Rapid Thermostabilization of Bacillus thuringiensis Serovar Konkukian

Aug 22, 2016 - Department of Chemistry, Northern Arizona University, P.O. Box 5698, Flagstaff, Arizona 86001, United States. § Chemistry Division, Lo...
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Rapid Thermostabilization of Bacillus thuringiensis Serovar Konkukian 97−27 Dehydroshikimate Dehydratase through a Structure-Based Enzyme Design and Whole Cell Activity Assay Lucas B. Harrington,†,# Ramesh K. Jha,†,# Theresa L. Kern,† Emily N. Schmidt,† Gustavo M. Canales,‡ Kellan B. Finney,‡ Andrew T. Koppisch,*,‡ Charlie E. M. Strauss,*,† and David T. Fox*,§ †

Bioscience Division, Los Alamos National Laboratory, P.O. Box 1663, MS M888, Los Alamos, New Mexico 87545, United States Department of Chemistry, Northern Arizona University, P.O. Box 5698, Flagstaff, Arizona 86001, United States § Chemistry Division, Los Alamos National Laboratory, P.O. Box 1663, MS E554, Los Alamos, New Mexico 87545, United States ‡

S Supporting Information *

ABSTRACT: Thermostabilization of an enzyme with complete retention of catalytic efficiency was demonstrated on recombinant 3-dehydroshikimate dehydratase (DHSase or wtAsbF) from Bacillus thuringiensis serovar konkukian 97−27 (hereafter, B. thuringiensis 97−27). The wtAsbF is relatively unstable at 37 °C, in vitro (t1/237 = 15 min), in the absence of divalent metal. We adopted a structure-based design to identify stabilizing mutations and created a combinatorial library based upon predicted mutations at specific locations on the enzyme surface. A diversified asbF library (∼2000 variants) was expressed in E. coli harboring a green fluorescent protein (GFP) reporter system linked to the product of wtAsbF activity (3,4dihydroxybenzoate, DHB). Mutations detrimental to DHSase function were rapidly eliminated using a high throughput fluorescence activated cell sorting (FACS) approach. After a single sorting round and heat screen at 50 °C, a triple AsbF mutant (Mut1), T61N, H135Y, and H257P, was isolated and characterized. The half-life of Mut1 at 37 °C was >10-fold higher than the wtAsbF (t1/237 = 169 min). Further, the second-order rate constants for both wtAsbF and Mut1 were approximately equal (9.9 × 105 M−1 s−1, 7.8 × 105 M−1 s−1, respectively), thus demonstrating protein thermostability did not come at the expense of enzyme thermophilicity. In addition, in vivo overexpression of Mut1 in E. coli resulted in a ∼60-fold increase in functional enzyme when compared to the wild-type enzyme under the identical expression conditions. Finally, overexpression of the thermostable AsbF resulted in an approximate 80−120% increase in DHB accumulation in the media relative to the wild-type enzyme. KEYWORDS: thermostabilization, shikimate pathway, flow cytometry, enzyme engineering, commodity chemicals

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upstream chemical precursors would significantly mitigate negative environmental impact on multiple levels.5 Over the previous decades, one rather appealing biosynthetic route to ADA and derivatives has come to the forefrontthe shikimate pathway. Pioneering studies by Frost and Draths2 revealed, through extensive genetic manipulation in Escherichia coli, the shikimate pathway intermediate, (−)-3-dehydroshikimate (DHS), could serve as the substrate for commodity chemical biosynthesis to numerous target molecules including 3, 4-dihydroxybenozoate (DHB), catechol, and cis,cis-muconic acid (ccMA) (Figure 1). Following nearly two decades of research, ccMA yields approached ∼30% mol/mol of glucose (∼60 g/L), which represents the highest yielding process to ccMA reported thus far. More recently, a promising alternative biosynthetic route to the identical target chemicals was suggested through genetic manipulation of Pseudomonas putida KT2440 (and related pseudomonads) using biological lignin

ydrocarbons derived from petroleum are the primary natural source of most organic chemicals. Recent estimates reveal that ∼98% of all chemicals produced in the U.S. are derived from petroleates and natural gas. A significant proportion of these compounds are high production volume (HPV) chemicals where it is estimated billions of pounds are generated domestically and trillions of pounds worldwide on a yearly basis.1,2 Many of the HPV chemicals encountered on a daily basis are derived directly from benzene, which is one of the top 20 most produced organic chemicals in the world, and is a known human carcinogen and toxic at low concentrations.3 Therefore, the production of HPV chemicals are directly linked to (1) depletion of our fossil fuel resources; (2) introducing carcinogenic agents into our environment; (3) generation of toxic organic and inorganic waste; and (4) production of teragrams of greenhouse gases (GHGs). As a representative example, millions of megatons (MMT) of adipic acid (ADA) are produced worldwide largely for Nylon-6.6 production. ADA synthesis begins with fossil fuels, proceeds through benzene, requires nitric acid, and produces teragrams of potent GHGs, largely as N2O.4 Clearly, any biological route to ADA and © XXXX American Chemical Society

Received: June 1, 2016

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Figure 1. Commodity chemical biosynthesis of 3,4-dihydroxybenzoate (DHB), catechol, cis,cis-muconic acid (ccMA) and derivatives in a bacterial host via the shikimate pathway.

