Directed evolution of the tryptophan synthase β-subunit for stand-alone function recapitulates allosteric activation

Article abstract of DOI:10.1073/pnas.1516401112

Enzymes in heteromeric, allosterically regulated complexes catalyze a rich array of chemical reactions. Separating the subunits of such complexes, however, often severely attenuates their catalytic activities, because they can no longer be activated by their protein partners. We used directed evolution to explore allosteric regulation as a source of latent catalytic potential using the β-subunit of tryptophan synthase from Pyrococcus furiosus (PfTrpB). As part of its native αββα complex, TrpB efficiently produces tryptophan and tryptophan analogs; activity drops considerably when it is used as a stand-alone catalyst without the α-subunit. Kinetic, spectroscopic, and X-ray crystallographic data show that this lost activity can be recovered by mutations that reproduce the effects of complexation with the α-subunit. The engineered PfTrpB is a powerful platform for production of Trp analogs and for further directed evolution to expand substrate and reaction scope.

Full text of DOI:10.1073/pnas.1516401112

Directed evolution of the tryptophan synthase
β-subunit for stand-alone function recapitulates
allosteric activation
Andrew R. Buller1, Sabine Brinkmann-Chen1, David K. Romney, Michael Herger, Javier Murciano-Calles,
and Frances H. Arnold2
Division of Chemistry and Chemical Engineering, California Institute of Technology, Pasadena, CA 91125
Edited by Alan R. Fersht, Medical Research Council Laboratory of Molecular Biology, Cambridge, United Kingdom, and approved October 16, 2015 (received
for review August 17, 2015)
associated with open, partially closed, and fully closed states
during the catalytic cycle (2, 4, 6).
Enzymes in heteromeric, allosterically regulated complexes cata-
lyze a rich array of chemical reactions. Separating the subunits of
such complexes, however, often severely attenuates their catalytic
activities, because they can no longer be activated by their protein
partners. We used directed evolution to explore allosteric regula-
tion as a source of latent catalytic potential using the β-subunit of
tryptophan synthase from Pyrococcus furiosus (PfTrpB). As part of
its native αββα complex, TrpB efficiently produces tryptophan and
tryptophan analogs; activity drops considerably when it is used as
a stand-alone catalyst without the α-subunit. Kinetic, spectro-
scopic, and X-ray crystallographic data show that this lost activity
can be recovered by mutations that reproduce the effects of com-
plexation with the α-subunit. The engineered PfTrpB is a powerful
platform for production of Trp analogs and for further directed
evolution to expand substrate and reaction scope.
TrpS is a naturally promiscuous enzyme complex: the model
system from S. typhimurium catalyzes its β-substitution reaction
with most haloindoles, methylindoles, and aminoindoles, along
with an assortment of nonindole nucleophiles for C–S, C–N, and
C–C bond formation (7). Such noncanonical amino acids (NCAAs)
have diverse applications in chemical biology (8), serve as inter-
mediates in the synthesis of natural products (9, 10), and are priv-
ileged scaffolds for the development of pharmaceuticals (11).
Despite its natural ability to produce these desirable compounds,
TrpS has enjoyed only limited application (7). Optimized methods
are restricted by low substrate concentrations and yields typically
below 50% (7, 12, 13). To produce NCAAs, researchers have used
the S. typhimurium TrpS complex (StTrpS), which suffers from poor
thermostability and low tolerance to organic solvents (14). Although
only the reactivity of the β-subunit, coupling of L-serine and indole,
is necessary and desirable for synthetic applications, using TrpB as
an isolated enzyme has not been feasible. Outside of its native
complex, TrpB loses up to 95% of its native activity and is subject to
inactivation (15, 16). These facts combine to present a compelling
challenge: can we use directed evolution to recover the activity lost
when TrpA is removed and create a highly active stand-alone TrpB
enzyme? If so, what are the structural and kinetics effects of the
mutations that harness this latent catalytic potential?
protein engineering allostery noncanonical amino acid PLP
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eteromeric enzyme complexes catalyzing a rich array of
useful reactions are often allosterically regulated by their
H
protein partners, such that the catalytic subunits are much less
active when isolated (1–3). Utilization of isolated enzyme sub-
units, however, is desirable for biosynthetic applications, where
expressing large complexes increases the metabolic load on the
host cell and complicates efforts to engineer activity, substrate
specificity, stability, and other properties. A solution to these prob-
lems is to engineer the isolated subunit to function independently.
However, it is unknown to what extent mutations in the amino acid
sequence can reproduce the complex structural and functional
changes that are induced by binding of a partner protein.
Significance
Many enzymes perform desirable biochemical transformations,
but are not suitable to use as biocatalysts outside of the cell. In
particular, enzymes from heteromeric complexes typically have
decreased activity when removed from their protein partners.
We used directed evolution to restore the catalytic efficiency of
the tryptophan synthase β-subunit (TrpB), which synthesizes
L-tryptophan from L-serine and indole, surpassing the activity
of the native complex. Experiments show that activating muta-
tions promote catalysis through the same mechanism as part-
ner protein binding, establishing that isolated subunits may be
readily reactivated through directed evolution. Engineering
TrpB for stand-alone function restored high activity with indole
analogs, providing a simplified enzyme platform for the bio-
catalytic production of noncanonical amino acids.
Tryptophan synthase (TrpS; EC 4.2.1.20) is a heterodimeric
complex that catalyzes the formation of L-tryptophan (Trp, 1)
from L-serine (Ser, 2) and indole glycerol phosphate (IGP, 3; Fig.
