Characterization of the metabolism of fenretinide by human liver microsomes, cytochrome P450 enzymes and UDP-glucuronosyltransferases

Article abstract of DOI:10.1111/j.1476-5381.2010.01104.x

Background and Purpose Fenretinide (4-HPR) is a retinoic acid analogue, currently used in clinical trials in oncology. Metabolism of 4-HPR is of particular interest due to production of the active metabolite 4-oxo 4-HPR and the clinical challenge of obtaining consistent 4-HPR plasma concentrations in patients. Here, we assessed the enzymes involved in various 4-HPR metabolic pathways. Experimental Approach Enzymes involved in 4-HPR metabolism were characterized using human liver microsomes (HLM), supersomes over-expressing individual human cytochrome P450s (CYPs), uridine 5-diphospho-glucoronosyl transferases (UGTs) and CYP2C8 variants expressed in Escherichia coli. Samples were analysed by high-performance liquid chromatography and liquid chromatography/mass spectrometry assays and kinetic parameters for metabolite formation determined. Incubations were also carried out with inhibitors of CYPs and methylation enzymes. Key Results HLM were found to predominantly produce 4-oxo 4-HPR, with an additional polar metabolite, 4-hydroxy 4-HPR (4-OH 4-HPR), produced by individual CYPs. CYPs 2C8, 3A4 and 3A5 were found to metabolize 4-HPR, with metabolite formation prevented by inhibitors of CYP3A4 and CYP2C8. Differences in metabolism to 4-OH 4-HPR were observed with 2C8 variants, CYP2C8 4 exhibited a significantly lower V_max value compared with 1. Conversely, a significantly higher V_max value for CYP2C8 4 versus 1 was observed in terms of 4-oxo formation. In terms of 4-HPR glucuronidation, UGTs 1A1, 1A3 and 1A6 produced the 4-HPR glucuronide metabolite. Conclusions and Implications The enzymes involved in 4-HPR metabolism have been characterized. The CYP2C8 isoform was found to have a significant effect on oxidative metabolism and may be of clinical relevance.

Full text of DOI:10.1111/j.1476-5381.2010.01104.x

DOI:10.1111/j.1476-5381.2010.01104.x
www.brjpharmacol.org
British Journal of
Pharmacology
BJP
Correspondence
RESEARCH PAPERbph_1104 989..999
Dr Gareth J. Veal, Northern
Institute for Cancer Research,
Paul O’Gorman Building, Medical
School, Framlington Place,
University of Newcastle upon
Tyne, Newcastle upon Tyne NE2
4HH, UK. E-mail:
Characterization of
the metabolism of
G.J.Veal@ncl.ac.uk
----------------------------------------------------------------
fenretinide by human liver
microsomes, cytochrome
P450 enzymes and
UDP-glucuronosyltransferases
NA Illingworth1, AV Boddy1, AK Daly2 and GJ Veal1
Keywords
fenretinide; metabolism;
microsomes; cytochrome
P450; CYP2C8; UDP-
glucuronosyltransferases
----------------------------------------------------------------
Received
21 July 2010
Revised
16 September 2010
Accepted
12 October 2010
1Northern Institute for Cancer Research and 2Institute of Cellular Medicine, Newcastle University,
Newcastle upon Tyne, UK
BACKGROUND AND PURPOSE
Fenretinide (4-HPR) is a retinoic acid analogue, currently used in clinical trials in oncology. Metabolism of 4-HPR is of
particular interest due to production of the active metabolite 4′-oxo 4-HPR and the clinical challenge of obtaining consistent
4-HPR plasma concentrations in patients. Here, we assessed the enzymes involved in various 4-HPR metabolic pathways.
EXPERIMENTAL APPROACH
Enzymes involved in 4-HPR metabolism were characterized using human liver microsomes (HLM), supersomes over-expressing
individual human cytochrome P450s (CYPs), uridine 5′-diphospho-glucoronosyl transferases (UGTs) and CYP2C8 variants
expressed in Escherichia coli. Samples were analysed by high-performance liquid chromatography and liquid
chromatography/mass spectrometry assays and kinetic parameters for metabolite formation determined. Incubations were also
carried out with inhibitors of CYPs and methylation enzymes.
KEY RESULTS
HLM were found to predominantly produce 4′-oxo 4-HPR, with an additional polar metabolite, 4′-hydroxy 4-HPR (4′-OH
4-HPR), produced by individual CYPs. CYPs 2C8, 3A4 and 3A5 were found to metabolize 4-HPR, with metabolite formation
prevented by inhibitors of CYP3A4 and CYP2C8. Differences in metabolism to 4′-OH 4-HPR were observed with 2C8 variants,
CYP2C8*4 exhibited a significantly lower Vmax value compared with *1. Conversely, a significantly higher Vmax value for
CYP2C8*4 versus *1 was observed in terms of 4′-oxo formation. In terms of 4-HPR glucuronidation, UGTs 1A1, 1A3 and 1A6
produced the 4-HPR glucuronide metabolite.
CONCLUSIONS AND IMPLICATIONS
The enzymes involved in 4-HPR metabolism have been characterized. The CYP2C8 isoform was found to have a significant
effect on oxidative metabolism and may be of clinical relevance.
Abbreviations
13-cisRA, 13-cis retinoic acid; 4-HPR, fenretinide; 4-MPR, 4-methoxy fenretinide; ATRA, all-trans retinoic acid; COMT,
catechol-O-methyltransferases; CYP, cytochrome P450; HIM, human intestinal microsomes; HLM, human liver
microsomes; PMT, phenol methyltransferase; SAM, S-adenosyl methionine; UGT, uridine 5′-diphospho-glucoronosyl
transferases
© 2011 The Authors
British Journal of Pharmacology © 2011 The British Pharmacological Society
British Journal of Pharmacology (2011) 162 989–999 989

NA Illingworth et al.
BJP
CYP supersomes and UGT supersomes were supplied by BD
Biosciences (Oxford, UK). Bactopeptone, bactotryptone, yeast
extract and bactoagar were purchased from Difco Laborato-
ries (East Mosely, UK). ECL Plus Western blotting detection
reagents were obtained from G E Healthcare (Buckingham-
shire, UK). Tris-glycine gels were supplied by Invitrogen
(Paisley, UK). JM109 competent cells were purchased from
Promega (Southampton, UK). Carbon monoxide was sup-
plied by BOC gases (Guildford, UK). high-performance liquid
chromatography (HPLC)-grade solvents were from Fisher Sci-
entific (Loughborough, UK). All other chemicals and reagents
were purchased from Sigma-Aldrich (Poole, UK).
Introduction
Fenretinide (4-HPR) is a synthetic analogue of retinoic acid,
currently used in clinical trials in children for the treatment of
neuroblastoma. In addition, Ewing’s sarcoma cell lines have
been shown to be sensitive to 4-HPR treatment in vitro (Lovat
et al., 2000; Myatt and Burchill, 2007) and 4-HPR has been
shown to be effective in pre-clinical animal studies (Ponthan
et al., 2003; Maurer et al., 2007). As both of these paediatric
cancers have 5-year survival rates of less than 60% (Gatta
et al., 2002), improvements in their clinical treatment are
clearly needed. Unlike other retinoid derivatives currently in
use in the clinic, such as all-trans retinoic acid (ATRA) and
13-cis retinoic acid (13-cisRA), 4-HPR induces apoptosis as
well as differentiation (Kitareewan et al., 1999), and is also
better tolerated than other retinoids (Garaventa et al., 2003).
