Synthetic Protein Scaffolding at Biological Membranes

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Review

Synthetic Protein Scaffolding at Biological Membranes James B.Y.H. Behrendorff,1,@ Guillem Borra`s-Gas,1 and Mathias Pribil1,@,* Protein scaffolding is a natural phenomenon whereby proteins colocalize into macromolecular complexes via specific protein–protein interactions. In the case of metabolic enzymes, protein scaffolding drives metabolic flux through specific pathways by colocalizing enzyme active sites. Synthetic protein scaffolding is increasingly used as a mechanism to improve product specificity and yields in metabolic engineering projects. To date, synthetic scaffolding has focused primarily on soluble enzyme systems, but many metabolic pathways for high-value secondary metabolites depend on membrane-bound enzymes. The compositional diversity of biological membranes and general challenges associated with modifying membrane proteins complicate scaffolding with membrane-requiring enzymes. Several recent studies have introduced new approaches to protein scaffolding at membrane surfaces, with notable success in improving product yields from specific metabolic pathways.

Challenges in Using Protein Scaffolding to Enhance Biological Membranes A vibrant field of research has emerged focused on studying and engineering the protein–protein interactions that drive post-translational formation of enzyme complexes. In nature, enzymes that belong to a common metabolic pathway and form complexes to improve metabolic flux (see Glossary) are known as metabolons. These naturally occurring complexes promote substrate channeling [1]: the transfer of metabolic intermediates between sequential enzyme active sites, minimizing the diffusive loss of those intermediates. An archetypical example from nature is the prokaryotic tryptophan synthase, which is a two-subunit enzyme complex that converts indole-3-glycerol-phosphate to tryptophan via an indole intermediate. The a and b subunits of the tryptophan synthase complex combine to form an enclosed hydrophobic tunnel between the first and second active sites [2]. This configuration facilitates extremely rapid transfer of the indole intermediate from the a to the b subunit [3] and results in conversion of indole-3-glycerol-phosphate to tryptophan with a yield close to 100%. Although we cannot yet engineer tailored proteinaceous tunnels between enzyme active sites, synthetic protein scaffolding aims to emulate these substrate-channeling effects in engineered metabolic pathways.

Highlights Protein scaffolding techniques are increasingly being used to create synthetic protein complexes bound to biological membranes. Scaffolding tags can be used to recruit soluble enzymes or complexes to specific membrane domains. Synthetic transport metabolons, where enzymes are scaffolded to transmembrane transporters, are a powerful means to control metabolite fate upstream of central carbon metabolism. New membrane-integral scaffolding tools facilitate the design of synthetic complexes of membranerequiring enzymes.

Early synthetic scaffolds improved the production of mevalonate and glucaric acid in Saccharomyces cerevisiae [4], and demonstrated that artificial enzyme colocalization to improve metabolic flux is a generalizable principle. Protein scaffolding is particularly relevant to metabolic engineering in cases where several metabolic pathways compete for a common intermediate, because the yield of the desired product can be increased by minimizing the loss of intermediates to metabolism by other enzymes, a key concept in substrate channeling. Substrate channeling requires that enzymes are organized such that an intermediate metabolite is more likely to interact with the next enzyme in the desired metabolic pathway than to diffuse into the bulk aqueous phase [5,6]. Preventing intermediates from diffusing throughout the cell prevents them from being metabolized by competing pathways, thereby increasing product yields. Given that the rate of metabolite diffusion in the bulk aqueous phase of the cell is typically greater than enzyme reaction rates [7], substrate channeling is most effective when the rate-limiting enzyme is not the final step in the scaffolded metabolic pathway. Typically, enzymes should be positioned within 1 nm of each other to optimize the likelihood of substrate channeling, and active site access channels of sequential enzymes should face toward each other. Substrate channeling is usually not observed over longer distances unless other factors, such as electrostatic interactions, create a pathway for facilitated diffusion of intermediates [6,8]. Tailoring the spatial arrangement and orientation of diverse enzymes to meet these constraints requires a robust set of molecular tools.

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1Copenhagen Plant Science Centre, Department of Plant and Environmental Sciences, University of Copenhagen, Thorvaldsensvej 40, 1871 Frederiksberg C, Denmark @Twitter: @JBehrendorff (J.B.Y.H. Behrendorff) and @LabPribil (M. Pribil).

*Correspondence: [email protected]

https://doi.org/10.1016/j.tibtech.2019.10.009 ª 2019 Elsevier Ltd. All rights reserved.

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There has been substantial progress in developing protein–protein interaction tags to organize soluble enzymes in defined ratios both in vitro [9] and in the cytoplasm of engineered microorganisms [4,10]. However, many useful enzymes, such as eukaryotic cytochrome P450 enzymes (P450s) [11] and a variety of terpene synthases [12,13], require membrane association. Engineering protein scaffolding systems for membrane-associated enzymes is inherently more complex than for soluble enzymes. Some enzymes have specific lipid requirements that limit which membranes they can interact with, constraining scaffolding designs. Additionally, enzyme–lipid interactions can occur in orientations that occlude protein surfaces or termini, constraining the design of synthetic protein–protein interactions, and overexpression of hydrophobic proteinaceous membrane anchors may cause misfolding or aggregation, particularly if the rate of protein synthesis exceeds the capacity of the cell for membrane insertion [14,15]. Properties of major biological membranes that could be targets for engineering are summarized in Table 1. Engineered biosynthesis of many valuable secondary metabolites could be improved if more protein scaffolding techniques were compatible with membrane-associated enzymes. Membranebound scaffolding systems would also allow for greater control over the spatial organization of metabolism within the cell by beneficially restricting synthetic pathways to defined organelles: the plasma membrane or mitochondrial outer membrane might provide better access to membrane-bound redox carriers, or useful metabolic pathways could be positioned at membrane domains where specific cellular functions are enriched, such as secondary metabolite transport across the vacuolar membrane. Recent studies have begun to tackle the challenge of membrane-associated protein scaffolding. In this review, we describe notable membrane-bound scaffolding systems from nature and discuss the diverse range of solutions emerging in synthetic membrane protein scaffolding.

Inspiration from Natural Systems Many recent examples of synthetic scaffolding systems borrow design elements from natural scaffolding systems. A deeper understanding of the factors that drive native membrane-bound metabolon formation will likely provide new tools and design templates for creating synthetic membrane-bound scaffolds. Three important membrane-bound protein scaffolding systems from nature are outlined here.