1A) (2). The mechanism of this transformation has been exten-
sively studied for TrpS from Escherichia coli and Salmonella
typhimurium, where it has been shown the enzyme consists of two
subunits, TrpA (α-subunit) and TrpB (β-subunit), both of which
have low catalytic efficiencies in isolation (4). The activities of
both subunits increase upon complex formation and are further
regulated by an intricate and well-studied allosteric mechanism
(2). IGP binding to the α-subunit stimulates pyridoxal phosphate
(PLP)-dependent aminoacrylate formation in the β-subunit
[E(A-A); Fig. 1B], which in turn promotes retro-aldol cleavage of
IGP in the α-subunit, releasing indole (4). This tightly choreo-
graphed mechanism serves to prevent the free diffusion of indole,
which is only released from the α-subunit when the complex is in a
closed conformation that forms a 25-Å tunnel through which indole
diffuses into the β-subunit (5). Here, indole reacts with E(A-A) in a
C–C bond-forming reaction, yielding L-tryptophan as product
(Fig. 1B) (2, 5). These allosteric effects are mediated through the
rigid-body motion of the communication (COMM) domain and
a monovalent cation (MVC) binding site within the β-subunit
(Fig. 1A), which undergo complex conformational transitions
Author contributions: A.R.B., S.B.-C., D.K.R., M.H., and J.M.-C. designed research; A.R.B.,
S.B.-C., D.K.R., M.H., and J.M.-C. performed research; A.R.B., S.B.-C., D.K.R., M.H., J.M.-C.,
and F.H.A. analyzed data; and A.R.B., S.B.-C., D.K.R., and F.H.A. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The atomic coordinates have been deposited in the Protein Data Bank,
www.pdb.org (PDB ID codes 5DVZ, 5DW0, 5DW3, and 5E0K).
1A.R.B. and S.B.-C. contributed equally to this work.
2To whom correspondence should be addressed. Email: frances@cheme.caltech.edu.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.
1073/pnas.1516401112/-/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.1516401112
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Fig. 1. Catalytic cycle of the native TrpS complex. (A) Indole (4) is released through a retro-aldol reaction in TrpA (red) and then diffuses through a 25-Å
tunnel into TrpB (black), where a PLP-mediated β-substitution reaction occurs with L-serine (2), yielding L-tryptophan (1). The COMM domain is indicated in
blue. Scheme is superimposed over PfTrpS with Lys-PLP internal aldimine, E(Ain), shown in green sticks. The native complex is an αββα heterotetramer; a single
αβ pair is shown for clarity. (B) Mechanism of L-tryptophan formation. Transimination of L-serine to form an external aldimine, E(Aex1), followed by de-
hydration across Cα–Cβ through a quinonoid intermediate, E(Q1), is designated stage I of the overall reaction. A mechanistically similar process occurs in
reverse for the addition of indole into E(A-A) and subsequent release of L-tryptophan, designated stage II of the reaction.
L-tryptophan took place at 75 °C for up to 1 h (generation 1, 1 h;
generations 2 and 3, 6 min). The reactions were then quenched
in an ice-water bath, and the production of L-tryptophan was
quantified at 290 nm with a plate reader. This procedure iden-
tified many activating mutations, with 3.8% (20/528) of the
screened variants of generation 1 showing at least 40% greater
product formation than the parent. A single mutation, T292S,
gave rise to a 3.5-fold increase in kcat compared with PfTrpB, which
completely recovered kcat and even exceeded the catalytic efficiency
(kcat/KM) of the PfTrpS complex (Table 1). Twenty-six other mu-
tations in 19 different variants also contributed activating effects
(Fig. S1).
We recombined 12 of the most activating mutations identified
in the first generation (SI Materials and Methods). Screening a
total of 1,408 clones, we identified PfTrpB4D11 containing mu-
tations E17G, I68V, F274S, T292S, and T321A. The kcat of
PfTrpB4D11 was 2.2 s−1, a sevenfold improvement over wild type
(Table 1). PfTrpB4D11 served as template for a final round of ran-
dom mutagenesis, for which we screened 1,144 clones. PfTrpB0B2
carried one additional mutation, P12L, and had a kcat that was 9.4-
fold higher than PfTrpB and threefold higher than PfTrpS. With
PfTrpB0B2 we had surpassed our goal of engineering an isolated
enzyme domain to be as active as in its native complex.
Results
Selection of the Parent Enzyme, TrpB, from Pyrococcus furiosus. We
focused the search for an engineering starting point on known
thermophilic TrpS enzymes for three reasons: (i) higher operating
temperatures afford increased solubility of the hydrophobic sub-
strates, which is useful for preparative reactions; (ii) thermostable
enzymes are more tolerant to the introduction of activating but
potentially destabilizing mutations (17); and (iii) thermostable en-
zyme variants can be screened efficiently (described below). We
compared published kinetic properties of TrpS from Thermotoga
maritima (18), Thermococcus kodakaraensis (19), and Pyrococcus
furiosus (20–22) and selected the last for its superior kinetic pa-
rameters and thermostability (Table S1). In our hands, PfTrpB
heterologously expressed and purified from E. coli has a kcat of
0.31 s−1 and experiences a 12-fold increase in catalytic efficiency
upon addition of purified P. furiosus TrpA (PfTrpA) to make the
PfTrpS complex (Table 1), similar to values reported previously
for E. coli TrpB (EcTrpB) (15). Notably, PfTrpB does not show
the mechanism-based inactivation that inhibits use of StTrpB for
preparation of NCAAs (23).
Directed Evolution of PfTrpB for Stand-Alone Function. We per-
formed random mutagenesis by error-prone PCR and screened a
small PfTrpB mutant library (528 clones) for increased Vmax
under saturating concentrations of substrate (L-serine and indole).
The extreme thermostability of the parent enzyme permitted a
1-h heat treatment of the lysates at high temperature (75 °C) that
precipitated the majority of E. coli proteins and ensured that any
activated variants retained significant stability. Formation of
Biochemical Comparison of Evolved PfTrpB Enzymes with PfTrpS.
