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Synthetic Protein Scaffolds for Biosynthetic Pathway Co-Localization on Lipid Droplet Membranes Jyun-Liang Lin, Jie Zhu, and Ian Wheeldon ACS Synth. Biol., Just Accepted Manuscript • Publication Date (Web): 12 May 2017 Downloaded from http://pubs.acs.org on May 13, 2017

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ACS Synthetic Biology

Synthetic Protein Scaffolds for Biosynthetic Pathway Co-Localization on Lipid Droplet Membranes

†,#

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Jyun-Liang Lin , Jie Zhu , and Ian Wheeldon*

Department of Chemical and Environmental Engineering, University of California, Riverside USA 92521

*To whom correspondence should be addressed: Phone: (951) 827-2471 Fax: (951) 827-5696 Email: [email protected]

ACS Paragon Plus Environment

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ABSTRACT Eukaryotic biochemistry is organized throughout the cell in and on membrane-bound organelles. When engineering metabolic pathways this organization is often lost, resulting in flux imbalance and a loss of kinetic advantages from enzyme co-localization and substrate channeling. Here, we develop a protein-based scaffold for co-localizing multienzyme pathways on the membranes of intracellular lipid droplets. Scaffolds based on the plant lipid droplet protein oleosin and cohesindockerin interaction pairs recruited upstream enzymes in yeast ester biosynthesis to the native localization of the terminal reaction step, alcohol-O-acetyltransferase (Atf1). The native localization is necessary for high activity and pathway assembly in close proximity to Atf1 increased pathway flux. Screening a library of scaffold variants further showed that pathway structure can alter catalysis and revealed an optimized scaffold and pathway expression levels that produced ethyl acetate at a rate nearly 2-fold greater than unstructured pathways. This strategy should prove useful in spatially organizing other metabolic pathways with key lipid droplet-localized and membrane-bound reaction steps.

Keywords: Biocatalysis, Cascade catalysis, Ester biosynthesis, Membrane localization, Metabolic engineering.


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INTRODUCTION A defining feature of eukaryotic biochemistry is the spatial organization of metabolic pathways in and on membrane-bound organelles. This organization provides potentially viable targets for improving the rates of microbial biosynthesis. Key steps in yeast long chain fatty acid synthesis and lipid storage are localized to the endoplasmic reticulum (ER),1-2 many plant secondary metabolite pathways that produce valuable pharmaceuticals are often organized on vacuole and ER membranes,3 and the focus of this study, yeast ester biosynthesis, localizes to lipid droplets (LDs).4-5 The intracellular organization of these and other membrane-bound pathways is catalytically useful. Native multienzyme structures with controlled interenzyme distances and ratios of active sites can concentrate metabolites and drive reactions counter to unfavorable thermodynamics, isolate toxic intermediates from the rest of the cell, and prevent the loss of substrates to alternative pathways.6-7 When engineering synthetic pathways, the benefits of spatial organization are often lost, as differences in eukaryote physiologies can disrupt localization and translating pathways to the common bacterial host E. coli or other prokaryotes results in a loss of native organelles. Metabolic engineering and synthetic biology use genome engineering and pathway refactoring tools to optimize flux along targeted biosynthetic pathways by disrupting nonessential competing pathways, increasing the activity of native and heterologous enzymes to alleviate kinetic bottlenecks, and modifying transcription to control pathway expression.8 Enzyme co-localization has emerged as an additional strategy to re-create some of the effects of natural spatial organization and introduce substrate (or metabolite) channeling in synthetic pathways.9-12 Molecular scaffolds with domains that selectively bind pathway enzymes can create multienzyme aggregates with high enzyme concentrations, minimal distance between


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active sites, controlled ratios of enzymes, and optimized spatial organization.13 Using intracellular scaffolds of protein-protein interaction domains, this strategy has been used to colocalize pathways for mevalonate, glucaric acid, and butyrate biosynthesis in E. coli, resulting in 77-, 5-, and 3-fold improvements in product titers, respectively.11,

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Resveratrol and 1,2

propanediol titers have also been improved (5 and 4.5-fold, respectively) with plasmid-based DNA scaffolds that recruit and co-localize zinc-finger modified multienzyme pathways in E. coli.16 Controlled intracellular assembly of multistep pathways has also been achieved with RNA scaffolds. Modified enzymes that bind to aptamers within the scaffold were used to created two-, three-, and four-step pathways that enhanced product titers and in some cases overall production rates.17-18 These examples of multienzyme co-localization are evidence that pathway structure can improve metabolite titers, but the scaffolding technologies have been limited to cytosolic organization and are not accessible to membrane-bound pathways. The lack of membrane-based scaffolding technologies is beginning to be addressed with engineered synthetic lipid-containing scaffolds. Lipid/protein particles based on proteins from the lipid-encapsulated bacteriophage φ6 have recently been used to assemble a two-step biosynthetic pathway in E. coli.19 Enhanced product titers from engineered microbes with spatially organized pathways are not necessarily the result of equivalent increases in biosynthetic rates. Analysis of in vivo scaffolded pathways point to a range of effects that can produce higher titers. Co-localization and controlled clustering of enzymes in engineered pathways can benefit microbial growth by limiting the concentrations of toxic pathway intermediates,11, 20 improve the underlying kinetics of bottleneck enzymes,14 and minimize metabolic burden by enabling low pathway expressionlevels.10-11 Overall reaction rates can also be increased: controlled spatial organization can limit parasitic side reactions, enhancing flux along the desired pathway;18 molecular scaffolds can


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increase local substrate concentrations driving higher rates at low substrate concentrations;21-22 and, enzyme proximity can enhance pre-steady state kinetics.22 The limits of rate acceleration have recently been modeled, suggesting that with optimized multienzyme cluster size, architecture, and number per cell, up to ~6-fold enhancements in the rate of two-step pathways are possible.12 To take advantage of the benefits of spatial organization with membrane-bound pathways we engineered a protein scaffold in yeast to co-localize the ester biosynthesis pathway on the outer membrane surface of intracellular LDs. In yeast fermentations, low concentrations of short and medium chain volatile esters are synthesized through the condensation of an alcohol and acetyl-CoA by alcohol-O-acetyltransferase (AATase; Figure 1a). The produced esters (e.g., ethyl and isoamyl acetate, among others) are responsible for the distinctive smells of yeast fermentations and are industrially useful as solvents and flavor and fragrance compounds. The predominant AATase for short chain ester synthesis in S. cerevisiae is Atf1, with contributions from Atf2.23 Atf1 and -2 are membrane-associated proteins that localize to the ER and traffic to LDs during the stationary phase.4 Heterologous expression of Atf1 and -2 as well as AATases from other yeasts and fruit results in a loss of membrane localization and the formation of low activity aggregates in the cytosol.24 Overexpression in S. cerevisiae does not disrupt native localization and maintains high Atf activity. Upstream enzymes in the acetyl-CoA branch of yeast ester biosynthesis, aldehyde dehydrogenase (Ald6) and acetyl-CoA synthetase (Acs1), are naturally segregated from Atf1, localizing to the cytosol (Ald6 and Acs1) and mitochondria (Acs1).25-27 We reasoned that the spatial organization of upstream enzymes around the native intracellular localization of the terminal reaction step, Atf1, would produce high local enzyme concentrations and limit crosstalk with other metabolic pathways, thus increasing pathway flux


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and the conversion of intracellular acetyl-CoA to ethyl acetate under ethanol producing fermentation conditions. The re-localization of enzymes from their native targets to LDs requires a localization tag that can be fused to target proteins. From a library of protein domains, we identified a terminal fusion that localizes to yeast LDs under fermentation conditions. The identified oleosin domain formed the basis of a synthetic protein scaffold that was able to co-localize native S. cereisiae Ald6 and Acs1 with Atf1 on LDs (Figure 1b). Enzyme co-localization was achieved using cohesion-dockerin protein-protein interactions pairs from Clostridia species: cohesion domains localized to LDs by fusion to the oleosin scaffold with the corresponding dockerin domains fused to Ald6 and Acs1. Fluorescence microscopy and Förster Resonance Energy Transfer (FRET) studies showed that the enzymes are tightly clustered on LDs and in vitro kinetic analysis revealed increased rates of ethyl acetate synthesis due to co-localization. To rapidly optimize scaffold and pathway expression and structure, we used a colorimetric assay for ethyl acetate biosynthesis and screened a combinatorial library of scaffolds with a series of different strength promoters to drive enzyme and scaffold expression. This approach enabled us to re-localize upstream pathway enzymes around the native LD-localization of Atf1 and demonstrate the effects of co-localization on pathway flux and ethyl acetate fermentation titers.

