Báo cáo khoa học: The role of cytochrome P450 monooxygenases in microbial fatty acid metabolism pdf

MINIREVIEW
The role of cytochrome P450 monooxygenases in
microbial fatty acid metabolism
Inge N. A. Van Bogaert
1
, Sara Groeneboer
2
, Karen Saerens
1
and Wim Soetaert
1
1 Department of Biochemical and Microbial Technology, Laboratory of Industrial Biotechnology and Biocatalysis, Ghent University,
Ghent, Belgium
2 Laboratory for Protein Biochemistry and Biomolecular Engineering, Ghent University, Ghent, Belgium
Introduction
P450 classification and nomenclature
Cytochrome P450s form a vast and divergent family of
enzymes. They are heme–thiolate proteins, bearing, in
a hydrophobic pocket, a protoporphyrin IX linked to
the apoprotein by a bond between the heme iron cen-
tre and the sulfur atom of a conserved cysteinyl resi-
due. In a typical reaction catalysed by a P450,
molecular oxygen binds to the heme iron for activation
before transfer to the substrate. Carbon monoxide can
also bind, leading to a reduced P450 producing a char-
acteristic CO-binding difference spectrum with an
absorbance maximum at 450 nm. This P450–CO com-
plex is inactive and has given the name to P450 (pig-
ment absorbing at 450 nm) [1]. Inhibition by CO and
reversion of this inhibition by 450 nm light are charac-
teristic for most reactions catalysed by P450s.

New genes are annotated as a P450 based on the
presence of typical conserved domains involved in
heme binding and proton transfer [2]. P450s have been
categorized in families and subfamilies. They belong to
the same family when they share ‡ 40% amino acid
identity and they belong to the same subfamily when
they share ‡ 55% amino acid identity [3]. For example,
Keywords
alkanes; bacteria; biosurfactants;
cytochrome P450 monooxygenase;
dicarboxylic acids; fatty acids; hydroxylation;
oxylipins; polyketides; yeast
Correspondence
I. N. A. Van Bogaert, Department of
Biochemical and Microbial Technology,
Laboratory of Industrial Biotechnology and
Biocatalysis, Faculty of Bioscience
Engineering, Ghent University, Coupure
Links 653, B-9000 Ghent, Belgium
Fax: +32 9 264 62 31
Tel: +32 9 264 60 34
E-mail:
Website:
(Received 22 June 2010, revised 19 August
2010, accepted 16 September 2010)
doi:10.1111/j.1742-4658.2010.07949.x
Cytochrome P450 monooxygenases (P450s) are a diverse collection of
enzymes acting on various endogenous and xenobiotic molecules. Most of
them catalyse hydroxylation reactions and one group of possible substrates
are fatty acids and their related structures. In this minireview, the signifi-

cance of P450s in microbial fatty acid conversion is described. Bacteria and
yeasts possess various P450 systems involved in alkane and fatty acid deg-
radation, and often several enzymes with different activities and specificities
are retrieved in one organism. Furthermore, P450s take part in the forma-
tion of fatty acid-based secondary metabolites. Finally, there are a substan-
tial number of microbial P450s displaying activity towards fatty acids, but
to which no biological role could be assigned despite the often quite intense
research.
Abbreviations
GPo1, alkane hydroxylase; P450, cytochrome P450 monooxygenase; psi factor, precocious sexual inducers.
206 FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS
CYP94A1 from the plant Vicia sativa is the first mem-
ber of subfamily A of family 94 (there is more about
the activity of this enzyme in the accompanying mini-
review by Pinot & Beisson [4]). Over 11 000 P450
genes are included in this internationally used nomen-
clature system, and probably an extensive number of
nonclassified genes are present in the emerging pile of
genome sequencing data. Besides being vast, the P450
superfamily is also highly divergent. For example, in
plant P450s, CYP94 identity may be as low as 16%
with CYP51 and even only 12% with CYP74. Diver-
gence is further illustrated by the huge number of fam-
ilies (977) and subfamilies (2519).
Phylogenetic studies have suggested that the unicel-
lular ancestor of plants had at least three distinct P450
branches [2]. The first, sharing a common ancestor
with animal P450s involved in xenobiotic metabolism,
gave rise to Group A, catalysing typical plants reac-
tions (i.e. synthesis of lignin, flavonoids, etc.). Group

non-A is composed of two branches. The first origi-
nates from a common ancestor with animal CYP4 and
fungal CYP52 fatty acid hydroxylases; plant fatty acid
hydroxylases of the families CYP86 and CYP94 origi-
nate from this branch. The second branch of Group
non-A shares a common ancestor with animal, fungal
and bacterial sterol oxidases (CYP51). This branch
gave rise to the plant obtusifoliol 14 demethylase
(CYP51) and brassinolide hydroxylases (CYP85 and
CYP90 families).
Microbial P450s and fatty acids
Fatty acids are simple, yet indispensable, molecules to
any living cell. Incorporated in phospholipids, they
make up the major part of the plasma membrane and
besides structural roles, they also function as a carbon
or energy source. Furthermore, modified fatty acids
are building blocks for other complex molecules or act
as signalling molecules to trigger physiological
changes. All these roles and processes require specific
enzymes, and cytochrome P450 monooxygenases
(P450s) make up a significant part of them. P450s are
heme–thiolate proteins involved in the hydroxylation
of a wide range of endogenous and xenobiotic com-
pounds. They are present in every eukaryotic organism
and in a substantial number of prokaryotes. More
than 3800 microbial P450s are known to date [5] and
we estimate that  10–17% of them display activity
towards fatty acids or related structures. On the one
hand, these activities are linked to the degradation of
fatty acids and alkanes; metabolization of these latter

compounds is inherently coupled to fatty acid degrada-
tion because the conversion of alkanes to fatty acids is
an essential step in the alkane assimilation process. On
the other hand, P450s are also involved in the synthe-
sis of special fatty acid-based molecules such as sec-
ondary metabolites or signal molecules. Although the
P450s described in this review all act on fatty acid sub-
strates, this is not reflected in their overall similarity;
according to Nelson’s classification system based on
amino acid identity they belong to various families [5].
Besides the involvement in different physiological
functions, P450s also differ in the position of the
hydroxylation; this may occur close to the carboxyl
group, giving rise to a-orb-hydroxylated fatty acids
(mediated by CYP152), in-chain (e.g. CYP1006) or at
the terminal or subterminal ending (e.g. CYP52). Sev-
eral classes of P450s involved in either metabolization
or biosynthesis processes and with different regiospeci-
ficities are discussed. Whenever possible, the interna-
tionally used nomenclature is applied [3].
Fatty acid metabolism
Alkane degradation
Microbial populations can break down almost every
natural organic compound. Even alkanes, which from
a chemical point of view are almost inert molecules,
can be degraded and utilized as a carbon source by
both bacteria and fungi. Traditional aerobic alkane
assimilation is initiated by terminal hydroxylation. In
the subsequent oxidation steps, the corresponding pri-
mary alcohol is converted via an aldehyde to a fatty

acid, which will enter b-oxidation [6]. Depending on
the particular microorganism, initial oxidation can be
governed by several unrelated alkane hydroxylases.
Although microbial alkane degradation was first dem-
onstrated about a century ago [7], research on this
topic was only really boosted in the 1950s and 1960s
when the production of single cell protein based on
paraffin or alkanes became a hot topic. The first alkane
hydroxylase (GPo1) was found in the early 1960s in
the soil bacteria Pseudomonas oleovorans (later
renamed Pseudomonas putida) and was shown to be an
integral membrane-bound nonheme di-iron monooxy-
genase [8]. More recently, related genes have been iso-
lated from a broad range of bacteria [9]. Whereas
various yeasts were also well established as single cell
protein producers, it took another few decades before
their corresponding alkane hydroxylase enzymes were
identified as cytochrome P450 monooxygenases, unre-
lated to the bacterial GPo1 system [10]. To date, all
yeast alkane hydroxylases belong to the CYP52 family
(membrane-bound class II). This family contains sev-
eral enzymes with demonstrated activity towards
I. N. A. Van Bogaert et al. The role of P450 in microbial fatty acid metabolism
FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS 207
alkanes and ⁄ or fatty acids, but also harbours numerous
genes which have not been explored (yet) or are proven
to be pseudogenes. Well-studied members of the
CYP52 family are the enzymes of the alkane-metabo-
lizing yeasts Candida tropicalis, Candida maltosa and
Yarrowia lipolytica. C. tropicalis has at least 18 genes

belonging to this family. Craft et al. [11] evaluated 10
of them by quantitative competitive reverse transcrip-
tion-PCR; this highly specific technique is required
because of the high identity between the genes. Five
genes were clearly induced by octadecane (CYP52A12,
-13, -14, -17 and -18), whereas no transcription was
detected for CYP52A15, -16, -19, -20 and D2 under
those and other conditions. Also, C. maltosa possesses
several so called P450alk genes: alk1 to alk8. Four
genes are considered to be the primary P450alk genes:
alk1, alk2, alk3 and alk5, corresponding to CYP52A3,
-A5, A4 and -A9, respectively. Quadruple mutants
were unable to grow on alkanes as a sole carbon
source, but complementation by one of the four genes
restored growth on hexadecane [12]. The corresponding
enzymes differ in their substrate chain-length specific-
ity, giving a possible reason for the multitude of
P450alk genes often present in one organism.
Although the induction of P450alk genes in response
to n-alkanes or fatty acids is a common feature among
alkane-assimilating yeasts, the underlying molecular
mechanisms remain largely unknown. C. maltosa alk2
turned out to be inducible by alkanes, as well as by
the peroxisome proliferator clofibrate. The respective
cis-acting elements in the alk2 promoter region were
identified and there are indications of similar motives
in other C. maltosa alk promoters [13].
Y. lipolytica is another well-studied alkanotrophic
yeast with 12 P450alk genes in its genome. Six of the
eight tested genes showed induction by alkanes.

