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Designing a new entry point into isoprenoid metabolism by exploiting fructose-6-phosphate aldolase side-reactivity of Escherichia coli Jason R King, Benjamin Michael Woolston, and Gregory Stephanopoulos ACS Synth. Biol., Just Accepted Manuscript • DOI: 10.1021/acssynbio.7b00072 • Publication Date (Web): 04 Apr 2017 Downloaded from http://pubs.acs.org on April 5, 2017

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ACS Synthetic Biology

Designing a new entry point into isoprenoid metabolism by exploiting fructose-6-phosphate aldolase side-reactivity of Escherichia coli Jason R. King, Benjamin M. Woolston, and Gregory Stephanopoulos*

ABSTRACT: The 2C-methyl-D-erythritol 4-phosphate (MEP) pathway in Escherichia coli has been highlighted for its potential to provide access to myriad isoprenoid chemicals of industrial and therapeutic relevance and discover antibiotic targets to treat microbial human pathogens. Here, we describe a metabolic engineering strategy for the de novo construction of a biosynthetic pathway that produces 1-dexoxy-D-xylulose-5-phosphate (DXP), the precursor metabolite of the MEP pathway, from the simple and renewable starting materials D-arabinose and hydroxyacetone. Unlike most metabolic engineering efforts in which cell metabolism is reprogrammed with enzymes that are highly specific to their desired reaction, we highlight the promiscuous activity of the native E. coli fructose-6-phosphate aldolase as central to the metabolic rerouting of carbon to DXP. We use mass spectrometric isotopomer analysis of intracellular metabolites to show that the engineered pathway is able to support in vivo DXP biosynthesis in E. coli. The engineered DXP synthesis is further able to rescue cells that were chemically inhibited in their ability to produce DXP and to increase terpene titers in strains harboring the non-native lycopene pathway. In addition to providing an alternative metabolic pathway to produce isoprenoids, the results here highlight the potential role of pathway evolution to circumvent metabolic inhibitors in the development of microbial antibiotic resistance. KEYWORDS: metabolic engineering, biocatalysis, MEP Pathway, isoprenoid, biosynthesis, promiscuity, antibiotic resistance.

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Demand for inexpensive, renewable sources of commodity chemicals and high-value small molecules has increased interest in the microbial biosynthesis of secondary metabolites.1, 2 Isoprenoids make up a class of plant and bacterial secondary metabolites with direct utility as replacements to petrol-derived commodity chemicals, drugs, and synthetic precursors of bioactive small molecules. Isoprenoids comprise well over 55,000 natural compounds including linear terpenes and cyclic terpenoids, carotenoids, and various other compounds. Given the attractive physico-chemical properties of isoprenoids as advanced biofuels, fragrances, flavoring agents, and pharmaceuticals, the appeal of microbial fermentations to generate isoprenoids is growing, as demonstrated by the microbial production of artemisinic acid, a chemical intermediate to the antimalarial drug artemisinin, in Saccharomyces cerevisiae and more recently the fermentation of cane syrup to β-farnesene (C15H24), a plant-based sesquiterpene with broad industrial application.3, 4 Aside from their potential industrial applications, isoprenoid biosynthesis pathways are of interest in the development of antimicrobials and the understanding and treatment of antimicrobial resistance.5, 6 Several antimicrobial targets have been discovered from bacterial isoprenoid biosynthesis, including 1-deoxy-d-xylulose-5-phosphate synthase (DXS) and 1deoxy-d-xylulose-5-phosphate reductoisomerase (DXR or IspC), which are inhibited by alkyl acetyl phosphonates and fosmidomycin analogues, respectively, to combat malaria (Plasmodium falciparum), tuberculosis (Mycobacterium tuberculosis), and other intestinal and respiratory infections resulting from gram-negative pathogens such as E. coli and Haemophilus influenza.6 Due to strong evolutionary pressure to survive, human pathogens are likely to develop resistance to MEP pathway antagonists, as has already been reported in a lab-evolved mutant strain of E. coli bearing the S222T mutation in DXR that conferred resistance to like fosmidomycin and its


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analogues.5 Generally, antimicrobial resistance results from genetic mutations that allow the microbe to remove a drug—via extracellular efflux or direct catabolism of a drug—or to evade a drug—via mutation of the target protein to reduce the binding of a drug, or alterations in the host metabolism to bypass the target pathways entirely.7-10 Thus, new biosynthetic pathways for the production of isoprenoids would not only impact our access to this valuable class of compounds, but also increase our understanding of the modes of resistance to isoprenoid-specific antimicrobials inherent in the potential re-routing of host metabolism.11

Figure 1. Metabolic map of DXP biosynthesis and catabolism. Arrows in gray denote native E. coli metabolism, except for the MEP pathway that is highlighted in green. Blue arrows and characters denote natural and evolved alternatives to the biosynthesis of DXP and isoprenoids.

Two native metabolic pathways exist for isoprenoid synthesis: the 2-C-methyl-D-erythritol-4phosphate (MEP) pathway found in bacteria and the plastids of plant cells, and the mevalonate (MVA) pathway from eukaryotes and archaea (Figure 1). While both pathways synthesize isoprenoids via common precursor molecules—the constitutional isomers isopentenyl diphosphate (IPP) and dimethylallyl diphosphate (DMAPP)—, they differ in the routes used to



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generate IPP and DMAPP from central carbon metabolites. The MVA pathway consumes two NADPH and three ATP to generate IPP/ DMAPP from 3 molecules of acetyl coenzyme A (AcCoA; Eq. 1). The MEP pathway consumes two NADPH and only two high-energy phosphates (CTP and ATP) to generate IPP/ DMAPP from pyruvate (Pyr) and glyceraldehyde-3phosphate (GAP; Eq. 2). Considering the varied ATP requirements of the pathways and the altered redox state of the cell arising from the differential use of AcCoA vs. pyruvate and GAP, the MEP pathway converts glucose to IPP/DMAPP at 18% greater stoichiometric efficiency than the MVA pathway and has garnered much interest in harnessing this increased efficiency for industrial applications through metabolic engineering of bacterial hosts. 12

3 + + 2 + 2 + 3 → + 3 + 2 + 3 + + (; )

+ + 2 + 2 + + → + 2 + + + + (!; ")

The MEP pathway begins via an enzyme-catalyzed decarboxylative condensation of pyruvate and GAP, yielding DXP, the entry point into isoprenoid, thiamine, and pyridoxal biosyntheses (Figure 1). DXP synthesis is catalyzed by DXS—a crucial control point of DXP productivity and potential antimicrobial target.6,

