Proximal ligand electron donation and reactivity of the cytochrome P450 ferric-peroxo anion

Article abstract of DOI:10.1021/ja211499q

CYP125 from Mycobacterium tuberculosis catalyzes sequential oxidation of the cholesterol side-chain terminal methyl group to the alcohol, aldehyde, and finally acid. Here, we demonstrate that CYP125 simultaneously catalyzes the formation of five other products, all of which result from deformylation of the sterol side chain. The aldehyde intermediate is shown to be the precursor of both the conventional acid metabolite and the five deformylation products. The acid arises by protonation of the ferric-peroxo anion species and formation of the ferryl-oxene species, also known as Compound I, followed by hydrogen abstraction and oxygen transfer. The deformylation products arise by addition of the same ferric-peroxo anion to the aldehyde intermediate to give a peroxyhemiacetal that leads to C-C bond cleavage. This bifurcation of the catalytic sequence has allowed us to examine the effect of electron donation by the proximal ligand on the properties of the ferric-peroxo anion. Replacement of the cysteine thiolate iron ligand by a selenocysteine results in UV-vis, EPR, and resonance Raman spectral changes indicative of an increased electron donation from the proximal selenolate ligand to the iron. Analysis of the product distribution in the reaction of the selenocysteine substituted enzyme reveals a gain in the formation of the acid (Compound I pathway) at the expense of deformylation products. These observations are consistent with an increase in the pK_a of the ferric-peroxo anion, which favors its protonation and, therefore, Compound I formation.

Full text of DOI:10.1021/ja211499q

Article
pubs.acs.org/JACS
Proximal Ligand Electron Donation and Reactivity of the Cytochrome
P450 Ferric−Peroxo Anion
Santhosh Sivaramakrishnan,† Hugues Ouellet,†,‡ Hirotoshi Matsumura,§ Shenheng Guan,†
Pierre Moenne-Loccoz,§ Alma L. Burlingame,† and Paul R. Ortiz de Montellano*,†
̈
†Department of Pharmaceutical Chemistry, University of California, 600 16th Street, San Francisco, California 94158-2517, United
States
§Division of Environmental & Biomolecular Systems, Oregon Health & Science University, 20,000 NW Walker Road, Beaverton,
Oregon 97006-8921, United States
S
* Supporting Information
ABSTRACT: CYP125 from Mycobacterium tuberculosis catalyzes sequential
oxidation of the cholesterol side-chain terminal methyl group to the alcohol,
aldehyde, and finally acid. Here, we demonstrate that CYP125 simultaneously
catalyzes the formation of five other products, all of which result from deformylation
of the sterol side chain. The aldehyde intermediate is shown to be the precursor of
both the conventional acid metabolite and the five deformylation products. The acid
arises by protonation of the ferric−peroxo anion species and formation of the ferryl−
oxene species, also known as Compound I, followed by hydrogen abstraction and
oxygen transfer. The deformylation products arise by addition of the same ferric−
peroxo anion to the aldehyde intermediate to give a peroxyhemiacetal that leads to
C−C bond cleavage. This bifurcation of the catalytic sequence has allowed us to examine the effect of electron donation by the
proximal ligand on the properties of the ferric−peroxo anion. Replacement of the cysteine thiolate iron ligand by a selenocysteine
results in UV−vis, EPR, and resonance Raman spectral changes indicative of an increased electron donation from the proximal
selenolate ligand to the iron. Analysis of the product distribution in the reaction of the selenocysteine substituted enzyme reveals
a gain in the formation of the acid (Compound I pathway) at the expense of deformylation products. These observations are
consistent with an increase in the pKa of the ferric−peroxo anion, which favors its protonation and, therefore, Compound I
formation.
is converted by protonation to a ferric−hydroperoxo (Fe3+-
INTRODUCTION
The cytochromes P450 (CYPs) are heme−thiolate enzymes
that use O2 and two reducing equivalents to insert an oxygen
atom into organic compounds, including fatty acids, steroids,
and xenobiotics. CYPs have been identified in all domains of
■
OOH) species named Compound 0. A second protonation of
the distal oxygen of Compound 0 results in immediate
fragmentation to give a molecule of water and a ferryl Fe4+
O species with a radical cation delocalized over the porphyrin
and the proximal thiolate iron ligand.22 On the basis of
peroxidase nomenclature, this intermediate can be designated
as Compound I. In a conventional hydroxylation, Compound I
abstracts a hydrogen atom from a substrate to give a carbon
radical and the equivalent of an iron-bound hydroxyl radical,
which then undergo radical recombination to give the
hydroxylated product.23,24 The factors governing the formation
and stability of Compound I have been extensively studied.25
The cysteine thiolate ligated to the heme iron atom is
believed to be important for the formation from molecular
oxygen and reactivity of Compound I in cytochromes P450.26
This feature distinguishes P450 enzymes from peroxidases,
catalases and other heme containing enzymes in which the
proximal iron ligand is either a histidine nitrogen or a tyrosine
hydroxyl. Electron donation from the proximal cysteine thiolate
ligand is proposed to facilitate protonation of the distal O−O
life and more than 11 500 distinct isoforms are now known.1 In
humans, some CYPs, (e.g., CYP19 or CYP51) are highly
specific and act on a single substrate, or a few closely related
substrates, whereas other isoforms (e.g., CYP3A4, CYP2D6 and
CYP2C9) are promiscuous and are responsible for the
oxidation of approximately 70% of all therapeutic drugs.2 In
contrast, the biological function(s) of the vast majority of
bacterial CYPs remain to be elucidated, but some (e.g., EryF,
EpoK, OxyB, OxyD, PikC, and PimD) have been shown to be
involved in the biosynthesis of antibiotics.3−12 Other bacterial
CYPs have been shown to participate in catabolic pathways,
including the degradation of terpenoids (e.g., CYP176A1
(P450cin),13 steroids (CYP125 and CYP142),14−19 and
explosives (XplA).20,21
The P450 catalytic cycle is initiated by one electron
reduction of the ferric heme iron (Fe3+) to the ferrous state
(Fe2+) which then binds molecular oxygen to give the ferrous
dioxy (Fe2+-O2) form. A second electron transfer to this
intermediate generates a ferric−peroxo anion (Fe3+-O-O−) that
Received: December 8, 2011
Published: March 23, 2012
© 2012 American Chemical Society
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Scheme 1. Partitioning of the Ferric−Peroxo Anion into the Compound I-Mediated Monooxygenation (a) and
Peroxyhemiacetal-Mediated Deformylation (b) Pathways in the Oxidation of an Aldehyde
bond in Compound 0, thus, channeling this intermediate
toward heterolytic formation of Compound I.27,28
increasing its pKa, and therefore promotes the formation of
Compound I.
Although Compound I is thought to be responsible for the
majority of P450-catalyzed oxidations, it is not responsible for
the oxidative C−C bond cleavage observed in the P450-
mediated oxidation of aldehydes. Nucleophilic addition of the
ferric−peroxo anion intermediate to the aldehyde to give a
peroxyhemiacetal is the generally accepted key step in
formation of the C−C bond cleavage products.29−31 The
actual C−C bond cleavage may occur by either concerted or
homolytic radical fragmentation of the peroxyhemiacetal.29
Thus, the ferric−peroxo anion intermediate can either undergo
two protonation steps to generate Compound I, resulting in
conventional monooxygenase activity, or it can add as a
nucleophile to an aldehyde carbonyl group, triggering a reaction
pathway that leads to C−C bond cleavage (Scheme 1).
