Discovery and Engineering of Pathways for Production of α-Branched Organic Acids

Article abstract of DOI:10.1021/jacs.7b07400

Cell-based synthesis offers many opportunities for preparing small molecules from simple renewable carbon sources by telescoping multiple reactions into a single fermentation step. One challenge in this area is the development of enzymatic carbon-carbon bond forming cycles that enable a modular disconnection of a target structure into cellular building blocks. In this regard, synthetic pathways based on thiolase enzymes to catalyze the initial carbon-carbon bond forming step between acyl coenzyme A (CoA) substrates offer a versatile route for biological synthesis, but the substrate diversity of such pathways is currently limited. In this report, we describe the identification and biochemical characterization of a thiolase-ketoreductase pair involved in production of branched acids in the roundworm, Ascaris suum, that demonstrates selectivity for forming products with an α-methyl branch using a propionyl-CoA extender unit. Engineering synthetic pathways for production of α-methyl acids in Escherichia coli using these enzymes allows the construction of microbial strains that produce either chiral 2-methyl-3-hydroxy acids (1.1 ± 0.2 g L^-1) or branched enoic acids (1.12 ± 0.06 g L^-1) in the presence of a dehydratase at 44% and 87% yield of fed propionate, respectively. In vitro characterization along with in vivo analysis indicates that the ketoreductase is the key driver for selectivity, forming predominantly α-branched products even when paired with a thiolase that highly prefers unbranched linear products. Our results expand the utility of thiolase-based pathways and provide biosynthetic access to α-branched compounds as precursors for polymers and other chemicals.

Full text of DOI:10.1021/jacs.7b07400

Article
pubs.acs.org/JACS
Discovery and Engineering of Pathways for Production of
α‑Branched Organic Acids
Michael R. Blaisse,† Hongjun Dong,† Beverly Fu,† and Michelle C. Y. Chang*,†,‡
†Department of Chemistry, University of California, Berkeley, Berkeley, California 94720-1460, United States
‡Department of Molecular and Cell Biology, University of California, Berkeley, Berkeley, California 94720-1460, United States
S
* Supporting Information
ABSTRACT: Cell-based synthesis offers many opportunities
for preparing small molecules from simple renewable carbon
sources by telescoping multiple reactions into a single
fermentation step. One challenge in this area is the
development of enzymatic carbon−carbon bond forming
cycles that enable a modular disconnection of a target
structure into cellular building blocks. In this regard, synthetic
pathways based on thiolase enzymes to catalyze the initial
carbon−carbon bond forming step between acyl coenzyme A
(CoA) substrates offer a versatile route for biological synthesis, but the substrate diversity of such pathways is currently limited.
In this report, we describe the identification and biochemical characterization of a thiolase-ketoreductase pair involved in
production of branched acids in the roundworm, Ascaris suum, that demonstrates selectivity for forming products with an α-
methyl branch using a propionyl-CoA extender unit. Engineering synthetic pathways for production of α-methyl acids in
Escherichia coli using these enzymes allows the construction of microbial strains that produce either chiral 2-methyl-3-hydroxy
acids (1.1 0.2 g L−1) or branched enoic acids (1.12 0.06 g L−1) in the presence of a dehydratase at 44% and 87% yield of fed
propionate, respectively. In vitro characterization along with in vivo analysis indicates that the ketoreductase is the key driver for
selectivity, forming predominantly α-branched products even when paired with a thiolase that highly prefers unbranched linear
products. Our results expand the utility of thiolase-based pathways and provide biosynthetic access to α-branched compounds as
precursors for polymers and other chemicals.
metabolic pathways. More recently, the promiscuity of thiolases
has been used to expand the scope of ω- and α-
functionalization of target compounds.11−13 Notably, the
incorporation of propionyl-CoA in a retro-degradative pathway
has been demonstrated to produce the α-substituted enoic acid,
tiglic acid.11 However, an efficient pathway for 2-methyl-3-
hydroxy acid production has not yet been reported despite the
important role that α-substituted hydroxy acids could serve as
new monomers for the production of biodegradable polymers.
Polyesters derived from 3-hydroxy acids in the poly-
(hydroxybutyrate) (PHB) family suffer from problems of
processability and instability to thermal decomposition.
Incorporation of an α-branched monomer, 2-methyl-3-hydrox-
ybutyrate, into PHB copolymers results in decreased melting
enthalpy without significantly lowering melting temperature
(Tm) and more facile crystallization related to the decreased
cold crystallization temperature (Tcc).14 These effects were not
observed in the other PHB-based copolymers tested.
