single electron to P450s, whereas both POR and Cytb5 can provide the electron for the second electron-transfer step. Cytb5 has been shown to stimulate, inhibit, or have no effect on various P450 reactions [77, 78]. Despite this seemingly elusive functional importance, Cytb5 has a significant role in forming hetero-dimers with P450s either to provide electrons or induce conformation changes augmenting interaction between POR and selected P450s [78]. NADH:cytochrome b5 reductase and Cytb5 may even serve as sole electron donors to the human CYP1A1 [79]. Most of the studies have focused on mammalian Cytb5 proteins. However, several isoforms of Cytb5 are also found in plants and most likely display similar roles [80]. Various approaches pointed to Cytb5 importance for P450 function in plant phenolic metabolism [81] and in fatty acid hydroxylations [82,83,84]. Cytb5 is not essential in glucosinolate biosynthesis but can modulate P450 activities and change the glucosinolate pattern [80]. Cytb5 isoforms have been found associated to the lignin metabolon [5] and the dhurrin metabolon [15••]. Arabidopsis thaliana Cytb5 proteins interact with a series of other proteins in addition to the P450s, such as a plasma membrane-localized sucrose transporter [85] and the ethylene-response regulator RTE [86]. This suggests that Cytb5 could play a broader important physiological role. Understanding the role of the Cytb5 isoforms may have huge implications for synthetic biology approaches to boost P450 metabolism [87••].
Reticulons contribute to ER tubule shaping [75, 88]. Vesicle-associated membrane protein (VAMP)-associated proteins (VAPs) are localized at the ER-plasma membrane contact sites (MCS), and VAPs serve to properly associate ER, cytoskeleton, plasma membrane and cell wall [89]. Plant SYTA is reported to be plasma membrane protein [75, 90], while other synaptotagmins in animal and yeast cells (called tricalbins in yeast) are ER-localized. Synaptotagmins form ER-plasma membrane junctions maintain ER morphology, stabilize the MCS and are involved in intracellular lipid trafficking, Ca2+ signaling and exo/endocytosis [89, 91]. Reticulons and VAPs effect endosome contact, endocytosis and even auxin homeostasis [92]. The band7 protein is a homolog of the ER lipid raft-associated proteins (erlins) found associated with high molecular mass protein complexes in mammalian cells [93, 94]. Accordingly, several of the proteins repeatedly found to be associated with P450s and metabolons are involved in ER membrane homeostasis and sterol and lipid homeostasis. This supports the hypothesis of the requirement of a specific environment for the establishment of ER membrane-anchored metabolons.
Membrane Environment: a Dynamic Regulatory Scaffold
Several biosynthetic pathways are distributed across different organelles and even different cells or have products excreted by cells [95,96,97,98,99,100]. MCS in animals have been shown to be involved in lipid trafficking and in other functions such as organelle dynamics, protein import and primary metabolism [101]. The metabolon formation theory is compatible to this organization of metabolic pathways. Subparts of biosynthetic pathways might be organized in different metabolons across different organelles with transporters and neighboring proteins in the metabolon environment making bridges between the different biosynthetic locations. The plant endoplasmic reticulum is an organelle connected to several compartments of the cell, cytoskeleton [102, 103], Golgi [104], chloroplast [105], plasma membrane [102, 106], peroxisomes and mitochondria [107, 108]. Physical contact sites, named plastid-associated membranes (PLAMs), have been demonstrated between the chloroplast and ER membranes [109]. PLAMs have specific lipid compositions which differ from chloroplast and ER [101]. Hemi-fused membranes at PLAMs facilitate inter-organellar interactions and allow enzymes on both membranes to process non-polar compounds from both organelle membranes in a transporter-independent manner [105, 110••]. This would allow deployment of metabolic intermediates from enzymes/metabolons directly into the leaflet of the adjunct organelle. Some metabolons could be settled in or around MCS and PLAMs. Proteins repeatedly observed to co-purify with P450s could be part of these structures.
The ER obviously functions as a highway for delivery of a diverse range of proteins and molecules [31••]. Quantified by in planta fluorescence correlation spectroscopy [15••] or fluorescence recovery after photobleaching [5], P450s and partner soluble enzymes are highly mobile at the ER membrane surface or in the cytoplasm. P450s and therefore also P450-containing metabolons associated to the ER are moving together with the dynamic and constantly remodeled ER structures [46]. Microenvironments containing enzymatic production sites, metabolons, for defense molecules could benefit from targeted actin-guided transport through ER dynamics in response to pathogen attack or for localization near pathogen entry sites to facilitate production of high local concentrations of defense compounds [5, 6, 14, 15••, 111]. Indeed, increase in ER structures on exposure to stress such as wounding has been suggested to have a role in defense against pests and pathogens [112,113,114,115,116]. Metabolons involved in production of cell polymer subunits may also depend on the ER dynamics, as ER structures increase during pollen tube growth [44].
Eukaryotic P450s and POR are both membrane anchored and thought to partly interact via their transmembrane N-amino acid terminus [51••, 71]. Local changes in lipid environment are therefore envisioned to play a key role in controlling and stabilizing metabolon assembly. The increasing understanding of the membrane during the last decade has indeed revealed that the membrane bilayer does not simply act as a passive 2D lattice but actively influences various cellular processes such as transport, protein trafficking and signaling. Independent sets of data show that the catalytic properties of P450s are also modulated by membrane composition [15••, 51••, 117•]. Lipids with varying acyl chain length and head groups may associate to generate lipid domains with different electrostatic surface charges. Lipid membranes can adopt two distinct phases referred to as non-fluid liquid-ordered domains and fluid liquid-disordered domains. The existence of lipid microdomains may direct the selective clustering of proteins into the same specific regions of the membrane increasing their propensity to interact. Modulation of membrane lipid composition under varying environmental conditions is an important part of plant adaptation [118], which would therefore also be expected to impact metabolon establishment. Liquid-ordered domains have been implicated in numerous cellular processes and could act as platforms for metabolon formation [119]. The catalytic properties of animal P450s and POR have also been found to be regulated by their presence in disordered or ordered domains [120]. Some lipid-mediated effects result from direct interactions of P450s and POR with phospholipids. These interactions can affect P450 function by altering their conformation, by affecting the interactions between P450s and their redox partners, or by altering the redox potential of the electron donor POR [51••, 117•, 121, 122]. Animal studies have suggested that P450s are enclosed in a “phospholipid halo”, possibly affecting the diffusion of substrates and products [123]. The specific membrane composition of liquid-ordered membranes has been shown to increase binding efficiency between CYP1A2 and POR [117•, 120]. In the absence of POR, P450s have been shown to possess a greater affinity for negatively charged phospholipids [117•]. CYP2B1 was shown to modulate membrane properties in reconstituted systems by inducing segregation of anionic phospholipids, which resulted in alterations in the activity and conformation of CYP2B1 [124]. The dhurrin P450, CYP71E1, was particularly affected by lipid charges displaying maximal activity in membranes containing 20–30% negatively charged phospholipids [15••]. These findings point to the important but still underestimated fact that plant ER membranes are heterogeneous and that the membrane itself is an integral part of the metabolon [6]. Lipid composition might just be the tip of the iceberg, and several other components of the microenvironment could have a role in metabolon formation and activity.
