PH-Rate profiles establish that polyketide synthase dehydratase domains utilize a single-base mechanism

Article abstract of DOI:10.1039/c8ob02637h

FosDH1 from module 1 of the fostriecin polyketide synthase (PKS) catalyzes the dehydration of a 3-hydroxybutyryl-SACP to the (E)-3-butenoyl-SACP. The steady-state kinetic parameters, k_cat and k_cat/K_m, were determined over the pH range 3.0 to 9.2 for the FosDH1-catalyzed dehydration of the N-acetycsteamine thioester, 3-hydroxybutyryl-SNAC (3), to (E)-3-butenoyl-SNAC (4). The pH rate profiles for both log(k_cat) and log(k_cat/K_m) each corresponded to a single pH-dependent ionization to give an active site general base, with a calculated pK_a 6.1 ± 0.2 for k_cat and pK_a 5.7 ± 0.1 for k_cat/K_m. These results are inconsistent with the commonly suggested "two-base" (base-acid) mechanism for the dehydratases of PKS and fatty acid biosynthesis and support a simple one-base mechanism in which the universally conserved active site His residue acts as the base to deprotonate C-2 of the substrate, then redonates the proton to the C-3 hydroxyl group to promote C-O bond-cleavage and elimination of water. The carboxylate of the paired Asp or Glu residue is thought to bind and orient the hydroxyl group of the substrate in the stereoelectonically favored conformation.

Full text of DOI:10.1039/c8ob02637h

Organic &
Biomolecular Chemistry
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PAPER
pH-Rate profiles establish that polyketide synthase
dehydratase domains utilize a single-base
mechanism†
Cite this: Org. Biomol. Chem., 2018,
16, 9165
Xinqiang Xie
and David E. Cane
*
FosDH1 from module 1 of the fostriecin polyketide synthase (PKS) catalyzes the dehydration of a 3-hydroxy-
butyryl-SACP to the (E)-3-butenoyl-SACP. The steady-state kinetic parameters, kcat and kcat/Km, were
determined over the pH range 3.0 to 9.2 for the FosDH1-catalyzed dehydration of the N-acetycsteamine
thioester, 3-hydroxybutyryl-SNAC (3), to (E)-3-butenoyl-SNAC (4). The pH rate profiles for both log(kcat
)
and log(kcat/Km) each corresponded to a single pH-dependent ionization to give an active site general
base, with a calculated pKa 6.1 0.2 for kcat and pKa 5.7 0.1 for kcat/Km. These results are inconsistent
with the commonly suggested “two-base” (base-acid) mechanism for the dehydratases of PKS and fatty
acid biosynthesis and support a simple one-base mechanism in which the universally conserved active
site His residue acts as the base to deprotonate C-2 of the substrate, then redonates the proton to the
C-3 hydroxyl group to promote C–O bond-cleavage and elimination of water. The carboxylate of the
paired Asp or Glu residue is thought to bind and orient the hydroxyl group of the substrate in the stereo-
electonically favored conformation.
Received 24th October 2018,
Accepted 13th November 2018
DOI: 10.1039/c8ob02637h
rsc.li/obc
display characteristic β + α double hotdog folds, in which each
active site harbours a universally conserved dyad of His and
Introduction
One or more dehydratase (DH) domains are present in all of Asp or Glu residues, typically separated by 4.1 Å, that are posi-
the dozens of biochemically characterized modular polyketide tioned above largely hydrophobic substrate binding
a
synthases (PKSs), as well as in the nearly 15 000 predicted pocket.5–12 Site-directed mutagenesis experiments have con-
modular PKSs that have been revealed by bacterial genome firmed that these paired His and Asp or Glu residues are each
mining.1–4 Each such DH domain is responsible for the de- essential to the catalytic activity of DH enzymes.11 DH
hydration of a specific 3-hydroxyacyl-acyl carrier protein (ACP) domains from several modular PKSs,6,13–16 as well as FabA and
or 2-alkyl-3-hydroxyacyl-ACP intermediate to give the corres- FabZ proteins17,18 and the analogous DH domain of the multi-
ponding enoyl-ACP or 2-alkylenoyl-ACP. These unsaturated domain yeast FAS,19 have all been shown to catalyse stereo-
intermediates are in turn either reduced by a paired enoyl specific syn dehydration of their respective 2-methyl-3-hydroxy-
reductase domain to the corresponding saturated acyl-ACP or, acyl-ACP or 3-hydroxyacyl-ACP substrates.