valorization as the carbon source.6 Adipic acid has also been formed directly by an E. coli heterologous host from acetyl-CoA and succinyl-CoA biosynthetic precursors.7 Our path to pursuing the biological production of commodity chemicals was realized through an unrelated research platform in siderophore biosynthesis. Petrobactin is a siderophore produced by several members of the Bacillus cereus sensu lato group, including the mesophiles Bacillus anthracis (causative agent of the anthrax disease), and the human pathogen B. thuringiensis 97−27.8−11 Our team kinetically characterized heterologously expressed AsbF and unambiguously determined the enzyme was responsible for the biosynthesis of the unique 3,4-catecholate moiety found as part of the petrobactin scaffold.12 Further, phylogenetic analysis at the amino acid level revealed AsbF bore weak homology to DHS dehydratases (DHSase) found in ∼25 mesophilic organisms, largely of fungal and soil bacteria origin, respectively,12 but the petrobactin biosynthetic enzyme was observed to have a greater catalytic efficiency than their related DHSase counterparts. In addition, the enzyme is relatively thermolabile in the absence of divalent metal (t1/246°C ∼ 10 min) with a modest increase in thermal stability when incubated in the presence of divalent magnesium, which is a feature shared with previously characterized DHSases.12−15 On the basis of an analysis of structural studies of AsbF,16 the divalent metal appears to serve two roles: coordination to acidic residues in the active site ostensibly to bring them in close proximity to the substrate and, possibly, to coordinate with a hydroxyl oxygen in DHS to assist with the aromatization during catalysis. At this time, no known thermophilic DHSases are reported in the literature. Given the biosynthetic potential of AsbF in commodity chemical production, and the thermolabile nature of the

enzyme, we opted to pursue a research platform to increase AsbF thermostability with net retention of catalytic activity. The rationale for this approach is centered on the hypothesis that in vivo enzyme stability in heterologous chemical production efforts directly affects (limits) product titers. Previous strategies to increase the thermostability of individual enzymes were pursued in an effort to increase the longevity and/or specific activity of the biocatalytic enzyme(s) within the host. This includes improvements in enzyme stability by identification and installation of stabilizing mutations through computationally assisted rational design17−19 or through directed evolution approaches,20−22 as representative examples. In this study, we report a method for thermostabilization of the B. thuringiensis 97−27 DHSase, which represents the first step in the commodity chemical biosynthetic pipeline to adipic acid via the well-studied bacterial shikimate pathway (Figure 1). This was accomplished through a combined structure-based enzyme design and high-throughput screening platform. Detailed kinetic characterization and half-life studies revealed nearly an order-of-magnitude increase in enzyme half-life, in vitro, at 37 °C with complete retention of catalytic activity relative to the wtAsbF. Furthermore, we also observed a 50−60 fold increase in intracellular accumulation of functional thermostable DHSase relative to the wtDHSase post β-D-1thiogalactopyranoside (IPTG) induction at 37 °C. Finally, a separate in vivo experiment revealed the thermostable DHSase produced an approximate 80−120% increase in DHB accumulation in the media relative to the wild-type enzyme.

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RESULTS AND DISCUSSION Stabilization of Dehydroshikimate Dehydratase through Rational Design. Our workflow for DHSase B

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Figure 2. Work flow for enzyme thermostabilization studies. (A) Three methods for surface point mutations to enhance enzyme thermostability were adopted. Visual selection (light gray), VIP protocol (dark gray) and pmut_scan (black). Regions shown in “magenta” and “cyan” were excluded for catalytic pocket or dimer interface. (B) Mutations suggested by the varying protocols. Multiple suggested mutations are shown in parentheses. Double mutations implicated to stabilize AsbF are depicted in square brackets. (C) Mutations selected for fixed-backbone design. The first amino acid represents the native sequence. (D) Mutations suggested by fixed-backbone design, native (green) and mutation (black). (E) Mutations based on fixed-backbone design. (F) Double codons or a degenerate codon that codes for native and one or more mutations. For example, at position 257, a single degenerate codon MMT encodes for H (native), T (mutation from fixed backbone), P (mutation suggested in pmut_scan) and N (an extra mutation). (G) A mutant isolated from the library screen. Native side chains (green) and mutation (black) are overlaid one another. (H) Residue specific calculated Rosetta energy compared for wild-type (wt) AsbF and the triple mutant (Mut1).

server24 was also utilized to aid our design, and this protocol suggested six mutations, largely to aid thermostability through space filling and an increase of van der Waals interactions with neighboring residues. Mutations that served to modify alkyl positions with other hydrophobic groups (such as V → L or L → F) were common outputs from this approach. In a final approach, we used a Rosetta “point mutation scan” protocol to search the entirety of the protein sequence to select for stabilizing mutations. Mutations were identified that encompassed the complete sequence of the protein, but further analysis was limited to a subset of the identified mutants, specifically those which were located on the surface and away from the dimer interface or catalytic pocket. Of these, nine mutations consisted of seven independent residues of each other and two (N27R/K264E) that were expected to coexist, ostensibly to form a novel salt-bridge. Next, all identified mutations were simultaneously modeled onto the structure of DHSase in order to select combinations that likely exerted the largest positive influence upon enzyme stability. Interestingly, only 10 out of a total of 24 positions revealed an amino acid substitution that varied from the native sequence. The combined effect of all mutations were calculated to have a ΔG of −10 Rosetta Energy Units (REU) over the native relaxed structure, which was based upon a newly parametrized energy function in Rosetta.25 Dehydroshikimate Dehydratase Library Design. Any mutations made to an enzyme, though predicted to be