Kinetic analysis of PfTrpB and PfTrpS established that the
12-fold increase in the catalytic efficiency for indole upon com-
plexation is driven by both an increase in kcat and decrease in KM
(Table 1). Despite screening the mutant libraries under saturating
Table 1. Biochemical characterization of tryptophan synthases
kcat/KM, mM-1s·−1 kcat change with
Enzyme
Mutations
kcat,s−1 KM, mM L-serine KM, μM indole
indole
TrpA*
T50 °C†
PfTrpS
PfTrpB
—
1.0
0.31
1.1
2.2
2.9
0.6
1.2
0.84
1.2
0.7
20
77
14
11
50
4
78
200
330
—
3.2
0.34
0.3
> 95
95
95
84
87
—
PfTrpB2G9
PfTrpB4D11
PfTrpB0B2
T292S
E17G, I68V, T292S, F274S, T321A
P12L, E17G, I68V, T292S, F274S, T321A
8.7
0.04
All kinetic data are measured at 75 °C using a previously described (15) continuous assay for Trp production. Additional mutations relative to their parent
enzyme are highlighted in bold.
*The effect of PfTrpA was measured by addition of a fivefold stoichiometric excess of PfTrpA under conditions saturating in each substrate.
†T50 is the temperature of half-maximal activity after incubation for 1 h. Assay parameters are described in Materials and Methods. SE for each measurement
is given in Table S2.
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conservation constitutes firm experimental support for applying
insights gleaned from StTrpS and EcTrpS to the function of
PfTrpB. Labeling studies have shown that β-substitution occurs
with retention of stereochemistry, indicating the hydroxyl of
E(Aex1) must rotate 180° from its crystallographically observed
state to eliminate from the same face to which indole is added (26).
The H-bond between E(Aex1) and Asp300 is, therefore, transient
during the catalytic cycle and suggests an important role for the
H-bond between Thr292 and Asp300 that we observed in the
ligand-free structure. Notably, T292S was one of the most activating
mutations identified in the first round of directed evolution
(Fig. 4 A and B and Table 1). When L-Ser is incubated with
PfTrpB2G9, the UV-vis spectrum clearly shows that the equilibrium
is shifted toward E(A-A) (Fig. S2B). We hypothesize that this
mutation alters the energetics of the Asp300-E(Aex1) interaction
to favor the fully closed conformational state of the enzyme and
thereby accelerate the reaction.
The structure of PfTrpB with L-tryptophan bound in the active
site at 1.74 Å resolution shows the product is not covalently
linked to the PLP cofactor, but is in a novel ligand-binding pose
(Fig. 3C). Residues Thr105 to His110 comprise a carboxylate
binding motif (4) that we report also forms H-bonds to the pri-
mary amine of L-tryptophan through the backbone N–H of Ala106.
This same residue also H-bonds with the Ser-hydroxyl of E(Aex1).
Asp300, however, is not observed to interact with L-tryptophan. A
hydrogen bond between Glu104 and the N-1 of L-tryptophan likely
also occurs when indole binds. This interaction may serve to in-
crease the nucleophilicity of indole by positioning C-3 close to the
acrylate of E(A-A) and by increasing electron density of the arene
(Fig. 1B) (2, 4). Glu104 is located in the COMM domain and shifts
closer to the active site upon closure (Fig. 3A). In each of the
substrate- and product-bound structures, we observed a rotameric
shift of Phe274 and His275 (Fig. 4C) that may function as a gate
within the indole tunnel (Fig. 4D). In the structure of PfTrpS, we
observe a hydrogen bond between His275 and Asp43 from the
α-subunit, indicating that PfTrpA binding stabilizes this open
conformation (Fig. S4A). In StTrpS, NMR analysis has shown
this motion is concerted, and our observation that it is conserved
across species is consistent with it having an important role for
regulating catalytic function (27). The location of the F274S
mutation is therefore quite striking, because it may alter the
energetics of this transition to favor higher activity.
Fig. 2. UV-vis absorption spectra of native and engineered PfTrpB. (A) The
PLP absorption spectrum of PfTrpB (black) has a λmax = 412 nm, characteristic
of E(Ain). Addition of 20 mM L-serine causes a red-shift to 428 nm (gray),
consistent with E(Aex1) formation. (B) Addition of L-serine to PfTrpS causes a shift
in λmax to 350 nm, which is attributed to the E(A-A). A residual peak at 428 nm
indicates population of E(Aex1). (C) Like PfTrpS, addition of 20 mM L-serine to
PfTrpB0B2 shows λmax = 350 nm as well as contributions from E(Aex1). All spectra
were collected with 20 μM of enzyme. See Materials and Methods for details.
conditions for both substrates, we observed a steady decrease in
the KM for indole in the enzymes with greater activity, which
mimics the behavior of the native complex. The KM for L-serine
fluctuated during evolution, and in the final round the values for
PfTrpB0B2 and PfTrpS were similar.
Though Michaelis–Menten kinetic analysis provides a simple
readout of overall catalytic performance, activity changes upon
complex formation and during directed evolution are associated
with a shift in the populations of the intermediates in the TrpB
catalytic cycle. The PLP cofactor absorbs in the UV-vis region,
and different chemical intermediates have characteristic peaks
that can be measured readily (Fig. 1B). Under turnover condi-
tions, the recorded spectrum reflects the Boltzmann-weighted
average of each of the intermediates in the catalytic cycle. In-
cubation of PfTrpB with L-serine results in a large increase in
absorption at 428 nm, consistent with accumulation of E(Aex1)
(Fig. 2A). A peak at 320 nm developed over several minutes,
whereas the absorption at 428 nm remained constant (Fig. S2),
which we attribute to the previously characterized serine deaminase
activity that is an artifact of not having a competing nucleophile
present (24). In contrast, L-serine-bound PfTrpS has a λmax near
350 nm, consistent with a shift in the equilibrium to populate the
E(A-A) (Fig. 2B) (2). With the engineered proteins, the external
aldimine absorbance at 428 nm decreases, and the absorbance
bands at 350 nm grow in intensity over the course of directed
evolution of Trp synthase activity (Fig. 2C and Fig. S2).