Metabolons with Membrane-Bound and Soluble Enzymes Membrane-bound metabolons can be governed either by direct interactions between enzymes or by interactions between enzymes and dedicated scaffolding proteins. Several examples of membranebound metabolons have been identified in plants, where active colocalization appears to be essential for attaining high yields of specific secondary metabolites among other metabolic pathways that compete for the same versatile precursors [16–19]. Efficient production of lignin, the heterogeneous polymer that provides rigidity to plant cell walls, depends on formation of an enzyme complex bound by membrane-integral scaffold proteins [20]. To produce the monomeric subunits of lignin (monolignols), cinnamic acid is hydroxylated by up to three endoplasmic reticulum (ER)-bound P450s, and the products of these reactions are further modified by a set of soluble enzymes that associate with the P450s to form the monolignol biosynthetic complex [21,22]. Two ER-bound structural proteins, membrane steroid-binding proteins 1 and 2 (MSBP1 and MSBP2), act as scaffolding hubs to colocalize the three P450s in the ER membrane [20] (Figure 1A). Isoflavonoids are a broad class of phytochemicals that are also derived from cinnamic acid, and isoflavonoid biosynthetic enzymes form similar metabolons, in which several soluble enzymes cluster with other ER-bound P450s [18,23]. Loss of MSBP1 and MSBP2 disrupts the monolignol biosynthetic metabolon and causes a decrease in lignin production, structurally weakening the plant. This is accompanied by an increase in isoflavonoid production [20]. MSBP-mediated scaffolding is clearly important for directing metabolic flux toward monolignol production and maintaining the balance between lignin and isoflavonoid production.

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Glossary Fusion protein: a protein comprising two or more domains from different proteins, genetically encoded by a single open reading frame. Interaction tags: protein domains or short peptides that can be fused to a protein of interest, and which interact with other specific interaction tags, facilitating complex formation between proteins that are fused to complementary pairs of interaction tags. Lipid droplet: eukaryotic subcellular compartment with a hydrophobic core of neutral lipids enclosed by a phospholipid monolayer. Lipid droplets bud from the ER during lipogenesis. Macromolecular complex: a highmolecular weight structure comprising multiple proteins that interact post translationally. Metabolic flux: relative rate at which a metabolite is processed through several catalytic steps. Metabolic intermediates: molecules produced by enzymes in a metabolic pathway that are precursors to the final product of the pathway. Metabolon: protein complex containing sequential enzymes of a metabolic pathway held together via noncovalent interactions. Oligomeric complex: complex formed by a small set of monomers (e.g., between three and ten monomers) from a larger pool. Protein oligomers could comprise different or identical monomers. Protein scaffolding: process where different proteins form a complex via specific protein–protein interactions, sometimes via intermediary structural proteins. Scaffold protein: structural protein that serves as an assembly hub for other proteins that bind to it via noncovalent interactions. Substrate channeling: process where a metabolic intermediate is directly transferred from one enzyme to the next without diffusing in the local environment Thylakoid: continuous network of membranes inside the chloroplast that hosts the photosynthetic light reactions. Thylakoids have a bilayer membrane that separates the internal thylakoid lumen from the bulk aqueous phase of the chloroplast (the stroma).

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Table 1. Properties of Potential Target Membranes for Protein Scaffolding

Membrane system

Composition

Metabolic role

Membrane protein targeting

Refs

Eukaryotic

Phospholipid asymmetric

Solute and protein translocation,

Membrane insertion occurs at the ER,

[57,58]

cytoplasmic

bilayer. The inner leaflet

endocytosis, and exocytosis

and protein-containing membranes

membrane

contains phosphatidylserine,

are exported to the cytosolic membrane

phosphatidylinositol, and

via the trans-Golgi network

phosphatidylethanolamine; the outer leaflet contains phosphatidylcholine and sphingolipids ER

Large interconnected network

Protein synthesis, glycosylation,

Insertion via the Sec61/SecY translocon.

delimited by a continuous

and transport, lipid and steroid

Tail-anchored proteins are inserted via

phospholipid bilayer membrane

biogenesis, carbohydrate

the GET pathway

[59–61]

metabolism, and calcium metabolism Lipid droplets

Spherical phospholipid monolayer

Lipid and energy homeostasis, and

Direct recruitment via hydrophobic

encompassing a hydrophobic core

fatty acid and sterol metabolism

amphipathic helices, and asymmetric

of mixed neutral lipids

[62–64]

partitioning from the ER during lipid droplet biogenesis

Chloroplast

The thylakoid network and the outer

Protein and metabolite transport

Nuclear-encoded proteins are imported

and inner envelopes are lipid bilayers

at the outer and inner envelopes.

across membranes by TOC-TIC complexes.

mainly comprising galactolipids. The

Light-dependent reactions of

Depending on the target membrane,

outer envelope membrane also

photosynthesis and energy

proteins are integrated by TOC, TIC,

contains a substantial proportion

metabolism in the thylakoids

SEC2, SRP, or TAT mechanisms, or can

of phosphatidylcholine Mitochondria

[65–71]

be inserted spontaneously

Outer and inner phospholipid

Redox and energy metabolism, and

Proteins are imported by the TOM-TIM

bilayer membranes. Cardiolipin is

metabolism of amino acids, lipids,

complex and integrated either by MIM,

only present in the inner membrane

and nucleotides

SAM, TIM23, TIM22, or Oxa1 complexes

Gram-negative

Phospholipid bilayer inner

Solute and protein translocation.

Outer membrane proteins are exported

bacterial

membrane. The outer membrane

Energy and redox metabolism and

by the Sec system and the TAT translocon,

membranes

is an asymmetric lipid bilayer:

lipid synthesis also occur at the

and inserted by the BAM complex. Inner

the periplasmic side comprises

inner membrane

membrane proteins are inserted via the

glycerophospholipids and the outer

[72,73]

[74–78]

Sec and SRP systems

leaflet comprises lipopolysaccharides Gram-positive

Phospholipid bilayer membrane

Metabolite transport, energy and

Membrane proteins are inserted via the

bacterial

that contains lipoteichoic acid

redox metabolism, lipid biosynthesis

Sec and SRP systems. Proteins displayed

membrane

[79–81]

on the outer surface of the cell can be translocated by the Sec and TAT translocon pathways

Dhurrin is a cyanogenic glucoside that can comprise up to 30% of the dry mass of etiolated sorghum seedlings [16], and the high pathway flux required to reach this concentration is enabled by metabolon formation. Dhurrin biosynthesis requires five sequential oxidations of tyrosine (catalyzed by two ER-bound P450s, CYP79A1 and CYP71E1) followed by O-glycosylation at the cyanohydrin group of the p-hydroxymandelonitrile intermediate by a specific soluble UDP-glucosyl transferase (UGT85B1) (Figure 1B). Complex formation has been demonstrated between P450s CYP79A1 and CYP71E1, and between the P450s and UGT85B1 [16], but the molecular mechanisms driving these interactions remain uncharacterized. MSBPs were identified in dhurrin metabolon-enriched ER membrane fractions [16] and it is possible that MSBPs have a similar role to that observed in the monolignol metabolon, but this is yet to be

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(A)

Associated soluble enzymes

CYP98A3

CYP73A5

CYP84A1

MSBP

MSBP

(B) UGT85B1

(C)

CPR

CYP71E1

CYP79A1

Glucose

Oligosaccharides

GH5 Cel48A

Scaffoldin

Anchoring protein

Cel8A Glucose

GH9

Oligosaccharides Trends in Biotechnology

Figure 1. Naturally Occurring Membrane-Bound Scaffolding Systems. (A) Metabolic flux toward production of lignin monomers is directed by scaffolding of three endoplasmic reticulum (ER)-bound cytochrome P450 enzymes (P450s), CYP73A5, CYP98A3, and CYP84A1. The enzymes are colocalized via interactions with ER membrane-integral scaffolding proteins (membrane steroid-binding proteins, MSBP1 and MSBP2, collectively represented here as MBSP). (B) The dhurrin biosynthetic metabolon comprises two ERbound P450s (CYP79A1 and CYP71E1) and a soluble glucosyl transferase (UGT85B1). Specific production of dhurrin depends on scaffolded colocalization of the enzymes, but the scaffolding mechanism has not yet been elucidated. The cytochrome P450 reductase (CPR) is required for P450 activity, but evidence suggests a transient interaction between CPR and the dhurrin metabolon. (C) The cellulosome of Clostridium thermocellum (Figure legend continued at the bottom of the next page.)