thermostabilization is depicted in Figure 2. In order to develop a comprehensive understanding of the structure based protein design to affect stability of our target, we applied multiple design approaches to identify positions for mutagenesis on a three-dimensional structure of DHSase. The DHSase sequence from B. thuringiensis 97−27 has 100% protein sequence identity with DHSase from Bacillus anthracis Sterne, and the threedimensional structure of the latter enzyme has been reported (PDB code 3DX5).16 The B. anthracis DHSase is an apparent homodimer, according to the crystal structure. Hence, residues within the dimer interface (4 Å cutoff) and side chains at the dimer interface (Figure 2 A, cyan) were excluded from consideration for mutational studies (Figure 2A, cyan). Similarly, amino acids in close proximity to the first shell residues surrounding the metal ion were also excluded (Figure 2A, magenta). Upon considering these exclusions, we selected nine positions that had potential to influence thermostability, namely those which were exposed to solvent and amenable to relatively conservative structural changes (such as mutations of smaller side chains to larger side chains or from neutral to charged side chains). Additionally, several hydrophobic residues were chosen for mutation to charged amino acids to enable installation of potential salt bridges with neighboring oppositely charged amino acids. For example, the mutations I15E and L75R were anticipated to create novel charge−charge interactions with other (basic and acidic, respectively) side chains in the protein. The VIP protocol23 on the ROSIE C

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ACS Synthetic Biology stabilizing via computational resources, may ultimately prove to be detrimental to enzyme expression, its ability to fold into the proper solution structure, or both. In terms of the enzyme’s catalytic competence, the point mutation(s) may inadvertently negatively affect any requisite allosteric requirements or, alternatively, increased stability gained from the substitutions may result in decreased enzyme thermophilicity (that is, increased Km for the substrate(s), decreased kcat due to improper substrate alignment in the active site pocket or slow product release, as representative examples). Hence, instead of screening a single AsbF variant with all ten identified substitutions, we opted to create a combinatorial library consisting of constructs containing one or more Rosettapredicted substitutions (Figure 2E). Wherever applicable, a single base “wobble” would represent a predicted substitution of the native amino acid that was readily incorporated using a single mutagenic primer. In cases where more than one base required modification, two different oligonucleotides were synthesized, each having a codon representing a mutant sequence and the wild-type sequence. For example, an oligonucleotide with two “wobbles” in a codon at position 257 was used in order to represent a mutation and retention of the wild-type sequence. The degenerate codon “MMT” coded for histidine (wild-type), threonine (computational mutation), proline (a suggested mutation from the point mutation scan protocol) and asparagine (often located in loops, as well as suggested with low frequency in the point mutation scan protocol) was also employed. The combinatorial focused asbF library was created with degenerate oligonucleotides specifically designed to introduce the desired amino acid substitutions on the surface of the enzyme, and was then assembled through gene splicing and overlap extension PCR (SOEing).26 Upon the basis of our library assembly design, the theoretical library size was estimated to contain 2048 independent variants. The corresponding list of degenerate overlapping primers and the asbF gene map are provided in the Supporting Information (Table S1 and Supplementary Figure S1, respectively). Upon assembly, the library was ligated into pRSF-1b, chemically transformed into E. coli DH5α and the library diversity was verified by the Sanger sequencing method. The corresponding asbF library was subsequently transformed into E. coli harboring the pGLO.pcaU.eRBS sensor plasmid previously developed in our laboratory.27 Briefly, the two-plasmid system provided a convenient handle for indirectly monitoring enzyme activity through flow cytometry using GFP fluorescence as a single-cell reporter. Upon induction of AsbF expression with IPTG, the endogenous shikimate pathway intermediate, DHS, is converted to DHB, which is readily detected at 290 nm by UV−vis spectroscopy.11,12 Expression of GFP under the control of the PcaU promoter is initiated upon binding of the intracellular DHB to the PcaU transcription factor. As no intracellular DHB is expected in the absence of AsbF enzyme activity, GFP expression must necessarily be linked to an active AsbF.27 Enzyme Thermostability, Heat Challenge and HighThroughput Screen. In our preliminary work, individual AsbF variants were overexpressed in a 96-well plate, the cells lysed, and the lysate subjected to a heat challenge (37, 46, or 50 °C) for 1 h. Residual enzyme activity was determined through an end-point titration at a saturating concentration of DHS (100 or 300 μM, with Mn(II) or Mg(II) in the assay buffer, respectively). Of the 279 variants assayed for residual activity following the 50 °C heat challenge, ∼95% were either inactive

or weakly active. The presence of active AsbF were 28% and 18% of the assayed clones from the 37 and 46 °C heat challenges, respectively. Due to the imperfections associated with computational enzyme redesign process, there was some expectation the library would contain a large percentage of AsbF variants with little to no demonstrable catalytic activity. Examples include intrinsic PCR errors, increased enzyme rigidity resulting in decreased substrate specificity in catalytic pocket, enzyme misfolding, increased protease susceptibility and/or negative long-range effects on catalytic competency. Certainly, the rational design approach decreases the sequence space to be explored relative to directed evolution strategies, the latter of which can exhibit an astronomically large theoretical sequence space (10130 for 100 amino acid sequence). Design of an effective screen of a library of this size for a desired physical property is beyond the capability of any known experimental technique. The focused library approach using computational design was implemented to avoid exploring this impossibly large sequence space and, instead, to discretely focus on specific amino acids that may be implicated in AsbF thermostabilization. Conversely, the computationally designed library can still expand to cover the sequence space upon combinatorial introduction of the mutations at ten amino acid positions. As such, adopting a 96-well format to screen the entire sequence space of the library (∼103−104) proved tedious. In order to expedite the library screening process, E. coli cells harboring active AsbF variants were separated from inactive or nonfunctional AsbF variants through a single-cell biosensor technology previously developed in our laboratory.27 Upon induction of the AsbF library with IPTG and subsequent fluorescenceactivated cell sorting (FACS) analysis, approximately 25% of the cells exhibited detectable GFP fluorescence comparable to the wild-type AsbF (Figure 3a). Within the GFP fluorescing cell