Controlling the Local Media Based on Intracellular Heterogeneity
Cells have generally been thought of as composed of two solvent environments, namely the hydrophilic aqueous phase and the hydrophobic lipid phase. Increasing evidence suggests that this is a too simplified view and that the cellular cytosol is a marbled soup of solvent microenvironments. This implies that local concentrations of solutes, salt and pH may differ and control metabolism in a hitherto unknown fashion [125••]. Intracellular heterogeneity may proceed from demixing of macromolecules and small molecules as exemplified by the formation of P granules of RNA and protein [126]. A few general metabolites including sugars, some amino acids, choline, and some organic acids such as citric acid, tartaric acid, succinic acid, lactic acid and malic acid are always present in considerable amounts in all microbial, mammalian and plant cells and cannot be only considered as intermediates in metabolic pathways [127•]. When mixed in precise ratios and in the absence of water, these simple molecules become viscous liquids, termed natural deep eutectic solvents (NADES) [128]. Thus, NADES have been proposed as a third cell phase [129]. NADES provide 50–100 times increased solubility of many poorly soluble specialized metabolites and may therefore enable solubilization and storage of various toxic and unstable metabolites in cells [127•, 128, 129] (Knudsen et al., submitted). The occurrence of NADES in the plant cell may explain that compounds like flavonoids and anthocyanins occur at levels above their solubility in both water and lipids [129]. NADES may provide stability towards several stresses such as light, temperature and time [130]. We have recently discovered that NADES provide an excellent inert environment for storing enzymes, which upon dilution recover their activity and pinpoint how plants may tolerate desiccation (Knudsen et al., submitted).
ER components might be associated with NADES, e.g., with the positively charged choline head groups in the ER membrane associating with sugars and acids and contributing to the formation of a NADES. P450 metabolons might be dissolved in this solvent together with substrates and intermediates. Thus, NADES could make a kind of shield around metabolons and create a microenvironment within the cytoplasm, making experimental studies to isolate the metabolon even more complex [5, 129]. NADES acting as a shield may explain the close interaction between enzymes in dynamic metabolons without the need for strong interactions.
Organization of Plant P450 Metabolons: a Case Study of the Dhurrin Metabolon
The plant ER is a multitasking membrane network connected to several organelles and involved in most cellular processes [44]. Consequently, P450s and related metabolons are in close vicinity to several other enzymes and non-catalytic proteins, all of which must be well organized. The precise organization and stoichiometry of individual components present remain poorly understood. Here, we propose several scenarios based on previously published data [15••] and considering the factors most likely to modulate metabolon assembly and activity (see above). The plausibility of each of these scenarios is evaluated pointing to the model 5 presented in Fig. 4b.
The ER membrane system can form diverse structures, e.g., sheets, cisternae, and tubules, but most commonly is tubular with a reported average ER tubule diameter of 50 nm [131,132,133,134]. With an ER tubule circumference of about 157 nm, an annulus at the circumference of an ER tubule membrane is envisioned to consist of an average of 31 proteins each with an averaged diameter of 5 nm. All cellular membranes including the ER are considered highly crowded by proteins [45]. Accordingly, we schematize the ER membrane proteins as a layer of contiguous proteins delimited by the circumference and the length of the ER structure where they reside. An illustration of 4 pixels in confocal images and movies [15••] of ER tubules would measure approximately 800 nm in length and 50 nm in diameter and contain an average of 4960 proteins (Fig. 2).
Representation of a protein array at the ER surface. ER-anchored protein molecules are each represented by a white dot. A total of 4960 protein molecules are displayed
For clarity of the further models, we do not display UGT85B1 and other soluble proteins or scaffold proteins that might mediate metabolon formation as discussed earlier.
Model Requirements and Constraints
In order to propose a model describing the current experimental data on the dhurrin metabolon, we have defined seven constraints essential for the validity of the model. We explore five models and validate them according to the constraints, where green “√” indicates that the constraint is fulfilled, the orange “(√)” indicates that the constraint may be satisfied in very specific conditions, and the red “X” indicates that the constraint cannot be explained by the configuration proposed.
Constraint #1: Based on quantitative proteomics data [15••], we calculated that for a total number of 100,000 ER protein molecules present on the ER, the number of protein molecules representing the dhurrin biosynthetic enzymes is as follows: 75 ± 3 POR2b, 238 ± 37 CYP79A1, 446 ± 181 CYP71E1 and 516 ± 95 other P450s. From S. bicolor total extract data, we calculated an abundance of 134 ± 2 UGT85B1 molecules within a total number of 100,000 proteins. Accordingly, we define constraint #1: ER membrane surfaces harboring a total number of 4960 proteins must display 4 ± 0 POR2b, 12 ± 2 CYP79A1, 22 ± 9 CYP71E1, 26 ± 5 other P450s and 4896 other ER proteins. P450s are in large excess compared to POR, whereas the UGT85B1 abundance is similar to that of CYP71E1. More than half of the total P450s are CYP79A1 and CYP71E1, as previously determined by spectroscopy [135]. Most importantly, all the dhurrin proteins are highly diluted in their native ER environment and even to a higher degree in context of total cellular protein. Even though CYP79A1, CYP71E1 and POR2b represented less than 0.8% of the total protein content of the ER membrane, they are able to interact specifically and form a dynamic metabolon.
Constraint #2: Using the Förster resonance energy transfer (FRET) equation with parameters of the eGFP-mRFP1 pair [15••], we have the relation between fluorophore distances and FRET values. FRET is calculated using the following equation:
with R 0 representing the Förster radius and R the distance between donor (eGFP) and acceptor (mRFP1). For the eGFP-mRFP1 couple, R 0 is equal to 4.7 nm [136]. Therefore, a measured FRET of 20% corresponds to an average distance between fluorescent proteins (center to center) of 5.9 nm, 5% FRET represents an average distance of 7.7 nm, 0.1% FRET represents an average distance of 14.9 nm, and finally, no FRET measured reflects distances above 15 nm. Considering proteins of an average diameter of 5 nm (average P450 diameter), we can record FRET between two proteins if no more than one protein is between the two targets. Above 5% FRET, no protein could interspace the target proteins. Thus, constraint #2a is as follows: High FRET values are measured only when proteins are interacting, forming oligomers (dimers or higher oligomers). In certain conditions, co-expression of a third partner protein enhanced the FRET values between the two other partners, e.g., increased FRET was observed between UGT85B1 and CYP79A1 fusion proteins upon co-expression of untagged CYP71E1 [15••]. This “additive” effect is constraint #2b. Increase in FRET average is the consequence of an increased spatial proximity of the two studied fluorescent proteins or an average of longer or more frequent interactions.
Constraint #3: Background FRET values measured between two unrelated ER proteins ranging from 0 to 5% [15••] define constraint #3. Non-interacting proteins display FRET values lower than 5%, when residing in the same micro-domain or meeting stochastically in a restricted volume as the ER membrane.
Constraint #4: When a protein A interacts with a protein B and concurrently protein B interacts with protein C, proteins A and C may be envisioned to be drawn very close to each other resulting in indirect FRET. High indirect FRET values were measured between non-partners at some very specific conditions [15••], and this phenomenon represents constraint #4.