Although the effect of proximal electron donation on the
reactivity and stability of Compound I has been studied, its
impact on the reactions involving the ferric−peroxo anion
intermediate remains obscure.
CYP125 from Mycobacterium tuberculosis (Mtb) is required
for degradation14,17−19 and incorporation of the cholesterol
side chain into the lipidic virulence factor, phthiocerol
dimycocerosate (PDIM).19 In addition, CYP125 is also
required to alleviate the toxic effect of cholest-4-en-3-one on
Mtb growth in vitro.19 CYP125 is a steroid C26-monoox-
ygenase that sequentially oxidizes the terminal methyl of the
side chain of both cholesterol and cholest-4-en-3-one to the
alcohol, aldehyde, and finally carboxylic acid.19 This same
oxidation of cholesterol is catalyzed by two other enzymes from
Mtb, CYP124 and CYP142,17 and by the human enzyme
CYP27A1.32
In this work, we have examined in greater detail the
enzymatic activity of CYP125 with cholesterol and cholest-4-
en-3-one. Liquid-chromatography mass spectrometry (LC−
MS) in combination with deuterium- and 18O2-labeling has led
to the identification of five additional metabolites produced by
this surprisingly versatile enzyme, all of which arise by carbon−
carbon bond cleavage stemming from attack of the ferric−
peroxo anion on the intermediate aldehyde. Furthermore, a
proximal selenocysteine substituted CYP125, SeCYP125, was
expressed, purified, and characterized using UV−vis, resonance
Raman, and EPR spectroscopy. The selenocysteine substitution
has made it possible to investigate the role of electron donation
by the proximal ligand on partitioning between the reaction
pathways that lead to formation of the carboxylic acid versus
carbon−carbon bond cleavage. Our results provide direct
evidence that increased proximal electron donation increases
protonation of the ferric−peroxo anion, presumably by
EXPERIMENTAL PROCEDURES
Materials. All solvents and reagents were of the highest purity
■
commercially available unless noted otherwise. Methyl-β-cyclodextrin
(mβCD), glucose-6-phosphate, glucose-6-phosphate dehydrogenase
(Saccharomyces cerevisiae), ferredoxin (Spinacea oleracea), ferredoxin
NADP+-reductase (S. oleracea), cholesterol oxidase (Streptomyces sp.),
cholesterol, cholest-4-en-3-one, progesterone and Dess-Martin reagent
were purchased from Sigma-Aldrich (St. Louis, MO). (25R)-26-
Hydroxycholesterol, (25S)-26-hydroxycholesterol and 1,4-cholesta-
diene-3-one were purchased from Research Plus (Barnegat, NJ).
Cholest-4-en-3-one-(25S)-26-acid was obtained from Avanti Polar
Lipids (Alabaster, AL).
Expression and Purification of CYP125C173S/C429L and
SeCYP125C173S/C429L. The C173S/C429L double mutant was used
to eliminate all cysteine residues in the protein other than the proximal
iron ligand. Amino acid substitution for the new construct
CYP125C173S/C429L was carried out using a QuikChange site-directed
mutagenesis kit (Stratagene) with the following primers: FC173S, 5′-
GTCGAGCAGGTTTCCTCTGAGCTGCCATTGCAG-3′; RC173S
5′-CTGCAATGGCAGCTCAGAGGAAACCTGCTCGAC-3′; FC429L
5′-CGACTACACCGGTAGACTCCCGGTTGCTCACT-3′; RC429L
,
,
,
5′-AGTGAGCAACCGGGAGTCTACCGGTGTAGTCG-3′. A con-
struct in which the N-terminus was truncated at Val17 was used as a
template. This construct was shown to improve crystallization and did
not exhibit any difference in enzymatic activity. The mutations were
verified using DNA sequencing. The CYP125 and CYP125C173S/C429L
variant were purified as described previously.19 The protein
preparations were assayed for their P450 Fe2+-CO difference spectrum
and the concentration of P450 was determined from the difference
spectrum using the extinction coefficient ε450−490 = 91 000 mM−1
cm−1.33 No difference in stability or enzymatic activity was observed
for the CYP125C173S/C429L variant relative to either the full-length or
the truncated form of CYP125. Expression of the selenocysteine
substituted CYP125C173S/C429L, SeCYP125, was carried out according
to the previously published protocol.34 The enzyme was purified in the
same way as the CYP125 protein19 and the concentration was
determined from the Fe2+-CO difference spectrum using the extinction
coefficient ε459−490 = 91 000 mM−1 cm−1. The total heme b content
was also quantitated by the pyridine hemochrome method.35 For
clarity, the CYP125C173S/C429L containing cysteine at the active site and
SeCYP125C173S/C429L proteins will be simply called WT* and
SeCYP125, respectively.
EPR and Resonance Raman Spectroscopy. The EPR and
resonance Raman (RR) analyses were performed with enzyme
concentrations ranging from 40 to 100 μM. EPR spectra were
obtained on a Bruker E500 X-band EPR spectrometer equipped with a
superX microwave bridge and a dual mode cavity with a helium flow
cryostat (ESR900, Oxford Instruments, Inc.). The microwave power,
modulation amplitude, magnetic field sweep, and the sample
temperature were optimized to ensure nonsaturating conditions for
all low spin Fe3+ EPR active species. The RR spectra were collected
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Figure 1. Reverse-phase HPLC separation of CYP125-catalyzed oxidation products of (A) cholest-4-en-3-one, (B) 26-hydroxycholest-4-en-3-one
and (C) cholest-4-en-3-one-26-aldehyde. The separation of the products was carried out using a reverse phase C8-A Polaris column (Varian), as
described in the Experimental Procedures. The elution was monitored at 240 nm. The retention times are: peak VI, 14.9 min; peak V, 15.8 min; peak
IV, 17.2 min; peak III, 18.3 min; peak II, 20.8; peak I, 29.6 min. The m/z values of the parent and daughter ions are listed in Table S1. Asterisk (*)
indicates contaminant from the cholest-4-en-3-one-26-aldehyde preparation.
using a 90° scattering geometry on room-temperature samples
mounted on a reciprocating translation stage with a custom
McPherson 2061/207 spectrograph (set at 0.67 m with variable
gratings) equipped with a Princeton Instruments liquid-N2-cooled
CCD detector (LN-1100PB). The 413-nm excitation was obtained
from an Innova 302 krypton laser (Coherent, Santa Clara CA) and the
442-nm excitation from a He/Cd laser (Liconix 4240NB). Kaiser
Optical supernotch filters were used to attenuate Rayleigh scattering.
Frequencies were calibrated relative to indene, CD3CN and NOg
standards and are accurate to 1 cm−1. Polarization conditions were
optimized using CCl4. The integrity of the RR samples, before and
after illumination, was confirmed by direct monitoring of their UV−vis
spectra in Raman capillaries.
