Structure and Spectroscopy of Alkene-Cleaving Dioxygenases Containing an Atypically Coordinated Non-Heme Iron Center

Article abstract of DOI:10.1021/acs.biochem.7b00251

Carotenoid cleavage oxygenases (CCOs) are non-heme iron enzymes that catalyze scission of alkene groups in carotenoids and stilbenoids to form biologically important products. CCOs possess a rare four-His iron center whose resting-state structure and inte

Full text of DOI:10.1021/acs.biochem.7b00251

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Article
Structure and spectroscopy of alkene-cleaving dioxygenases
containing an atypically coordinated non-heme iron center
Xuewu Sui, Andrew C. Weitz, Erik R. Farquhar, Mohsen Badiee, Surajit Banerjee, Johannes
von Lintig, Gregory Tochtrop, Krzysztof Palczewski, Michael P. Hendrich, and Philip D Kiser
Biochemistry, Just Accepted Manuscript • Publication Date (Web): 11 May 2017
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Structure and spectroscopy of alkeneꢀcleaving
dioxygenases containing an atypically coordinated
nonꢀheme iron center
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‡
2‡
3,4‡
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Xuewu Sui , Andrew C. Weitz , Erik R. Farquhar , Mohsen Badiee , Surajit Banerjee ,
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1,7
2*
Johannes von Lintig , Gregory P. Tochtrop , Krzysztof Palczewski , Michael P. Hendrich and
1,8*
Philip D. Kiser
1
Department of Pharmacology, School of Medicine, Case Western Reserve University, 10900
Euclid Ave., Cleveland, OH 44106, USA
2
Department of Chemistry, Carnegie Mellon University, 4400 Fifth Ave., Pittsburgh, PA 15213,
USA
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National Synchrotron Light SourceꢀII, Brookhaven National Laboratory, Upton, NY 11973,
USA
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Center for Proteomics and Bioinformatics, Center for Synchrotron Biosciences, School of
Medicine, Case Western Reserve University, 10900 Euclid Ave., Cleveland, Ohio 44106–4988,
USA
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Department of Chemistry, Case Western Reserve University, 2080 Adelbert Rd., Cleveland,
OH, 44106, USA
Department of Chemistry and Chemical Biology, Cornell University, Ithaca, NY 14850, and
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Northeastern Collaborative Access Team, Argonne National Laboratory, Argonne, IL 60439,
USA
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Cleveland Center for Membrane and Structural Biology, Case Western Reserve University,
st
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819 E 101 Street, Cleveland, OH 44106, USA
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Research Service, Louis Stokes Cleveland VA Medical Center, 10701 East Boulevard,
Cleveland, OH 44106, USA
KEYWORDS Carotenoid cleavage oxygenase, retinal, oxygenase, resveratrol, crystallography,
dioxygen, Mössbauer spectroscopy, nitric oxide, electron paramagnetic resonance
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ABSTRACT: Carotenoid cleavage oxygenases (CCOs) are nonꢀheme iron enzymes that
catalyze scission of alkene groups in carotenoids and stilbenoids to form biologically important
products. CCOs possess a rare 4ꢀHis iron center whose restingꢀstate structure and interaction
with substrates are incompletely understood. Here, we address this knowledge gap through a
comprehensive structural and spectroscopic study of three phyletically diverse CCOs. The crystal
structure of a fungal stilbenoidꢀcleaving CCO, CAO1, reveals strong similarity between its iron
center and those of carotenoidꢀcleaving CCOs, but with a markedly different substrateꢀbinding
cleft. These enzymes all possess a 5ꢀcoordinate highꢀspin Fe(II) center with resting state FeꢀHis
bond lengths of ~2.15 Å. This ligand set generates a more electropositive iron environment
relative to other nonꢀheme iron dioxygenases as observed by Mössbauer isomer shifts. Dioxygen
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(O ) does not coordinate iron in the absence of substrate. Substrates bind away (~4.7 Å) from the
2
iron and have little impact on its electronic structure thus excluding coordinationꢀtriggered O2
binding. However, substrate binding does perturb the spectral properties of CCO FeꢀNO
derivatives indicating proximate organic substrate and O binding sites, which might influence
2
FeꢀO interactions. Together, these data provide a robust description of the CCO iron center and
2
its interactions with substrates and substrate mimetics that illuminates commonalities as well as
subtle and profound structural differences within the CCO family.
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INTRODUCTION
CCOs constitute a family of nonꢀheme iron enzymes, present in all life domains, that typically
catalyze oxidative cleavage of carotenoid substrates (Fig. 1A and B). This activity is critical for a
number of biological processes including vitamin metabolism, hormone production and
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generation of retinaldehyde chromophore for visual and nonꢀvisual light detection . The CCO
family also encompasses enzymes expressed in bacteria and fungi that catalyze cleavage of
Figure 1. Phylogeny and enzymatic activities of carotenoid and stilbenoidꢀcleaving CCOs. (A) Maximum likelihood
phylogeny of biochemically characterized stilbenoidꢀcleaving CCOs (azure box) and two cyanobacterial apocarotenoid
cleavage enzymes used as an outgroup to root the tree (grey box). Proteins in red were examined in the present study. The
scale bar represents the average number of amino acid substitutions over the indicated branch length. Numbers on the
branches are bootstrap percentages from 2000 pseudoreplicates. (B) Carotenoid cleavage reaction catalyzed by ACO. (C)
Stilbenoid cleavage reaction catalyzed by CAO1 and NOV2. The red wavy line indicates the position of the scissile bond in
7ꢀ9
stilbenoids such as resveratrol (Fig. 1A and C) . Presently, the physiological functions of
stilbenoidꢀcleaving CCOs are poorly understood. Retinal pigment epithelium 65 kDa protein
(RPE65) is a vertebrate CCO that exhibits retinoid isomerase rather than alkene cleavage
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activity, which is necessary for visual cycle function .
The diversity of important biological processes in which alkeneꢀcleaving CCOs are involved
has prompted interest in characterizing their enzymatic properties including substrate specificity
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and mode of catalysis. CCOs catalyzing cleavage of monoꢀ, biꢀ and acyclic carotenoids as well
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, 5, 8, 9, 11, 12
as various stilbenoids have been identified, e.g.
. Despite this variability in substrate
specificity, individual CCO enzymes are often highly stereoꢀ and regioꢀselective. Crystal
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structures of apocarotenoid oxygenase from Synechocystis (ACO) , RPE65 from Bos taurus
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and 9ꢀcisꢀepoxy carotenoid dioxygenase from Zea mays, viviparousꢀ14 (VP14) have revealed
plasticity in the CCO substrateꢀbinding cavity that has resulted from the need to accommodate
the diverse substrates of these enzymes and control active site accessibility.