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demonstrated directly. The P450s also interact with the membrane-bound cytochrome P450 reductase (CPR), which supplies electrons to support the P450 catalytic cycle, but evidence suggests that this is a transient interaction [24]. Characterization of membrane scaffolding mechanisms from nature will likely help in developing synthetic membrane-bound scaffolding.

Membrane-Anchored Soluble Enzyme Complexes The cellulosome is a complex of soluble cellulolytic enzymes bound to the outer membrane of some prokaryotes, such as Clostridium thermocellum. Cellulosome enzymes work synergistically to degrade extracellular cellulose and hemicellulose, releasing utilizable monosaccharides [25]. The extracellular enzymes bind to a secreted structural protein, the scaffoldin, which in turn binds an anchoring protein on the outer membrane of the bacterial cell. Tethering the enzymes to the surface of the cell ensures that the bacterium is near the sugars liberated from cellulose degradation. The secreted enzymes attach to the scaffoldin protein via interaction domains (type I dockerins) that bind to complementary domains on the scaffoldin protein (type I cohesins) [26] (Figure 1C). The scaffoldin has many type I cohesin domains for binding cellulolytic enzymes, and one type II dockerin domain that specifically interacts with a type II cohesin on the bacterial outer membrane. The specific pairwise interactions mediated by cohesins and dockerins have been repurposed in several synthetic scaffolding studies, as discussed later.

Approaches to Synthetic Membrane-Bound Scaffolding Generic Tools Adapted from Soluble Scaffolding Systems Protein scaffolding can be used to create synthetic complexes of membrane-requiring enzymes or interactions between soluble and membrane-bound enzymes. The primary challenge of membrane-bound protein scaffolding is to have at least one element of the complex bound to the target membrane without loss of function. Beyond this, many approaches are available for colocalizing other proteins with the anchoring module. While it is possible to combine enzymes as genetically encoded fusion proteins, a more flexible approach is to fuse short interaction domains or ‘tags’ to the proteins of interest to drive post-translational colocalization. Tag-mediated protein scaffolding has several advantages over the fusion protein approach: not all enzymes can be fused in a single open reading frame without negatively affecting protein folding; more proteins can be assembled via tag-mediated scaffolding than can efficiently be cloned as a fusion protein; and tag-mediated post-translational scaffolding allows for the design of larger complex assemblies that positively affect the overall metabolic flux through a synthetic metabolon. Noncovalent post-translational scaffolding could also allow for the dynamic assembly and disassembly of complexes in response to environmental or intracellular signals. Several sets of small peptide tags have been developed for creating scaffolds via specific protein– protein interactions. These tags are fused to proteins at the N or C terminus, or can even be introduced into loop regions. Proteins that bear complementary tags form complexes via noncovalent interactions. The binding affinities of protein scaffolding tags are an important design consideration: complexes built using high-affinity tags (usually characterized in terms of low nanomolar dissociation constants) will exist primarily in the scaffolded state. Scaffolds that use low-affinity tags may not exhibit enhanced substrate channeling due to increased time spent in the dissociated state. Some popular interaction tags are adapted from naturally occurring protein–protein interactions, such as Src homology 3 (SH3) ligand and binding domain pairs [4], or

Figure 1. Continued is an extracellular enzymatic complex where a mixture of cellulolytic enzymes, such as exo- and endocellulases (e.g., Cel48A and Cel8A) and glucosyl hydrolases (e.g., GH9 and GH5), are bound to an extracellular scaffolding protein, the scaffoldin. The scaffoldin has several type I cohesin domains, to which the cellulolytic enzymes bind via noncatalytic type I dockerin domains. The scaffoldin binds a cell surface-anchoring protein via a type II cohesin–dockerin interaction, thus maintaining proximity between the cell surface and the cellulolytic activity of the cellulosome. A simplified cartoon of the cellulosome is shown here with only four enzymes bound; in reality, the scaffoldin has many more binding sites and the mass of the complex exceeds 1 MDa. Cohesin domains are shown in gray, while dockerin domains are the same color as the proteins to which they are fused.

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cohesin and dockerin domains [27]. Sets of synthetic interaction tags based on interacting coiled-coils (zippers) [28–30] and other engineered interaction domains, such as affibodies [31], provide a range of orthogonal tags with specific interaction partners and a range of binding affinities, with dissociation constants spanning the micromolar to nanomolar range. Examples of tags used to colocalize soluble and membrane-integral components are described in several of the examples detailed later, and their properties are summarized in Table 2.

Membrane-Anchored Scaffolding for Soluble Proteins Soluble protein complexes can be assembled at membrane surfaces by tag-mediated scaffolding with a membrane-anchored structural protein. The choice of membrane anchor used then dictates which membrane the soluble enzymes are recruited to. This approach was used to enhance the production of ethyl acetate in Saccharomyces cerevisiae. Ethyl acetate biosynthesis is normally limited by spatial segregation between alcohol-O-acetyltransferase (Atf1), which is bound to the surface of lipid droplets, and the cytosolic upstream enzymes aldehyde dehydrogenase (Ald6) and acetyl-CoA synthase (Acs1) [32]. To colocalize the ethyl acetate biosynthetic enzymes, Lin and colleagues scaffolded Ald6 and Acs1 in a synthetic complex that was bound to the surface of lipid droplets via a heterologous oleosin domain [32]. Oleosins are small (16–24 kDa) structural proteins embedded in the surface of oil bodies in higher plants [33,34]. They comprise N- and C-terminal amphipathic helices linked by a hydrophobic hairpin that anchors the protein in lipid membranes. The hydrophobic hairpin is buried in the lipid phase and the N- and C- terminal amphipathic helices are exposed to the cytosol. When overexpressed in S. cerevisiae, oleosins bind to lipid droplets and increase the rate of lipid droplet production from the ER [35]. A lipid droplet-binding scaffold protein was created by fusing two different cohesin domains from Clostridium perfringens and C. thermocellum to the C terminus of the Zea mays oleosin. Complementary dockerin domains were fused to Ald6 and Acs1 to enable their recruitment to the oleosin-cohesin scaffold (Figure 2A). When these proteins were co-expressed in S. cerevisiae, an oleosin-Ald6-Acs1 complex colocalized with Atf1 at the surface of lipid droplets, doubling ethyl acetate production [32]. The synthetic oleosin-cohesin protein could be used to scaffold any dockerin-fused soluble protein to lipid droplets, and the specificity of cohesin–dockerin interactions provides a mechanism for controlling the stoichiometry of the recruited enzymes. Oleosins themselves can also form homo-oligomeric complexes in lipid droplets, and adjusting the amino acid composition of the oleosin alters lipid droplet morphology [36]. This suggests that there is an opportunity to create supercomplexes of engineered oleosin scaffolds, and perhaps even to further tune the spatial arrangement of scaffolded enzymes via customized lipid droplet morphologies.

