Aurone synthase is a catechol oxidase with hydroxylase activity and provides insights into the mechanism of plant polyphenol oxidases

Article abstract of DOI:10.1073/pnas.1523575113

Tyrosinases and catechol oxidases belong to the family of polyphenol oxidases (PPOs). Tyrosinases catalyze the o-hydroxylation and oxidation of phenolic compounds, whereas catechol oxidases were so far defined to lack the hydroxylation activity and catalyze solely the oxidation of o-diphenolic compounds. Aurone synthase from Coreopsis grandiflora (AUS1) is a specialized plant PPO involved in the anabolic pathway of aurones. We present, to our knowledge, the first crystal structures of a latent plant PPO, its mature active and inactive form, caused by a sulfation of a copper binding histidine. Analysis of the latent proenzyme's interface between the shielding C-terminal domain and the main core provides insights into its activation mechanisms. As AUS1 did not accept common tyrosinase substrates (tyrosine and tyramine), the enzyme is classified as a catechol oxidase. However, AUS1 showed hydroxylase activity toward its natural substrate (isoliquiritigenin), revealing that the hydroxylase activity is not correlated with the acceptance of common tyrosinase substrates. Therefore, we propose that the hydroxylase reaction is a general functionality of PPOs. Molecular dynamics simulations of docked substrate-enzyme complexes were performed, and a key residue was identified that influences the plant PPO's acceptance or rejection of tyramine. Based on the evidenced hydroxylase activity and the interactions of specific residues with the substrates during the molecular dynamics simulations, a novel catalytic reaction mechanism for plant PPOs is proposed. The presented results strongly suggest that the physiological role of plant catechol oxidases were previously underestimated, as they might hydroxylate their - so far unknown - natural substrates in vivo.

Full text of DOI:10.1073/pnas.1523575113

Aurone synthase is a catechol oxidase with
hydroxylase activity and provides insights into the
mechanism of plant polyphenol oxidases
Christian Molitora, Stephan Gerhard Maurachera, and Annette Rompela,1
aInstitut für Biophysikalische Chemie, Fakultät für Chemie, Universität Wien, 1090 Vienna, Austria
Edited by Richard A. Dixon, University of North Texas, Denton, TX, and approved February 19, 2016 (received for review December 3, 2015)
Tyrosinases and catechol oxidases belong to the family of poly-
phenol oxidases (PPOs). Tyrosinases catalyze the o-hydroxylation
and oxidation of phenolic compounds, whereas catechol oxidases
were so far defined to lack the hydroxylation activity and catalyze
solely the oxidation of o-diphenolic compounds. Aurone synthase
from Coreopsis grandiflora (AUS1) is a specialized plant PPO in-
volved in the anabolic pathway of aurones. We present, to our
knowledge, the first crystal structures of a latent plant PPO, its
mature active and inactive form, caused by a sulfation of a copper
binding histidine. Analysis of the latent proenzyme’s interface be-
tween the shielding C-terminal domain and the main core provides
insights into its activation mechanisms. As AUS1 did not accept
common tyrosinase substrates (tyrosine and tyramine), the en-
zyme is classified as a catechol oxidase. However, AUS1 showed
hydroxylase activity toward its natural substrate (isoliquiritigenin),
revealing that the hydroxylase activity is not correlated with the
acceptance of common tyrosinase substrates. Therefore, we pro-
pose that the hydroxylase reaction is a general functionality of
PPOs. Molecular dynamics simulations of docked substrate–enzyme
complexes were performed, and a key residue was identified that
influences the plant PPO’s acceptance or rejection of tyramine.
Based on the evidenced hydroxylase activity and the interactions
of specific residues with the substrates during the molecular dynam-
ics simulations, a novel catalytic reaction mechanism for plant PPOs
is proposed. The presented results strongly suggest that the physi-
ological role of plant catechol oxidases were previously underesti-
mated, as they might hydroxylate their—so far unknown—natural
substrates in vivo.
biosynthetic processes was shown only in a few cases (1, 7–11).
Recently, Sullivan (7) wondered whether there exist more PPOs
that are involved in the plant’s secondary metabolism and whether
the few so far known specialized PPOs represent only “the tip of
the iceberg.”
More than one decade ago, a vacuole-localized PPO, aureusidin
synthase, was identified to be a key enzyme in the anabolic aurone
(yellow pigments) formation in petals of Antirrhinum majus (12, 13).
Recently, it has been reported that aurone biosynthesis in Aster-
aceae species (4-deoxyaurone formation) (14) differs in various
aspects from the 4-hydroxyaurone biosynthesis in A. majus. A PPO
homolog, termed aurone synthase (AUS1, Uniprot A0A075DN54),
was assigned to be involved in 4-deoxyaurone biosynthesis in petals
of Coreopsis grandiflora (tickseed, Asteraceae) (15, 16). As with
most plant PPOs, this enzyme possesses an N-terminal chloroplast
transit peptide and a thylakoid transfer domain and is hence pre-
dicted to be transported to the thylakoid lumen (16). AUS1 com-
prises the conserved motifs of common plant PPOs—for example, a
thioether bridge between a cysteine and a CuA binding histidine
and a phenylalanine (“blocker” or “gate” residue) above the active
site. AUS1 exhibits an insertion in a loop region near the active site
and is therefore classified as a member of the group 2 PPOs (15).
So far uniquely, it possesses a disulfide linkage between the
shielding C-terminal domain and the catalytically active main core.
The latent proenzyme has been shown to undergo an allosteric
activation mechanism, caused by in situ-formed, highly reactive
o-quinones (15). Mechanistic considerations regarding the differences
Significance
crystal structure tyrosinase catechol oxidase type III copper protein
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mechanism
Catechol oxidases and tyrosinases belong to the family of poly-
phenol oxidases (PPOs). In contrast to tyrosinases, catechol oxi-
dases were so far defined to lack hydroxylase activity toward
monophenols. Aurone synthase (AUS1) is a plant catechol oxi-
dase that specializes in the conversion of chalcones to aurones
(flower pigments). We evidence for the first time, to our knowl-
edge, hydroxylase activity for a catechol oxidase (AUS1) toward
its natural monophenolic substrate (chalcone). The presented first
crystal structure of a plant pro-PPO provides insights into its ac-
tivation mechanisms, and based on biochemical and structural
studies of AUS1, we propose a novel catalytic reaction mecha-
nism for plant PPOs. The proven hydroxylase functionality of
AUS1 suggests that other catechol oxidases might also be in-
volved in the plant’s secondary metabolism.
yrosinases and catechol oxidases are polyphenol oxidases
(PPOs) and belong to the family of type III copper proteins.
