Download Effects of phytanic acid on the vitamin E status, lipid composition and

Clinical Science (2001) 101, 697–705 (Printed in Great Britain)
Effects of phytanic acid on the vitamin E status,
lipid composition and physical properties
of retinal cell membranes: implications
for adult Refsum disease
S. P. YOUNG, A. W. JOHNSON and D. P. R. MULLER
Biochemistry, Endocrinology and Metabolism Unit, Institute of Child Health, University College London, 30 Guilford Street,
London WC1N 1EH, U.K.
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Adult Refsum disease is an inherited disorder in which phytanic acid accumulates in tissues and
serum. Two hypotheses have been proposed to explain the pathogenesis of this condition. The
molecular distortion hypothesis suggests that phytanic acid may alter membrane composition
and structure, thereby affecting membrane function(s). The anti-metabolite hypothesis suggests
that an accumulation of phytanic acid in membranes may interfere with vitamin E function. These
two hypotheses were investigated by studying the effects of modulating phytanic acid and
α-tocopherol concentrations on the fatty acid composition and certain physical parameters of
cultured retinal cells. Results showed that (a) the phospholipid fraction of retinal cells readily
incorporated phytanic acid, (b) the incorporation of phytanic acid increased membrane fluidity,
(c) there was no competition for uptake between phytanic acid and α-tocopherol, and (d) the
incorporation of phytanic acid did not increase the susceptibility of membranes to lipid
peroxidation in vitro. These results obtained with cultured retinal cells suggest that the
molecular distortion hypothesis, but not the anti-metabolite hypothesis, could explain the
pathogenesis of adult Refsum disease. In vitro tissue culture models can, however, only
approximate to the much more complex situation that occurs in vivo.
INTRODUCTION
Classical or adult Refsum disease (ARD) is an autosomal
recessive disorder in which the major clinical features are
degenerative changes in the retina and the central and
peripheral nervous systems. The disease is caused by an
isolated deficiency of the peroxisomal enzyme phytanoylCoA α-hydroxylase [1,2], which catalyses the first step in
the degradative pathway of the exogenous fatty acid,
phytanic acid. A number of inactivating mutations have
been reported in the gene encoding phytanoyl-CoA
α-hydroxylase, which is localized to chromosome 10p
[3,4]. The resulting accumulation of phytanic acid in the
serum and tissues of patients with ARD is thought to
contribute to the clinical features of this condition.
The earliest clinical manifestation of ARD is typically
night blindness, which suggests that the retina is more
sensitive to an accumulation of phytanic acid than other
tissues. The relative vulnerability of the retina compared
with other neural tissues may be a consequence of the
high rate of turnover of specialized membranes in the
photoreceptor outer segments, resulting in increased
Key words: lipid peroxidation, membrane fluidity, phytanic acid, Refsum disease, retina, vitamin E.
Abbreviations: ARD, adult Refsum disease ; DPH, diphenylhexatriene ; FAME, fatty acid methyl ester ; FBS, foetal bovine serum ;
MDA, malondialdehyde ; PUFA, polyunsaturated fatty acid(s) ; RPE, retinal pigment epithelium.
Correspondence: Dr D. P. R. Muller (e-mail D.Muller!ich.ucl.ac.uk).
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S. P. Young, A. W. Johnson and D. P. R. Muller
commercially available retinoblastoma (Y79) cell line
were used in these studies. Methods were validated for
the reproducible supplementation of these cell lines with
phytanic acid, vitamin E and linolenic acid, and the
following investigations were carried out : (a) examination of the fate of phytanic acid taken up by cells, (b)
the effects of phytanic acid uptake on membrane fluidity,
(c) the effects of phytanic acid supplementation on
α-tocopherol uptake, and (d) the susceptibility of
retinal membranes to lipid peroxidation.
Figure 1
Structures of α-tocopherol and phytanic acid
MATERIALS AND METHODS
Tissue culture
uptake of phytanic acid into this tissue. Phytanic acid
concentrations equivalent to 35.4 % and 9.6 % of the total
fatty acid content of the retina have been reported in two
patients with ARD [5].
Two principal hypotheses (the molecular distortion
hypothesis and the anti-metabolite hypothesis) have been
proposed to explain the pathogenesis of ARD [6–8]. The
molecular distortion hypothesis proposes that infiltration
of cellular fatty acid pools by phytanic acid will result in
its incorporation into membrane lipid structures at the
expense of other fatty acids. This, in turn, may affect the
physicochemical properties of membranes and their
associated functions. It has been suggested, for example,
that the demyelination that occurs in ARD is a result
of myelin destabilization due to incorporation of
phytanic acid into its structure [7]. An accumulation
of phytanic acid in the retina may, therefore, perturb
retinal membrane functions, such as that of the photoreceptor outer membranes or the apical membranes of
the retinal pigment epithelium (RPE).