13, 14

DXS is regulated at the genetic and protein levels so

endeavors to increase flux through this step will have to overcome feedback inhibition of DXS by IPP and DMAPP, which cannot be combatted by genetic approaches (e.g. promoter engineering and DXS overexpression) alone.15, 16 Indeed, Banerjee et al. have recently shown


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that enzyme engineering of the recombinant Populus trichocarpa DXS active site may, to an extent, alleviate this feedback inhibition of DXS.17 Alternatively, one could envision a biosynthetic bypass of DXS to produce DXP. Indeed, an alternative metabolic shunt to DXP was found in the photosynthetic proteobacterium Rhodospirillum rubrum, which produced DXP via the demethylsulfurylation of 1-methylthio-Dxylulose-5-phosphate (MTXP) an intermediate in a sulfur salvage pathway (Figure 1).18-20 Through evolutionary studies, a few other alternatives to DXS have been identified, including a mutation in E. coli aceE encoding pyruvate dehydrogenase subunit E1 with the mutation E636Q and native E. coli yajO gene, which encodes for a putative xylose reductase that may be involved in DXP biosynthesis under thiamine starvation conditions.11,

18, 21

Rodriguez-Concepcion,

Keasling, and co-workers imported the MVA pathway into E. coli and established a ∆dxs mutant strain with mevalonate auxotrophy.18 Using this system, the team evolved a ribB mutant strain, encoding the mutant riboflavin biosynthetic enzyme RibB G108S, that reduced ribulose-5phosphate directly to DXP through an unknown mechanism and yet-identified reductant (Figure 1).21 We hypothesized that a systematic approach could allow for the de novo biosynthesis of DXP and bypass DXS with well-defined chemistries. Here we describe the design and implementation of a synthetic pathway to produce DXP in vivo from native metabolites other than pyruvate, GAP, or AcCoA. The approach is based on the pentose catabolism of ethylene glycol- and propylene glycol-producing E. coli strains that have elevated concentrations of the reactive twocarbon substrate glycolaldehdye (GA) and three-carbon ketone hydroxyacetone (HA).22-24 Stable isotope mass spectrometry metabolomics demonstrated that D-arabinose-derived GA and exogenous HA can be combined via an enzyme-catalyzed aldol addition to produce 1-



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deoxyxylulose as a precursor of DXP in vivo. The ability of this new pathway to overcome DXS chemical inhibition and, separately, augment the production of the nonnative isoprenoid molecule lycopene in E. coli was demonstrated. Additionally, combinatorial biosynthesis was used to establish that HA could be produced in vivo through the methylglyoxal pathway in E. coli, which would enable the consolidated fermentation of pentoses such as D-arabinose to isoprenoids in a single cell. Aside from offering a new, direct route to produce DXP from HA and D-arabinose, our results highlight potential engineering targets in the future development of this pathway for increased productivity of isoprenoids from pentoses and add credence to the broad application of chemical logic to the de novo design of metabolic pathways in live cells.

RESULTS AND DISCUSSION Retro-biosynthetic design of a DXP pathway In order to alleviate any product inhibition at DXS by downstream MEP metabolites IPP and DMAPP, one could circumvent DXS entirely in the design of DXP biosynthesis. We hypothesized that retro-biosynthetic analysis could enable de novo DXP production as shown in Figure 2a. Retro-biosynthetic analysis applies the logic of retrosynthesis from organic chemistry to the modular reconstruction of metabolic pathways.25 Our analysis began with the insight of Wungsintaweekul et al. that DXP could be generated in vivo by the aberrant activity of the enzyme xylulose kinase (XK), encoded by the native E. coli gene xylB.26 We hypothesized that a non-native reaction with an E. coli aldolase might produce the 1-deoxyxylulose scaffold by a syn-aldol addition of the donor hydroxyacetone (HA) into the acceptor molecule glycolaldehyde (GA). The native E. coli fructose-6-phosphate aldolase (FSA), encoded by the gene fsaA, was previously found to catalyze the reversible aldol cleavage of fructose-6-phosphate to


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dihydroxyacetone and glyceraldehyde-3-phosphate, with a preference for the aldol addition reaction (∆G°’ = + 32 kJ•mole-1).27 Upon in vitro characterization of FSA, Schürmann, Sprenger, and co-workers indicated some substrate plasticity allowing for substitution of HA and GA as aldol donor and acceptor, respectively.27 Surprisingly, this aberrant activity occurs at 18% of the efficiency of the reaction between native donor dihydroxyacetone and acceptor glyceraldehyde3-phosphate.27,

28

Crucially, Schürmann et al. detailed the stereoconfiguration of the aldol

addition products of the FSA-catalyzed additions, including HA to GA, and they concluded that the product diols exclusively formed the syn-diasteromers, including 1-deoxy-D-xylulose (DX), rather than the anti-diastereomers, such as the C-3 epimer of DX, 1-deoxy-D-ribulose.28 Notably, DHAP was found to be an unacceptable substrate of FSA.27 Together, these characteristics detail the utility of FSA as a native E. coli catalyst for the “non-native” addition of HA to GA to form DX.

Figure 2. A) Retro-biosynthetic analysis of 1-deoxy-D-xylulose-5-phosphate (DXP); B) the engineering strategy for the de novo biosynthesis of DXP in a glycolate deficient E. coli host begins with the deletion of the native aldehyde dehydrogenase gene (aldA) to preclude glycolaldehyde (GA) oxidation to glycolate (denoted by the “X” symbol). Overexpressed enzymes include methylglyoxal synthase (MGS) and an aldehyde reductase (AR) to convert dihydroxyacetone phosphate (DHAP) to hydroxyacetone (HA) in module 1 coupled to the subsequent overexpression of fructose-6-phosphate aldolase (FSA) and xylulose kinase (XK) to convert glycolaldehyde (GA) and HA into DXP in module 2.