In contrast to this substrate and substrateꢀbinding cleft diversity, all CCOs contain a conserved
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metal center consisting of an iron cofactor coordinated by 4ꢀHis residues
. Two cisꢀlocalized
sites in the iron coordination sphere remain unoccupied by protein ligands and are potentially
available for ligand binding, although one is rendered inaccessible to many ligands by the
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presence of a nearby Val or Thr residue in most CCOs
. Three conserved and functionally
critical Glu residues serve as a second coordination sphere for the iron cofactor by forming
hydrogen bonding interactions with the His residues. This mode of iron coordination is rare and
differs from that of the wellꢀcharacterized facialꢀtriad nonꢀheme iron dioxygenases wherein iron
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is bound by a 2ꢀhistidine,1ꢀcarboxylate (2Hꢀ1C) motif with three sites open for ligand binding
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Because CCOs require Fe(II) for their catalytic activity, it is assumed that the iron center
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mediates dioxygen (O ) activation as opposed to substrate activation . CCOs were previously
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thought to function as monooxygenases, but recent data obtained using improved assay systems
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suggest that dioxygenase activity is common to the entire family
. The carotenoid and
stilbenoid substrates of CCOs are electronꢀrich, redoxꢀactive molecules that provide all reducing
equivalents needed for their oxidative cleavage. However, little is known about the specific
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mechanisms of O activation by CCOs. Crystal structures of VP14 and Novosphingobium
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aromaticivorans oxygenase (NOV) 1 have suggested that the iron center can form stable adducts
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with O in the absence of organic substrate
. This contrasts with the gating mechanism of O2
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activation by oxygenases wherein organic substrate and O binding are tightly coupled to avoid
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generation of detrimental reactive oxygen species . In the 2Hꢀ1C dioxygenases this
synchronization is often accomplished by direct coordination of organic substrate to the iron
center, which lowers the iron redox potential, enables electron transfer between substrate and O2,
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and positions the scissile bond for cleavage
. In the case of pterinꢀdependent
dioxygenases, the ironꢀO interaction is facilitated by organic cofactor binding near but not in
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direct coordination with the Fe(II) ion
. As typical CCO substrates lack functionalities for
metal coordination (Fig. 1B and C), it is clear that an alternative mechanism of O activation
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must be used by these enzymes.
Further progress in elucidating the CCO alkene cleavage mechanism has been slowed by an
incomplete understanding of the CCO nonꢀheme iron center and its interaction with substrates.
Studies addressing this knowledge deficit were not previously feasible due to a lack of suitable
highꢀyield expression systems for these enzymes. Recently, we developed methods for producing
quantities of ACO, a prototypical carotenoidꢀcleaving CCO, sufficient for structural and
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spectroscopic studies . These methods have also been applicable to a stilbenoidꢀcleaving CCO,
which we anticipated would be more amenable to structural and spectroscopic characterization
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due to the greater aqueous solubility of its substrates relative to that of carotenoids
.
Carotenoids and stilbenoids are both redoxꢀactive polyenes whose CCOꢀcatalyzed cleavage
reactions are closely analogous, both occurring at nonꢀaromatic alkene groups (Fig. 1B and C).
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Therefore, information gleaned from stilbenoidꢀcleaving CCOs could be applicable to the
broader group of alkeneꢀcleaving CCOs.
Here, we characterize the 4ꢀHis iron centers of ACO and two stilbenoidꢀcleaving CCOs,
NOV2 and Neurospora crassa carotenoid oxygenase 1 (CAO1) using a combination of Xꢀray
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crystallography, Xꢀray absorption spectroscopy (XAS) and FeꢀMössbauer and electron
paramagnetic resonance (EPR) spectroscopies. The data allow detailed comparisons of these
enzymes in terms of their substrate binding clefts, iron center resting state structures and
interactions with substrates and substrate analogs. Synthesis of these high-resolution data provide
new insights into catalysis by CCOs as well as their biological functions.
MATERIALS AND METHODS
Reagents. Except as noted below, chemical reagents were purchased from SigmaꢀAldrich (St.
Louis, MO) in the highest purity form available. LB media was purchased from USB (Cleveland,
OH). Ironꢀ57 metal (95.93% isotope enrichment) was obtained from Cambridge Isotope
Laboratories, Inc. (Andover, MA). Water from a MilliꢀQ purification system (resistivity = 18.2
mꢁ × cm) (ED Millipore, Billerica, MA) was used to prepare all reagents and buffer solutions.
Synthesis of β-fluororesveratrol
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βꢀfluororesveratrol was prepared as described previously
. Further purity was acquired by
o
prepꢀHPLC [Luna® 15 µm PREP silica (2) 100 A column, nꢀhexanes/ethyl acetate 75:25, 20
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mL/min, 280 nm; Retention time: 46.0 min (Zꢀisomer)]. H NMR (500 MHz, Acetoneꢀd ) δ 8.67
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(
s, 1H), 8.20 (s, 2H), 7.55 (d, J = 7.8 Hz, 2H), 6.91 (d, J = 8.3 Hz, 2H), 6.68 (s, 2H), 6.28 (s, 1H),
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6.24 (d, J = 41.5 Hz, 1H). F NMR (470 MHz, Acetoneꢀd ) δ ꢀ114.24 (d, J = 41.3 Hz). C NMR
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(
125 MHz, Acetoneꢀd ) δ 159.45 (s), 158.30 (d, J = 256.2 Hz), 136.79 (d, J = 2.6 Hz),
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26.84 (d, J = 7.7 Hz), 125.20 (d, J = 28.4 Hz), 116.51 (d, J = 2.2 Hz), 108.35 (d, J =
CF CF CF CF
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.1 Hz), 104.71 (d, J = 9.6 Hz), 102.69 (d, J = 1.9 Hz)
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CF
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Scheme 1. Synthetic route for production of βꢀfluororesveratrol
Protein production
Cloning, expression and purification of CAO1. The coding sequence of CAO1 (GI:759001907)
from Neurospora carssa OR74A was synthesized (GenScript, Piscataway, NJ) and cloned into
the pET3a expression vector without fusion tags. The integrity of the expression plasmid was
confirmed by sequencing.
The plasmid was transformed into the T7 express BL21 Escherichia coli strain (New England
Biolabs, Ipswich, MA) for protein expression. Oneꢀliter cultures of LB media containing 100 ꢂg
of ampicillin/mL were grown at 37 °C to an A600 nm of 0.5 – 0.6 at which time protein expression
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was induced by the addition of isopropyl βꢀDꢀ1ꢀthiogalactopyranoside (IPTG) (Roche Applied
Science, Indianapolis, IN) to a final concentration of 0.1 mM. The temperature was lowered to
28 °C and the culture was supplemented with an additional 100 ꢀg/mL of ampicillin. After
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overnight growth (12–14 h) at 230 rpm, cells were harvested by centrifugation, suspended in 20
mM TrisꢀCl, pH 8.5, and either stored at −80 °C or used immediately.