The anti-metabolite hypothesis suggests that an accumulation of phytanic acid may interfere with the
function of vitamin E because of the structural similarity
between these two molecules [7] (see Figure 1). In support
of this hypothesis, a severe deficiency of vitamin E results
in a spectrum of features similar to those observed in
ARD, including peripheral and central neuropathy and
retinal degeneration [9]. Vitamin E is considered to be the
major lipid-soluble antioxidant [10]. It is able to break
the chain reaction of lipid peroxidation by reacting with
lipid peroxyl free radicals, and thus plays an important
role in protecting cell membranes from lipid peroxidation. It is, therefore, possible that an accumulation of
phytanic acid in tissues and cellular membranes may
perturb the function of vitamin E and induce a vitamin E
deficiency-like state.
In view of the susceptibility of the retina to phytanic
acid accumulation, this tissue was chosen for an investigation of the mechanism(s) involved in the pathogenesis
of ARD. An immortalized rat RPE cell line and a
# 2001 The Biochemical Society and the Medical Research Society
An immortalized rat RPE cell line [11] was kindly
supplied by Dr J. Greenwood (The Institute of Ophthalmology, London, U.K.). The cells were cultured in
Ham’s F10 medium containing 20 mM Hepes, 20 %
(v\v) foetal bovine serum (FBS), 0.09 % (w\v) Na CO ,
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1 mM glutamate and penicillin (100 units\ml)\
streptomycin (100 µg\ml) in a 5 % CO humidified
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atmosphere at 37 mC. The medium was replaced twice
weekly. Confluent cells were passaged and split 1 : 3 each
week by treatment with 0.05 % trypsin\EDTA in
Ca#+\Mg#+-free Hanks balanced salt solution for 5 min
at room temperature.
Y79, an established human retinoblastoma cell line [12]
that grows in suspension, was obtained from the
European Collection of Animal Cell Cultures (Porton
Down, Salisbury, Wilts., U.K.). The cells were maintained in RPMI-1640 medium containing 10 % (v\v)
FBS, 2 mM glutamine and penicillin (100 units\ml)\
streptomycin (100 µg\ml), at a density of (4–9)i10&
cells\ml in humidified 5 % CO at 37 mC. Fresh medium
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was added to the cultures twice weekly, replacing onethird of the suspension, which was either reseeded or
discarded.
Cells supplemented with phytanic acid and α-tocopherol were routinely harvested by trypsinization and
washed once with warmed FBS (37 mC) and then three
times with warmed (37 mC) PBS. Membranes were prepared by sonicating cell suspensions with 3i20 s bursts at
10 A using a probe sonicator (Soniprep 150 MSE). The
suspensions were cooled on ice between each burst to
minimize overheating. Sonicates were then centrifuged at
100 000 g for 1 h at 4 mC in a fixed-angle Beckman rotor
(70 Ti). After removing the supernatant, the membrane
pellet was washed with 3i1 ml of deionized water.
Lipid supplementation
The method used for supplementation of the culture
medium with fatty acids was based on that of Calder et al.
[13]. In summary, the medium was supplemented with
phytanic acid and\or linolenic acid (C : ,n− ) complexed
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to fatty acid-free BSA. Potassium salts of the fatty acids
Phytanic acid and retinal cell membranes
were prepared by incubating equimolar ratios (typically
250 µmol) of fatty acid and KOH at 70 mC for 1 h. After
cooling, the potassium salt was added to medium
containing fatty acid-free BSA, such that the final fatty
acid-free BSA concentration was 5 % (w\v), and the
mixture was rotated for 2 h at room temperature. The pH
was adjusted to 7.4 with 1 M KOH and the mixture was
sterilized through a 0.22 µm filter (Falcon) and stored at
k20 mC. The concentration of the fatty acid in the
medium was determined using GC with flame ionization
detection as described below, and the solution was diluted
with unsupplemented medium to achieve the desired
concentration of fatty acid.
Vitamin E (all-rac-α-tocopherol) from Sigma-Aldrich
Co. Ltd was dispersed in FBS as described by Tran and
Chan [14] before its addition to the medium. An aliquot
of 80 µl (or less depending upon the final concentration
required) of 500 mM α-tocopherol in acetone was added
to 30 ml of FBS at 37 mC. The acetone was removed with
nitrogen before mixing the tocopherol\FBS by rotation
at room temperature for 2 h. The tocopherol\FBS was
sterilized by passing it through a 0.22 µm filter. Aliquots
of 50 µl of the filtrate were extracted and the concentration of α-tocopherol was determined as described
below. Medium supplemented with α-tocopherol was
prepared fresh on the day of each feed.