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In order to provide HA and GA for the aldol formation of DX, we looked to known microbial sugar fermentations to two and three carbon alcohols. HA has been previously shown to be an intermediate in the fermentation of hexoses to 1,2-propanediol in E. coli and Saccharomyces cerevisiae.29-32 Additionally, GA accumulation was used by Pereira et al. to augment ethylene glycol production from pentoses such as D-xylose and D/L-arabinose in E. coli.22-24 At this point, a forward de novo metabolic pathway of D-arabinose conversion to DXP was parsed into an upstream module (module 1; Fig. 2b) and a downstream module (module 2; Fig 2b), and the modules were separately evaluated in parallel in two E. coli strains. A combinatorial approach enables HA production Much work exists on the fermentation of pentoses to two- and three-carbon glycols via the intermediate overproduction of GA and HA. Pereira and co-workers previously showed that GA could be efficiently obtained from D-arabinose in E. coli lacking in the general aldehyde reductase activity of the enzyme AldA, encoded by aldA (Figure 2A). E. coli ∆aldA strains with elevated GA pools produced increased titers of ethylene glycol (EG) up to 3.4 g•L-1 and a corresponding yield on D-arabinose of 0.35 g•g-1 (0.85 mol•mol-1 or 34 % mole carbon basis).23 Notable adaptations of the system allowed for the reconstruction of other pentoses like D-xylose and L-arabinose metabolism to produce EG and glycolate.22,

23

Others have shown that the

natural tendency of E. coli to ferment excess dihydroxyacetone phosphate (DHAP) into the methylglyoxal (MG) shunt can be used to overproduce HA en route to the high titer synthesis of propylene glycol.33-36


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Figure 3. Hydroxyacetone (HA) titers from module 1 transformed into E. coli MG1655(DE3). On the left axis, blue bars show the HA titer in mM normalized to the culture optical density (OD) at 600 nm. On the right axis, the orange line and points show the mean HA titer of the cultures, demonstrating that HA toxicity (~100 mM, Figure SI-3b) is not limiting production. Error bars represent standard deviations from three biological replicates. Geneotypes of the strains are given in Table 1.

Given the prior conversions of sugars to GA and HA, module 1 construction began with a combinatorial screen for the two step conversion of DHAP to HA using either E. coli MG synthase (EcMGS) or Bacillus subtilis MG synthase (BsMGS) in combination with one of three native E. coli reductases: NADH-dependent MG reductase (YdjG), NADPH-dependent MG reductase (YeaE), and NADPH-dependent aldehyde reductase (YqhD) (Table 1). Six combinations of MG synthases and reductases were constructed under control of the T7/lac promoter system using the commercial pET28 vector (EMD Millipore, Billerica, MA). The constructs were transformed into the E. coli K12 MG1655 (DE3) ∆aldA base strain from Pereira et al.24 (hereafter E. coli ∆aldA), and the ability to ferment glucose into HA, as determined by HPLC analysis in comparison to an external standard of HA (Figure SI-2), was compared among the strains HA-01 – HA-06 (Figure 3 and Table 1). In comparison to the un-induced control, the engineered strains showed a significant increase in the basal HA titers from 50 µM up to 110 µM after 24 hours of growth on glucose in LB broth at 37 °C. A seven-fold increase in titer to 760 µM was observed when the E. coli reductase YqhD was used with BsMGS, and another 2.4-fold increase to 1.83 mM HA was observed when the MGS was exchanged for the E. coli isoform.



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Further evaluation of pathway induction temperature at 22 °C and 30 °C failed to increase HA titers above those observed at 37 °C. (Table SI-8). From the variable temperature tests, the major factor affecting HA titers was the choice of methylglyoxal reductase, with YqhD generating the highest titers near 2 mM. Given the observed HA titers, HA production was deemed sufficient to continue with examination of HA conversion to DXP in E. coli strains transformed with module 2. In vivo coupling of HA and GA to form DXP In parallel with the construction of HA producers, module 2 (Figure 2b) was used to convert Darabinose-derived GA and exogenous HA to DXP. To test whether FSA and XK could catalyze a chemical bypass of DXS, the genes were cloned on a pET28a backbone, yielding plasmids pDXP010 (fsa-, xylB+), pDXP020 (fsa+, and xylB-), and pDXP030 (fsa+, xylB+) (Table 1), which were used to overexpress the proteins FSA and XK (Figure SI-1). The genes fsaA and xylB were induced while growing the cells on uniformly 13C-labeled D-glucose in the presence of un-labeled (12C)-D-arabinose and un-labeled (12C)-HA (Figure 4a). Under these conditions, the cells were initially grown on heavy D-glucose, containing only 13C. When the cells reached midlog-phase growth, the induction mixture (IPTG for the pET genes and L-fucose to induce the endogenous fucIKA pathway for D-arabinose metabolism)23, 37 was added, which allowed for the overexpression of the synthetic pathway. At that point, light carbon substrates were also added. We hypothesized that incorporation of the light substrates (12C-Ara and

12

C-HA) into DXP

would present as a shift in the labeling ratio of heavy: light (H/L) DXP, as determined by LCMS/MS metabolite profiling, owing to the competing fluxes of carbon through the native DXS vs. our synthetic pathway. As native pathways also exist to allow the cells to consume the light arabinose through central carbon metabolism and form DXP through the DXS node, a basal


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degree of DXP lightening was expected in the presence of light carbon substrates. Given the empirical ease of observing phosphorylated metabolites via LC-MS/MS, the central carbon metabolites DHAP and PEP were monitored for the purpose of evaluating the differential use of the native vs. engineered pathways for DXP production. Assuming heavy carbon remains in the system, one would expect a differential shift in the H/L ratio of carbon isotopomers of DXP in comparison to DHAP and PEP as flux through the engineered pathway increases relatively to that of native metabolism for DXP biosynthesis, upon induction.

Figure 4. A) Labeling scheme for mass spectrometric determination of DXP origin in engineered E. coli. Metabolites are shown with circles to indicate number and connectivity of carbons, and the m/z [M-H]- of the metabolites is given in the upper left corner of the three-letter abbreviations. Shaded circles indicate 13C labeling (from glucose), and hollow circles indicate unlabeled 12 C (from arabinose and HA). Phosphate groups are indicated in blue. Native metabolism is shown in gray arrows, and the engineered pathway is shown in green. B - C) Ratio of heavy:light metabolites for engineered E. coli strains upon feeding un-



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labeled D-arabinose and HA to (U-13C)-D-glucose grown cells. Differential inclusion of IPTG and HA for panel C is indicated below the graph. Abbreviated genotypes are given in the inset boxes: A= EcmgsA; D = yqhD; F = fsaA; X = xylB.