Constraint #5: At the resolution of the confocal microscope, a uniform fluorescence emission was observed from the ER membrane system expressing the fluorescent-labeled dhurrin pathway enzymes [15••], although brighter areas indicated higher local concentrations of the tagged proteins. Hence, the dhurrin metabolon assembles in multiple clusters along the ER network. Thus, constraint #5 is as follows: the continuous fluorescence detected across the ER network, including local brighter areas.
Constraint #6: FCS experiments highlighted high dynamics of the target proteins, high variability of diffusion speeds, and decreased speeds of each target protein when partner proteins were co-expressed [15••], which defines constraint #6.
Constraint #7: Metabolomic data demonstrated efficient channeling upon metabolon formation without leakage of intermediates or by-products [15••] and describes constraint #7.
Model 1 (Fig. 3a) assumes a random distribution of all proteins and only fully meets the criteria of constraint #1. Part of other constraints could be fulfilled if proteins are highly dynamic and partners display strong affinity for each other. These two conditions increase the probability of protein-protein interactions without release of intermediates. Nevertheless, protein associations driven by strong protein-protein interactions will inevitably lead to formation of classical stable enzyme complexes and high-order aggregates. The entire dhurrin metabolon has never been purified in its ensemble, which points to transient and weak protein-protein interactions [15••]. Finally, formation of large aggregates would result in observable ER membrane surface areas depleted in target proteins, which disagrees with the continuous fluorescence observed. Consequently, this model was discarded.
Models 1, 2, and 3. a Model 1 of a random distribution of dynamic proteins. All the proteins are freely moving and only interact by random “hit-and-run” mechanism. b Model 2 of stable metabolons. This model involves pre-formed metabolons that are exchanged to the POR acting as a charging station. c Models 3a, b and c of proteins organized in large structures. Model 3a, proteins organized in highways across the ER tubule with POR trapped inside the highways. Model 3b, proteins organized in highways across the ER tubule with POR freely moving around the highways. Model 3c, proteins organized in aggregates with freely moving POR
Model 2 (Fig. 3b) represents an exchange of metabolons close to POR, which behaves as a hub for providing electrons for specific P450s. P450s not interacting with POR are not functional. To fulfill constraint #6, not all proteins of the same type can be permanently forming a metabolon, requiring non-associated proteins and thus an exchange between free and bound proteins. Constraint #4 requires functionally distinct metabolons in close proximity or large mixed complexes. However, such organization would inevitably lead to depletion of fluorescence or high heterogeneity in confocal images. Such large complexes are not in compliance with the required dynamic nature. Again, the dhurrin metabolon has never been purified in its ensemble, which points to weak protein-protein interactions. Consequently, this model was discarded.
Models 3a, b, and c display three different configurations of P450s organized in highways of large clusters (Fig. 3c). The constraints #1 and #5 cannot be satisfied at the same time if studied proteins are organized in large structures. With these kinds of large complexes composed of mixed metabolons, high FRET values would be recorded from unrelated proteins. To fulfill the constraint #6, some of the proteins studied have to exist in a free, non-metabolon-associated state. As explained previously, the dhurrin metabolon has never been purified in its ensemble, pointing to the formation of non-stable complexes. Consequently, this model was discarded.
Model 4 displays two models (Fig. 4a) revisiting the earlier model of P450s encircling PORs [137]. These models do not meet three of the experimental constraints. Firstly, if P450s involved in competitive metabolons are permanently associated around the same POR, this will result in high FRET between unrelated P450s, which was not detected in Laursen et al. [15••]. Secondly, free, non-associated proteins are required to satisfy constraint #6. Thus, continuous exchange of P450s around the PORs is expected. Thirdly, a repetitive similar protein organization across the ER network is a requisite for the observed continuous fluorescence. Consequently, we discard these models.
Models 4 and 5 of proteins organized as dynamic metabolons. a Model 4a, mixed metabolons are formed around the POR acting as a charging dock. Model 4b, POR encircled by one or mixed metabolons. b Model 5 of formation of specific transient and dynamic metabolon around the POR. Only one metabolon is present at a time t around the POR. This model implies a dynamic exchange of metabolon components around a POR acting as a charging station. Metabolons could be formed by oligomers of each component. Association and dissociation of metabolons are controlled upon cellular needs. Three arbitrary time points are shown in this figure
Model 5 proposes that specific P450s organize in dynamic metabolons around POR (Fig. 4b). To satisfy constraint #4, a continuous exchange of P450s around PORs is required, provoking association and dissociation of competitive metabolons. The exchange of P450 interactions between non-partners will, as observed, result in increased background FRET [15••]. Furthermore, restriction of PORs and P450s to micro-domains in the membrane will increase the probability of FRET between non-partners at the surface of ER. All the other constraints were satisfied; thus, we favor this model. This model satisfies observations of homo- and hetero-oligomerization of CYP79A1, CYP71E1 and UGT85B1: that CYP79A1 and CYP71E1 recruit UGT85B1, and that UGT85B1 interacts with both CYP79A1 and CYP71E1 but is not necessary for CYP79A1-CYP71E1 complex formation. UGT85B1 is situated close to the non-partner ER membrane proteins, CYP98A1 and POR2b, during association and dissociation of the metabolon, when CYP79A1 and CYP71E1 are co-expressed.
The model proposed here suggests that P450s and soluble partner enzymes interact with POR, which acts as a charging station possibly in concert with Cytb5. Lipid composition and non-catalytic proteins might guide metabolon formation and control P450 activity as discussed above. The existence of NADES might also limit leakage of metabolic intermediates by isolating the metabolon components from the cytosol.
Conclusion
On-demand formation of specialized metabolites enables plants to respond to environmental stresses through the sequential action of multiple enzymes. Fine-tuning of the metabolism is accomplished through dynamic metabolons to provide the necessary plasticity for plants to survive the continuum of biotic and abiotic challenges. The question remains, how such dynamic metabolons are organized, despite the facts that (1) P450 metabolons are localized at the dynamic ER surface, (2) metabolon components are highly diluted in plant cells, (3) the ER is highly crowded, and (4) the ER is a multifunctional and dynamic organelle site of protein production, lipid biosynthesis [138], auxin regulation [139], calcium homeostasis [140], and oil and protein body formation [141]. In this review, we argue that all proteins involved in metabolon formation must be tightly regulated and organized facilitating substrate channeling without release of metabolic intermediates. Our proposed model (model 5, Fig. 4b) integrates the current knowledge of the dhurrin metabolon. We propose that P450s interact and associate with POR to form transient platforms for recruitment of soluble partner enzymes. The lifetime of metabolons still remains largely unknown mainly due to technological limitations. A metabolon with changing composition and/or numbers of proteins has the potential to provide additional regulatory power under changing environmental and developmental situations.
Cellular metabolism is highly modular utilizing relatively few enzyme classes to generate an impressive diversity of metabolites. Therefore, we hypothesize that other biosynthetic pathways, especially those involved in formation of specialized metabolites, may be organized in a similar fashion as the dhurrin metabolon. Understanding the mechanisms that govern metabolon assembly has huge implications for synthetic biology approaches aiming to produce high-value plant specialized metabolites in heterologous hosts, which often suffer from low yield and release of toxic intermediates [6, 15••, 142, 143••]. Optimizing yield and titers may require co-expression of additional non-catalytic proteins, changing the lipid environment and intracellular metabolite profile.