T
They are widespread among plants, fungi, and bacteria (1, 2).
Plant PPOs display a high diversity, and often several PPO genes
are present in one organism (2, 3). Tyrosinases catalyze the hydrox-
ylation and oxidation of monophenols (e.g., tyrosine) to o-diphenols
and the corresponding o-quinones. Due to the fact that catechol
oxidases are unable to hydroxylate common tyrosinase substrates
(e.g., tyrosine and tyramine), they were so far defined to lack the
hydroxylation (monophenolase) activity and to catalyze solely the
oxidation of o-diphenols to o-quinones (diphenolase activity).
Plant PPOs are expressed as latent proenzymes (∼64 kDa), in
which the active site of the enzyme is shielded by the C-terminal
domain (∼19 kDa). Presumably, the C-terminal domain is pro-
teolytically cleaved during the maturation process, resulting in a
fully active enzyme (4, 5). The latent proenzymes can be acti-
vated in vitro by proteases, an acidic pH, fatty acids, or deter-
gents (e.g., SDS) (6). PPOs are found in almost all plants and
have been assigned primarily to enzymatic browning reactions
and to the protection of organisms against biotic and abiotic
stress (1, 7). However, in most cases, their individual physio-
logical role and especially their natural substrates remain un-
known. Beside aurone formation, their specific involvement in
Author contributions: A.R. designed research; C.M. performed research; C.M. and S.G.M.
analyzed data; and C.M. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Freely available online through the PNAS open access option.
Data deposition: The crystallography, atomic coordinates, and structure factors have been
deposited in the Protein Data Bank, www.pdb.org (PDB ID codes 4Z0Y, 4Z0Z, and 4Z11–4Z13).
1To whom correspondence should be addressed. Email: annette.rompel@univie.ac.at.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.
1073/pnas.1523575113/-/DCSupplemental.
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between tyrosinases and catechol oxidases were so far based on
the comparison of structures of plant catechol oxidases (17, 18)
linker region of approximately 10 amino acids, located in the
vicinity of the interdomain disulfide linkage (Fig. 1C). The
with structures of several bacterial (19, 20) and fungal (21, 22) C-terminal domain displays seven β-strands that form a jelly roll
tyrosinases. However, the comparability of fungal and bacterial barrel motif with the four loops being arranged in a staggered
with plant PPOs is very limited due to the existence of the plant manner above the active site. The latent enzyme displayed very
PPOs’ bulky gate residue, which will have a general high influence
in substrate coordination. Recently, the first crystal structure
of a plant tyrosinase from walnut leaves was reported, and it was
low activity (toward butein) and an allosteric activation (in the
presence of in situ-formed o-quinones) (15). Therefore, the ac-
tive site of a small portion of latent enzyme molecules has to be
revealed that the residues located at the entrance of the active site accessible for substrates. A loop region of the jelly roll motif as
are more important for the acceptance of tyramine and tyrosine
than the previously suggested restriction of the active site (23).
However, the differences in the enzymatic activity and the
mechanism of catechol oxidases and tyrosinases still remain
striking questions.
In the following, to our knowledge, the first crystal structures
of a latent plant pro-PPO as well as its mature activated and an
inactive form, caused by a sulfation of a copper binding histidine,
are presented. Insights into the in vitro activation and the allo-
steric activation of the latent enzyme were obtained by the analysis
of the interdomain interactions of the proenzyme. A combination
of molecular dynamics (MD) simulations of docked natural sub-
strates to active AUS1 and its hydroxylase activity toward its
natural substrate leads to the proposal of a novel catalytic reaction
mechanism of plant PPOs.
well as the loop that carries the insertion show weak or even
completely missing electron density. This demonstrates the flexi-
bility of this region possibly being an entrance for the substrates
(Fig. S2B). The interface between the catalytically active main
core and the shielding C-terminal domain is fairly large and
composed of several hydrogen bonds, hydrophobic interactions,
and water bridges. However, the interdomain interactions are
concentrated at two different locations (Fig. 2). One area is lo-
cated near the linker region and the interdomain disulfide linkage,
whereas the other is located in the vicinity of the active site. The
residue Ile456 is of crucial importance, as it blocks the entrance to
the active site and functions similar to a “plug” (Fig. 1D). This
plug residue is located outside of the active site and on top of the
gate residue (Phe273). It is stabilized by its hydrophobic interac-
tions with the gate residue, one CuB binding histidine (His252),
and Thr253.
Results
Copper Binding Sites of met-, oxy-, and Inactive Forms of AUS1. Active,
latent, and recombinantly expressed latent AUS1 crystallized in
the resting met-form, in which the copper ions (CuII–CuII distance,
∼4 Å) are bridged by a hydroxide ion or a water molecule (Fig.
3A). The geometry of the met-form of AUS1 is fully consistent
with previously published structures (18, 23). To generate the oxy-
form, crystals of recombinantly expressed AUS1 were soaked in
cryoprotectant solution containing hydrogen peroxide (24). The
copper ions (CuII) of the obtained crystal structure are bridged by a
peroxide ion, and the Cu–Cu distance is reduced to ∼3.4 Å (Fig. 3B).
The O–O distance of 1.9–2.0 Å (not restrained during refinements)
indicates a Cu2O2 geometry that lies between the bis-μ-oxo and the
μ-η2:η2-peroxo geometry, which exists in equilibrium. Interestingly,
the complex exhibits a “butterfly” distortion (distortion of the Cu–O–
O–Cu torsion angles). The Cu–O distances seem to slightly differ
between chain A and chain B. However, the resolution (∼1.8 Å;
Tables S1 and S2) of the obtained dataset is not high enough to
provide an unambiguous statement about the precise Cu–O distance.
The crystal structures of mature AUS1 (Fig. 3 C and D), pu-
rified from the natural source, displayed an unexpected electron
density at the CuB binding His252. This was particularly the case
in crystal structures from enzyme samples that exhibited a high
portion of sulfation or phosphorylation (24), making it impossi-
ble to place a histidine-coordinated copper atom into the density.