As shown in Figure 4b, metabolite isotopomer profiling of strain E. coli ∆aldA at 24 h postinduction revealed a H/L ratio of 0.24 ± 0.31 for DXP and 1.1 ± 1.2 for DHAP indicating some flux of unlabeled arabinose and HA into DXP through the native metabolism. Similar H/L metabolite ratios of DXP (0.18 ± 0.22) and DHAP (2.4 ± 2.1) were observed when only XK was overexpressed in strain DXP-01. Upon overexpression of the FSA, a significant shift to lighter H/L ratios was observed in the DXP pool (H/L = 2.5 ± 3.2 × 10-3), while the DHAP metabolite resembled the negative control (H/L = 4.3 ± 5.2). That FSA alone enabled a lightening of the DXP pool in the absence of XK overexpression indicates that native XK activity of the cells was sufficient to allow phosphorylation of 1-deoxyxlyulose to DXP. This observation is consistent with prior reports in which 1-deoxyxylulose is able to complement an otherwise lethal knockout of the dxs gene.14,

26

When the FSA and XK were combined in strain DXP-03 a 105-fold

lightening of the DXP pool (H/L = 6.4 ± 4.9 × 10-5) was observed while significant heavy carbon remained in central metabolism as indicated in a DHAP ratio (H/L = 7.1 ± 10.7) that was similar with the negative controls (Figure 4b). This indicated a significant, 104-fold switch away from the use of DXS and glucose to form DXP in the engineered strains DXP-02 and DXP-03. Notably, the differential lightening of the DXP pool of strains DXP-02 and DXP-03 in comparison to the wild type strain confirmed that DXP was formed from the unlabeled substrates HA and arabinose-derived GA. Following in vivo confirmation of the FSA-mediated DXP synthesis with exogenous HA in strain DXP-03, a one-step conversion of D-arabinose into DXP was tested in which individual plasmids for pathway modules 1 (pHA011) and 2 (pDXP030)) were co-transformed to form strain DXP-04. The metabolite labeling with light carbon substrates in the presence of


13

C-

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glucose-grown cells was repeated with the EV-01 strain as the negative control, and the H/L metabolite ratios were measured for PEP, DXP, and 2-C-methyl-D-erythritol 2,4cyclodiphosphate (MEC)—a down-stream derivative of DXP. The metabolite labeling experiment again revealed significant DXP lightening (H/L = 3.87 ± 0.95 x 10-4) upon induction of the pathways with IPTG, but only in the presence of exogenously fed HA (Figure 4c). The MEC pool metabolite ratio (H/L = 4.46 ± 3.9 x 10-4) showed a similar profile to DXP, indicating that the DXP formed from HA and GA was further metabolized through the MEP pathway. While the results reaffirmed the use of FSA and XK to bypass DXS for DXP biosynthesis, the expected lightening of the DXP and MEC pools was only observed when HA was added exogenously, revealing a pathway limitation in the production of HA within strain DXP-04. Indeed, comparison of HA titers for strains DXP-04 to HA-01 that were grown in M9 media containing 2.5 g•L-1 glucose indicated no observable concentration of HA by strain DXP-04, while HA-01 generated 367 ± 0.5 µM HA (Figure SI-10). The reduced HA titers can be partially explained by the switch from LB-Glucose media to M9-Glucose that resulted in a decrease in HA specific titers of strain HA-01 from 0.59 mM•OD cells-1 in LB to 0.28 mM•OD-1 after 24 h in M9 media. However, strain DXP-05, which harbors the genes mgsA, yqhD, fsaA, and xylB on a single pET vector (pDXP040), produced a HA titer of 950 ± 60 µM in M9-Glucose media with a specific HA titer similar to the HA-01 strain grown in LB-Glucose media (0.67 mM•OD-1). That strain DXP-05 produced measureable HA while DXP-04 did not could also indicate insufficient MG and YqhD expression by the low-copy number plasmid pHA011. In vivo utility of a DXS bypass



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Figure 5. Fractional Growth of E. coli DXP strains in presence of the antibiotic butyl acetyl phosphonate (BAP). A) Schematic detailing BAP inhibition of DXS and the mode for rescue of engineered cells with hydroxyacetone (HA) and D-arabinose (Ara); B) graph of the fractional growth of engineered E. coli strain in the presence of BAP, HA, Ara and the pathway inducer IPTG. Fractional growth is normalized to the EV-01 positive control (far left bar) and error bars represent the standard deviation of triplicate cultures. Abbreviated genotypes are given in the inset box: A= EcmgsA; D = yqhD; F = fsaA; X = xylB.

We next sought to test the ability of the engineered pathway to complement a loss of DXS activity in the cell. The dxs gene is essential for growth in E. coli,38 and, due to its absence from human cells, DXS has been identified as an antibiotic target in human pathogens such as E. coli, Plasmodium falciparum, and Mycobacterium tuberculosis.6, 13, 14 As such, the deletion of dxs is of interest to define engineering strategies to enhance isoprenoid metabolism and inform metabolic routes to evolve resistance to DXS-inhibiting antibiotics. Previous efforts to disrupt DXS activity for engineering purposes have achieved moderate success and usually require ∆dxs complementation with a DXS-encoding plasmid or supply of an auxotrophic metabolite associated with isoprenoid or thiamine production.14, 18 Our initial efforts to complement a ∆dxs mutant E. coli strain with HA and pDXP030 failed. We hypothesized that


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a “gentler” approach than the “hard” deletion of the dxs gene might enable the gradual switch in metabolism to support cell growth and demonstrate the effective use of the DXS bypass. We thus synthesized butyl acetyl phosphonate (BAP), which was shown to be a selective inhibitor of E. coli DXS and effective antibiotic in thiamine-free minimal media when used in combination with a cell-weakening agent like ampicillin.14 Using 1.5 mM BAP and a sub-lethal dose of ampicillin (8 µg•L-1) in thiamine-free M9-glucose media, growth of the empty vector control strain (EV-01) was completely inhibited, and a combination of arabinose, HA, and IPTG did not rescue growth (Figure 5, lanes 1—4 ).50 Strain DXP-03 was also growth inhibited by BAP when only glucose and/or arabinose was fed, but the growth was rescued to 50% with HA alone or 54% with a mixture of HA and IPTG (Figure 5, lanes 5—8). That growth rescue for strain DXP-03 was observed with HA alone and that IPTG did not substantially improve the rescue suggest that basal pathway expression is enough to allow some conversion through the engineered pathway, which we believe is due to low level induction of FSA and XK in minimal media without IPTG. The lack of BAP rescue in the EV-01 strain supports this hypothesis. Strains DXP-04 and DXP-05 were also tested for the ability to rescue BAP-induced growth inhibition. Like the other strains, the cells did not grow in the absence of arabinose (Figure 5, lanes 11 & 15), nor did arabinose alone rescue the growth (Figure 5, lanes 12 & 16). Strain DXP04 harbors plasmid pHA011, which encodes pathway module 1 on an orthogonal expression vector to module 2 (pDXP030). This strain encoded a HA pathway that should allow for growth rescue without exogenous HA; however, substantial growth rescue for strain DXP-04 was only observed when HA was fed to the cells (Figure 5, lane 12). A similar, but greater, rescue was observed for strain DXP-05, which contains the full, engineered pathway (modules 1 and 2) on a single vector: pDXP040 (Figure 5, lane 16). From these results, it is clear that HA is able to



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rescue cell death caused from chemical-induced DXS inhibition only when the de novo DXP pathway has been transformed into the cells.