Synthesis of Cholesterol-26-aldehyde. To a stirred solution of
(25S)-26-hydroxy cholesterol (4.2 mg, 0.01 mmol) in CH2Cl2 (2 mL)
was added Dess-Martin reagent (16 mg, 0.035 mmol). The resulting
mixture was stirred at room temperature under an inert atmosphere
for ∼2 h. Diethyl ether (6 mL) was added and the reaction was
quenched by the addition of NaOH (1N, 3 mL). The organic phase
was separated, the aqueous phase was extracted with diethyl ether (3 ×
5 mL), and the combined organic extracts were washed with NaOH
(10 mL), water and brine solution. The resulting organic phase was
dried over anhydrous Na2SO4, filtered and evaporated, giving an oil
that was purified by preparative TLC (2% methanol in chloroform, Rf
= 0.35) to yield ∼2 mg (4.5 μmol, 45%) of the product. The identity
of the product was confirmed by LC−MS analysis, which showed an
m/z of 383 amu for the [(M + H) − H2O]+ ion. The presence of an
aldehyde was further confirmed by a characteristic chemical shift at 9.6
ppm, corresponding to one aldehyde proton, in the 1H NMR
spectrum.
Enzyme Activity Assays. Substrate oxidation reactions were
carried out in glass tubes at room temperature (23 °C) in 50 mM
potassium phosphate buffer containing 0.05% (w/v) mβCD and 10
mM MgCl2. In a typical 50 μL reaction, CYP125, WT*, or SeCYP125
(0.5 μM) was preincubated with the appropriate substrate (0.1 mM)
for 5 min before the addition of spinach ferredoxin (1 μM), spinach
ferredoxin-NADP+ reductase (0.1 U/mL), bovine liver catalase (0.1
mg/mL), glucose-6-phosphate (4 mM), glucose-6-phosphate dehy-
drogenase (0.4 U/mL), and NADP+ (0.1 mM). At appropriate time
points, 50 μL aliquots of the reactions were quenched by the addition
of 150 μL of acetonitrile containing 0.1% formic acid. The reaction
mixtures were centrifuged at 11 000 rpm for 4 min and the
supernatants were directly analyzed by LC−MS as described below.
When necessary, cholesterol and other oxygenated cholesterol
substrates were preincubated for 60 min at room temperature in the
presence of 2.5 U/mL of cholesterol oxidase and 0.1 mg/mL of bovine
liver catalase to convert the metabolites into their cholest-4-en-3-one
derivatives. The UV−visible absorbance at 240 nm of the α, β-
unsaturated ketone in cholest-4-en-3-one facilitates detection of the
metabolites.
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Figure 2. Mass spectra of the CYP125 oxidation products. Black trace, cholest-4-en-3-one under an atmosphere of 16O2; green trace, cholest-4-en-3-
one under an atmosphere of 18O2; red trace; 25,26,26,26,27,27,27-d7-cholest-4-en-3-one under an atmosphere of 16O2. For clarity, only the m/z range
showing the parent ion is presented. The proposed structures of the metabolites are also shown, where R represents the rest of the cholest-4-en-3-
one structure.
Identification of Reaction Products by LC−MS. For qualitative
analysis and identification of the metabolites, the reaction mixtures
were analyzed by LC−MS using a Waters Micromass ZQ coupled to a
Waters Alliance HPLC system equipped with a 2695 separations
module, a Waters 2487 Dual λ Absorbance detector, and a reverse
phase C8 column (Varian Polaris C8-A, 5 μm, 4.6 × 250 mm). The
column was eluted isocratically at a flow rate of 0.5 mL/min (35%
buffer A, H2O + 0.1% formic acid (FA); 65% buffer B, CH3CN + 0.1%
FA). The elution was monitored at 240 nm. The mass spectrometer
settings were as follows: mode, ES+; capillary voltage, 3.5 kV; cone
voltage, 25 V; desolvation temperature, 250 °C. The HPLC peaks of
the individual metabolites were collected separately and analyzed by
MS using a quadrupole instrument in the positive ion mode. The MS/
MS analysis was performed at an Orbitrap XL instrument in the
Higher Energy Collision Dissociation (HCD) mode.
For quantitation of the relative amounts of oxidation products,
cholest-4-en-3-one (25 μM) was added as an internal standard for the
oxidation of cholesterol, 26-hydroxycholesterol and cholesterol-26-
aldehyde substrates. Similarly, 1,4-cholestadiene-3-one (25 μM) was
added for the oxidation of cholest-4-en-3-one. The samples were
analyzed by LC−MS as described above, but using a reverse phase
C18 column (Waters Xterra C18 column, 3.5 μm, 2.1 × 50 mm). The
contents were eluted at a flow rate of 0.5 mL/min (solvent A, H2O +
0.1% formic acid (FA); solvent B, CH3CN + 0.1% FA) with a gradient
starting at 70% B up to 1 min and the solvents ramped up to 100% B
over 12 min. The elution was maintained at 100% B up to 14 min and
then ramped back to 70% B within 1 min, followed by equilibration at
the same composition for 2 min before the next run. The cholesterol
and its derivatives were analyzed using the total ion current signal
obtained for the specific mass with selective ion monitoring. The
starting materials showed a clear concentration dependent increase in
the ion current signal. The individual metabolites were then
quantitated as a ratio of the integrated ion current signal of the
corresponding metabolite to the signal from the known concentration
of the internal standard.
Enzyme Kinetics. The assays for determining the steady-state
kinetic parameters were conducted as described above, but stopped at
earlier time points to measure the initial rates. The reaction conditions
were carefully optimized for the generation of a single product. All
assays contained 0.25 μM of the corresponding enzyme in a final
reaction volume of 300 μL, unless mentioned otherwise. For the first
hydroxylation step, 0.45% of mβCD was used as reported before.17
The kinetics for the second oxidation step was measured with 0.05%
mβCD as reported above for the end-point assays.
RESULTS
■
Oxidative Activities of CYP125. We previously reported
that the incubation of cholest-4-en-3-one with purified,
recombinant Mtb CYP125, surrogate redox partners from
spinach, and NADPH in the presence of 0.45% (w/v) mβCD
resulted in the formation of three major products correspond-
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a
Scheme 2. Possible Mechanisms for the Deformylation Reactions Catalyzed by CYP125
a
The important steps involve addition of the ferric−peroxo anion of CYP125 to the C26 carbonyl and subsequent radical fragmentation of the
peroxyhemiacetal adduct. The radical fragmentation of the peroxyhemiacetal adduct led to (a) the formation of an alkene (M1) or (b) the formation
of a one-carbon deficient alcohol (M4). The Compound I-catalyzed oxidation of M1 generates a diol (M5) via the acid-catalyzed ring-opening of an
epoxide intermediate. A C25 cation may also derive from the single-electron oxidation of the C25 radical (c). Trapping of the cation by formate or
water (e) results in the formation of the C25-oxyformyl (M2) and the one-carbon reduced alcohol (M4), respectively. Loss of a proton from the
C25 cation may also generate M1 (d). The Compound I-catalyzed oxidation of M4 produces a gem-diol intermediate that dehydrates to a keto
compound (M3).
Table 1. Molecular Weight, Mass of the Parent Ion, and Numbers of Retained Deuterium and 18O Atoms for the Metabolites
Obtained from the CYP125-Catalyzed Oxidation of Cholest-4-en-3-one
metabolite
peak mol. mass (Da) parent ion mass (m/z) number of retained D atoms number of retained 18O atoms
Cholest-4-en-3-one
S
384.6
400.6
398.6
414.6
368.6
414.6
384.6
386.6
402.6
385.3 [MH]+
401.3 [MH]+
399.3 [MH]+
415.3 [MH]+
369.3 [MH]+
415.3 [MH]+
385.3 [MH]+
387.3 [MH]+
385.3 [MH-H2O]+
7
6
5
4
3
5
3
4
3
n.a.