RESULTS Identification of a yeast fermentation lipid droplet-targeting domain To identify a suitable LD-targeting domain, we screened a series of five protein domains known to localize to mammalian cell LDs and the plant LD-protein oleosin. The screened domains included terminal hydrophobic sections of a putative methyltransferase AAM-B,28 17β-


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hydroxysteriod dehydrogenase (17βHSD),29 the antiviral protein viperin,30 and a hepatitis C virus core protein (HCVcp).31 The library also included an internal domain of a guanine nucleotide exchange factor GBF-132 and Zea mays oleosin.33 Fluroescence microscopy of Cterminally modified cyan fluorescent protein (CFP) revealed that AAM-B-CFP and HCVcp-CFP co-localized with the ER protein marker Sec61 with C-terminally fused red fluorescent protein (Sec61-DsRed) during exponential growth. Trafficking to LDs during stationary phase was observed by co-localization with the LD protein marker Erg6 (Erg6-DsRed) after 24 hours (Figure 2a). The LD localization domains of 17βHSD, viperin, and GBF-1 were found to be primarily cytosolic in S. cerevisiae and therefore not suitable as targeting domains (Figure S1). Fluorescence microscopy of cells overexpressing oleosin with C-terminally fused CFP (OleCFP) showed vesicle-like structures adjacent to Sec61-DsRed, suggesting oleosin localization to premature LDs. Similar to AAM-B-CFP and HCVcp-CFP, Ole-CFP co-localized with Erg6DsRed during stationary phase, indicating LD targeting. Although Ole-CFP, AAM-B-CFP, and HCVcp-CFP could localize to LDs under aerobic condition, only Ole-CFP maintain LD localization under fermentation conditions to 72 hours (Figure 2b). In anaerobic cultures both AAM-B-CFP and HCVcp-CFP were dispersed in cytosol. As such, the plant protein oleosin was used in subsequent experiments for ester pathway localization. Fluorescence microscopy of overexpressed Acs1 and Ald6 with C-terminal fusions of Ole-CFP (Acs1-Ole-CFP and Ald6-Ole-CFP) showed that oleosin was able to re-localize both enzymes to LDs (Figure 2c). In the absence of oleosin, overexpressed Acs1-CFP and Ald6-CFP localized to the cytoplasm. Functional characterization was less successful as LD-targeted Acs1Ole-CFP and Ald6-Ole-YFP exhibited whole cell lysate activities equal to the cell background (Figure 2d). Acs1-Ole-CFP produced acetyl-CoA from CoA-SH and acetate at a rate of


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0.19±0.01 µmol min-1mg-1 of total protein, a value equal to the empty vector control (0.21±0.02 µmol min-1mg-1). Ald6-Ole-CFP converted acetaldehyde to acetate at a rate of 0.82±0.04 µmol min-1mg-1 of total protein, with the vector control limited to 0.75±0.02 µmol min-1mg-1. Nterminal fusions of oleosin were also found to have limited activity (Figure S2). Control experiments suggested that the low activities were not due to C-terminal fusions as cell lysates containing Acs1-CFP and Ald6-CFP exhibited activities significantly greater than the cell backgrounds (Acs1-CFP, 0.37±0.01 µmol min-1mg-1 of total protein, and Ald6-CFP, 1.32±0.12 µmol min-1mg-1). It is possible that re-localization to the membrane surface of LDs resulted in poor enzyme orientation restricting substrate access to the active sites. For example, the crystal structure of Acs1 shows a positively charged patch around the active site that could interact with the negatively charged phosphate end-groups of the LD membrane, thus limiting catalysis.34

Functional localization of enzymes to lipid droplet membranes Our strategy to recover Acs1 and Ald6 function focused on an oleosin-based scaffold with protein-protein interaction domains to selectively recruit each enzyme to the outer surface of LDs. We selected cellulosomic cohesion-dockerin interaction pairs because there are no natural homologs in S. cerevisiae and they have previously been used to selectively co-localize multienzyme structures in vitro35 and on yeast cell surfaces.36 The protein parts of the scaffold system are shown in Figure 3a including, oleosin (Ole) and two dockerin-cohesin pairs (D1-C1 from Clostridium perfringens37 and D2-C2 from Clostridium thermocellum38). The scaffold and modified enzymes also contained a protein identification tag (e.g., HA, His, or V5) for western blot analysis. The putative assembly of dockerin-modified Acs1 and Ald6 with an Ole-C1-C2 LD-localized scaffold is schematically depicted in Figure 3b (left). To demonstrate pathway co-


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localization Ald6-YFP-D1, Acs1-CFP-D2, and Atf1-DsRed were co-expressed with the Ole-C1C2 scaffold. S. cerevisiae harboring the pathway expression vector was cultured to stationary phase and imaged to confirmed pathway localization. In the absence of the scaffold, Ald6-YFPD1 and Acs1-CFP-D2 were observed in the cytoplasm, while Atf1-DsRed localized to LDs (Figure 3b, right). Both Ald6-YFP-D1 and Acs1-CFP-D2 co-localized to LDs with expression of the scaffold. Control experiments of YFP tagged scaffold confirmed LD targeting by colocalization with the Erg6-DsRed LD marker (Figure S3) and culture conditions did not affect oleosin function (Figure S4). It is interesting to note that measurable levels of the dockerin modified enzymes were not observed in the cytosol as judged by fluorescence microscopy (Figure 3). This suggests that the dockerin-cohesin interactions may occur early in the secretory pathway and/or there is a surplus of scaffold for enzyme attachment. Acs1 and Ald6 activities above cell lysate backgrounds were observed when enzymes were targeted to LDs by the synthetic protein scaffold. Localized Acs1-D2 produced acetyl-CoA at a rate of 0.19±0.01 µmol min-1mg-1 of total protein in cell lysate assays, while a vector control expressing Atf1 and the Ole-C1-C2 scaffold exhibited an activity of 0.16±0.01 µmol min-1mg-1 of total protein (Figure 3c). Functional localization of Ald6-D1 was also successful with measured activity reaching 0.82±0.05 µmol min-1mg-1 of total protein, a value greater than the background activity of 0.62±0.06 µmol min-1mg-1. Based on the kinetic results of overexpressed Ald6-D1 and Acs1-D2 along with our previous analysis of Atf1 kinetics,24 the order of activity of pathway enzymes was assumed to be Ald6>Atf1>Acs1. Of note is the reduced activities of LD-targeted Ald6-D1 and Acs1-D2 in comparison to cytosolically targeted enzymes (compare Figures 3c and 2d). Western blot analysis suggests that the reduction in measured activities was due to reduced expression levels (Figure S5).