Figure 6. Fractional growth of engineered cells in presence of 1.5 mM BAP at 46 h and 37°C as function of hydroxyacetone concentration (left axis, blue bars) and predicted FSA activity as a function of [HA] (right axis, orange line and points). EV-01 controls were grown the absence (-) or presence (+) of BAP, and DXP-03 cells (fsaA+, xylB+) were grown with BAP and increasing concentrations of HA from 0 to 300 mM. Modeled FSA velocity was generated using Michaelis-Menten kinetics with constants obtained from Garrabou et al.39 for HA addition to G3P (Km = 17.4 mM; Vmax = 33.7 µmol min-1 mg protein-1. HA titer for HA-01 strain (1.8 mM in LB + Glucose) is represented by the black dotted line, and the toxic limit of HA for strain DXP03 is given in the red dotted line (>150 mM).

From the metabolite profiling and BAP rescue experiments, it is clear that HA feeding is necessary to enable de novo DXP biosynthesis. We hypothesized that poorly tuned FSA kinetics, rather than insufficient HA productivity, could be the cause of the apparent HA limitation in the pathway. This hypothesis is borne out by the observation by Garrabou et al. that native E. coli FSA catalyzes the addition of the aldol donor HA to aldehyde acceptor glyceraldehyde-3phosphate (GAP) with Km for HA of 17.4 mM.39 Given that our best HA producing strains (HA01 in LB-Glucose media) only reached HA concentrations of 1.8 mM (Figure 3), we hypothesized that the DXP strains were below the optimal range of HA concentrations for FSAcatalyzed addition to glycolaldehyde to generate substantial levels of DXP for cell growth under conditions of BAP inhibition of DXS. To test this hypothesis, the BAP rescue experiment was repeated using strain DXP-03, which lacks the HA module encoded by mgsA and yqhD, with HA feeding across the dynamic range of HA predicted from the Michaelis-Menten kinetics of FSA


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acting on HA and GAP (Vmax = 34 µmol•min-1mg-1 and Km(HA) = 17.4 mM).39 Figure 6 shows that extent of BAP rescue after 46.5 h in response to varying extracellular HA concentrations. Addition of 5 mM HA allowed for growth rescue of strain DXP-03 from BAP inhibition to 42 ± 4% that of the EV-01 control strain without BAP. This amount of growth rescue was similar to strain DXP-05 (50 ± 3 %), which produced an extracellular HA titer of 0.96 mM after 16 h of incubation in M9-Glucose media at 37°C (Figure SI-10). As the HA concentration was increased from 15 mM to 45 mM and 150 mM (near and above the predicted FSA Km for HA), the fractional growth increased with a strong correlation to the predicted velocity of FSA, which is plotted on the right axis of Figure 6. There is a qualitative shift in the curve of fractional growth of the DXP-03 cells and the FSA model as a function of HA concentration such that the apparent half-saturation point of DXP-03 response is near 45 mM HA. This difference could be explained either by the different partitioning of HA in and out of the cell such that the intracellular HA concentration is about 1/3 the extracellular concentration, or by a 3-fold increase in the FSA Km for HA as the acceptor aldehyde GAP is replaced here by glycolaldehyde. As the HA concentration was increased to 300 mM, no cell growth was observed, indicating the cells had crossed the toxic limit whereby HA began to inhibit growth. Interestingly, WT E. coli cells become growth inhibited at 50 – 100 mM HA, indicating that the DXP cells were tolerant to 1.5 – 3-fold higher concentrations of HA (Figure SI-3B). Due to the toxic nature of HA near the levels of maximal FSA velocity, the optimal engineering target to improve pathway productivity of the DXP strains would be a FSA mutant with a lower Km for HA, as has proven effective in the adaptation of FSA for other non-native aldehydes and ketones.40-42

To further probe the in vivo utility of the engineered DXP biosynthesis pathway, we looked to the ability of de novo biosynthesized DXP to augment carbon flux through the MEP pathway en



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route to isoprenoids. A common reporter system of isoprenoid pathway flux relies on the production of the tetraterpene molecule lycopene (C40H56), which is a chromophore with maximal visible light absorbance at 475 nm known for the red color it brings to tomatoes.43 The constitutive lycopene synthesis plasmid pAC-LYCipi was reported by Cunningham et al. to synthesize lycopene from IPP/DMAPP using the genes crtBEI and idi from the Gram-negative bacterium Pantoea agglomerans (formerly Erwinia herbicola).44 Strains EV-01, DXP-02, DXP03, and DXP-05 were transformed with pAC-LYCipi forming strains LYC-01 – LYC-04 (Table 1). The strains were grown on arabinose and HA with and without pathway induction, and cellular lycopene content was determined from dried cells by correlating visible light absorbance of organic cell extracts to a lycopene standard curve (Figures 7 and SI-9). The negative control strain, EV-01, showed no significant lycopene content. Strains LYC-01 and LYC-02 showed similar basal levels of lycopene content of 224 ± 21 ppm and 189 ± 19 ppm, respectively, with no change upon addition of IPTG. Strain LYC-03 showed similar basal lycopene content as strains LYC-01 and LYC-02 without induction (216 ± 14 ppm), but a 4-fold increase in lycopene content to 882 ± 74 ppm was observed upon induction of both fsaA and xylB with IPTG. This indicates that while FSA activity alone was enough to observe DXP biosynthesis by strain DXP02 in metabolite labeling and growth rescue experiments, it did not support lycopene production; however, when the full module 2 pathway was induced in strain LCY-03, the pathway flux through DXP and the MEP metabolites was enough to support lycopene synthesis.


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Figure 7. Lycopene content of engineered E. coli cells. A) schematic of the engineered pathway for the conversion of D-arabinose and hydroxyacetone to lycopene. B) lycopene content in ppm is shown in all strains at 24 h after induction of the engineered DXP biosynthesis pathway either without (teal) or with (orange) IPTG. Abbreviated genotypes are given in the inset box: L = pAC-LYCipi; A= EcmgsA; D = yqhD; F = fsaA; X = xylB.