Opportunities to Combine Soluble and Membrane-Bound Enzymes in Synthetic Complexes Interplay between soluble enzymes and membrane-bound P450s is essential for the biosynthesis of valuable terpenoid compounds, such as monoterpene indole alkaloids [37], and many flavors and fragrances [38]. Monoterpene indole alkaloids are a class of natural products that include the anticancer drugs vinblastine, vincristine, and vinflunine, all of which are derived from a common precursor, strictosidine. Strictosidine biosynthesis in S. cerevisiae requires at least 11 proteins, including soluble enzymes and four nonsequential membrane-bound P450s, which are considered to be rate-limiting [37]. While ambitious compared with the current state-of-the-art, it may soon be possible to test whether the efficiency of the strictosidine pathway can be improved by scaffolding the entire biosynthetic pathway in an ER membrane-bound mega-complex. To create larger scaffolded complexes with more enzymes, a single membrane-anchored tag can be used to recruit an intermediate scaffold protein with many tag-binding interfaces, akin to the cellulosome scaffoldin protein (Figure 1C). The size of the intermediate scaffold protein is ultimately limited by the same constraints as engineered fusion proteins, but these constraints can be overcome by using a scaffold protein that itself assembles into a macromolecular complex. Recently, a self-assembling macromolecular scaffold was developed based on PduA, a structural subunit of the bacterial propanediol utilization microcompartment. A modified version of PduA (PduA*) assembles into

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Table 2. Properties of Example Scaffolding Modules

Scaffolding module

Description

Size (amino acid residues)

GBD ligand/binding

GTPase-binding domain from N-WASP, an actin

Binding domain: 79

domain pair

polymerization switch

Ligand: 32

PDZ ligand/binding

PSD95/DlgA/Zo-1 domain from syntrophin, an

Binding domain: 95

domain pair

adaptor protein

Ligand: 7

SH3 ligand/binding

Src homology 3 domain from CRK, an adaptor

Binding domain: 57

domain pair

protein

Ligand: 11

WH1 ligand/binding

WASP homology 1 domain from rat neural

Binding domain: 114

domain pair

Wiskott–Aldrich syndrome protein (N-WASP)

Ligand: 12

Cohesin-dockerin pairs

Binding domains from the extracellular

Cohesin: 140

cellulosome of cellulolytic bacteria such as

Dockerin: typically 60–70

Affinity (Kd) 1310

–6

Refs

M

[4]

8310–6 M

[4]

1310–7 M

[4]

To be determined

[45,46]

1310–8 to 1310–11 M

[27,32,82,83]

116

To be determined

[10]

21, 24, or 28

3310–6 to <1310–11 M

[84]

30–50

>4310–4 to <1310–8 M

[28]

RIDD domain (derived from a cAMP-dependent

RIDD: 44

1.2310–9 M

[30]

protein kinase) forms homodimers that can

RIAD: 18

58

9310–7 to 5310–8 M

[31]

Membrane-integral TatB and TatC subunits of the

TatB: 260

To be determined

[48]

twin-arginine translocase (Tat) complex. TatB and

TatC: 340

To be determined

[52]

Clostridium thermocellum PduA*

PduA bacterial microcompartment protein with an additional 23 residues at the C terminus forms filaments when overexpressed in Escherichia coli

CC-Di series of

Pairs of synthetic coiled-coil domains, with

coiled-coil domains

different binding affinities available depending on which domains are paired

SYNZIP domains

Pairs of synthetic coiled-coil domains, with different binding affinities available depending on which domains are paired

RIDD-RIAD tags

then bind RIAD (an amphipathic helix from an A-kinase anchoring protein). This results in a 2:1 binding stoichiometry (two RIDD-fused proteins: one RIAD-fused protein) Affibodies

Libraries of binding domains based on Fc-binding domain of Staphylococcus aureus protein A

TatBC complex

TatC form TatBC heterodimers, and a complete Tat complex contains six or seven TatBC heterodimers CURT1A

A membrane-integral hydrophobic hairpin with

103

amphipathic helices exposed to the aqueous phase. CURT1A forms homo-oligomeric complexes in a variety of membranes. Dimeric, trimeric, and tetrameric CURT1A complexes have been demonstrated, and higher-order complexes may also be possible.

filaments when overexpressed in Escherichia coli [10,39], and these PduA* filaments can be localized to the inner membrane of E. coli by adding a compatible scaffolding tag to a membrane-anchoring domain (Figure 2B). In this example, the membrane-anchoring domain was the hydrophobic C terminus of a cell division-associated protein, minD, from Bacillus subtilis. A membrane-anchored

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(A)

Ald6

Acs1 Atf1

Oleosin

(B)

PduA* subunits

Basic coiled-coil Acidic coiled-coil minD anchor

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Figure 2. Membrane-Bound Synthetic Soluble Complexes. (A) A scaffold protein for targeting enzymes to lipid droplets in Saccharomyces cerevisiae was created by fusing two different cohesin domains to an oleosin protein. Oleosins integrate into lipid droplet membranes when expressed in S. cerevisiae. This scaffold was used to enhance ethyl acetate biosynthesis by targeting dockerin-fused aldehyde dehydrogenase (Ald6) and acetyl-CoA synthase (Acs1) to lipid droplets, colocalizing the first two enzymes in ethyl acetate biosynthesis with Atf1, which natively localizes to lipid droplets. Cohesin domains (fused to the C terminus of the oleosin) are shown in gray, while dockerin domains are the same color as the proteins to which they are fused. (B) A soluble multimeric ultrastructure can be targeted to the inner membrane of Escherichia coli. PduA* is a modified bacterial microcompartment shell protein that self-assembles into macromolecular filaments when expressed in E. coli. Synthetic coiled-coil domains were used to bind the filament to the E. coli inner membrane. The membrane-integral C terminus of the minD protein was fused to an acidic coiled-coil domain, which bound to a basic coiled-coil fused to the PduA* monomer.

PduA* filament could be decorated with enzymes by co-expressing PduA* monomers bearing different interaction tags.