more commonly, passed directly to the proximal downstream
PKS module for a further round of polyketide chain elongation β-hydroxydecanoyl dehydratase-isomerase (FabA) and the
and modification.
closely related dehydratase (FabZ) of E. coli, Schwab18,20,21
In early speculation on the mechanism of the
Polyketide synthase DH domains exhibit significant levels extended the original ideas of Rose and Hanson on the
of mutual amino acid sequence similarity and protein struc- relationship between enzyme stereospecificity and mecha-
tural homology to the DH domains of type I fatty acid nism22 to point out that the demonstrated overall syn (supra-
synthases (FASs), as well as to the closely related discrete DH facial) stereochemistry of the dehydratase-catalyzed reaction
enzymes, FabA and FabZ, of type II bacterial fatty acid biosyn- favors a one-base mechanism (while not excluding an alterna-
thesis. All known PKS and FAS dehydratase protein structures tive “two-base” (more correctly, base-acid) process), whereas
the alternative anti (antarafacial) steric course would be com-
patible only with a “two-base” (base-acid) mechanism (barring
rotation of the reaction intermediate in the dehydratase active
site) (Scheme 1). Nonetheless, there have been no reported
Department of Chemistry, Box H, Brown University, Providence, Rhode Island,
02912-9108, USA. E-mail: David_cane@brown.edu
†Electronic supplementary information (ESI) available: LC-MS data, steady-state
experiments directed at distinguishing between these canoni-
cal one-base and base-acid mechanisms. Indeed, since 1996,
kinetic plots, table of pH-dependent kinetic parameters. See DOI: 10.1039/
c8ob02637h
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Scheme 2 FosDH1-catalyzed dehydration of (3R)-1 to (E)-2 by the fos-
triecin PKS.
Scheme 1 Dehydratase mechanistic models. a. One-base (B:)
mechanism. b. Base-acid (B: – AH) mechanism.
following the structural characterization of FabA and FabZ pro-
teins from a variety of microbial sources, a base-acid dehydra-
tion mechanism has come to be commonly invoked,8–13
inspired largely by the observed spatial relationship between
the substrate and the His – Asp/Glu dyads, in which the univer-
sally conserved active site His residue is proposed to serve as
the general base to remove the H-2 proton of the 3-hydroxy-
acyl-ACP substrate, with the paired Asp or Glu side chain car-
boxyl thought to act as the general acid to protonate the
3-hydroxyl oxygen and thus promote the requisite cleavage of
the C–O bond (Scheme 1b). As superficially appealing as such
proposals might seem, however, the base-acid mechanism has
an intrinsic flaw, since it would require that the pKa of the nor-
mally acidic Asp carboxylate (solution pKa ∼ 4) exceed that of
the His imidazolium/imidazole pair (solution pKa ∼ 6).
Although the observed pKa of an active site amino acid side
chain can differ by 2–3 pH units from those of the corres-
ponding free amino acids (and in one case an active site Asp
has been reported to have a pKa > 9),23 to date there has been
no direct experimental evidence to support the requisite per-
turbations of the pKa of the active site Asp/Glu or His residues
of any PKS or FAS dehydratase. Although some investigators
have pointed out this apparent conundrum,8,24 to the best of
our knowledge there have been no published studies of the pH
dependence of any FAS dehydratase, and only a single determi-
nation of the pH rate profile for the kcat/Km of a PKS DH
domain.25 We now report that the pH-rate behavior of both kcat
and kcat/Km of a prototypical PKS DH domain corresponds to a
single ionization to an active basic form, consistent only with
a single-base dehydration mechanism, and therefore inconsist-
ent with an alternative base-acid mechanism requiring two
pH-dependent ionizations.
Scheme 3 Reversible FosDH1-catalyzed dehydration of (3R)-3 to (E)-4.