The ability of the combination strain LYC-04 to combine the engineered pathway modules 1 and 2 for fermentation of arabinose to lycopene without HA was also tested (Figure 7). While only basal levels of lycopene were observed on arabinose alone (with or without IPTG), addition of exogenous HA (30 mM) upon induction of the plasmid pDXP040 presented a significant increase in lycopene content to ~2-fold the basal level (394 ± 60 ppm). The results mirrored the metabolite profiling and cell rescue experiments in which HA feeding was necessary to supplement insufficient flux through pathway module 1 for DXP biosynthesis, which further highlighted FSA-catalyzed HA conversion to DX as the principle target for future engineering efforts for increased DXP and isoprenoid production. It is interesting to note that lycopene titers



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of LYC-04 were ~2-fold less than the LYC-03 strain, even with HA supplementation. A possible explanation is reduced productivity in LYC-04 in comparison to LYC-03 owing to the increased burden of added gene expression.

In summary, we have demonstrated the synthetic potential of enzyme promiscuity and aberrant reactions in cellular metabolism. By utilizing the known promiscuity of native E. coli enzymes FSA and XK, we were able to show a redirection of DXP biosynthesis away from the native DXS node. The de novo DXP pathway was used to engineer resistance in E. coli to the DXS specific antibiotic BAP, and to augment non-native lycopene production in the cell by 4fold. The implications of this work are broader than the access of the isoprenoid precursor DXP, alone, as it opens the door to explore myriad carbon re-shuffling reactions in the cell by using native enzyme promiscuity to bypass critical control points of metabolism. We believe such approaches will add to the metabolic engineering and synthetic biology toolkits for unlocking the potential of secondary metabolism in the upgrading of renewable feedstocks with engineered biocatalysts. In addition to the synthetic utility of re-routing central carbon metabolism with aberrant reactions, the specific application to circumvent a potential antibiotic target in E. coli underscores the need for further insight into the role of metabolic plasticity in the evolution of microbial antibiotic resistance.

METHODS General

Agarose was purchased from Amresco (Solon, OH, USA). DNA manipulation was monitored by standard electrophoresis techniques using ethidium bromide (Sigma-Aldrich, St. Louis, MO,


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USA) to visualize the DNA under UV irradiation. Gel electrophoresis was powered by a BioRad Powerpac Basic power supply (Hercules, CA, USA), and gel documentation was performed with a ProteinSimple AlphaImager HP camera (San Jose, CA, USA). PCR reactions were performed on a Bio-Rad C1000 Touch Dual-block gradient thermocycler. Plasmids were synthesized using standard Gibson45 (isothermal type) or golden gate46 enzyme-assisted assembly techniques; whereby the requisite enzymes, buffers, and reagents were purchased from NEB, and synthetic primers were purchased from Sigma-Aldrich (see Table SI-9). Primer designs for Gibson assemblies were facilitated using the online tool NEBuilder (NEB), and primer designs for Golden Gate assembly were obtained using the built-in suite of the web-based cloning software from Benchling (San Francisco, CA, USA). Plasmid transformants were screened by colony PCR (standard PCR using an E. coli colony, rather than purified DNA, as the template) with plasmid-specific T7 or pACYC primers. Plasmid DNA was extracted from positive clones using the Zyppy plasmid miniprep kit (Zymo Research, Irvine, CA, USA), and insert sequences were confirmed by Sanger DNA sequencing (Quintara Biosciences, Boston, MA, USA). Genomic DNA from WT E. coli and Bacillus subtilis (type strain 168) was obtained from growing cultures using the Mo Bio UltraClean Microbial DNA Isolation Kit as per manufacturer instructions (Mo Bio Laboratories, Carlsbad, CA, USA).

Strains and Plasmids

Table 1. Plasmids and strains used in this study Name Plasmids pET28a pACYCDuet-1 pHA001 pHA010

Description

Reference

pBR322_T7-lacI-neo p15A_T7-lacI-cat pET28a_Ec.mgsA pET28a_Ec.mgsA-yqhD

Novagen Novagen This study This study



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pHA011 pHA020 pHA030 pHA040 pHA050 pHA060 pDXP010 pDXP011 pDXP020 pDXP030 pDXP040 pDXP041 pAC-LYCipi Strains WT HA-01 HA-02 HA-03 HA-04 HA-05 HA-06 ∆aldA EV-01 HA-07 DXP-01 DXP-02 DXP-03 DXP-04 DXP-05 DXP-06 LYC-01 LYC-02 LYC-03 LYC-04

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pACYCduet_Ec.mgsA-yqhD pET28a_Ec.mgsA-yeaE pET28a_Ec.mgsA-ydjG pET28a_Bs.mgsA-yqhD pET28a_Bs.mgsA-yeaE pET28a_Bs.mgsA-ydjG pET28a_xylB pET28a_xylB-His6 pET28a_fsaA pET28a_fsaA-xylB pET28a_Ec.mgsA-yqhD-fsaA-xylB pACYCduet_Ec.mgsA-yqhD-fsaA-xylB p15A_Pa.crtE-idi-crtI-crtB-cat

This study This study This study This study This study This study This study This study This study This study This study This study Cunningham et al. 200644

E. coli K-12 MG1655 (DE3) ∆endA ∆recA WT pHA010 WT pHA020 WT pHA030 WT pHA040 WT pHA050 WT pHA060 WT ∆aldA WT ∆aldA/pET28a WT ∆aldA/pHA011 WT ∆aldA/pDXP010 WT ∆aldA/pDXP020 WT ∆aldA/pDXP030 WT ∆aldA/pDXP030/pHA011 WT ∆aldA/pDXP040 WT ∆aldA/pDXP041 WT ∆aldA/pAC-LYCipi/pET28a WT ∆aldA/pAC-LYCipi/pDXP020 WT ∆aldA/pAC-LYCipi/pDXP030 WT ∆aldA/pAC-LYCipi/pDXP040

Pereira et al. 201623 This study This study This study This study This study This study Pereira et al. 201623 This study This study This study This study This study This study This study This study This study This study This study This study

Escherichia coli K-12 MG1655 (DE3) ∆endA ∆recA was used as the parent strain for metabolic engineering of HA producing strains and is designated hereafter as the WT strain. E. coli WT ∆aldA strain was reported by Pereira et al.23,