Scaffolding Soluble Proteins to Transmembrane Transporters Transport proteins that regulate the movement of specific molecules from one side of a membrane to the other are a defining feature of biological membranes. Naturally occurring transport metabolons scaffold intracellular metabolic functions onto transport proteins so that substrates can be metabolized immediately upon import to the intracellular environment [40]. This phenomenon may improve the efficiency of transport by preventing back-flux of the imported substrate. Synthetic scaffolding between cytosolic metabolic pathways and transmembrane transporters creates an opportunity to increase yields by capturing imported molecules in preferred metabolic pathways before they

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interact with native metabolism. Similarly, scaffolding toxic metabolite pathways to a relevant export channel could relieve toxicity. GABA, a precursor to nylon, can be produced by enzymatic decarboxylation of glutamate. However, microbial hosts, such as E. coli, can also catabolize glutamate as a carbon source [41] or use it for protein synthesis. To minimize the loss of glutamate to central metabolism and improve conversion of glutamate to GABA, a glutamate decarboxylase was scaffolded to gadC, an E. coli transmembrane channel that imports glutamate and exports GABA [42] (Figure 3A). In an optimized configuration, SH3-mediated scaffolding between the glutamate decarboxylase and the glutamate/GABA transporter resulted in conversion of glutamate to GABA with a yield >97% and increased the production rate 3.5-fold compared with unscaffolded expression of glutamate decarboxylase. This near-complete conversion of fed glutamate to GABA is a compelling demonstration of the advantage gained by spatially reconfiguring cellular biochemistry to divert substrates away from central metabolism, and hints at opportunities to design more complex high-yield whole-cell biocatalysts. Microorganisms can be used to upgrade organic acids to alcohols for use in biofuels [43,44], but typical pathways for these reactions proceed via CoA-thioester intermediates that can be metabolized by a range of competing reactions, including assimilation into central carbon metabolism. Import channel-anchored scaffolding could improve the conversion of acids to alcohols at the site of import, limiting the loss of CoA-thioester intermediates to competing reactions. A similar approach has been used to improve utilization of an alternative feedstock (xylose) in S. cerevisiae [45]. The first step toward incorporating xylose into the pentose phosphate pathway is isomerization to xylulose by xylose isomerase, but S. cerevisiae also contains nonspecific aldose reductases that convert xylose to xylitol. Xylitol accumulation inhibits xylose isomerase, decreasing both the yield and the rate of xylose utilization. To minimize xylitol formation, Thomik and colleagues scaffolded xylose isomerase to a galactose permease channel that imports xylose [45]. The galactose permease was tagged with a synthetic coiled-coil zipper domain [28] (SYNZIP SZ2) that specifically binds a complementary SYNZIP domain, SZ1. However, xylose isomerase activity was lost when fused to the SZ1 tag, meaning that direct SYNZIP-mediated scaffolding was not possible. The authors found that xylose isomerase retained function when tagged with a WH1 [46] ligand domain; thus, they developed a synthetic scaffold protein to mediate interactions between the SZ2-tagged galactose permease and the WH1-tagged xylose isomerase (Figure 3B). This intermediary scaffold protein (comprising SZ1 and a WH1-binding domain) recruited xylose isomerase to the galactose permease, doubling the rate of xylose uptake while decreasing xylitol production and increasing the ethanol titer 2.5-fold. Fusing new binding domains onto enzymes can have unpredictable effects on folding and function, meaning that identifying a compatible scaffolding tag for a given enzyme can seem like a stochastic process. This study highlights the value of testing diverse sets of interaction tags and shows that solutions can be engineered for connecting incompatible tags. Linking the transporter and isomerase via a structural scaffold protein also creates an opportunity for greater regulatory control over metabolism. For example, recruitment of xylose isomerase to the galactose permease could be dynamically regulated by independently inducing or repressing expression of the scaffold protein.

Membrane-Integral Scaffolds for Complexes with Multiple Membrane Proteins The examples discussed earlier depend on one membrane protein serving as an anchoring point for the recruitment of other enzymes via soluble protein–protein interactions. This approach is limited to complexes involving only one membrane protein. Membrane-integral scaffolding modules are necessary to create synthetic complexes with multiple membrane-requiring enzymes (e.g., multiple eukaryotic P450s). Early success in this area has focused on fusing enzymes to proteins that natively form multimeric membrane-integral complexes. The need for adaptable membrane scaffolding modules was highlighted when Gnanasekaran and colleagues attempted to transfer the dhurrin biosynthetic metabolon to the chloroplast [47].

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(A)

Glutamate/ GABA antiporter

Glutamate decarboxylase

(B)

Galactose/ xylose permease

SZ1 WH1-binding domain

Xylose isomerase Trends in Biotechnology

Figure 3. Scaffolding Enzymes to Transmembrane Transporters. (A) Glutamate decarboxylase was scaffolded to the GABA/glutamate antiporter to support high yield conversion of fed glutamate to GABA in Escherichia coli. A Src homology 3 (SH3)-binding domain was fused to the cytosolic N terminus of the GABA/glutamate antiporter, which recruited an SH3 ligand domain fused to a glutamate (Figure legend continued at the bottom of the next page.)

