REVIEW ARTICLE
a-Methylacyl-CoA racemase – an ‘obscure’ metabolic
enzyme takes centre stage
Matthew D. Lloyd
1
, Daniel J. Darley
1
, Anthony S. Wierzbicki
2
and Michael D. Threadgill
1
1 Department of Pharmacy & Pharmacology, Medicinal Chemistry, University of Bath, UK
2 Department of Chemical Pathology, St Thomas’ Hospital, London, UK
Introduction
Branched-chain fatty acids and related compounds are
important components of the human diet and are also
used as drug molecules. Owing to the presence of
methyl groups on the carbon chain, the majority can-
not be immediately metabolized within mitochondria,
and instead undergo initial metabolism in peroxisomes
[1–4]. A consequence of the presence of methyl groups
on the carbon chain is that many of these fatty acids
contain chiral centres. Methyl groups can be located
on both the two and three carbon positions, and this
has consequences for metabolism. The oxidation of
these fats is stereoselective [1], and this has conse-
quences for the regulation of metabolism.
Branched-chain fatty acids can arise from several dif-
ferent sources. Humans endogenously synthesize bile
acids, which are oxidized cholesterol derivatives. These
acids possess the methyl group on carbon 2 (relative to

a-oxidation, b-oxidation, and x-oxidation. Dietary branched-chain lipids
(especially phytanic acid) have attracted much attention in recent years,
due to their link with prostate, breast, colon and other cancers as well as
their role in neurological disease. A central role in all the metabolic path-
ways is played by a-methylacyl-CoA racemase (AMACR), which regulates
metabolism of these lipids and drugs. AMACR catalyses the chiral inver-
sion of a diverse number of 2-methyl acids (as their CoA esters), and regu-
lates the entry of branched-chain lipids into the peroxisomal and
mitochondrial b-oxidation pathways. This review brings together advances
in the different disciplines, and considers new research in both the meta-
bolism of branched-chain lipids and their role in cancer, with particular
emphasis on the crucial role played by AMACR. These recent advances
enable new preventative and treatment strategies for cancer.
Abbreviations
ACOX, acyl-CoA oxidase; AMACR, a-methylacyl-CoA racemase; CYP, cytochome P450; FALDH, fatty aldehyde dehydrogenase; FAR and
MCR, a-methylacyl-CoA racemase from Mycobacterium tuberculosis; PhyH, phytanoyl-CoA 2-hydroxylase; PPAR, peroxisome proliferation-
activated receptor.
FEBS Journal 275 (2008) 1089–1102 ª 2008 The Authors Journal compilation ª 2008 FEBS 1089
Phytanic acid is its 3-methyl dietary precursor, with
stereochemistry identical to that of pristanic acid.
Phytanic acid is originally derived from the isoprenoid
side-chain of chlorophyll A, phytol, although it is gen-
erally believed that phytol cannot be cleaved from
chlorophyll in plant-derived foods and that phytanic
acid comes directly from animal products. Foods that
are particularly rich in phytanic acid include beef,
other meats and dairy products. A typical daily intake
of phytanic acid in a Western diet has been estimated
to be 50–100 mg [2]. Finally, anti-inflammatory drugs
such as ibuprofen are 2-methyl acids [1]. These drugs

enzyme is discussed.
Branched-chain fatty acid metabolism
The a-oxidation pathway for phytanic acid (and pre-
sumably other 3-methyl acids) was finally elucidated
about 10 years ago [1–4]. Significant further progress
has been made, including considerable advances in
understanding the conversion of free phytol to phyte-
noyl-CoA, which can be converted to pristanic acid.
Further progress has also been made in understanding
x-oxidation, a secondary degradation pathway for
phytanic acid (Scheme 1). The presence of 3-methyl
groups in phytanic acid prevents b-oxidation, as a qua-
ternary alcohol is produced from this substrate. Hence,
phytanic acid undergoes preliminary a-oxidation, in
which chain shortening from the carboxyl group
occurs. This pathway produces pristanic acid, which
has a 2-methyl group, and hence b-oxidation is not
blocked.
The a-oxidation pathway consists of four steps [1],
the first being conversion of phytanic acid to its
CoA ester and peroxisomal import (Scheme 2). This
is followed by hydroxylation by a nonhaem iron(II)
and a 2-oxoglutarate-dependent oxygenase, phyta-
noyl-CoA 2-hydroxylase (PhyH). Adult Refsum’s dis-
ease is a result of inactivating mutations in this
enzyme [13,14] or of defects in the system responsi-
ble for importing this protein into peroxisomes [15].
The X-ray crystal structure of PhyH has recently
been solved [13], and this demonstrates that the
majority of clinical mutations cluster around the