All purification procedures were conducted at 4 °C. Harvested cells were lysed by two passes
through a French press. The lysate was clarified by centrifugation at 186,000g for 30 min. The
supernatant was then loaded onto a HiTrap Q HP 5 mL anionꢀexchange column (GE Healthcare,
Chicago, IL) equilibrated with buffer A consisting of 20 mM HEPESꢀNaOH, pH 7.0. The
column was then washed with 60 mL buffer A, followed by linear increase in buffer B
containing 20 mM HEPESꢀNaOH, pH 7, and 1 M NaCl. Fractions containing CAO1 were
pooled, concentrated, and further purified by gel filtration chromatography. Specifically, the
sample was injected into a 120ꢀml Superdex 200 gel filtration column (GE Healthcare)
equilibrated with gel filtration buffer consisting of 20 mM HEPESꢀNaOH, pH 7.0 and 200 mM
NaCl. Fractions containing purified CAO1 (≥90% pure as judged by SDSꢀPAGE analysis) were
pooled, concentrated to 20 mg/ml, flash frozen and stored in liquid nitrogen or placed on ice for
immediate use.
Production of cobaltꢀsubstituted CAO1 and FeꢀCAO1 in minimal media. Cobaltꢀsubstituted
CAO1 (CoꢀCAO1) and FeꢀCAO1 were produced by cultivating E. coli cultures in M9 minimal
media (1X M9 salts, 2 mM MgSO , 0.4% w/v glucose, and 0.1 mM CaCl ) in the presence of
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cobalt or iron. E. coli colonies freshly transformed with the pET3aꢀCAO1 plasmid were grown
on an LB agar plate containing 100 ꢀg/mL ampicillin and inoculated into 2 mL of LB media
containing the same concentration of antibiotic and grown for ~ 5 h at 37 °C. Cells were
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collected by centrifugation at 3,220g for 15 min at 37 °C. Cell pellets were then suspended in 1
mL of M9 minimal media and centrifuged again to remove residual LB media. The cell pellet
was then suspended in 1 mL minimal media and transferred into 1 L of M9 minimal media
containing 100 ꢀg/mL ampicillin. The cell culture was grown at 37 °C with 230 rpm shaking to
an A600 nm of 0.5–0.6 at which time IPTG was added to a final concentration of 0.1 mM to induce
protein expression at 28 °C. Simultaneously, 15 or 30 mg of CoCl ·6H O or
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(
NH ) Fe(SO ) ·6H O was added to the culture, the former quantity for activity studies and the
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latter for crystallization studies. After overnight growth, cells were harvested, and protein
purification was conducted as described for the native protein from LB culture. All glassware
and protein purification instrumentation were washed with MilliꢀQ water containing 1 mM
EDTA to reduce adventitious metal contamination. The purity of CAO1 expressed in minimal
media was similar to that of native protein expressed in LB culture.
Expression and purification of ACO and NOV2 – Synechocystis ACO and Novosphingobium
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NOV2 were expressed and purified as previously described
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Expression and purification of Fe labeled CAO1, NOV2 and ACO – E. coli cells (T7 express
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BL21 strain, New England Biolabs) harboring the pET3aꢀNOV2 or pET3aꢀACO plasmids
were grown for 3 h at 37 ºC in LB media containing 100 ꢀg/mL of ampicillin. Cells were
pelleted by centrifugation at 37 ºC and used to inoculate 2 L of M9 minimal media containing
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.16% v/v glycerol, 100 ꢀg/mL ampicillin and 5 mg of Fe (delivered as a 25 mg/mL solution in
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N HCl). The culture was grown at 37 ºC for 9 h at which time the culture optical density at 600
nm was ~0.6ꢀ0.8. Protein expression then was induced by the addition of IPTG to a final
concentration of 100 ꢀM, and the culture was supplemented with an additional 100 mg/mL
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ampicillin and 5 mg of Fe. Protein expression was conducted overnight at 28 ºC with 235 RPM
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shaking. Cells were pelleted by centrifugation the next morning and suspended in 50 mL of 20
mM BisꢀTrisꢀHCl, pH 7 in the case of ACO and 20 mM TrisꢀHCl, pH 7.5 in the case of CAO1
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and NOV2. Protein purification was conducted either as previously described
described above for CAO1.
or as
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Protein crystallization and formation of enzyme-substrate complexes
Crystallization of both FeꢀCAO1 and CoꢀCAO1 was performed by the hanging drop vapor
diffusion method by mixing purified protein sample at 20 mg/mL with a crystallization cocktail
containing 42ꢀ43% (w/v) sodium polyacrylate 2100 and 0.1 M HEPESꢀNaOH, pH 6.5 in a 1 : 1
ratio at room temperature and incubating the drops over 0.5 mL of the same crystallization
solution at 8 °C. Rodꢀshaped crystals, hexagonal in crossꢀsection, appeared within 1ꢀ2 weeks and
continued to grow to full size within another 1ꢀ2 weeks. Crystals of FeꢀCAO1 in complex with
βꢀfluororesveratrol were prepared by addition of a 200 mM DMSO stock solution of the purified
protein samples to a final concentration of 3 mM followed by an overnight incubation at 4 °C.
Samples were centrifuged at 71,680g for 15 min to remove any aggregates prior to initiation of
crystallization trails. Crystals of CoꢀCAO1 in complex with resveratrol and piceatannol were
obtained in a similar manner except that the crystals were soaked in synthetic mother liquor
containing 10 mM of the target ligand overnight at 8 °C prior to harvesting to ensure full ligand
occupancy. Mature crystals were harvested using Mitegen dual thickness loops (Mitegen, Ithaca,
NY) and flashꢀcooled in liquid nitrogen before Xꢀray exposure.
Diffraction data collection, phasing, and structural refinement
Data collection and processing. Diffraction data sets were collected at the NEꢀCAT 24ꢀIDꢀC/E
beamlines of the Advanced Photon Source and beamline 12ꢀ2 of the Stanford Synchrotron
Radiation Lightsource. Data for cobaltꢀsubstituted CAO1 crystals were collected at Xꢀray
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energies above and below the cobalt Kꢀedge based on an Xꢀray fluorescence scan to assess the
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active site cobalt occupancy. Data reduction was conducted with XDS (Table S1).