Furthermore, a major pathway for thermal degradation of
PHB-type polymers proceeds through α-deprotonation and
elimination, which is reduced by α-functionalization.15
INTRODUCTION
■
Living systems offer unique advantages as multistep catalysts for
chemical synthesis. Indeed, cellular chemistry has evolved to
take simple carbon sources, such as glucose and carbon dioxide,
as universal building blocks to create an enormous array of
metabolites and natural products, enabling the industrial-scale
production of a broad range of small molecule targets in a
single fermentation step from renewable carbon sources.1−7
Surprisingly, cell metabolism depends mainly on just two
related carbon−carbon bond forming reactions, the aldol and
Claisen condensations. The Claisen condensation catalyzed by
thiolase enzymes is particularly versatile, allowing the coupling
of two acyl coenzyme A (CoA) thioesters (Figure 1).8,9 This
step serves as an initiation point into a reaction cycle of C−C
bond formation, followed by ketoreduction, dehydration, and
enoyl reduction, which can be used in principle to disconnect
any small molecule target.
Despite the large number of members in the thiolase
superfamily (>50 000), a major challenge in their utilization for
this purpose is that the predominant substrate selectivity
characterized has been the condensation of simple acetyl-CoA
monomers (R = H).10 Broader substrate selectivity has been
identified based on the participation of thiolases in diverse
metabolic pathways, but the relatively small number of
characterized thiolases still limits the construction of synthetic
Received: July 16, 2017
© XXXX American Chemical Society
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Figure 1. Thiolase-based reaction cycle for assembly of small molecule targets. Both decarboxylative and nondecarboxylative Claisen condensations
are involved in the biosynthesis of a broad range of small molecule targets, including fatty acids, isoprenoids, and polyketides. However, the
predominant substrate selectivity of these enzymes involves utilizing the unfunctionalized monomers (R = H), acetyl-CoA and malonyl-CoA (left).
Utilizing the nondecarboxylative Claisen condensation catalyzed by thiolases, a reaction cycle of condensation followed by ketoreduction,
dehydration, and reduction could be used in principle to disconnect any small molecule as long as the R groups of interest are accommodated
(right).
Figure 2. Characterization of thiolases from A. suum. (A) Activity screen of Acat1−5 thiolases and a canonical acetyl-CoA selective PhaA from
C. necator. Activity assayed by CoA release in the presence of either 1 mM acetyl- or propionyl-CoA using DTNB. Data are mean s.e. (n = 3). (B)
Structures of 3-oxoacyl-CoA substrates. (C) Steady-state kinetic characterization of Acat2,3,5 thiolases in the thiolysis or degradative direction
monitoring formation of the thioester bond in the presence of 200 μM CoA. In cases where saturation was not reached, the kcat/KM parameter was fit
directly and a lower bound is provided for individual kcat and KM values. Data are mean s.e. (n = 3).
In this context, we present the discovery and characterization
of a new thiolase and ketoreductase pair from the roundworm,
Ascaris suum that is selective for biosynthesis of α-branched
acids. We also report the use of these enzymes to engineer
pathways for the production of α-methyl branched 3-hydroxy
and enoic acids in Escherichia coli. A. suum has been reported to
accumulate short branched organic acids as part of an anaerobic
respiration pathway, providing an unusual example of a
biosynthetic thiolase-based pathway that condenses both
acetate and propionate units.16,17 We have identified thiolase,
ketoreductase, and dehydratase candidates, finding two
thiolases that prefer propionyl-CoA as well as a ketoreductase
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Figure 3. Characterization of the AsHadh ketoreductase. (A) Steady-state kinetic characterization of AsHadh reduction of various 3-oxoacyl-CoA
substrates as well as the oxidation of 3S- and 3R-hydroxybutyryl-CoA produced in situ. Data are mean s.e. (n = 3). (B) Assay for examining the
stereochemical preference of AsHadh.
whose preference for an α-methyl substituent serves as an
important selectivity filter. We have used these enzymes to
produce α-methyl acids in E. coli at 87% and 44% conversion of
propionate for the enoic and 3-hydroxy acids, respectively, to
provide a useful route to chemical precursors and chiral
building blocks.18
thiolytic directions. Its function remains unknown, but a Ser
residue is found in place of the essential Cys that is conserved
in all other members of the thiolase superfamily26 (Supporting
Information Figure S1). These enzymes were also screened
against the four-carbon butyryl-CoA, but no activity was
detected with the longer substrate with any of the Acat
thiolases.
RESULTS AND DISCUSSION
With this simple screen of Acat1−5 activity in hand, we
decided to carry out a more detailed characterization of the
substrate selectivity of Acat2 and 3. To carry out this
experiment, we chemically synthesized a set of 3-oxoacyl-CoA
substrates to examine thiolysis since the CoA release assay is
not adequate for characterizing mixed substrate condensation in
the synthetic direction (Figure 2B, Supporting Information
Figures S4−S11). The catalytic efficiencies of Acat2 and Acat3
with all substrates formed from combinations of 2- or 3-carbon
units were of similar magnitude, but individual kcat and KM
values were both higher for α-methyl substrates compared to
their linear counterpart (Figure 2C). This behavior is similar to
that observed for human T2 thiolase, although the observed KM
values for Acat 2 and 3 were significantly higher than that
observed in T2.21 These higher KM values may reflect a
preference to release 3-oxoacyl-CoA product since biosynthesis
is expected to be the physiologically relevant direction in
A. suum mitochondria. Some substrates showed substrate
inhibition behavior at higher concentrations. Initial rate data for
these substrates fit poorly to the standard Michaelis−Menten
equation but fit well to the modified form (Supporting
Information Figure S12). Substrate inhibition was most
prominent with the six-carbon chain substrates, 3-oxohex-
anoyl-CoA and 2-methyl-3-oxohexanoyl-CoA, with which
Acat2, 3, and 5 also demonstrated lower catalytic efficiencies.