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Targeting expression of the dhurrin pathway enzymes to chloroplasts resulted in a loss of metabolon formation. CYP79A1 and CYP71E1 both anchored to the thylakoid but did not form a complex [48], and UGT85B1 was soluble in the stroma. This nonscaffolded configuration resulted in glycosylation of the intermediate metabolites by nonspecific chloroplast glucosyl transferases, substantially decreasing the yield of dhurrin. It is not clear why complex formation was lost when the enzymes were targeted to the tobacco chloroplast rather than the ER of sorghum. Interactions between P450s could be lost due to differences in membrane lipid composition or the lack of ER-resident scaffolding proteins such as MSBPs [16], and P450-UGT85B1 interactions might be sensitive to changes in the composition of the aqueous phase. A thylakoid membrane-bound dhurrin metabolon was created by fusing enzymes to subunits of the twin-arginine translocase (Tat) complex [48]. TatB and TatC are transmembrane proteins that interact to form a heterodimer (TatBC), and the complete Tat complex comprises seven or eight TatBC heterodimers arranged in a membrane-embedded ring [49,50]. CYP79A1 and CYP71E1 were expressed as independent TatB-P450 fusions where TatB replaced the N-terminal membrane anchor of the P450. UGT85B1 was fused to TatC via a flexible linker (Figure 4A). Glycosylation of intermediate metabolites was minimized when all three enzymes were colocalized in synthetic TatBC heterocomplexes, resulting in a fivefold increase in the yield of dhurrin. The Tat-based system can be used to create complexes with multiple soluble and membraneassociated enzymes, but its deployment is probably limited to membranes that naturally contain Tat complexes (thylakoids and bacterial inner membranes) and it is not known whether enzyme– TatB/C fusion proteins affect the function of the Tat complex. A recent study discovered that CURT1A, a structural protein that constrains the shape of thylakoid membranes [51], can be repurposed as an anchor that forms oligomeric complexes in membranes of diverse expression hosts [52]. CURT1A has a similar structure to oleosins, with two amphipathic helices linked by a hydrophobic hairpin, except CURT1A is natively found in the galactolipid-rich thylakoid bilayer, whereas oleosins are found in phospholipid monolayers. Reporter proteins fused to CURT1A formed dimeric, trimeric, and tetrameric complexes not only in thylakoid membranes, but also in the ER of tobacco and S. cerevisiae, and the inner membrane of E. coli when expressed heterologously (Figure 4B). CURT1A-based scaffolding had distinct properties in each expression system. For example, in S. cerevisiae, CURT1A-fused proteins had enhanced expression and clustered in distinct membrane subdomains in the cortical ER, creating an additional level of spatial control over the subcellular distribution of enzymes. In addition to scaffolding reporter proteins, CURT1A fusion was compatible with expression of a human P450 enzyme, CYP2C19. CURT1A was fused to CYP2C19 either as a replacement for the hydrophobic N-terminal membrane anchor of CYP2C19 or to the full-length protein, and, in both cases, the P450 could be expressed in E. coli without CURT1A interfering with the folding and maturation of the holoenzyme. Membrane-integral protein scaffolding is the least-developed aspect of membrane-bound scaffolding, but it demands attention for its potential to improve metabolic engineering with some of most interesting yet challenging enzymes in nature. In particular, scaffolding could improve substrate channeling in pathways that require multiple sequential membrane-bound P450s, and could be developed to regulate the stoichiometry between P450s and their corresponding CPR [53]. Biosynthesis of several chemicals of environmental, economic, and social importance depends on P450s, including production of sandalwood fragrance compounds [54], artemisinin (an antimalarial), and taxol (a chemotherapeutic agent against several cancers). Currently, the Tat and CURT1A fusion Figure 3. Continued decarboxylase. (B) A synthetic scaffold protein was used to connect xylose isomerase to a galactose/xylose permease channel. Attempts to directly connect xylose isomerase to the permease channel via synthetic zipper (SZ) domains caused a loss of xylose isomerase function; thus, an intermediary scaffolding protein comprising an SZ1-domain and a WH1-binding domain was created (labeled and shown in gray). This scaffold protein mediated colocalization of a WH1 ligand-fused xylose isomerase and an SZ2-fused permease channel.

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UGT85B1

UGT85B1

CYP71E1

CYP79A1

TatB

CYP79A1

CYP71E1

TatC

(B) mCitrine CURT1A

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Figure 4. Membrane-Integral Synthetic Scaffolding. (A) The catalytic enzymes of the dhurrin pathway were fused to twin-arginine translocase (Tat)B and TatC subunits of the Tat complex. Expression of the fusion proteins was targeted to the chloroplast, where they formed chimeric Tat complexes in the thylakoid membrane, consequently colocalizing the dhurrin pathway enzymes. A complete Tat complex forms a channel comprising six TatB subunits and six or seven TatC subunits. (B) A soluble fluorescent reporter protein (mCitrine) fused to CURT1A formed multimeric complexes in a variety of biological membranes.

protein-based approaches are limited by a lack of control over the stoichiometric composition of their complexes: heterocomplex composition is probably governed partly by the translation efficiency of each of the different fusion proteins. New techniques to control the stoichiometric composition of membrane-integral enzyme complexes would be a significant achievement.

Concluding Remarks and Future Perspectives The benefit of creating enzyme complexes to drive specific metabolic fluxes is increasingly apparent. Metabolons found in nature show that scaffolded complexes are particularly important for producing secondary metabolites when multiple pathways compete for common intermediates. Currently, synthetic membrane-bound protein scaffolding mostly depends on combining soluble protein scaffolding tools with pre-existing membrane proteins, including structural proteins and transmembrane transporters.

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Some membrane-integral scaffolding approaches have been demonstrated, but more tools are required to mediate membrane-integral complex formation between unrelated enzymes. A better understanding of the protein–protein interactions that underpin naturally occurring membrane metabolons will expand this set of scaffolding modules and provide added flexibility when designing synthetic membrane metabolons. Modern genome-editing techniques have simplified the work required to substantially modify the lipid composition of microbial membranes [55,56], and this raises the possibility that membrane lipid profiles or new membrane domains could be engineered in microbial hosts to accommodate the specific lipid requirements of some orthogonal membrane enzymes and scaffolding systems (see Outstanding Questions). Additional membrane engineering may be particularly important for implementing membrane-integral scaffolds in prokaryotes, the survival of which is dependent on not disrupting the integrity of their one available inner membrane. Ultimately, fully orthogonal membrane-bound scaffolding systems should permit the design of synthetic metabolons that emulate the level of complexity observed in nature.

Acknowledgments This work was supported, in part, by funding from the Novo Nordisk Foundation (project grant number NNF15OC0016586) and the Independent Research Fund Denmark (project grant number DFF-7017-00122). We thank Francesca Toselli and Mareike Bongers for their constructive feedback on this manuscript.

References 1. Miles, E.W. et al. (1999) The molecular basis of substrate channeling. J. Biol. Chem. 274, 12193– 12196 2. Dunn, M.F. et al. (2008) Tryptophan synthase: the workings of a channeling nanomachine. Trends Biochem. Sci. 33, 254–264 3. Anderson, K.S. et al. (1995) Kinetic characterization of channel impaired mutants of tryptophan synthase. J. Biol. Chem. 270, 29936–29944 4. Dueber, J.E. et al. (2009) Synthetic protein scaffolds provide modular control over metabolic flux. Nat. Biotechnol. 27, 753–759 5. Castellana, M. et al. (2014) Enzyme clustering accelerates processing of intermediates through metabolic channeling. Nat. Biotechnol. 32, 1011– 1018 6. Lin, J.-L. et al. (2014) Design and analysis of enhanced catalysis in scaffolded multienzyme cascade reactions. ACS Catal. 4, 505–511 7. Sweetlove, L.J. and Fernie, A.R. (2018) The role of dynamic enzyme assemblies and substrate channelling in metabolic regulation. Nat. Commun. 9, 2136 8. Bauler, P. et al. (2010) Channeling by proximity: the catalytic advantages of active site colocalization using Brownian dynamics. J. Phys. Chem. Lett. 1, 1332–1335 9. Schoffelen, S. and van Hest, J.C.M. (2012) Multienzyme systems: bringing enzymes together in vitro. Soft Matter 8, 1736–1746 10. Lee, M.J. et al. (2017) Engineered synthetic scaffolds for organizing proteins within the bacterial cytoplasm. Nat. Chem. Biol. 14, 142–147 11. Barnaba, C. and Ramamoorthy, A. (2018) Picturing the membrane-assisted choreography of cytochrome P450 with lipid nanodiscs. ChemPhysChem 19, 2603–2613 12. Yang, Y. et al. (2018) Identification and characterization of a membrane-bound sesterterpene cyclase from Streptomyces somaliensis. J. Nat. Prod. 81, 1089–1092 13. Tang, M.-C. et al. (2017) Late-stage terpene cyclization by an integral membrane cyclase in the biosynthesis of isoprenoid epoxycyclohexenone natural products. Org. Lett. 19, 5376–5379