fatty aldehyde dehydrogenase (FALDH) [25], the
enzyme that is deficient in Sjo
¨
gren–Larsson syndrome
[1,28]. Studies on recombinant FALDH showed that it
was also able to oxidize alcohols to aldehydes [29].
a-Methylacyl-CoA racemase and cancer M. D. Lloyd et al.
1090 FEBS Journal 275 (2008) 1089–1102 ª 2008 The Authors Journal compilation ª 2008 FEBS
Although conversion of phytol was not demonstrated,
the use of a bifunctional oxidoreductase would prevent
the release of the highly reactive allylic aldehyde, phyt-
enal. Phytenic acid is converted to its CoA ester and
reduced to phytanic acid by an NADPH-dependent
oxidoreductase [26,30]. It is not clear how much plant-
derived phytol is converted into phytanic acid in
humans, as humans are not supposed to be able to
cleave this side-chain from chlorophyll, although some
contribution from gut bacteria cannot be excluded
Scheme 1. Metabolism of branched-chain fatty acids and related compounds. *Peroxisomes contain more than one fatty acyl-CoA synthe-
tase, and it is not clear which specific enzyme is responsible for the phytenic acid-to-phytenoyl-CoA conversion. Enzymes, cosubstrates and
cofactors [1,2]: 1, phytanoyl-CoA 2-hydroxylase, iron(II), 2-oxoglutarate, O
2
; 2, 2-hydroxyphytanoyl-CoA lyase (also known as 2-hydroxyacyl-
CoA lyase), Mg
2+
-thiamine diphosphate; 3, FALDH-V, CYPs; 4, very-long-chain fatty acyl-CoA synthetase, Mg
2+
-ATP, CoA-SH; 5, 6, unidenti-
fied oxidoreductases or CYP enzyme – Reactions will go via aldehydes and acid intermediates; 7, branched-chain acyl-CoA oxidase, FAD;
8, 9, D-bifunctional protein, NAD

1092 FEBS Journal 275 (2008) 1089–1102 ª 2008 The Authors Journal compilation ª 2008 FEBS
FALDH [37] has been identified (FALDH-V), and
very recently it has been shown to localize in peroxi-
somal membranes [38]. Two further splice variants
(FALDH-V2 and FALDH-V3) were also identified
[38], although these appear not to be localized in
peroxisomes. The authors propose that FALDH-V
catalyses the conversion of pristanal to pristanic
acid, and this is supported by the observation that
overexpression of FALDH-V but not FALDH-N
protects cells against phytanic acid-induced damage.
Production of all four protein splice variants of
FALDH are induced by peroxisome proliferation-
activated receptor (PPAR)a agonists, and increased
expression of FALDH-N and FALDH-V protects
against lipid peroxidation. The low level of residual
pristanal dehydrogenase activity in Sjogren–Larsson
syndrome fibroblasts was attributed to incomplete
loss of activity in FALDH mutants [34,38]. How-
ever, PPARa agonists were also shown to induce
several other genes in addition to aldh3a2 (the
gene encoding for the FALDH splice variants),
including several cytochome P450 (CYP) enzymes
[39]. It could be that one or more CYP enzymes
play a secondary role in the pristanal-to-pristanic
acid conversion.
Although a-oxidation is the primary metabolic
pathway for phytanic acid, some metabolism can also
occur by x-oxidation [40–44]. Clinically, x-oxidation
is important in patients deficient in a-oxidation, such

entry into the b-oxidation pathway. The b-oxidation
pathway chain shortens the fatty acids by two car-
bons during each cycle. In the case of pristanic acid,
b-oxidized fragments, such as acetyl-CoA and pro-
pionoyl-CoA, and chain-shortened intermediates are
exported into mitochondria for final metabolism via
the acyl-carnitine shuttle [1]. As these chain-shortened
intermediates also contain chiral methyl groups with
the (R)-configuration, AMACR is also required
within mitochondria (see below) for b-oxidation to
occur. It is not known whether chain-shortened bile
acids are similarly exported to the mitochondria.
Patients deficient in AMACR exhibit neurological
symptoms [49] with some similarities to adult Ref-
sum’s disease [2] but with later onset and a more
peripheral than central neurological phenotype. They
exhibit the expected biochemical profile, with accumu-
lation of bile acids and dietary (2R)-branched acids
[47,50]. A ‘knockout’ mouse model is also available,
and this shows a similar metabolic profile, with
upregulation of expression for several genes, including
those encoding CYP enzymes that may be involved in
x-oxidation [51].
Ibuprofen is a 2-methyl acid, and is generally given
as a racemic mixture of (2R)- and (2S)-enantiomers.
Activation as the CoA ester and chiral inversion [52–
56] have been implicated in both pharmacological
activity and toxic side-effects. The enzyme responsible
for this is ‘ibuprofenoyl-CoA epimerase’ [52], which,
upon cloning, proved to be identical to AMACR