Acat5 was characterized to compare a canonical thiolase from
the same host organism. In contrast to Acat2 and Acat3, Acat5
had high catalytic efficiency with linear 3-oxoacyl-CoA
substrates but a catalytic defect of 100-fold in kcat/KM upon
introduction of an α-methyl substituent to 3-oxobutyryl-CoA,
mostly due to a large increase in KM (Figure 2C). Extending the
main chain by one carbon (2-methyl-3-oxopentanoyl-CoA)
lowered the relative kcat/KM by an additional 30-fold. Notably,
when comparing the preference between the linear and
corresponding branched substrate, Acat5 displayed a preference
of two to three orders of magnitude for the linear analogue.
This observation is quite different from Acat2 and 3, which
displayed only a 1.5 to 4.8-fold deviation between the kcat/KM
values for the linear and branched analogues.
■
Identification and Characterization of Thiolase En-
zymes from A. suum. We first set out to identify gene
candidates responsible for the branched acid production
reported in A. suum from the available Transcriptome Shotgun
Assembly database in GenBank.19,20 On the basis of the report
that these short-chain branched acids were produced in the
mitochondria of A. suum muscle tissue, we used the human
mitochondrial thiolase T2 responsible for branched amino acid
degradation (UniProtKB P24752)21 as a search query and
obtained seven thiolase candidates, six of which had sequence
identity >50% (Supporting Information Figure S1). Five of
these thiolases (Acat1−5) were predicted to be targeted to
mitochondria22,23 and were selected for further biochemical
characterization. The five synthetic genes were then assembled
using a truncated sequence for the predicted mature polypetide
using standard codon optimization for heterologous expression
in E. coli (Supporting Information Table S1). The Acat1−5
proteins were purified by Ni affinity chromatography and then
cleaved with Tobacco Etch Virus (TEV) protease to remove
the N-terminal His6-tag (Supporting Information Table S1,
Figure S2). All proteins were further purified by size-exclusion
chromatography and appeared to elute as active tetramers
(Supporting Information Figure S3), as is typically found for
biosynthetic thiolases in solution.21
Purified Acat1−5 thiolases were then screened for activity
with the two- and three-carbon acyl-coenzyme A (CoA)
substrates, acetyl-CoA and propionyl-CoA. Activity was
monitored by CoA release using Ellman’s reagent (Dithio-
bis(2-nitrobenzoic acid), DTNB) and compared to the
canonical thiolase (PhaA) from Cupriavidus necator, which
selectively assembles two acetyl-CoA monomers.24,25 From this
initial screen, Acat5 resembles the canonical PhaA thiolase
involved in poly(hydroxyl)alkanoate (PHA) synthesis in
bacteria, with high activity for acetyl-CoA and no detectable
activity for propionyl-CoA (Figure 2A). Interestingly, Acat2
and Acat3 showed higher activity with respect to propionyl-
CoA compared to acetyl-CoA. Acat1 also displayed activity for
propionyl-CoA condensation, but to a lesser degree. Acat4
demonstrated no activity as a thiolase in the synthetic or
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Identification and Characterization of Ketoreductase
AsHadh. In addition to the thiolase, we also sought to identify
a ketoreductase capable of accepting the α-branched substrates.
Searching the RNASeq database with human mitochondrial
short chain hydroxyacyl-CoA dehydrogenase HADH227 as
query, we identified just one ketoreductase candidate in
A. suum, AsHadh (ADY43626.1). His10-AsHadh was expressed
in E. coli from a synthetic gene and purified (Supporting
Information Table S1 and Figure S13). Steady-state kinetic
characterization of the enzyme revealed a ∼20-fold preference
for α-methyl substrates, 2-methyl-3-oxobutyryl-CoA and 2-
methyl-3-oxopentanoyl-CoA, compared to the linear substrate
3-oxobutyryl-CoA (Figure 3A). This selectivity arose mostly
from significantly lower KM values for the α-methyl substrates.