Figure 3. (a) Flow cytometry analysis of the AsbF library using whole cell biocatalysis and biosensing. The top panel depicts the histograms for the controls (empty vector and wtasbF). The bottom panel depicts a histogram for the cells expressing AsbF variants and shows a clear bimodality. The cells that fall within the box representing the top 50% of the bright peak were sorted, regrown, then subjected to the library screen. (b) Comparative analysis of the AsbF variants in presorted population and postsorted population that were active post heat challenge at different temperatures.

population, the top 50% were collected and subjected to the aforementioned heat challenge and residual enzyme activity assessed. Encouragingly, a significant enrichment in AsbF activity was observed across all three heat challenge temperatures (Figure 3b, Table S2). The most striking enrichment was observed at 50 °C where nearly one-third (31%) of all variants tested (93 total) exhibited AsbF activity after the 1 h heat challenge versus only 5% in the presorted library. Wild-type D

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t1/2 (37 °C, min)a

kcat (min−1)b

KM (μM)

wtAsbF

15 ± 0.75

Mut1

169 ± 22

130 ± 3 29 ± 2 150 ± 4 42 ± 2

± ± ± ±

2.2 4.6 3.2 7.4

0.4 1.4 0.5 0.1

kcat/KM (M−1 s−1) (9.88 (1.05 (7.84 (9.60

± ± ± ±

1.82) 0.37) 1.43) 0.53)

× × × ×

Ea (kJ mol−1) 5

10 105 105 104

60.8 ± 1.9 59.0 ± 1.9

Incubation time at 37 °C in the absence of metal at which the enzymatic activity reduces to 50% bMichaelis−Menten kinetic parameters at 37 °C (top row) and 20 °C (bottom row) for AsbF variants.

a

Figure 4. Thermal denaturation profiles of wild-type (blue trace) and Mut1 (red trace) in (a) the absence of metal, or the presence of (b) magnesium(II) or (c) manganese(II).

Figure 5. In vivo assessment of functional enzyme over a 5 h period for (a) Mut1 and (b) wtAsbF, and (c) SDS-PAGE analysis depicting accumulation of soluble Mut1 relative to wtAsbF.

AsbF has no detectable activity after a 1 h incubation at 50 °C in the absence of divalent metal cofactors.12,16 Following the 50 °C heat challenge on the FACS-enriched AsbF library, the crude lysate from two AsbF variants rapidly converted DHS into DHB in less than 15 min. Sequence analysis on the two constructs revealed three and six mutations in Mut1 and Mut2, respectively, and were introduced at the predicted locations on the surface of AsbF. Quite unexpectedly, both variants harbored the identical mutations, T61N, H135Y and H257P with three additional mutations present in Mut2, L75R, N123D and R244 K. A convergence in sequence at three locations was a pleasant surprise after a single round of flow sorting and heat challenge. Preliminary steady-state kinetic analysis revealed nearly identical catalytic efficiencies and similar thermal tolerance profiles. With no apparent beneficial contribution from the three additional point mutations (Mut2), Mut1 was therefore selected for steady-state characterization, thermal stability, and screened in vivo for biocatalytic potential toward DHB production in E. coli. We also confirmed the Mut1 secondary structure to be nearly identical to the wtAsbF solution structure as judged by circular dichroism (CD) spectroscopy (Supplementary Figure S2). Future experiments to determine the individual contribution for each point mutation on the Mut1 stability are being developed. However,

based upon crystal structure analysis, we speculate the H257P point mutation may result in tighter surface packing due to the increased hydrophobicity of proline to avoid the water solvation likely afforded by the charged histidine residue. Certainly, solving the Mut1 crystal structure concomitant with performing thermostability studies on the individual point mutations would provide the requisite insight. Steady-State Kinetics and Half-Life Determination of wtAsbF and Mut1. The comparative analysis of Mut1 and wtAsbF steady-state kinetics at both 37 and 20 °C revealed small differences in the respective Michaelis−Menten parameters (Table 1) and comparable activation energy for catalysis (Supplementary Figure S3). Further, the data revealed an approximately 30-fold lower apparent KM for the substrate, DHS, when 10 μM Mn(II) was employed in the activity assay when compared to 5 and 7.5 mM Mg(II) as previously reported by us and others, respectively.12,16 To resolve this difference, an activity assay containing both 7.5 mM Mg(II) and 10 μM Mn(II) were incubated with AsbF and varying DHS concentrations. The apparent KMDHS containing the divalent cation mixture was in good agreement with the apparent KMDHS in the presence of 10 μM Mn(II) alone (∼5 μM). The apparent KMDHS observed in the presence of 7.5 mM Mg(II) (∼160 μM) approximated the previously published values for AsbF.12,16 E