also occur in breast [69], colon [63], renal [70,71] and
other cancers [61,72], although there is considerable
heterogeneity in the degree of overproduction (for
example, Jiang et al. [61] reported that only 27% of
gastric adenocarcinomas overproduce AMACR).
Since then, a large body of evidence has linked die-
tary branched-chain lipid intake (especially phytanic
acid), AMACR overproduction [11,12], and cancer.
Xu et al. [73] reported that dietary phytanic acid
intake and levels in the blood directly correlate with
prostate cancer risk, whereas Mobley et al. [74] showed
that dietary branched-chain fatty acids increased pro-
duction of AMACR in prostate cancer cells, with cata-
lytic activity also being increased [66,75]. AMACR
overproduction appears to be mediated by a nonclassic
C ⁄ EBP-binding motif in the promoter region [76].
Other enzymes involved in the peroxisomal b-oxidation
of branched-chain fatty acids are also overproduced
[e.g. acyl-CoA oxidase (ACOX)2, also known as
D-bifunctional protein] [77], and that the relative levels
of production of enzyme subtypes can also change (for
example, ACOX3 expression is increased [77]), presum-
ably due to increased levels of the substrates. Certain
AMACR polymorphisms leading to single amino acid
substitutions are also associated with increased pros-
tate [78,79] and colon [80] cancer risk. In the case of
prostate cancer, the strongest correlation is for the
M9V polymorphism [79], with the minor allele over-
represented in unaffected men. Inactivating mutations
in AMACR give rise to an adult-onset neurological

AMACR is colocalized in both peroxisomes and mito-
chondria in both humans [84,85] and rats [86]. The
enzyme localized in both organelles is derived from a
single transcript [84,86]. The enzyme possesses an
N-terminal mitochondrial targeting signal and a C-ter-
minal peroxisomal targeting sequence-1 variant, the
final four amino acids, KASL [49]. These studies were
performed before the existence of the minor splice vari-
ants [81–83] was known, and therefore refer to AMA-
CR 1A, the major form of the enzyme in ‘normal’ cells.
Examination of the minor splice variant sequences
[81–83] reveals a common N-terminus containing the
mitochondrial targeting signal. The C-terminal peroxi-
somal targeting sequence-1 signal is missing in all splice
variants, implying that they will be exclusively mito-
chondrial, although this has yet to be verified.
The racemase-catalysed reaction requires no cofac-
tors or cosubstrates [1,52,87,88], and involves stereo-
specific removal and addition of a proton. The
formation of the CoA ester facilitates this process by
increasing the basicity of the 2-proton (a-proton) by
reducing the pKa from  34 to 21 [89]. Although this
simple reaction could be theoretically performed with-
out an enzyme, in practice the rates would be prohibi-
tively slow and the alkali pH values would bring about
hydrolysis of the CoA ester in preference to racemiza-
tion. The reaction is reversible, and for the substrate
containing a single chiral centre, the in vitro equilib-
rium constant has been measured as  1.5 (ibuprofe-
noyl-CoA with the rat enzyme) [52] in favour of the

mologues of AMACR, MCR [90] and FAR [95], have
been reported, which possessed the same overall fold.
The structure of MCR was reported in conjunction
with a site-directed mutagenesis study that identified
some of the catalytic residues [90]. The study also
looked at the effects of the equivalent mutations (I56P
and M111P [90]) to those giving rise to AMACR defi-
ciency in humans (S52P and L107P [49]). As expected,
the M111P mutation led to a significant reduction in
catalytic activity (to  1.6% of wild-type activity).
Unexpectedly, the I56P mutant had 76% activity as
compared to the wild-type enzyme, when almost com-
plete abolition of activity was expected. This anoma-
lous result could reflect differences in the structures
between the human and mycobacterial enzymes, or it
may be that the S52P human mutant is significantly
active and that racemase deficiency results from some
other mechanism, e.g. reduced transcription or transla-
tion, or mRNA or protein instability.
The structural and mutagenic data enable some
mechanistic details about the human AMACR-cataly-
sed reaction to be predicted. However, the primary
sequence identity of human AMACR 1A with these
other enzymes is quite low, e.g.  30% with MCR
[90] and  25% with YfdW [93], so any predictions
should be treated with caution. It is noteworthy that
the four important residues identified in MCR [90]
are in regions of relatively high conservation. The
equivalent residue to MCR Arg91 in AMACR 1A is
Lys87; the MCR mutant displays an increased K