26-Hydroxycholest-4-en-3-one
III
II
1
Cholest-4-en-3-one-26-aldehyde
0
a
a
Cholest-4-en-3-one-26-acid
IV
I
1 or 2
27-Nor-cholest-4-en-3-one-26-ene (M1)
27-Nor-25-oxyformyl-cholest-4-en-3-one (M2)
27-Nor-25-oxo-cholest-4-en-3-one (M3)
27-Nor-25-hydroxy-cholest-4-en-3-one (M4)
27-Nor-25,26-dihydroxy-cholest-4-en-3-one (M5)
0
II
1 or 2
III
V
1
1
1
VI
a
Although two oxygen atoms are added, one can be exchanged with an oxygen of water, resulting in incorporation of only one labeled oxygen from
18O2.
ing to stepwise oxidation of the side-chain to the cholest-4-en-
3-one-26-acid.17,19 Interestingly, we observed the formation of
several additional products when the concentration of mβCD
was decreased to 0.05% (w/v). Lowering the mβCD
concentration decreases its competition for binding of the
sterol metabolites. As shown in Figure 1, at least six peaks (I−
VI) could be distinctly seen from a reverse-phase liquid
chromatographic analysis of the reactions of CYP125 with
cholest-4-en-3-one. Importantly, the fact that analysis of the
oxidations of authentic 26-hydroxy-cholest-4-en-3-one and
cholest-4-en-3-one-26-aldehyde by CYP125 yielded nearly
identical elution profiles (Figure 1), but that of cholest-4-en-
3-one-26-acid did not (data not shown), indicates that the
unknown metabolites originated from competitive, concom-
itant reactions of the 26-aldehyde intermediate.
Mass Spectrometric Characterization of the Unknown
Metabolites. We used LC−MS and 25,26,26,26,27,27,27-d7-
cholest-4-en-3-one to help determine the structures of the
metabolites. We also carried out the reactions under an
atmosphere of 18O2 to help elucidate the mechanisms of their
formation. Figure 2 displays the mass spectra of the three
expected products as well as for one major (M3) and four
minor (M1, M2, M4 and M5) unidentified metabolites
separated by liquid chromatography from the different
incubations. The proposed structures of these metabolites are
also presented. The masses of the parent and daughter ions of
all the metabolites, which were confirmed by MS/MS, are
summarized in Supporting Information Table S1. As expected,
three of the products correspond to the previously charac-
terized side chain oxidation metabolites, that is, 26-hydroxy-
cholest-4-en-3-one (peak III, m/z 401 [M + H]+), cholest-4-en-
3-one-26-aldehyde (peak II, m/z 399 [M + H]+), and cholest-
4-en-3-one-26-acid (peak IV, m/z 415 [M + H]+). The
identities of these metabolites were previously confirmed using
authentic standards. The identities of the new metabolites
(M1−M5) were established as discussed below.
27-Nor-25-oxo-cholest-4-en-3-one (M3). Close inspec-
tion of the mass spectrum from peak III (Figure 1) revealed the
presence of a second parent ion at m/z 385.3 (Figure 2). The
mass difference (16 Da) argues that the species at m/z 385.3 is
not a fragment ion of the 401.3 species. The mass difference of
three when the heptadeuterated substrate is used, indicating
only three deuterium atoms are retained, and of two when 18O2
is used, indicating the incorporation of one oxygen atom
(Figure 2), strongly suggest the metabolite is 27-nor-25-oxo-
cholest-4-en-3-one.
The formation of M3 can be rationalized by addition of the
ferric−peroxo anion to the 26-aldehyde followed by homolytic
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fragmentation to give Compound II and an alkoxy radical that
secondarily fragments to a demethylated carbon radical.
Recombination of the oxygen of Compound II with the carbon
radical yields the 27-nor-25-hydroxy product (pathway b,
Scheme 2). A second, conventional 25-hydroxylation of this
compound by CYP125 then yields a gem diol that dehydrates to
give the final 27-nor-25-keto metabolite. The intermediate 25-
hydroxy compound is also detected (M4, see below). In accord
with the data, this metabolite has lost four deuterium atoms and
incorporates an atom of molecular oxygen (Table 1).
27-Nor-25-oxyformyl-cholest-4-en-3-one (M2). Close
inspection of the mass spectrum of peak II revealed the
presence of two parent ions at m/z 399.3 and 415.3 (Figure 2).
The parent ion at m/z 399.3 corresponds to the 26-aldehyde
with which M2 co-elutes. Although the parent ion mass of m/z
415.3 is the same as that for the 26-acid, the large difference in
the retention time (>2 min) precludes its identification as a 26-
acid diastereomer. Moreover, the m/z value of the parent ion of
the deuterated metabolite is 420.3, which is 5 Da rather than 4
Da higher, so that one of the three deuterium atoms of the
methyl group that is oxidized is retained. This can be explained
if the metabolite is a 27-nor-25-oxoformyl derivative, as shown.
However, this metabolite has two oxygen atoms, but the 18O2
data suggests that only one 18O-atom is incorporated. This is
not unexpected, as the 26-aldehyde from which the formyl is
derived, like most aldehydes, can readily exchange its oxygen
with water from the medium, resulting in loss of one of the two
originally labeled oxygens.36
The formation M2 could occur via the trapping of a C25
carbocation by the formate ion generated in the deformylation
step. In turn, the carbocation could be generated by single-
electron oxidation of the C25 radical generated by homolytic
decomposition of a peroxyhemiacetal intermediate (pathway c,
Scheme 2). Alternatively, the formation of M2 could result
from a rearrangement of the peroxyhemiacetal adduct via an
ionic Baeyer−Villiger mechanism, similar to that proposed for
CYP51-mediated lanosterol demethylation (not shown).37
27-Nor-25-en-cholest-4-en-3-one (M1). A single metab-
olite M1 with an m/z of 369.3 was detected in peak I (Figure
1). Its low polarity, its retention of three deuterium atoms in
the reaction with the heptadeuterated sterol substrate, and the
absence of a mass increase in the presence of 18O2 suggest that
it is the 27-nor-25-ene derivative (Figure 2). Furthermore, the
presence of three atoms of deuterium in the deuterated product
(m/z 372.3) argues against formation of the isomeric 27-nor-
cholest-4-en-3-one-24-ene, a metabolite containing an internal
double bond between the C24 and C25 positions, as it would
retain four, and not three, deuterium atoms.
The formation of this product is readily rationalized by the
same manifold used to explain the formation of M2 and M3.
Homolytic fragmentation of the peroxyhemiacetal generated by
addition of the ferric−peroxo anion to the 26-aldehyde
generates a carbon radical that, by transfer of one electron to
the enzyme, is converted to a C25-cation (pathway c, Scheme
2). Loss of a proton (or deuterium) then produces the terminal
olefin (pathway d, Scheme 2). Alternatively, abstraction of a
terminal proton from the C25 radical intermediate by Cpd II
could also result in the direct formation of M1 (pathway a,
Scheme 2).
mechanism would require introduction of a double bond in our
sterol molecule between C24 and C25, which is not observed.