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wtAsbF on a per cell basis. We speculate Mut1 may be more protease resistant than wtAsbF under the defined growth conditions but this hypothesis will require more extensive experimental design to correctly assess. Commodity Chemical Production. The functionality of Mut1 in heterologous commodity chemical production relative to the wild-type enzyme was ascertained using two methods. First, overexpression plasmids (kanr) harboring Mut1 and the wild-type gene were introduced into E. coli BL21 (DE3) cells, overnight enzyme expression (14 h, 30 °C), and resuspension of the wet cell paste in M9 salts supplemented with glucose, the rate of DHB synthesis and secretion into the medium was quantified by absorbance at 290 nm relative to a control E. coli BL21 strain. The time course was carried out at 6, 20, 30, and 37 °C, respectively, for both DHB production strains. Second, both aforementioned plasmids were also introduced into E. coli BL21 (DE3) strains transformed with ampicillin-resistant overexpression vectors encoding both subunits of the dihydroxybenzoate dioxygenase enzyme from Pseudomonas putida (PcaG/H).28 Strains containing AsbF and PcaG/H synthesize β-carboxy-cis,cis-muconic acid (β-CMA), which is a novel polymer precursor.29 Production of β-CMA by a heterologous host is similarly quantified in the medium via UV-spectroscopy (255 nm) relative to a control E. coli BL21 strain.30 Similar assays using cells harboring a plasmid carrying the inactive form of AsbF, H144A, also verified no appreciable absorbance attributable to background aromatic amino acid side products were observed. While we did not observe the optimal temperature for DHB production in Mut1/wtAsbF transformants to change appreciably throughout a 24-h incubation, the DHB titers in Mut1 expression strains during this time frame were elevated relative to wtAsbF strains at each observed condition. The exception was the 6 °C incubation where DHB production was approximately constant (∼25 μM DHB) throughout the entire assay. However, the 20, 30, and 37 °C incubations with each Mut1 expression strain produced on average 80−120% more DHB than the wtAsbF expressing strains at the 6, 18, and 24 h time points (Figure 6), respectively. Production at earlier time points (e.g., 2 h), however, revealed an approximately equal DHB titer at all temperatures examined. In addition, the 2 h time point data approximates the in vivo enzyme concentrations and activities as determined immediately after an overnight induction at 20 °C (Figure S5). Therefore, it would appear the increase in DHB titers (except for the 6 °C incubation) for the Mut1 relative to the wtAsbF harboring strains occur after prolonged incubation at 20, 30, or 37 °C. We speculate that this observation is likely due to a loss of in vivo wtAsbF activity over time that is more than that incurred for Mut1 under comparable conditions. Enzymes lose activity over time upon exposure to temperatures that approximate or exceed their optimal temperature for catalysis, however a similar rate of decline of activity for thermostable enzymes exposed to temperatures significantly below their optimum is generally not observed.31 In the in vivo experiments, 20 °C (and to a lesser extent, 30 °C) is closer to the optimal functional temperature for the wtAsbF and much further away from the Mut1 optimal operating temperature (see Table 1). To accurately correlate in vivo DHB production with temperature for both strains, a full profile of in vitro half-life and enzyme kinetic studies at multiple operating temperatures will be required. This information will be communicated at a later date.

The divalent cation [Mg(II) or Mn(II)] did not markedly influence Vmax (refer Supplementary Figure S4). Overall, the data corroborate the reported crystal structure with bound Mn(II) and DHB as the likely in vivo Mn(II)-AsbF complex.16 To ascertain the stabilizing effect of T61N, H135Y and H257P in Mut1, the half-life determination at 37 °C (t1/237) was assessed in the absence or presence of either divalent metal cation, Mg(II) or Mn(II). The data revealed that wtAsbF was quite unstable in the absence of divalent cation but the three surface mutations in Mut1 resulted in a greater than 10-fold increase in enzyme half-life at 37 °C (Figure 4a). The presence of saturating concentrations of Mg(II) only resulted in a subtle effect on thermostability for both wtAsbF and Mut1 (Figure 4b) whereas the presence of Mn(II) resulted in substantial retention of DHSase activity in both variants after a 5 h incubation at 37 °C. Most notably, Mut1 retained greater than 80% of the original activity during this time frame. In Vivo DHSase Overexpression Analysis: wtAsbF versus Mut1. A comparative overexpression and functional enzyme assay between the wtAsbF and Mut1 was assessed in order to determine how in vitro thermostability translates into in vivo enzyme expression and activity. As such, each E. coli strain (wtAsbF or Mut1) was grown to mid log phase in rich media and enzyme expression induced with IPTG at 37 °C. Aliquots of each strain were harvested and gently lysed under nondenaturing conditions over 5 h. Soluble protein fraction content was assessed by SDS-PAGE and functional enzyme, normalized for dry cell weight, was determined using the aforementioned DHSase activity assay. Interestingly, during the first 5 h postinduction, the cell line harboring Mut1 appeared to exhibit a nearly 60-fold increase in functional enzyme relative to the line harboring the wild-type enzyme. (Figures 5a and 5b for wtAsbF and Mut1, respectively). The observed activity for each enzyme appeared to qualitatively correlate with total enzyme concentration (Figure 5c) as monitored by denaturing SDSPAGE. Further, the data revealed a jump in functional wtAsbF during the first hour postenzyme induction followed by an approximate 2-fold decrease in hour two. All points thereafter revealed wtAsbF activity to be relatively constant. During the time course postinduction, little visual evidence of wtAsbF expression on the denaturing gel was observed. In contrast, and under identical expression conditions (in parallel), functional Mut1 activity was observed to be nearly 60-fold greater than the wild-type counterpart within 2 h post-IPTG induction. Therefore, given similar in vitro kinetic parameters for both wtAsbF and Mut1, we believe the apparent 60-fold increase in functional Mut1 activity is likely due to an approximate 60-fold higher intracellular enzyme concentration relative to the wildtype enzyme. Clearly, the data revealed Mut1 is more thermotolerant, with net retention of functional activity, at 37 °C, than wtAsbF under the employed growth conditions and also correlates with our in vitro kinetic data. However, we observed a discrepancy with the half-life determination, in vitro, in both the presence and absence of divalent metal and the apparent difference in functional enzyme from the in vivo studies. Specifically, the half-life determination for both the wtAsbF and Mut1 in the absence of metal revealed an approximate 10−12-fold difference at 37 °C. Furthermore, in the presence of Mn(II), the wtAsbF half-life was observed to be approximately 4-fold lower than Mut1. Although the intracellular divalent metal concentrations in the in vivo experiments were not directly measured, there would appear to be other factors leading to higher Mut1 concentrations relative to the F