The metabolism of both ATRA and 13-cisRA have previously
been characterized, with cytochrome P450 enzymes (CYPs)
including 2C8, 3A7, 3A5, 2C18, 3A4 and 2C9 having been
identified in the metabolism of ATRA, and CYPs including
2C8, 3A7, 4A11, 1B1, 2B6 and 2C9 responsible for the metabo-
lism of 13-cisRA (Marill et al., 2000; 2002; McSorley and Daly,
2000). While 4-HPR metabolism is similar to that of ATRA and
13-cis retinoic acid in many respects, 4-HPR produces an
active metabolite, 4′-oxo fenretinide (4′-oxo 4-HPR), which is
able to act synergistically with 4-HPR and is also active against
some 4-HPR-resistant cell lines (Villani et al., 2006). 4-HPR is
also metabolized to an inactive metabolite in the form of
4-methoxy fenretinide (4-MPR) (Mehta et al., 1998), with sig-
nificant metabolism to both of these metabolites observed in
clinical trials (Villani et al., 2004; Villablanca et al., 2006;
Formelli et al., 2008). Although the formation of glucuronide
metabolites of fenretinide has not previously been reported,
this metabolic pathway may be applicable to fenretinide as
glucuronidation of other retinoids has been described by our
group and others (Czernik et al., 2000; Samokyszyn et al.,
2000; Rowbotham et al., 2010a). As one of the main limita-
tions to the clinical development of orally administered
4-HPR is the achievement of effective and consistent plasma
concentrations in patients (Maurer et al., 2007), metabolism
of the parent drug is clearly an important issue. The aim of the
current study was to characterize the in vitro metabolism of
4-HPR in human liver microsomes (HLM), supersomes over-
expressing individual human CYPs, CYP2C8 variants ex-
pressed in Escherichia coli and a panel of recombinant human
uridine 5′-diphospho-glucoronosyl transferases (UGTs).
Liquid chromatography/mass spectrometry
(LC/MS) analysis of 4-HPR and metabolites
Separation of 4-HPR and its metabolites was achieved using a
Perkin Elmer LC (Beaconsfield, UK) system, consisting of a
vacuum degasser, two series 200 pumps, a thermostatically
controlled series 200 autosampler and a Waters 2487 UV
absorbance detector. Reverse-phase chromatography was per-
formed using a Luna 3 mm C18 50 ¥ 2 mm column with a
flow rate of 250 mL·min-1 and an injection volume of 10 mL.
Mobile phases consisted of (A) 40% acetonitrile/60% (0.2%)
acetic acid and (B) acetonitrile/0.2% acetic acid. A linear
gradient ran from 100% A at 0 min to 100% B at 3 min, at
which it was maintained for 2.5 min before returning to
100% A. An Applied Biosystems (Warrington, UK) API-2000
liquid chromatography/mass spectrometry/mass spectro-
metry (LC/MS/MS) triple Q (quadrupole) mass spectrometer
with electrospray ionization source, controlled by Analyst
software, was operated in single quadrupole negative mode
for the detection of 4-HPR [multiple reaction monitoring
(MRM) of 392/283], 4′-hydroxy 4-HPR (4′OH 4-HPR) (MRM
407/299) and 4′-oxo 4-HPR (MRM 406/297).
HPLC analysis of 4-HPR and metabolites
HPLC analysis was carried out using a Waters 2690 Separa-
tions Module and 996 photodiode array (PDA) detector
(Waters Ltd, Elstree, UK), with Waters Millennium software
for data acquisition, based on the previously published
method of Formelli et al. (1993). Separation of 4-HPR, 4′-OH
4-HPR, 4′-oxo 4-HPR and 4-MPR was achieved using a Waters
Symmetry C18 3.5 mm (4.6 ¥ 150 mm) column, with mobile
phases (A) 70% acetonitrile/30% (0.2%) acetic acid and (B)
acetonitrile/0.2% acetic acid.
A linear gradient ran at
1 mL·min-1 from 100% A at 0 min to 100% B at 20 min,
returning to 100% A for 10 min to re-equilibrate the column.
A sample volume of 50 mL was injected onto the column for
analysis. Separation of 4-HPR and its glucuronide metabolites
was achieved using a Luna C18 (2) column (50 ¥ 2.0 mm,
3 mm) (Phenomenex, Torrance, CA, USA), was used with
mobile phases (A) 0.1% acetic acid, pH 5.0 and (B) 100%
acetonitrile. A linear gradient ran at 0.2 mL·min-1 from 60% A
at 0 min to 100% B at 6 min, maintaining at 100% B for
4 min and returning to 60% A for 10 min to re-equilibrate the
column. A sample volume of 20 mL was injected onto the
column for analysis. The limit of quantification of the assay
was 0.01 mg·mL-1 and the inter-assay coefficients of variation
were 4.3–6.5% for 4-HPR and metabolites.
Methods
Chemicals
CYP2C8 clones were kindly provided by Dr Frank J Gonzalez
of the National Cancer Institute, Bethesda, MD, USA. The
human P450 reductase clone was kindly provided by Dr
Thomas Friedberg of the University of Dundee. 4-HPR and
4-MPR were generously provided by Cancer Research UK and
4′-oxo-4-HPR was provided by High Force Research Ltd.
(Durham, UK). Anti-CYP reductase, anti-CYP 2C8 and horse-
radish peroxidase-linked donkey anti-rabbit IgG antibodies
were purchased from Millipore (Watford, UK). HLM, HIM,
990 British Journal of Pharmacology (2011) 162 989–999

Characterization of fenretinide metabolism
BJP
rabbit anti-CYP2C8 and rabbit anti-P450 reductase antibody,
followed by horseradish peroxidase-linked donkey anti-rabbit
IgG secondary antibody. Detection was by enhanced chemi-
luminescence. CYP content was measured using Fe2+-CO
versus Fe2+ difference spectra and P450 reductase content was
measured using the cytochrome C reductase assay as previ-
ously described (Rowbotham et al., 2010b).
Kinetic parameters for the formation of 4′-OH 4-HPR and
4′-oxo 4-HPR were determined following incubations of
0–1 mg·mL-1 protein of CYP2C8 isoform membrane fractions
with 50 mM 4-HPR, or 0–100 mM 4-HPR with 0.25 mg·mL-1
protein. Because of the lack of an authentic standard for
4′-OH 4-HPR, Vmax results are given in peak area U·min-1.
One-way analysis of variance was used to compare Km and
Vmax data, with P < 0.05 used to determine significance. Indi-
vidual CYP2C8 variants were then compared with wild type
by use of Student’s t-test. All calculations were performed
with GraphPad Prism version 4.0 software (GraphPad Soft-
ware Inc.).