Among them, alk1 (CYP52F1) displayed the highest
expression and although single disruptions in the other
genes did not result in yeasts unable to metabolize alk-
anes, Dalk1 mutants are unable to grow on decane. In
addition, a Dalk1Dalk2 double mutant cannot grow on
hexadecane either. Therefore, it is suggested that the
primary P450alk gene alk1 is required for assimilation
of decane and dodecane, whereas alk2 (CYP52F2) is
involved in the assimilation of molecules longer than
dodecane. The other alk genes possibly act on even
longer alkanes or other types of carbon chains [14].
Recently, Ohta’s group shed light on the transcrip-
tional induction of the alk1 gene by alkanes [15]; in
the presence of alkanes, Yas1p and Yas2p, two basic
helix–loop–helix proteins form heterodimers and bind
to the cis-acting alkane-responsive element 1 in the
alk1 promoter. The protein complex also binds to
other promoters of genes interfering in alkane degrada-
tion such as the acetoacetyl-CoA thiolase gene and the
yas1 gene itself. This latter binding creates a positive
autoregulatory feedback which results in a quick and
profound transcriptional response to alkanes. Further-
more, a third regulatory protein was identified. The
repressor Yas3p binds specifically to Yas2p when no
alkanes are present, but when the yeast is exposed to
alkanes, Yas3p is translocated from the nucleus to the
endoplasmatic reticulum. Conserved motives of the
alkane-responsive element 1 sequence were retrieved in
the P450alk promoters of C. maltosa, C. tropicalis and
Debaryomyces hansenii, suggesting a common mecha-

nism for alkane-responsive induction.
Although Cardini and Jurtshuk [16] provided strong
indications for the involvement of a bacterial P450 in
the hydroxylation of octane in Rhodococcus rhodoch-
rous, the role of bacterial P450s in alkane degradation
in addition to the well-established alkane hydroxylase
system has long been underestimated. The first bacte-
rial P450alk was cloned from Acinetobacter calcoaceti-
cus in 2001, this class I P450 was assigned to the new
family CYP153 [17]. Recently, similar enzymes were
found in other alkane-utilizing species such as Sphingo-
monas sp. HXN200, Mycobacterium sp. HXN1500 and
Alcanivorax borkumensis. Van Beilen et al. [18] were
able to demonstrate the functionality of seven of ele-
ven genes (CYP153A6, -A7, A11, A13, A14 and -D1)
by functional expression in a DGPo1 P. putida strain
and its restored ability to grow on alkanes. Most alk-
anotrophes attack various or mixed substrates with
different and often length-specific enzymes, reflected in
the variation among yeast CYP52 genes and the bacte-
rial alkane hydroxylase system. A similar trend can be
observed for the bacterial P450alk enzymes. For exam-
ple, Sphingomonas sp. HXN200 possesses five CYP153
genes, three of which show activity towards C5–C10
alkanes, whereas no affinity for these substrates was
observed for the other two genes, suggesting that these
genes are either pseudogenes or act on different sub-
strates such as long-chain alkanes. Other organisms
possess several types of enzymes; Al. borkumensis con-
tains two alkane hydroxylases and two CYP153s [19]

and whereas the first alkane hydroxylase is essential
for the degradation of C6, no clear role could be
assigned to the second, but double knockouts resulted
in deficient growth on C8–C16. Because this organism
is, thanks to its efficient and broad-spectrum hydrocar-
bon-degrading capacities, a dominant microbe in oil-
polluted waters, the CYP153 enzymes are postulated
to cover the rest of the alkane length range.
In general, one associates alkane-degrading organ-
isms with oil-polluted environments, but in fact
The role of P450 in microbial fatty acid metabolism I. N. A. Van Bogaert et al.
208 FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS
alkane-degrading enzymes are found in many more
organisms than the ones strictly appearing in oil-
related niches. Indeed, alkanes are persistent molecules
and this property makes them the perfect components
for natural barriers such as plant and insect cuticles.
The major component of plant cuticles is cutin, a poly-
mer consisting of omega hydroxy fatty acids cross-
linked by ester and epoxide bonds, which is impreg-
nated and covered with waxes (more information on
the constitution and biosynthesis of plant envelopes
can be found in the accompanying minireview by Pinot
& Beisson [4]). These cuticular and epicuticular waxes
are a mixture of long-chain alkanes (C16–C30) and
related structures. For example, in the rice blast fungus
Magnoporthe oryzae, a putative alkane-degrading cyto-
chrome P450 (MGG_05908.5 or CYP584A2) is upreg-
ulated upon the first stages of infection, together with
other genes regarding the utilization of nonconvention-

al carbon sources [20]. These findings suggest that, on
the one hand, alkane degradation is required for
breaking the plant’s defence but, on the other hand,
alkanes serve as nutritional input during the initial col-
onization steps. Insect cuticle consists of protein and
chitin, and is covered by a highly resistant lipid layer:
the epicuticle. This epiculticle is composed of hydro-
carbons, wax esters, fatty alcohols and fatty acids.
Hydrocarbons are the prevalent component and
include alkanes, alkenes and methyl-branched chains
in various ratios, depending on the insect species. In
general, alkanes make up the biggest part of the
hydrocarbon moiety and their chain length ranges
between C21 and C35, with a particular preference for
odd-numbered chains [21]. The entomopathogenic fun-
gus Metarhizium anisopliae infects a broad range of
insects by direct penetration of the host cuticle and
hence can be exploited as a biological control agent of
pests. cDNA microarray analysis of the fungus grown
on cuticular extracts revealed a clear upregulation of
an alkane-inducible cytochrome P450 (AJ273607)
during the first hours of incubation [22]. According to
amino acid similarity, this gene should be classified in
the CYP52 family. Several other expressed sequence
tag (EST) or microarray cDNA analysis regarding en-
tomopathogenic infection concealed involvement of
P450s supposed to hydroxylate alkanes and ⁄ or fatty
acids.
Fatty acid degradation – omega oxidation
Although cytochrome P450s do not intervene in the

degradation of fatty acids in the b-oxidation cycle
itself, they do take part in the steps preceding b-oxida-
tion (Fig. 1). As mentioned in the previous section,
alkanes can be converted to common fatty acids by
initial cytochrome P450 interference. However, the
same enzymes responsible for terminal alkane oxida-
tion often also mediate subterminal oxidation. The
corresponding secondary alcohols are oxidized to ketones
and a Baeyer–Villiger monooxygenase converts them
to esters, which are cleaved to give rise to a primary
alcohol and a common fatty acid.
Furthermore, common fatty acids can be terminally
oxidized by a cytochrome P450 monooxygenase and
the resulting hydroxy-fatty acid is further converted to
a dicarboxylic acid, which then enters b-oxidation.
This so-called x-oxidation occurs both in bacteria and
yeasts, and yet is far more documented for this latter
group because medium- and long-chain dicarboxylic
acids are commercially produced by yeast fermenta-
tions to serve as building blocks of, for example, per-
fume, polymers, high-quality lubricants or macrolide
antibiotics.
In this respect, the previously discussed fungal
CYP52 enzymes are versatile enzymes; several isoforms
exhibit different activities and specificities towards alk-
anes, as well as towards fatty acids or related struc-
tures. An individual CYP52 not only demonstrates a
distinct substrate specificity regarding chain lengths,
Fig. 1. Assimilation of alkanes and fatty
acids. The (putative) involvement of cyto-

chrome P450 monooxygenases is indicated
by grey arrows.
I. N. A. Van Bogaert et al. The role of P450 in microbial fatty acid metabolism
FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS 209
but in addition has preferences for either alkanes or
fatty acids, and similarly, compared with the chain
lengths, alkane ⁄ fatty acid specificities of the various
enzymes within species are often overlapping. The
C. maltosa P450Cm1 (ALK1 or CYP52A3), for exam-
ple, prefers alkanes, whereas P450Cm2 (ALK3 or
CYP52A4) shows highest affinity towards fatty acids.
Nevertheless, both enzymes are able to hydroxylate
both substrates. However, the C. maltosa CYP52A10
and CYP52A11 enzymes are only able to convert fatty
acids [23,24]. It is not completely clear which gene(s)
exactly mediate dicarboxylic acid formation, but Kog-
ure et al. [25] demonstrated that the alk5 gene
(CYP52A9) was highly induced when comparing a
C. maltosa dicarboxylic acid overproducing mutant
with a reference strain. Interestingly, P450alk5 is
indeed the isozyme with the strongest tendency to
x-hydroxylate fatty acids. By contrast, in vitro experi-
ments after heterologous expression demonstrated that
the highly active CYP52A3, although alkane prefer-
ring, not only converts hexadecane to its primary
product 1-hexadecanol, but also further oxidizes this
component to hexadecanal, hexadecanoic acid, 1,16-
hexadecandiol, 16-hydroxyhexadecanoic acid and even
1,16-hexadecanedioic acid, in this way bypassing the
two enzymes normally involved in x-oxidation [26].