14. Zhang, Z. et al. (2015) High-level production of membrane proteins in E. coli BL21(DE3) by omitting the inducer IPTG. Microb. Cell Fact. 14, 142 15. Gialama, D. et al. (2017) Functional requirements for DjlA- and RraA-mediated enhancement of recombinant membrane protein production in the engineered Escherichia coli strains SuptoxD and SuptoxR. J. Mol. Biol. 429, 1800–1816 16. Laursen, T. et al. (2016) Characterization of a dynamic metabolon producing the defense compound dhurrin in sorghum. Science 354, 890–893 17. Gou, M. et al. (2018) The scaffold proteins of lignin biosynthetic cytochrome P450 enzymes. Nat. Plants 4, 299–310 18. Dastmalchi, M. et al. (2016) Twin anchors of the soybean isoflavonoid metabolon: evidence for tethering of the complex to the endoplasmic reticulum by IFS and C4H. Plant J 85, 689–706 19. Pollmann, S. et al. (2019) Substrate channeling in oxylipin biosynthesis through a protein complex in the plastid envelope of Arabidopsis thaliana. J. Exp. Bot. 70, 1483–1495 20. Gou, M. et al. (2018) The scaffold proteins of lignin biosynthetic cytochrome P450 enzymes. Nat. Plants 4, 299–310 21. Chen, H.-C. et al. (2011) Membrane protein complexes catalyze both 4- and 3-hydroxylation of cinnamic acid derivatives in monolignol biosynthesis. Proc. Natl. Acad. Sci. U. S. A. 108, 21253–21258 22. Bassard, J.-E. et al. (2012) Protein-protein and protein-membrane associations in the lignin pathway. Plant Cell 24, 4465–4482 23. Mameda, R. et al. (2018) Involvement of chalcone reductase in the soybean isoflavone metabolon: identification of GmCHR5, which interacts with 2-hydroxyisoflavanone synthase. Plant J. 96, 56–74 24. Bassard, J.-E. et al. (2017) Assembly of dynamic P450mediated metabolons-order versus chaos. Curr. Mol. Biol. Rep. 3, 37–51 25. Smith, S.P. et al. (2017) Continually emerging mechanistic complexity of the multi-enzyme cellulosome complex. Curr. Opin. Struct. Biol. 44, 151–160 26. Barth, A. et al. (2018) Dynamic interactions of type I cohesin modules fine-tune the structure of the

Outstanding Questions What are the protein components that drive complex formation in native membrane metabolons, and can they be repurposed for creating synthetic membranebound heterocomplexes? What are the rules underpinning interactions between membrane protein scaffolds and different biological lipid bilayers and monolayers? How might we design scaffolds to control the stoichiometric composition of membrane-integral enzyme complexes? How can we design dynamic assembly and disassembly of membrane-bound protein complexes in response to environmental or intracellular signals? Can membrane-integral scaffolding modules be used to create new membrane subdomains for concentrating specific metabolic functions? How can we combine membrane protein scaffolding with lipid engineering to produce new synthetic membranous vesicles or organelles?

Trends in Biotechnology, --, Vol. --, No. --

13

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27.

28.

29. 30. 31. 32. 33.

34. 35.

36. 37. 38. 39.

40. 41. 42.

43.

44.

45. 46. 47.

14

cellulosome of Clostridium thermocellum. Proc. Natl. Acad. Sci. U. S. A. 115, E11274–E11283 Carvalho, A.L. et al. (2003) Cellulosome assembly revealed by the crystal structure of the cohesindockerin complex. Proc. Natl. Acad. Sci. U. S. A. 100, 13809–13814 Thompson, K.E. et al. (2012) SYNZIP protein interaction toolbox: in vitro and in vivo specifications of heterospecific coiled-coil interaction domains. ACS Synth. Biol. 1, 118–129 Anderson, G.P. et al. (2018) Orthogonal synthetic zippers as protein scaffolds. ACS Omega 3, 4810– 4815 Kang, W. et al. (2019) Modular enzyme assembly for enhanced cascade biocatalysis and metabolic flux. Nat. Commun. 10, 4248 Tippmann, S. et al. (2017) Affibody scaffolds improve sesquiterpene production in Saccharomyces cerevisiae. ACS Synth. Biol. 6, 19–28 Lin, J.-L. et al. (2017) Synthetic protein scaffolds for biosynthetic pathway colocalization on lipid droplet membranes. ACS Synth. Biol. 6, 1534–1544 Napier, J.A. et al. (1996) The structure and biogenesis of plant oil bodies: the role of the ER membrane and the oleosin class of proteins. Plant Mol. Biol. 31, 945–956 Abell, B.M. et al. (2004) Membrane topology and sequence requirements for oil body targeting of oleosin. Plant J 37, 461–470 Jacquier, N. et al. (2013) Expression of oleosin and perilipins in yeast promotes formation of lipid droplets from the endoplasmic reticulum. J. Cell Sci. 126, 5198–5209 Vargo, K.B. et al. (2012) Self-assembly of tunable protein suprastructures from recombinant oleosin. Proc. Natl. Acad. Sci. U. S. A. 109, 11657–11662 Brown, S. et al. (2015) De novo production of the plant-derived alkaloid strictosidine in yeast. Proc. Natl. Acad. Sci. U. S. A. 112, 3205–3210 Schempp, F.M. et al. (2018) Microbial cell factories for the production of terpenoid flavor and fragrance compounds. J. Agric. Food Chem. 66, 2247–2258 Pang, A. et al. (2014) Structural insights into higher order assembly and function of the bacterial microcompartment protein PduA*. J. Biol. Chem. 289, 22377–22384 Moraes, T.F. and Reithmeier, R.A.F. (2012) Membrane transport metabolons. Biochim. Biophys. Acta 1818, 2687–2706 Marcus, M. and Halpern, Y.S. (1969) The metabolic pathway of glutamate in Escherichia coli K-12. Biochim. Biophys. Acta 177, 314–320 Somasundaram, S. et al. (2017) Introduction of synthetic protein complex between Pyrococcus horikoshii glutamate decarboxylase and Escherichia coli GABA transporter for the improved production of GABA. Biochem. Eng. J. 120, 1–6 Doud, D.F.R. et al. (2017) Metabolic engineering of Rhodopseudomonas palustris for the obligate reduction of n-butyrate to n-butanol. Biotechnol. Biofuels 10, 178 Scully, S.M. et al. (2019) Biotransformation of organic acids to their corresponding alcohols by Thermoanaerobacter pseudoethanolicus. Anaerobe 57, 28–31 Thomik, T. et al. (2017) An artificial transport metabolon facilitates improved substrate utilization in yeast. Nat. Chem. Biol. 13, 1158–1163 Zettl, M. and Way, M. (2002) The WH1 and EVH1 domains of WASP and Ena/VASP family members bind distinct sequence motifs. Curr. Biol. 12, 1617–1622 Gnanasekaran, T. et al. (2016) Transfer of the cytochrome P450-dependent dhurrin pathway from Sorghum bicolor into Nicotiana tabacum

Trends in Biotechnology, --, Vol. --, No. --

48.