Incubation of 4-HPR with HLM, HIM, CYP
supersomes and UGT supersomes
HLM or a panel of CYP supersomes (up to 1 mg·mL-1 protein)
over-expressing individual human CYPs were incubated with
50 mM 4-HPR in 0.1 M phosphate buffer, pH 7.4, containing
1 mM MgCl2 and 2 mM nicotinamide adenine dinucleotide
phosphate in a final volume of 200 mL for 3 h. CYPs 1A6, 2B6,
2E1, 3A4, 3A5, 2C8, 2C9 and 2C19 were used. For experiments
investigating 4-HPR glucuronidation, HLM, HIM or a panel of
UGT supersomes over-expressing individual UGTs were incu-
bated with 200 mM 4-HPR in an incubation mixture consisting
of alamethecin (25 mg·mL-1), MgCl2 (8 mM), uridine diphos-
phate glucuronic acid (2 mM) and methanol (0.25%), made up
to a final volume (50 mL) with Tris-HCl buffer (50 mM, pH 7.5).
UGTs 1A1, 1A3, 1A4, 1A6, 1A7, 1A8, 1A9, 1A10, 2B4, 2B5, 2B15
and 2B17 were used. Incubations were carried out at 37°C for
3 h. Reactions were initiated by the addition of enzyme and
terminated with ice-cold acetonitrile (100 mL). To confirm that
the metabolite produced was a glucuronide, b-glucuronidase
(800 U) in potassium phosphate buffer (75 mM, pH 6.8) was
added to samples prepared from the incubations as described
above, and the mixture was incubated at 37°C for 30 min. All
reactions were terminated by the addition of ice-cold acetoni-
trile (400 mL) and samples were centrifuged at 10 000¥ g for
5 min to remove all protein. Supernatant was then retained for
HPLC analysis. Experiments were carried out in parallel, with
all comparative samples for a defined experiment being analy-
sed within a single assay.
Inhibition of 4-MPR
Known inhibitors or competitive substrates of several methy-
lating enzymes were added to HLM incubations. Compounds
used were: 4-nitrocatechol, nialamide, pargyline [inhibitors
of catechol-O-methyltransferases (COMT)], acetaminophen
[inhibitor of phenol methyltransferase (PMT)] and imidazole
(competitive substrate for amino-N-methyltransferase). HLM
(0.5 mg·mL-1) were incubated with 50 mM 4-HPR, 0.2 mM
SAM and 0–5 mM inhibitor in 0.1 M phosphate buffer, pH
7.4, containing 1 mM MgCl2 and 2 mM NADPH in a final
volume of 200 mL for 3 h. The reaction was stopped by addi-
tion of 400 mL acetonitrile, and samples were centrifuged at
10 000¥ g for 5 min to remove all protein. Supernatant was
then retained for HPLC analysis.
Determination of kinetic parameters for
4Ј-OH 4-HPR, 4Ј-oxo 4-HPR, 4-MPR and
4-HPR glucuronide formation
Kinetic parameters for the formation of 4′-OH 4-HPR, 4′-oxo
4-HPR and 4-MPR were determined following incubations
of 0–1 mg·mL-1 protein of HLM or CYP3A4, 3A5 or 2C8
supersomes with 50 mM 4-HPR, or 0–100 mM 4-HPR with
0.25 mg·mL-1 protein. Incubations to determine 4-MPR for-
mation also had 0.2 mM S-adenosyl methionine (SAM) added
as a methylation co-factor. Kinetic parameters for the forma-
tion of the glucuronide metabolite of 4-HPR were determined
following incubations of 0–1.5 mg·mL-1 protein of HLM, HIM
or UGTs 1A1, 1A3 or 1A6 with 200 mM 4-HPR, or 0–2 mM
4-HPR with 1 mg·mL-1 protein. Calculations were performed
with GraphPad Prism version 4.0 software (GraphPad Soft-
ware Inc., San Diego, CA, USA). Because of the lack of authen-
tic standards for 4′-OH 4-HPR and 4-HPR glucuronide, Vmax
results were calculated in peak area U·min-1.
Results
Chromatographic analysis of 4-HPR
and its metabolites
Following incubation of 4-HPR with HLM, the metabolites
produced were analysed by LC/MS/MS and HPLC. Peaks were
identified for 4-HPR and its two main polar metabolites,
4′OH-4-HPR and 4′-oxo 4-HPR, in addition to 4-HPR glucu-
ronide, as shown in Figure 1. 4-HPR and 4′-oxo 4-HPR were
identified by co-elution with authentic standards and by
their mass spectrometry/mass spectrometry (MS/MS) profiles.
4′-OH 4-HPR was identified by its MS/MS profile alone and
4-HPR glucuronide was identified by peak removal following
incubation of samples with b-glucuronidase, as no authentic
standards were available for either of these metabolites.
Retention times (RT) for the HPLC assay were 3.5 min,
3.9 min and 13.5 min for 4′-OH 4-HPR, 4′-oxo 4-HPR and
4-HPR respectively. RT for the HPLC assay used for identifi-
cation of 4-HPR glucuronide were 6.9 min for 4-HPR glucu-
ronide and 11.1 min for 4-HPR. Additional peaks seen at
similar RT to 4-HPR were isomers of 4-HPR. LC/MS/MS analy-
sis identified 4-HPR with an MRM of 392/283 (RT 5. 8 min),
4′-OH 4-HPR with an MRM of 407/299 (RT 4.3 min), and
4′-oxo 4-HPR with an MRM of 406/297 (RT 5.1 min).
Determination of kinetic parameters for
4Ј-OH 4-HPR and 4Ј-oxo 4-HPR formation
by CYP2C8 variants
JM109 high competency E. coli cells were transfected with
plasmids for CYP variants *1 (wild type), *3, *4 or an empty
plasmid (control), and co-transfected with CYP reductase
prior to the generation of membrane fractions as previously
described (Rowbotham et al., 2010b). The level of CYP2C8
and P450 reductase expression was assessed by sodium
dodecyl sulphate-polyacrylamide gel electrophoresis on
4–20% tris-glycine gels followed by Western blotting with
British Journal of Pharmacology (2011) 162 989–999 991

NA Illingworth et al.
BJP
Figure 1
Representative chromatograms showing separation of fenretinide (4-HPR) and metabolites by LC/MS/MS (A) and reversed-phase high-
performance liquid chromatography (B, C and D). Metabolites were generated following a 3-h incubation of 20 mM 4-HPR with 0.5 mg·mL-1
human liver microsomes (HLM; A, B and C) or following a 3-h incubation of 200 mM 4-HPR with 0.5 mg·mL-1 HLM (D). Generation of methoxy
fenretinide (4-MPR) is shown in (C) in the presence of 0.2 mM S-adenosyl methionine (SAM).
4-HPR concentration as shown in Figure 3, with kinetic
parameters for all three CYPs and HLM presented in Table 1.
Incubation of 4-HPR with CYP supersomes
4-HPR 50 mM was incubated with a panel of supersomes over-
HLM were the most effective at metabolizing 4-HPR to the
known active metabolite 4′-oxo 4-HPR, with a Vmax of 131
peak area U·min-1, as compared with Vmax values of 30, 25 and
0 peak area U·min-1 for of the individual CYPs 2C8, 3A4 and
3A5 respectively. In terms of 4′-OH 4-HPR production, HLM
exhibited a Vmax of 28 peak area U·min-1, as compared with
Vmax values of 115–282 peak area U·min-1 for supersomes
expressing individual CYPs. Results expressed as values nor-
malized for CYP content are also shown for the individual
human CYPs in Table 1. Significant differences were observed
in Vmax values between CYP2C8 and CYP3A4 for both 4′-oxo
and 4′-OH metabolite formation (P = 0.0001 and P = 0.0005,
respectively) and between CYP2C8 and CYP3A5 for 4′-OH
formation (P < 0.0001). Because of the level of metabolism
seen with CYP2C8, and as 2C8 is known to be polymorphic,
the effect of CYP2C8 variants on metabolism was then
investigated.
expressing individual human CYPs, to identify the enzymes
responsible for the oxidation and hydroxylation of 4-HPR.