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Spatial organization of a multienzyme pathway co-localized to LDs Theoretical and experimental analysis of substrate channeling in coupled enzyme reactions suggests that the distance between enzyme anchor points should be less than ~5 nm to produce significant enhancements in catalysis.39 The flexibility of the cohesion-based protein scaffold prevents accurate calculation of the distance between immobilized Ald6-D1, Acs1-D2, and LD-localized Atf1, as such we quantified pathway organization by FRET microscopy (Figure 4). Acceptor photobleaching experiments showed that scaffold-bound Ald6-YFP-D1 and Acs1-CFP-D2 maintained a FRET efficiency of 13±4%, while a CFP-tagged scaffold and YFPtagged Atf1 had an efficiency of 15±2% (< 4.5 nm average interenzyme distance).40-41 Control experiments with cytosolic YFP and CFP tagged scaffold did not produce a positive FRET signal. The interenzyme distances achieved with the LD scaffold are sufficient to produce an effect of metabolic channeling and are comparable to a LD-localized CFP-YFP fusion, which produced a FRET efficiency of 37±4% (< 3 nm average interenzyme distance).

Enzyme co-localization enhances acetyl-CoA and ethyl acetate biosynthesis rates Initial evaluation of the LD-localization strategy was accomplished using the genetic constructs schematically shown in Figure 5a. The scaffold Ole-C1-C2, Acs1-D2, and Ald6-D1 were co-expressed from a single copy number expression vector, while ATF1 was integrated in the genome at the yprc∆15 locus on Chr XVI, a previously characterized high expression integration site.42 PTEF1, PTDH2, PTDH3, and PADH1 constitutive promoters were used to transcribe ACS1-D2, ALD6-D1, ATF1, and OLE-C1-C2, respectively. Preliminary experiments testing the


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effect of episomal pathway expression showed reduced Atf1 expression when Ole-C1-C2 was coexpressed (Figure S6). Variable enzyme expression levels in the presence and absence of the scaffold would complicate analysis of the co-localization effect, thus ATF1 was integrated in the genome resulting in normalized pathway expression levels. To demonstrate the effects of pathway co-localization we first evaluated acetyl-CoA and ethyl acetate biosynthesis from two-step pathways (Ald6/Acs1 and Acs1/Atf1, respectively). Cell lysates prepared from stationary phase cultures were equilibrated with pathway substrates and cofactors (ATP, CoA-SH, and NADP+ for Ald6/Acs1; ethanol, ATP, and CoA-SH for Acs1/Atf1) and reactions were initiated by the addition of acetaldehyde (Ald6/Acs1) and acetate (Acs1/Atf1). To confirm that LDs remained intact, cell lysates containing Ole-CFP were prepared in parallel and LD morphology was confirmed by fluorescence microscopy (Figure S7). S. cerevisiae with integrated ATF1 produced acetyl-CoA at a rate of 85±20 nmol min-1mg-1 of total protein from natively expressed Ald6 and Acs1 (Figure 5b). The rate of acetyl-CoA synthesis was 93±15 nmol min-1mg-1 of total protein when Ald6-D1 and Acs1-D2 were overexpressed. Co-localization of the pathway enzymes on LDs with the co-expression of the Ole-C1-C2 scaffold increased the rate of acetyl-CoA production to 159±8 nmol min-1mg-1 of total protein, 1.8-fold enhancement over the control strain and unassembled pathway. Similarly, the rate of the last two reaction steps of ethyl acetate biosynthesis, Acs1/Atf1, increased 2.9 and 1.9fold over the control strain and unassembled pathway, respectively (Figure 5c; ATF1 integrated control, 14±1 nmol min-1mg-1 of total protein; unassembled control, 21±0.4 nmol min-1mg-1; and, co-localized pathway, 40±2 nmol min-1mg-1). Western blot analysis of the expressed pathway and scaffold revealed that enzyme levels were consistent across the tested strains (Figure S8).


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A traditional test for substrate channeling (i.e., the transfer of an intermediate from one active site to another without first diffusing to the bulk environment) is the competition for pathway intermediates by an orthogonal reaction.6-7 Competing side reactions reduce the overall pathway reaction rate by consuming pathway intermediates that diffuse to the bulk solution. The effect of substrate channeling is protection against competing reactions and an increase in the rate of the co-localized pathway. The cell lysate assays shown in Figure 5 approximates such a test because natively expressed enzymes (e.g., pyruvate dehydrogenase, citrate synthase, and acetyl-CoA synthetase-2, among others) compete for the acetate, CoA-SH, and acetyl-CoA intermediates of the Ald6/Acs1 and Acs1/Atf1 coupled reactions. The rates of the Ald6/Acs1 and Acs1/Atf1 reactions increased when co-localized on the LD-membrane while enzyme levels remained constant in comparison to the unassembled pathways, thus suggesting the possibility of substrate channeling along the structured ethyl acetate biosynthesis pathway. Additional channeling assays (e.g., controlled challenge assays or the tracking of radiolabeled intermediates; see ref. 4) are necessary to draw a conclusion of substrate channeling.

Optimizing pathway architecture and expression enhances ester biosynthesis To optimize pathway flux during fermentation, we created a combinatorial library of the pathway by varying scaffold design and promoter strength. Kinetic analysis of Acs1-D2 and Ald6-D1 suggested that Acs1 activity was limiting (Figure 3). Western blot analysis of the expressed pathway supports this conclusion. Comparison of Ald6-D1, Acs1-D2, and Atf1 expression levels relative to a GFP standard revealed low Acs1-D1 expression (Figure S9). In light of these results, our optimization strategy focused on increasing Acs1-D2 expression and varying the ratio of Acs1-D2 to Ald6-D1 on the LD-localization scaffold. A library of 60 pathway


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variants was created by homologous recombination in S. cerevisiae as previously described.43 The design of the genetic constructs and pathway variants are shown in Figure 6a. A second copy of Acs1-D2 with different strength promoters (PPGK1, PCCW12, PHXT7, and PADH1) was included to increase expression. LD-localized pathway stoichiometry was varied by expressing scaffolds with one, two, and three copies of C2 (cohesin from Clostridium thermocellum that corresponds to the D2 dockerin fused to Acs1), while scaffold expression was altered with a second promoter series (PRNR1, PPGI1, PRPL188, PREV1, and PADH1). Pathway naming followed the convention of PxPySz where Px refers to the promoter of Acs1-D2, Py refers to the scaffold promoter, and Sz refers to scaffold design. The genetic constructs of each library member were transformed into S. cerevisiae with the previously integrated copy of Atf1 and five randomly selected colonies of each pathway variant were independently screened (Tables S1). Screening was accomplished with a UV-Vis based ester quantification assay (Figure S10).44 Analysis of the screened library supports our hypothesis that Acs1 activity was likely limiting. Colonies that produced ethyl acetate three or more standard deviations above the library average of 46.6 mg L-1 had scaffolds with two or three C2 domains (Figure 6b). This subset included P2P5S3, P4P3S2, P4P5S3, P1P3S3, P2P4S3, and two colonies of P4P4S3. The highest producing colony, P2P5S3, expressed Ole-C1-3C2 from the strong Adh1 promoter and Acs1-D2 from a PGI1 promoter, and generated 84.5 mg L-1 of ethyl acetate over a 24 hour period. The control pathway, P0P5S1 (the construct evaluated in vitro; Figure 5) produced 33.6 mg L-1 of ethyl acetate, two standard deviations below the library average. To better evaluate the optimized pathway, the high producing P2P5S3 construct was isolated, amplified, and re-transformed to the ATF1 integrated strain of S. cerevisiae. Sequencing