In order to determine the pH dependence of the FosDH1-
catalyzed dehydration reaction, we chose, as surrogates for the
natural -S-FosACP1 substrate and product, the corresponding
N-acetylcysteamine analogues, (3R)-3-hydroxybutyryl-SNAC (3)
and (E)-2-butenoyl-SNAC (4), respectively (Scheme 3). The com-
monly used SNAC thioesters lack ionizable substituents, unlike
the natural -SFosACP1 thioester or the -SCoA analog, thereby
avoiding any confounding pH-dependent effects on substrate
binding to the FosDH1 active site or perturbation of established
DH–ACP charge–charge surface interactions.9,26 The FosDH1-
catalyzed dehydration of 3 to 4, as well as the reverse hydration
reaction, were conveniently monitored by high performance
liquid chromatography – mass spectrometry (LC-MS) analysis of
ethyl acetate extracts of periodically withdrawn samples of each
incubation mixture (Fig. S1–S3†). The identity of the individual
substrate and product peaks was confirmed by MS–MS ana-
lysis. At pH 7.2, FosDH1 exhibited kcat for 3 of 24 2 min−1
and Km 65 7 mM, corresponding to a kcat/Km 370 M−1 min−1
.
The reverse reaction, the FosDH1-catalyzed hydration of
2-butenoyl-SNAC (4) to 3-hydroxybutyryl-SNAC (3), had kcat 12
2 min−1 and Km 23 7 mM (kcat/Km 520 M−1 min−1).
To determine the pH dependence of the FosDH1-catalyzed
dehydration of (3R)-3-hydroxybutyryl-SNAC (3), the steady state
kinetic parameters were determined in a series of incubations
carried out from pH 3.0 to 9.2 (Fig. S4, Table S1†).
FosDH1 had negligible dehydratase activity below pH 5.0, with
increasing activity at pH 6.0 that reached a maximum from pH
7.0–9.2 (Fig. 1 and S5, S6†). The observed pH dependence of
kcat was fit to a single acid–base ionization27,28 with a calcu-
Results
lated pKa 6.1
reflected a single ionization, pKa 5.7 0.1.
0.1, while the pH dependence of kcat/Km
We have recently established that FosDH1, from module 1 of
the fostriecin polyketide synthase, catalyzes the reversible
stereospecific dehydration of (3R)-3-hydroxybutyryl-SFosACP1
(1) to (E)-2-butenoyl-SFosACP1 (2) (Scheme 2). FosDH1 is a
prototypical dehydratase, as typified by the presence of the
universally conserved active site His-Asp dyad, housed within
Discussion
characteristic DHXXXGXXXXP and DXXXH motifs that are The observation of a single ionization in the pH–rate profiles
common to PKS and FAS DH domains and proteins.
for both log(kcat) and log(kcat/Km) is inconsistent with a base-
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A recently published study of PicDH2, the DH domain from
module 2 of the picromycin PKS, included determination of
the pH dependence of the log (Vmax/Km) of PicDH2, which
exhibited a half-bell-shaped profile.25 The observed pKa 7.0
0.1 was assigned to the deprotonation of the active site His
residue. In spite of the evidence for only a single ionization
over the pH range 6.6–9.0, the authors invoked the commonly
proposed “two-base” DH mechanism, suggesting that the
active site Asp residue would serve as the proton source. In any
case, the pH dependence of kcat/Km reflects only the properties
of the free enzyme at the limit of zero substrate concentration.
By contrast, the pH dependence of kcat reflects the catalytically
relevant pKa values of the enzyme with bound substrate.
In light of the compelling evidence for a single-base DH
mechanism, what then might be the role of the carboxylate of
the essential active site Asp residue? We suggest that rather
than Asp acting as a general acid, the primary role of the nega-
tively charged Asp side chain is to bind and orient the sub-
strate by an essential H-bond to the hydroxyl group of the
3-hydroxyacyl-SACP substrate (Scheme 4a). In the reverse
hydration reaction, the carboxylate of this Asp side chain
would similarly H-bond to the nucleophilic water. Strong
experimental analogy supporting this mechanistic model for
DH domains can be found in the results of extensive protein
structural, site-directed mutagenesis, pH rate, kinetic, and
spectroscopic studies of the analogous enoyl-SCoA hydratase
(ECH) reaction of the fatty acid oxidation pathway.29–31
Although enoyl-SCoA hydratase has a protein structure and
primary amino acid sequence that are entirely distinct from
those of the FAS and PKS DH domains, the underlying bio-
chemical reaction mechanism for the Michael addition of
water to an 2-enoyl-SCoA substrate bears striking similarities
to the reverse of the DH-catalyzed dehydration reaction
(Scheme 4b). In place of the conserved His – Asp/Glu dyad of
the dehydratases, the active site of enoyl-CoA hydratase har-
bours a pair of catalytically essential Glu residues that play an
Fig. 1 pH dependence of FosDH1 (replicate 1). A. log(kcat) vs. pH. B. log
(kcat/Km) vs. pH.