Interestingly, the α-ethyl substituted substrate, 2-ethyl-3-
oxohexanoyl-CoA, is also accepted by this enzyme with similar
efficiency as the linear substrate. Potent substrate inhibition was
observed for AsHadh (Supporting Information Figure S14),
which is common in NAD(P)H oxidoreductases28 based on
their ordered-binding reaction pathway.29
We next investigated the stereoselectivity of AsHadh with
respect to the 3-position in a coupled assay testing its ability to
oxidize the two enantiomers of 3-hydroxybutyryl-CoA. Starting
with crotonyl-CoA, either 3S-hydroxybutyryl-CoA or 3R-
hydroxybutyryl-CoA can be formed by stereospecific hydration
with the crotonase or PhaJ hydratases, respectively (Figure
3B).4 We then monitored the ability of AsHadh to oxidize each
enantiomer of in situ-generated 3-hydroxybutyryl-CoA. In these
experiments, we found that AsHadh had 40-fold higher catalytic
efficiency when coupled to crotonase compared to PhaJ,
corresponding to a preference for 3S- over 3R-hydroxybutyryl-
CoA (Figure 3A). Although the stereochemical preference of
AsHadh at the α-branch site remains cryptic, we would predict
a (2S,3S) configuration based on the behavior of murine
HADH230 and the expectation that the subsequent dehydration
proceeds by a syn-elimination mechanism.31
Production of Branched Enoic Acids in E. coli. We next
set out to use A. suum enzymes to produce branched organic
acids through construction of a synthetic pathway in E. coli
(Figure 4A, Supporting Information Figure S15). To produce
enoic acids, we identified a gene from A. suum encoding
dehydratase AsEch based on its similarity to the human
mitochondrial hydratase ECHS132 and predicted mitochondrial
localization. Biochemical characterization of His6-AsEch
purified from heterologous expression in E. coli showed that
it prefers linear substrates but is permissive of the α-methyl
substrate (Supporting Information Figures S16−S17A,B).
Plasmids containing a thiolase (Acat2, 3, or 5), AsHadh, and
AsEch were then constructed from the pTrc99a base vector. A
second set of pTrc33-based plasmids was also constructed with
a panel of four thioesterases from E. coli (TesA, TesB, YciA, or
YdiI)33 to release free organic acids from the corresponding
acyl-CoA intermediates. Dual-plasmid systems were trans-
formed into E. coli BAP1, a modified BL21(de3) strain
engineered to increase the intracellular concentration of
propionyl-CoA.34 E. coli strains 1−16 were cultured in LB
media supplemented with 2 g L−1 propionate and tested for
their ability to produce different enoic acids (Figure 4B).
Product titer was observed to be highly dependent on the
thioesterase, with YciA and YdiI both being competent for
enoic acid production. The highest titers were obtained using
YdiI, which is in agreement with previous studies.33 Strain 12
containing Acat3, AsHadh, and AsEch in combination with YdiI
Figure 4. Production of enoic acids in E. coli BAP1 strains. (A) A two-
plasmid system was constructed for production of enoic acids. The
first plasmid consists of an operon containing a thiolase (Acat2, 3, or 5;
A2, 3, or 5), a ketoreductase (AsHadh), and a dehydratase (AsEch)
driven by a single promoter. The second plasmid contains different
thioesterases (TesA, TesB, YciA, or YdiI). (B) Production of enoic
acids in select strains. E. coli BAP1 were cotransformed with both
plasmids. Strains were cultured for 2 d at 30 °C in LB broth containing
1% (w/v) glucose and 0.2% (w/v) sodium propionate before collecting
samples for analysis. Data are mean s.d. of biological replicates (n =
3). Data are not displayed for Strains 1−4 (*) as no product was
detected.
produced a total of 1120
60 mg L−1 branched enoic acid
products, tiglic acid and 2-methylpentenoic acid, resulting in
the incorporation of 87% of supplemented propionate into
these products. The dominant product was 2-methylpentenoic
acid resulting from the condensation of 2 equiv of propionyl-
CoA.
Interestingly, branched enoic acids were produced almost
exclusively in all strains, including those containing Acat5 (15
and 16), a thiolase that had poor activity with branched
substrates in vitro. This product distribution may reflect the
importance of the preference of the AsHadh ketoreductase for
the α-methyl branch in driving the pathway equilibrium
forward given the reversibility of the thiolase reaction. Another
factor that may affect enoic acid product distribution, however,
is the thermodynamic difference in dehydration of linear or
branched hydroxyacyl-CoA substrates. Using component
contribution methods to estimate the pathway thermodynamics
with eQuilibrator,35 we noted that pathway energetics diverged
greatly for the dehydration of linear and branched inter-
mediates (Supporting Information Figure S17C,D). As an
experimental measure, we then determined Keq values for the
hydration of the linear crotonyl-CoA and corresponding
branched tigloyl-CoA to be 4.68
0.06 and 0.164
0.001,
respectively. The unfavorable hydration of α-branched
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substrates and their corresponding stability in the enoic form is
likely related to the chemical stability of the trisubstituted
alkene compared to a disubstituted alkene and may also
contribute to the observed product distribution.
lower than for enoic acids. We thus set out to optimize flux
through branched hydroxy acid synthesis by modulating
pathway enzyme expression levels with different gene
promoters and ribosome-binding site (RBS) sequences.