24

and was used as the parent strain of DXP-


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producing strains. NEB 5-alpha competent E. coli (New England Biolabs, Ipswich, MA, USA) was for routine cloning and plasmid maintenance. Plasmid pAC-LYCipi was a gift from Francis X Cunningham Jr (Addgene plasmid # 53279).44 Plasmid pHA001 was constructed by enzyme assisted, Gibson assembly45 of the gene fragments mgsA, which was PCR amplified from E. coli genomic DNA with primers JRKI151p001 and JRKI151p015 by Phusion high-fidelity DNA polymerase (NEB), and the protein expression vector pET28a, which was linearized by a digest reaction with endonucleases NcoI and XhoI in cutsmart buffer. Plasmid pHA011 was constructed by a three-fragment Gibson assembly with EcoRI/ SalI linearized expression vector pACYCduet, E. coli mgsA (amplified with primers JRKI151p067 & JRKI151p018), and yqhD (amplified with primers JRKI151p019 and JRKI151p068). Plasmids pHA010, pHA020, and pHA030 were prepared by 3-fragment Gibson assemblies of NcoI/ XhoI-linearized pET28a with E. coli mgsA—amplified with the common forward primer JRKI151p001 and variable reverse primers JRKI151p018 (pHA010), JRKI151p026 (pHA020), or JRKI151p013 (pHA030)—and E. coli yqhD (pHA010; primers JRKI151p019 & JRKI151p010), yeaE (pHA020; primers JRKI151p017 & JRKI151p008), or ydjG (pHA030; primers JRKI151p014 & JRKI151p006). Plasmids pHA040, pHA050, and pHA060 were prepared by three-fragment Gibson assemblies of NcoI/ XhoI-linearized pET28a with B. subtilis mgsA—amplified with the common forward primer JRKI151p003 and variable reverse primers JRKI151p022 (pHA040), JRKI151p020 (pHA050), or JRKI151p012 (pHA060)—and E. coli yqhD (pHA040; primers JRKI151p023 & JRKI151p010), yeaE (pHA050; primers JRKI151p021 & JRKI151p008), or ydjG (pHA060; primers JRKI151p011 & JRKI151p006).



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Plasmids pDXP010 and pDXP011 were prepared by Gibson assembly of BamHI/ SalI linearized pET28a and E. coli xylB amplified from genomic DNA with the common forward primer JRKI44p001 and different reverse primers JRKI44p002 (pDXP010) and JRKI144p003 (pDXP011). Plasmid pDXP020 was prepared by Gibson assembly of NcoI/ BamHI linearized pET28a and E. coli fsaA amplified using primers JRKI44p004 and JRKI44p005. Plasmid pDXP030 was prepared by a three-fragment Gibson assembly of NcoI/ SalI linearized pET28a, fsaA—amplified using primers JRKI44p004 and JRKII48p002—, and xylB—amplified with primers JRKII48p003 and JRKI44p002. Plasmid pDXP041 was prepared by Gibson assembly of BglII/ XhoI linearized pHA011 and the fsaA-xylB dual gene fragment that was amplified from pDXP030 using the primers JRKI151p069 and JRKI151p070. Due to difficulty with Gibson assembly arising from homology between ribosome binding sites in each fragment, plasmid pDXP040 was prepared by BsaI-mediated Golden Gate46 assembly of fragments pHA001— amplified with primers JRKI151p073 and JRKI151p074—, yqhD—amplified with primers JRKI151p075 and JRKI151p076—, fsaA—amplified with primers JRKI151p077 and JRKI151p078—, and xylB—amplified with primers JRKI151p079 and JRKI151p080. Media preparation, cell culture, and protein expression and analysis. LB and agar were purchased from Bacto Laboratories (Mount Pritchard, NSW, Australia) and prepared by manufacturer’s instructions. Isopropylthio-β-D-galactopyranoside (IPTG) was purchased from Gold Biotechnology (St. Louis, MO, USA), reconstituted as a 100 mM stock solution, sterile filtered through 0.22 µm syringe filter (Millipore, Billerica, MA, USA), and stored at -20 °C. Other media components, solvents, and antibiotics were purchased from SigmaAldrich. M9-Glucose media was prepared by combining the following components in a 1 L solution and sterile filtering with a bottle top vacuum filter (0.22 µm; VWR, Radnor, PA, USA):


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Bacto M9 Salt base (200 mL of 5X stock), D-glucose (2.5 g), MgSO4 (2 mL of 1 M stock), CaCl2 (0.5 mL of 0.1 M stock), ATCC vitamin mixture (10 mL; ATCC, Manassas, VA, USA), ATCC trace minerals solution (10 mL), and 18  Milli-Q water to 1 L. Thiamine-free M9 media was prepared as above while substituting the ATCC vitamin mixture with vitamin B12 (0.5 mL of 0.037 mM stock) and 0.5 mL of a synthetic vitamin mixture comprised of 10 mM NaPhosphate (pH = 7.1), sodium 4-aminobenzoate (0.26 mM), D(+)-biotin (41 µM), nicotinic acid (0.81 mM), calcium D(+)-pantothenate (0.21 mM), and pyridoxine hydrochloride (0.73 mM). Where specified, sterile antibiotic stocks were used to supplement the media with kanamycin sulfate (50 µg•mL-1) and chloramphenicol (30 µg•mL-1). Cultures were transformed with synthetic plasmids using the Inoue heat-shock method47 and grown overnight at 37°C on antibiotic LB-agar plates to select for positive transformants. Single clones were subsequently grown overnight in 3 mL LB or M9 media in a 14 mL Falcon tube (BD Biosciences, San Jose, CA, USA) at 37°C in an incubator shaker. The starter cultures were subcultured at 1:100 dilution ratio into pre-warmed media for growth and/or protein expression. When induced, the cells were grown to and optical density (OD; 600 nm observed wavelength) between 0.4 and 0.6 and IPTG and other additives were added. The cells were then cultured while shaking at 250 rpm at the specified temperature and time-lengths before collection of the cells or media by centrifugation or filtration. When necessary, protein expression was analyzed by SDS-PAGE analysis of B-Per Complete cell extracts that were prepared as per manufacturer’s instructions (Thermo Fisher Scientific, Waltham, MA, USA). SDS-PAGE was performed under standard conditions using BioRad power supply and pre-cast mini-protein 12% acrylamide gels and Instant-Blue protein stain (Expedeon, San Diego, CA, USA). Protein expression for DXP cells was optimal at 100 - 400 µM IPTG and 37°C for 12 – 24 h (Supporting Information).