acid (“two-base”) mechanism for the FosDH1-catalyzed de-
hydration of (3R)-3-hydroxybutyryl-SNAC (3), which would have
given rise to a bell-shaped pH rate profile. The observed pH
dependence fully supports a single-base mechanism, with the
observed pKa 6.1 for kcat, which reflects the properties of the
enzyme under saturating conditions, consistent with the
action of the basic active site His residue. The fact that the
observed dehydratase activity shows no decrease up to pH 9.2
is incompatible with any mechanism in which the active site
Asp serves as a general acid, thereby excluding the commonly
invoked “two-base” (base-acid) mechanism for the DH-cata-
lyzed dehydration.
Scheme 4 One-base dehydration and hydration mechanisms. (a) Dehydration by PKS and FAS DH domains. (b) Enoyl-CoA hydratase.
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analogous catalytic role. Thus the E164 carboxylate has been and a Phenomenex Jupiter C16 column (150 mm × 2 mm,
shown to serve as the single active site base for the deprotona- 5.0 μm) was utilized for analysis of SNAC esters.
tion of the nucleophilic water and reprotonation at C2 of the
Protein expression and purification. Recombinant FosDH1
resulting enolate, while the carboxylate of E144 binds and was expressed and purified by procedures similar to those pre-
orients this active site water. Consistent with this mechanism, viously described.24 Competent E. coli BL21(DE3) cells were
it has been established that both protons of the nucleophilic transformed with the pET28a vector containing FosDH1 with
water are incorporated into the 3-hydroxybutyryl-SCoA product.30 codons optimized for expression in E. coli and the resultant
recombinants were cultured under standard conditions. A
single recombinant clone was inoculated into 10 ml LB
medium containing 50 μg ml−1 kanamycin and grown at 37 °C
Conclusions
overnight. The overnight culture was transferred into 500 ml of
The pH dependence of both kcat and kcat/Km for the dehydra- Super Broth with 50 μg ml−1 kanamycin in a 2.5 L flask and
tion of 2-hydroxybutyryl-SNAC (3) to 2-butenoyl-SNAC (4), cata- grown until an OD600 of 0.4–0.8. The cultures were cooled to
lysed by FosDH1, a prototypical PKS dehydratase, each exhibi- 18 °C and 0.2 mM IPTG was added to induce protein
ted a single ionization from inactive to active form with a expression. The cell culture was continuously grown for an
pKa ∼ 6. Combined with the syn stereochemistry established additional 40–48 h at 18 °C and cells were then harvested by
for many DH-catalyzed eliminations, these results are incom- centrifugation at 4200g for 20 min. The resulting cell pellet was
patible with the commonly invoked two-base (base-acid) de- dissolved in 35 ml lysis buffer (1 M NaCl, 50 mM sodium phos-
hydratase mechanism, and are instead fully consistent with a phate, 75 mM imidazole, 1 mg ml−1 of lysozyme, pH 7.8) and
one-base mechanism in which the active site His residue stored at −80 °C. Frozen cells in lysis buffer were thawed at
serves as the base that first removes the H-2 proton of the sub- room temperature, followed by sonication. The cell supernatant
strate, then transfers this proton directly to the hydroxyl group and pellet were separated by centrifugation at 23 000g for
of the intermediate enolate, thereby facilitating cleavage of the 30 min. The supernatant was filtered using a 0.8 μm filter and
C–O bond with net elimination of water. The negatively loaded onto a pre-charged 5 ml HisTrap™ FF column. The
charged carboxylate of the active site Asp is unable to serve as column was washed with 25 mL lysis buffer and then 25 mL
a general acid, but instead binds and orients the hydroxyl washing buffer (50 mM sodium phosphate, 1 M NaCl, 75 mM
group of the 3-hydroxyacyl thioester substrate in the preferred imidazole, pH 7.6). Proteins were eluted by elution buffer
stereo-electronic conformation for elimination of water.
(150 mM NaCl, 50 mM phosphate, 150 mM imidazole, pH 7.5).