The authors did not verify this phenomenon for other
fatty acid-oxidizing C. maltosa isozymes, but one can
postulate that CYP52A9 generates a similar oxidation
cascade. This assumption is supported by data from
another dicarboxylic acid-producing strain. Just like
C. maltosa, C. tropicalis mutant strains are used in
industrial dicarboxylic acid production, the strains are
among others blocked in b-oxidation by inactivation
of the POX genes, resulting in higher dicarboxylic acid
yields. Upon exposure to oleic acid, CYP52A13 and
CYP52A17 are strongly and consistently induced.
Again, enzymatic tests after heterologous expression
revealed the enzyme’s abilities to synthesize dicarbox-
ylic acids. CYP52A17 shows the greatest overoxidizing
capacities not only regarding substrate chain length,
but also concerning activity; the conversion of oleic
acid to its diacid occurs twice as quickly as the forma-
tion of x-hydroxy oleic acid [27]. Despite the in vitro
evidence, there is no clear answer to the question of
which role this P450 oxidation cascade plays in vivo.
One can assume that the prevalence of different
enzyme systems contributes to the yeast capacities to
grow efficiently on a broad chain-length range of alk-
anes and fatty acids (e.g. C7–C40 for C. maltosa).
Another advantage of the P450 bypass is the circum-
vention of H
2
O
2
formation by the fatty alcohol oxi-

dase. However, the overoxidation cascade requires
three molecules of NADPH, potentially creating
metabolic limitations. In classical x-oxidation, only one
NADPH molecule is used, whereas the alcohol dehy-
drogenase delivers one reducing equivalent (NADH).
Members of the CYP52 family are supposed to be
all linked to alkane and ⁄ or fatty acid hydroxylation,
yet these assumptions are made based on the amino
acid sequence and some enzymes might be involved in
unrelated biological processes. CaAlk8, for example, is
the only CYP52 member originating from C. albicans
(CYP52A21). Although it has been shown that the
enzyme terminally and subterminally hydroxylates lau-
ric, myristic and palmitic acid [28], and is involved in
alkane degradation, Panwar et al. [29] suggested that
CYP52A21 takes part in conferring multidrug resis-
tance to the opportunistic pathogen C. albicans. Dis-
ruption of CYP52A21 in the wild-type strain did not
lead to a drug-sensitive strain, probably attributed to
the presence of several other multidrug resistance
mechanisms. Nevertheless, the role of CYP52A21 in
multidrug resistance was demonstrated by overexpres-
sion in Saccharomyces cerevisiae and in a hypersensi-
tive C. albicans host, rendering the latter resistant to
fluconazole, itraconazole and 4-nitroquinoline oxide.
In addition, experiments with C. albicans microsomes
indicate that resistance is caused by CYP52A21-medi-
ated drug modification.
Besides the yeast species discussed above, Coryne-
bacterium sp. is also known as a producer of dicarbox-

ylic acids [30]. P450s are probably involved, but the
exact pathway remains unrevealed. By contrast, a spe-
cific P450 could be put forward as a candidate for
x-oxidation in the cyanobacteria Anabaena variabilis.
Although CYP110 is induced by alkanes such as hex-
adecane, CYP110 does not participate in alkane degra-
dation; findings which are supported by the fact that
alkanes are toxic for Anabaena variabilis. Based on the
sequence similarity and binding affinity experiments
with fatty acids, it was suggested that CYP110 is
related to fatty acid x-oxidation of saturated and
(poly)unsaturated fatty acids and subsequent forma-
tion of dicarboxylic acids which then undergo b-oxida-
tion [31]. The alkane inducibility of the cyp110 mRNA
was used to design a hexadecane biosensor system [32].
Biosynthesis of a- and b-hydroxylated
fatty acids
Not only are hydroxylated fatty acids intermediates in
alkane and fatty acid metabolization, they can also be
useful components as such. a-Hydroxylated long-chain
fatty acids, for example, are important constituents of
sphingolipids. These lipids are essential components of
The role of P450 in microbial fatty acid metabolism I. N. A. Van Bogaert et al.
210 FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS
mammalian cell membranes, but can also be found in
some bacterial and fungal cell membranes. Interest-
ingly, bacterial sphingolipid synthesis predominantly
occurs in anaerobic species. One such anaerobe is
Sphingomonas paucimobilis. Its sphingolipids are rich
in a-hydroxymyristic acid and the enzyme responsible

for the conversion of myristic acid has been identified
as a member of the P450 superfamily: P450
SPa
or
CYP152B1 [33]. However, the enzyme utilizes H
2
O
2
instead of O
2
and does not require reducing equiva-
lents, comparable with the peroxide shunt reaction for
common P450s [34]. Indeed, P450
SPa
lacks the critical
residues compulsory for the fulfilment of the typical
monooxygenation reaction; in the distal I helix the
Asp ⁄ Glu–Thr proton delivery system is substituted by
Arg–Pro and, albeit the heme-binding cysteine was
retrieved, the preceding consensus motive involved in
electron transfer was modified [35,36]. Furthermore, it
sounds quite logical that anaerobic organisms try to
circumvent the use of molecular oxygen. The enzyme
is highly specific towards fatty acids; no alkanes, fatty
alcohols or fatty aldehydes are hydroxylated. In addi-
tion to myristic acid, the enzyme shows activity to
slightly shorter or longer saturated fatty acids and ara-
chidonic acid [37]. Based on a database similarity
search with the P450
SPa

gene, Matsunaga’s group were
able to identify another fatty acid hydroxylase in Bacil-
lus subtillis. However, this P450
Bsb
or CYP152A1
enzyme is less regiospecific; myristic acid is converted
to a mixture of a- and b-hydroxymyristic acid, with
a slightly higher amount of the b-hydroxylated
product [38]. A few years ago, P450
CLA
(CYP152A2)
from the anaerobe Clostridium acetobutylicum was
characterized: like P450
Bsb
, this enzyme performs both
a- and b-hydroxylations of saturated and unsaturated
fatty acids, but in this particular case the a-position is
preferred [39].
Although there is a link between the occurrence of
fatty acid a-hydroxylation and sphingolipids, some
questions remain: what is the biological role of
P450
Bsb
in the non-sphingolipid-producing B. subtillis
and why is a mixture of a- and b-forms produced? It
was suggested that peroxide-utilizing P450s might be
involved in the oxygen-detoxification system of Closti-
dium acetobutylicum [39]. Although anaerobic, Clos-
tridium species can tolerate microoxic conditions
(< 5% O

2
). Besides the classical oxygen detoxification
systems, heme oxygenases, oxidases and lipid per-
oxidase scavenging enzymes are involved in the estab-
lishment of the anoxic microenvironment [40].
Furthermore, P450
SPa
turned out to be capable of
hydroxylating phytanic acid (3,7,11,15-tetramethyl
hexadecanoic acid), a degradation product of chlorophyll
[41]. This branched fatty acid cannot undergo
b-oxidation because of methylation at the b-position.
In humans, metabolization occurs by an initial
a-oxidation step, resulting in the removal of one car-
bon instead of two [42]. The subsequent pristanic acid
will be entirely degraded by b-oxidation. Oxidation of
phytanoyl-CoA is mediated by phytanoyl-CoA dioxy-
genase, an iron requiring non-heme oxidoreductase.
Homologue enzymes can be retrieved in a wide variety
of bacteria, questioning the role of P450 in bacterial
a-oxidation of phytanic acid.
Fatty acid hydroxylating P450s
involved in secondary metabolite
synthesis
Biosurfactants
Biosurfactants are surface-active compounds capable
of reducing interfacial tension between liquids thanks
to their amphiphilic properties. Amphiphilic molecules
consist of a hydrophilic and a hydrophobic moiety that
interacts with the phase boundary in heterogeneous

systems, allowing them to, for example, act as a deter-
gent, wetting agent or emulsifier of oil ⁄ water mixtures.
In general, common fatty acids or b-hydroxy fatty
acids originating from b-oxidation act as the hydro-
phobic part. However, in some particular cases, P450
hydroxylated fatty acids make up the hydrophobic tail.
One such example are the cellobiose lipids produced
by several yeast-like fungi (Fig. 2A). Ustilagic acids
from the plant-pathogen Ustilago maydis contain
15,16-dihydroxypalmitic acid or 2,15,16-trihydroxypal-
mitic acid. Teichmann and co-workers [43] elucidated
the biosynthetic gene cluster harbouring two P450
genes. Upon disruption of the first P450 gene, cyp1,no
cellobiose lipids could be detected. CYP1 catalyses the
conversion of palmitic acid to juniperic acid and this
terminal hydroxylation is essential for the covalent
binding of the hydroxylated fatty acid to the cellobiose
moiety. By contrast, Dcyp2 mutant strains retain their
capacities to secrete cellobiose lipids. Yet, these mole-
cules lack the typical hydroxylation at the subterminal
position. These findings prove that CYP1-dependent
x-hydroxylation does not depend on prior subterminal
hydroxylation and that both enzymes are highly selec-
tive for either the terminal or subterminal position.
Surprisingly, CYP1 and CYP2 share only 15% amino
acid identity and despite their activity towards the ter-
minus of fatty acids, they are not classified into the
CYP52 family but, based on their amino acid sequence,
are assigned CYP5025A1 and CYP5030A1, respectively.
No P450 activity is linked to a-hydroxylation; this