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HA production and analysis in E. coli cultures Strains HA-01 HA-06, harboring plasmids pHA010 – pHA060, and WT E. coli were grown in starter cultures (LB with 50 µg•mL-1 kanamycin; 3 mL) over night at 37 °C before subculturing in to LB media (3 mL x 3 cultures) with kanamycin (except for WT) and 2 g•L-1 Dglucose in 14 mL Falcon tubes. The cultures were grown at 37 °C while shaking at 250 rpm until the OD reached 0.7 – 0.8; then, IPTG was added to 0.4 mM. After 6 h, the cells were removed by syringe filtration, and the filtrate was analyzed for presence of hydroxyacetone (HA) by ion exchange HPLC using an Agilent-1200 HPLC system with a G1362A RID detector while running a 20 min isocratic flow of 14 mM sulfuric acid (0.7 mL/min; 50 °C) through a BioRad Aminex HPX-87H ion exchange column (300 mm x 7.8 mm). Data collection and peak integration was performed using Agilent’s OpenLAB CDS ChemStation software (version A.02.08 SP1). HA eluted at 15.1 min and was quantified in comparison to an external standard and calibration curve (Figure SI-3). DXP synthesis, metabolite extraction, and labeling analysis in engineered cells. Strains EV-01, DXP-01, DXP-02, DXP-03, DXP-04, and DXP-05 were prepared by transformation of strain E. coli ∆aldA with the respective plasmids listed in Table 1. The strains were cultured over night at 37°C with shaking (250 rpm) in 14 mL culture tubes containing 3 mL M9 media with 2.5 g•L-1 (12C)-glucose. The next day, 75 – 100 µL of the starter was used to inoculate 3 mL of labeled M9 media containing 2.5 g•L-1 U-(13C)-D-glucose (Cambridge Isotope Laboratories, Tewksbury, MA). The cultures were grown at 37 °C with shaking at 250 rpm until the OD = 0.6 – 0.8. The cultures were then induced with the indicated mixture of (12C)-Darabinose (15 mM, final concentration), (12C)-hydroxyacetone (15 mM, final concentration), IPTG (80 µM, final concentration), and (12C)-L-fucose (1 mM, final concentration). Cultures


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continued shaking at 37 °C for 20 – 24 h before collecting the cells for metabolite extraction. Intracellular metabolites were extracted by a variation of the method described by Müller et al.51 3 mL of cell culture were rapidly filtered onto a 47 mm 0.45 µm Nylon Filter (Product #WHA7404004, Sigma Aldrich) and washed with 10 mL DI water. The filter was immediately immersed in 5 mL of -20 °C extraction solution (Acetonitrile:Methanol:Water 40:40:20) in a 50 mL Falcon tube and vortexed vigorously for 30 seconds. After 1 h lysis at -20 °C, the suspension was centrifuged (3750 RPM, 10 min), and the supernatant transferred to a glass test tube and dried overnight under air. The dried metabolites were resuspended in 120 µL DI H2O, centrifuged once more (20,000 RPM, 20 min) and the supernatant taken for LC-MS/MS analysis. LC-MS/MS analysis was conducted using a API 4000 triple quadrupole mass spectrometer (SCIEX, Framingham, MA) with ESI running in negative MRM mode with an Agilent 1100 Series HPLC (Agilent Technologies, Santa Clara, CA). The LC conditions were as described previously48, with an injection volume of 20 µL. Source parameters were as described by Luo et al.49 Metabolite-specific ionization and fragmentation voltages were obtained during infusion of 1 µM standard solutions using the automatic optimization features of the software (Analyst 1.6) and are listed in Supplemental Table SI-1. Integration of peaks was performed using the manufacturer’s software (Analyst 1.6) BAP inhibition of E. coli growth and rescue of DXP strains. Lithium butylacetylphosphonate (BAP) was synthesized as described by Smith et al.13 and stored desiccated in powder form at -20 °C until needed (see Supporting Information). Strains EV-01, and DXP-02 DXP-05 were grown overnight in 3 mL thiamine-free M9 media with 50 µg•mL1

kanamycin (plus 30 µg•mL-1 chloramphenicol for DXP-04) at 37 °C in 14 mL Falcon tubes.

The cultures were sub-cultured into 50 mL modified M9 media (2.5 g•L-1 D-glucose, no



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thiamine, 8 µg•mL-1 ampicillin, 25 µg•mL-1 kanamycin) and shaking continued at 37 °C and 250 rpm in 250 mL baffled flasks. At OD = 0.4, the cultures were transferred to 3 x 3 mL cultures in 14 mL Falcon tubes. Buffer additives and/or inducers were added from stock solutions to the following final concentrations according to the specifications in Figures 5 & 6: 1.5 mM BAP, 30 mM D-arabinose, 1 mM L-fucose, 30 mM hydroxyacetone, 80 µM IPTG, and 13 µg•mL-1 chloramphenicol. The cultures continued to shake at 250 rpm and 37 °C in a standing test tube rack over 2 days with periodic monitoring of culture ODs in a 96-well plate (part# 89131-504, VWR, Radnor, PA, USA) using SpectraMax M3 plate reader (Molecular Devices, Sunnyvale, CA, USA) with manufacturer’s software (SoftMax Pro 6.5). The culture ODs after 28.5 h postinduction were normalized to the BAP-free EV-01 control. Analysis of cell lycopene content Strains EV-01 and LYC-01 LYC-04 were grown overnight in 3 mL M9 media with DGlucose (2.5 g•L-1) and antibiotics (50 µg•mL-1 kanamycin, 30 µg•mL-1 chloramphenicol) at 37 °C while shaking at 250 rpm. The cells were sub-cultured into 3 x 5 mL cultures (6 x 5 mL for strain LYC-04), returned to 37 °C with shaking, and induced at OD = 0.5 – 0.6 with a mixture of D-arabinose (30 mM), L-fucose (1 mM), IPTG (5 µM for strains EV-01, LYC-01, and LYC-04; 25 µM for strains LYC-02 and LYC-03), and HA (30 mM; except none added to half of the LYC-04 cultures). After 20 h, the cells were collected by centrifugation (3750 rpm x 10 min), washed with water, and pelleted once more. The cells were frozen on liquid nitrogen and dried in pre-weighed Eppendorf tubes with a lyophilizer (Labconco, Kansas City, MO, USA). After recording the dry cell mass, lycopene was extracted from the cells. To extract lycopene, the cells were resuspended in a 1 mL of extraction solution (1:4:5 mixture of EtOH: EtOAc: CHCl3) and incubated at room temperature for 1 h with exclusion of light and intermittent mixing. The cell


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debris was pelleted at 15,000 rpm for 20 min, and the supernatant was carefully transferred to a fresh Eppendorf tube. The solvents were removed under a gentle flow of air for

Designing a New Entry Point into Isoprenoid ... - ACS Publications

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