The eluted fractions were collected, concentrated with an
Amicon filter (MWCO 30 000), and the buffer was exchanged
with exchange buffer (50 mM sodium phosphate, 10% glycerol,
100 mM NaCl, pH 7.2), concentrated, and stored at −80 °C until
use. Protein purity was assessed as >90% by 12% acrylamide
Experimental section
Materials and methods
Isopropylthio-β-D-galactopyranoside (IPTG) and ampicillin SDS-PAGE, and the N-terminal His-tagged proteins were utilized
were purchased from Thermo Scientific. E. coli BL21(DE3) was without further modification.
purchased from New England BioLabs. All other chemical
Determination of initial velocity conditions. A typical assay
reagents were purchased from Sigma-Aldrich and utilized consisted of FosDH1 (1.25, 2.5 or 5.0 μM) and 10 mM substrate
without further purification. Amicon Ultra Centrifugal Filter 3 or 4 in a total volume of 700 μL of 50 mM phosphate buffer
Units (Amicon Ultra-15 and Amicon Ultra-4, 30 000 MWCO) (2.5 mM Tris(2-carboxyethyl)phosphine (TCEP), pH 7.2). At 5,
were purchased from Millipore. Pre-charged 5 mL HisTrap™ 10 and 15 min time points, 100 μL of reaction mixture was
FF columns were purchased from GE Healthcare Life Sciences. withdrawn and added to 100 μL of 0.5 M HCl to quench the
(3R)-3-hydroxybutyryl-SNAC (3) was synthesized from (3R)- reaction, then extracted with ethyl acetate (3 × 500 μL). After
hydroxybutyric acid and SNAC, as previous described.32 (E)-2- evaporation of the solvent, the concentrated organic extract
Butenoyl-SNAC (4) was synthesized from (E)-2-butenoyl chlor- was dissolved in 200 μL methanol and analyzed by HPLC-MS.
ide and N-acetylcysteamine, as previous described.32,33 General HPLC was carried out at a flow rate of 0.2 ml min−1 at room
methods were as previously described.34 Growth media and temperature. Eluent A was 0.1% formic acid in water, and
conditions used for E. coli strains and standard methods for eluent B was 100% acetonitrile. HPLC conditions used were
handling E. coli in vivo and in vitro were those described pre- 5%–95% buffer B for 10 min, 95% buffer B for another 1 min,
viously,34 unless otherwise noted. All proteins were handled at 100%–5% buffer B for 3 min and then 5% buffer B for 4 min.
4 °C unless otherwise stated. Protein concentrations were Each reaction was performed in duplicate. Progress curves at
determined according to the method of Bradford,35 using a varying enzyme concentrations were generated and the initial
Tecan infinite M200 Microplate Reader with bovine serum velocity at each enzyme concentration was obtained (Fig. S1
albumin as the standard. Protein purity and size was estimated and S2†). The amount of enzymatic product formed at each
using SDS-PAGE, visualized using Coomassie Blue stain, and time point was calculated using eqn (1):
analyzed with a Bio-Rad ChemiDoc MP System. A Thermo LXQ
LC-mass spectrometer equipped with Surveyor HPLC system
Ap ¼ As ꢀ Pp=ðPs þ PpÞ
ð1Þ
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where Ap: amount of product; As: initial amount of substrate;
Pp: relative MS intensity of product in MS spectra of retention
Conflicts of interest
time between 1 min and 10 min; Ps: relative MS intensity for There are no conflicts to declare.
substrate in MS spectra of retention time between 1 min and
10 min.
Kinetic analysis of FosDH1-catalyzed dehydration and
hydration reactions by LC-MS/MS. Enzymatic reactions were
carried out in a total volume of 250 μL under initial velocity
Acknowledgements
The work was supported by a grant from the U. S. National
Institutes of Health, GM022172, to D. E. C. We would also like
to thank one of the referees for valuable comments and for
suggesting an alternative statistical analysis of the pH-rate
data, as described in the ESI.†
conditions containing FosDH1 (2.5 μM), reaction buffer
(50 mM sodium phosphate, 100 mM NaCl, pH 7.2) and sub-
strates 3 or 4 at variable concentrations (0.5, 1, 4, 10, 20,
60 mM). The incubation mixture also contained ∼1% each of
glycerol from the protein solution and DMSO from the stock
solutions of 3 or 4. After incubation at room temperature for
15 min, 100 μL reaction mixture was transferred into a new
Eppendorf tube, acidified with 100 μL of 1 M HCl, and
extracted with ethyl acetate (3 × 500 μL). The concentrated
organic extract was dissolved in 200 μL methanol and analyzed
by HPLC-MS. Parallel control incubations for each concen-
tration of substrate were performed without the addition of
enzyme. Each reaction and analysis were performed in dupli-
cate (Fig. S3†). The steady-state kinetic parameters were deter-
mined by fitting the observed rate and substrate concentration
data to the Michaelis–Menten equation by non-linear least
squares regression using GraphPad Prism 7.0d. Reported stan-
dard deviations in the steady-state kinetic parameters rep-
resent the calculated statistical errors in the non-linear, least
squares regression analysis.