Alternatively, the hydroxylation could take place at the other
methyl group of the side chain, but only one of the products,
the terminal olefin M1, can be accommodated by this
mechanism. Furthermore, this mechanism is unlikely as it
involves an energetically unfavorable four-membered ring in the
transition state. Another mechanism that has been proposed
involves formation of the gem diol from the aldehyde (or from
the alcohol) followed by oxidation of this species by Cpd I-
mediated hydrogen abstraction, one electron oxidation, and C−
C bond cleavage to produce the alkene.38 However, the support
for this mechanism drawn from gas phase computational
studies has been negated by more recent studies by the same
group with a more sophisticated computational model.39
Another Cpd I mediated mechanism that involves initial
hydroxylation 3 carbons away from the aldehyde carbon to
form an alkene29 is untenable as it fails to generate the desired
product.
27-Nor-25-hydroxy-cholest-4-en-3-one (M4). A metab-
olite eluted in low abundance was detected in peak V. The MS
(Figure 2) and MS/MS (Table S1) of this metabolite revealed,
respectively, a parent ion [M + H]+ at m/z 387 and a daughter
ion [(M + H) − H2O]+ at m/z 369, in accord with a structure
containing an alcohol group. The lower mass and the shorter
retention time, compared to 26-hydroxy-cholest-4-en-3-one
(peak IV) suggest that the substrate underwent oxidative loss of
one of the two terminal methyl groups with incorporation of a
hydroxyl group at C25. We thus propose that this metabolite is
27-nor-25-hydroxy-cholest-4-en-3-one. This structure is con-
firmed by the retention of 4 deuterium atoms from the
heptadeuterated cholest-4-en-3-one substrate and by the
incorporation of one atom of 18O2 (+2 Da) (Table 1).
Interestingly, the MS analysis of the reaction performed in the
presence of 18O2 revealed the additional presence of this
metabolite without incorporation of the heavier oxygen isotope
(Figure 2). This observation could be explained by
contamination of the 18O2 with 16O2, but this is unlikely as
this contamination was not observed with any of the other
oxygenated metabolites.
The formation of M4 is most consistent with radical
decomposition of a peroxyhemiacetal adduct to form the C25
radical species that then recombines with the Compound II
ferryl oxygen (pathway b, Scheme 2). The detection of 16O2-
containing M4 metabolite when the reaction was performed
under an atmosphere of 18O2 is readily explained by oxidation
of the C25 radical to the cation, as already postulated above for
formation of the formyl adduct, followed by trapping by a water
molecule rather than formate ion (pathway e, Scheme 2).
27-Nor-25,26-dihydroxy-cholest-4-en-3-one (M5). Fi-
nally, in peak VI, we detected a major species with m/z value of
385 (Figure 2). This product retained three deuterium atoms of
the original seven in heptadeutero cholest-4-en-3-one and, from
the shift of 2 units with 18O2, incorporated one atom of
molecular oxygen (Table 1). We propose that this metabolite is
27-nor-25,26-dihydroxy-cholest-4-en-3-one for which the major
ion, as observed, corresponds to the loss of a water molecule
[(M + H) − H2O]+. This compound would result from
CYP125-catalyzed epoxidation of M1 followed by acid
catalyzed epoxide hydrolysis, which would introduce the
second oxygen from water rather than oxygen gas. In principle,
it could arise from hydroxylation of M4, but that would
introduce two atoms of molecular oxygen. Finally, CYP125-
Alternate mechanisms for the formation of an alkene
involving a Cpd I intermediate were considered for human
aromatase CYP19.29,38 One such mechanism involves hydrox-
ylation two atoms away from the aldehyde carbon, but this
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catalyzed oxidation of a synthetic, authentic sample of 27-nor-
25-en-cholesterol confirmed the epoxidation of the terminal
alkene and its subsequent acid-catalyzed ring opening to yield a
vicinal diol with a retention time similar to that of M5.
Expression, Purification, and UV−Vis Spectrophoto-
metric Characterization of SeCYP125. As CYP125
possesses three cysteines, we prepared the C172S/C429L
mutant to remove the two cysteines other than the one
coordinated to the heme iron atom. The selenium substituted
CYP125, SeCYP125, was expressed and purified from this
construct using minimal media in the presence of selenocys-
teine, as described elsewhere.34 About 3−4 mg/L of the
SeCYP125 was obtained after affinity purification, which is ∼4
fold less than the yield of the WT* protein expressed using the
media containing cysteine. Our selenocysteine incorporation
approach resulted in introduction of a selenocysteine only at
the iron coordinating position.
SeCYP125 exists as a mixture of ferric high spin (HS) and
low spin (LS) complexes with absorption maxima at 393 and
417 nm, respectively, along with broader peaks at 532 and 570
nm corresponding to the α and β bands, and a CT band at 643
nm. This contrasts with the WT* protein, which is
predominantly in the high spin form with a large peak at 395
nm bearing only a small shoulder at 416 nm (Figure 3A). On
the basis of the UV−vis spectrum, SeCYP125 appears to
contain about 40% of the heme in the low-spin form.
Replacement of the cysteine by selenocysteine clearly leads to
tighter binding of the distal water molecule coordinated to the
heme iron atom and an increased stability of the low-spin form.
The Fe(II)−CO complex of SeCYP125 with a Soret band at
459 nm (Figure 3B) exhibits a 9 nm red shift relative to that of
the WT* protein at 450 nm. As reported earlier for selenium
substituted CYP119 and CYP101,40,41 this red shift of the
Fe(II)−CO spectrum is consistent with introduction of the
electron rich selenolate as the proximal ligand.
Characterization of SeCYP125 by Resonance Raman
and EPR Spectroscopy. The resonance Raman (RR) spectra
of the ferric forms of WT* and SeCYP125 clearly indicate that
selenium substitution has stabilized the distal water bound low
spin species, in agreement with what was observed in the UV−
vis spectra. The RR spectrum of WT* CYP125 shows ν4, ν3,
and ν2 at 1373, 1488, and 1574 cm−1, respectively, and confirms
the predominance of the HS species over LS species identified
by ν3 and ν2 at 1502 and 1584 cm−1 (Figure 4). In contrast, the
Figure 4. Resonance Raman spectra of Fe3+ WT* and SeCYP125.
Room temperature RR spectra obtained with a 413 nm excitation.
LS marker bands are dominant contributions in the RR
spectrum of SeCYP125 compared to the WT* protein.
Substituting the heme iron axial sulfur with selenium in
SeCYP125 downshifts the ν4 mode 1 cm−1 which may reflect
an increased electron density on the antibonding porphyrin π*
orbitals, as previously observed with SeCYP119.40 The ν4
downshift in SeCYP125 is smaller than the 2-cm−1 downshift
observed in SeCYP119, but it may be due to the weak
dependence of this mode on the Fe3+ spin state.
The similarity of the impact of the sulfur to selenium
substitution in SeCYP125 and SeCYP119 is also evident from
the EPR spectra of the ferric low-spin species.40 The EPR
spectrum of WT* shows g values at 2.4, 2.25 and 1.94, as
previously reported.18 New resonances with increased g values
at 2.45, 2.29, and 1.98 dominate the EPR spectrum of
SeCYP125 (Figure 5). Minor contributions corresponding to
the WT* resonances indicate that the cysteine substituted
protein is present as a contaminant; While these minor signals
vary in intensity between different preparations, they
consistently remain below 20% of the total low spin signal.