However, the density matched very well to the density of a
phosphohistidine or sulfohistidine. High-resolution mass spec-
trometric analysis revealed that the histidine residue was sulfated
(Fig. S4). Due to the sulfation, the histidine loses its ability to
participate in copper coordination. This leads to the loss of the
CuB atom (Fig. 3D) and to the loss of enzymatic activity of the
modified enzyme molecules.
Overall Structure of Active AUS1. Mature AUS1 (41.6 kDa) crystal-
lized in space groups P212121 and P1211, both with four monomers
in the asymmetric unit, yielding crystals of comparable quality (24)
(diffracting to ∼1.6 Å; Tables S1 and S2). The enzyme’s structure
possesses a very high structural similarity to the plant catechol
oxidases from Ipomoea batatas (18) [ibCO; Protein Data Bank
(PDB) ID code 1BT3; rmsd of 0.81 Å between 296 atom
pairs] and Vitis vinifera (17) (vvCO; PDB ID 2P3X; rmsd of 0.79 Å
between 284 atom pairs) as well as to the plant tyrosinase from
J. regia (23) (jrTYR; PDB ID code 5CE9; rmsd of 0.82 Å between
287 atom pairs). Mature AUS1 shares ∼47% sequence identity with
each of the active catechol oxidases and ∼48% with the active ty-
rosinase. Structural divergences to these PPOs are the unique fea-
tures (15) of AUS1 (Fig. 1A and Figs. S1 and S2A): the remaining
C-terminal peptide (Asp338 to Gly452)—connected to the catalyt-
ically active main core by a disulfide bond (Cys206–Cys445)—and
the insertion of four amino acids (V237ANG240) that form a cavity near
the active site. Both crystal forms possess homodimeric assemblies
of their monomers [analyzed with PISA (Proteins, Interfaces,
Structures and Assemblies)] (25), and a portion of mature AUS1
was found in a dimeric state by size exclusion chromatography
(Fig. S3), evidencing that the dimer exists in solution. The biological
assembly reveals a so far unknown role of the residual C-terminal
peptide, as it forms a main building block of the homodimeric in-
terface (Fig. 1B). The quaternary structure is stabilized by four
hydrogen bonds, 11 bridging water molecules, and several hydro-
phobic interactions between the residues of the C-terminal peptide
of one monomer and the α-helix of the second monomer that
carries the C-terminal peptide. These interactions result in a buried
surface area ranging from 1,523 to 2,323 Å2 and a free energy of
dissociation of the dimer ranging from 2.6 to 8.9 kcal/mol. The
active sites of the dimeric monomers are facing opposite directions,
due to their twofold rotational symmetry relation.
Reactivity of AUS1 and Insights into the Reaction Mechanism by MD
Simulations. Recently, we reported that AUS1 lacks monophenolase
activity (o-hydroxylation of monophenolic compounds) and displays
exclusively diphenolase activity (oxidation of diphenolic compounds
to the corresponding o-quinones) (15, 16). However, a modification
of the activity assay by adding ascorbic acid as a reducing agent
revealed that AUS1 in fact possesses hydroxylase activity toward
Overall Structure of Latent AUS1. Latent AUS1 was found to be
monomeric, and the overall structure of the main core of the
latent proenzyme is almost identical to the overall structure of
the mature enzyme (rmsd of 0.47 Å between 338 atom pairs),
except for the location of one loop, which gives way for the
shielding C-terminal domain (Fig. S2B). Interestingly, this isoliquiritigenin (Fig. S5 A and B) and that consecutive reactions
loop contains the characteristic insertion (V237ANG240) of the
group 2 PPOs (15). The C-terminal domain begins with a short
were responsible for the previously reported false-negative state-
ments. In the absence of a reducing agent, the highly reactive
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Fig. 1. Overall structures of latent and mature aurone synthase. (A) Top view of mature aurone synthase. The coloring was performed according to the
secondary structure features (α-helices, green; 310-helices, yellow; β-sheets, blue), and the characteristic features of AUS1 are colored red (loop carrying the
insertion V237ANG240) and magenta (residual C-terminal peptide). (B) Side view of the dimeric biological assembly of mature AUS1. The interacting residues
are colored orange (monomer chain B) and magenta (monomer chain D). (C) Overall structure of latent aurone synthase. The features of the C-terminal
domain are as follows: the three proteolytic cleavage sites, colored magenta and labeled as (1), (2), and (3); the linker region (orange) that connects the
catalytically active domain (gray) and the shielding C-terminal domain (green); and the residual peptide of the C-terminal domain (blue) and the active site
shielding Ile456 (plug residue) (red). (D) The active site is shielded by the C-terminal residue Ile456, which is responsible for the latency of the proenzyme and
functions like a plug [2Fo-Fc electron density map (blue mesh) contoured at 1.0 σ].
quinoid intermediates and products polymerize and are therefore
undetectable by HPLC analysis. Furthermore, the reaction product
sulfuretin has been reported to be a suicide substrate for AUS1
The chalcone isoliquiritigenin and the chalcone–aurone pairs of
butein and lanceoletin (corresponding aurones, sulfuretin and lep-
tosidin) naturally occur in Coreopsis species [summarized in Molitor
(15), for which the utilization of isoliquiritigenin strongly depended et al. (15)]. To validate obtained molecular docking results of butein to
on the amount of enzymes used in the activity assay. Additionally, AUS1, two different poses were subjected to MD simulations with the
the difference spectra of isoliquiritigenin differed between AUS1 met-form of AUS1. During the simulation of one docking pose, the
and abTYR (Fig. S5 A and B). However, in the presence of a re- reactive B-ring of the chalcone remained in the active site, whereas
ducing agent that traps the highly reactive quinoid intermediates,
the difference spectra show identical absorption changes, which is
evidence that the differences observed in the absence of a reducing
agent are related not to the hydroxylation reaction but to the
complex consecutive reactions. Furthermore, the enzyme showed
hydroxylation activity toward kaempferol (Fig. S5C). In contrast to
tyrosinases, AUS1 did not accept tyramine and tyrosine as sub-
strates (10 μg active AUS1; 2 mM tyramine in 1 mL total volume;
recordings up to 1 h; the experiments were performed with and
without the addition of hydrogen peroxide and ascorbic acid).
the A-ring was very flexible and interacted with several different resi-
dues. The other docking pose resulted in a stable substrate enzyme
complex, where the diphenolic substrate rapidly moved deeper into the
cavity, which is formed by the extended loop region (V237ANG240
)
(compare with Fig. 1A), and remained there relatively stably during the
simulation time. The substrate’s oxygen atoms are almost centered
between the two copper atoms (Fig. 4B and Fig. S6A). To mimic the
substrate position during the electron transfer, the simulation was re-
peated with restrained Cu–O–substrate bonds (26), resulting in a
considerably reduced fluctuation of the unreactive A-ring due to its
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lanceoletin and the monophenolic substrates isoliquiritigenin and
tyramine. The interactions of lanceoletin with AUS1 were almost
identical with those of butein, but with additional stabilization of
the methoxyl group by hydrogen bonds with Gly240 and hydro-
phobic interactions arising from Val244, Ala249, and Glu248.