Improvements in titers were achieved by using a TesB
expression construct with an alternative RBS (pTrc33-TesB2,
strains 29 and 30). Although pTrc33-TesB2 was predicted to
have ∼11-fold higher expression level (110 K) than the initial
RBS (10 K),36 lower level of soluble TesB was observed
(Supporting Information Figure S19). Titration of upstream
enzymes Acat3 and AsHadh (Supporting Information Figure
S20) gave only slight improvement in selective production of α-
methyl products (strain 38).
A titration of Acat3 in conjunction with CnHadh
(WP_011614598), a homologue of AsHadh (52% sequence
identity) from C. necator, showed these strains were capable of
producing higher titers of branched hydroxy acids (up to 140
20 mg L−1, strain 46). However, this improved productivity was
accompanied by a loss of selectivity with an additional 500
100 mg L−1 of 3-hydroxybutyrate and 3-hydroxypentanoate
products being formed. In vitro characterization of CnHadh
showed that it did indeed accept α-methyl substituted
substrates with high catalytic efficiency, although with 5-fold
lower preference for the branched substrate compared to
AsHadh (Supporting Information Figure S21). In vitro assays
for 3-hydroxy acid production comparing AsHadh and CnHadh
to two well-characterized ketoreductases PhaB and Hbd with
strong linear substrate preference reflect the product
distribution observed in vivo (Supporting Information Figure
S22). These data indicate that the ketoreductase is indeed a
strong determinant of the product distribution between
branched and linear 3-hydroxy acids.
Production of Branched Hydroxy Acids in E. coli.
While the production of branched enoic acids has been
previously engineered,11 robust pathways for the production of
their corresponding 3-hydroxy acids have yet to be reported. In
order to construct a pathway for the production of hydroxy
acids, the same panel of thiolases and thioesterases were
assembled with AsHadh in the absence of AsEch and tested in
E. coli BAP1 cultured in LB media supplemented with 2 g L−1
propionate (Figure 5, Supporting Information Figure S18,
strains 17−28). Even in strains containing the thioesterase
TesB, which has been shown to be effective for producing short
linear hydroxy acids in E. coli,12 productivity was considerably
The improved titers seen in strains 29 and 30 compared to
22 and 26 suggested a critical role for TesB in controlling
pathway flux, suggesting that overly high TesB levels can derail
successful hydroxy acid production (Supporting Information
Figure S19). To further explore the origin of pathway sensitivity
to TesB, we purified TesB and tested its hydrolytic activity
against various acyl-CoA substrates and found that TesB
accepted propionyl-CoA and 2-methyl-3S-hydroxybutyryl-CoA
with similar efficiency (Supporting Information Figure S23).
Taken together, these data are consistent with a model that the
pathway is limited by hydrolysis of the propionyl-CoA building
block by the thioesterase.
Given the importance of thioesterase selectivity in control-
ling pathway yield, we screened a panel of hydrolases that could
have improved performance with 3-hydroxyacyl thioesters.
These candidates included putative 3-hydroxyisobutyryl-
CoA37−39 and hydroxybenzoyl-CoA hydrolases,40−42 as well
as the thioesterase domain of the 6-deoxyethronolide
polyketide synthase (Supporting Information Figure S24).43
The most promising candidate of those tested was the 3-
hydroxyisobutyryl-CoA hydrolase from Bacillus cereus, BcBch.
The resulting strains 56 containing BcBch thioesterase
outperformed TesB-expressing strain 38 and allowed branched
hydroxy acid titers of440 70 mg L−1 in TB media (Figure 5).
In vitro characterization of purified BcBch showed low catalytic
efficiency on both propionyl-CoA and 2-methyl-3-oxobutyryl-
CoA (Supporting Information Figure S25), so we reasoned that
high BcBch expression could further improve hydroxy acid
production. Strain 57 expressing BcBch from a T7 promoter
Figure 5. Production of 3-hydroxy acids acids in E. coli BAP1 strains.
(A) A two-plasmid system was constructed for production of 3-
hydroxy acids. The first plasmid consists of an operon containing a
thiolase (Acat2, 3, or 5; A2, 3, or 5) and a ketoreductase (AsHadh or
CnHadh) driven by a single promoter. The second plasmid contains
different thioesterases (TesA, TesB, YciA, YdiI, or BcBch). pTrc33-
TesB and pTrc33-TesB2 contain different RBS sequences, with
reduced TesB expression from pTrc33-TesB2 leading to increased
product titer. Strain 58 also contains a propionyl-CoA transferase
(Pct) in an operon with the thioesterase. (B) Production of 3-hydroxy
acids in select strains. E. coli BAP1 were cotransformed with both
plasmids. Strains were cultured for 2 d at 30 °C in LB broth containing
1% (w/v) glucose or TB broth containing 2.5% (w/v) glucose. Media
was supplemented with 0.2% (w/v) sodium propionate. Data are mean
s.d. of biological replicates (n = 3). Data are not displayed for Strains
17−25 (*) as no product was detected.