pH dependence profile of FosDH1. The effect of pH on the
steady-state kinetic parameters for FosDH1-catalyzed dehydra-
tion of 3 was determined using a series of 50 mM buffers of
increasing pH: citric acid-sodium citrate (pH 3.0, 4.0, 5.0 and
6.0), HEPES (pH 7.0 and 8.0), and sodium carbonate (pH 9.2).
At each pH value (3.0, 4.0, 5.0, 6.0, 7.0, 8.0, and 9.2), the incu-
bations were conducted with 3-hydroxybutyryl-SNAC (3, 0.5, 1,
4, 10, 20, 60 mM) under the conditions described above. The
kcat and kcat/Km values at each pH were obtained by fitting the
Notes and references
1 D. A. Hopwood, Methods in Enzymology. Complex enzymes in
microbial natural product biosynthesis, part B: polyketides,
aminocoumarins and carbohydrates, Academic Press, United
States, 2009.
2 K. J. Weissman, Nat. Chem. Biol., 2015, 11, 660–670.
3 C. Khosla, D. Herschlag, D. E. Cane and C. T. Walsh,
Biochemistry, 2014, 53, 2875–2883.
4 R. V. O’Brien, R. W. Davis, C. Khosla and
M. E. Hillenmeyer, J. Antibiot., 2014, 67, 89–97.
5 A. Keatinge-Clay, J. Mol. Biol., 2008, 384, 941–953.
6 D. Gay, Y. O. You, A. Keatinge-Clay and D. E. Cane,
Biochemistry, 2013, 52, 8916–8928.
7 D. L. Akey, J. R. Razelun, J. Tehranisa, D. H. Sherman,
W. H. Gerwick and J. L. Smith, Structure, 2010, 18, 94–105.
8 L. Moynie, S. M. Leckie, S. A. McMahon, F. G. Duthie,
A. Koehnke, J. W. Taylor, M. S. Alphey, R. Brenk,
A. D. Smith and J. H. Naismith, J. Mol. Biol., 2013, 425,
365–377.
9 M. Leesong, B. S. Henderson, J. R. Gillig, J. M. Schwab and
J. L. Smith, Structure, 1996, 4, 253–264.
observed initial rate and substrate concentration data to the 10 D. Kostrewa, F. K. Winkler, G. Folkers, L. Scapozza and
Michaelis–Menten equation by non-linear least squares R. Perozzo, Protein Sci., 2005, 14, 1570–1580.
regression using GraphPad Prism 7.0d. The pKa value of the 11 M. S. Kimber, F. Martin, Y. Lu, S. Houston, M. Vedadi,
ionizable group was obtained by fitting the observed values of
A. Dharamsi, K. M. Fiebig, M. Schmid and C. O. Rock,
kcat and kcat/Km at each pH 5.0–9.2 to eqn (2) and (3), respect-
J. Biol. Chem., 2004, 279, 52593–52602.
ively, corresponding to the observed half-bell shape with a 12 D. T. Wagner, J. Zeng, C. B. Bailey, D. C. Gay, F. Yuan,
single acidic break (Fig. S4–S6, Table S1†):
H. R. Manion and A. T. Keatinge-Clay, Structure, 2017, 25,
1045–1055.