While all CYPs tested were able to generate 4′-OH 4-HPR, as
shown in Figure 2, CYPs 3A4, 3A5 and 2C8 clearly produced
the highest levels. Only CYPs 3A4 and 2C8 were able to
generate 4′-oxo 4-HPR, the known active metabolite of
4-HPR.
Determination of kinetic parameters for
4Ј-OH 4-HPR, 4Ј-oxo 4-HPR and
4-MPR formation
4-HPR was incubated with HLM or supersomes over-
expressing individual human CYPs to determine enzyme
kinetic parameters. CYPs 3A4, 3A5 and 2C8 were used, as
these had been shown to metabolize 4-HPR to 4′-OH 4-HPR
and 4′-oxo 4-HPR. Metabolite production was related to
992 British Journal of Pharmacology (2011) 162 989–999

Characterization of fenretinide metabolism
BJP
Figure 2
Formation of 4′-OH 4-HPR and 4′-oxo fenretinide (4′-oxo 4-HPR)
metabolites of 4-HPR by a panel of supersomes over-expressing
individual human cytochrome P450s. Metabolite formation was
determined by high-performance liquid chromatography analysis.
Control supersomes were from cells transfected with an empty vector.
Metabolites were generated by 3-h incubation of 50 mM 4-HPR with
1 mg·mL-1 of each supersome for 3 h. Results are mean Ϯ SD from
three independent experiments. HLM, human liver microsomes.
Determination of kinetic parameters for
4Ј-OH 4-HPR and 4Ј-oxo 4-HPR formation
by CYP2C8 variants
The CYP content of E. coli transfectants varied from 175–
295 pmol·mg-1 of protein, and was comparable to CYP2C8
supersomes, which contained 293 pmol of CYP mg-1
protein. CYP reductase concentrations ranged from 180–
213 nmol·mg-1·min-1, again comparable to CYP2C8 super-
somes (176 nmol·mg-1·min-1). 4-HPR was incubated with each
CYP2C8 variant to determine enzymatic kinetic parameters.
Metabolite production with increasing 4-HPR concentration is
shown in Figure 4, with kinetic parameters provided in
Table 2. Differences in metabolism to 4′-OH 4-HPR and 4-oxo
4-HPR were observed with the 2C8 variants, most notably with
CYP2C8*4. In terms of 4′-OH 4-HPR production, although
there were no significant differences in Km values for *1, *3 and
*4 (5.6, 8.6 and 4.3 mM, respectively), Vmax values for *3 (0.24
peak area U·min-1·pmol-1 CYP) and *4 (0.11 peak area
U·min-1·pmol-1 CYP) were significantly different from *1,
which had a Vmax of 0.20 peak area U·min-1·pmol-1 CYP (P =
0.03 and 0.0025 respectively). Vmax/Km ratios for 4′-OH 4-HPR
were 0.028 and 0.026 for *3 and *4, respectively, as compared
with 0.036 for *1. Formation of 4′-oxo 4-HPR by CYP2C8 *1
was characterized by a Vmax of 0.04 peak area U·min-1·pmol-1
CYP and a Km of 19.3 mM. These results were comparable to
Vmax and Km values of 0.05 and 18.8 mM, respectively, for
CYP2C8*3. However, for CYP2C8*4, the Vmax was 0.128 peak
area U·min-1·pmol-1 CYP, and the Km was 59.8 mM, signifi-
cantly different from CYP2C8 *1 (P = 0.004).
Figure 3
Determination of kinetic parameters for the formation of (A) 4′-oxo
fenretinide (4′-oxo 4-HPR) and (B) 4′-OH 4-HPR metabolites by a
panel of supersomes over-expressing individual human CYPs. The
major CYPs (0.5 mg mL-1) found to metabolize 4-HPR (human cyto-
chrome P450s 3A4, 3A5 and 2C8) were incubated with 0, 2.5, 5, 10,
15, 20, 50 and 100 mM 4-HPR for 3 h. Metabolite formation was
determined by high-performance liquid chromatography analysis.
Results are mean Ϯ SD from three independent experiments.
(inhibitor of CYP3A4), either alone or in combination (data
not shown). Ketoconazole (100 mM) considerably inhibited
the production of both metabolites, with 75% inhibition of
4′-OH 4-HPR and 85% inhibition of 4′-oxo 4-HPR. Omepra-
zole alone at a concentration of 100 mM had very little effect
on the production of either metabolite, but in combination
with 100 mM ketoconazole resulted in complete inhibition of
the production of both metabolites.
Production of 4-MPR from HLM
Addition of the methylation co-factor SAM to incubations of
HLM and 4-HPR resulted in production of 4-MPR (Figure 1C),
with 4-MPR peak area increasing with increasing SAM con-
centrations up to 0.2 mM. This concentration of SAM was
added to all microsomal reactions investigating the produc-
tion of 4-MPR. Figure 5A shows the extent of formation of
4-MPR following incubations of 4-HPR at concentrations of
0–100 mM. A Km value of 236 mM was determined for 4-MPR
formation, markedly higher than those observed for 4′-OH
CYP inhibition
4-HPR was incubated with HLM and CYP inhibitors omepra-
zole (inhibitor of CYP2C8 and 2C9) and ketoconazole
British Journal of Pharmacology (2011) 162 989–999 993

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Table 1
Determination of kinetic parameters for the formation of the 4′-OH 4-HPR and 4′-oxo 4-HPR metabolites of 4-HPR by a panel of supersomes
over-expressing individual human CYPs and HLM
4Ј-oxo 4-HPR
Km (mM)
4Ј-OH 4-HPR
Km (mM)
Vmax
Vmax
(peak area
(peak area
(peak area
(peak area
U·min-1
)
U·min-1·pmol-1 CYP)
U·min-1
)
U·min-1·pmol-1 CYP)
HLM
3A4
3A5
2C8
9.3 Ϯ 3.2
2.8 Ϯ 1.1
N/D
131 Ϯ 14
25 Ϯ 2
N/D
N/A
17.8 Ϯ 9.4
4.8 Ϯ 1.2
19.1 Ϯ 5.0
2.2 Ϯ 1.1
28 Ϯ 6
N/A
0.43 Ϯ 0.03
N/D
115 Ϯ 7
118 Ϯ 6
282 Ϯ 24
2.05 Ϯ 0.1
1.84 Ϯ 0.1
1.45 Ϯ 0.1
5.0 Ϯ 3.1
30 Ϯ 5
0.18 Ϯ 0.03
The major CYPs found to metabolize 4-HPR (CYPs 3A4, 3A5 and 2C8) were incubated with 0, 2.5, 5, 10, 15 and 20 mM 4-HPR for 3 h.