49. 50. 51.

52.

53. 54.

55.

56.

57.

58.

59.

60. 61.

62. 63. 64. 65. 66. 67.

chloroplasts for light-driven synthesis. J. Exp. Bot. 67, 2495–2506 Henriques de Jesus, M.P.R. et al. (2017) Tat proteins as novel thylakoid membrane anchors organize a biosynthetic pathway in chloroplasts and increase product yield 5-fold. Metab. Eng. 44, 108–116 Patel, R. et al. (2014) Protein transport by the bacterial Tat pathway. Biochim. Biophys. Acta 1843, 1620– 1628 Celedon, J.M. and Cline, K. (2012) Stoichiometry for binding and transport by the twin arginine translocation system. J. Cell. Biol. 197, 523–534 Armbruster, U. et al. (2013) Arabidopsis CURVATURE THYLAKOID1 proteins modify thylakoid architecture by inducing membrane curvature. Plant Cell 25, 2661–2678 Behrendorff, J.B.Y.H. et al. (2019) Membranebound protein scaffolding in diverse hosts using thylakoid protein CURT1A. ACS Synth. Biol. 8, 611–620 Urlacher, V.B. and Girhard, M. (2019) Cytochrome P450 monooxygenases in biotechnology and synthetic biology. Trends Biotechnol. 37, 882–897 Celedon, J.M. et al. (2016) Heartwood-specific transcriptome and metabolite signatures of tropical sandalwood (Santalum album) reveal the final step of (Z)-santalol fragrance biosynthesis. Plant J. 86, 289–299 Wu, T. et al. (2017) Membrane engineering - a novel strategy to enhance the production and accumulation of b-carotene in Escherichia coli. Metab. Eng. 43, 85–91 Teixeira, P.G. et al. (2018) Engineering lipid droplet assembly mechanisms for improved triacylglycerol accumulation in Saccharomyces cerevisiae. FEMS Yeast Res. 18, foy060 Casares, D. et al. (2019) Membrane lipid composition: effect on membrane and organelle structure, function and compartmentalization and therapeutic avenues. Int. J. Mol. Sci. 20, E2167 Chevalier, A.S. and Chaumont, F. (2015) Trafficking of plant plasma membrane aquaporins: multiple regulation levels and complex sorting signals. Plant Cell Physiol. 56, 819–829 Schwarz, D.S. and Blower, M.D. (2016) The endoplasmic reticulum: structure, function and response to cellular signaling. Cell. Mol. Life Sci. 73, 79–94 Rapoport, T.A. et al. (2017) Structural and mechanistic insights into protein translocation. Annu. Rev. Cell Dev. Biol. 33, 369–390 Mateja, A. and Keenan, R.J. (2018) A structural perspective on tail-anchored protein biogenesis by the GET pathway. Curr. Opin. Struct. Biol. 51, 195–202 Pol, A. et al. (2014) Biogenesis of the multifunctional lipid droplet: lipids, proteins, and sites. J. Cell Biol. 204, 635–646 Olzmann, J.A. and Carvalho, P. (2019) Dynamics and functions of lipid droplets. Nat. Rev. Mol. Cell Biol. 20, 137–155 Pre´vost, C. et al. (2018) Mechanism and determinants of amphipathic helix-containing protein targeting to lipid droplets. Dev. Cell 44, 73–86 Ho¨lzl, G. and Do¨rmann, P. (2019) Chloroplast lipids and their biosynthesis. Annu. Rev. Plant Biol. 70, 51–81 Kim, J. et al. (2019) Biogenesis of chloroplast outer envelope membrane proteins. Plant Cell Rep. 38, 783–792 Mechela, A. et al. (2019) A brief history of thylakoid biogenesis. Open Biol. 9, 180237

Please cite this article in press as: Behrendorff et al., Synthetic Protein Scaffolding at Biological Membranes, Trends in Biotechnology (2019), https://doi.org/10.1016/j.tibtech.2019.10.009

Trends in Biotechnology

68. Gutbrod, K. et al. (2019) Phytol metabolism in plants. Prog. Lipid Res. 74, 1–17 69. Li, Y. et al. (2017) Identification of putative substrates of SEC2, a chloroplast inner envelope translocase. Plant Physiol. 173, 2121–2137 70. Yang, X. et al. (2019) Targeted control of chloroplast quality to improve plant acclimation: from protein import to degradation. Front. Plant Sci. 10, 958 71. Frain, K.M. et al. (2016) Protein translocation and thylakoid biogenesis in cyanobacteria. Biochim. Biophys. Acta 1857, 266–273 72. Mejia, E.M. and Hatch, G.M. (2016) Mitochondrial phospholipids: role in mitochondrial function. J. Bioenerg. Biomembr. 48, 99–112 73. Pfanner, N. et al. (2019) Mitochondrial proteins: from biogenesis to functional networks. Nat. Rev. Mol. Cell Biol. 20, 267–284 74. Henderson, J.C. et al. (2016) The power of asymmetry: architecture and assembly of the Gramnegative outer membrane lipid bilayer. Annu. Rev. Microbiol. 70, 255–278 75. Koebnik, R. et al. (2000) Structure and function of bacterial outer membrane proteins: barrels in a nutshell. Mol. Microbiol. 37, 239–253 76. Facey, S.J. and Kuhn, A. (2010) Biogenesis of bacterial inner-membrane proteins. Cell. Mol. Life Sci. 67, 2343–2362

77. Rollauer, S.E. et al. (2015) Outer membrane protein biogenesis in Gram-negative bacteria. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 370, 20150023 78. De Geyter, J. et al. (2019) Inner membrane translocases and insertases. Subcell. Biochem. 92, 337–366 79. Rajagopal, M. and Walker, S. (2017) Envelope structures of Gram-positive bacteria. Curr. Top. Microbiol. Immunol. 404, 1–44 80. Silhavy, T.J. et al. (2010) The bacterial cell envelope. Cold Spring Harb. Perspect. Biol. 2, a000414 81. Anne´, J. et al. (2017) Protein secretion in Grampositive bacteria: from multiple pathways to biotechnology. Curr. Top. Microbiol. Immunol. 404, 267–308 82. Shimon, L.J. et al. (1997) A cohesin domain from Clostridium thermocellum: the crystal structure provides new insights into cellulosome assembly. Structure 5, 381–390 83. Slutzki, M. et al. (2012) Indirect ELISA-based approach for comparative measurement of highaffinity cohesin-dockerin interactions. J. Mol. Recognit. 25, 616–622 84. Thomas, F. et al. (2013) A set of de novo designed parallel heterodimeric coiled coils with quantified dissociation constants in the micromolar to subnanomolar regime. J. Am. Chem. Soc. 135, 5161–5166

Trends in Biotechnology, --, Vol. --, No. --

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