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of the isolated plasmid confirmed that the pathway was assembled as designed and western blot analysis confirmed expression of all parts of the pathways (Figure S11). The re-transformed P2P5S3 pathway produced 14.8 mg L-1 OD-1 of ethyl acetate, productivity significantly greater than the control pathways with and without the expression of the scaffold (Figure 6c). Expression of an oleosin-only scaffold (P2P5Sole) that did not bind Ald6-D1 and Acs1-D2 produced 8.5±0.8 mg L-1 OD-1 of ethyl acetate. Similar ethyl acetate productivity was observed in the absence of the scaffold (P2P5S0; 8.5±0.9 mg L-1 OD-1) and with the un-optimized scaffold (P0P5S1; 8.3±0.1 mg L-1 OD-1). With the exception of Acs1 in the P2P5Sole control (the non-functional scaffold control), enzyme expression levels remained consistent across the tested pathway variants (Figure S11). In the case P2P5Sole, Acs1 expression was higher than other samples. But despite the higher expression of the putative bottleneck enzyme, ethyl acetate production remained low. It is interesting to note that the co-localization effect was significantly enhanced with the optimized scaffold architecture. A comparison of the scaffold-less controls and their corresponding scaffold samples (i.e., P2P5S0 vs. P2P5S3 and P0P5S0 vs. P0P5S1) revealed that the optimized scaffold increased ethyl acetate productivity by 1.7-fold, while the effect was limited in the un-optimized case. The enhanced production of ethyl acetate during fermentation was, in this case, mirrored by a similar enhancement observed in lysate assays. In vitro, the rate of the optimized pathway P2P5S3 was found to be ~2.0-fold greater than P2P5Sole and P2P5S0 (Figure 6d). Combined, the library screening results, GC analysis of the optimized and control pathways, along with the results of the in vitro analysis of the pathway in the presence and absence of the scaffold support the conclusion that the re-localization of upstream enzymes to the native localization of Atf1 increased the rate of ester biosynthesis.


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DISCUSSION Enzyme co-localization is proving to be an effective tool for metabolic engineering. By mimicking substrate channeling effects that occur in natural multienzyme complexes and bifunctional enzymes (e.g., malate dehydrogenase-citrate synthase, key steps in the TCA cycle that electrostatically channel pathway intermediates between active sites45) engineered intracellular scaffolding can enhance the rates and titers of microbial biosynthesis.11-12, 14-18, 20 In this work, we open this concept to pathway spatial organization on LD membranes by enhancing the productivity and rate of ethyl acetate biosynthesis in S. cerevisiae. Re-localizing upstream enzymes that naturally localize to the cytosol (Ald6 and Acs1) and mitochrondria (Acs1) to the outer surface of LDs —the natural localization of the terminal ester biosynthesis step, Atf1— produced a clustered three-step pathway that channels key intermediates. We targeted ester biosynthesis as the LD-localization of Atf1 is necessary for high alcohol acetyltransferase activity,24 but we anticipate that this and similar technologies may be applicable to other biosynthetic and signaling pathways with essential steps localized to intracellular membranes and compartmentalized in organelles. FRET microscopy and in vitro kinetic analysis of the LD-assembled pathway revealed that the nanometer spacing between co-localized enzymes (Figure 4) increased pathway rate (Figure 6). When measured in vitro, the rates of both the up- and downstream coupled enzyme reactions (Ald6/Acs1 and Acs1/Atf1) increased nearly 2-fold when assembled on LDs (Figure 5). Due to the presence of natively expressed enzymes competing for pathway intermediates (CoA-SH, acetyl-CoA, and acetate), the cell lysate assays are suggestive of substrate channeling: pathway rates increased when co-localized in the presence of competing reaction.6-7 Additional


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analyses are required to draw a conclusion of substrate channeling in both in the in vitro and in vivo environments. It is important to note here that while the in vitro lysate assays demonstrate an effect of enhanced kinetics due to colocalization, these results do not necessarily translate to increased production during fermentation. Lysate assays were performed in dilute solution and differences in reaction kinetics in comparison to the fermentation experiments could be due to differences in enzyme concentrations, the distance between LDs in dilute solution versus intracellular conditions, in vivo enzyme dynamics, and the presence of active cellular metabolism during fermentation. A comparison of the in vitro rate (Figure 5) and fermentation data (Figure 6) is an example of the disparity between lysate assays and in vivo synthesis. The kinetics of P0P5S1 are superior to the unassembled control P0P5S0 in vitro, but not in vivo. With optimized scaffold architecture and pathway expression levels (P2P5S3) the rate of ethyl acetate production on a per cell basis was 1.7-fold above the unassembled controls (P2P5S0 and P2P5Sole), and in this case lysate assays produced a similar enhancement in cascade rate. The optimized LD colocalized pathway achieved kinetic enhancements similar to those produced by cytosolic scaffolding. For example, protein scaffolding of a three-step glucaric acid pathway in E. coli increased rate by 5-fold, in part due to pathway co-localization and in part due to activation of the bottleneck enzyme (myo-inostiol oxygenase; MIOX).14 Cytosolic scaffolding of a threonine pathway in E. coli enhanced the production rate by ~50% while improving growth kinetics.20 Comparison to other systems that use cohesion-dockering pairs for enzyme colocalization is also relevant. In one engineered cellulosome example, colocalizing various cellulases to the outer surface of S. cerevisiae increased the rate of cellulose hydrolysis by more than 4-fold in comparison to an unassembled control.46


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With respect to rate enhancement, the example of mevalonate production in E. coli stands out.13 Co-localization of acetoacetyl-CoA thiolase (AtoB), hydroxy-methylglutaryl-CoA synthase (HMGS), and hydroxymethylglutaryl-CoA reductase (HMGR) produced a 77-fold increase in mevalonate titer over a 96-hour period in comparison to an unassembled control. This increase in titer resulted from a combination of effects including improved growth kinetics and improved rate. As a point of comparison, a 50-fold increase in reaction rate of a typical enzyme catalyzed reaction (Ea ~50 kJ mol-1) requires a temperature input of ~75 °C equivalent to fermentations above 100 °C, suggesting that in this case rate enhancement is only part of the overall co-localization effect. In our ester biosynthesis example, localizing ethyl acetate production to LDs did not affect cell growth (Table S2) but by controlling the flux of central metabolites (acetate, acetyl-CoA and CoA-SH) along an artificially structured pathway we were able to increase the rate of production on a per cell basis by ~2-fold. There is strong evidence that this increase in rate is due to colocalization: the optimized scaffold (P2P5S3) produced more ethyl acetate than the unassembled controls (P2P5S0 and P2P5Sole); and, quantitative western blot analysis showed that expression was consistent across strains and enzyme levels in the optimized strain were equivalent or less than expression in the control strains (Figure S11). Yeast fermentation naturally produces small quantities of esters including ethyl acetate as well as isoamyl and phenylethyl acetate.23 Translating Atf1-based pathways from yeast to E. coli has enabled the biosynthesis of a range of short and medium chain volatile esters including isobutyl acetate, isoamyl acetate, and butyrate esters.47-49 Ethyl acetate has also been produced, but rates and specific productivities have been limited.24,

47

Fermentation for 24-hours with

overexpressed Atf1 in E. coli produced 1.8 mg L-1OD-1.23 Chromosomal overexpression of ATF1 in S. cerevisiae produced only ~0.2 mg L-1OD-1 over the same time period (Figure 6c).