did indeed achieve a titer of 600
although SDS-PAGE did not indicate an obvious difference in
100 mg L−1 (Figure 5),
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soluble BcBch expression compared to strain 56 (Supporting
Information Figure S26). Unconsumed propionate remaining
in spent media could account for nearly all of the propionate
not incorporated into product for strain 57, resulting in a 97%
yield of consumed propionate. Finally, the additional over-
expression of propionyl-CoA transferase Pct from Clostridium
It is also important to note that the thiolase selectivity does
appear to greatly affect substitution at the ω-position, as the
pathways with Acat2 and Acat3 preferentially use propionyl-
CoA as a starter to produce the 6-carbon product, 2-methyl-
pentenoic acid, rather than the 5-carbon product, tiglic acid, as
reported for the FadA thiolase.11 As previously observed,
thioesterase selectivity plays an important role in pathway yield
as nonspecific hydrolysis of the precursor acyl-CoAs limits
carbon flux into the pathway.33 In this study, we find that enoic
acids are generated at high yield due to the high selectivity of
the thioesterase for the enoyl-CoA functional group. In
comparison, entry into the 3-hydroxy acids appears to be
more limited, which we attribute to the hydrolysis of the
propionyl-CoA starter but can be improved by addition of a
propionyl-CoA transferase. This large fraction of unconsumed
propionate lowers yields of the 3-hydroxy acids to 44% of the
total propionate added despite being produced with high
selectivity (>90% of consumed propionate). Taken together,
these studies provide insight into the use of thiolase-based
pathways for developing carbon−carbon bond forming cycles
using enzymes. In addition, the enzymes identified in this study
can be used to construct high-yielding pathways for the
production of α-substituted compounds, particularly 2-methyl-
3-hydroxy acids that may be used as monomers for controlling
the material properties of plastics.46
propionicum44 afforded 1.1
0.2 g L−1 of 2-methyl-3-
oxobutyrate in strain 58, corresponding to 44% conversion of
supplemented propionate (Supporting Information Figure
S27).
CONCLUSION
■
The noncanonical branched acid fermentation of A. suum serves
as a starting point to identify new enzymes involved in
physiological thiolase-based pathways. Characterization of this
pathway also yields new genetic components for engineering
synthetic metabolic pathways for production of α-branched
acids. Of the five thiolases expressed in A. suum, two (Acat2 and
Acat3) were found to demonstrate a preference for propionyl-
CoA over acetyl-CoA. Upon further characterization in the C−
C bond cleaving or degradative direction, we observed that
Acat2 and Acat3 demonstrated up to 640-fold increase in
accommodation of branched substrates in comparison to the
control thiolase, Acat5. Interestingly, sequence alignments show
that Acat5 contains a Phe residue that was reported to serve as
the gatekeeper residue to allow for substrate branching,21
highlighting the need to identify and characterize more thiolase
enzymes to increase our understanding of sequence-substrate
selectivity relationships. We have also examined a ketoreduc-
tase, AsHadh, which we assign to this pathway based on its
preference for branched substrates. Further stereochemical
characterization indicates that AsHadh prefers the 3S- rather
than the 3R-hydroxy enantiomer, despite falling in the same
superfamily of ketoacyl reductases as the 3R-hydroxy acid-
forming ketoreductases exemplified by PhaB in the PHA
pathway.45 Characterization of the dehydratase, AsEch, shows
that it can accommodate branched substrates with a 15-fold
preference for the linear congener.
METHODS
■
Detailed procedures for plasmid construction, protein purification,
acyl-CoA substrate synthesis and characterization, cell culture,
biochemical assays, and metabolite analysis are provided in Supporting
Information.
ASSOCIATED CONTENT
* Supporting Information
■
S
Supporting Information includes a full description of . The
Supporting Information is available free of charge on the ACS
Publications website at DOI: 10.1021/jacs.7b07400.
Materials and methods; DNA and protein sequences;
SDS-PAGE, NMR spectra, HPLC and size-exclusion
chromatography traces; biochemical and production data
(PDF)
These enzymes have been used to demonstrate a cellular
route for the production of branched organic acids starting
from glucose and propionate, including α-methyl enoic acids
(1120 60 mg L−1) and chiral 2-methyl-3-hydroxy acids (1100
200 mg L−1) at 87% and 44% yield from fed propionate,
respectively. Based on biochemical and thermodynamic
analysis, we propose that the product distribution of these
pathways is controlled both by the ketoreductase selectivity as
well as difference in reaction equilibrium for dehydration of
linear and branched 3-hydroxy acids. Given the reversibility of
the thiolase-catalyzed condensation, our data suggest that the
20-fold preference of AsHadh for branched substrates leads to
the dominant production of branched products even when the
thiolase demonstrates a strong preference against these
pathways (Acat5). This model is consistent with the
observation that the selectivity for branched 3-hydroxy acids
is significantly diminished with CnHadh, which displays only a
4-fold branched substrate preference in vitro. However, even
with a 1000-fold selectivity difference, the Acat5 thiolase retains
a catalytically competent kcat/KM (∼104) for branched
substrates. This effect appears to be further amplified in the
case of enoic acids, possibly because of a change of equilibrium
of 3-hydroxy acid dehydration conferred by the α-methyl
substituent to favor the enoic acid product.