Phasing and structure refinement. The structure of CAO1 was solved by molecular
replacement augmented by single wavelength anomalous diffraction data (MRꢀSAD) and density
modification. A 2.16 Å resolution data set from a ligandꢀfree FeꢀCAO1 crystal (Table S1) was
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submitted to the MrBUMP molecular replacement pipeline
through the CCP4 Online web
server. The top solution in space group P3 21 with four monomers in the asymmetric unit was
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identified with a Sculptorꢀmodified version of Synechocystis apocarotenoid oxygenase ACO
35
(
PDB accession code: 4OU9) in the program PhaserMR . The solution featured a final
translational Zꢀscore of 21.5, firmly indicating its correctness, but was not readily amenable to
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direct refinement efforts with an Rfree after REFMAC refinement of 48.7% . This MR solution
was then applied as a partial model in the PhaserEP phasing pipeline by using a high multiplicity
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FeꢀCAO1 data set collected just above the iron Kꢀedge . Four iron atoms and four sulfur atoms
were located in the Phaser combined realꢀimaginary log likelihood gradient maps. The MRꢀSAD
38
phases were then improved by solvent flipping in the program SOLOMON . The coordinates of
the MR solution were then refined against the iron SAD data set in REFMAC using experimental
phase restraints giving an Rfree value of 43.8%. The updated coordinates were then fed back into
PhaserEP for a second round of anomalous scatterer identification and SAD phasing, followed
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by density modification in Parrot . The coordinates and updated phases were used for
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automated model building in Buccaneer with interspersed cycles of REFMAC refinement with
HendricksonꢀLattman coefficients used as phase restraints, resulting in an Rfree of 34%. The
updated model was then refined against the 2.16 Å resolution data set used originally for
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molecular replacement resulting in an Rfree of 24.9%. ARP/wARP was then used to correct and
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extend several regions of the model resulting in an Rfree of 21.8%. The model was further
improved by multiple rounds of refinement in REFMAC using local NCS restraints together with
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manual adjustments performed in COOT
based on difference map searches and
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MOLPROBITY geometry analyses. Metal ion temperature factors were refined anisotropically
in REFMAC. No distance restraints were applied to the metalꢀligand bond lengths during
refinement. Subsequent structures were solved either by direct refinement or by MR. Residues 1ꢀ
2
6 or 1ꢀ30, depending on the subunit and specific structure, were not resolved in the electron
density maps and hence were omitted. Identification of anomalous scatterers was accomplished
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through the use of imaginary LLG maps produced by Phaser . Residues Phe91 and Lys164
were harmonically restrained in the unliganded structures to prevent the side chains from being
pulled into the unidentified active site electron density found in these structures. Atomic
coordinates and geometric restraints for the ligands were generated using the GRADE server
(
Global Phasing Ltd.). Atomic models were validated using Molprobity and the wwPDB
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structure validation server . A summary of the Xꢀray data collection and refinement statistics
are shown in Table S1. All structure figures were prepared with PyMOL (Schrödinger).
CCO multiple sequence alignment
Protein sequences were retrieved from the NCBI protein database. A structureꢀbased amino
acid sequence alignment of CAO1 and ACO was generated with the DaliLite server using the
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pairwise option . Amino acid sequence alignment for the other resveratrolꢀcleaving CCOs was
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performed with MUSCLE . These two multiple sequence alignments and the NsACO sequence
were successively aligned in MUSCLE using the profile option.
Phylogenetic analysis
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Protein sequences for Synechocystis sp. PCC 6803 ACO (P74334.1), Nostoc sp. 7120 ACO
BAB75983.1) Ustilago maydis RCO1 (UM05084), Botryotinia fuckeliana RCO1
XP_001548426), Chaetomium globosum RCO1 (XP_001219451), Neurospora carassa CAO1
NCU07008), Aspergillus fumigatus RCO1 (XP_746307), Novosphingobium aromaticivorins
(
(
(
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NOV2 (YP_498079), Pseudomonas nitroreducens isoeugenol monooxygenase (IEM)
ACP17973.1), Novosphingobium aromaticivorins NOV1 (YP_496081), Sphingomonas
(
paucimobilis lignostilbene dioxygenase isoenzyme 1 (LSD1) (AAC60447) and Sphingomonas
paucimobilis lignostilbene dioxygenase isoenzyme 3 (LSD3) (AAB35856) were retrieved from
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the NCBI protein database and aligned with Clustal Omega . Phylogeny inference was
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performed with PhyML
using the LG substitution matrix with empirical equilibrium
frequencies along with the estimated proportion of invariant sites and gamma distribution
parameter (six categories). The starting neighborꢀjoining tree topology was improved by surface
pruning and regrafting. Branch lengths also were optimized. Tree robustness was assessed with
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000 bootstrap pseudoꢀreplicates. The maximum likelihood tree had a logꢀlikelihood value of ꢀ
2,232 with good bootstrap support for most branches (Fig. 1A). The tree was rooted with the
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Synechocystis and Nostoc ACO sequences used as outgroups. Visualization of the tree was
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performed with the Interactive Tree of Life (iTOL) v3 .
Enzyme kinetics
Stilbenoid cleavage activity assay. The enzymatic activity of CAO1 and NOV2 were assessed
17
as previously described using either resveratrol or piceatannol as substrates . For both
substrates, absorbance at 305 nm was monitored over time using a FlexStation 3 plate reader.
Activity levels were determined using the initial linear regions of each reaction progress curve.
Determination of enzyme transition metal content and enzyme-metal stoichiometry
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The transition metal content of the enzyme samples was measured by inductively coupled
plasma optical emission spectroscopy (Soil Research Analytical Laboratory, University of
Minnesota). Samples for ICPꢀOES analysis were prepared by digesting each enzyme sample in
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% v/v HNO . The proteins precipitated immediately upon addition of HNO and the samples
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were incubated for 3 h at room temperature with gentle rocking. Protein precipitates were then
removed by filtration through a 0.22 ꢀm filter (Millipore) and the aqueous solutions containing
dissociated metals were analyzed by ICPꢀOES. The buffer solution was treated identically to
determine the background metal content, which was below the limits of detection for all metals
of interest. Amino acid analysis of the purified enzyme samples was conducted to enable
accurate determination of the enzyme concentrations (Texas A&M University, Protein
Chemistry Laboratory). The molar absorptivities of ACO, NOV2 and CAO1 at 280 nm in units
ꢀ1
ꢀ1
of M ·cm were 75,248, 69,786 and 64,924, respectively.
X-ray absorption spectroscopy.
Sample Preparation. Samples of ACO at ~ 58 mg/mL (equal to ~ 1.2 mM) in 50 mM Trisꢀ
HCl, pH 8, 200 mM NaCl, 1 mM DTT and 20% glycerol, NOV2 at ~ 97 mg/mL (1.7 mM) in 20
mM TrisꢀHCl, pH 8, 200 mM NaCl and 20% v/v glycerol and CAO1 at ~ 95 mg/mL (1.2 mM) in
16 mM HEPESꢀNaOH, pH 7, 160 mM NaCl and 20% v/v glycerol were used for XAS
experiments. Samples were flash cooled in liquid nitrogen and stored in dry ice before use.