I. N. A. Van Bogaert et al. The role of P450 in microbial fatty acid metabolism
FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS 211
reaction is governed by AHD1, a non-heme diiron oxi-
do-reductase. cyp1 homologues were retrieved in two
other cellobiose lipids producing organisms, Pseu-
odozyma flocullosa and Pseuodozyma fusiformata [44].
One was not able to provide direct evidence for cyp1
contribution to the biosurfactant synthesis, but a posi-
tive correlation between cyp1 expression and flocculo-
sin synthesis was demonstrated.
Sophorolipids are another type of hydroxyfatty acid
containing biosurfactants. They consist of a x-orx-1-
hydroxylated fatty acid etherified via its hydroxylgroup
to a sophorose unit (Fig. 2B). The fatty acid can be
palmitic, palmitoleic, stearic, oleic or linoleic acid.
Typical sophorolipid-producing organisms are the
yeasts Candida bombicola and C. apicola. P450 involve-
ment in sophorolipid biosynthesis was suggested from
a simultaneous increase of cellular P450 content in
C. apicola. Two cyp52 genes were cloned from C. api-
cola (CYP52E1 and CYP52E2). Yet, Southern hybrid-
ization results indicated the existence of additional
cyp52 sequences, making it hard to draw conclusions
on the role of CYP52E1 and -E2 [45]. C. bombicola,
another sophorolipid-producing yeast closely related to
C. apicola, harbours at least eight cyp52 genes. One of
these, CYP52M1, was highly induced upon sophoroli-
pid production, suggesting its participation in the
sophorolipid biosynthesis pathway [46].
Polyketides

Polyketides are a structurally very diverse family of
secondary metabolites with different biological activi-
ties occurring in bacteria, plants and animals. They are
synthesized by polymerization of acetyl and propionyl
in a similar process to fatty acid synthesis and can
undergo extensive derivatization; in many cases, macr-
olidic structures are formed which are further modified
by, for example, several hydroxylation steps (Figs 3
and 4). These hydroxylation steps are very often medi-
ated by cytochrome P450 monooxygenases.
Discussing biosynthesis of all microbial polyketides
would take us too far from the scope of this review,
but we focus on two well-described polyketides of
which the backbone structure displays fatty acid simi-
larity. Fumonisin is a mycotoxin produced by several
Fusarium species, among others Fusarium verticilloides
and Fusarium proliferatum, both widespread plant-
pathogens infecting maize and other grains, rendering
this mycotoxin a common contaminant of corn.
Fumonisin is hepatotoxic and nephrotoxic, but its
acute toxicity is low. The long-term effect of low con-
centrations is less clear, but fumonisin is suggested to
be carcinogenic. The 17 fumonisin biosynthetic genes
are located in a gene cluster and three of them are
P450 enzymes. The FUM6 protein (CYP505B1) inter-
venes in one of the first steps in fumonisin synthesis by
hydroxylation of the polyketide-amino acid at C-14
and C-15 (Fig. 3). Fum6 deletion mutants of F. verticil-
loides are unable to produce fumonisin-like compounds
because the hydroxylgroups are required for the esteri-

fication of tricarballylic moieties downstream of the
biosynthesis pathway. FUM6 is a self-sufficient P450
containing a NADPH-dependent reductase domain
and belongs to the same family (CYP505) as the first
discovered self-sufficient eukaryotic P450: P450foxy
from F. oxysporum. The second cytochrome P450
monooxygenase, FUM2 (CYP65AH1), most likely
catalyses fumonisin C-10 hydroxylation, whereas the
third, FUM15 (CYP617F1), is suggested to be respon-
sible for the synthesis of low levels of a fumonisin iso-
form [47].
Well-studied examples of bacterial polyketides are
the antifungal components typically produced by soil
actinomycetes. Nystatins produced by Streptomyces
A
B
Fig. 2. Structure of (A) cellobiose lipids produced by Ustilago may-
dis; n = 2 or 4. (B) A common sophorolipid molecule in the acidic
form. R=H or COCH
3
.
The role of P450 in microbial fatty acid metabolism I. N. A. Van Bogaert et al.
212 FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS
noursei are commercialized as an antibiotic to treat
Candida sp. and Cryptococcus sp. infections (Fig. 4A).
The nystatin backbone is composed of a 38-membered
macrolactone ring which can be further modified by
cytochrome P450 enzymes; NysN (CYP105H1) oxi-
dizes the methyl group at C-16 and NysL (CYP161A1)
performs a hydroxylation at the C-10 position. DnysL

mutants produce 10-deoxynystatin, but despite the
absence of the hydroxyl group, the product retains its
antifungal activity [48]. Streptomyces nodosus synthe-
sizes amphotericin; in addition to its antifungal proper-
ties, this antibiotic is also active against human
immunodeficiency virus, Leishmania parasites and
prion diseases. Amphothericin has a 38-membered
macrolacton backbone similar to the one of nystatin
(Fig. 4B) and also this biosynthetic cluster contains
two P450 genes, amphL and amphN or CYP161A3 and
CYP105H4 respectively. AmphL mediates C-8 hydrox-
ylation, whereas AmphN oxidizes the C-16 methyl
group [49]. The biosynthetic gene clusters for a number
of polyketides have been elucidated (amphotericin,
nystatin, candicidin, pimaricin and rimocidin). All
these clusters contain a P450 gene homologue to amp-
hN and NysN associated to the C-16 oxidation. These
polyketide-specific P450 sequences can be used for
screening purposes. This strategy led to the identification
of a nystatin-like gene cluster in Pseudonocardia
autrophica containing the typical nppL and nppN genes
[50] and the isolation of putative polyketide producing
actinomycetes [51].
Oxylipins
Fatty acids have an established role as building
blocks of membranes and triacylglycerols, acting as a
structural component or energy reservoir. Besides
their classical roles, fatty acid derivates act as signal-
ling molecules with great physiological significance.
One such example are oxylipins, molecules originating

from oxidized unsaturated fatty acids. They are wide-
spread in aerobic organisms such as plants, animals
and fungi, but also occur in certain bacteria. Also
the well-described mammal prostaglandins and leuko-
trienes belong to the oxylipin family. Although oxyli-
pin synthesis in mammals and plants is well
documented, far less information can be found on
microbial oxylipins and reports on the cloning of
responsible genes are scarce. Oxylipin-forming
enzymes are structurally very diverse; in plants they
belong to an atypical cytochrome P450 subfamily,
whereas most other lipoxygenases are non-heme iron-
containing proteins.
Fig. 3. Part of the biosynthetic pathway of
fumonisin in Fusarium sp. R
1
= H or OH
(hydroxylation is rare and is supposed to be
governed by Fum15p), R
2
= H or OH. Steps
with P450 involvement are marked with an
arrow.
I. N. A. Van Bogaert et al. The role of P450 in microbial fatty acid metabolism
FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS 213
Oxylipins of fungal species are believed to be
involved in signalling, but more research is required
to assign specific functions. Aspergillus nidulans is a
model organism for the understanding of fungal
development because it has a defined sexual and asex-

ual development cycle. Oxylipins regulate the balance
between both cycles. Furthermore, they regulate sec-
ondary metabolism and are in this way important for
plant–host colonization and mycotoxin production
[52]. The involved oxylipins are called precocious sex-
ual inducers (psi-factors) and are derived from unsat-
urated C18 fatty acids. One such psi factor-producing
oxygenases or Ppo enzyme is PpoA, a bifunctional
protein with a fatty acid heme dioxygenase ⁄ peroxi-
dase domain in its N-terminal region and a P450
heme–thiolate domain in its C-terminal region. The
enzyme first oxidizes linoleic acid to (8R)-hydrox-
yperoxyoctadecadienoic acid and transfers this
product in the second reaction step to 5,8-dihydro-
xyoctadecadienoic acid by means of the P450 domain.
PpoA acts in vitro on unsaturated C16 and C20 fatty
acids as well, and was assigned as CYP6001A1 [53].
Several CYP6001 homologues can be retrieved among
fungi such as other Aspergillus sp., Neurospora sp.,
Fusarium sp. and Ustilago maydis, although their
function is not confirmed in these latter species
[54,55].
Another enzyme taking part in oxylipin biosynthesis
is PpoC (CYP1006C1). Like PpoA, two different
heme-containing regions are present, but in contrast to
PpoA the P450 heme–thiloate domain is degenerated;
the conserved cysteine residue known as the fifth heme
iron ligand is replaced by a glycine or phenylalanine in
A. nidulans or A. fumigatus PpoC, respectively. This
is reflected in the enzymes’ activity; whereas PpoA