Metabolite formation was determined by HPLC analysis. Results are expressed as mean Ϯ SD from n Ն 3 experiments. Vmax for 4′-oxo 4-HPR
significantly different for CYP3A4 and CYP2C8 (P = 0.0001); Vmax for 4′-OH 4-HPR significantly different for CYP3A4 and CYP2C8 (P = 0.0005)
and for CYP3A5 and CYP2C8 (P < 0.0001); Km for 4′-OH 4-HPR significantly different for CYP3A4 and CYP3A5 (P = 0.0085) and for CYP3A5
and CYP2C8 (P = 0.0046). Statistical analysis carried out on supersome data only, with Vmax results standardized to pmol CYP used for analysis.
4-HPR, fenretinide; 4′-OH 4-HPR, 4′-hydroxy fenretinide; 4′-oxo 4-HPR, 4′-oxo fenretinide; CYP, cytochrome P450; HIM, human intestinal
microsomes; HLM, human liver microsomes; N/A, no data available; N/D, not detected.
and 4′-oxo 4-HPR. Similarly, a calculated Vmax value of 823
peak U·min-1 for 4-MPR was higher than those observed for
the 4′-OH and 4′-oxo metabolites.
Inhibition of 4-MPR formation
Known inhibitors or competitive substrates of several methy-
lating enzymes were added to HLM incubations. Although
inhibitors of COMT and PMT had no effect on 4-MPR pro-
duction, imidazole (a competitive substrate for amine-N-
methyltransferase) inhibited 80% of 4-MPR production, but
only at a concentration of 5 mM, as shown in Figure 5B.
Incubation of 4-HPR with UGT supersomes
4-HPR (200 mM) was incubated with a panel of supersomes
expressing individual UGT enzymes, as well as HIM and
HLM, to identify the enzymes responsible for the production
of the glucuronide metabolite of 4-HPR (Figure 1D). Of the
UGTs included in the screen, only UGTs 1A1, 1A3 and 1A6, in
addition to HIM and HLM, produced 4-HPR glucuronide
(Figure 6). Incubation of samples with b-glucuronidase
resulted in complete removal of the metabolite peak, con-
firming it as a glucuronide.
Determination of kinetic parameters for
formation of 4-HPR glucuronide
4-HPR was incubated with HLM, HIM or individual UGT
enzymes to determine enzyme kinetic parameters. UGTs 1A1,
1A3 and 1A6 were used, as these had been shown to metabo-
lize 4-HPR to its glucuronide metabolites. Glucuronide pro-
duction was related to 4-HPR concentration as shown in
Figure 4
Figure 7, with kinetic parameters provided in Table 3. Vmax
Determination of kinetic parameters for the formation of (A) 4′-oxo
values of 0.62, 2.11 and 0.02 (peak area U·min-1·pmol-1 UGT)
fenretinide (4′-oxo 4-HPR) and (B) 4′-OH 4-HPR metabolites by
were determined for UGTs 1A1, 1A3 and 1A6, respectively,
with significant differences in Vmax values observed between
all three UGTs. Km values were high for all UGTs and
microsomes investigated, ranging from 389 mM for UGT 1A3
to 1212 mM for HIM.
CYP2C8 variants. E. coli membrane fractions (0.5 mg·mL-1
)
co-expressing CYP2C8 variants and P450 reductase were incubated
with 0, 2.5, 5, 10, 15 and 20 mM 4-HPR for 3 h. Metabolite formation
was determined by high-performance liquid chromatography analy-
sis. Results are mean Ϯ SD from three independent experiments.
994 British Journal of Pharmacology (2011) 162 989–999

Characterization of fenretinide metabolism
BJP
Table 2
Determination of kinetic parameters for the formation of the 4′-OH 4-HPR and 4′-oxo 4-HPR metabolites of 4-HPR by CYP2C8 variants
4Ј-oxo 4-HPR
Km (mM)
4Ј-OH 4-HPR
Km (mM)
Vmax (peak area
Vmax (peak area
U·min-1·pmol-1 CYP)
Vmax/Km
U·min-1·pmol-1 CYP)
Vmax/Km
2C8 *1
2C8 *3
2C8 *4
19.3 Ϯ 5.9
18.8 Ϯ 8.3
59.8 Ϯ 22.5
0.04 Ϯ 0.002
0.05 Ϯ 0.001
0.128 Ϯ 0.001
0.002
0.003
0.002
5.6 Ϯ 1.7
8.6 Ϯ 2.5
4.3 Ϯ 2.4
0.2 Ϯ 0.007
0.24 Ϯ 0.01
0.11 Ϯ 0.007
0.036
0.028
0.026
Individual CYP2C8 variants co-expressing P450 reductase, were incubated with 0, 2.5, 5, 10, 15 and 20 mM 4-HPR for 3 h. Metabolite
formation was determined by high-performance liquid chromatography analysis. Results are expressed as mean Ϯ SD from n Ն 3 experi-
ments. Vmax for 4′-oxo 4-HPR significantly different for *1 and *4 (P = 0.0037) and for *3 and *4 (P = 0.01); Km for 4′-oxo 4-HPR significantly
different for *1 and *4 (P = 0.0035) and for *3 and *4 (P = 0.039); Vmax for 4′-OH 4-HPR significantly different for *1 and *3 (P = 0.03),*1 and
*4 (P = 0.0025) and for *3 and *4 (P = 0.0004).
4-HPR, fenretinide; 4′-OH 4-HPR, 4′-hydroxy fenretinide; 4′-oxo 4-HPR, 4′-oxo fenretinide; CYP, cytochrome P450.
Figure 6
Formation of glucuronide metabolites of fenretinide (4-HPR) by a
panel of uridine 5′-diphospho-glucoronosyl transferases (UGT)
enzymes, human intestinal microsomes (HIM) and human liver
microsomes (HLM). Metabolite formation was determined by HPLC
analysis. Metabolites were generated by 3-h incubation of 200 mM
4-HPR with 1 mg·mL-1 of each UGT/microsome for 3 h. Results are
mean Ϯ SD from three independent experiments.
Discussion and conclusions
The in vitro metabolism of 4-HPR was investigated to charac-
terize the key metabolic pathways and the enzymes involved.
Elucidation of these enzymes is particularly important as
4-HPR is significantly metabolized in vivo to both active and
inactive moieties, in the form of 4′-oxo 4-HPR and 4-MPR
respectively (Villani et al., 2004; Villablanca et al., 2006;
Formelli et al., 2008). We have also identified 4′-OH 4-HPR as
an additional polar metabolite of 4-HPR formed in vitro. This
metabolite is likely to be one of several additional unidenti-
fied polar metabolites previously observed by Formelli et al.
(1989) in breast cancer patients treated with 4-HPR. 4-HPR
metabolism is of particular relevance to its clinical utility as
the achievement of effective, and consistent plasma concen-
trations of parent drug in patients has been a key limitation
to its clinical development (Maurer et al., 2007).
Figure 5
Formation of methoxy fenretinide (4-MPR) following a 3-h incuba-
tion of 0.5 mg·mL-1 human liver microsomes (HLM) with 0–100 mM
fenretinide (4-HPR) and 0.2 mM S-adenosyl methionine (SAM) (A)
and following a 3 h incubation of 0.5 mg·mL-1 HLM with 50 mM
4-HPR and 0.2 mM SAM in the presence of 0–5 mM imidazole (B).