Samples were loaded into copper sample cells (NSLS) or Delrin cuvettes (SSRL) wrapped in
Kapton tape and flash frozen in liquid nitrogen immediately prior to XAS measurements.
Data collection. XAS data were obtained at the National Synchrotron Light Source (NSLS)
beamline X3B (ACO) and the Stanford Synchrotron Radiation Lightsource beamline 9ꢀ3
(NOV2, CAO1). At NSLS, the storage ring operated at 2.8 GeV and 180ꢀ300 mA. A Si(111)
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double crystal monochromator with sagittal focusing of the second crystal provided energy
selection and horizontal focusing, with a downstream Niꢀcoated cylindrically bent mirror
rejecting higher harmonics and vertically focusing the beam. At SSRL, the storage ring operated
at 3.0 GeV and 500 mA in topꢀoff mode. A Si(220) double crystal monochromator provided
energy selection, with an upstream Rhꢀcoating collimating mirror rejecting higher harmonics
prior to the monochromator and a downstream Rhꢀcoated toroidal mirror focusing the
monochromatic radiation at the sample. Temperature control was provided by a He Displex
cryostat (NSLS, 15ꢀ20 K sample temperature) or Oxford liquid He cryostat (SSRL, 10K sample
temperature). XAS data were acquired in fluorescence mode using Canberra solid state
germanium detectors (NSLS: 31 element discrete; SSRL: 100 pixel monolithic) with XIA digital
data acquisition electronics. Data were typically acquired in 10 eV steps in the preꢀedge region (1
sec. acquisition time), 0.3 eV steps along the edge (2 sec. acquisition time), and 0.05k steps in
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the EXAFS up to 14ꢀ15k (acquisition time increasing from 2ꢀ9 sec in k weighted fashion).
Samples were monitored for evidence of radiation damage as indicated by redꢀshifts in either the
preꢀedge or edge energies, and new spots were exposed as needed. An iron metal foil placed
between ion chambers downstream of the sample was used for energy calibration, with the first
inflection point of the edge set to 7112.0 eV.
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Data analyses. XAS data were processed and averaged using Athena for NSLS X3B data,
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whereas EXAFSPAK was used for SSRL 9ꢀ3 data. XANES peakꢀfitting was conducted using
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Fityk with the rising edge modeled using pseudo—Voigt functions and the preꢀedge modeled
with Gaussian functions. Fits were carried out over several different energy ranges spanning
7104ꢀ7118 eV, and reported preꢀedge peak fit parameters are averages of at least 3 fits. EXAFS
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calculated for simple models of the ACO active site using FEFF6L. From this, relevant paths
were incorporated into the fitting model and evaluated for the significance of their contributions.
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For a given shell in all simulations, the coordination number n was fixed, while r and σ were
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allowed to float. The amplitude reduction factor S0 was fixed at 0.9, while the edge shift
parameter ꢃE was allowed to float at a single common value for all shells. Histidine multiple
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scattering was evaluated using a method where the imidazole moiety is represented by four sets
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of grouped paths representing the dominant scattering contributions . The fit was evaluated in
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k ꢀweighted Rꢀspace, and fit quality was judged by the reported Rꢀfactor and reduced χ . Bond
valence sums were used as an additional test to identify the best fit to the EXAFS data, using
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reported parameters for Fe(II) .
Sample preparation for EPR and Mössbauer spectroscopy
E and ES sample preparation. Samples of ACO (0.5 mM ACO in 20 mM HEPESꢀNaOH, pH
7
, containing 20% Glycerol and 0.02ꢀ0.05% v/v Triton Xꢀ100), CAO1 (1.2 mM CAO1 in 16 mM
HEPESꢀNaOH, pH 7.5, 160 mM NaCl and 20% v/v glycerol) and NOV2 (1.9 mM NOV2 in 20
mM TrisꢀHCl, pH 7.5 or 8.5, 200 mM NaCl and 20% glycerol with or without 1 mM Tris
carboxylethyl phosphine) were made anaerobic by passing Ar(g) over the top of the sample as it
was stirred at 5 °C for 30 min in a sealed vial. Following this, the protein vials were transferred
into an anaerobic N gasꢀfilled glove box. The vials were opened in the glove box and stirred
2
while open to the glove box environment for 1 h. Resveratrol and piceatannol substrate stock
solutions (200 mM) were prepared by preꢀweighing an amount of solid substrate and transferring
the powder into a sealed vial. The vial was then transferred into the glove box to equilibrate with
the glove box environment overnight. Prior to its addition to enzyme samples, the substrate was
dissolved in degassed DMSO that had also been stored in the glove box overnight. Typically, 1ꢀ5
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ꢂL of substrate in DMSO was delivered to the protein solution, to a maximum concentration of
5% v/v DMSO, which did not affect protein activity. Stock solutions of 4 mM allꢀtransꢀ8’ꢀ
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apocarotenol, generated by reduction of allꢀtransꢀ8’ꢀapocarotenal as previously described
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were prepared by dissolving solid apocarotenoid in a solution of degassed ethanol. For O2
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binding experiments, a 1 mM sample of Feꢀenriched NOV2 in a Mössbauer cell at 4 °C was
exposed to pure O gas at a pressure of 40 psi for 5 minutes. While maintaining 40 psi at 4 C,
2
the sample was frozen in liquid nitrogen.
Preparation of Feꢀnitrosyl complexes. Samples of ACO (0.2 mM protein in 20 mM HEPESꢀ
NaOH, pH 7, 20% Glycerol and 0.02ꢀ0.05% Triton Xꢀ100) and NOV2 (0.1 mM protein in 20
mM TrisꢀHCl, pH 7.5, and 20% glycerol) and substrates were prepared anaerobically as
described above. A stock solution of diethylammonium salts (DEA NONOate) was prepared by
dissolving a preꢀweighed amount of NONOate powder in degassed 5 mM NaOH, to give a
concentration of 10ꢀ15 mM DEA NONOate. The vial containing dissolved DEA NONOate was
transferred to the glove box and equilibrated in the glove box overnight. This solution was stored
in the glovebox for no longer than 24 h before use. Over this period, the DEA NONOate is stable
in the NaOH stock solutions. An aliquot of substrate stock solution was added to the protein
solutions in the glove box (when applicable), and left to incubate for 2 min. An aliquot of the
DEA NONOate solution then was added to each protein solution. The equivalents of NO
delivered were determined based on the calculated rate of decay of the DEA NONOate (halfꢀlife
of 16 min at pH 7.4 and 22 °C) and were generally in excess (3 eq. or more) relative to the
protein concentration . Following a 5 min incubation, the samples were transferred to either an
EPR tube or Mössbauer cup, sealed, and quickly removed from the glove box and frozen in
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EPR and Mössbauer Spectroscopy
XꢀBand EPR spectra were recorded on a Bruker Elexsys spectrometer equipped with an
Oxford ESR 910 cryostat and a Bruker bimodal cavity. The modulation amplitude and frequency
were 1 mT and 100 kHz for all spectra. The microwave frequency was measured with a
frequency counter and the magnetic field was calibrated with a NMR gaussmeter. The
temperature was calibrated with a carbon glass resistor (Lakeshore CGRꢀ1ꢀ1000). EPR signals
were quantified by relative to a 1 mM Cu(II)EDTA standard in 10% glycerol. The Cu(II)
concentration was quantified by inductively coupled plasma mass spectrometry. Species
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concentrations were determined with the software SpinCount developed by M. P. Hendrich .