further converts (8R)-hydroxyperoxyoctadecadienoic
acid to 5,8-dihydroxyoctadecadienoic acid, PpoC only
performs the first reaction step [56].
Oxylipins are also present in bacteria and might take
part in stress responses and host–pathogen interactions.
Most bacterial lipoxygenases are non-heme iron pro-
teins, but few plant CYP74-like proteins can be
retrieved, for example, in the rhizobacterium Methylo-
bacterium nodulans. This bifunctional protein possesses
an N-terminal peroxidase region and a C-terminal
CYP74-like P450 region. Because Methylobacteri-
um nodulans is a root-nodule-forming and nitrogen-fix-
ing symbiont of Crotalaria (plants belonging to the
Fabaceae family), it is plausible that this bacterial lipox-
ygenase originated from horizontal gene transfer [57].
Fatty acid-acting P450s with unclear
biological function
Self-sufficient P450
The majority of the P450 monooxygenases obtain the
necessary electrons for oxygen cleavage and substrate
hydroxylation via one or two redox partners. There
are several P450 redox systems which can be classified
into different groups according to the components
involved (reviewed in [58]). Most eukaryotic micro-
somal P450s – among them the previously discussed
CYP52 family – use the class II redox system. They
form a small electron transfer chain together with the
NADPH cytochrome P450 reductase. Both enzymes
O
O

H
3
C
H
3
C
H
2
N
H
3
C
H
3
C
H
3
C
H
2
N
H
3
C
CH
3
CH
3
O
A

B
OH
OH
OHOHOH
OH
OH
O
OH
O
HO
O
OH
OH
NysL
NysN
AmphL
O
OH
OH
O
OOH
OH
OH
OH
OH
O
O
HO
OH
O

OH
HO
AmphN
Fig. 4. Structure of (A) nystatin produced by
Streptomyces noursei, (B) amphotericin pro-
duced by Streptomyces nodosus. P450-
mediated hydroxylation or oxidations are
marked with an arrow.
The role of P450 in microbial fatty acid metabolism I. N. A. Van Bogaert et al.
214 FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS
are N-terminally anchored to the endoplasmatic reticu-
lum and derived structures, and the body of the pro-
tein is located in the cytoplasmatic space [59].
Cytochrome P450 reductase is a flavoprotein contain-
ing the flavin cofactors FAD and FMN. It transfers
the hydride ion of NADPH to the lower redox poten-
tial FAD. FAD then transfers single electrons to
FMN, which in turn reduces the cytochrome P450
monooxygenase heme centre as required to activate
molecular oxygen [60]. By contrast, most prokaryotic
P450s are cytosolic and communicate with two sepa-
rate redox partners: a NAD(P)H-binding and FAD-
containing reductase and a ferrodoxin or flavodoxin
that transfers the electrons from the reductase to the
P450 heme. These redox systems are referred to as
class I. Yet, in the 1980s, a catalytically self-sufficient
119 kDa protein was characterized and purified from
Bacillus megaterium: BM-3 or CYP102A1, now posi-
tioned in the class VIII redox family. This large pro-
tein is a gene fusion product between a P450 and a

cytochrome P450 reductase, rendering electron transfer
extremely efficient and resulting in a catalytic activity
of 4600 nmol fatty acid per nmol P450 per minute,
whereas most class II systems have catalytic activities
of two or even three orders of magnitude lower [61].
The self-sufficient and soluble properties simplify the
enzyme’s overexpression and purification, and turn it
into an ideal model for spectroscopic and structural
studies. The derived models for the heme–substrate
interactions indeed gave substantial input for the
understanding of mammalian P450 systems [62,63].
Despite the available data on the enzyme’s structure
and in vitro substrates, its proper biological role and
natural substrate remain to be revealed. BM-3
hydroxylates fatty acids with a chain length between
12 and 18 carbon atoms at the x-1, x-2 or x-3 posi-
tion and has highest affinity towards pentadecanoic
and palmitic acid (K
m
of 2 lm). Unsaturated fatty
acids are even better substrates and besides the typical
x-1, x-2 or x-3 hydroxylation, additional epoxidation
of double bonds can occur. Turnover rates of
> 15 000 have been reported for arachidonic acid
(C20:4) hydroxylation. Saturated fatty acid amides and
fatty alcohols are hydroxylated as well, yet at lower
efficiencies (reviewed in [64]).
BM-3 not only displays structural similarity with
mammalian P450s, it also shows an induction profile
very similar to mammalian CYP4A enzymes. These

P450s are fatty acid x-hydroxylases which are induced
by barbirturates and other peroxisomal proliferators.
Also in the 5¢-flanking region of the BM-3 gene a
so-called Barbie-box is retrieved; motives occurring in
all barbiturate-inducible genes. The regulatory system
includes the positive transcription factor BM3P1, the
autoregulated repressor Bm3R1 and several regulatory
sites. English et al. [65] found that the branched fatty
acid phytanic acid is not only an inducer of BM-3, but
also a substrate that is converted to x-1 hydroxyphy-
tanic acid. B. megaterium is a soil bacterium and the
authors state that many plant-derived unsaturated
fatty acids are extremely toxic to this bacteria; the
induction of BM-3 by phytanic or other fatty acids
may contribute to a metabolization or detoxification
system. However, it must be mentioned that phytanic
acid is not toxic to B. megaterium, although it is a
major vegetative breakdown product occurring in the
soil. Furthermore, branched chain fatty acids make up
80% of the fatty acid content of the Bacillus sp. mem-
branes and when the hydroxylation of these substrates
was studied in more detail, they were shown to be at
least as good substrates as their straight chain ana-
logues, having a higher regio- and stereospecific
hydroxylation pattern. Therefore, it is possible that
BM-3 takes part in the oxidative degradation of
branched chain fatty acids [66].
Its high catalytic activity, elucidated protein struc-
ture and ease of expression and use in in vivo experi-
ments have made BM-3 an attractive target for protein

engineering with possible biotechnological applications.
Various mutants are described as being able to act on
shorter fatty acids, polycyclic aromatic hydrocarbons
and even gaseous alkanes [67–69] or displaying a shift
in the hydroxylation pattern towards the terminal or
internal positions [70,71].
Meanwhile, the self-sufficient CYP102A family has
been extended with > 10 members, mainly originating
from soil bacteria. CYP102A2 and -A3 from B. subtil-
lis, CYP102A5 from B. cereus and CYP102A7 from
B. licheniformis have been characterized and in general
hydroxylate the same substrates as BM-3, sometimes
with even higher activities [72]. This group of proteins
even harbours a sequence of a noncultured soil bacte-
rium obtained by screening a metagenome database
[73]. Again, the biological roles in the different organ-
isms remain to be discovered, but it has been demon-
strated that CYP102A2 and -A3 are nonessential genes
and are not involved in the adaptive response concern-
ing fatty acid detoxification [74]. The same conclusion
can be drawn for CYP102B1. CYP102B1 is a cofactor
requiring arachidonic acid hydroxylating and epoxydiz-
ing P450 from Streptomyces coelicolor. No differences
concerning cell development or antibiotic production
were observed when comparing Dcyp102b1 strains
with wild-type strains, but the lipid profiles of both
strains were quite different, suggesting the involve-
ment of CYP102B1 in lipid biochemical pathways.
I. N. A. Van Bogaert et al. The role of P450 in microbial fatty acid metabolism
FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS 215

Unfortunately, it has not yet been possible to identify
CYP102B1-mediated products because of the complex
lipid fingerprint of Streptomyces coelicolor [75].
In the 1990s, a membrane-bound eukaryotic BM-3
counterpart was found in the fungus Fusarium oxyspo-
rum. The enzyme was first called P450foxy, but is
nowadays also referred to as CYP505A1. P450foxy
resembles BM-3 in its regiospecificity (x-1, x-2 or x-3)
and catalytic turnover, but differs slightly in fatty acid
preferences: saturated fatty acids are favoured over
unsaturated ones and the highest turnover is observed
for lauric acid [76]. Homologous sequences have been
found in other fungi such as Aspergillus, Neurospora
and Fusarium species, and the previously discussed
polyketide hydroxylases also belong to the CYP505
family.
Cofactor requiring P450s
Some P450s acting on fatty acids are hidden between
related proteins acting on totally different substrates.
CYP106A1 or BM-1 for B. megaterium, for example,
hydroxylates fatty acids, but information about the
CYP106 family is dominated by the steroid-hydroxyl-
ating capacities of CYP106A2 or BM-2. Another
example is CYP105D5 from Streptomyces coelicolor,
belonging to a large family merely constituted of
actinomycetes enzymes involved in various biological
processes such as vitamin D3 hydroxylation, degrada-
tion of xenobiotics and synthesis of polyketide antibi-
otics. Most enzymes can handle a broad range of
substrates, although CYP105D5 activity is restricted to