References
Papers of particular interest, published recently, have been highlighted as: • Of importance •• Of major importance
•• Sweetlove LJ, Fernie AR. The spatial organization of metabolism within the plant cell. Annu Rev Plant Biol. 2013;64:723–46. An excellent review dealing with the aspects of macro- and micro-compartmentations of the plant metabolism.
• Laursen T, Moller BL, Bassard JE. Plasticity of specialized metabolism as mediated by dynamic metabolons. Trends Plant Sci. 2015;20:20–32. This review focuses on the factors that facilitate and stimulate the assembly of metabolons.
Moller BL. Dynamic metabolons. Science. 2010;330:1328–9.
Srere PA. Complexes of sequential metabolic enzymes. Annu Rev Biochem. 1987;56:89–124.
Bassard JE, Richert L, Geerinck J, Renault H, Duval F, Ullmann P, Schmitt M, Meyer E, Mutterer J, Boerjan W, et al. Protein-protein and protein-membrane associations in the lignin pathway. Plant Cell. 2012;24:4465–82.
Dastmalchi M, Facchini PJ. Plant metabolons assembled on demand. Science. 2016;354:829–30.
Dueber JE, Wu GC, Malmirchegini GR, Moon TS, Petzold CJ, Ullal AV, Prather KL, Keasling JD. Synthetic protein scaffolds provide modular control over metabolic flux. Nat Biotechnol. 2009;27:753–9.
Farre G, Blancquaert D, Capell T, Van Der Straeten D, Christou P, Zhu C. Engineering complex metabolic pathways in plants. Annu Rev Plant Biol. 2014;65:187–223.
Singleton C, Howard TP, Smirnoff N. Synthetic metabolons for metabolic engineering. J Exp Bot. 2014;65:1947–54.
Wlodarczyk A, Gnanasekaran T, Nielsen AZ, Zulu NN, Mellor SB, Luckner M, Thofner JF, Olsen CE, Mottawie MS, Burow M, et al. Metabolic engineering of light-driven cytochrome P450 dependent pathways into Synechocystis sp. PCC 6803. Metab Eng. 2016;33:1–11.
• Lassen LM, Nielsen AZ, Ziersen B, Gnanasekaran T, Moller BL, Jensen PE. Redirecting photosynthetic electron flow into light-driven synthesis of alternative products including high-value bioactive natural compounds. ACS Synth Biol. 2014;3:1–12. This review presents approaches taken to channel the reducing power of photosystem I into driving the catalytic cycle of cytochrome P450s.
Jensen K, Jensen PE, Moller BL. Light-driven chemical synthesis. Trends Plant Sci. 2012;17:60–3.
Stafford HA. The metabolism of aromatic compounds. Annu Rev Plant Physiol. 1974;25:459–86.
Jorgensen K, Rasmussen AV, Morant M, Nielsen AH, Bjarnholt N, Zagrobelny M, Bak S, Moller BL. Metabolon formation and metabolic channeling in the biosynthesis of plant natural products. Curr Opin Plant Biol. 2005;8:280–91.
•• Laursen T, Borch J, Knudsen C, Bavishi K, Torta F, Martens HJ, Silvestro D, Hatzakis NS, Wenk MR, Dafforn TR, et al. Characterization of a dynamic metabolon producing the defense compound dhurrin in sorghum. Science. 2016;354:890–3. This study shows that association of two P450s enables recruitment of a soluble enzyme to form a dynamic metabolon and demonstrates a functional link between protein interactions and metabolite production. This study also points to the importance of the lipid environment on metabolon activity.
Achnine L, Blancaflor EB, Rasmussen S, Dixon RA. Colocalization of L-phenylalanine ammonia-lyase and cinnamate 4-hydroxylase for metabolic channeling in phenylpropanoid biosynthesis. Plant Cell. 2004;16:3098–109.
Burbulis IE, Winkel-Shirley B. Interactions among enzymes of the Arabidopsis flavonoid biosynthetic pathway. Proc Natl Acad Sci U S A. 1999;96:12929–34.
Crosby KC, Pietraszewska-Bogiel A, Gadella Jr TW, Winkel BS. Forster resonance energy transfer demonstrates a flavonoid metabolon in living plant cells that displays competitive interactions between enzymes. FEBS Lett. 2011;585:2193–8.
• Dastmalchi M, Bernards MA, Dhaubhadel S. Twin anchors of the soybean isoflavonoid metabolon: evidence for tethering of the complex to the endoplasmic reticulum by IFS and C4H. Plant J. 2016;85:689–706. This study provides evidence for an isoflavonoid metabolon. Interaction studies identified several candidates of various biosynthetic and structural elements that may be involved in isoflavonoid production pointing to the complex regulation of the flavonoid pathway.
Waki T, Yoo D, Fujino N, Mameda R, Denessiouk K, Yamashita S, Motohashi R, Akashi T, Aoki T, Ayabe S, et al. Identification of protein-protein interactions of isoflavonoid biosynthetic enzymes with 2-hydroxyisoflavanone synthase in soybean (Glycine max (L.) Merr.). Biochem Biophys Res Commun. 2016;469:546–51.
Lallemand B, Erhardt M, Heitz T, Legrand M. Sporopollenin biosynthetic enzymes interact and constitute a metabolon localized to the endoplasmic reticulum of tapetum cells. Plant Physiol. 2013;162:616–25.
Bulutoglu B, Garcia KE, Wu F, Minteer SD, Banta S. Direct evidence for metabolon formation and substrate channeling in recombinant TCA cycle enzymes. ACS Chem Biol. 2016;11:2847–53. This study explores structural determinants of substrate channeling in vitro via the use of several mutants of the TCA cycle enzymes.
• Wu F, Minteer S. Krebs cycle metabolon: structural evidence of substrate channeling revealed by cross-linking and mass spectrometry. Angew Chem Int Ed Engl. 2015;54:1851–4. This study demonstrates the existence of the Krebs cycle metabolon and proposes the organization of the enzymes involved.
Graciet E, Lebreton S, Gontero B. Emergence of new regulatory mechanisms in the Benson-Calvin pathway via protein-protein interactions: a glyceraldehyde-3-phosphate dehydrogenase/CP12/phosphoribulokinase complex. J Exp Bot. 2004;55:1245–54.
Li H, Ban Z, Qin H, Ma L, King AJ, Wang G. A heteromeric membrane-bound prenyltransferase complex from hop catalyzes three sequential aromatic prenylations in the bitter acid pathway. Plant Physiol. 2015;167:650–9.
Kwiatkowska M, Polit JT, Stepinski D, Poplonska K, Wojtczak A, Domiotanguez E, Heredia A. Lipotubuloids in ovary epidermis of Ornithogalum umbellatum act as metabolons: suggestion of the name ‘lipotubuloid metabolon’. J Exp Bot. 2015;66:1157–63.