British Journal of Pharmacology (2011) 162 989–999 995

NA Illingworth et al.
BJP
Initial experiments were carried out to identify the
metabolites produced from incubations of 4-HPR with HLM.
The individual enzymes responsible for production of these
metabolites were then investigated by incubation of 4-HPR
with supersomes over-expressing individual human CYP
enzymes. The panel of CYPs tested contained those known to
be involved in drug metabolism, including CYP3A4, as well as
those known to metabolize other retinoids (Marill et al.,
2000; 2002; McSorley and Daly, 2000; Zhang et al., 2000).
Metabolism to the newly identified metabolite, 4′-OH 4-HPR,
was catalysed by all CYPs tested, but to the greatest extent by
CYPs 3A4, 3A5 and 2C8. Metabolism to the active metabolite
4′-oxo 4-HPR was only achieved following incubations with
CYPs 2C8 and 3A4. This is a similar profile of CYP isoforms to
those previously identified as being involved in the metabo-
lism of 9-cis retinoic acid, 13-cis retinoic acid and ATRA to the
corresponding 4-oxo metabolites (Marill et al., 2000; 2002;
McSorley and Daly, 2000). It is possible that higher levels of
4-oxo production in HLMs may involve a degree of metabo-
lism not mediated by CYP. Metabolism by CYP2C8 was
further investigated, as it is known to be highly polymorphic
in the general population (Bahadur et al., 2002) and has pre-
viously been shown to play a role in the metabolism of cancer
drugs, including paclitaxel (Dai et al., 2001) and other retin-
oids (Rowbotham et al., 2010b).
Analysis of kinetic parameters of CYP2C8 isoforms dem-
onstrated differences in the metabolism of 4-HPR in terms
of metabolism to 4′-oxo 4-HPR and 4′-OH 4-HPR. Most
notably, the Vmax for the formation of 4′-OH 4-HPR was
higher for wild-type CYP2C8*1 (Vmax 0.2 peak area
U·min-1·pmol-1 CYP) as compared with CYP2C8*4 (Vmax 0.11
peak area U·min-1·pmol-1 CYP), with Vmax/Km ratios of 0.028
and 0.026 determined for *3 and *4, respectively, as com-
pared with 0.036 for *1. Conversely, the Vmax for the forma-
tion of 4′-oxo 4-HPR was lower for wild-type CYP2C8*1
(Vmax 0.04 peak area U·min-1·pmol-1 CYP) as compared with
CYP2C8*4 (Vmax 0.128 peak area U·min-1·pmol-1 CYP). These
data suggest that CYP2C8 genotype may have a consider-
able impact on the metabolism of 4-HPR, and consequently
on the clinical effects of this drug. This pathway of metabo-
lism may be particularly important in tumour cells which
are resistant to 4-HPR, but sensitive to the 4′-oxo metabolite
(Villani et al., 2006). Of interest, lower activity for
CYP2C8*3 and *4, as compared with wild type, has previ-
ously been reported in vitro for the metabolism of arachi-
donic acid to epoxyeicosatrienoic acids, with a greater risk
of renal toxicity associated with CYP2C8*3 genotype in
patients treated with calcineurin inhibitors (Smith et al.,
2008). As different expression systems were used, it was not
possible to directly compare kinetic parameters obtained
Figure 7
Determination of kinetic parameters for the formation of glucuronide
metabolites of fenretinide (4-HPR) by
a
panel of uridine
5′-diphospho-glucoronosyl transferases (UGT) enzymes, human
intestinal microsomes (HIM) and human liver microsomes (HLM);
1 mg·mL-1 of the major UGTs found to metabolize 4-HPR (1A1, 1A3
and 1A6, as well as HIM and HLM) were incubated with 0, 5, 25, 50,
75, 100, 150, 200, 250, 300, 500, 1000 and 2000 mM 4-HPR for 3 h.
Metabolite formation was determined by high-performance liquid
chromatography analysis. Results are mean Ϯ SD from three inde-
pendent experiments.
Table 3
Determination of kinetic parameters for the formation of the glucuronide metabolite of 4-HPR by a panel of uridine 5′-diphospho-glucoronosyl
transferases (UGT) enzymes, HIM and HLM
Vmax (peak area
U·min-1
Vmax (peak area
Km (mM)
)
U·min-1·pmol-1 UGT)
UGT 1A1
UGT 1A3
UGT 1A6
HIM
716 Ϯ 87
389 Ϯ 53
422 Ϯ 46
1212 Ϯ 202
540 Ϯ 69
577 Ϯ 34
317 Ϯ 18
124 Ϯ 6
0.62 Ϯ 0.04
2.11 Ϯ 0.12
0.02 Ϯ 0.001
N/A
418 Ϯ 38
679 Ϯ 39
HLM
N/A
The major UGTs found to metabolize 4-HPR (1A1, 1A3 and 1A6, as well as HIM and HLM) were incubated with 0, 5, 25, 50, 75, 100, 150,
200, 250, 300, 500, 1000 and 2000 mM 4-HPR for 3 h. Metabolite formation was determined by HPLC analysis. Results are expressed as mean
Ϯ SD from n Ն 3 experiments. Vmax significantly different for UGT1A1 and UGT1A3 (P < 0.0001), for UGT1A1 and UGT1A6 (P < 0.0001) and
for UGT1A3 and UGT1A6 (P < 0.0001); Km significantly different for UGT1A1 and UGT1A3 (P = 0.0051) and for UGT1A1 and UGT1A6 (P =
0.0066). Statistical analysis carried out on supersome data only, with Vmax results standardized to pmol UGT used for analysis.
4-HPR, fenretinide; 4′-OH 4-HPR, 4′-hydroxy fenretinide; 4′-oxo 4-HPR, 4′-oxo fenretinide; CYP, cytochrome P450; HIM, human intestinal
microsomes; HLM, human liver microsomes; N/A, no data available.
996 British Journal of Pharmacology (2011) 162 989–999

Characterization of fenretinide metabolism
BJP
from CYP2C8 variants expressed in E. coli with those
obtained from supersomes over-expressing CYP2C8.
However, it is reassuring that Vmax/Km ratios of comparable
magnitude were generated from experiments using the two
different approaches.
that this pathway of metabolism will play a significant role in
determining 4-HPR disposition in patients.
Retinoids are also known to be significantly metabolized
by CYP26, though the contribution of this metabolism
cannot be determined by the same methods as for the other
CYPs investigated, as supersomes over-expressing CYP26 are
not currently available. However, as ATRA is known to induce
CYP26 expression, pretreatment of cell lines with ATRA prior
to 4-HPR treatment, in conjunction with an assay to deter-
mine CYP26 expression would enable investigation into this
aspect of 4-HPR metabolism (Armstrong et al., 2005). Once
the role played by CYP26 in determining 4-HPR metabolism
has been fully ascertained, further ways to modulate this
metabolism through the use of retinoic acid metabolism
blocking agents may be investigated. Such an approach has
already been extensively studied with other retinoids and has
been found to consistently reduce retinoid metabolism
(Huynh et al., 2006; Njar et al., 2006; Armstrong et al., 2007).