fatty acids only. x-1, x-2, x-3 and x-4 hydroxylation
products are formed, with the x-1 compound being
most prominent [77]. The same trend can be observed
in the bacterial CYP107 family: whereas most P450s
intervene in xenobiotic degradation, CYP107H1 from
B. subtilis is active towards myristic and palmitic acid.
The x-1, x-2 and x-3 hydroxylation of myristic acid is
believed to be required for the generation of pimelic
acid equivalents for biotin biosynthesis. Pimelic acid is
formed by P450-mediated in-chain cleavage via alcohol
and threo-diol intermediates [78].
Another so-called orphan P450 is the thermostable
CYP119A1 from archaebacteria Sulfolobus acidocalda-
rius. The strong hydrogen bonds and salt link net-
works, shortened loops and optimal aromatic stacking
safeguard the enzyme activity up to 85 °C. Despite
extensive structural analysis, the physiological function
of the enzyme is yet unclear. Initially, styrene was put
forward as a (poor) substrate, but recently a tight
binding affinity was demonstrated towards lauric acid,
resulting in the synthesis of mainly x-1 hydroxylated
lauric acid, suggesting a role in the lipid oxidative
metabolism. Interestingly, the percentage of the
x-hydroxylated product increases from 2.5 to 12%
when increasing the temperature from 24 to 80 °C [79].
Phylogenetic relationship of the
discussed enzymes
Figure 5 depicts the phylogenetic tree of the P450s
described in this minireview. Obviously, members of
the same family cluster together. The only exception is

CYP584A2, which is located in the CYP52 cluster
because of its high similarity with these molecules (30–
36% amino acid identity). P450s are classified in the
same family if their amino acid identity is at least
40%. In general, the criteria work quite well, but for
some proteins with high similarity instead of identity,
correct classification is not always that straightfor-
ward. Furthermore, the self-sufficient P450s, the
CYP102 and the CYP505 families, are found on a
common branch, indicating a mutual ancestor. One
would expect a close phylogenetic relationship between
the alkane-hydroxylating enzymes of bacteria
(CYP153) and yeasts (CYP52). Nevertheless, these
families are located quite distant from each other and
only share low amino acid identities (8–10%; Table
S1); convergent evolution has led to two types of
enzymes that in the end could fulfil identical biological
functions.
The proteins displaying the lowest similarity to the
other P450s are those involved in oxylipin biosynthesis
(2–6% amino acid identity). This can be explained in
part by their bifunctional structure.
Conclusion
In November 2009, 1015 bacterial and 2780 fungal
P450s were listed on the cytochrome P450 homepage
from Nelson [5], making up about one-third of the
total P450 database.
The majority of these enzymes catalyse hydroxyl-
ation reactions and although various endogenous and
xenobiotic compounds such as steroids and complex

aromatic structures can act as substrates, a significant
fraction of the P450s shows activity towards simple
molecules such as fatty acids and alkanes. The termi-
nal methyl groups of such molecules are quite inert
from a chemical point of view, yet P450s are able to
activate this thermodynamically disfavoured position
by hydroxylation. It is suggested that x-hydroxylation
is associated with a narrow substrate-access channel
governing restricted sterical activity. CYP52A21,
for example, possesses a small access channel and
The role of P450 in microbial fatty acid metabolism I. N. A. Van Bogaert et al.
216 FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS
predominantly hydroxylates the x-position [28]. BM-3,
however, is characterized by a large access channel and
is unable to perform hydroxylations at the terminal
ending. However, one must keep in mind that the
x ⁄ x-1 ratio can be influenced by other parameters
such as the substrate itself and more specifically its
absolute length, in vitro conditions and temperature as
has been demonstrated for CYP119.
This latter enzyme is referred to as an orphan
enzyme; its native substrates and biochemical func-
tion remain unclear. Yet, CYP119A1 is not the only
P450 without biochemical connotation; in the grow-
ing pile of data generated by genome-sequencing pro-
jects numerous genes are annotated as putative P450s
and although sequence similarity can give a clue
about the enzymes function, this still can be surpris-
ingly different as described for CYP105D1,
CYP106A1 and CYP107H1. Even for comprehen-

sively studied P450 systems such as BM-3, there are
merely assumptions regarding its physiological role.
Most P450 activities are studied by heterologous
expression. This approach is indeed convenient to
determine the enzyme’s potential activity, substrates
and specificities because there is a high expression
level and no background activity, but the approach
fails to provide information on the enzyme’s physio-
logical role and natural substrates. Therefore, more
direct experiments such as knockout studies and tran-
scription analysis are required.
A common characteristic of P450s are the many
gene duplication and conversion events. The fatty acid
and ⁄ or alkane-hydroxylating enzymes nicely illustrate
this feature within one organism, isoenzymes with dif-
ferent substrate specificities and expression levels are
Fig. 5. Phylogenetic tree of the P450s discussed in this minireview. The tree was constructed using the Protein Maximum Likelihood
(ProML) algorithm. The marker bar denotes the integer branch length. Ac ca, Acinetobacter calcoaceticus; Al bo, Alcanivorax borkumensis;
An va, Anabaena variabilis; As fu, Aspergillus fumigatus; As ni, Aspergillus nidulans; Ba ce, Bacillus cereus; Ba li, B. licheniformis; Ba me,
B. megaterium; Ba su, B. subtillus; Ca al, Candida albicans; Ca ap, C. apicola; Ca bo, C. bombicola; Ca ma, C. maltosa; Ca tr, C. tropicalis,
Cl ac, Clostridium acetobutylicum; Fu ox, Fusarium oxysporum; Fu ve, F. verticilloides; Ma or, Magnoporthe oryzae; My, Mycobacterium sp.;
No ar, Novosphingobium aromaticivorans; St co, Streptomyces coelicolor; St nod, S. nodosus; St nou, S. noursei; Su ac, Sulfolobus acido-
caldarius; Sp, Sphingomonas sp.; Sp pa, Sphingomonas puacimobilis; Us ma, Ustilago maydis; Ya li, Yarrowia lipolytica.
I. N. A. Van Bogaert et al. The role of P450 in microbial fatty acid metabolism
FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS 217
jointly able to degrade a whole range of substrates. In
the long-term, gene duplication and conversion pro-
gressing over different species result in a huge P450
diversity, in this way leading to various enzymes which
in the end could fulfil related or even identical reac-

tions (e.g. the fungal CYP52 and bacterial CYP153
families).
Throughout this review it has become clear that
there are numerous poorly studied or even orphan
P450s. However, there are quite a lot of biochemical
processes that require hydroxylation steps and that are
not associated with a specific enzyme. Hence, it is sug-
gested that several of these hydroxylating roles are ful-
filed by uncharacterized or yet to be discovered P450s.
Acknowledgements
The authors wish to thank the Flemish Agency or
Innovation by Science and Technology (IWT) for
financial support (grants IWT80050 and IWT090104)
and Mrs. Barabara Toch for carefully reviewing the
manuscript.
References
1 Omura T & Sato R (1964) Carbon monoxide-binding
pigment of liver microsomes. I. Evidence for its hemo-
protein nature. J Biol Chem 239, 2370–2378.
2 Kahn RA & Durst F (2000) Function and evolution of
plant cytochrome P450. In Evolution of Metabolic Path-
ways (Romeo JT, Ibrahim R, Varin L & DeLuca V,
eds), Pergamon, Oxford. pp. 151–189.
3 Nelson DR (1998) Cytochrome P450 nomenclature.
Methods Mol Biol 107, 15–24.
4 Pinot F & Beisson F (2010) Cytochrome P450-metabo-
lizing fatty acids in plants: characterization and physio-
logical roles. FEBS J 278, 195–205.
5 Nelson DR (2009) The cytochrome P450 homepage.
Hum Genome 4, 59–65.

6 Britton LN (1984) Microbial degradation of aliphatic
hydrocarbons. In Microbial Degradation of Organic
Compounds (Gibson DT, ed). Marcel Dekker, New
York. pp. 89–129.
7So
¨
hngen NL (1913) Benzin, petroleum, paraffino
¨
l und
paraffin als kohlenstoff- und energiequelle fu
¨
r mikro-
ben. Zentr Bacteriol Parasitenk 37, 595–609.
8 Baptist JN, Gholson RK & Coon MJ (1963) Hydro-
carbon oxidation by a bacterial enzyme system. 1.
Products of octane oxidation. Biochim Biophys Acta
69, 40–47.
9 Van Beilen JB, Li Z, Duetz WA, Smits THM & Witholt
B (2003) Diversity of alkane hydroxylase systems in the
environment. Oil Gas Sci Technol 58, 427–440.
10 Sanglard D, Chen C & Loper JC (1987) Isolation of the
alkane inducible cytochrome-P450 (P450alk) gene from
the yeast Candida tropicalis . Biochem Biophys Res
Commun 144, 251–257.
11 Craft DL, Madduri KM, Eshoo M & Wilson CR
(2003) Identification and characterization of the CYP52
family of Candida tropicalis ATCC 20336, important
for the conversion of fatty acids and alkanes to alpha,
omega-dicarboxylic acids. Appl Environ Microbiol 69,
5983–5991.