Troncoso-Ponce MA, Nikovics K, Marchive C, Lepiniec L, Baud S. New insights on the organization and regulation of the fatty acid biosynthetic network in the model higher plant Arabidopsis thaliana. Biochimie. 2016;120:3–8.
Panicot M, Minguet EG, Ferrando A, Alcazar R, Blazquez MA, Carbonell J, Altabella T, Koncz C, Tiburcio AF. A polyamine metabolon involving aminopropyl transferase complexes in Arabidopsis. Plant Cell. 2002;14:2539–51.
Szecowka M, Heise R, Tohge T, Nunes-Nesi A, Vosloh D, Huege J, Feil R, Lunn J, Nikoloski Z, Stitt M, et al. Metabolic fluxes in an illuminated Arabidopsis rosette. Plant Cell. 2013;25:694–714.
Hayashi K, Nakamura S, Fukunaga S, Nishimura T, Jenness MK, Murphy AS, Motose H, Nozaki H, Furutani M, Aoyama T. Auxin transport sites are visualized in planta using fluorescent auxin analogs. Proc Natl Acad Sci U S A. 2014;111:11557–62.
•• Hawes C, Kiviniemi P, Kriechbaumer V. The endoplasmic reticulum: a dynamic and well-connected organelle. J Integr Plant Biol. 2015;57:50–62. This review explores all the major features of the ER network acting as a dynamic intracellular highway connected to all the organelles of the cell.
Kriechbaumer V, Botchway SW, Hawes C. Localization and interactions between Arabidopsis auxin biosynthetic enzymes in the TAA/YUC-dependent pathway. J Exp Bot. 2016;67:4195–207.
Luu W, Hart-Smith G, Sharpe LJ, Brown AJ. The terminal enzymes of cholesterol synthesis, DHCR24 and DHCR7, interact physically and functionally. J Lipid Res. 2015;56:888–97.
• Kyoung M, Russell SJ, Kohnhorst CL, Esemoto NN, An S. Dynamic architecture of the purinosome involved in human de novo purine biosynthesis. Biochemistry. 2015;54:870–80. Quantitative analyses of the purinosome enzymes provide clues for the dynamic architecture of the purinosome metabolon assembly.
Cheung CW, Cohen NS, Raijman L. Channeling of urea cycle intermediates in situ in permeabilized hepatocytes. J Biol Chem. 1989;264:4038–44.
Shatalin K, Lebreton S, Rault-Leonardon M, Velot C, Srere PA. Electrostatic channeling of oxaloacetate in a fusion protein of porcine citrate synthase and porcine mitochondrial malate dehydrogenase. Biochemistry. 1999;38:881–9.
Vertessy B, Ovadi J. A simple approach to detect active-site-directed enzyme-enzyme interactions. The aldolase/glycerol-phosphate-dehydrogenase enzyme system. Eur J Biochem. 1987;164:655–9.
Clegg JS, Jackson SA. Glucose metabolism and the channeling of glycolytic intermediates in permeabilized L-929 cells. Arch Biochem Biophys. 1990;278:452–60.
Araiza-Olivera D, Chiquete-Felix N, Rosas-Lemus M, Sampedro JG, Pena A, Mujica A, Uribe-Carvajal S. A glycolytic metabolon in Saccharomyces cerevisiae is stabilized by F-actin. FEBS J. 2013;280:3887–905.
Srere PA. The metabolon. Trends Biochem Sci. 1985;10:109–10.
Winkel BSJ. Metabolic channeling in plants. Annu Rev Plant Biol. 2004;55:85–107.
Ralston L, Yu O. Metabolons involving plant cytochrome P450s. Phytochem Rev. 2006;5:459–72.
Bak S, Beisson F, Bishop G, Hamberger B, Höfer R, Paquette S, Werck-Reichhart D. Cytochromes P450. The Arabidopsis Book / American Society of Plant Biologists. 2011;9:e0144.
Griffing LR, Lin C, Perico C, White RR, Sparkes I: Plant ER geometry and dynamics: biophysical and cytoskeletal control during growth and biotic response. Protoplasma 2016.
Guigas G, Weiss M: Effects of protein crowding on membrane systems. Biochim Biophys Acta 2015.
Hrazdina G, Wagner GJ. Metabolic pathways as enzyme complexes: evidence for the synthesis of phenylpropanoids and flavonoids on membrane associated enzyme complexes. Arch Biochem Biophys. 1985;237:88–100.
Rasmussen S, Dixon RA. Transgene-mediated and elicitor-induced perturbation of metabolic channeling at the entry point into the phenylpropanoid pathway. Plant Cell. 1999;11:1537–52.
Ro DK, Douglas CJ. Reconstitution of the entry point of plant phenylpropanoid metabolism in yeast (Saccharomyces cerevisiae): implications for control of metabolic flux into the phenylpropanoid pathway. J Biol Chem. 2004;279:2600–7.
Chen HC, Li Q, Shuford CM, Liu J, Muddiman DC, Sederoff RR, Chiang VL. Membrane protein complexes catalyze both 4- and 3-hydroxylation of cinnamic acid derivatives in monolignol biosynthesis. Proc Natl Acad Sci U S A. 2011;108:21253–8.
Nielsen KA, Tattersall DB, Jones PR, Moller BL. Metabolon formation in dhurrin biosynthesis. Phytochemistry. 2008;69:88–98.
•• Reed JR, Backes WL. The functional effects of physical interactions involving cytochromes P450: putative mechanisms of action and the extent of these effects in biological membranes. Drug Metab Rev. 2016;48:453–69. This review focuses on the formation of P450-P450 complexes suggesting that P450s can be influenced by the interaction with other P450s.
Backes WL, Kelley RW. Organization of multiple cytochrome P450s with NADPH-cytochrome P450 reductase in membranes. Pharmacol Ther. 2003;98:221–33.
Ro DK, Ehlting J, Douglas CJ. Cloning, functional expression, and subcellular localization of multiple NADPH-cytochrome P450 reductases from hybrid poplar. Plant Physiol. 2002;130:1837–51.
Denisov IG, Baas BJ, Grinkova YV, Sligar SG. Cooperativity in cytochrome P450 3A4: linkages in substrate binding, spin state, uncoupling, and product formation. J Biol Chem. 2007;282:7066–76.
Miwa GT, Lu AY. The association of cytochrome P-450 and NADPH-cytochrome P-450 reductase in phospholipid membranes. Arch Biochem Biophys. 1984;234:161–6.
Jamakhandi AP, Kuzmic P, Sanders DE, Miller GP. Global analysis of protein-protein interactions reveals multiple CYP2E1-reductase complexes. Biochemistry. 2007;46:10192–201.
Reed JR, Cawley GF, Backes WL. Inhibition of cytochrome P450 1A2-mediated metabolism and production of reactive oxygen species by heme oxygenase-1 in rat liver microsomes. Drug Metab Lett. 2011;5:6–16.
Estabrook RW, Franklin MR, Cohen B, Shigamatzu A, Hildebrandt AG. Biochemical and genetic factors influencing drug metabolism. Influence of hepatic microsomal mixed function oxidation reactions on cellular metabolic control. Metabolism. 1971;20:187–99.