The Fe3+−NO complexes of WT* and SeCYP125 were also
characterized by RR spectroscopy with 442-nm Soret
Figure 3. UV−vis spectrophotometric characterization of SeCYP125.
(A) Comparison of the native ferric spectrum of the WT* and
SeCYP125 proteins in 100 mM potassium phosphate buffer, pH 7.4.
(B) Fe2+−CO difference spectrum of the SeCYP125 measured over 12
min, at 1 min intervals, after the addition of 1 mM dithionite.
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Figure 5. EPR spectra of Fe3+ WT* and SeCYP125. EPR spectra of
Fe3+ WT* and SeCYP125 at 10 K. Microwave frequency, 9.66 GHz;
microwave power, 0.01 mW; modulation amplitude, 10 G.
excitation. In the low-frequency RR spectra of WT*, intense
vibrations at 521 and 549 cm−1 downshift with 15NO and are
assigned to mixtures of bending and stretching modes, that is,
ν(Fe−NO) and δ(Fe−N−O), respectively (Figure 6). While
substrate-free P450 heme Fe3+−NO complexes typically exhibit
a single band assigned to the ν(Fe−NO), substrate binding
promotes the enhancement and mixing of δ(Fe−N−O) mode
as the Fe−N−O unit deviates from linearity; this trend has
been particularly well documented in P450cam by Hu and
Kincaid.42 Presumably, distal pocket residues in CYP125
promote a bent Fe−N−O geometry even in the absence of
substrate. While the low-frequency RR spectra of the Fe3+−NO
complex in SeCYP125 are remarkably similar to those of WT*,
the sulfur to selenium substitution downshifts the ν(Fe−NO)
and δ(Fe−N−O) vibrations from 521 to 519 cm−1 and from
549 to 541 cm−1 (Figure 6). These modest downshifts are
comparable to those observed earlier with the nitrosyl complex
of SeCYP119.40 In the high-frequency region of the RR
spectrum of WT*, the ν(N−O) mode is identified at 1835
cm−1 as its 34-cm−1 15N-downshift matches the calculated value
based on an isolated diatomic oscillator. The sulfur to selenium
substitution downshifts the ν(N−O) mode to 1819 cm−1; an
equivalent downshift of the ν(N−O) mode was observed in
CYP119 upon substitution of the trans ligand.40 In addition to
the ν(N−O) modes, a second isotope sensitive signal is
observed above 1900 cm−1 in WT* and SeCYP125, but the
small 15N-isotope shifts these bands exhibits identified them as
combination mode between ν(Fe−NO)/δ(Fe−N−O) and the
ν4 mode which occur at 1375 cm−1 in the ferric−nitrosyl
complexes. The concomitant decreases of all vibrations
involving the NO upon substitution of the sulfur ligand with
selenium can be interpreted in terms of competing σ-trans
effect between the two axial ligands. As with our earlier study of
SeCYP119, the RR results support the notion that the sulfur to
selenium substitution increased electron donation from the
proximal ligand to the heme iron.
Figure 6. Resonance Raman spectra of Fe3+−NO complexes. Room
temperature RR spectra of the Fe3+−NO complexes of WT* and
SeCYP125 prepared with 14NO and 15NO. All spectra were obtained
with a 442 nm excitation. The low-frequency spectra were normalized
on the ν7 at 674 cm−1. The inset shows 14NO minus 15NO high-
frequency difference spectra obtained after normalization on the ν4
mode at 1375 cm−1.
typically results in displacement of the water molecule
coordinated to the heme iron and consequently in a
spectroscopically detectable low-spin to high-spin transition
known as a type-I shift. The KD values of WT* and SeCYP125
for cholest-4-en-3-one calculated from titration curves (Figure
7A) are 0.71 0.05 and 2.28 0.11 μM, respectively. A 3-fold
decrease in the binding affinity for SeCYP125 is consistent with
the greater stability of the water-bound low-spin state.
The enzymatic activity of SeCYP125 was analyzed in vitro
and was compared to that of the WT* enzyme. As shown in
Figure 7B, SeCYP125 also efficiently catalyzes the oxidation of
cholest-4-en-3-one, as judged by the appearance of six peaks in
the HPLC chromatogram. The activity of SeCYP125 toward
cholest-4-en-3-one is very similar to that of the WT* with 82%
of the substrate consumed after 60 min, compared to 86% for
WT*. MS analysis also confirmed that the metabolites
produced by SeCYP125 were identical to those generated by
the WT* enzyme (not shown). These findings clearly indicate
that replacement of the cysteine by a selenocysteine has not
altered the nature of the active site, as the substrate binding and
the net reactivity of the SeCYP125 remain highly comparable
to those of the WT* protein.
SeCYP125 Binds and Catalyzes the Oxidation of
Cholest-4-en-3-one. The binding and catalytic activities of
SeCYP125 were tested using cholest-4-en-3-one as a substrate.
The binding of a substrate-like molecule to P450 enzymes
Kinetic studies show that the kcat for formation of 26-hydroxy
cholest-4-en-3-one, calculated as 49 min−1 for the WT*
enzyme, is approximately 2-fold faster than the rate observed
for SeCYP125 (Table S3, Supporting Information). In contrast,
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the monooxygenation versus deformylation pathways. For this
purpose, we compared the oxidation of the cholesterol 26-
aldehyde itself by the WT* and SeCYP125 proteins. Focusing
on the oxidation of the cholesterol 26-aldehyde is desirable
because the aldehyde sits at the branching point between the
two pathways. The metabolites were quantitated as the ratio of
the total mass spectrometric ion current of the individual
metabolites to the signal from the known amount of internal
standard (Experimental Procedures). The effect of increased
electron donation was then determined by comparing the
amount of the 26-acid produced to the sum of the products
originating from the deformylation pathway for the WT* and
SeCYP125-mediated oxidations.
The oxidation of the cholesterol-26-aldehyde by the WT*
and SeCYP125 proteins generated similar products (Figure S1,
Supporting Information). The oxidation reaction was very
efficient as more than 90% of the substrate was oxidized by
both enzymes. More importantly, the oxidation of the 26-
aldehyde by SeCYP125 produced a significantly higher amount
of the 26-acid and lesser amounts of the deformylation
products, with 27-nor-25-oxo-cholesterol formed as the major
deformylation product. Overall, the ratio of the 26-acid to the
sum of the deformylated products was found to be ∼1.3-fold
higher with SeCYP125 than with the WT* protein (Figure 8).
It should be stressed that not only was the difference in the
ratio between the two enzymes highly reproducible, but
virtually identical differences in the ratio were observed when
either 26-hydroxycholesterol or cholesterol was used as the
substrate (Figure 8).
Figure 7. Binding and catalytic activities of SeCYP125 toward cholest-
4-en-3-one. (A) Binding of cholest-4-en-3-one to recombinant
SeCYP125. The concentration dependence of ligand binding was
deduced from the difference in absorption changes obtained from
titration of the protein (2.5 μM P450) with increasing concentrations
of cholest-4-en-3-one. The data was fit using a tight binding
equation.19 (B) LC chromatograms with absorbance detection at
240 nm of the WT* and SeCYP125 oxidation products of cholest-4-
en-3-one, separated using a reverse phase C-18 Xterra column
(Waters), as described in the Experimental Procedures. A control
reaction carried out in the absence of NADPH is also included. Inset:
zoom-in showing the HPLC elution of the minor products. S and I.S.
stand for substrate and internal standard, respectively.
the steady-state rate constant measured for the second
oxidation from the 26-hydroxy to 26-aldehyde was approx-
imately 3-fold faster for SeCYP125 than for the WT*. Although
the rates are comparable between the WT* and SeCYP125 for
each step, they are significantly different for the individual
enzymes. In contrast, the Km values for binding of the two
substrates to the WT and SeCYP125 enzymes are very similar.