The software diagonalizes the spin Hamiltonian, H = β B•g •S + S•D•S, where all parameters
e
e
have their usual definitions. The quantitative simulations are leastꢀsquares fits of the
experimental spectra generated with consideration of all intensity factors, which allows the
computation of simulated spectra for a specified sample concentration.
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Fe Mössbauer spectra were recorded with two spectrometers using Janis Research dewars.
Isomer shifts are reported relative to iron metal at 298 K. The simulations of Mössbauer spectra
were calculated with leastꢀsquares fitting using the program SpinCount and the standard spin
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Hamiltonian :
ꢀ
ꢁ
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H = β B•g •S + D[Sz – S + E/D(S + S ) +S•A•I – g β B•I +
[3I – I +η[I – I ]]
e
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RESULTS
Crystal structure of a fungal stilbenoid-cleaving CCO
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To provide a structural framework for our spectroscopic investigations, we aimed to elucidate
the structure of a stilbenoidꢀcleaving CCO. We focused our search on those enzymes with
defined substrate specificity. In our expression system, the originally characterized stilbenoidꢀ
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cleaving CCO from Pseudomonas paucimobilis, lignostilbeneꢀα,βꢀdioxygenase (LSD) 1 , was
Figure 2. Crystal structure of CAO1 and its
comparison to ACO. (A) Cartoon representation of
CAO1 (left) and ACO (right). The protein chain is
colored blue to red from the Nꢀ to Cꢀtermini. The first
and second sphere His and Glu iron ligands,
respectively, are shown as sticks with the bound iron
shown as a brown sphere. (B) Closeꢀup view of the
CAO1 (left) and ACO (right) iron centers. (C)
Comparison of the cavities leading to the iron center
in CAO1 (left) and ACO (right). The prominent apolar
tunnel (indicated by the arrow) present in ACO is
absent in CAO1. The CAO1 active site is instead
accessible through a narrow tunnel that is found in
roughly the same location as the hydrophilic tunnel
found in ACO. The entrances to these tunnels are
marked with orange asterisks. (D) CAO1 (left) and
ACO (right) in surface representation as viewed down
the axes of the arrows shown in panel (C). The
entrance to the hydrophobic tunnel in ACO
(
delineated by a dashed line) is surrounded by a
cluster of hydrophobic residues (represented as sticks
and ribbons) that allow the protein to interact with
membranes where hydrophobic carotenoids are
dissolved. The corresponding region of CAO1 is more
polar due to multiple amino acid substitutions.
Additionally, segments of CAO1 that are homologous
to the hydrophobic patch residues adopt
conformations that occlude the potential tunnel and
thus prevent active site accessibility from this region
of the protein.
found in inclusion bodies. NOV2, by
contrast, was soluble, highly expressed
1
7
and could be purified in an active form
but failed to crystallize after extensive
trials. A third stilbenoidꢀcleaving CCO
12
from the fungus Neurospora crassa, referred to as CAO1 , was also active when purified (Fig.
S1) and readily formed rodꢀshaped trigonal crystals that diffracted Xꢀrays to ~2 Å resolution
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(
Table S1). The crystal structure of this enzyme was solved in space group P3 21 by molecular
2
replacement in combination with anomalous scattering data.
CAO1, like other CCOs, adopts a 7ꢀbladed βꢀpropeller covered on its top face by a set of loops
and αꢀhelices that form a dome structure housing the active site (Fig. 2A). The iron cofactor
resides on the top face of the propeller coordinated by His residues 197, 248, 313 and 510 (Fig.
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B). Despite the low sequence identity between CAO1 and other CCOs of known structure, their
overall threeꢀdimensional similarity is relatively high with rootꢀmeanꢀsquare (RMS) differences
between matched Cα atoms of 1.6, 2.1, 2.3 and 2.6 Å for NOV1, ACO, VP14 and RPE65
4
5
respectively . The asymmetric unit of the CAO1 crystals is composed of a nearly D symmetric
2
dimer of dimers. The four monomers are practically identical in structure except for residues
2
65ꢀ285 and 334ꢀ355, which differ between subunits by a small rigid body displacement. One of
2
the assemblies is a twoꢀfold symmetric dimer with a buried surface area of ~1,415 Å per
monomer (Fig. S2A). The dimer interface is comprised of an antiparallel βꢀstrand interaction
formed between residues 51ꢀ54 of the outer βꢀstrand of blade VII together with polar and nonꢀ
polar interactions involving residues 35ꢀ37 and 47ꢀ62 within the Nꢀterminal segment of the
protein along with residues 80, 81, 105ꢀ108, 126 and 500ꢀ504 (Fig. S2B). Both the positive
5
6
calculated ꢁG of interface association as well as its modest shape complementary statistic of
57
0
.613 suggest the dimer may not be stable. However, CAO1 migrates faster on size exclusion
media than expected based on its calculated molecular weight, which could be attributable to
dimer formation in solution (Fig. S1A and C). LSD isozymes also form homoꢀ and heteroꢀ
58
dimeric assemblies in solution . Interestingly, a dimeric assembly like that of CAO1 is also
24
present in the NOV1 crystal structure suggesting a conserved interface .
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The most notable differences between CAO1 and structurally characterized carotenoid/retinoid
metabolizing CCOs, for which ACO is considered the prototype, occur in the substrateꢀbinding
cavity and region corresponding to the membraneꢀinteracting nonꢀpolar patch (Fig. 2C and D).