Watanabe J, Asaka Y, Fujimoto S, Kanamura S. Densities of NADPH-ferrihemoprotein reductase and cytochrome P-450 molecules in the endoplasmic reticulum membrane of rat hepatocytes. J Histochem Cytochem. 1993;41:43–9.
Shephard EA, Phillips IR, Bayney RM, Pike SF, Rabin BR. Quantification of NADPH: cytochrome P-450 reductase in liver microsomes by a specific radioimmunoassay technique. Biochem J. 1983;211:333–40.
Gomes AM, Winter S, Klein K, Turpeinen M, Schaeffeler E, Schwab M, Zanger UM. Pharmacogenomics of human liver cytochrome P450 oxidoreductase: multifactorial analysis and impact on microsomal drug oxidation. Pharmacogenomics. 2009;10:579–99.
Laursen T, Singha A, Rantzau N, Tutkus M, Borch J, Hedegard P, Stamou D, Moller BL, Hatzakis NS. Single molecule activity measurements of cytochrome P450 oxidoreductase reveal the existence of two discrete functional states. ACS Chem Biol. 2014;9:630–4.
Laursen T, Jensen K, Moller BL. Conformational changes of the NADPH-dependent cytochrome P450 reductase in the course of electron transfer to cytochromes P450. Biochim Biophys Acta. 1814;2011:132–8.
Wadsater M, Laursen T, Singha A, Hatzakis NS, Stamou D, Barker R, Mortensen K, Feidenhans’l R, Moller BL, Cardenas M. Monitoring shifts in the conformation equilibrium of the membrane protein cytochrome P450 reductase (POR) in nanodiscs. J Biol Chem. 2012;287:34596–603.
Andersen TB, Hansen NB, Laursen T, Weitzel C, Simonsen HT. Evolution of NADPH-cytochrome P450 oxidoreductases (POR) in Apiales—POR 1 is missing. Mol Phylogenet Evol. 2016;98:21–8.
Parage C, Foureau E, Kellner F, Burlat V, Mahroug S, Lanoue A, Duge de Bernonville T, Londono MA, Carqueijeiro I, Oudin A, et al. Class II cytochrome P450 reductase governs the biosynthesis of alkaloids. Plant Physiol. 2016;172:1563–77.
Koopmann E, Hahlbrock K. Differentially regulated NADPH:cytochrome P450 oxidoreductases in parsley. Proc Natl Acad Sci U S A. 1997;94:14954–9.
Mizutani M, Ohta D. Two isoforms of NADPH:cytochrome P450 reductase in Arabidopsis thaliana. Gene structure, heterologous expression in insect cells, and differential regulation. Plant Physiol. 1998;116:357–67.
Schwarz H, Liu B, Peters S, Barillas W, Beerhues L. Purification, cDNA cloning and functional expression of NADPH-cytochrome P450 reductase from Centaurium erythraea cell cultures. Plant Biol (Stuttg). 2009;11:300–6.
Rana S, Lattoo SK, Dhar N, Razdan S, Bhat WW, Dhar RS, Vishwakarma R. NADPH-cytochrome P450 reductase: molecular cloning and functional characterization of two paralogs from Withania somnifera (L.) Dunal. PLoS One. 2013;8:e57068.
Szczesna-Skorupa E, Mallah B, Kemper B. Fluorescence resonance energy transfer analysis of cytochromes P450 2C2 and 2E1 molecular interactions in living cells. J Biol Chem. 2003;278:31269–76.
Szczesna-Skorupa E, Kemper B. BAP31 is involved in the retention of cytochrome P450 2C2 in the endoplasmic reticulum. J Biol Chem. 2006;281:4142–8.
Ducret A, Nguyen M, Breckenridge DG, Shore GC. The resident endoplasmic reticulum protein, BAP31, associates with gamma-actin and myosin B heavy chain. Eur J Biochem. 2003;270:342–9.
Mo C, Bard M. Erg28p is a key protein in the yeast sterol biosynthetic enzyme complex. J Lipid Res. 2005;46:1991–8.
Kriechbaumer V, Botchway SW, Slade SE, Knox K, Frigerio L, Oparka K, Hawes C. Reticulomics: protein-protein interaction studies with two plasmodesmata-localized reticulon family proteins identify binding partners enriched at plasmodesmata, endoplasmic reticulum, and the plasma membrane. Plant Physiol. 2015;169:1933–45.
Hildebrandt A, Estabrook RW. Evidence for the participation of cytochrome b5 in hepatic microsomal mixed-function oxidation reactions. Arch Biochem Biophys. 1971;143:66–79.
Zhang H, Gao N, Liu T, Fang Y, Qi B, Wen Q, Zhou J, Jia L, Qiao H. Effect of cytochrome b5 content on the activity of polymorphic CYP1A2, 2B6, and 2E1 in human liver microsomes. PLoS One. 2015;10:e0128547.
Bhatt MR, Khatri Y, Rodgers RJ, Martin LL: Role of cytochrome b5 in the modulation of the enzymatic activities of cytochrome P450 17alpha-hydroxylase/17,20-lyase (P450 17A1). J Steroid Biochem Mol Biol 2016.
Stiborova M, Indra R, Moserova M, Frei E, Schmeiser HH, Kopka K, Philips DH, Arlt VM. NADH:cytochrome b5 reductase and cytochrome b5 can act as sole electron donors to human cytochrome P450 1A1-mediated oxidation and DNA adduct formation by benzo[a]pyrene. Chem Res Toxicol. 2016;29:1325–34.
Vik D, Crocoll C, Andersen TG, Burow M, Halkier BA. CB5C affects the glucosinolate profile in Arabidopsis thaliana. Plant Signal Behav. 2016;11:e1160189.
de Vetten N, ter Horst J, van Schaik HP, de Boer A, Mol J, Koes R. A cytochrome b5 is required for full activity of flavonoid 3′, 5′-hydroxylase, a cytochrome P450 involved in the formation of blue flower colors. Proc Natl Acad Sci U S A. 1999;96:778–83.
Kumar R, Wallis JG, Skidmore C, Browse J. A mutation in Arabidopsis cytochrome b5 reductase identified by high-throughput screening differentially affects hydroxylation and desaturation. Plant J. 2006;48:920–32.
Kumar R, Tran LS, Neelakandan AK, Nguyen HT. Higher plant cytochrome b5 polypeptides modulate fatty acid desaturation. PLoS One. 2012;7:e31370.
Nagano M, Ihara-Ohori Y, Imai H, Inada N, Fujimoto M, Tsutsumi N, Uchimiya H, Kawai-Yamada M. Functional association of cell death suppressor, Arabidopsis Bax inhibitor-1, with fatty acid 2-hydroxylation through cytochrome b(5). Plant J. 2009;58:122–34.
Li Y, Li LL, Fan RC, Peng CC, Sun HL, Zhu SY, Wang XF, Zhang LY, Zhang DP. Arabidopsis sucrose transporter SUT4 interacts with cytochrome b5-2 to regulate seed germination in response to sucrose and glucose. Mol Plant. 2012;5:1029–41.
Chang J, Clay JM, Chang C. Association of cytochrome b5 with ETR1 ethylene receptor signaling through RTE1 in Arabidopsis. Plant J. 2014;77:558–67.