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to the oppositely charged counterparts (for example, I → E), relatively small charged amino acids to larger charged amino acids (e.g., D → E or K → R) or neutral to charged (Q → E) were considered in the design process. The second structurebased method adopted the protein design server, ROSIE24 and the VIP-based protein design23 protocol. This software was utilized to identify voids and suggest mutations that may yield improved surface packing. Finally, a standalone protocol “point mutant scan” under the Rosetta macromolecular modeling suite35 was utilized to predict all stability enhancing mutations on DHSase. In order to remove any crystallographic defects due to crystal packing, nonideal backbone angles and atomic clashes, we preceded our design with an all-atom structure refinement of the protein.36 An optimized (or relaxed) structure was then used for the point mutation design via Rosetta. The aforementioned approach considers each position on the protein, simulates saturation mutagenesis and calculates the difference in all-atom score of the resulting mutants relative to the wild-type enzyme. A default filter of −1 REU (Rosetta Energy Units) was used to select stability enhancing mutations. During the course of the point mutation design, extensive rotamer sampling (included additional ±1 standard deviation from mean chi angle for all chi1, chi2, chi3 and chi4 torsional angles) was used. The rotamer substitution was carried out on a fixed protein backbone. The mutations identified from these three approaches were visually assured using PyMOL (Shrodinger, LLC) to be on surface of AsbF, away from the dimer interface, and outside the catalytic center of the enzyme. A total of 24 positions with a wild-type amino acid identity and one or more favorable mutations were examined to reconfirm the most favorable combination of mutations using a fixed backbone protein design approach in Rosetta. Library Construction and Screening. The asbF combinatorial library was constructed through gene splicing and overlap extension PCR (SOEing) methodologies.26 At specific positions in the oligonucleotide, degenerate codons were used to code for multiplicity, and where “noise” due to unwanted mutations was inevitable, more than one oligonucleotide was used, each carrying a single codon for the mutagenesis position of interest. A table of the degenerate primers as the corresponding gene map identifying the mutation sites can be found in the Supporting Information (SI, Table S1, Figure S1). After PCR amplification of the asbF library with the T7 promoter and T7 terminator primer pair, the library was digested with either NcoI/HindIII (for subsequent ligation into pET-28a(+); Novagen/EMD Millipore) or SacI/XhoI (for ligation into pRSF-1b; Novagen/EMD Millipore). Following ligation, double transformation with the library plasmids were performed along with the sensor-reporter plasmid, pGLO.pcaU.eRBS.27 Expression and Purification. For protein expression, the pRSF plasmids containing the selected mutants (Mut1 and Mut2) were transformed into BL21(DE3) (Agilent Technologies) and grown in Super Optimal Broth (SOB) supplemented with 50 μg/mL kanamycin at 37 °C with shaking at 350 rpm. When OD600 reached 0.4, the temperature was dropped to 18 °C and the cells were allowed to grow to OD600 = 0.7 at which point 0.5 mM IPTG was added. The cells were grown for 14 h at 18 °C and harvested by centrifugation at 4000g and 4 °C, yielding about 2 g wet cell paste per 100 mL culture. Protein purification was conducted as described previously, with modification. In summary, the wet cell paste

Figure 6. DHB production in E. coli expressing (A) wtASbF or (B) Mut1 at 37 °C (blue), 30 °C (red), 20 °C (green) or 6 °C (purple).

Inclusion of Mut1 into a de novo β-CMA biosynthetic pathway did not appear to offer the same advantages over wtAsbF as those realized in DHB production. The optimal temperature for production and the absolute quantities of βCMA produced were not significantly different between the two strains (Supplementary Figure S6). In this case, it is likely that the rate limiting step in β-CMA production occurs after DHB biosynthesis. P. putida PcaG/H is well-known to have properties which limit its utility in a biosynthetic scaffold, including substantial product inhibition28 and a sensitivity of the biocatalyst to oxygen.32 Our observations are consistent with PcaG/H acting as the likely rate limiting step in β-CMA formation as well, although little is known about export of this molecule from an E. coli heterologous host. It is worth noting that transcriptional regulators responsive to β-carboxymuconolactone (which forms readily from β-CMA both biosynthetically and nonenzymatically) are known,33 and thus an improvement to the thermostability/catalytic efficiency of PcaG/H to realize higher product titers may also be envisioned using an identical approach to that employed for the thermostabilization of AsbF reported here.

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METHODS Structure Based Protein Design for Stability. Three parallel approaches to identify mutations on DHSase were pursued. The structure of DHSase/AsbF from Bacillus anthracis (PDB code 3DX5)16 is available in the protein data bank.34 The sequence of DHSase from B. thuringiensis is 100% identical to the sequence for which a 3D structure was solved, thus making it suitable for structure based design. In the first approach, surface residues were identified where introduction of a charged amino acid may initiate favorable charge−charge interactions with proximal amino acids of opposite charge. For example, surface residues that included arginine, lysine, glutamate and/or aspartate were candidates to create new salt bridges. As such, proximal surface mutations that transformed nonpolar residues G