Two mutations, E17G and P12L, map onto the α/β interface of
a previously determined 3.0-Å structure of PfTrpS (22), but this
low resolution precluded confident assessment of the side-chain
interactions for these residues. We solved a higher-resolution
One question motivating this study was how mutational reac-
tivation alters the allosteric regulation of PfTrpB by its TrpA pro-
tein partner. Adding purified PfTrpA to each TrpB variant gave a
surprising result: instead of enhancing activity (or doing nothing),
PfTrpA inhibited each improved TrpB, an effect that was stronger
with each evolutionary step, to the point where PfTrpB0B2 had just
4% of its activity when PfTrpA was added (Table 1).
Structural Analysis of PfTrpB. Whereas the residues that perform
the TrpB chemistry are conserved throughout evolution, the
conformational motions associated with allosteric signaling have
not been characterized outside of StTrpS or in the absence of the
α-subunit. We obtained high-resolution structures of wild-type
PfTrpB in its ligand-free state, bound to its L-serine substrate,
and bound to its L-tryptophan product (Table S3). Comparison
of the ligand-free and Ser-bound structures (at 1.69 Å and 2.0 Å
resolution, respectively) reveals motion of the COMM domain
into a partially closed state upon serine binding, as well as a
hydrogen bond between Asp300 and the Ser-hydroxyl of E(Aex1)
(Fig. 3 A and B). This large conformational rearrangement is
structurally equivalent to that characterized for StTrpS (25), with
0.5 Å rmsd between the E(Aex1) forms of TrpB (Fig. S3), despite
the modest (59%) sequence identity and lack of an α-subunit.
Combined with their similar biochemical properties, this structural
Fig. 3. Structural transitions upon ligand binding in PfTrpB. (A) Superim-
position of PfTrpB-E(Ain) and PfTrpB-E(Aex1) in gray and cyan, respectively.
Overlay shows the 2.1-Å displacement of the COMM domain upon E(Aex1)
formation. This closure moves the side chain of Glu104 by 3.7 Å, toward its
catalytic orientation (orange dashes). (B) Structure of Ser-bound PfTrpB with
a Fo–Fc map of E(Aex1) contoured at 3.0 σ (green). (C) Structure of L-tryp-
tophan–bound PfTrpB with Fo–Fc map of Trp ligand contoured at 3.0 σ.
Buller et al.
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panel of substrate analogs to study promiscuous activity in the na-
tive and engineered proteins (Fig. 5A). The StTrpS complex reacts
with most halogenated and methylated indoles, which only modestly
change the steric and electronic properties of the indole ring and
afford reactivity at C3. Alternatively, indazole (5) and indoline (6),
which have substantially altered electronic properties, react at N1
for C–N bond formation. We measured the relative rate of PfTrpB
compared with PfTrpS and observed much larger rate enhance-
ments for TrpB on the indole derivatives than on the native sub-
strate (Fig. 5B), up to a 100-fold increase in the rate of NCAA
synthesis with 5-bromoindole (7). These results raised an important
question about our objective to engineer PfTrpB for function as an
independent subunit by directed evolution: will screening for higher
activity with indole translate into increased activity on alternative
substrates, or will evolution generate an enzyme that is highly active
on indole but less active toward other substrates, as has been ob-
served in directed evolution of other enzymes (31, 32)?
We measured the activity of PfTrpB0B2 on the substrate panel
(Fig. 5B) and were pleased to observe a substantial increase in
activity relative to PfTrpB for all nucleophiles tested. The activity
profile is broadly similar to that of PfTrpS, with a few exceptions.
The PfTrpS complex reacts approximately eightfold faster than
PfTrpB0B2 with bromoindole 7, whereas PfTrpB0B2 reacts ap-
proximately sixfold faster than PfTrpS with indoline 6 and
threefold faster with indazole 5. The identity of each product was
confirmed by mass spectrometry and NMR in reactions using
PfTrpB0B2. Because we screened for increases in Vmax during
laboratory evolution, the variants we selected also have improved
expression levels relative to PfTrpB. E. coli cultures produce
threefold more soluble and active PfTrpB0B2 than PfTrpB, ex-
ceeding 230 mg enzyme per liter culture, facilitating preparative
reactions and future use of PfTrpB0B2 as a biocatalyst.
Fig. 4. Distribution of PfTrpB0B2 mutations and interaction networks al-
tered by mutational reactivation. (A) PfTrpB residues within 5 Å of PfTrpA
(red) are colored cyan and the COMM domain is colored blue. (B) A hydro-
gen bond between D300 and E(Aex1) in the Ser-bound structure (orange) is
formed transiently during the catalytic cycle. When this H-bond is severed,
D300 may interact with T292 (no ligands, red, or Trp-bound, blue). This
complex network is centered on a monovalent cation cofactor, shown here
as Na+, which is known to mediate allosteric interactions between the α- and
β-subunits (6). (C) Residues F274 and H275 undergo a rotameric shift upon
substrate or product binding into an open state (blue). (D) In the closed state
(red, no ligands bound), H275 blocks access to the active site.