now possible, but surprisingly, only one paper has thus
far appeared in this area [96]. The paper reported com-
petitive inhibitors with K
i
values of 0.9–20 lm when
tested against enzyme purified from rat liver, with the
Fig. 1. Active site residues of human
AMACR 1A identified from the Mycobacte-
rium tuberculosis enzyme, MCR [90,91].
The catalytic residue is in green; the oxyan-
ionic intermediate stabilization and proton
acceptor residues are in red; the CoA-bind-
ing residue is in blue. The protonation state
is for the (2S)-substrate to (2R)-substrate
conversion.
M. D. Lloyd et al. a-Methylacyl-CoA racemase and cancer
FEBS Journal 275 (2008) 1089–1102 ª 2008 The Authors Journal compilation ª 2008 FEBS 1095
most active compounds inhibiting growth of cancer
cell lines. The potency of inhibition in cells is directly
correlated with levels of AMACR protein in the cells.
These results are encouraging, but a greater under-
standing of the roles of all the human splice variants is
required in order for this approach to be fully
exploited.
Unanswered questions and future work
Dietary branched-chain fatty acids represent a signifi-
cant risk factor for prostate cancer, and the metabolic
pathways responsible for degradation of these fatty
acids are upregulated in cancers. AMACR acts as a
‘gate-keeper’ for b-oxidation. The identification of

late expression of fat-metabolizing enzymes and brown
fat tissue [118]. PPAR-a and PPAR-c receptor agonists
protect against cancer, whereas PPAR-d agonists pro-
mote cancer in some animal models [119]. Phytanic
acid [109] and pristanic acid [113] are agonists of
PPAR-a, but their effects on PPAR-d are unknown.
Support for this model was recently provided by the
observation that increased expression of FALDH-V
protects cells against phytanic acid-induced damage in
rodents [38]. This splice variant of FALDH performs
the pristanal-to-pristanic acid conversion in the a-oxi-
dation pathway, thus facilitating detoxification of phy-
tanic acid and its phytol precursor. However, this area
is complicated by the considerable differences between
rodent and human PPAR pathways as well as between
tissues. For example, phytol [111,114] may be a
PPAR-a ligand in human cell lines, whereas phytanic
acid is a PPAR-a ligand in mice [103] but its effects in
humans are controversial. It could be that branched-
chain fatty acids or their metabolites are agonists for
PPAR-d or antagonists for PPAR-c, and this is the
molecular basis for cancer formation, at least in some
model systems. These theories merit further investiga-
tion and are attractive in the sense that they explain
why particular cancers appear to be promoted, as
prostate and breast tissues are particularly active in fat
metabolism.
Selective inhibition of specific splice variants could
lead to new anticancer therapies. The use of AMACR
inhibitors is particularly attractive, as protein expres-

a-Methylacyl-CoA racemase and cancer M. D. Lloyd et al.
1096 FEBS Journal 275 (2008) 1089–1102 ª 2008 The Authors Journal compilation ª 2008 FEBS
intakes are associated with lower rates of prostate can-
cer [73,121]. Diets with low phytanic acid have been
available for many years for the treatment of adult
Refsum’s disease [122–124]. A recent study was per-
formed as part of an EU project on adult Refsum’s
disease, and the website contains a phytanic acid calcu-
lator for various foodstuffs in resources for both
patients and clinicians ().
A reduced phytanic acid diet could be of benefit to
men at risk of developing prostate cancer and be of
use for prevention of other major cancers, such as
those of breast and colon. Plasma phytanic acid levels
are strongly associated with dairy fat intake [32], with
the levels found in meat eaters, lacto-ovo-vegetarians
and vegans being 5.77, 3.93 and 0.87 lm, respectively.
Restriction of intake of dairy fats, animal fats and fish
oils is a simple and effective method of reducing phy-
tanic acid intake.
In the wider context, branched-chain fatty acid
metabolism could have wide-reaching implications.
The number of structures that could be theoretically
metabolized by this route is large (in some cases, preli-
minary metabolism by x-oxidation is required). These
include fat-soluble vitamins such as vitamin E and
many plant sterols and fats. This implies that a large
number of dietary fats could be either protective or
procarcinogenic.
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