The achievement of effective and consistent plasma con-
centrations in patients has been a major limitation to the
clinical development of 4-HPR. Clinical trials in neuroblas-
toma patients have shown huge variability in peak plasma
concentrations of 4-HPR, varying from 1–20 mM at higher
doses (Villablanca et al., 2006). Reformulation of 4-HPR from
the currently used oil-based capsules to a lipid matrix has
been shown to increase plasma concentrations up to seven-
fold in a mouse model (Maurer et al., 2007) and to reduce
inter-patient variation in a phase I trial in neuroblastoma
patients (Marachelian et al., 2009). Although this largely rep-
resents a formulation and bioavailability issue, it is clearly not
advantageous for a significant percentage of 4-HPR to be
metabolized to inactive metabolites. Indeed, as 4-HPR bio-
availability is improved, the level of metabolism to its active,
synergistic 4′-oxo 4-HPR metabolite, as compared with the
inactive 4-MPR metabolite, may become a more significant
factor in determining its overall effectiveness. Identification
of the enzymes responsible for 4-HPR metabolism in the
current study provides a stepping stone for future studies to
determine the potential role of these enzymes in determining
the efficacy of 4-HPR.
The effect of known CYP inhibitors was also investigated
to further corroborate the involvement of CYPs 3A4 and
2C8 in 4-HPR metabolism. 4-HPR was incubated with HLM
in the presence and absence of ketoconazole and/or ome-
prazole. Ketoconazole considerably inhibited the production
of both metabolites, with 75% inhibition of 4′-OH 4-HPR
and 85% inhibition of 4′-oxo 4-HPR at 100 mM. Omeprazole
alone, at a concentration of 100 mM, had very little effect on
either metabolite, but in combination with 100 mM keto-
conazole, complete inhibition of both metabolites was
observed. No effect was seen with omeprazole in the absence
of ketoconazole, as HLM have approximately 25 times more
CYP3A4 activity than CYP2C8, meaning that any inhibition
of CYP2C8 alone may be masked by the greater activity of
CYP3A4 as compared with CYP2C8 in this system. It should
be noted that these CYP inhibitors may be relatively non-
selective at the concentrations used in these experiments.
For example, ketoconazole has been shown to markedly
inhibit CYP2C8, as well as CYP3A4, at concentrations
>10 mM in a previous study (Ong et al., 2000). Similarly,
omeprazole has been shown to inhibit CYPs 2C9 and 2C19
(Li et al., 2004), although the role played by these two
enzymes in the metabolism of fenretinide would appear to
be minimal. The results obtained in the current study are
comparable to the effect of CYP inhibitors on other retin-
oids; for example, ATRA metabolism has previously been
shown to be inhibited approximately 90% by ketoconazole
(Schwartz et al., 1995).
In addition to 4′-OH 4-HPR and 4′-oxo 4-HPR, 4-MPR
has previously been identified as a major metabolite of
4-HPR (Swanson et al., 1980) and its pharmacokinetic prop-
erties have subsequently been investigated (Hultin et al.,
1990; Mehta et al., 1998; Vratilova et al., 2004). However,
the enzymes responsible for this methylation reaction have
not previously been characterized. The identification of
SAM as a necessary cofactor for 4-HPR methylation allowed
investigations into the formation of all three major 4-HPR
metabolites in a single in vitro system. Of several candidate
methylation enzymes, which require SAM as a cofactor,
Acknowledgements
This work was supported by the Medical Research Council
(MRC) and Research Councils UK (RCUK).
only
a limited number are microsomal. The current
study showed that 4-MPR production was not affected by
inhibitors of COMT or PMT, but was inhibited up to 80%
by imidazole,
a
competitive substrate for amine-N-
Conflicts of interest
methyltransferases, which are known to be microsomal and
are involved in the metabolism of many drugs and carcino-
gens (Ansher and Jakoby, 1986). It was not possible to
definitively identify the enzyme involved in this important
pathway of 4-HPR metabolism.
The authors declare no conflicts of interest.
Although the formation of 4-HPR glucuronide metabo-
lites has not previously been reported, we investigated the
potential for 4-HPR to be metabolized by a panel of recom-
binant human UGTs as this pathway has previously been
shown to be relevant to the metabolism of other retinoid
drugs (Czernik et al., 2000; Samokyszyn et al., 2000; Row-
botham et al., 2010a). Although UGTs 1A1, 1A3 and 1A6 were
shown to generate 4-HPR glucuronide, it would seem unlikely
References
Ansher SS, Jakoby WB (1986). Amine N-methyltransferases from
rabbit liver. J Biol Chem 261: 3996–4001.
Armstrong JL, Ruiz M, Boddy AV, Redfern CPF, Pearson ADJ,
Veal GJ (2005). Increasing the intracellular availability of all-trans
retinoic acid in neuroblastoma cells. Br J Cancer 92: 696–704.
British Journal of Pharmacology (2011) 162 989–999 997

NA Illingworth et al.
BJP
Armstrong JL, Taylor GA, Thomas HD, Boddy AV, Redfern CPF,
Veal GJ (2007). Molecular targeting of retinoic acid metabolism in
neuroblastoma: the role of the CYP26 inhibitor R116010 in vitro
and in vivo. Br J Cancer 96: 1675–1683.
Marachelian A, Kang MH, Hwang K, Villablanca JG, Groshen S,
Matthay KK et al. (2009). Phase I study of fenretinide (4-HPR) oral
powder in patients with recurrent or resistant neuroblastoma: New
Approaches to Neuroblastoma Therapy (NANT) Consortium trial. J
Clin Oncol 27 (15S): 10009.
Bahadur N, Leathart JBS, Mutch E, Steimel-Crespi D, Dunn SA,
Gilissen R et al. (2002). CYP2C8 polymorphisms in Caucasians and
their relationship with paclitaxel 6a-hydroxylase activity in human
liver microsomes. Biochem Pharmacol 64: 1579–1589.
Marill J, Cresteil T, Lanotte M, Chabot G (2000). Identification of
human cytochrome P450s involved in the formation of
all-trans-retinoic acid principal metabolites. Mol Pharmacol 58:
1341–1348.
Czernik PJ, Little JM, Barone GW, Raufman JP,
Radominska-Pandya A (2000). Glucuronidation of estrogens and
retinoic acid and expression of UDP-glucuronosyltransferase 2B7 in
human intestinal mucosa. Drug Metab Dispos 28: 1210–1216.
Marill J, Capron CC, Idres N, Chabot GG (2002). Human
cytochrome P450s involved in the metabolism of 9-cis- and
13-cis-retinoic acids. Biochem Pharmacol 63: 933–943.
Dai D, Zeldin DC, Blaisdell JA, Chanas B, Coulter SJ, Ghanayem BI
et al. (2001). Polymorphisms in human CYP2C8 decrease
metabolism of the anticancer drug paclitaxel and arachidonic acid.
Pharmacogenetics 11: 597–607.
Maurer BJ, Kalous O, Yesair DW, Wu X, Janeba J, Maldonado V
et al. (2007). Improved oral delivery of N-(4-hydroxyphenyl)
retinamide with a Novel LYM-X-SORB organized lipid complex.
Clin Cancer Res 13: 3079–3086.
Formelli F, Carsana R, Costa A, Buranelli F, Campa T, Dossena G
(1989). Plasma retinol level reduction by the synthetic retinoid
fenretinide: a one year follow-up study of breast cancer patients.
Cancer Res 49: 6149–6152.