•• Paddon CJ, Westfall PJ, Pitera DJ, Benjamin K, Fisher K, McPhee D, Leavell MD, Tai A, Main A, Eng D, et al. High-level semi-synthetic production of the potent antimalarial artemisinin. Nature. 2013;496:528–32. This research article demonstrates the complete biosynthetic pathway leading to artemisinic acid and presents several parameters (cytochrome b5 , chemicals, promoters) which impact specialized metabolite production in heterologous hosts.
Sparkes I, Tolley N, Aller I, Svozil J, Osterrieder A, Botchway S, Mueller C, Frigerio L, Hawes C. Five Arabidopsis reticulon isoforms share endoplasmic reticulum location, topology, and membrane-shaping properties. Plant Cell. 2010;22:1333–43.
Levy A, Zheng JY, Lazarowitz SG. Synaptotagmin SYTA forms ER-plasma membrane junctions that are recruited to plasmodesmata for plant virus movement. Curr Biol. 2015;25:2018–25.
Uchiyama A, Shimada-Beltran H, Levy A, Zheng JY, Javia PA, Lazarowitz SG. The Arabidopsis synaptotagmin SYTA regulates the cell-to-cell movement of diverse plant viruses. Front Plant Sci. 2014;5:584.
Siao W, Wang P, Voigt B, Hussey PJ, Baluska F. Arabidopsis SYT1 maintains stability of cortical endoplasmic reticulum networks and VAP27-1-enriched endoplasmic reticulum-plasma membrane contact sites. J Exp Bot. 2016;67:6161–71.
Stefano G, Renna L, Lai Y, Slabaugh E, Mannino N, Buono RA, Otegui MS, Brandizzi F. ER network homeostasis is critical for plant endosome streaming and endocytosis. Cell Discov. 2015;1:15033.
Browman DT, Resek ME, Zajchowski LD, Robbins SM. Erlin-1 and erlin-2 are novel members of the prohibitin family of proteins that define lipid-raft-like domains of the ER. J Cell Sci. 2006;119:3149–60.
Hoegg MB, Browman DT, Resek ME, Robbins SM. Distinct regions within the erlins are required for oligomerization and association with high molecular weight complexes. J Biol Chem. 2009;284:7766–76.
St-Pierre B, Vazquez-Flota FA, De Luca V. Multicellular compartmentation of Catharanthus roseus alkaloid biosynthesis predicts intercellular translocation of a pathway intermediate. Plant Cell. 1999;11:887–900.
Pollard M, Beisson F, Li Y, Ohlrogge JB. Building lipid barriers: biosynthesis of cutin and suberin. Trends Plant Sci. 2008;13:236–46.
Bonawitz ND, Chapple C. The genetics of lignin biosynthesis: connecting genotype to phenotype. Annu Rev Genet. 2010;44:337–63.
Pulido P, Perello C, Rodriguez-Concepcion M. New insights into plant isoprenoid metabolism. Mol Plant. 2012;5:964–7.
Sun P, Schuurink RC, Caissard JC, Hugueney P, Baudino S. My way: noncanonical biosynthesis pathways for plant volatiles. Trends Plant Sci. 2016;21:884–94.
Qin M, Tian T, Xia S, Wang Z, Song L, Yi B, Wen J, Shen J, Ma C, Fu T, et al. Heterodimer formation of BnPKSA or BnPKSB with BnACOS5 constitutes a multienzyme complex in tapetal cells and is involved in male reproductive development in Brassica napus. Plant Cell Physiol. 2016;57:1643–56.
Block MA, Jouhet J. Lipid trafficking at endoplasmic reticulum-chloroplast membrane contact sites. Curr Opin Cell Biol. 2015;35:21–9.
Wang P, Richardson C, Hawkins TJ, Sparkes I, Hawes C, Hussey PJ. Plant VAP27 proteins: domain characterization, intracellular localization and role in plant development. New Phytol. 2016;210:1311–26.
Sparkes IA, Frigerio L, Tolley N, Hawes C. The plant endoplasmic reticulum: a cell-wide web. Biochem J. 2009;423:145–55.
Sparkes IA, Ketelaar T, de Ruijter NC, Hawes C. Grab a Golgi: laser trapping of Golgi bodies reveals in vivo interactions with the endoplasmic reticulum. Traffic. 2009;10:567–71.
Mehrshahi P, Johnny C, DellaPenna D. Redefining the metabolic continuity of chloroplasts and ER. Trends Plant Sci. 2014;19:501–7.
Wang P, Hawkins TJ, Richardson C, Cummins I, Deeks MJ, Sparkes I, Hawes C, Hussey PJ. The plant cytoskeleton, NET3C, and VAP27 mediate the link between the plasma membrane and endoplasmic reticulum. Curr Biol. 2014;24:1397–405.
Jaipargas EA, Barton KA, Mathur N, Mathur J. Mitochondrial pleomorphy in plant cells is driven by contiguous ER dynamics. Front Plant Sci. 2015;6:783.
Sinclair AM, Trobacher CP, Mathur N, Greenwood JS, Mathur J. Peroxule extension over ER-defined paths constitutes a rapid subcellular response to hydroxyl stress. Plant J. 2009;59:231–42.
Andersson MX, Goksor M, Sandelius AS. Optical manipulation reveals strong attracting forces at membrane contact sites between endoplasmic reticulum and chloroplasts. J Biol Chem. 2007;282:1170–4.
•• Mehrshahi P, Stefano G, Andaloro JM, Brandizzi F, Froehlich JE, DellaPenna D. Transorganellar complementation redefines the biochemical continuity of endoplasmic reticulum and chloroplasts. Proc Natl Acad Sci U S A. 2013;110:12126–31. This study redefines the understanding of the biochemical continuity of the ER-chloroplast and suggests formation of a hemi-fused membrane between both organelles for sharing of metabolic intermediates of organelle-spanning pathways.
Chuong SD, Good AG, Taylor GJ, Freeman MC, Moorhead GB, Muench DG. Large-scale identification of tubulin-binding proteins provides insight on subcellular trafficking, metabolic channeling, and signaling in plant cells. Mol Cell Proteomics. 2004;3:970–83.
Matsushima R, Hayashi Y, Yamada K, Shimada T, Nishimura M, Hara-Nishimura I. The ER body, a novel endoplasmic reticulum-derived structure in Arabidopsis. Plant Cell Physiol. 2003;44:661–6.
Yamada K, Nagano AJ, Nishina M, Hara-Nishimura I, Nishimura M. NAI2 is an endoplasmic reticulum body component that enables ER body formation in Arabidopsis thaliana. Plant Cell. 2008;20:2529–40.
Ogasawara K, Yamada K, Christeller JT, Kondo M, Hatsugai N, Hara-Nishimura I, Nishimura M. Constitutive and inducible ER bodies of Arabidopsis thaliana accumulate distinct beta-glucosidases. Plant Cell Physiol. 2009;50:480–8.
Takemoto D, Jones DA, Hardham AR. GFP-tagging of cell components reveals the dynamics of subcellular re-organization in response to infection of Arabidopsis by oomycete pathogens. Plant J. 2003;33:775–92.