Discussion
We evolved PfTrpB for independent function by selecting mu-
tations that increased its Vmax in heat-treated E. coli lysates. We
found that activating mutations were unusually common, which
could in part be attributed to the high thermal stability of the
parent enzyme that increases tolerance to the introduction of
random mutations (17). However, it is clear that many mutations
have the potential to recover activity lost when TrpB is removed
from its native complex. The single T292S mutation raised the
catalytic efficiency of PfTrpB on indole by almost 20-fold and
completely restored activity to TrpS-like levels (Table 1). Fur-
ther evolution resulted in the identification of PfTrpB0B2, which
retained T292S as well as five additional mutations. This variant
has an 83-fold increase in catalytic efficiency on indole compared
with PfTrpB. The T292S and F274S mutations occur in dynamic
regions of TrpB related to catalytic function, as discussed above.
However, the majority of the mutations in PfTrpB0B2, and 40%
structure at 2.76 Å that revealed a salt bridge between Glu17 of
PfTrpB and Arg148 of PfTrpA (Fig. S4B). Pro12 of PfTrpB is an
evolutionarily conserved residue that lies along the indole tunnel
between the two subunits, where previous studies have found
that mutation to bulkier residues inhibits substrate channeling
(28). Such interactions are clearly disrupted at the α/β interface
(Fig. S4C), but the strong inhibition of PfTrpB0B2 upon PfTrpA
addition demonstrates that these proteins still associate. Finally,
two mutations present in PfTrpB0B2, I68V and T321A, are distal
to sites that undergo an observable structural change upon substrate
binding or complex formation, and their contribution to rate en-
hancement is difficult to rationalize. Overall, 60% of the activating
mutations identified through random mutagenesis were located
within 5 Å of the α/β interface, the COMM domain, or regions that
undergo observable motion upon transition to the closed confor-
mation (Fig. S1). These positions comprise just 31% of the protein
sequence, indicating modest enrichment within residues in the im-
mediate spatial route between the α/β interface and the β-subunit
active site. In contrast to the many mutations that have been
identified as deleterious to the allosteric communication in TrpS
(29, 30), the mutations identified here using directed evolution
are the first reported to affect allosteric communication and
increase the activity of TrpB in isolation.
PfTrpB0B2 Is a Stand-Alone Catalyst for Production of NCAAs. Our
selection of TrpB to evolve for high activity outside of its native
complex was motivated by practical considerations. Previous
studies have shown that StTrpS is a promiscuous enzyme,
capable of synthesizing diverse analogs of L-tryptophan (7, 13).
These analogs require subtly different transition state stabiliza-
tion for catalysis, but the role of allostery in promoting this de-
sirable chemistry has not been studied. We selected a representative
Fig. 5. Substrate profile of native and engineered TrpB enzymes. (A) Indole
analogs that have been reported to react with StTrpS were tested for reactivity
with the PfTrpB, PfTrpS, and PfTrpB0B2 enzymes. The nucleophilic atom is in-
dicated with a gray circle. (B) Relative activities of enzyme complex PfTrpS (black)
and PfTrpB0B2 (gray) compared with PfTrpB. Reactions performed in duplicate
with 20 mM of each substrate and varying enzyme concentrations to ensure
incomplete conversion after 1 h. Products were later confirmed in scaled-up
reactions using PfTrpB0B2. See Materials and Methods for details.
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of all activating mutations that we identified, are at positions that
do not undergo any observable structural changes upon forma-
tion of the partially closed state, nor do the corresponding res-
idues in StTrpS move upon formation of the fully closed state
(2). The location of many activating mutations outside of any
observed allosteric site raises the central question of this study:
did directed evolution increase activity through the same mecha-
nism(s) through which TrpA binding regulates function? If so, we
posit that increases in kcat, which was under selective pressure
during directed evolution, would be coupled to other kinetic and
spectroscopic characteristics of the native complex for which there
was no selective pressure.
To a first approximation, the open–close equilibrium of TrpB
can be inferred from its KM for indole because TrpA binding
stabilizes the closed conformation (2) and decreases the KM
(Table 1). Each generation of evolution of PfTrpB produced a
lower KM for indole, to values below those in PfTrpS, despite
screening under saturating conditions (Table 1). UV-vis spec-
troscopy affords a more-sensitive probe of the active site envi-
ronment because the steady-state distribution of intermediates in
the catalytic cycle can be directly observed. Measurement of the
steady-state population of the stage I intermediates in PfTrpB
shows that the electrophilic E(A-A) state is stabilized relative to
E(Aex1) when PfTrpA binds. This same stabilization of E(A-A)
is observed in TrpB enzymes evolved for independent function
(Fig. 2). These data reflect the properties of PfTrpB in its native
reaction. Activity on indole analogs is also significantly increased
by PfTrpA binding, with the rate enhancements differing by two
orders of magnitude between diverse indole analogs (Fig. 5).
Because each of these substrates requires slightly different
transition state stabilization, the rate enhancement upon effector
addition is a sensitive metric for changes in the conformational
ensemble that are relevant to catalysis. Consistent with the data
measured on the native reaction, we observed that increases in
activity with indole analogs caused by mutation were similar to
the effects of TrpA addition (Fig. 5). Taken together, these data
support the hypothesis that increases in PfTrpB activity from
directed evolution arose through the same mechanism by which
effector binding activates catalysis.
A perplexing observation about mutational reactivation is that
addition of PfTrpA to the engineered proteins did not result in
allosteric activation, but instead inhibited catalysis. A simple
open–closed model of the ensemble might imply that TrpA
binding and mutations that stabilize the closed state would be
additive in their effects on catalysis. Though the introduction of
mutations may change the nature of the allosteric signaling that
results from TrpA binding, no such assumption is required to
explain the inhibition. It is well documented that stabilization of
the closed state by effector binding alters the energetics of
multiple transition states of TrpB (2). These changes in transi-
tion-state energy result in a lower overall barrier to the reaction.