These CCOs all possess a hydrophobic patch on their surfaces surrounding the opening to an
apolar cavity that serves as both a passageway and a binding site for lipophilic
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carotenoid/retinoid substrates . The hydrophobic patch on the surface allows the enzymes to
interact with membranes and extract their lipophilic substrates, which then traverse the apolar
cavity to reach the catalytic center. In CAO1, this route is absent owing to multiple structural
alterations. The metal center is instead accessible only through a single narrow entry port whose
opening locates to an alternative region of the protein surface (Fig. 2C). This tunnel is
topographically equivalent to the hydrophilic tunnel identified in ACO through which one of its
2
two apocarotenoid products is thought to exit the active site . Similar structural features were
observed in the bacterial stilbenoidꢀcleaving CCO, NOV1, although this enzyme has a broader
entry port structure than CAO1 due in part to an unstructured loop in this region of the protein.
Many of the residues in CAO1 equivalent to those comprising the hydrophobic patch of ACO
are either charged, polar or adopt nonꢀsolvent exposed conformations. This together with the
lack of a hydrophobic patch around the mouth of the substrate entry port demonstrates that
membraneꢀintegrating capability is absent in the stilbeneꢀcleaving CCO lineage. The structure
reveals a striking rearrangement of residues equivalent to those of the hydrophobic patch in ACO
such that active site accessibility from this region of the enzyme is completely occluded (Fig. 2D
and Fig. S3). The tunnel is further blocked by residues Arg166, Asp166, Glu213 and His277
whose charged side chains protrude into the potential cavity to form ionic or dipolar interactions.
Crystallographic description of the CCO iron resting state
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In contrast to these marked differences in the substrate binding tunnel and active site
accessibility, the CAO1 4ꢀHis iron center closely resembles that of other structurally
characterized CCOs. His residues 197, 248, 313 and 510 directly coordinate the iron cofactor
with bond lengths of 2.4, 2.2, 2.3, and 2.3 Å, respectively, averaged over the four monomers
(Fig. 2B). Notably, the iron Bꢀfactors in this structure were somewhat elevated as compared to
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ε
those of the surrounding N atoms suggesting subꢀmaximal iron occupancy. The His ligands are
ε
ε
arranged with the N ꢀFeꢀN angles all near 90°. A solvent molecule is bound to the iron trans to
His197 with a bond length of ~2.4 Å, which suggests it is a coordinated water, whereas the sixth
site trans to His304 is unoccupied by ligand due to the nearby Thr151 methyl group. The iron is
thus 5ꢀcoordinate with a distorted square pyramidal geometry, similar to that of ACO (Fig. 2B)
15
24
but unlike VP14 and NOV1 which contain O modeled in place of the solvent atom. Glu
2
residues 165, 383 and 453 hydrogen bond with His 248, 313 and 510, respectively, forming an
anionic second coordination sphere similar to that of ACO.
XAS structures of the CCO 4-His iron center resting state
Crystal structures of CCOs provide a general description of the 4ꢀHis iron center, but they lack
the ability to provide high precision metalꢀligand bond lengths, detect subtle differences in active
site geometry, and may exhibit structural differences that arise from the conditions required for
crystallization. To characterize the CCO 4ꢀHis Fe(II) structure in greater detail and under a more
uniform environment we carried out iron Kꢀedge Xꢀray absorption spectroscopy studies on ACO,
NOV2 and CAO1.
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Xꢀray absorption nearꢀedge spectroscopy (XANES) provides insight into the metal oxidation
state and site symmetry. All three asꢀisolated enzymes exhibit similar edge shapes, with
inflections observed at 7121.4 – 7121.8 eV and 7125.3 – 7126.1 eV, consistent with previous
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observations for Fe(II)ꢀRPE65
(Fig. 3A and Table S2). There was
Figure 3. XAS analysis of asꢀisolated CCO
iron centers. (A) Normalized XANES
spectra of asꢀisolated Fe(II)ꢀACO (——),
Fe(II)ꢀNOV2 (– – –), and Fe(II)ꢀCAO1
(•••). The inset shows an expansion of the
preꢀedge region. The first derivative
XANES spectrum is shown in the bottom
panel. (B) Representative best fits (bolded
3
entries in Tables S3 – S5) to k ꢀweighted
EXAFS data of Fe(II)ꢀACO (top), Fe(II)ꢀ
NOV2 (middle), and Fe(II)ꢀCAO1
(bottom). Experimental data is shown as a
dotted line, whereas the best fit is shown as
a solid line.
minimal to no photoreduction in the Xꢀray beam upon
extended exposure. Both observations are consistent with
an Fe(II) center in all three asꢀisolated enzymes. The preꢀedge peaks, associated with a parityꢀ
forbidden 1sꢀtoꢀ3d transition that gains intensity by mixing with 4p orbitals, provide additional
understanding of metal site structure. Fe(II)ꢀACO exhibits a single, broad preꢀedge peak centered
at ~7113.0 eV with an area of 12.7(1) units (Table S2 and Fig. S4). The other enzymes,
measured with a higherꢀresolution Si(220) monochromator, show evidence for two preꢀedge
peaks with an approximate energy splitting of 1.9 eV. Specifically, Fe(II)ꢀNOV2 has two preꢀ
edge peaks at 7112.6 eV and 7114.5 eV with areas of 9.5(7) and 4.7(3) units, respectively, while
the Fe(II)ꢀCAO1 preꢀedge peaks are found at 7112.8 eV (10.6(5) units) and 7114.6 eV (5.9(9)
units) (Table S2 and Fig. S4). The energies of these peaks are consistent with the expected Fe(II)
state. The peak areas, on the high side for Fe(II), are most consistent with an Fe coordination
6
0, 61
number of four or five
. We note that the Fe(II)ꢀNOV2 preꢀedge spectrum and intensity
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pattern is quite similar to previously reported 5ꢀcoordinate, C Fe(II) sites in a model complex
3
v
6
3
and the mineral grandidierite . We favor a five coordinate assignment based on the EXAFS and
1
3, 28
Mössbauer analyses described below as well as the ACO
which both feature 5ꢀcoordinate iron.
and CAO1 crystal structures,
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Extended Xꢀray absorption fine structure (EXAFS) analyses provided additional details of the
iron coordination environment. All three Fe(II)ꢀenzyme preparations can be fit with a single shell
of 5 N/O scatterers at 2.12 – 2.13 Å (Fig. 3B, and Tables S3ꢀ5). For ACO, this shell can be
further split into two subshells consisting of 4 N/O scatterers at 2.15 Å and 1 O/N scatterer at
1
.98 – 2.01 Å, the latter being attributable to an Fe(II)ꢀhydroxo unit. These distances are in good
2
8
agreement with the bond lengths determined by crystallography . CAO1 required a more
elaborate scattering model consisting of 4 NHis scatterers at 2.15 Å, a shorter N/O scatterer at
1
.99 Å and two additional light atom (C/N/O) scatterers at 2.5 Å. In the CAO1 crystal structure,
the short scatterer is not apparent, possibly because of the lower pH of the crystallization solution
compared to the solution used for XAS studies, which could alter the bound solvent protonation
state. The more distant scattering is at least partially attributed to the ~2.4 Å solventꢀiron
interaction observed crystallographically (Fig. 2B). Notably, the relatively low DebyeꢀWaller
factor suggests a stable association of these longerꢀrange scatterers with the iron center, and is
consistent with nearly complete occupancy of these sites by bound ligand. The requirement for
2.5 Å FeꢀC/N/O units in the CAO1 scattering model bears resemblance to the scattering
59
observed for RPE65 . In the latter enzyme, the longer scatterer was attributed to the bridging
carbon of a bound carboxylate. Our inability to resolve a short Fe–O moiety in Fe(II)ꢀNOV2
would suggest that solvent water is not deprotonated in the asꢀisolated form of the enzyme.