Product Distributions in the Turnover of Cholesterol-
26-aldehyde by WT* and SeCYP125. WT* and SeCYP125
clearly produce a 26-aldehyde intermediate that can undergo
monooxygenation to generate the corresponding 26-acid. As
shown here, the aldehyde is also oxidized via a peroxyhemia-
cetal intermediate to the C−C bond cleavage products M1−
M5, providing an opportunity to investigate the influence of
increased electron donation from the proximal iron ligand on
Figure 8. Comparison of the ratio of the 26-acid over the sum of the
deformylation products produced during the oxidation of various
substrates by WT* and SeCYP125 after 60 min incubation.
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In contrast, although the ratios between WT and SeCYP125
were identical for cholesterol and its 26-alcohol and 26-
aldehyde products, a smaller increase of ∼1.2-fold was observed
when cholest-4-en-3-one was used as the substrate. Further-
more, the ratios of the acid to deformylation products were not
identical for the same enzyme when compared for different
substrates. Thus, ratios of 1.6, 1.4, and 1.2 were observed for
the WT* enzyme with the cholesterol, 26-hydroxy and 26-
aldehyde substrates, respectively (Figure 8).
the aldehyde is provided as a substrate (Table S2) as when the
aldehyde is generated internally from cholesterol or 26-hydroxy
cholesterol through initial oxidative turnovers. In this system,
the formation of products from the Compound I-mediated
monooxygenation and peroxyhemiacetal-mediated deformyla-
tion pathways is controlled by the intrinsic reactivity of the
ferric−peroxo anion intermediate. To investigate the effect of
increased electron donation from the proximal ligand on
partitioning of the aldehyde intermediate into the two reaction
pathways, we expressed CYP125 with a selenocysteine
replacing the cysteine that provides the proximal thiolate iron
ligand.
The introduction of selenocysteine (SeCys) as the proximal
iron ligand in a cytochrome P450 enzyme has recently been
reported.40,41 Unlike other proximal ligand substitutions, the
SeCys proteins retain spectroscopic and catalytic properties
comparable to those of their cysteine counterparts, although
increased electron donation by the selenolate versus thiolate
ligand does cause some differences. These include a shift to
longer wavelengths of the Soret maxima of the Fe2+−CO
complexes and a lowering of the redox potentials of the
enzymes.34,41 In agreement with these earlier findings, the Soret
maximum of the SeCys-substituted CYP125 utilized in these
studies was at 459 nm rather than the 450 nm found for the
WT* protein. The position of this Soret indicated a high degree
of incorporation of SeCys into the protein, an inference
supported by the near absence of EPR resonances from the
WT* enzyme in the EPR spectrum of SeCYP125 (Figure 5).
Furthermore, comparison of the resonance Raman spectra of
the ferric and ferric-nitrosyl hemes in WT* and SeCYP125
proteins show that the SeCys substitution increases electron
donation to the metal iron.
DISCUSSION
■
As shown here, oxidation of the cholesterol side-chain by
CYP125 leads to the formation not only of the C26-acid, as
previously reported,19 but also of five additional products
(M1−M5) resulting from C−C bond cleavage. The formation
of these additional products was uncovered by varying the
incubation conditions and carefully analyzing the product
HPLC peaks. The identities of the metabolites were established
by comparison with authentic standards or by mass
spectrometry in combination with deuterium labeling and the
use of 18O2 in the oxidation reaction. The mechanisms
proposed for formation of the C−C bond cleavage products
(Scheme 2) are fully consistent with the deuterium and oxygen
labeling results and with the general mechanisms proposed for
such reactions.29−31,43 None of the reactions leading to the
observed products were entirely unprecedented, but the co-
occurrence of this diversity of products in the catalytic turnover
of a single substrate by a P450 enzyme is highly unusual. This
product diversity has provided us with a tool with which to
examine the effect of electron donation from the proximal
ligand on formation of the Compound I species.
The role of Compound I in monooxygenation reactions is
well documented. Similarly, growing evidence points to the
ferric−peroxo anion intermediate as the active oxidant in C−C
bond cleavage reactions. All this evidence clearly supports the
role of the ferric−peroxo anion in the generation of
deformylation products. Thus, the feasibility of the reaction is
established by nonenzymatic deformylation of the 19-aldehyde
of androstenedione by a porphyrin ferric−peroxo anion model
complex.44 Trapping of the ferric−peroxo anion in CYP19
using cryo-freeze quench EPR methods confirms the biological
existence of this intermediate and bolsters its potential role in
deformylation reactions.45 More directly, the use of radio-
labeled substrates and 18O2 has clearly established the
involvement of the ferri−peroxo anion in the formation of
acyl-carbon bond cleavage products by CYP5131 and CYP17,43
results that specifically rule out the participation of a
Compound I intermediate. Furthermore, consistent with
nucleophilic attack of the peroxo anion on the aldehyde, a
distal H-bonding mutant in CYP2B4 that blocks the
protonation of the peroxy anion, generated more deformylated
metabolites relative to the Compound I-mediated monooxyge-
nation product.46 The peroxy anion mechanism was recently
supported by a detailed theoretical analysis.39 Finally, in the
present case, the total number of deuterium atoms retained by
the metabolites after the oxidation of cholesterol with the
heptadeuterated side chain, excludes alternative mechanisms
involving a Compound I intermediate. The branchpoint
intermediate in generating the acid and all the C−C bond
cleavage products is the C26 aldehyde (Scheme 1). On the
basis of literature precedent, it is likely that the 26-aldehyde acts
as the common precursor of all the end products since the same
products, and similar product distributions, are obtained when
More importantly, SeCYP125 binds cholest-4-en-3-one with
a similar affinity to that of the WT* enzyme and oxidizes this
substrate to the same products, albeit not in exactly the same
ratios. When the reaction is run with cholesterol 26-aldehyde as
the substrate, both enzymes consume the substrate to
essentially the same degree after 60 min, so the differences in
product ratios are not due to differential extents of substrate
utilization. This result also argues that factors such as
differential uncoupling of the two enzymes is not a factor in
determining the differences in the product ratios, as higher
uncoupling would lead to slower consumption of the aldehyde.
It should also be emphasized that the ratio of the products
starting with the aldehyde is the most relevant to the
conclusions of this paper, as it directly compares the reactions
at the branchpoint without any confounding factors that might
arise from a requirement for multiple turnovers prior to
generating the aldehyde. With both enzymes, all the C−C bond
cleavage products are formed, but the 26-acid metabolite is the
major product. The primary finding is that the SeCYP125-
mediated oxidation produces more cholesterol 26-acid relative
to the sum of the C−C bond cleavage products than the WT*.