Doubling these energetic perturbations (e.g., TrpA and the
mutations combined) would lower the energy of the original rate-
limiting step, but could end up impeding catalysis by increasing the
energy of a different step above the original reaction barrier (Fig.
S5). With this explanation, the inhibitory effect of TrpA becomes
consistent with the demonstration that the mutations and effector
addition accelerate catalysis through the same mechanism.
Allostery is common in enzymes and has been targeted in
previous protein engineering endeavors (vide infra). To compare
the role of allostery in these diverse efforts, we define the change
in activity that arises through effector binding as an enzyme’s
allosteric potential. In this view, it has been shown that allosteric
potential can be decreased by using mutagenesis to disrupt
specific interactions between fructose-1,6-bisphosphatase and its
effector, adenosine monophosphate (33). Recent studies have
also shown that engineering can increase allosteric potential,
enhancing inhibition of phosphofructokinase by phosphoenol-
pyruvate (34). An allosteric potential can even be engineered
into enzymes where previously one did not exist (35). Nature has
found mutations that constitutively activate allosterically regulated
enzymes, as is well known in phosphorylation cascades contrib-
uting to tumorigenesis (36). Finding such mutations for engi-
neering purposes has proved challenging. The native substrate of
the acyl-transferase LovD is covalently bound to an acyl carrier
protein, LovF, which also functions as an allosteric activator.
LovD was subjected to nine rounds of directed evolution for al-
tered substrate specificity, thermal stability, and tolerance to or-
ganic solvent to generate a biocatalyst for production of the drug
simvastatin (37). Insight into whether increases in activity were
related to allosteric regulation was gleaned from microsecond
molecular dynamics simulations, which suggested that LovF pro-
motes the stability of a closed and catalytically active conformation
of LovD and that directed evolution identified mutations that
increased activity in a similar fashion (37). Our kinetic, structural,
and spectroscopic data provide firm experimental support for a
similar effect in TrpB, demonstrating that allosteric potential can be
readily converted into catalytic activity through directed evolution.
We advocate that future efforts to recoup allosteric potential
not be limited to making mutations within a hypothetical allo-
steric pathway (38). Residues whose interactions are not changed
upon effector addition can nonetheless tune the cooperative
network of interactions that transfer allosteric signals (39). More
generally, allostery enables proteins to respond to environmental
cues, and often only small energetic differences separate active
conformations from inactive or less-active ones. The protein
ensembles of such enzymes are likely to be sensitive to the effects
of widely distributed mutations. Indeed, a single conservative
mutation (T292S) was sufficient to raise the catalytic efficiency
of PfTrpB to the level of the TrpS complex, and many other
mutations contributed smaller activating effects. Therefore, we
anticipate that recovering allosteric potential with directed evo-
lution will be broadly achievable.
These insights into the modulation of protein allosteric regu-
lation are of both fundamental and practical interest. Our ob-
servation that the activity of PfTrpB was enhanced on a range of
substrates is consistent with the hypothesis that the mutations
alter transition-state stabilization in a manner similar to PfTrpA
binding; it also establishes a greatly simplified enzyme platform
for production of NCAAs. This demonstration that isolated
subunits may be readily reactivated through directed evolution
provides a useful tool for the chemical biology community and
expands the scope of enzymes accessible for biocatalysis.
Materials and Methods
Detailed experimental methods are presented in SI Materials and Methods.
Cloning, Expression, and Purification of PfTrpA and PfTrpB. The genes encoding
PfTrpB (UNIPROT ID Q8U093) and PfTrpA (UNIPROT ID Q8U094) were
obtained as gBlocks and cloned into pET22(b)+ for expression in E. coli BL21
E. cloni EXPRESS cells (Lucigen). Heterologous protein expression of PfTrpA
and PfTrpB was performed in Terrific Broth with 100 μg/mL ampicillin
(TBamp) and induced with 500 mM IPTG (final concentration 1 mM). PfTrpB
was purified via a HisTrap HP column. PfTrpA was purified via a Q HP HiTrap
column (flow-through), followed by ammonium sulfate precipitation, and
hydrophobic interaction chromatography on a phenyl Sepharose HP HiTrap
column (Fig. S6).
Library Construction and High-Throughput Screening. Error-prone PCR libraries
were constructed using standard protocols with either MnCl2 or Mutazyme II
(Stratagene). DNA shuffling and site-directed mutagenesis by overlap ex-
tension (SOE) PCR were performed to recombine activating mutations. The
resulting libraries were cloned into pET22(b)+ with the C-terminal his-tag for
expression in E. coli BL21 E. cloni EXPRESS cells. High-throughput expression
and screening were performed on 96-well scale. Formation of L-tryptophan
was recorded at 290 nm.
Kinetics and UV-Vis Spectroscopy. Data were collected between 550 and 250 nm
on a UV1800 Shimadzu spectrophotometer (Shimadzu) using 0.25–20 μM
of enzyme in 200 mM potassium phosphate (pH 8.0) in a quartz cuvette.
Samples were incubated at 75 °C for >3 min to ensure a stable temperature
was reached. PfTrpB activity (kcat) was measured by monitoring tryptophan
formation at 290 nm using Δe290 = 1.89 mM−1·cm−1 (15).
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Substrate Selectivity. The relative rate of NCAA production was measured
using 20 mM L-serine and 20 mM indole analog (Fig. 5A) in 200 mM potas-
sium phosphate (pH 8.0) with 5% (vol/vol) DMSO. Reactions were incubated
at 75 °C for 1 h, quenched, and the relative rate of production formation
was measured by comparing the ratio of the product peaks measured via
ultra HPLC-MS (UHPLC-MS) Agilent 1290 with 6140 MS detector at 280 nm
and then normalizing for the enzyme concentration.