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debris were collected by centrifugation at 4750g for 20 min. Soluble cell lysate was distributed between three flat bottom 96well plates (10 μL per well) containing 190 μL of Buffer F (20 mM citrate, 20 mM phosphate, 20 mM borate at pH 8.6, 50 mM NaCl, 7.5 mM MnCl2, 100 μM β-ME, and 0.1 μg/mL BSA). Plates were individually challenged by incubation at 37, 46, or 50 °C for 1 h. An absorbance reading for each cell was taken and 100 μM DHS was added. The reaction was allowed to proceed at 37 °C for 30 min. Finally, a reading was taken again at 290 nm and wells which had a change in Abs290 greater than or equal to wild-type AsbF control challenged at 37 °C were recorded as active. For analysis of the library following the HTS, a modified assay was also conducted in addition to this analysis where the reaction was allowed to proceed at 37 °C for 1 h, with readings taken every 15 min. Steady-State Measurements. Data for Michaelis−Menten model was made as previously described with modification. Reaction volume was increased to 0.5 mL and Buffer F was used for the reaction, replacing the 7.5 mM MgCl2 used in the past work with 100 μM MnCl2. Measurements were made at 20 and 37 °C. Data were fit and KM and kcat were calculated using Prism 6.0 by GraphPad. The 1 μM DHS measurements were excluded from the Lineweaver−Burk plot because their relatively large reciprocal value drove the regression. Arrhenius plots were constructed using the same reaction conditions as the Michaelis−Menten data. Temperature was varied from 20 to 50 °C in 5° steps. DHS concentration was held constant at 300 μM to approximate saturating conditions. The data were inverted and fit using a linear regression in Prism 6.0 by GraphPad. Functional Protein In Vivo. Mut1 and wild type AsbF were transformed into BL21(DE3) and grown in SOB containing 50 μg/mL kanamycin at 37 °C. When the cultures reached an OD600 of 0.7, they were induced with 0.5 mM IPTG. Starting at the time of induction, t0, 1 mL aliquots were taken out of the cultures and OD600 were measured at each hour. The aliquots were lysed using 50 μL BugBuster (Novagen) as described for the MTS. Soluble cell lysate was then assayed as described in Steady-State Measurements using 100 μM DHS and 37 °C. The intracellular AsbF concentration was estimated using the known kinetic parameters calculated for the purified enzyme and the grams dry cell weight (gDCW) of E. coli estimated using published conversion factors from the observed OD600 at the specified time point. Long-Term Stability at 37 °C. Purified protein was diluted to a concentration of 0.06 ng/μL in TRIS, pH 7.5 with no additional metal, 7.5 mM MgCl2 or 10 μM MnCl2. The samples were then aliquoted into thin-walled PCR tubes with 30 μL each. These tubes were incubated at 37 °C for 5 h, taking readings every 30 min using 100 μM DHS and 37 °C. The data were normalized to the activity of the protein before heat treatment and fit to either a linear or exponential decay curve. The curve with the highest R2 is shown in the figures below. Heterologous Production of DHB and β-Carboxycis,cis-muconic Acid. For DHB production, E. coli BL21(DE3) cells transformed with plasmids containing the gene for wild-type AsbF or Mut1, respectively, were grown at 37 °C with shaking (200 rpm) in LB containing kanamycin (50 μg/mL). Overexpression of AsbF or Mut1 followed as previously described with minor modifications. Cells were grown to an OD600 of 0.4 at which time overexpression was induced with IPTG (0.5 mM), the temperature reduced to 30 °C, and growth continued for 14 h. Upon harvesting by centrifugation,

was lysed with 5 mL of BugBuster (Novagen) and 1000 units benzonase per gram cell paste for 20 min at room temperature before removing cellular debris by centrifugation at 4500 × g and 4 °C for 45 min. All purification steps following this were done on ice and using buffers at 4 °C. The soluble portion of the cell lysate was filtered through a 0.22 μm syringe filter and buffer exchanged into Buffer A (50 mM TRIS at pH 7.5, 10 mM β-ME and 20 mM imidazole) using a 10K MWCO centrifugal filter for three dilution and centrifugation cycles. The filtered soluble lysate was batch loaded onto Ni-NTA resin (0.25 mL resin per gram wet cell paste) equilibrated with Buffer A at 4 °C for 1 h with constant stirring. The resin protein mixture was packed, washed and eluted with 3 column volumes Buffer B (50 mM TRIS at pH 7.5, 10 mM β-ME, 50 mM imidazole) followed by 3 column volumes Buffer C (50 mM TRIS at pH 7.5, 10 mM β-ME, 200 mM imidazole). Fractions which were greater than 95% AsbF, assessed by SDS-PAGE, were pooled and buffer exchanged into Buffer D (50 mM TRIS at pH 7.5, 10 mM β-ME, 10% Glycerol). The protein was diluted to 0.6 mg/mL with Buffer D, aliquoted, flash frozen with liquid nitrogen and stored at −80 °C until needed. High-Throughput Screening (HTS). The mixture of pRSF-1b plasmids containing the diversified asbf was transformed into BL21-Gold(DE3) cells (Agilent Technologies) containing the plasmid pGLO.pcaU.eRBS, which is our optimized pcaU-gf p sensor-reporter system.9 Transformed cells were plated and allowed to grow for 8 h before the entire population (∼10 000 colonies) was resuspended in liquid media. The cell suspension was then diluted 1:1000 and grown in SOB with 50 μg/mL kanamycin and 100 μg/mL carbenicillin in a 6-well plate at 37 °C. Once the cultures reached an OD600 of 0.7, the temperature was reduced to 20 °C and 0.5 mM IPTG was added followed by growth for 14 h. As a control BL21-Gold(DE3) cells containing pcaU-gf p plasmid and pRSF1b empty vector or pRSF-1b with wtasbF gene were also grown under similar conditions. The samples were analyzed for GFP expression on a BD FACSAria II using a 488 nm blue laser for excitation and 530 nm emission filter. The top 50% of the peak showing increased fluorescence were sorted and plated on LB containing 50 μg/ mL kanamycin and 100 μg/mL carbenicillin. Enrichment was assessed using the MTS described below. Medium-Throughput Screening (MTS). Individual asbF variants were transferred into a 96 deep-well plate containing Luria−Bertani (LB) growth media, enzyme expression induced with IPTG and incubated overnight at 20 °C. The following day, the cells were harvested by centrifugation, growth media removed and pellet gently lysed with BugBuster reagent. An aliquot of the crude lysate was transferred to a new, UVtransparent 96-well plate containing AsbF activity assay buffer, plate sealed, and incubated at 37, 46, or 50 °C, respectively, for 1 h. Following the heat challenge, saturating DHS (300 μM) was added to each well, incubated for 30 min at 37 °C, and DHB detected by UV−vis spectroscopy at 290 nm. MTS was accomplished using a 96-well plate format. Individual colonies were selected from a plate and grown in a deepwell, round-bottom 96-well plate with 1 mL LB containing 100 μM IPTG and the appropriate antibiotics. Cultures were grown for 14 h at 20 °C, at which point the plate was replicated and cells were harvested at room temperature (∼22 °C) using a swinging bucket rotor at 4000g for 15 min. The cell pellet was resuspended in 50 μL per well BugBuster (Novagen) and incubated on a rocker at room temperature for 20 min. Cellular H