The 1.3-fold increase in the ratio of the 26-acid to
deformylation products indicates that selenocysteine substitu-
tion favors the formation of Compound I. The oxidation of
cholesterol and 26-hydroxycholesterol, the two precursors of
the aldehyde, by SeCYP125 also yielded an ∼1.3-fold increase,
relative to the WT*, in the ratio of the acid to homolysis
products (Figure 8). Finally, oxidation of 25,26,26,26,27,27,27-
d7-cholesterol by the WT* enzyme was examined. The results
show that even with deuterated cholesterol, the amount of acid
produced is enhanced relative to the sum of deformylation
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products. This result argues against any mechanism in which
both carboxylic acid formation and decarboxylation are
mediated by the ferryl species.
observed here. To our knowledge, this is the first direct
evidence emphasizing the role of increased electron donation
from the proximal ligand leading to enhanced electron density
on the distal oxygen of the ferric-peroxo anion intermediate.
Protonation of the ferric−peroxo anion depends, of course,
on the availability of a suitable proton donor on the distal side
of the heme. Thus, a number of studies have shown that
replacement of a highly conserved distal threonine thought to
be involved in this process leads to decreased catalysis and
increased uncoupling.46,49 However, in the present instance, the
distal Thr272 and its environment are unchanged and cannot
be the cause of the difference in protonation of the ferric−
peroxo anion.
The change of 1.2- to 1.3-fold in the ratio of the acid to
deformylation products is significant but modest. Importantly,
identical ratios are reproducibly observed with both cholesterol
and cholest-4-en-3-one, as well as when the 26-alcohol and 26-
aldehyde derivatives are used instead of the parent steroids as
the substrates. It is difficult to predict the change in the pKa
value of the distal oxygen of the ferric−peroxo anion
intermediate when a selenolate replaces the thiolate ligand,
but the change is not likely to be large. Thus, the expected
changes in product ratios should be modest, as observed.
Moreover, as the increased electron density on the peroxy
anion increases its nucleophilic character, it may promote its
addition to the aldehyde and lower the acid to deformylation
product ratios.
Although the differences in the product ratios obtained with
the WT* and SeCYP125 enzymes are essentially identical
among the cholesterol derivatives, the product ratios for the
same enzyme varied significantly for the different substrates.
The steady-state amounts of the cholesterol 26-aldehyde
remaining at the end of the 60 min incubation of WT*
enzyme with cholesterol and 26-hydroxy cholesterol were
calculated to be ∼5 and 2.8 μM, respectively. Similarly for
SeCYP125, the amounts of cholesterol 26-aldehyde remaining
were ∼5.8 and 3.1 μM. Although more aldehyde reacted when
26-hydroxy cholesterol was the substrate, overall the amounts
of aldehyde reacted with WT* and SeCYP125 at the 60 min
time point were comparable.
The ratio of the products differed when the substrate was
changed from cholesterol or its 26-oxidized products to cholest-
4-en-3-one. This finding is not surprising, as there is no reason
to expect that the 3-keto-26-aldehyde generated from cholest-4-
en-3-one will bind in the active site and will interact with the
protein residues in exactly the same manner as the 3-hydroxy-
26-aldehyde of cholesterol. Nevertheless, it is clear that with
this substrate the carboxyl-forming branch of the reaction is
also favored by selenolate ligation to the iron. More surprising
is the finding that the ratio of products exhibited some variation
for a given enzyme when comparing the degradation of
cholesterol, 26-hydroxycholesterol, and the cholesterol 26-
aldehyde. A constant ratio would be expected if no factors
intervened other than the intrinsic partitioning of the product
pathways at the aldehyde stage. There is no evidence for
allosteric effects of cholesterol or its oxidation products on
CYP125 catalysis. It is possible that this variation results from a
channeling effect that occurs when the substrate is cholesterol
or 26-hydroxycholesterol. As shown by the UV−vis binding
experiments, a water molecule is displaced from the heme iron
on binding of the substrate. This water molecule may slightly
perturb the hydrogen bonding pattern in the active site during
catalytic turnover, but more importantly, this perturbation may
not be the same when a substrate such as cholesterol binds to
the enzyme two catalytic turnovers prior to the one that
controls the product ratio. Of course, this argument only
applies if, at least to some extent, the substrate is processed
through multiple turnovers without dissociating from the
protein. Interestingly, channeling in the oxidation of aldehydes
generated in situ from precursor molecules has been reported
in the reactions of CYP2A6.47
The extensive partitioning between monooxygenation and
deformylation of the C26 aldehyde seen in the oxidation of
cholesterol by CYP125 is unique among the known steroid
C26-monooxygenases, as CYP27A1, CYP124, and CYP142
produce the alcohol, aldehyde, and acid metabolites without
detectably forming the deformylation products.16,17,19,32,50
Thus, the ferric−peroxo anion intermediate in these latter
enzymes is either protonated much more quickly than in
CYP125, or its possible addition to the aldehyde carbonyl
group is suppressed within the active site.
CONCLUSION
■
In conclusion, we demonstrate the unique ability of CYP125A1
to catalyze the oxidation of the cholesterol side chain not only
to the acid, but also to five other metabolites stemming from
the deformylation of the intermediate aldehyde. Furthermore, a
proximal selenocysteine substituted CYP125 enzyme was
expressed, purified, and characterized using various spectro-
scopic methods. This enzyme was utilized to probe the effect of
excess electron donation by the proximal ligand on the
partitioning of the products between heterolytic (Compound
I) and homolytic (deformylation) pathways. These experiments
reveal that the increased electron donation from the proximal
ligand to the iron favors protonation of the ferric−peroxo anion
intermediate to form Compound I.
Partitioning of the ferric−peroxo anion intermediate between
conversion to Compound I (leading to acid formation) and
addition to the aldehyde to give a peroxyhemiacetal (leading to
C−C bond cleavage) hinges on the extent to which the
intermediate is present as the peroxy anion versus the
hydroperoxide. Increased electron donation from the proximal
thiolate to the iron bound dioxygen is proposed to increase the
pKa of Compound 0.27 Computational studies suggest that the
more electron-donating selenolate proximal ligand should
enhance the rate of formation of Compound I.48 In our
studies, an increase in the pKa of the peroxy anion would result
in a shift from the anionic toward the hydroperoxo form, and
this shift would result in increased formation of Compound I
and decreased formation of the peroxyhemiacetal intermediate.
This predicts an increased oxidation of the aldehyde to the
carboxylic acid at the expense of the deformylation products, as
ASSOCIATED CONTENT
■
S
* Supporting Information
Tables with LC−MS and MS/MS data, kinetic constants for
the reactions, LC spectrum of the 26-aldehyde derivative, and a
representative kinetic plot are available. This material is
available free of charge via the Internet at http://pubs.acs.org.
6683
dx.doi.org/10.1021/ja211499q | J. Am. Chem. Soc. 2012, 134, 6673−6684

Journal of the American Chemical Society
Article
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AUTHOR INFORMATION
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Corresponding Author
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ortiz@cgl.ucsf.edu
Present Address
‡Department of Biological Sciences and The Border Biomedical
Research Center, University of Texas, 500 W. University, El
Paso, Texas 79968
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Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS
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We thank Arti Singh and Prof. James De Voss of the University
of Queensland for kindly providing us with the synthetic
standard of 27-nor-25-en-cholesterol and Dr. Jonathan
Johnston for helpful discussions and suggestions. This research
was supported by National Institutes of Health Grants
GM25515 and AI074824 (to P.R.O.M.) and GM74785
(P.M.-L.), and by National Center for Research Resources
Grants P41RR001614 and RR019934 [to the Bio-Organic
Biomedical Mass Spectrometry Resource at the University of
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