Furthermore, the residues Gly271 and Asn271 form water bridges
with the carbonyl oxygen of the chalcone. Interestingly, when the
bridging water molecule is missing (water molecules in the simu-
lation box were placed randomly), the highly conserved residue
Glu248 is capable of directly coordinating one copper-coordinating
hydroxyl group of the substrate (Fig. 4C and Fig. S6B). This
residue has been proposed to participate in the water-mediated
deprotonation of substrates (17, 20, 23). However, the MD
simulations indicate that it might even be directly involved in
substrate deprotonation (Fig. 4C).
For the simulation of the monophenols isoliquiritigenin and
tyramine, the oxy-form of the enzyme was used. Even without a Cu–
O–substrate distance restraint, isoliquiritigenin remained very sta-
ble within the cavity, with identical enzyme–substrate interactions
as in the case of the diphenols butein and lanceoletin under re-
strained conditions. The distances of both copper atoms to the
reactive hydroxyl group ranged mainly between 3.5 and 4.5 Å (Fig.
4D and Fig. S6C). In contrast, tyramine rapidly left the binuclear
copper site of AUS1 (within 75 ps) due to attractive interactions
with Arg257 (Fig. 4E and Fig. S6D), which is in accordance with the
absence of monophenolase activity of AUS1 toward tyramine. The
characteristic insertion (V237ANG240) of AUS1 might additionally
interact with tyramine. To obtain more generalized information
about the role of Arg257, simulations of tyramine in complex with
jrTYR and with the corresponding L244R mutant of jrTYR were
Fig. 2. Interactions between the shielding C-terminal domain and the cat-
alytically active domain of latent AUS1. The latent structure is segmented by
its domains (left, catalytically active domain; right, shielding C-terminal do-
main), and their solvent-accessible surface is presented. The interdomain
interactions are located at two distinct areas: in the vicinity of the active site
(colored red) and in the vicinity of the linker region (colored green).
stronger interaction with protein residues of the cavity. Butein is
stabilized by hydrogen bonds with the residues Asn117, Asn239,
Arg257, and Gly240 (backbone); by hydrophobic interactions with
the residues Asn239, Val244, Ala249, Thr253, His256, Phe273,
and Ala276; and by residue Glu248 through a water bridge (Fig.
4B and the substrate binding residues are highlighted in Fig. S1).
The unrestraint but stable docking position of butein was then
taken as a basis for the simulation of the diphenolic substrate
Fig. 3. Copper binding site of AUS1. The 2Fo-Fc electron density map (blue mesh) is contoured at 1.0 σ, and the anomalous Fourier difference map in A, C,
and D (green mesh) is contoured at 3.0 σ. (A) Native met-copper center of recombinantly expressed AUS1. (B) Oxy-copper center of recombinantly expressed
AUS1 (μ-η2:η2-peroxo geometry; crystal soaked in H2O2). The oxygen atoms are slightly unsymmetrically bound to the copper atoms, and the μ-η2:η2 peroxo
complex displays a butterfly distortion. (C) Copper binding site of active AUS1 (occupancy CuB, ∼0.55). For clarity, the sulfohistidine (occupancy, ∼0.45) is not
shown. (D) Inactive AUS1 (occupancy sulfohistidine, 0.9).
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Fig. 4. Reactivity and in silico-obtained substrate–enzyme complexes of AUS1. (A) Reaction scheme of the hydroxylation of isoliquiritigenin in the presence
of ascorbic acid to trap highly reactive quinoid intermediates and products. (B) MD simulation snapshot of AUS1 in complex with butein. The reactive oxygen
atoms at the B-ring of butein (O3, O4) are almost symmetrically located between CuA and CuB. The enzyme–substrate interactions are visualized by magenta
lines (hydrogen bonding), yellow dashes (hydrophobic interactions), cyan dashed lines (π–π stacking), red dashes (cation–π interactions), and gray lines (water
bridges). (C) Superimposition of snapshots obtained from different MD simulations of AUS1 with lanceoletin as the substrate. Residue Glu248 coordinates
an oxygen atom of the A-ring through the formation of a water bridge (blue sticks; water bridges, gray lines) or one reactive oxygen atom (O4, B-ring)
directly (white sticks; hydrogen bonding, magenta lines). (D) MD simulation snapshot of AUS1 in complex with isoliquiritigenin. (E) Three snapshots [0 ps,
green (1); 44 ps, cyan (2); and 344 ps, magenta (3)] of the MD simulation of AUS1 with tyramine. Hydrophobic interactions of tyramine with Arg257 are
visualized by yellow dashes.
performed. Tyramine remained within the active site of jrTYR,
whereas it left the active site of the L244R mutant rapidly (Fig. S6E).
These results indicate a crucial role of the residues in this position
reveal that the main function of the interdomain linkage is the
retention of the residual peptide, as it is a main building block of
the homodimeric interface (Fig. 1B). The structure of the latent
(Fig. S1) for the acceptance or refusal of tyramie as a substrate for proenzyme exhibits two domains: the catalytically active domain
and the C-terminal domain that is responsible for the latency of
the proenzyme. The dominating structural motif of the C-terminal
domain is a jelly roll barrel. Similar motifs are also present in the
shielding domains of fungal (21, 27) (C-terminal), arthropod (28)
(N-terminal), and insect (29) (N-terminal) PPOs. This taxa-span-
ning structural homology might indicate another, so far unknown,
function of the shielding domains. Due to the interdomain disul-
fide linkage of the main core with the C-terminal domain, the
proteolytic activation of the latent enzyme has to occur at three
different sites (compare with Fig. S1) to result in the active en-
plant PPOs.