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ACKNOWLEDGMENTS D.T.F. was supported in this work by the Los Alamos National Laboratory under the U.S. Department of Energy, Laboratory Directed Research and Development grant [LDRD ER20100182ER] and Office of Energy Efficiency & Renewable Energy, Bioenergy Technologies Office Annual Operating Plan. L.H. was supported by the DOE Summer Undergraduate Laboratory Internships (SULI) program. R.K.J and C.E.M.S were supported by the Defense Threat Reduction Agency [CBCALL12-LS-6-0622]. Computational work was supported by the LANL Institutional Computing grant [W13_SynBio]. A.T.K. was supported in this work by the NAU College of Forestry, Natural Science and Engineering Dean’s office through faculty startup funding, and G.C. was supported by grant NIH- 5R25GM056931, Initiative to Maximize Student Development in the Biomedical Sciences. We thank Drs. Ron Jacak and Brian Kuhlman (UNC Chapel Hill) for Rosetta point mutation scan (p_mut scan) protocol and Dr. Dung Vu (LANL) for assistance with the circular dichroism experiments.

the spent media was decanted and the wet cell paste (approximately 0.5 g) was washed with M9 medium supplemented with glucose (4 g/L) to remove any aromatic contaminants in the LB. Cells were washed (2×) by resuspension in a small volume of medium (10 mL) by vortexing, followed by harvesting by centrifugation and decanting of the spent medium. After the washing, the cell paste was resuspended into M9 with glucose to a final OD600 of 0.5. Further incubation of the suspended cultures was conducted for 24 h at defined temperatures (6, 20, 30, and 37 °C, respectively). At defined time points (2, 6, 18, and 24 h), aliquots (1 mL) were removed, the cells pelleted by centrifugation (1 min, 12 000 rpm), and production of DHB were quantified via absorbance at 290 nm. All DHB production experiments were performed in triplicate. Production of β-carboxy-cis,cis-muconic acid was performed in a similar manner. In this case, dual transformants were constructed by introducing plasmids containing either wild-type AsbF or Mut1 (kanamycin resistant) into an E. coli BL21(DE3) host maintaining a plasmid containing the P. putida pcaG and pcaH genes, respectively, as a bicistronic construct (ampicillin resistant). Growth of strains containing AsbF (strain KFβ1) or Mut1 (strain GCβ1) were conducted in LB containing kanamycin (50 μg/mL) and ampicillin (100 μg/mL). Overexpression and product analysis proceeded as previously described, with the exception that β-carboxy-cis,cis-muconic acid concentration was quantified at 255 nm.13 All production experiments were performed in triplicate. Production of DHB or β-CMA, respectively, was quantified relative to a E. coli BL21(DE3) control strain. An authentic standard of DHB was purchased from Sigma-Aldrich and used without further purification. Authentic standards of β-CMA were synthesized by oxidation of vanillin using the method of Husband et al.37 Circular Dichroism. CD data were obtained using a Jasco J715 spectropolarimeter fitted with a temperature controller. Protein was buffer exchanged into 20 mM TRIS, 0.5 mM TCEP-HCl and 10 μM MnCl2 using 10K MWCO dialysis tubing. Buffer exchanged samples were loaded into a 1 mm cuvette equilibrated at the starting temperature. Far-UV CD scans (260−196 nm) were performed at 50 mdeg sensitivity and 5 nm/min scanning speed. Scans were collected in triplicates and averaged. All ellipticity data were corrected using a buffer blank.

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ABBREVIATIONS ADA, Adipic acid; AsbF/wtAsbF, B. thuringiensis 3-dehydroshikimate dehydratase; β-CMA, β-carboxy-cis,cis-muconic acid; ccMA, cis,cis-muconic acid; DHB, 3,4-dihydroxybenzoic acid; DHS, 3-dehydroshikimic acid; DHSase, 3-dehydroshikimate dehydratase; FACS, Fluorescence-activated cell sorting; GFP, Green fluorescent protein; GCβ1, Heterologous E. coli host expressing Mut1 and PcaG/H; IPTG, β-D-1-thiogalactopyranoside; KFβ1, Heterologous E. coli host expressing wtAsbF and PcaG/H; PcaG/H, Subunits of dihydroxybenzoate dioxygenase; PCR, Polymerase chain reaction; Mut1, Triple mutant construct of AsbF; REU, Rosetta energy units

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REFERENCES

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acssynbio.6b00159. Supporting Tables, Figures and Graphs (PDF)

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Research Article

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. *E-mail: [email protected]. Author Contributions #

L.B.H. and R.K.J. contributed equally to this work.

Notes

The authors declare no competing financial interest. I

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