Takemoto D, Jones DA, Hardham AR. Re-organization of the cytoskeleton and endoplasmic reticulum in the Arabidopsis pen1-1 mutant inoculated with the non-adapted powdery mildew pathogen, Blumeria graminis f. sp. hordei. Mol Plant Pathol. 2006;7:553–63.
• Brignac-Huber LM, Park JW, Reed JR, Backes WL. Cytochrome P450 organization and function are modulated by endoplasmic reticulum phospholipid heterogeneity. Drug Metab Dispos. 2016;44:1859–66. This review presents data on P450s and POR that are selectively associated with specific phospholipids and that these lipid-protein interactions influence P450 activity.
Li Q, Shen W, Zheng Q, Fowler DB, Zou J. Adjustments of lipid pathways in plant adaptation to temperature stress. Plant Signal Behav. 2016;11:e1058461.
Zajchowski LD, Robbins SM. Lipid rafts and little caves. Compartmentalized signalling in membrane microdomains. Eur J Biochem. 2002;269:737–52.
Brignac-Huber L, Reed JR, Backes WL. Organization of NADPH-cytochrome P450 reductase and CYP1A2 in the endoplasmic reticulum—microdomain localization affects monooxygenase function. Mol Pharmacol. 2011;79:549–57.
Das A, Sligar SG. Modulation of the cytochrome P450 reductase redox potential by the phospholipid bilayer. Biochemistry. 2009;48:12104–12.
Reed JR, Backes WL. Formation of P450.P450 complexes and their effect on P450 function. Pharmacol Ther. 2012;133:299–310.
Stier A, Sackmann E. Spin labels as enzyme substrates. Heterogeneous lipid distribution in liver microsomal membranes. Biochim Biophys Acta. 1973;311:400–8.
Kim KH, Kim DH, Jang HH, Kim M, Kim DH, Kim JS, Kim JI, Chae HZ, Ahn T, Yun CH. Lateral segregation of anionic phospholipids in model membranes induced by cytochrome P450 2B1: bi-directional coupling between CYP2B1 and anionic phospholipid. Arch Biochem Biophys. 2007;468:226–33.
•• Hyman AA, Weber CA, Julicher F. Liquid-liquid phase separation in biology. Annu Rev Cell Dev Biol. 2014;30:39–58. This review points to the evidence that non-membrane compartments are formed by phase separation from the cytoplasm.
Hyman AA, Simons K. Cell biology. Beyond oil and water—phase transitions in cells. Science. 2012;337:1047–9.
• Dai Y, van Spronsen J, Witkamp GJ, Verpoorte R, Choi YH. Natural deep eutectic solvents as new potential media for green technology. Anal Chim Acta. 2013;766:61–8. This study explores the chemical and physical properties of NADES and tests NADES as a green solvent.
Dai Y, Witkamp GJ, Verpoorte R, Choi YH. Natural deep eutectic solvents as a new extraction media for phenolic metabolites in Carthamus tinctorius L. Anal Chem. 2013;85:6272–8.
Choi YH, van Spronsen J, Dai Y, Verberne M, Hollmann F, Arends IW, Witkamp GJ, Verpoorte R. Are natural deep eutectic solvents the missing link in understanding cellular metabolism and physiology? Plant Physiol. 2011;156:1701–5.
Dai Y, Verpoorte R, Choi YH. Natural deep eutectic solvents providing enhanced stability of natural colorants from safflower (Carthamus tinctorius). Food Chem. 2014;159:116–21.
Griffing LR. Networking in the endoplasmic reticulum. Biochem Soc Trans. 2010;38:747–53.
Hu J, Shibata Y, Voss C, Shemesh T, Li Z, Coughlin M, Kozlov MM, Rapoport TA, Prinz WA. Membrane proteins of the endoplasmic reticulum induce high-curvature tubules. Science. 2008;319:1247–50.
Park SH, Blackstone C. Further assembly required: construction and dynamics of the endoplasmic reticulum network. EMBO Rep. 2010;11:515–21.
Bouchekhima AN, Frigerio L, Kirkilionis M. Geometric quantification of the plant endoplasmic reticulum. J Microsc. 2009;234:158–72.
Sibbesen O, Koch B, Halkier BA, Moller BL. Isolation of the heme-thiolate enzyme cytochrome P-450TYR, which catalyzes the committed step in the biosynthesis of the cyanogenic glucoside dhurrin in Sorghum bicolor (L.) Moench. Proc Natl Acad Sci U S A. 1994;91:9740–4.
Peter M, Ameer-Beg SM, Hughes MK, Keppler MD, Prag S, Marsh M, Vojnovic B, Ng T. Multiphoton-FLIM quantification of the EGFP-mRFP1 FRET pair for localization of membrane receptor-kinase interactions. Biophys J. 2005;88:1224–37.
Peterson JA, Ebel RE, O’Keeffe DH, Matsubara T, Estabrook RW. Temperature dependence of cytochrome P-450 reduction. A model for NADPH-cytochrome P-450 reductase:cytochrome P-450 interaction. J Biol Chem. 1976;251:4010–6.
Wallis JG, Browse J. Lipid biochemists salute the genome. Plant J. 2010;61:1092–106.
Friml J, Jones AR. Endoplasmic reticulum: the rising compartment in auxin biology. Plant Physiol. 2010;154:458–62.
Bonza MC, Loro G, Behera S, Wong A, Kudla J, Costa A. Analyses of Ca2+ accumulation and dynamics in the endoplasmic reticulum of Arabidopsis root cells using a genetically encoded cameleon sensor. Plant Physiol. 2013;163:1230–41.
Herman EM. Endoplasmic reticulum bodies: solving the insoluble. Curr Opin Plant Biol. 2008;11:672–9.
Wilson SA, Cummings EM, Roberts SC. Multi-scale engineering of plant cell cultures for promotion of specialized metabolism. Curr Opin Biotechnol. 2014;29:163–70.
•• Galanie S, Thodey K, Trenchard IJ, Filsinger Interrante M, Smolke CD. Complete biosynthesis of opioids in yeast. Science. 2015;349:1095–100. This study presents the complete biosynthesis of opiates in a heterologous host using more than 20 heterologous genes. This work also demonstrates the complexity to produce specialized metabolites in heterologous hosts and the need to engineer enzymes and optimize pathway and strain.
Acknowledgments
This research was supported by the VILLUM Research Center “Plant Plasticity” and by the Center for Synthetic Biology “bioSYNergy” (UCPH Excellence Programme for Interdisciplinary Research). T.L. is the recipient of a fellowship awarded by the VILLUM Foundation (Project No: 95-300-73023).
Author information
Authors and Affiliations
Corresponding author
Ethics declarations
Conflict of Interest
The authors declare that they have no conflict of interest.
Human and Animal Rights and Informed Consent
This article does not contain any studies with human or animal subjects performed by any of the authors.
Additional information
This article is part of the Topical Collection on Enhancing Agricultural Production
Rights and permissions
Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.
About this article
Cite this article
Bassard, JE., Møller, B.L. & Laursen, T. Assembly of Dynamic P450-Mediated Metabolons—Order Versus Chaos. Curr Mol Bio Rep 3, 37–51 (2017). https://doi.org/10.1007/s40610-017-0053-y
Published:
Issue Date:
DOI: https://doi.org/10.1007/s40610-017-0053-y