12 Ohkuma M, Zimmer T, Iida T, Schunck WH, Ohta A
& Takagi M (1998) Isozyme function of n-alkane-induc-
ible cytochromes P450 in Candida maltosa revealed by
sequential gene disruption. J Biol Chem 273, 3948–3953.
13 Kogure T, Takagi M & Ohta A (2005) n-Alkane and
clofibrate, a peroxisome proliferator, activate transcrip-
tion of ALK2 gene encoding cytochrome P450alk2
through distinct cis-acting promoter elements in Can-
dida maltosa. Biochem Biophys Res Commun 329, 78–86.
14 Lida T, Sumita T, Ohta A & Takagi M (2000) The
cytochrome P450ALK multigene family of an n-alkane-
assimilating yeast, Yarrowia lipolytica
: cloning and char-
acterization of genes coding for new CYP52 family
members. Yeast 16, 1077–1087.
15 Hirakawa K, Kobayashi S, Inoue T, Endoh-Yamagami
S, Fukuda R & Ohta A (2009) Yas3p, an Opi1 family
transcription factor, regulates cytochrome P450 expres-
sion in response to n-alkanes in Yarrowia lipolytica.
J Biol Chem 284, 7126–7137.
16 Cardini G & Jurtshuk P (1970) Enzymatic hydroxyl-
ation of normal-octane by Corynebacterium sp. strain
7e1c. J Biol Chem 245, 2789–2796.
17 Maier T, Forster HH, Asperger O & Hahn U (2001)
Molecular characterization of the 56-kDa CYP153 from
Acinetobacter sp EB104. Biochem Biophys Res Commun
286, 652–658.
18 Van Beilen JB, Funhoff EG, van Loon A, Just A,
Kaysser L, Bouza M, Holtackers R, Rothlisberger M,
Li Z & Witholt B (2006) Cytochrome P450 alkane

hydroxylases of the CYP153 family are common in
alkane-degrading eubacteria lacking integral membrane
alkane hydroxylases. Appl Environ Microbiol 72, 59–65.
19 Sabirova JS, Ferrer M, Regenhardt D, Timmis KN &
Golyshin PN (2006) Proteomic insights into metabolic
adaptations in Alcanivorax borkumensis induced by
alkane utilization. J Bacteriol 188, 3763–3773.
20 Oh Y, Donofrio N, Pan HQ, Coughlan S, Brown DE,
Meng SW, Mitchell T & Dean RA (2008) Transcrip-
tome analysis reveals new insight into appressorium
formation and function in the rice blast fungus
Magnaporthe oryzae. Genome Biol 9, R85.
21 Pedrini N, Crespo R & Juarez MP (2007) Biochemistry
of insect epicuticle degradation by entomopathogenic
fungi. Comp Biochem Physiol C 146, 124–137.
The role of P450 in microbial fatty acid metabolism I. N. A. Van Bogaert et al.
218 FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS
22 Freimoser FM, Hu G & St Leger RJ (2005) Variation
in gene expression patterns as the insect pathogen
Metarhizium anisopliae adapts to different host cuticles
or nutrient deprivation in vitro. Microbiology 151, 361–
371.
23 Scheller U, Zimmer T, Kargel E & Schunck W-H
(1996) Characterization of the n-alkane and fatty acid
hydroxylating cytochrome P450 forms 52A3 and 52A4.
Arch Biochem Biophys 328, 245–254.
24 Zimmer T, Scheller U, Takagi M & Schunck W-H
(1998) Mutual conversion of fatty-acid substrate speci-
ficity by a single amino-acid exchange at position 527 in
P-450Cm2 and P-450Alk3A. Eur J Biochem 256, 398–

403.
25 Kogure T, Horiuchi H, Matsuda H, Arie M, Takagi M
& Ohta A (2007) Enhanced induction of cytochromes
P450alk that oxidize methyl-ends of n-alkanes and fatty
acids in the long-chain dicarboxylic acid-hyperproduc-
ing mutant of Candida maltosa. FEMS Microbiol Lett
271, 106–111.
26 Scheller U, Zimmer T, Becher D, Schauer F & Schunck
WH (1998) Oxygenation cascade in conversion of n-alk-
anes to alpha,omega-dioic acids catalyzed by cyto-
chrome P450 52A3. J Biol Chem 273, 32528–32534.
27 Eschenfeldt WH, Zhang YY, Samaha H, Stols L,
Eirich LD, Wilson CR & Donnelly MI (2003) Transfor-
mation of fatty acids catalyzed by cytochrome P450
monooxygenase enzymes of Candida tropicalis. Appl
Environ Microbiol 69, 5992–5999.
28 Kim D, Cryle MJ, De Voss JJ & de Montellano PRO
(2007) Functional expression and characterization of
cytochrome P450 52A21 from Candida albicans. Arch
Biochem Biophys 464, 213–220.
29 Panwar SL, Krishnamurthy S, Gupta V, Alarco AM,
Raymond M, Sanglard D & Prasad R (2001) CaALK8,
an alkane assimilating cytochrome P450, confers multi-
drug resistance when expressed in a hypersensitive strain
of Candida albicans. Yeast 18, 1117–1129.
30 Broadway NM, Dickinson FM & Ratledge C (1993)
The enzymology of dicarboxylic-acid formation by
Corynebacterium sp strain 7e1c grown on n-alkanes.
J Gen Microbiol 139, 1337–1344.
31 Torres S, Fjetland CR & Lammers PJ (2005) Alkane-

induced expression, substrate binding profile, and
immunolocalization of a cytochrome P450 encoded on
the nifD excision element of Anabaena 7120. BMC
Microbiol 5, doi:10.1186/1471-2180-5-16.
32 Asai R, Nakamura C, Ikebukuro K, Karube I &
Miyake J (2003) A bioassay to detect contaminant-
induced messenger RNA using a transcriptomic
approach: detection of RT-PCR-amplified single-
stranded DNA based on the SPR sensor in
cyanobacteria. Anal Lett 36, 1475–1491.
33 Matsunaga I, Yamada M, Kusunose E, Miki T &
Ichihara K (1998) Further characterization of
hydrogen peroxide dependent fatty acid alpha-hydroxy-
lase from Sphingomonas paucimobilis. J Biochem 124,
105–110.
34 Sono M, Roach MP, Coulter ED & Dawson JH (1996)
Heme-containing oxygenases.
Chem Rev 96, 2841–2887.
35 Matsunaga I, Yokotani N, Gotoh O, Kusunose E,
Yamada M & Ichihara K (1997) Molecular cloning
and expression of fatty acid alpha-hydroxylase from
Sphingomonas paucimobilis. J Biol Chem 272, 23592–
23596.
36 Matsunaga I & Shir Y (2004) Peroxide-utilizing biocata-
lysts: structural and functional diversity of heme-con-
taining enzymes. Curr Opin Chem Biol 8, 127–132.
37 Matsunaga I, Sumimoto T, Ueda A, Kusunose E &
Ichihara K (2000) Fatty acid-specific, regiospecific, and
stereospecific hydroxylation by cytochrome P450
(CYP152B1) from Sphingomonas paucimobilis : substrate

structure required for alpha-hydroxylation. Lipids 35,
365–371.
38 Matsunaga I, Ueda A, Fujiwara N, Sumimoto T &
Ichihara K (1999) Characterization of the ybdT gene
product of Bacillus subtilis: novel fatty acid beta-
hydroxylating cytochrome P450. Lipids 34, 841–846.
39 Girhard M, Schuster S, Dietrich M, Durre P & Urlach-
er VB (2007) Cytochrome P450 monooxygenase from
Clostridium acetobutylicum: a new alpha-fatty acid
hydroxylase. Biochem Biophys Res Commun 362, 114–
119.
40 Kawasaki S, Watamura Y, Ono M, Watanabe T,
Takeda K & Niimura Y (2005) Adaptive responses to
oxygen stress in obligatory anaerobes Clostridium acet-
obutylicum and Clostridium aminovalericum . Appl
Environ Microbiol 71, 8442–8450.
41 Matsunaga I, Sumimoto T, Kusunose E & Ichihara K
(1998) Phytanic acid alpha-hydroxylation by bacterial
cytochrome P450. Lipids 33, 1213–1216.
42 Wanders RJA, Komen J & Kemp S (2010) Fatty acid
omega-oxidation as a rescue pathway for fatty acid oxi-
dation disorders in humans. FEBS J 278, 182–194.
43 Teichmann B, Linne U, Hewald S, Marahiel MA &
Bolker M (2007) A biosynthetic gene cluster for a
secreted cellobiose lipid with antifungal activity from
Ustilago maydis. Mol Microbiol 66, 525–533.
44 Marchand G, Remus-Borel W, Chain F, Hammami W,
Belzile F & Belanger RR (2009) Identification of genes
potentially involved in the biocontrol activity of Pseud-
ozyma flocculosa. Phytopathology 99, 1142–1149.