These 5ꢀcoordinate solutions from EXAFS are consistent with the XANES analyses and are also
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supported by bond valence sum calculations . In addition, multiple scattering analyses provide
evidence for three to four Hisꢀderived imidazole units bound to the iron center in each enzyme,
consistent with the known 4ꢀHis coordination motif of CCOs.
Characterization of the CCO iron center by 57Fe-Mössbauer spectroscopy
Mössbauer spectroscopy is sensitive to the
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primary coordination sphere of iron active sites in
enzymes and provides information about the
electronic structure of the metal center. As the
57
natural abundance of Fe is low (2%), enrichment
5
7
with Fe significantly improves the spectral content.
We developed methods for highꢀyield expression of
each CCO in ironꢀdeficient M9 minimal media in
5
7
which Fe can be added to produce enriched
enzyme samples. Of the three enzymes, NOV2 was
expressed at the highest levels, which allowed the
most detailed spectroscopic characterization for
this protein. The Mössbauer spectrum of NOV2
recorded at 4.2 K (Fig. 4A) showed a broad
doublet with parameters given in Table 1 that are
Figure 4. Mössbauer spectra recorded at 4.2 K in an
applied field of 45 mT. (A) NOV2, (B) NOV2 in the
presence of 10 eq. piceatannol, (C) NOV2 after
turnover of 10 eq. piceatannol, (D) NOV2 after
turnover of 10eq. piceatannol and subsequent
anaerobic addition of 10eq piceatannol. (E) NOV2 in
the presence of 10 eq. resveratrol. Protein (~ 2 mM)
was prepared in 20 mM TrisꢀHCl, pH 7.5, 200 mM
NaCl, 20% v/v glycerol. The experimental and
simulated spectra are shown as blue and black lines,
respectively.
indicative of nonꢀheme S = 2 Fe(II). The width of the doublet (Γ = 0.5 mm/s) is compatible with
the presence of two or more Fe(II) species. Approximately 20% of iron in the sample was
adventitious Fe(III), indicated by the broad paramagnetic features extending to ±8 mm/sec. This
adventitious Fe(III) was unreactive to reductants or substrate turnover and the amount was
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preparation dependent. The Mössbauer spectrum of CAO1 recorded at 4.2 K in a small applied
field (Fig. S5C) showed a nonꢀheme Fe(II) quadrupole doublet similar to that of NOV2 with
parameters given in Table 1. As with NOV2, the broad line width (Γ = 0.5 mm/s) indicates
multiple conformations of the active site.
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Notably, the isomer shifts of these two enzymes are comparable to each other suggesting
similar active site environments, but significantly lower than other nonꢀheme iron enzymes
(Table 1), including bacterial and plant reaction centers which possess 4ꢀHis nonꢀheme iron
centers similar in structure to CCOs. The low isomer shifts for NOV2 and CAO1 indicate a
comparatively more electropositive Fe(II) environment. The 4ꢀHis, 1 water/hydroxide iron
coordination of CCOs contrasts with the more common 2 His, 1 carboxylate and 3 water
coordination of 2Hꢀ1C dioxygenases. The difference in coordination of two additional His
residues versus a carboxylate and 2 waters is expected to lower the ligand charge donation to the
iron, thus causing the observed lower isomer shift.
The Mössbauer spectrum of ACO recorded at 4.2 K in a small applied field (Fig. S5A) showed
a broad (Γ = 0.5 mm/s) quadrupole doublet, similar to NOV2/CAO1, with parameters given in
Table 1. However, the isomer shift of ACO is significantly higher than that of CAO1 and NOV2
and comparable to other nonꢀheme iron proteins. This finding indicates a different primary
coordination sphere for the ACO Fe(II) compared to NOV2 and CAO1. Considering the near
uniform FeꢀHis bond lengths and coordination geometry observed for these three enzymes, the
isomer shift difference may arise from differences in ligands occupying exchangeable sites, such
as a change in the protonation state of water or an additional water coordinating to the iron.
In the absence of substrate, the Fe(II) sites of the CCOs studied here were stable in air at room
temperature. Previous crystallographic studies on VP14 and NOV1 reported a stable O ꢀiron
2
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species
. In an attempt to generate such a species, a sample of uncomplexed NOV2 was
exposed to pure O gas at the highest pressure tolerable by the sample cell and preparatory
2
equipment (40 psi). O coordination to Fe(II) centers results in significant and characteristic
2
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changes in the Mössbauer parameters . However, the Mössbauer spectrum before and after O2
exposure was unchanged. Assuming a minimum detection level of 10% of the total sample iron
for an ironꢀoxy species gives a dissociation constant for O of greater than 50 mM in the absence
2
of substrate. This result, in conjunction with the structural data presented above, demonstrates
that the enzymes of the present study do not have O bound to iron in the absence of organic
2
substrate.
Mössbauer analysis of the Fe(II) center interaction with substrates
With the resting state of the CCO Fe(II) center structurally and electronically defined we next
turned our attention to its interactions with substrates and substrateꢀmimetics. We used
previously established substrates for enzymes in these studies: resveratrol and piceatannol for
8
, 12
11
NOV2 and CAO1
and allꢀtransꢀapoꢀ8’ꢀcarotenol for ACO . The direct interaction of these
57
organic substrates with the metal of each enzyme was first investigated by FeꢀMössbauer
spectroscopy, which probes ligandꢀinduced perturbations in the iron electronic structure. For
nonꢀheme Fe(II) enzymes in which the substrate binds to the iron, the addition of substrate
causes a distinct change in the Mössbauer parameters (Table 1).
Table 1. Mössbauer parameters of non-heme Fe(II) complexes.
a
a
Sample
ACO
ACO + apocarotenal
NOV2
NOV2 + resveratrol
NOV2 + piceatannol
δ
ꢃEq
3.05
2.92
3.47
%
reference
tw
tw
tw
tw
tw
b
1.21
1.18
1.09
1.08
1.08
100
100
100
30/70
30/70
2.90/3.47
2.27/3.42
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