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Optimization of pathway expression levels and localization to LDs in yeast increased productivity to nearly 15 mg L-1OD-1. An unintended effect of pathway localization to LDs may be the partitioning of pathway products from the cytosol to the LD hydrophobic core, an effect that has been observed with the overproduction of carotenoids.50 In the case of toxic intermediates this could be beneficial, but in the case of ethyl acetate it would likely represent a parasitic loss. Given the high volatility of ethyl acetate, we suspect that there is minimal partitioning to LDs. By using cell localization and protein-protein interactions domains we engineered an enzyme co-localization scaffold that assembles the final three reactions steps of yeast ester biosynthesis on intracellular LDs. Re-organizing pathway enzymes and controlling the architecture of co-localization and optimizing expression level increased the rate of ethyl acetate fermentation. This is a possible strategy for engineering the flux of metabolites down natural and synthetic metabolic pathways with controlled intracellular localizations and targeting lipid modifying enzymes to the intracellular location of lipid storage, LDs.

Methods Strains and culture conditions All E. coil and S. cerevisiae strains used in this work are summarized in Table S3. All modified yeast strains were derived from BY4742. E. coli DH5α and BL21(DE3) used for molecular cloning and protein expression, respectively. E. coli was grown in Luria Broth (LB) (Sigma) medium containing 50 µg mL-1 ampicillin (Fisher) for cloning experiment and 50 µg mL-1 kanamycin (Fisher) for HA-myc-GFP-(His)6 expression. All S. cerevisiae BY4742 strains


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were grown in synthetic complete medium (SC) containing 0.67% yeast nitrogen base (BactonDickinson), amino acid supplements (Sunrise and Sigma), and 2% glucose (Sigma).

Plasmid and yeast strain construction The plasmids used in this study are listed in Table S4. All heterologous genes were codon optimized for expression in S. cerevisiae by IDT codon optimization tool and synthesized by a gene assembly method.51 DNA sequences of heterologous genes are provided in Table S5. Yeast genes, promoters, and terminators were amplified from genomic DNA. The peptide tags used for western blot were incorporated into proteins by including the DNA sequence in the appropriate PCR amplification primers. Cellular localizations constructs of potential LD targeting motifs were made by fusing a CFP gene to the 3’ of heterologous genes by overlap extension PCR (OE-PCR). The resulting fusions were cloned into pRS426-PPGK1-TPGK1 at NotI and SpeI sites. To analyze the cellular localization of oleosin, Ald6, Acs1, and oleosin-tagged Ald6 and Acs1, pRS426-PPGK1-CFPTPGK1 was first created with CFP at the NotI and SpeI sites of pRS426-PPGK1-TPGK1. Oleosin, Ald6, Acs1, and Ald6/Acs1-Oleosin were inserted at the SacII and NotI sites of pRS426-PPGK1CFP-TPGK1. S. cerevisiae gene-tagged strains, ERG6::DsRed and SEC61::DsRed, were constructed as previously described.4 Synthetic scaffolds, Ole-C1-C2 and its variants, were assembled from oleosin,33 X82 (C1),37 and cohesin (C2).38 Enzyme-dockerin and enzyme-fluorescent protein-dockerin fusions were assembled via OE-PCR. The resulting fragments were cloned into either pRS426-PPGK1TPGK1 at SacII and SpeI sites or p413-PGPD at SpeI and XhoI sites. pRS426-PPGK1-OleosinCohesin (C2)-YFP-TPGK1 was constructed with an insertion of YFP at NotI and SpeI sites and


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Oleosin-Cohesin (C2) at SacII and NotI sites. pRS426-PPGK1-ATF1-DsRed-TPGK1 was constructed with an insertion of ATF1 at SacII and NotI sites and DsRed at NotI and SpeI sites of pRS426PPGK1-TPGK1. Plasmids for overexpression of the pathways were assembled in either pRS316 or pRS426 by transforming promoters, genes, and terminators to S. cerevisiae through DNA assembler.43 The combinatorial library for pathway optimization included synthetic scaffolds with 2and 3-cohesin (C2) domains and a second copy of Acs1. Both the scaffold and Acs1 were combined with a series of differed promoters. Oleosin-X82 (C1)-cohesin (C2) was extended with 1- or 2-copies of cohesin (C2) at 3’ end by Gibson assembly. Casettes that included the additional copy of Acs1 and the scaffolds were also assembled by Gibson assembly. The resulting genetic components were transformed into S. cerevisiae for assembly by the DNA assembler moethod.43 Pathway vectors constructed by yeast homologous recombination were isolated by ZymoprepTM Yeast Plasmid Miniprep I kit. Isolated plasmids were transformed into E. coli, amplified, and sequenced. A second chromosomal copy of ATF1 was integrated at the yprc∆15 locus of chromosome 16 by homologous recombination. The cassette included the LEU2 marker amplified from pRS315, PTDH3, ATF1, and TTEF2, which were assembled into pRS316. Finally, epitope-tagged GFP was constructed with a forward primer containing HA and myc tags and a reverse primer containing His tag. The resulted fragment was cloned into pET29 between XbaI and XhoI sites. Fluorescence microscopy and FRET analysis Cells were imaged using an Olympus BX51 microscope (UPlanFL 100X 1.30 oilimmersion objective lens, mercury lamp) and fluorescence micrographs were captured by a QImaging Retiga Exi CCD camera. Images were processed using CellSens Dimension 1.7


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software (Olympus). In all cases, at least three cell populations were analyzed and images representative of the cell populations are shown. FRET analysis was accomplished using confocal laser microscopy (Leica SP5; Bannockburn, IL) on cells that had reached stationary phase by the quantitative acceptor photobleaching method. Pre- and post-bleaching images were captured with CFP excitation at 458 nm and YFP excitation at 514 nm using an argon laser and appropriate emission bands. The acceptor was bleached with high intensity at 514 nm and donor intensity in the region of interest was compared before and after acceptor bleaching. FRET efficiency (EFRET) was calculated on a pixel-by-pixel basis as EFRET = 1-(Dpre/Dpost). Positive FRET occurred if acceptor bleaching resulted in increased donor fluorescence. Preparation of cell lysates Yeast cells harboring expression plasmids were cultured to early stationary phase in 50 mL SC-Ura in 250 ml baffled flasks. Cells were harvested by centrifugation at 3,500 rpm for 5 min at 4 °C, and washed twice with 100 mM potassium phosphate buffer (pH 7.4) containing 2 mM MgCl2. Equal volumes of wet cell pellets and 425-600 µm diameter acid-washed glass beads (Sigma-Aldrich) were added to a 15 mL tube and resuspended in 1 mL ice-cold lysis buffer (100mM potassium phosphate buffer, 2 mM MgCl2, 2 mM dithiothreitol (DTT), and protease inhibitor cocktail (Roche), pH7.4). Cells were disrupted by vortexing 10 times for 30 s. After each vortexing, the suspension was kept on ice for 30 s. Beads were removed by centrifugation at 500g for 5 min at 4 °C and the supernatant was transferred to a pre-cooled 1.5 mL tube. Cell lysate protein concentrations were determined by a commercially available assay (Pierce 660 nm Protein Assay, Thermo Fisher) and quantified with Nanodrop 2000c. Enzyme and pathway activity assays