Exemplary video files of the simulation of AUS1 in complex
with butein (Movies S1–S3) are available.
Discussion
Overall Structures of Mature AUS1 and of the First Latent Plant PPO.
The most relevant differences to the plant PPO homologs ibCO (17),
vvCO (18), and jrTYR (23) are the characteristic four-amino-acid
insertion of the group 2 PPOs and the short residual C-terminal
peptide, connected to the main core by a disulfide bond. The in-
sertion forms a binding pocket for the natural substrates (chal- zyme (15). Two proteolytic cleavage sites are exposed on the
cones) of AUS1 (discussed below). The disulfide linkage was surface (1 and 2 in Fig. 1C). In contrast, the third is covered by the
proposed to stabilize the C-terminal domain in the latent pro-
C-terminal domain as well as by the main core but becomes
enzyme (15). However, the crystal structures of mature AUS1 accessible after the proteolytic cleavage at the first two cleavage
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sites. This suggests, and is supported by the purification of
“semicleaved” latent forms from the natural source (15), that the
proteolytic activation occurs stepwise and is targeted. The sep-
aration of the interdomain interactions into two distinct areas
(Fig. 2) reflects specific roles of each area. The interactions close
to the linker region will be responsible for the general structural
integrity of the proenzyme. This is supported by the fact that
the interdomain disulfide linkage is also part of this area. The
strength of the interactions near the active site will affect the
latency of the proenzyme. Furthermore, we propose that the in
vitro activation (acidic activation and SDS activation) (6) as well
as the allosteric activation (15) of latent plant PPOs are pri-
marily caused by a structural rearrangement in this region. The
transfer of the detailed knowledge of the interdomain interac-
tions (Fig. S1) to other plant PPO homologs will enable the
targeted modulation of the kinetic properties of recombinantly
expressed proenzymes (30).
The active site of fungal (21, 27), bacterial (31), insect (29),
and arthropod (28) pro-PPOs is blocked by a substrate mim-
icking the “placeholder” residue (tyrosine or phenylalanine,
originating from the shielding domain) and located within the
active site. However, due to the presence of the bulky gate res-
idue above the active site (phenylalanine, originating from the
main core), the shielding strategy in plant PPOs has to be dif-
ferent. The gate residue would have to swing away to make space
for the placeholder residue (compare with Figs. 4 and 5). As this
is energetically slightly unfavorable, other stabilizing interactions
would be necessary for compensation. The crystal structures of
the first latent plant pro-PPO reveal that plant PPOs overcome
this issue by a smaller hydrophobic plug residue (Ile456 in AUS1;
valine, leucine, and isoleucine in other plant PPOs) located
above the gate residue and outside of the active site (Fig. 1D).
iii) The purification has been performed independently two
times with approximately 1 y between them. The sulfohisti-
dine occupancy varied from ∼0.45 (first batch, 6 kg petal
tissue) to ∼0.9 (second batch, 9 kg petal tissue) and provides
hints that the modification is either caused by sulfated com-
pounds of petals of C. grandiflora or occurred in vivo.
Other sources of this modification might be a reaction of
AUS1 with sulfated compounds within the petals of C. grandi-
flora during the isolation and purification procedure or an in vivo
modification of AUS1 by a membrane-bound sulfotransferase.
The latter possibility is very unlikely, as a controlled in vivo PPO
inactivation has not been reported so far and we do not have any
hints that this occurred in petals of C. grandiflora. In contrast, fla-
vonoid sulfate esters are common in plants, especially in Aster-
aceae species (36). Four flavonol sulfotransferases were identified
in Flaveria chloraefolia and Flaveria bidentis and catalyze sequential
sulfation of quercetin to quercetin 3,7,3′,4′-tetrasulfate (37, 38).
Interestingly, it has been crystallographically evidenced that a direct
sulfuryl group transfer from p-nitrophenylsulfate to histidine, resulting
in an identical sulfohistidine as observed in AUS1, represents an
intermediate stage in the reaction mechanism of a bacterial sulfo-
transferase (39). This suggests that a direct sulfuryl transfer might also
have taken place in the case of AUS1, as several flavonoids (e.g.,
quercetin and fisetin) (15) represent substrates for PPOs. So far, the
preparation of samples of crude extracts (without the use of ammo-
nium sulfate) from petals of C. grandiflora, suitable for analysis by
HPLC–ESI–MS, failed due to the extremely high amount of pig-
ments and polyphenols that interfere with performed SDS/PAGE
analysis. However, the development of effective tyrosinase inhibitors
has become increasingly important in the cosmetic, medicinal, and
agricultural industries for application as antibrowning and depig-
menting agents, and the inactivation of a PPO by a sulfonated sub-
strate would have a high impact in this field of research.
Copper Binding Site of AUS1. The reactive oxy-form of AUS1 ex-
hibits a butterfly distortion (Fig. 3B) similar to the distortion found
in the oxy-form of Streptomyces castaneoglobisporus tyrosinase
(scTYR) (31). Because no indications of a distortion were found
in ligand field molecular dynamical simulations of the oxy-form of
tyrosinase (32), it has been suggested that the distortion in scTYR
might be caused by a hydrogen bonding interaction of the place-
holder residue (tyrosine) with the oxygen atoms. However, the
oxy-form of AUS1 indicates that the butterfly distortion is a gen-
eral characteristic of the oxy-form of PPOs, as AUS1 does not
possess a placeholder residue that might cause this distortion.