Identification of Nonnatural Amino Acid Products. Preparative-scale reactions
were conducted using PfTrpB0B2, which was prepared as a heat-treated ly-
sate. Products were purified directly on C-18 silica, and their identities con-
firmed by 1H NMR and low-resolution mass spectrometry (LRMS). The optical
purity of the products was estimated by derivatization with N-(5-fluoro-2,
4-dinitrophenyl)alanamide (FDNP-alanamide).
ACKNOWLEDGMENTS. The authors thank Jackson Cahn, Dr. Robert Trachman,
Dr. Mike Chen, and Belinda Wenke for helpful discussions and comments on
the manuscript. We thank the staff of the Caltech Molecular Observatory,
Dr. Jens Kaiser, Dr. Julie Hoy, and Pavle Nikolovski for crystallographic support.
The Molecular Observatory is supported by the Gordon and Betty Moore
Foundation, the Beckman Institute, and the Sanofi-Aventis Bioengineering
Research Program at Caltech. This work was funded through the Jacobs In-
stitute for Molecular Engineering for Medicine; Ruth Kirschstein NIH Post-
doctoral Fellowship F32GM110851 (to A.R.B.); and the Alfonso Martín Escudero
Foundation (J.M.C.).
Crystallography. Crystals of PfTrpB and PfTrpS were grown using the sitting-
drop vapor diffusion method, and cryoprotected before diffraction at
the Stanford Synchrotron Radiation Laboratories on beamline 12-2. Ligand-
bound crystals of PfTrpB were prepared by soaking preformed crystals
with a concentrated solution of L-Ser or L-Trp. Structures were deter-
mined by molecular replacement and models were built using standard
procedures.
1. Du L, Lou L (2010) PKS and NRPS release mechanisms. Nat Prod Rep 27(2):255–278.
2. Dunn MF (2012) Allosteric regulation of substrate channeling and catalysis in the
tryptophan synthase bienzyme complex. Arch Biochem Biophys 519(2):154–166.
3. Davidsen JM, Bartley DM, Townsend CA (2013) Non-ribosomal propeptide precursor
in nocardicin A biosynthesis predicted from adenylation domain specificity dependent
on the MbtH family protein NocI. J Am Chem Soc 135(5):1749–1759.
4. Niks D, et al. (2013) Allostery and substrate channeling in the tryptophan synthase
bienzyme complex: Evidence for two subunit conformations and four quaternary
states. Biochemistry 52(37):6396–6411.
5. Hyde CC, Ahmed SA, Padlan EA, Miles EW, Davies DR (1988) Three-dimensional
structure of the tryptophan synthase alpha 2 beta 2 multienzyme complex from
Salmonella typhimurium. J Biol Chem 263(33):17857–17871.
6. Dierkers AT, Niks D, Schlichting I, Dunn MF (2009) Tryptophan synthase: Structure and
function of the monovalent cation site. Biochemistry 48(46):10997–11010.
7. Phillips RS (2004) Synthetic applications of tryptophan synthase. Tetrahedron
Asymmetry 15(18):2787–2792.
8. Lang K, Chin JW (2014) Cellular incorporation of unnatural amino acids and bio-
orthogonal labeling of proteins. Chem Rev 114(9):4764–4806.
9. Barry SM, et al. (2012) Cytochrome P450–catalyzed L-tryptophan nitration in thax-
tomin phytotoxin biosynthesis. Nat Chem Biol 8(10):814–816.
10. Kieffer ME, Repka LM, Reisman SE (2012) Enantioselective synthesis of tryptophan
derivatives by a tandem Friedel-Crafts conjugate addition/asymmetric protonation
reaction. J Am Chem Soc 134(11):5131–5137.
11. Patel RN (2013) Biocatalytic synthesis of chiral alcohols and amino acids for devel-
opment of pharmaceuticals. Biomolecules 3(4):741–777.
24. Miles EW, McPhie P (1974) Evidence for a rate-determining proton abstraction in the
serine deaminase reaction of the beta 2 subunit of tryptophan synthetase. J Biol
Chem 249(9):2852–2857.
25. Ngo H, et al. (2007) Allosteric regulation of substrate channeling in tryptophan syn-
thase: Modulation of the L-serine reaction in stage I of the beta-reaction by alpha-site
ligands. Biochemistry 46(26):7740–7753.
26. Miles EW, Houck DR, Floss HG (1982) Stereochemistry of sodium borohydride re-
duction of tryptophan synthase of Escherichia coli and its amino acid Schiff’s bases.
J Biol Chem 257(23):14203–14210.
27. McDowell LM, Lee M, McKay RA, Anderson KS, Schaefer J (1996) Intersubunit com-
munication in tryptophan synthase by carbon-13 and fluorine-19 REDOR NMR.
Biochemistry 35(10):3328–3334.
28. Schlichting I, Yang XJ, Miles EW, Kim AY, Anderson KS (1994) Structural and kinetic
analysis of a channel-impaired mutant of tryptophan synthase. J Biol Chem 269(43):
26591–26593.
29. Rowlett R, et al. (1998) Mutations in the contact region between the alpha and beta
subunits of tryptophan synthase alter subunit interaction and intersubunit commu-
nication. Biochemistry 37(9):2961–2968.
30. Raboni S, Bettati S, Mozzarelli A (2005) Identification of the geometric requirements
for allosteric communication between the alpha- and beta-subunits of tryptophan
synthase. J Biol Chem 280(14):13450–13456.
31. Aharoni A, et al. (2005) The ‘evolvability’ of promiscuous protein functions. Nat Genet
37(1):73–76.
32. Bloom JD, Romero PA, Lu Z, Arnold FH (2007) Neutral genetic drift can alter pro-
miscuous protein functions, potentially aiding functional evolution. Biol Direct 2:17.