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Aldehyde dehydrogenase (Ald6) and acetyl-CoA synthetase (Acs1) activity assays were conducted as previously described.52 Aldehyde dehydrogenase activity reaction mixtures containing 0.4 mM DTT, 10 mM MgCl2, 0.4 mM NADP+, 15 mM pyrazole, and 100 mM potassium phosaphate buffer (pH 8.0) were used with 1-5 µg cell lysate protein. Reactions were initiated with 0.1 mM acetaldehyde. To determine the activity of acetyl-CoA synthetase, reactions consisting of 10 mM malate, 8 mM ATP, 1 mM NAD+, 10 mM MgCl2, 3 U malate dehydrogenase (Sigma), 0.4 U citrate synthase (Sigma), and 100 mM potassium phosphate (pH 7.4) were used with 1-5 µg cell lysate protein. Reactions were initiated with 0.2 mM CoA-SH and 10 mM potassium acetate. To evaluate pathway efficiency from acetyl-CoA synthetase (Acs1) to alcohol acetyltransferase (Atf1), reaction mixtures containing 10 mM MgCl2, 500 mM ethanol, 8 mM ATP, 1 mM CoA, and 100 mM potassium phosphate (pH 7.4) were used with 5 µg cell lysate protein. Reactions were initiated with 100 mM potassium acetate and were allowed to continue for 5 min. 1 g NaCl was used to stop the reaction and produced ethyl acetate was quantified by headspace gas chromatography (GC). Liquid transfers between the reaction mixtures and GC analysis tubes were accomplished rapidly to ensure minimal ethyl acetate losses due to evaporation. To evaluate pathway efficiency from aldehyde dehydrogenase (Ald6) to acetyl-CoA synthetase (Acs1), reaction mixtures containing 10 mM MgCl2, 0.4 mM DTT, 0.4 mM NADP+, 8 mM ATP, 15 mM pyrazole, 0.5 mM CoA and 100 mM potassium phosphate (pH 7.4) were used with 5 µg cell lysate protein. Reactions were initiated by adding 0.1 mM acetaldehyde and stopped by adding perchloric acid to a final concentration of 1 M. Samples were neutralized with KOH and the resulting solutions were analyzed by the acetyl-CoA Assay Kit (Sigma-Aldrich).


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All reaction assays were completed with a minimum of three replicates. In all cases the mean and standard deviation are presented. Protein expression analysis Western blots were performed using standard procedure. Briefly, 5 µg cell lysate protein were loaded on Any kDTM or 7.5% Mini-PROTEAN® TGXTM Gel (Bio-Rad) and run at 150 V for 30 to 60 min. Samples were electrophoretically transferred overnight to a PVDF membrane at 25 V. Membranes were blocked with 5% non-fat milk in Tris buffer saline with TWEEN-20 (TBST) buffer for 1 h at room temperature, washed, and incubated with primary antibody in TBST with 1% non-fat milk for 1 h. Secondary antibody in TBST with 1% non-fat milk was applied, and incubated at room temperature for 0.5 h. After washing with TBST, Immobilon Western HRP Substrate (Millipore) was used for signal detection and UN-SCAN-IT gel 6.1 software was used to quantify band intensity. The following antibodies were used: rabbit antiHA (71-5500; Invitrogen), mouse anti-His (A00186-100; Genescript), rabbit anti-V5 (V8137, Sigma), rabbit anti-myc (C3956; Sigma), mouse anti-GAPDH (MA5-15738; Thermo Fisher Scientific), goat anti-rabbit IgG-HRP (65-6120; Invitrogen), goat anti-mouse IgG-HRP (31430; Thermo Fisher Scientific). UV-Vis ester detection assay and library screening A colorimetric assay based on the stoichiometric conversion of esters to a ferric chelate complex was developed for rapid library screening.44 Briefly, hexane extracted ethyl acetate was first reacted with alkaline hydroxylamine to produce hydroxamic acid and ethanol. The hydroxamic acid product was subsequently reacted with ferric ions to form a ferric chelate complex with strong absorbance at 520 nm. Alkaline hydroxylamine solutions were prepared by combining equal volumes of 2.5% (w/v) solutions hydroxylamine•HCl and sodium hydroxide in


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95% ethanol. Ferric perchlorate stock solutions (0.8% v/v) were prepared by combining ferric chloride dissolved in 75% perchloric acid and deionized water. Ferric perchlorate working solutions were prepared from stock solutions by diluting 20 times in absolute ethanol. The colorimetric assay was performed by mixing 150 µL of hexane-extracted ethyl acetate and with 50 µl of alkaline hydroxylamine solution and allowing the reaction to proceed for 10 mins at room temperature. The produced hydroxamic acid was reacted with ferric perchlorate working solutions for 5 min to form ferric chelate complex. Ethyl acetate quantification was accomplished by measuring the absorbance of the ferric chelate complex at 520 nm and comparing to a standardized calibration curve. Single colonies were cultured overnight at 250 rpm and 30 °C in 2 mL of SC -Ura with 2% glucose in 14 mL culture tubes. Overnight cultures were transferred to 3 mL SC -Ura with 10% glucose in 15 mL conical tubes with a starting OD600 of 0.05. Fermentations continued for 24 h at 250 rpm and 30 °C. Post fermentation, 50 µL of culture was withdrawn for OD600 measurement and ethyl acetate was extracted with 3 mL hexane. Extraction was conducted with a rotary shaker (MAXQ 2508) for 10 min with vigorous mixing. 1.4 mL of the mixtures was transferred to microcentrifuge tubes and 13,000 rpm was used to separate the hexane and aqueous phases. Ethyl acetate extracted in the top layer was withdrawn and quantified using the developed colorimetric assay in 96-well microplates. Ethyl acetate fermentations Single colonies were inoculated into 2 mL SC -Ura and with 2% glucose in 14 mL culture tubes and grown overnight at 250 rpm and 30 °C. Overnight cultures were transferred to 20 mL SC -Ura with 2% glucose in 50 mL conical tubes. The cultures were initiated with OD600 of 0.05 cells, and fermented for 24 h at 250 rpm and 30 °C. 1.4 mL fermentation culture was


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transferred to a microcentrifuge tube, centrifuged at 13,000 rpm, and 1 mL fermented broth was harvested and transferred to a 10 mL glass vial (Fisher) for GC analysis. 1 g NaCl was added to the sample and 50 mg L-1 pentanol was used as the internal standard. Gas chromatography analysis Ethyl acetate was quantified by a headspace gas chromatography (GC) with a flame ionization detector (FID) (Agilent Technologies 7890A GC with CTC-PAL headspace mode injector). The separation of volatile compounds was carried out by Rtx®-WAX column (30 m, 0.32 mmID, 5 mm film thickness; RESTEK) with helium as carrier gas. The GC oven temperature was initially held at 30 °C for 7 min and increased with a gradient of 50 °C/min until 200 °C, and the held for 4 min. The FID was held at 250 °C. One-mL headspace gas was injected to the GC with 10:1 split ratio, and 1-pentanol was used as internal standard.

ASSOCIATED CONTENT The Supporting Information is available free of charge on the ACS Publications website. Strains, plasmids, and protein sequences used in this study; microscopy images; other figures described in the text (PDF).

AUTHOR INFORMATION Corresponding Author *E-mail: [email protected]

Current address #

Jyun-Liang Lin, Chemical Engineering, University of Texas at Austin


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Author Contributions †

JLL and JZ contributed equally. JLL and IW conceived the study. JLL, JZ, and IW designed

and analyzed the experiments. JLL and JZ conducted the experiments. All authors wrote the manuscript.

Notes The authors declare no competing financial interest.