45 Lottermoser K, Schunck WH & Asperger O (1996)
Cytochromes P450 of the sophorose lipid-producing
yeast Candida apicola: heterogeneity and polymerase
chain reaction-mediated cloning of two genes. Yeast 12,
565–575.
46 Van Bogaert INA, Demey M, Develter D, Soetaert W
& Vandamme EJ (2009) Importance of the cyto-
chrome P450 monooxygenase CYP52 family for the
I. N. A. Van Bogaert et al. The role of P450 in microbial fatty acid metabolism
FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS 219
sophorolipid-producing yeast Candida bombicola.
FEMS Yeast Res 9, 87–94.
47 Alexander NJ, Proctor RH & McCormick SP (2009)
Genes, gene clusters, and biosynthesis of trichothecenes
and fumonisins in Fusarium. Toxin Rev 28, 198–215.
48 Volokhan O, Sletta H, Ellingsen TE & Zotchev SB
(2006) Characterization of the P450 monooxygenase
NysL, responsible for C-10 hydroxylation during bio-
synthesis of the polyene macrolide antibiotic nystatin in
Streptomyces noursei. Appl Environ Microbiol 72, 2514–
2519.
49 Carmody M, Murphy B, Byrne B, Power P, Rai D,
Rawlings B & Caffrey P (2005) Biosynthesis of ampho-
tericin derivatives lacking exocyclic carboxyl groups.
J Biol Chem 280, 34420–34426.
50 Kim BG, Lee MJ, Seo J, Hwang YB, Lee MY, Han K,
Sherman DH & Kim ES (2009) Identification of func-
tionally clustered nystatin-like biosynthetic genes in a
rare actinomycetes, Pseudonocardia autotrophica . J Ind
Microbiol Biotechnol 36, 1425–1434.

51 Hwang YB, Lee MY, Park HJ, Han K & Kim ES
(2007) Isolation of putative polyene-producing actino-
mycetes strains via PCR-based genome screening for
polyene-specific hydroxylase genes. Process Biochem 42,
102–107.
52 Tsitsigiannis DI & Keller NP (2006) Oxylipins act as
determinants of natural product biosynthesis and seed
colonization in Aspergillus nidulans. Mol Microbiol 59,
882–892.
53 Brodhun F, Gobel C, Hornung E & Feussner I (2009)
Identification of PpoA from Aspergillus nidulans as a
fusion protein of a fatty acid heme dioxygenase ⁄ peroxi-
dase and a cytochrome P450. J Biol Chem 284, 11792–
11805.
54 Jerneren F, Garscha U, Hoffmann I, Hamberg M &
Oliw EH (2010) Reaction mechanism of 5,8-linoleate
diol synthase, 10R-dioxygenase, and 8,11-hydroperoxide
isomerase of Aspergillus clavatus. Biochim Biophys Acta
1801, 503–507.
55 Jerneren F, Hoffmann I & Oliw EH (2010) Linoleate
9R-dioxygenase and allene oxide synthase activities of
Aspergillus terreus. Arch Biochem Biophys 495, 67–73.
56 Brodhun F, Schneider S, Gobel C, Hornung E & Feussner
I (2010) PpoC from Aspergillus nidulans is a fusion pro-
tein with only one active haem. Biochem J 425, 553–565.
57 Lee DS, Nioche P, Hamberg M & Raman CS (2008)
Structural insights into the evolutionary paths of oxyli-
pin biosynthetic enzymes. Nature 455, 363–368.
58 Hannemann F, Bichet A, Ewen KM & Bernhardt R
(2007) Cytochrome P450 systems – biological variations

of electron transport chains. Biochim Biophys Acta
1770, 330–344.
59 Edwards RJ, Murray BP, Singleton AM & Boobis AR
(1991) Orientation of cytochromes-P450 in the endopla-
smicreticulum. Biochemistry 30
, 71–76.
60 Nebert DW & Gonzalez FJ (1987) P450 genes: struc-
ture, evolution, and regulation. Annu Rev Biochem 56,
945–993.
61 Narhi LO & Fulco AJ (1986) Characterization of a cat-
alytically self-sufficient 119,000-dalton cytochrome P450
monooxygenase induced by barbiturates in Bacil-
lus megaterium. J Biol Chem 261, 7160–7169.
62 Li HY & Poulos TL (1997) The structure of the cyto-
chrome p450BM-3 haem domain complexed with the
fatty acid substrate, palmitoleic acid. Nat Struct Biol 4,
140–146.
63 Li HY & Poulos TL (1995) Modeling protein substrate
interactions in the heme domain of cytochrome
P450(Bm-3). Acta Crystallogr D Biol Crystallogr 51,
21–32.
64 Hilker BL, Fukushige H, Hou C & Hildebrand D
(2008) Comparison of Bacillus monooxygenase genes for
unique fatty acid production. Prog Lipid Res 47, 1–14.
65 English N, Palmer CNA, Alworth WL, Kang L,
Hughes V & Wolf CR (1997) Fatty acid signals in
Bacillus megaterium are attenuated by cytochrome
P-450-mediated hydroxylation. Biochem J 327, 363–368.
66 Cryle MJ, Espinoza RD, Smith SJ, Matovic NJ & De
Voss JJ (2006) Are branched chain fatty acids the natu-

ral substrates for P450(BM3)? Chem Commun 22, 2353–
2355.
67 Lentz O, Qing-Shang LI, Schwaneberg U, Lutz-Wahl S,
Fischer P & Schmid RD (2001) Modification of the
fatty acid specificity of cytochrome P450BM-3 from
Bacillus megaterium by directed evolution: a validated
assay. J Mol Catal B 15, 123–133.
68 Li QS, Ogawa J, Schmid RD & Shimizu S (2001) Engi-
neering cytochrome P450BM-3 for oxidation of polycy-
clic aromatic hydrocarbons. Appl Environ Microbiol 67,
5735–5739.
69 Lewis JC & Arnold FH (2009) Catalysts on demand:
selective oxidations by laboratory-evolved cytochrome
P450 BM3. Chimia 63, 309–312.
70 Meinhold P, Peters MW, Hartwick A, Hernandez AR
& Arnold FH (2006) Engineering cytochrome P450BM3
for terminal alkane hydroxylation. Adv Synth Catal
348, 763–772.
71 Dietrich M, Do TA, Schmid RD, Pleiss J &
Urlacher VB (2009) Altering the regioselectivity of the
subterminal fatty acid hydroxylase P450 BM-3
towards gamma- and delta-positions. J Biotechnol 139,
115–117.
72 Dietrich M, Eiben S, Asta C, Do TA, Pleiss J &
Urlacher VB (2008) Cloning, expression and charac-
terisation of CYP102A7, a self-sufficient P450
monooxygenase from Bacillus licheniformis. Appl Micro-
biol Biotechnol 79, 931–940.
73 Kim BS, Kim SY, Park J, Park W, Hwang KY, Yoon
YJ, Oh WK, Kim BY & Ahn JS (2007) Sequence-based

screening for self-sufficient P450 monooxygenase from
The role of P450 in microbial fatty acid metabolism I. N. A. Van Bogaert et al.
220 FEBS Journal 278 (2011) 206–221 ª 2010 The Authors Journal compilation ª 2010 FEBS
a metagenome library. J Appl Microbiol 102, 1392–
1400.
74 Gustafsson MCU, Palmer CNA, Wolf CR & von
Wachenfeldt C (2001) Fatty-acid-displaced transcrip-
tional repressor, a conserved regulator of cytochrome
P450 102 transcription in Bacillus species. Arch
Microbiol 176, 459–464.
75 Lamb DC, Lei L, Zhao B, Yuan H, Jackson CJ,
Warrilow AGS, Skaug T, Dyson PJ, Dawson ES, Kelly
SL et al. (2010) Streptomyces coelicolor A3(2) CYP102
protein, a novel fatty acid hydroxylase encoded as a
heme domain without an N-terminal redox partner.
Appl Environ Microbiol 76 , 1975–1980.
76 Nakayama N, Takemae A & Shoun H (1996) Cyto-
chrome P450foxy, a catalytically self-sufficient fatty acid
hydroxylase of the fungus Fusarium oxysporum. J Bio-
chem 119, 435–440.
77 Chun YJ, Shimada T, Sanchez-Ponce R, Martin MV,
Lei L, Zhao B, Kelly SL, Waterman MR, Lamb DC &
Guengerich FP (2007) Electron transport pathway for a
Streptomyces cytochrome P450 – cytochrome P450
105D5-catalyzed fatty acid hydroxylation in
Streptomyces coelicolor A3(2). J Biol Chem 282, 17486–
17500.
78 Cryle MJ & De Voss JJ (2004) Carbon-carbon bond
cleavage by cytochrome P450(BioI) (CYP107H1). Chem
Commun 1, 86–87.

79 Lim Y-R, Eun C-Y, Park HG, Songhee H, Han J-S,
Cho KS, Chun Y-J & Kim D (2010) Regioselective
oxidation of lauric acid by CYP119, an orphan
cytochrome P450 from Sulfolobus acidocaldarius.
J Microbiol Biotechnol 20, 574–578.
Supporting information
The following supplementary material is available:
Table S1. Amino acid sequence identities (%) between
the P450s discussed in this review.
This supplementary material can be found in the
online version of this article.
Please note: As a service to our authors and readers,
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I. N. A. Van Bogaert et al. The role of P450 in microbial fatty acid metabolism
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