Irreversible inactivation of tyrosinase from Agaricus bisporus
(PPO3; abTYR) has been reported to occur by incubation with
sulfite ions (33). The authors evidenced the sulfation of a CuB-
coordinating histidine and speculated that the sulfation (ac-
companied by the loss of the CuB atom) presumably occurred at
His263 (His256 in AUS1). The crystal structure of AUS1, how-
ever, reveals that the sulfuryl group is transferred to the Ne atom
of His252 (His259 in PPO3; abTYR). The consistent discovery of
this inactivation in both PPOs might indicate an infiltration of
sulfite ions during the purification procedure of AUS1. However,
there exist several matters that evoke doubts about this:
Reactivity, Enzyme–Substrate Simulations, and Reaction Mechanism
of Plant PPOs. AUS1 is a catechol oxidase type of PPO, as no
reactivity toward the common tyrosinase substrates tyramine and
tyrosine was detectable. However, AUS1 showed monophenolase
activity toward its natural substrate isoliquiritigenin (chal-
cone) and the flavonol kaempferol (Fig. S5). A similar reactivity
has been reported for catechol oxidase from Aspergillus oryzae
(aoCO4) (40). This fungal PPO showed activity toward amino-
phenol and guaiacol but not toward tyrosine. Historically, the
definition that catechol oxidases lack the hydroxylase activity was
primarily based on the acceptance of common tyrosinase sub-
strates (e.g., p-cresol, tyrosine, tyramine) and not on the hy-
droxylation of their natural substrates (or at least a wider range
of potential substrates). Due to the evidenced hydroxylase ac-
tivity of AUS1 and aoCO4 (40), the question arises as to whether
this definition is still valid. There is no doubt that catechol oxi-
dases are incapable of hydroxylating tyramine or tyrosine, how-
ever AUS1 and aoCO4 reveal that they might hydroxylate
various other monophenols (e.g., their potential natural sub-
strates). Furthermore, the comparison of the crystal structures of
plant catechol oxidases (PDB ID codes 2P3X and 1BT3) with the
structure of a plant tyrosinase (PDB ID code 5CE9) did not
reveal relevant structural differences except in the potential
substrate binding residues (23). In this study, we identified a
substrate binding residue (by performing MD simulations, dis-
cussed below) that is able to stabilize or destabilize the tyrosinase
substrate tyramine within the active site of plant PPOs. This
supports the presumption that no general structural limitation
regarding the hydroxylase activity of plant PPOs exists, except the
prerequisite of a substrate stabilization in a way that a hydrox-
ylation reaction can occur. However, further investigations will
be necessary to prove a putative general hydroxylase activity of
PPOs. Therefore, we suggest using a substantially broader range
i) The chemicals used to purify AUS1 are not only commonly
used in protein purification, the identical chemicals were also
used in the purification of tyrosinases from J. regia (34) and
A. bisporus (35) and catechol oxidase from V. vinifera (24) in
either similar or even identical methods. Mass spectrometric
analyses [HPLC–ESI–MS (HPLC electrospray ionization mass
spectrometry) of digested protein samples and ESI–Q-TOF–
MS (electrospray ionization quadrupole time-of-flight mass
spectrometry) of intact protein samples] were performed
routinely, and none of the PPO samples exhibited any in-
dication of such a modification (34, 35).
ii) The latent form of AUS1 also did not display any modification.
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Fig. 5. Proposed catalytic reaction mechanism of plant PPOs. (A) Monophenolase cycle: The monophenolic substrate is guided to the binuclear copper site
[oxy-form, Cu(II)2O2, μ-η2:η2 peroxide complex] by displaced π–π interactions with the tilt out gate residue Phe273 and cation–π interaction with the CuB
binding histidine His256 (M1). When the substrate binds to the copper atoms, hydrophobic interactions with the substrate’s para-substituent become im-
portant additionally. The reactive substrate oxygen atom binds equally to both copper ions (M2), and the peroxide ligand is transferred to an inverse butterfly
distortion (ligand field molecular dynamical simulations, the Cu–O–O–Cu torsion angles of the peroxo ligand fluctuate by 20°) (32). Nucleophilic attack of
the Cu2O2 moiety by the substrate results either in an o-diphenolic product and the met-form [M4D, Cu(II)2OH] or in an o-quinone and the deoxy-form [M4Q,
Cu(I)2]. During product release, the gate residue swings back to its preferred position. Finally, oxygen uptake closes the catalytic cycle (M5). (B) Diphenolase
cycle: The principles of the diphenolase cycle are similar to the monophenolase cycle described in A. The reactive substrate oxygen atoms bind equally to both
copper ions of the met-form (D1, D2), and the o-quinone is released after electron transfer to the binuclear copper site resulting in the deoxy-form (D3).
Oxygen uptake results in the oxy-form (D4) and substrate binding results in an inverse butterfly distortion in the transition state (D5). The catalytic cycle is
closed by the release of an o-quinone resulting in the met-form of the binuclear copper site.
of compounds (including the class of flavonoids—e.g., chalcones, toward compounds that were previously reported to be tyrosinase
flavonols, flavanols) in activity assays for plant PPOs. Furthermore,
we suggest varying the conditions by adding reducing agents (to
avoid polymerization or enzyme inactivation) or hydrogen perox-
ide. The latter is useful not only in omitting the lag phase of the
hydroxylation reaction but also in detecting hydroxylase activity
inhibitors but in fact are hydroxylized and oxidized by tyrosinase
(41) [e.g., arbutin and isoeugenol (42), phloridzin and phloretin (43)].
The very specific interactions observed during the MD simu-
lations of AUS1 with the naturally occurring chalcones strongly
support the hypothesis that the crystallized enzyme represents a
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specialized enzyme involved in aurone biosynthesis (15, 16).
strates that in the long term the classification and the nomenclature
Furthermore, the proposed importance of the characteristic in- of plant PPOs will have to change to avoid misunderstandings.
As a consequence of the evidenced hydroxylation activity of
AUS1 toward its natural substrate in combination with the con-
sistent proposed novel reaction mechanism, another complexion is
put on the function of plant PPOs. So far, the role of plant PPOs
has primarily been assigned to enzymatic browning reactions, with
only few exceptions [e.g., aureusidin synthase (12, 13) and aurone
synthase (15, 16)]. However, we demonstrated that the mono-
phenolase activity of plant PPOs seems to depend only on whether
the monophenol can be stabilized within the active site or not.
There is no doubt that the hydroxylase activity represents the
primary functionality of tyrosinases (e.g., in the hydroxylation of
tyrosine in the melanin biosynthetic pathway). If catechol oxi-
dases also possess hydroxylase activity, it seems reasonable that
the above statement is also true for this type of enzyme. As a
consequence, it could be assumed that the catechol oxidases are
involved in anabolic pathways by hydroxylating their—so far
unknown—natural substrates. Hence, it can be concluded that
the identification of the few plant PPOs involved in the sec-
ondary metabolism does indeed represent just the “tip of the
iceberg” (7).