AUTHOR INFORMATION
Corresponding Author
*mcchang@berkeley.edu
■
ORCID
Michael R. Blaisse: 0000-0002-2336-0223
Michelle C. Y. Chang: 0000-0003-3747-7630
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
We thank Dr. Jeffery Hanson for purification of PhaA, PhaB,
PhaJ, Hbd, and Crt enzymes and Tristan de Rond for advice on
chemical synthesis. M.R.B. acknowledges the support of an
NSF Graduate Research Fellowship. H.D. thanks the U.C.
Berkeley Tang Distinguished Scholars Program for a
postdoctoral fellowship. This work was funded by the generous
support of the Center for Sustainable Polymers, a National
Science Foundation-supported center for Chemical Innovation
(Grant CHE-1413862). The College of Chemistry NMR
F
DOI: 10.1021/jacs.7b07400
J. Am. Chem. Soc. XXXX, XXX, XXX−XXX

Journal of the American Chemical Society
Article
(31) Willadsen, P.; Eggerer, H. Eur. J. Biochem. 1975, 54, 247.
Facility at U.C. Berkeley is supported in part by the National
Institutes of Health (1S10RR023679 and 1S10RR16634-01).
(32) Agnihotri, G.; Liu, H. W. Bioorg. Med. Chem. 2003, 11, 9.
(33) McMahon, M. D.; Prather, K. L. J. Appl. Environ. Microbiol.
2014, 80, 1042.
(34) Pfeifer, B. A.; Admiraal, S. J.; Gramajo, H.; Cane, D. E.; Khosla,
C. Science 2001, 291, 1790.
(35) Flamholz, A.; Noor, E.; Bar-Even, A.; Milo, R. Nucleic Acids Res.
2012, 40, D770.
(36) Espah Borujeni, A.; Channarasappa, A. S.; Salis, H. M. Nucleic
Acids Res. 2014, 42, 2646.
REFERENCES
■
(1) Clomburg, J. M.; Crumbley, A. M.; Gonzalez, R. Science 2017,
355, 355.
(2) Liao, J. C.; Mi, L.; Pontrelli, S.; Luo, S. Nat. Rev. Microbiol. 2016,
14, 288.
(3) Zhou, K.; Edgar, S.; Stephanopoulos, G. In Methods in
Enzymology; Sarah, E. O. C., Ed.; Academic Press: 2016; Vol. 575, p
225.
(4) Bond-Watts, B. B.; Bellerose, R. J.; Chang, M. C. Nat. Chem. Biol.
2011, 7, 222.
(37) Lee, S.-H.; Park, S. J.; Lee, S. Y.; Hong, S. H. Appl. Microbiol.
Biotechnol. 2008, 79, 633.
(38) Redondo-Nieto, M.; Barret, M.; Morrisey, J. P.; Germaine, K.;
́
Martínez-Granero, F.; Barahona, E.; Navazo, A.; Sanchez-Contreras,
(5) Dellomonaco, C.; Clomburg, J. M.; Miller, E. N.; Gonzalez, R.
Nature 2011, 476, 355.
M.; Moynihan, J. A.; Giddens, S. R.; Coppoolse, E. R.; Muriel, C.;
Stiekema, W. J.; Rainey, P. B.; Dowling, D.; O’Gara, F.; Martín, M.;
Rivilla, R. J. Bacteriol. 2012, 194, 1273.
(6) Jiang, M.; Zhang, H. Curr. Opin. Biotechnol. 2016, 42, 1.
(7) Meadows, A. L.; Hawkins, K. M.; Tsegaye, Y.; Antipov, E.; Kim,
Y.; Raetz, L.; Dahl, R. H.; Tai, A.; Mahatdejkul-Meadows, T.; Xu, L.;
Zhao, L.; Dasika, M. S.; Murarka, A.; Lenihan, J.; Eng, D.; Leng, J. S.;
Liu, C.-L.; Wenger, J. W.; Jiang, H.; Chao, L.; Westfall, P.; Lai, J.;
Ganesan, S.; Jackson, P.; Mans, R.; Platt, D.; Reeves, C. D.; Saija, P. R.;
Wichmann, G.; Holmes, V. F.; Benjamin, K.; Hill, P. W.; Gardner, T.
S.; Tsong, A. E. Nature 2016, 537, 694.
(39) Wong, B. J.; Gerlt, J. A. J. Am. Chem. Soc. 2003, 125, 12076.