ACKNOWLEDGEMENTS This work was supported by the Army Research Office MURI (#W911NF1410263) and NSF CBET-1403264. We thank Harvey Blanch for his comments and discussion. REFERENCES 1. Ivessa, A. S.; Schneiter, R.; Kohlwein, S. D., Yeast acetyl-CoA carboxylase is associated with the cytoplasmic surface of the endoplasmic reticulum. Eur J Cell Biol 1997, 74 (4), 399406. 2. Markgraf, D. F.; Klemm, R. W.; Junker, M.; Hannibal-Bach, H. K.; Ejsing, C. S.; Rapoport, T. A., An ER Protein Functionally Couples Neutral Lipid Metabolism on Lipid Droplets to Membrane Lipid Synthesis in the ER. Cell Rep 2014, 6 (1), 44-55. 3. Ziegler, J.; Facchini, P. J., Alkaloid biosynthesis: Metabolism and trafficking. Ann Rev Plant Biol 2008, 59, 735-769. 4. Lin, J. L.; Wheeldon, I., Dual N- and C-Terminal Helices Are Required for Endoplasmic Reticulum and Lipid Droplet Association of Alcohol Acetyltransferases in Saccharomyces cerevisiae. PLoS One 2014, 9 (8), e104141. 5. Tiwari, R.; Koffel, R.; Schneiter, R., An acetylation/deacetylation cycle controls the export of sterols and steroids from S-cerevisiae. EMBO J 2007, 26 (24), 5109-5119. 6. Spivey, H. O.; Ovadi, J., Substrate channeling. Methods 1999, 19 (2), 306-21. 7. Wheeldon, I.; Minteer, S. D.; Banta, S.; Barton, S. C.; Atanassov, P.; Sigman, M., Substrate channelling as an approach to cascade reactions. Nat Chem 2016, 8 (4), 299-309. 8. Lee, J. W.; Na, D.; Park, J. M.; Lee, J.; Choi, S.; Lee, S. Y., Systems metabolic engineering of microorganisms for natural and non-natural chemicals. Nat Chem Biol 2012, 8 (6), 536-546.


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41. Sahoo, H., Forster resonance energy transfer - A spectroscopic nanoruler: Principle and applications. J Photoch Photobio C 2011, 12 (1), 20-30. 42. Flagfeldt, D. B.; Siewers, V.; Huang, L.; Nielsen, J., Characterization of chromosomal integration sites for heterologous gene expression in Saccharomyces cerevisiae. Yeast 2009, 26 (10), 545-551. 43. Shao, Z. Y.; Zhao, H. M., Construction and Engineering of Large Biochemical Pathways via DNA Assembler. ACS Synth Biol 2013, 1073, 85-106. 44. Löbs, A.-K.; Lin, J.-L.; Cook, M.; Wheeldon, I., High throughput, colorimetric screening of microbial ester biosynthesis reveals high ethyl acetate production from Kluyveromyces marxianus on C5, C6, and C12 carbon sources. Biotechnol J 2016, 11 (10), 1274-1281. 45. Wu, F.; Minteer, S., Krebs Cycle Metabolon: Structural Evidence of Substrate Channeling Revealed by Cross-Linking and Mass Spectrometry. Angew Chem 2015, 54 (6), 1851-1854. 46. Tsai, S. L.; DaSilva, N. A.; Chen, W., Functional Display of Complex Cellulosomes on the Yeast Surface via Adaptive Assembly. ACS Synth Biol 2013, 2 (1), 14-21. 47. Rodriguez, G. M.; Tashiro, Y.; Atsumi, S., Expanding ester biosynthesis in Escherichia coli. Nat Chem Biol 2014, 10 (4), 259-265. 48. Layton, D. S.; Trinh, C. T., Engineering modular ester fermentative pathways in Escherichia coli. Metab Eng 2014, 26 (0), 77-88. 49. Tai, Y. S.; Xiong, M. Y.; Zhang, K. C., Engineered biosynthesis of medium-chain esters in Escherichia coli. Metab Eng 2015, 27, 20-28. 50. Gao, S.; Tong, Y.; Zhu, L.; Ge, M.; Zhang, Y.; Chen, D.; Jiang, Y.; Yang, S., Iterative integration of multiple-copy pathway genes in Yarrowia lipolytica for heterologous β-carotene production. Metab Eng 2017 accepted 04/13/2017 (in press). 51. Hoover, D. M.; Lubkowski, J., DNAWorks: an automated method for designing oligonucleotides for PCR-based gene synthesis. Nucleic Acids Res 2002, 30 (10), e43. 52. Postma, E.; Verduyn, C.; Scheffers, W. A.; Vandijken, J. P., Enzymic Analysis of the Crabtree Effect in Glucose-Limited Chemostat Cultures of Saccharomyces-Cerevisiae. Appl Environ Microbiol 1989, 55 (2), 468-477.


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Figures Legends

Figure 1. Pathway co-localization on intracellular LD membranes in S. cerevisiae. (a) Ethyl acetate biosynthesis in S. cerevisiae and the subcellular localization of aldehyde dehydrogenase (Ald6), acetyl-coA synthetase (Acs1), and alcohol-O-acetyltransferase (Atf1). Abbreviations: Cyt, cytosol; Mt, mitochondria; ER, endoplasmic reticulum; and, LD, lipid droplet. (b) A schematic diagram of the scaffolding strategy, co-localizing Ald6 and Acs1 with Atf1 on LDs via a membrane bound protein scaffold.

Figure 2. Identification of synthetic LD targeting domains in S. cerevisiae. Fluorescence microscopy analysis of subcellular localization of three LD tags including oleosin (Ole) and


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terminal domains of AAM-B(1-38) and hepatitis C virus core protein (HCVcp(118-164)) at 10 and 24 hours (a) and under fermentation conditions (b). Sec61-DsRed and Erg6-DsRed are ER and LD protein markers, respectively. Cell morphology is shown by phase contrast (false colored blue). CFP: cyan fluorescent protein; and, DsRed: red fluorescent protein. Scale bar: 1 µm. (c) Fluorescence microscopy analysis of subcellular localization of Acs1-Ole and Ald6-Ole. Scale bar: 1 µm. (d) Enzyme activity of modified Acs1 and Ald6, and empty vector controls in units of µmol min-1 mg-1 of total lysate protein. Cells overexpressing fusion constructs were cultured to stationary phase and cell lysates were prepared for kinetic assays (n=3). Schematic diagrams of LD localized Acs1-Ole-CFP and Ald6-Ole-CFP are shown above the activity data.

Figure 3. Functional re-localization of enzymes to LDs via synthetic membrane scaffolds. (a) Protein parts of the synthetic membrane scaffold and abbreviations used in the figure. C1 and D1 are X82 (a cohesin-like protein) and dockerin from C. perfringens, while C2 and D2 are cohesin and dockerin from C. thermocellum, respectively. (b) Schematic representation the LD assembled pathway (left). Fluorescence microscopy of the subcellular localization of Ald6-D1YFP, Acs1-D2-CFP, and Atf1-DsRed in the presence and absence of the scaffold at stationary phase (right). Scale bar: 3 µm. (c) Enzyme activity of Acs1-D2 and Ald6-D1 in units of µmol min-1 mg-1 of total lysate protein. Cell lysates from overexpressed Ald6-D1, Acs1-D2, Atf1, and the scaffold Ole-C1-C2 were analyzed for Acs1 and Ald6 activity. The mean and standard deviation are shown (n=3).