13 C. R. Valenzano, Y. O. You, A. Garg, A. Keatinge-Clay,
C. Khosla and D. E. Cane, J. Am. Chem. Soc., 2010, 132,
14697–14699.
logðkcatÞ ¼ log C=ð1 þ ½Hþꢁ=KaÞ
logðkcat=KmÞ ¼ log C=ð1 þ ½Hþꢁ=KaÞ
ð2Þ
ð3Þ
where, C is the pH-independent plateau value of kcat or kcat/Km, 14 X. Guo, T. Liu, C. R. Valenzano, Z. Deng and D. E. Cane,
[H+] is the hydrogen ion concentration, and Ka is the dis-
J. Am. Chem. Soc., 2010, 132, 14694–14696.
sociation constant of the acid.27,28 Two independent replicates 15 X. Xie, A. Garg, C. Khosla and D. E. Cane, J. Am. Chem. Soc.,
of each pH rate series were carried out. The data for each repli- 2017, 139, 3283–3292.
cate were also analysed over the entire experimental pH range 16 X. Xie, A. Garg, C. Khosla and D. E. Cane, J. Am. Chem. Soc.,
3.0–9.2 using an alternative formulation of the Logistical 2017, 139, 9507–9510.
equations for the pH behaviour of kcat or kcat/Km, as described 17 J. M. Schwab, J. B. Klassen and A. Habib, J. Chem. Soc.,
in the ESI,† with each plot displaying a single pKa (Fig. S6†).
Chem. Commun., 1986, 357–358.
This journal is © The Royal Society of Chemistry 2018
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View Article Online
Paper
Organic & Biomolecular Chemistry
18 J. M. Schwab, A. Habib and J. B. Klassen, J. Am. Chem. Soc., 27 H. Serrano and J. S. Blanchard, Biochemistry, 2013, 52,
1986, 108, 5304–5308. 5009–5015.
19 B. Sedgwick, C. Morris and S. J. French, J. Chem. Soc., 28 L. A. Reinhardt, J. B. Thoden, G. S. Peters, H. M. Holden
Chem. Commun., 1978, 193–194. and W. W. Cleland, FEBS Lett., 2013, 587, 2876–2881.
20 J. M. Schwab and J. B. Klassen, J. Am. Chem. Soc., 1984, 29 H. A. Hofstein, Y. Feng, V. E. Anderson and P. J. Tonge,
106, 7217–7227. Biochemistry, 1999, 38, 9508–9516.
21 J. M. Schwab and B. S. Henderson, Chem. Rev., 1990, 90, 30 B. J. Bahnson, V. E. Anderson and G. A. Petsko,
1203–1245.
Biochemistry, 2002, 41, 2621–2629.
22 K. R. Hanson and I. A. Rose, Acc. Chem. Res., 1975, 8, 1–10.
23 N. A. Wilson, E. Barbar, J. A. Fuchs and C. Woodward,
Biochemistry, 1995, 34, 8931–8939.
24 D. D. Shah, Y.-O. You and D. E. Cane, J. Am. Chem. Soc.,
2017, 139, 14322–14330.
31 R. L. D’Ordine, J. Pawlak, B. J. Bahnson and
V. E. Anderson, Biochemistry, 2002, 41, 2630–2640.
32 S. K. Piasecki, C. A. Taylor, J. F. Detelich, J. Liu, J. Zheng,
A. Komsoukaniants, D. R. Siegel and A. T. Keatinge-Clay,
Chem. Biol., 2011, 18, 1331–1340.
25 Y. Li, G. J. Dodge, W. D. Fiers, R. A. Fecik, J. L. Smith and 33 O. Vergnolle, F. Hahn, A. Baerga-Ortiz, P. F. Leadlay and
C. C. Aldrich, J. Am. Chem. Soc., 2015, 137, 7003–7006. J. N. Andexer, ChemBioChem, 2011, 12, 1011–1014.
26 C. Nguyen, R. W. Haushalter, D. J. Lee, P. R. Markwick, 34 J. Sambrook, E. F. Fritsch and T. Maniatis, Molecular
J. Bruegger, G. Caldara-Festin, K. Finzel, D. R. Jackson,
F. Ishikawa, B. O’Dowd, J. A. McCammon, S. J. Opella,
S. C. Tsai and M. D. Burkart, Nature, 2014, 505, 427–431.
Cloning, A Laboratory Manual, Cold Spring Harbor
Laboratory Press, Cold Spring Harbor, NY, 2nd edn, 1989.
35 M. Bradford, Anal. Biochem., 1976, 72, 248–254.
9170 | Org. Biomol. Chem., 2018, 16, 9165–9170
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