There is no doubt that there will be a long interdisciplinary
way to go (including metabolomics and transcriptomic and pro-
teomic analyses) until the role of plant PPOs is clarified, partic-
ularly with regard to their complex diversity. The substrate
binding residues (compare Fig. 4 and Fig. S1) around the CuA
and CuB binding sites differ considerably within the family of
PPOs (2, 3), indicating the occurrence of individual natural sub-
strates. Therefore, the upcoming challenges will be to cluster the
PPOs according to their substrate binding residues, to identify the
potential natural substrates (or substrate scaffolds), and to vali-
date the involvement of the corresponding PPOs in the plants’
secondary metabolism.
sertion of the group 2 PPOs in substrate binding (15) is confirmed
(Fig. 4B and Movies S1–S3). However, although the substrates are
strongly stabilized, it should be mentioned that the diphenols do
not “have” to bind within this binding pocket. PPOs accept an
enormous variety of diphenolic substrates (e.g., chlorogenic acid,
4-tert-butylcatechol, 3,5-di-tert-butylcatechol), suggesting that even
unspecific binding to the copper center can result in product
formation. Apart from the characteristic insertion of AUS1, the
involvement of several residues at the copper binding sites (Fig. 4B
and Fig. S1) in substrate coordination will be transferrable to
other plant PPOs.
It has been shown that in a bacterial tyrosinase (from Bacillus
megaterium; bmTYR) the substrates bind to the CuA atom, and
therefore, it was concluded that the monophenols will perform a
substrate rotation during the hydroxylation reaction (20). The
authors proposed that the substrates will bind to the same copper
atom in plant catechol oxidases. Based on these assumptions, it
was further concluded that the hydroxylation reaction is im-
possible in plant catechol oxidases, as the bulky gate residue in
combination with the rigidity of the CuA site (thioether bond) will
hinder the substrate rotation. However, the proposed mechanism
is incompatible with the hydroxylation activity of jrTYR (34,
44) as well as the observed hydroxylation activity of AUS1, as
both possess a bulky gate residue and a thioether bond. Fur-
thermore, no explanation for the lack of monophenolase activity
of aoCO4 toward tyrosine as a substrate (40) was given.
Naturally, classical MD simulations are not suitable to obtain
information about the geometry of the substrate–copper binding
scenario, as they do not take into account quantum mechanical
effects or the Jahn–Teller distortion. However, the strong hy-
drophobic interactions of the substrate with the gate residue
(Phe273) and His256 direct the B-ring to the active site, where the
substrate’s o-diphenolic oxygen atoms are almost equally positioned
between both copper ions (Fig. 4 B and C and Fig. S6). The posi-
tions of the oxygen atoms are in a way comparable with the position
of the butterfly distorted oxygen atoms of the oxy-form, demon-
strating the general possibility of this coordination. Further-
more, the monophenol isoliquiritigenin is stabilized in a position
that allows the hydroxylation in the ortho position to the copper-
coordinating hydroxyl group (Fig. 4D). These observations in
combination with the detected hydroxylation activity of AUS1
(Fig. S5) and jrTYR (23, 34, 45) led us to propose a novel
mechanism for the hydroxylation and oxidation of phenolic sub-
strates by plant PPOs (Fig. 5). In this mechanism, no substrate
rotation is required, as the substrate is coordinated equally to both
copper ions. This removes the contradiction of the previously
proposed mechanisms with the experimentally proven mono-
phenolase activities of plant PPOs. Furthermore, the MD simu-
lations of tyramine in the active site of AUS1, jrTYR, and the
L244R mutant of jrTYR strongly suggest that the residue at the
position of Arg257 (compare with Fig. S1) is one of the key res-
idues for the acceptance of tyramine as a substrate for plant PPOs.
It should be noted that the interactions will be different in fungal
and bacterial PPOs, as they do not possess a bulky gate residue
and the substrates are bound to the CuA atom (20). However,
during MD simulations the classical tyrosinase substrate tyramine
is destabilized in the active center of plant PPOs by an arginine in
this position (AUS1, L244R mutant of jrTYR), whereas it is sta-
bilized by a leucine in this position (jrTYR). It has been reported
recently that vvCO, possessing a lysine in this position, showed
weak activity toward tyrosine and tyramine, and vvCO was
therefore classified as a tyrosinase (46). However, a comparison of
the kinetic data of vvCO and jrTYR indicates that tyramine will
presumably not be the natural substrate of vvCO [Km(vvCO) =
7.7 mM (46); Km(jrTYR) = 0.274 mM (23)]. This example demon-
Methods
Protein Purification, Crystallization, Data Collection, Structure Determination,
and Refinement. The protein purification of active and latent AUS1 from
petals of C. grandiflora is described in Molitor et al. (15), and the purification
of recombinantly expressed AUS1 is described in Kaintz et al. (16). The
crystallization of the obtained enzyme samples and the data collection, us-
ing synchrotron radiation, of the obtained crystals have been described in
Molitor et al. (24). The corresponding data collections of the presented
crystal structures were performed using synchrotron radiation at 100 K (PDB ID
code 4Z0Y—P14, PETRA III, EMBL, DESY; λ = 1.23953 Å; PDB ID code 4Z0Z—I04-1,
DIAMOND; λ = 0.9173 Å; PDB 4Z11—ID23-1, ESRF; λ = 0.972499 Å; PDB 4Z12—
P11, PETRA III, DESY; λ = 1.0247 Å; PDB 4Z13—I04-1, DIAMOND; λ = 0.9173 Å).
Automated refinements of all models (obtained as described below) were
performed by phenix.refine from the PHENIX suite (47) followed by manual
structure completion and correction using COOT (48).
Initial phases for the active AUS1 form (space group P212121) were obtained
by molecular replacement using the BALBES webserver (49), which used the
crystal structure of catechol oxidase from I. batatas (PDB ID code 1BT3, se-
quence identity ∼47%) as a search model. The refined structure of AUS1 was
then used as a search model for structure determination of the remaining
obtained datasets by molecular replacement using PHASER (MR) from the
PHENIX program suite. The structure of the latent AUS1 form was obtained by
automated model building using AUTOBUILD from the PHENIX suite, after
placing the refined catalytically active main core with PHASER (MR) in the
asymmetric unit. Restraints for the polyoxometalate hexatungstotellurate (VI),
used as a cocrystallization agent for recombinantly expressed AUS1 (24, 50),
were obtained with phenix.reel and phenix.elbow as previously reported (27,
51, 52) using the crystal structure of hexatungstotellurate (VI) (53). The quality
of the models was validated by MOLPROBITY (54). All models were refined to
excellent stereochemistry without any Ramachandran outliers, and the re-
spective refinement statistics are presented in Tables S1 and S2.