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Figure 4. FRET microscopy and quantitative analysis of multienzyme co-localization. Stationary phase cells expressing scaffold-bound Ald6-YFP-D1 and Acs1-CFP-D2, Ole-C1-C2-CFP and Atf1-YFP, Ole-C1-C2 with C-terminally fused CFP-YFP, and Ole-C1-C2-CFP with cytosolically expressed YFP were imaged and analyzed. Phase contrast micrographs and fluorescence images of the CFP donor, YFP acceptor, and raw FRET signal are shown. CFP signal is shown in green (Ex/Em: 426-450nm/502-538nm), YFP signal is shown in yellow (Ex/Em: 488-512nm532554nm), and the FRET signal is shown in blue (Ex/Em: 426-450nm/532-554nm). FRET efficiency was quantified using the Leica SP5 FRAP (fluorescence recovery after photobleaching) protocol with acceptor fluorescence photobleached to ~10% of its initial value. Presented data represents the mean and standard deviation of five samples. The scale bar represents 1 µm.

Figure 5. In vitro evaluation of pathway activity from Ald6 to Acs1 and Acs1 to Atf1. (a) Genetic constructs used to overexpress pathway genes. (b) Catalytic reactions of Ald6 and Acs1, and the overall pathway activity converting acetaldehyde to acetyl-CoA. (c) Catalytic reactions of Acs1 and Atf1, and the overall pathway activity converting acetate to ethyl acetate. The mean and standard deviation are shown (n=3). Activity is reported in units of µmol min-1 mg-1 of total lysate protein.

Figure 6. Optimizing ester biosynthesis pathway structure. (a) Genetic constructs and nomenclature of the combinatorial library used for pathway expression. (b) Histogram of ethyl acetate titers from the pathway library. The titers were quantified by the colorimetric assay described in the Method section. (c) Specific productivity of ethyl acetate from 24 hr


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fermentations with strains expressing the pathway without a scaffold (P0P5S0), with scaffold (P0P5S1), with optimized scaffold (P2P5S3), without the optimized scaffold (P2P5S0), and with an oleosin-only scaffold (P2P5Sole). The titers were quantified by GC-FID. The mean and standard deviation are shown (n=3). (d) The rate of ethyl acetate production from in vitro assays of lysates containing P2P5S3, P2P5Sole, and P2P5S0. The mean and standard deviation are shown (n=3). Activity is reported in units of µmol min-1 mg-1 of total lysate protein.


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TOC graphic 83x39mm (300 x 300 DPI)


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Figure 1. Pathway co-localization on intracellular LD membranes in S. cerevisiae. (a) Ethyl acetate biosynthesis in S. cerevisiae and the subcellular localization of aldehyde dehydrogenase (Ald6), acetyl-coA synthetase (Acs1), and alcohol-O-acetyltransferase (Atf1). Abbreviations: Cyt, cytosol; Mt, mitochondria; ER, endoplasmic reticulum; and, LD, lipid droplet. (b) A schematic diagram of the scaffolding strategy, colocalizing Ald6 and Acs1 with Atf1 on LDs via a membrane bound protein scaffold. 86x72mm (300 x 300 DPI)



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Figure 2. Identification of synthetic LD targeting domains in S. cerevisiae. Fluorescence microscopy analysis of subcellular localization of three LD tags including oleosin (Ole) and terminal domains of AAM-B(1-38) and hepatitis C virus core protein (HCVcp(118-164)) at 10 and 24 hours (a) and under fermentation conditions (b). Sec61-DsRed and Erg6-DsRed are ER and LD protein markers, respectively. Cell morphology is shown by phase contrast (false colored blue). CFP: cyan fluorescent protein; and, DsRed: red fluorescent protein. Scale bar: 1 m. (c) Fluorescence microscopy analysis of subcellular localization of Acs1-Ole and Ald6-Ole. Scale bar: 1 m. (d) Enzyme activity of modified Acs1 and Ald6, and empty vector controls in units of µmol min-1 mg-1 of total lysate protein. Cells overexpressing fusion constructs were cultured to stationary phase and cell lysates were prepared for kinetic assays (n=3). Schematic diagrams of LD localized Acs1-Ole-CFP and Ald6-Ole-CFP are shown above the activity data. 151x119mm (300 x 300 DPI)


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Figure 3. Functional re-localization of enzymes to LDs via synthetic membrane scaffolds. (a) Protein parts of the synthetic membrane scaffold and abbreviations used in the figure. C1 and D1 are X82 (a cohesin-like protein) and dockerin from C. perfringens, while C2 and D2 are cohesin and dockerin from C. thermocellum, respectively. (b) Schematic representation the LD assembled pathway (left). Fluorescence microscopy of the subcellular localization of Ald6-D1-YFP, Acs1-D2-CFP, and Atf1-DsRed in the presence and absence of the scaffold at stationary phase (right). Scale bar: 3 m. (c) Enzyme activity of Acs1-D2 and Ald6-D1 in units of µmol min-1 mg-1 of total lysate protein. Cell lysates from overexpressed Ald6-D1, Acs1-D2, Atf1, and the scaffold Ole-C1-C2 were analyzed for Acs1 and Ald6 activity. The mean and standard deviation are shown (n=3). 139x106mm (300 x 300 DPI)



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Figure 4. FRET microscopy and quantitative analysis of multienzyme co-localization. Stationary phase cells expressing scaffold-bound Ald6-YFP-D1 and Acs1-CFP-D2, Ole-C1-C2-CFP and Atf1-YFP, Ole-C1-C2 with Cterminally fused CFP-YFP, and Ole-C1-C2-CFP with cytosolically expressed YFP were imaged and analyzed. Phase contrast micrographs and fluorescence images of the CFP donor, YFP acceptor, and raw FRET signal are shown. CFP signal is shown in green (Ex/Em: 426-450nm/502-538nm), YFP signal is shown in yellow (Ex/Em: 488-512nm532-554nm), and the FRET signal is shown in blue (Ex/Em: 426-450nm/532-554nm). FRET efficiency was quantified using the Leica SP5 FRAP (fluorescence recovery after photobleaching) protocol with acceptor fluorescence photobleached to ~10% of its initial value. Presented data represents the mean and standard deviation of five samples. The scale bar represents 1 µm. 77x62mm (300 x 300 DPI)


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Figure 5. In vitro evaluation of pathway activity from Ald6 to Acs1 and Acs1 to Atf1. (a) Genetic constructs used to overexpress pathway genes. (b) Catalytic reactions of Ald6 and Acs1, and the overall pathway activity converting acetaldehyde to acetyl-CoA. (c) Catalytic reactions of Acs1 and Atf1, and the overall pathway activity converting acetate to ethyl acetate. The mean and standard deviation are shown (n=3). Activity is reported in units of µmol min-1 mg-1 of total lysate protein. 88x93mm (300 x 300 DPI)



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Figure 6. Optimizing ester biosynthesis pathway structure. (a) Genetic constructs and nomenclature of the combinatorial library used for pathway expression. (b) Histogram of ethyl acetate titers from the pathway library. The titers were quantified by the colorimetric assay described in the Method section. (c) Specific productivity of ethyl acetate from 24 hr fermentations with strains expressing the pathway without a scaffold (P0P5S0), with scaffold (P0P5S1), with optimized scaffold (P2P5S3), without the optimized scaffold (P2P5S0), and with an oleosin-only scaffold (P2P5Sole). The titers were quantified by GC-FID. The mean and standard deviation are shown (n=3). (d) The rate of ethyl acetate production from in vitro assays of lysates containing P2P5S3, P2P5Sole, and P2P5S0. The mean and standard deviation are shown (n=3). Activity is reported in units of µmol min-1 mg-1 of total lysate protein. 153x76mm (300 x 300 DPI)


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