Mehta RR, Hawthorne ME, Graves JM, Mehta RG (1998).
Metabolism of N-[4-hydroxyphenyl]retinamide (4-HPR) to
N-[4-methoxyphenyl]retinamide (4-MPR) may serve as a biomarker
for its efficacy against human breast cancer and melanoma cells.
Eur J Cancer 34: 902–907.
Formelli F, Clerici M, Campa T, Di Mauro MG, Magni A,
Mascotti G et al. (1993). Five-year administration of fenretinide:
pharmacokinetics and effects on plasma retinol concentrations. J
Clin Oncol 11: 2036–2042.
Myatt SS, Burchill SA (2007). The sensitivity of the Ewing’s sarcoma
family of tumours to fenretinide-induced cell death is increased by
EWS-Fli1-dependent modulation of p38MAPK activity. Oncogene
27: 985–986.
Formelli F, Cavadini E, Luksch R, Garaventa A, Villani MG,
Appierto V et al. (2008). Pharmacokinetics of oral fenretinide in
neuroblastoma patients: indications for optimal dose and dosing
schedule also with respect to the active metabolite 4-oxo-
fenretinide. Cancer Chemother Pharmacol 62: 655–665.
Njar VCO, Gediya L, Purushottamachar P, Chopra P, Vasaitis TS,
Khandelwal A et al. (2006). Retinoic acid metabolism blocking
agents (RAMBAs) for treatment of cancer and dermatological
diseases. Bioorg Med Chem 14: 4323–4340.
Garaventa A, Luksch R, Piccolo MSL, Cavadini E, Montaldo PG,
Pizzitola MR et al. (2003). Phase I trial and pharmacokinetics of
fenretinide in children with neuroblastoma. Clin Cancer Res 9:
2032–2039.
Ong C-E, Coulter S, Birkett DJ, Bhasker CR, Miners JO (2000). The
xenobiotic inhibitor profile of cytochrome P4502C8. Br J Clin
Pharmacol 50: 573–580.
Ponthan F, Lindskog M, Karnehed N, Castro J, Kogner P (2003).
Evaluation of anti-tumour effects of oral fenretinide (4-HPR) in rats
with human neuroblastoma xenografts. Oncol Rep 10: 1587–1592.
Gatta G, Capocaccia R, Coleman MP, Gloeckler LA, Berrino F
(2002). Childhood cancer survival in Europe and the United States.
Cancer 95: 1767–1772.
Rowbotham SE, Illingworth NA, Daly AK, Veal GJ, Boddy AV
(2010a). Role of UDP-glucuronosyltransferase isoforms in 13-cis
retinoic acid metabolism in humans. Drug Metab Dispos 38:
1211–1217.
Hultin TA, Filla MS, McCormick DL (1990). Distribution and
metabolism of the retinoid N-(4-Methoxyphenyl)-all-trans-
retinamide, the major metabolite of N-(4-Hydroxyphenyl)-all-trans-
Retinamide, in female mice. Drug Metab Dispos 18: 175–179.
Rowbotham SE, Boddy AV, Redfern CPF, Veal GJ, Daly AK (2010b).
Relevance of non-synonymous CYP2C8 polymorphisms to 13-cis
retinoic acid and paclitaxel hydroxylation. Drug Metab Dispos 38:
1261–1266.
Huynh CK, Brodie AMH, Njar VCO (2006). Inhibitory effects of
retinoic acid metabolism blocking agents (RAMBAs) on the growth
of human prostate cancer cells and LNCaP prostate tumour
xenografts in SCID mice. Br J Cancer 94: 513–523.
Samokyszyn VM, Gall WE, Zawada G, Freyaldenhoven MA,
Chen G, Mackenzie PI et al. (2000). 4-hydroxyretinoic acid,
a novel substrate for human liver microsomal UDP-
glucuronosyltransferase(s) and recombinant UGT2B7. J Biol Chem
275: 6908–6914.
Kitareewan S, Spinella MJ, Allopenna J, Reczek PR, Dmitrovsky E
(1999). 4HPR triggers apoptosis but not differentiation in retinoid
sensitive and resistant human embryonal carcinoma cells through
an RAR independent pathway. Oncogene 18: 5747–5755.
Li X-Q, Andersson TB, Ahlström M, Weidolf L (2004). Comparison
of inhibitory effects of the proton pump-inhibiting drugs
omeprazole, esomeprazole, lansoprazole, pantoprazole, and
rabeprazole on human cytochrome P450 activities. Drug Metab
Dispos 32: 821–827.
Schwartz EL, Hallam S, Gallagher RE, Wiernik PH (1995). Inhibition
of all-trans-retinoic acid metabolism by fluconazole in vitro and in
patients with acute promyelocytic leukemia. Biochem Pharmacol
50: 923–928.
Smith HE, Jones JP, Kalhorn TF, Farin FM, Stapleton PL, Davis CL
et al. (2008). Role of cytochrome P450 2C8 and 2J2 genotypes in
calcineurin inhibitor-induced chronic kidney disease.
Pharmacogenet Genomics 18: 943–953.
Lovat PE, Ranalli M, Annichiarrico-Petruzzelli M, Bernassola F,
Piacentini M, Malcolm AJ et al. (2000). Effector mechanisms of
fenretinide-induced apoptosis in neuroblastoma. Exp Cell Res 260:
50–60.
McSorley LC, Daly AK (2000). Identification of human cytochrome
P450 isoforms that contribute to all-trans-retinoic acid
4-hydroxylation. Biochem Pharmacol 60: 517–526.
Swanson BN, Zaharevitz DW, Sporn MB (1980). Pharmacokinetics
of N-(4-hydroxyphenyl)-all-trans-retinamide in rats. Drug Metab
Dispos 8: 168–172.
998 British Journal of Pharmacology (2011) 162 989–999

Characterization of fenretinide metabolism
BJP
Villablanca JG, Krailo MD, Ames MM, Reid JM, Reaman GH,
Reynolds CP (2006). Phase I trial of oral fenretinide in children
with high-risk solid tumors: a report from the Children’s Oncology
Group (CCG 09709). J Clin Oncol 24: 3423–3430.
identified fenretinide metabolite, induces marked G2-M cell cycle
arrest and apoptosis in fenretinide-sensitive and fenretinide-
resistant cell lines. Cancer Res 66: 3238–3247.
Vratilova J, Frgala T, Maurer BJ, Reynolds PC (2004). Liquid
chromatography method for quantifying
N-(4-hydroxyphenyl)retinamide and
N-(4-methoxyphenyl)retinamide in tissues. J Chrom B 808:
125–130.
Villani MG, Appierto V, Cavadini E, Valsecchi M, Sonnino S,
Curley RW et al. (2004). Identification of the fenretinide metabolite
4-oxo-fenretinide present in human plasma and formed in human
ovarian carcinoma cells through induction of cytochrome P450
26A1. Clin Cancer Res 10: 6265–6275.
Zhang Q, Dunbar D, Kaminsky L (2000). Human cytochrome P-450
metabolism of retinals to retinoic acids. Drug Metab Dispos 28:
292–297.
Villani MG, Appierto V, Cavadini E, Bettiga A, Prinetti A,
Clagett-Dame M et al. (2006). 4-oxo-fenretinide, a recently
British Journal of Pharmacology (2011) 162 989–999 999