33. Yang J-S, Seo SW, Jang S, Jung GY, Kim S (2012) Rational engineering of enzyme
allosteric regulation through sequence evolution analysis. PLOS Comput Biol 8(7):
e1002612.
34. McGresham MS, Reinhart GD (2015) Enhancing allosteric inhibition in Thermus
thermophilus Phosphofructokinase. Biochemistry 54(3):952–958.
35. Lee J, et al. (2008) Surface sites for engineering allosteric control in proteins. Science
322(5900):438–442.
12. Smith DRM, et al. (2014) The first one-pot synthesis of L-7-iodotryptophan from
7-iodoindole and serine, and an improved synthesis of other L-7-halotryptophans.
Org Lett 16(10):2622–2625.
13. Goss RJM, Newill PLA (2006) A convenient enzymatic synthesis of L-halotryptophans.
Chem Comm 47:4924–4925.
14. Ahmed SA, Miles EW (1994) Aliphatic alcohols stabilize an alternative conformation
of the tryptophan synthase alpha 2 beta 2 complex from Salmonella typhimurium.
J Biol Chem 269(23):16486–16492.
36. Torkamani A, Verkhivker G, Schork NJ (2009) Cancer driver mutations in protein ki-
nase genes. Cancer Lett 281(2):117–127.
37. Jiménez-Osés G, et al. (2014) The role of distant mutations and allosteric regulation
on LovD active site dynamics. Nat Chem Biol 10(6):431–436.
38. Reynolds KA, McLaughlin RN, Ranganathan R (2011) Hot spots for allosteric regula-
tion on protein surfaces. Cell 147(7):1564–1575.
39. Hilser VJ, Dowdy D, Oas TG, Freire E (1998) The structural distribution of cooperative
interactions in proteins: Analysis of the native state ensemble. Proc Natl Acad Sci USA
95(17):9903–9908.
40. Gibson DG, et al. (2009) Enzymatic assembly of DNA molecules up to several hundred
kilobases. Nat Methods 6(5):343–345.
41. Stemmer WPC (1994) DNA shuffling by random fragmentation and reassembly: In vitro
recombination for molecular evolution. Proc Natl Acad Sci USA 91(22):10747–10751.
42. Kunkel TA, Roberts JD, Zakour RA (1987) Rapid and efficient site-specific mutagenesis
without phenotypic selection. Methods Enzymol 154:367–382.
43. Kabsch W (2010) XDS. Acta Crystallogr D Biol Crystallogr 66(Pt 2):125–132.
44. Evans PR, Murshudov GN (2013) How good are my data and what is the resolution?
Acta Crystallogr D Biol Crystallogr 69(Pt 7):1204–1214.
45. Karplus PA, Diederichs K (2012) Linking crystallographic model and data quality.
Science 336(6084):1030–1033.
46. McCoy AJ, et al. (2007) Phaser crystallographic software. J Appl Cryst 40(Pt 4):658–674.
47. Winn MD, et al. (2011) Overview of the CCP4 suite and current developments. Acta
Crystallogr D Biol Crystallogr 67(Pt 4):235–242.
15. Lane AN, Kirschner K (1983) The catalytic mechanism of tryptophan synthase from
Escherichia coli. Kinetics of the reaction of indole with the enzyme–L-serine com-
plexes. Eur J Biochem 129(3):571–582.
16. Lane AN, Kirschner K (1983) The mechanism of binding of L-serine to tryptophan
synthase from Escherichia coli. Eur J Biochem 129(3):561–570.
17. Bloom JD, Labthavikul ST, Otey CR, Arnold FH (2006) Protein stability promotes
evolvability. Proc Natl Acad Sci USA 103(15):5869–5874.
18. Hettwer S, Sterner R (2002) A novel tryptophan synthase beta-subunit from the hy-
perthermophile Thermotoga maritima. Quaternary structure, steady-state kinetics,
and putative physiological role. J Biol Chem 277(10):8194–8201.
19. Hiyama T, Sato T, Imanaka T, Atomi H (2014) The tryptophan synthase β-subunit
paralogs TrpB1 and TrpB2 in Thermococcus kodakarensis are both involved in tryp-
tophan biosynthesis and indole salvage. FEBS J 281(14):3113–3125.
20. Yamagata Y, et al. (2001) Entropic stabilization of the tryptophan synthase alpha-
subunit from a hyperthermophile, Pyrococcus furiosus. X-ray analysis and calorimetry.
J Biol Chem 276(14):11062–11071.
21. Hioki Y, et al. (2004) The crystal structure of the tryptophan synthase beta subunit
from the hyperthermophile Pyrococcus furiosus. Investigation of stabilization factors.
Eur J Biochem 271(13):2624–2635.
22. Lee SJ, et al. (2005) Conformational changes in the tryptophan synthase from a hy-
perthermophile upon alpha2beta2 complex formation: Crystal structure of the
complex. Biochemistry 44(34):11417–11427.
48. Emsley P, Cowtan K (2004) Coot: Model-building tools for molecular graphics. Acta
Crystallogr D Biol Crystallogr 60(Pt 12 Pt 1):2126–2132.
23. Ahmed SA, Ruvinov SB, Kayastha AM, Miles EW (1991) Mechanism of mutual acti-
vation of the tryptophan synthase alpha and beta subunits. Analysis of the reaction
specificity and substrate-induced inactivation of active site and tunnel mutants of the
beta subunit. J Biol Chem 266(32):21548–21557.
49. Winn MD, Murshudov GN, Papiz MZ (2003) Macromolecular TLS refinement in
REFMAC at moderate resolutions. Methods Enzymol 374:300–321.
50. Chen VB, et al. (2010) MolProbity: All-atom structure validation for macromolecular
crystallography. Acta Crystallogr D Biol Crystallogr 66(Pt 1):12–21.
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