(40) Ismail, W. Arch. Microbiol. 2008, 190, 451.
(41) Song, F.; Thoden, J. B.; Zhuang, Z.; Latham, J.; Trujillo, M.;
Holden, H. M.; Dunaway-Mariano, D. Biochemistry 2012, 51, 7000.
(42) Song, F.; Zhuang, Z.; Dunaway-Mariano, D. Bioorg. Chem. 2007,
35, 1.
(43) Yuzawa, S.; Eng, C. H.; Katz, L.; Keasling, J. D. Biochemistry
2013, 52, 3791.
(44) Selmer, T.; Willanzheimer, A.; Hetzel, M. Eur. J. Biochem. 2002,
269, 372.
(45) Cantu, D. C.; Dai, T.; Beversdorf, Z. S.; Reilly, P. J. Protein Eng.,
Des. Sel. 2012, 25, 803.
(8) Haapalainen, A. M.; Merilainen, G.; Wierenga, R. K. Trends
̈
Biochem. Sci. 2006, 31, 64.
(9) Fox, A. R.; Soto, G.; Mozzicafreddo, M.; Garcia, A. N.;
Cuccioloni, M.; Angeletti, M.; Salerno, J. C.; Ayub, N. D. Gene 2014,
533, 5.
(10) Pereto,
65.
́ ́
J.; Lopez-García, P.; Moreira, D. J. Mol. Evol. 2005, 61,
(46) Schneiderman, D. K.; Hillmyer, M. A. Macromolecules 2016, 49,
2419.
(11) Cheong, S.; Clomburg, J. M.; Gonzalez, R. Nat. Biotechnol. 2016,
34, 556.
(12) Martin, C. H.; Dhamankar, H.; Tseng, H.-C.; Sheppard, M. J.;
Reisch, C. R.; Prather, K. L. J. Nat. Commun. 2013, 4, 1414.
(13) Sheppard, M. J.; Kunjapur, A. M.; Wenck, S. J.; Prather, K. L. J.
Nat. Commun. 2014, 5, 5031.
(14) Watanabe, Y.; Ishizuka, K.; Furutate, S.; Abe, H.; Tsuge, T. RSC
Adv. 2015, 5, 58679.
(15) Tijsma, E. J.; Does, L. V. D.; Bantjes, A.; Vulic, I. J. Macromol.
Sci., Polym. Rev. 1994, 34, 515.
(16) de Mata, Z. S.; Lizardo, R.; Diaz, F.; Saz, H. J. Arch. Biochem.
Biophys. 1991, 285, 158.
(17) Komuniecki, R.; Campbell, T.; Rubin, N. Mol. Biochem.
Parasitol. 1987, 24, 147.
(18) Dunn, E. W.; Coates, G. W. J. Am. Chem. Soc. 2010, 132, 11412.
(19) Altschul, S. F.; Gish, W.; Miller, W.; Myers, E. W.; Lipman, D. J.
J. Mol. Biol. 1990, 215, 403.
(20) Benson, D. A.; Clark, K.; Karsch-Mizrachi, I.; Lipman, D. J.;
Ostell, J.; Sayers, E. W. Nucleic Acids Res. 2015, 43, D30.
(21) Haapalainen, A. M.; Merilainen, G.; Pirila, P. L.; Kondo, N.;
Fukao, T.; Wierenga, R. K. Biochemistry 2007, 46, 4305.
(22) Claros, M. G.; Vincens, P. Eur. J. Biochem. 1996, 241, 779.
(23) Nielsen, H.; Engelbrecht, J.; Brunak, S.; von Heijne, G. Protein
Eng., Des. Sel. 1997, 10, 1.
(24) Kim, E.-J.; Kim, K.-J. Biochem. Biophys. Res. Commun. 2014, 452,
124.
(25) Sato, S.; Maruyama, H.; Fujiki, T.; Matsumoto, K. J. Biosci
Bioeng 2015, 120, 246.
(26) Jiang, C.; Kim, S. Y.; Suh, D. Y. Mol. Phylogenet. Evol. 2008, 49,
691.
(27) Ofman, R.; Ruiter, J. P. N.; Feenstra, M.; Duran, M.; Poll-The,
B. T.; Zschocke, J.; Ensenauer, R.; Lehnert, W.; Sass, J. O.; Sperl, W.;
Wanders, R. J. A. Am. J. Hum. Genet. 2003, 72, 1300.
(28) Soler, J.; de Arriaga, D.; Cadenas, Q.; Cadenas, E. Exp. Mycol.
1981, 5, 357.
(29) Cooper, W. C.; Jin, Y.; Penning, T. M. J. Biol. Chem. 2007, 282,
33484.
(30) Luo, M. J.; Mao, L. F.; Schulz, H. Arch. Biochem. Biophys. 1995,
321, 214.
G
DOI: 10.1021/jacs.7b07400
J. Am. Chem. Soc. XXXX, XXX, XXX−XXX