The models were deposited in the PDB (www.rcsb.org) as ID codes 4Z0Y,
4Z0Z, 4Z11, 4Z12, and 4Z13.
Protein Digestion and HPLC–ESI–MS Measurements of Inactive AUS1. The ex-
periments were performed by the company Proteome Factory. The protein
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Molitor et al.

sample was denatured by incubation in 8 M urea for 30 min at room temperature.
For reduction, a final concentration of 5 mM Tris(2-carboxyethyl)phosphine
was added, followed by incubation for 20 min at room temperature. Then
iodoacetamide was added to a final concentration of 10 mM. After in-
cubation for another 20 min in the dark, the samples were diluted to 0.8 M
urea and subsequently digested by two endoproteases (trypsin and AspN)
with an enzyme:protein ratio of 1:50 (wt/wt) according to the Proteome
Factory’s protein digestion standard operating procedures. The acidified
peptide populations were applied to nanoLC–ESI–MS (LTQ-FT, Thermo Fin-
nigan) analyses using a 60-min nanoLC gradient (Agilent 1100 nanoLC system)
with solvent A [0.1% (vol/vol) formic acid in 5% (vol/vol) acetonitrile] and
solvent B [0.1% (vol/vol) formic acid in 99.9% acetonitrile].
The mass accuracy was better than 3 ppm for MS data and 0.02 Da for
MS2 data. MS2 data analyses were done using MASCOT (Matrix Science),
whereas the MS data were analyzed using Qualbrowser (Thermo Finnigan).
The induced modification carbamidomethylation (C) was allowed during
MS2 data searches, along with possible modifications such as oxidation (M),
deamidation (NQ), and phosphorylation/sulfation (H). The precursor show-
ing the predicted modification and a strong neutral loss signal was subject to
dedicated MS3 experiments.
box (10 Å edge distance) of TIP3PBOX water molecules. The charge of the
system was neutralized by replacing water molecules by the respective num-
ber of sodium ions. Energy minimization was performed with the steepest step
algorithm with a maximum of 3,000 steps and an energy tolerance of 1,000
kJ·mol−1·nm−1. After convergence, the system was equilibrated in two phases:
NVT ensemble (isothermal-isochoric: constant number of particles, volume, and
temperature) and NPT ensemble (isothermal-isobaric: constant number of
particles, pressure, and temperature), each for 100 ps at 300 K. MD simulations
were carried out by applying periodic boundary conditions. The LINCS (Linear
Constraint Solver) algorithm was used to constrain all bond lengths, and the
particle-mesh Ewald method was used to calculate long-range electrostatic
interactions. The Berendsen coupling algorithm was used for temperature, and
the Parrinello–Rahman coupling algorithm was used for pressure control.
After the two equilibration runs, a 5-ns simulation was carried out with,
unless otherwise stated, no position restraints.
Structural Analysis and Graphical Presentation. The quaternary structure of
the obtained crystal structures was analyzed with PISA (Proteins, Interfaces,
Structures and Assemblies) (25). Protein interfaces and ligand interactions
were analyzed and visualized with LigPlot+ (61) and PLIP (Protein–Ligand
Interaction Profiler) (62). Molecular graphic images and movies were gen-
erated with PyMOL (www.pymol.org/) and VMD (Visual Molecular Dynamics)
(63), respectively. The secondary structure features were assigned using DSSP
(Dictionary of Secondary Structures of Proteins) (64).
Molecular Docking and MD Simulations. The structures of active AUS1 (PDB ID
code 4Z0Y) and jrTYR (PDB ID code 5CE9) were prepared for molecular
docking studies and MD simulations by the addition of missing side chain
atoms using COOT and the removal of the residual C-terminal peptide of
AUS1. Molecular docking of butein was performed with AutoDock VINA (55)
by the use of the deoxy-form of active AUS1 and setting the gate residue
PHE273 as a flexible residue. The search exhaustiveness was set to the maximal
value of 2,000 (default 8). MD simulations of obtained enzyme–substrate
complexes were performed with GROMACS (56) version 4.6.7 package apply-
ing the AMBER99SB force field (57). The ligand topologies were generated
using the SWISSPARAM webserver (58). RESP (restrained electrostatic poten-
tial) charges were obtained from the RED webserver (59) by applying the HF/6–
31G* level of theory (Gaussian09) to calculate the electrostatic potential. For
the simulation of o-diphenols in complex with AUS1, the met-form was used
and new residues were defined for the copper atoms, the copper-coordinating
histidines, and the thioether forming cysteine, according to Bochot et al. (60).
For the simulation of monophenols in complex with AUS1, the oxy-form was
used and parameters of the dinuclear copper site were adopted from Deeth
and Diedrich (32). The protein–ligand complex was solvated in a dodecahedral
ACKNOWLEDGMENTS. We thank Kristina Djinovic-Carugo and Georg Mlynek
[Max F. Perutz Laboratories (MFPL), Vienna Biocenter] for their kind support
during the early stages of this research. We thank beamline scientists Elspeth
Gordon (ESRF ID23-1, mx1450), Anja Burkhardt (DESY P11, I-20120633 EC), and
Alice Douangamath (Diamond Light Source I04-1, MX8476) for their generous
support during the allocated beam times. We give special thanks to Gleb Bour-
enkov and Victor S. Lamzin (DESY/EMBL) for the opportunity of data collection
at beamline P14 during “European School for Macromolecular Crystallography
(ESMAX) 2012.” For cultivating C. grandiflora and for taking care of the plant
fields, we thank the Horticultural Department of Molecular Systems Biology,
UZA1-Glashaus1, University of Vienna—in particular Thomas Joch and Andreas
Schröfl—and the gardeners of the experimental garden Augarten—especially
Miroslav Crep and Erich Wagner. We also thank Aleksandar Bijelic, Matthias
Pretzler, and Ioannis Kampatsikas for valuable discussions concerning this
work. The research was funded by the Austrian Science Fund (FWF): P25217
and the Deutsche Forschungsgemeinschaft (Ro 1084/8-1).
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