Miscellaneous procedures
Lipid extraction
Lipids were extracted from the medium, disrupted cell
suspensions and crude membrane homogenates by the
method of Folch et al. [15]. Harvested cells were
resuspended in 300 µl of PBS (pH 7.4) and were disrupted
using a probe sonicator as described above. Crude
membranes were homogenized in a total of 500 µl of PBS,
using a 0.1 ml glass homogenizer. The lipid extract was
typically reconstituted in 40 µl of chloroform\methanol
(1 : 1, v\v).
Separation of neutral lipids
Lipids were separated by TLC using 20 cmi20 cm silica
gel plates (Supelco) and a solvent system of hexane\
diethyl ether\acetic acid (60 : 40 : 2 or 80 : 20 : 1, by vol.),
the latter being used for the separation of triacylglycerol
from cholesterol ester. After developing and drying at
room temperature, the lipid fractions were visualized by
exposure of the plate to iodine vapour and identified
by comparing their RF values with those of appropriate
standards.
Staining for neutral lipids
Confluent RPE cells and Y79 cells were harvested,
washed once with FBS at 37 mC and three times with PBS,
and then resuspended in PBS. Cytospin preparations of
the cells were made and the cells were fixed for 1–2 min
in formol\calcium acetate. After rinsing with water, the
cells were stained in Oil Red O solution at room
temperature for 1 h. The slides were then rinsed briefly in
70 % (v\v) ethanol, washed in water and counter-stained
with Carazzi’s haematoxylin for 4–5 min before being
mounted in glycerin jelly.
Synthesis of phytanic acid
Phytanic acid was synthesized from phytol (SigmaAldrich Co. Ltd) using the method of Jellum et al. [16],
which results in the formation of both phytanic acid and
dihydrophytol. Phytanic acid was separated and purified
using a column of 100 g of silica gel 60 (Sigma-Aldrich
Co. Ltd) prepared in 30 ml of hexane. The crude
preparation was dissolved in 150 ml of 2 % ethyl acetate
in hexane and applied to the column. The phytanic acid,
which could be seen as a yellow band, was washed
through the column with 6 % ethyl acetate. The first
170 ml of effluent was discarded and the phytanic acid
was collected in the next 350 ml. It was necessary to
purify this product further by repeating the chromatographic step. After two cycles of purification, the product
was analysed by GC-MS (see below) and found to be
96.4 % pure.
In vitro lipid peroxidation
Cells were resuspended and sonicated in 40 mM Trizma
buffer (pH 7.4) and adjusted to a protein concentration
of 5 mg\ml with the Trizma buffer. The sonicate was
then added to 10 mM ice-cold H O and the reaction was
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started by the addition of 0.2 mM CuSO , such that the
%
final proportions of sonicate\CuSO \H O were 5 : 1 : 1
% # #
(by vol.). Both the CuSO and H O were prepared in
%
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40 mM Trizma buffer. Following incubation for fixed
times at 37 mC, the reaction was stopped by the addition
of an equal volume of chilled acetonitrile, followed by
vortexing for 15 s. The protein precipitate was pelleted
by centrifugation at 13 000 g for 5 min. The samples were
then kept on ice and the free malondialdehyde (MDA),
which was used as a measure of lipid peroxidation, was
analysed (see below) within 3 h. Measurement of free
MDA concentrations in samples without oxidative stress
involved addition of the sonicate to the chilled acetonitrile prior to the addition of H O and CuSO .
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Analyses
Fatty acids
The direct one-step transesterification method of Lepage
and Roy [17], as modified by Liebich et al. [18], was used
to prepare total fatty acid methyl esters (FAMEs) from
the stock medium, lipid fractions, cells and crude
membrane preparations. FAMEs were analysed on a Pye
Unicam 4550 gas chromatograph with flame ionization
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S. P. Young, A. W. Johnson and D. P. R. Muller
detection, and were separated using an SP2330 fused
silica capillary column (30 m ; 0.32 mm internal diameter ;
0.2 µm film thickness) supplied by Supelco. The flow rate
of the helium carrier gas was 20 cm\s at a pressure of 0.5
kg\cm#. Injections of typically 1 µl were made in split
mode with a split ratio of 2 : 1. The injector port
temperature was 220 mC and that of the detector was
240 mC. FAMEs were separated using a temperature
programme starting at 140 mC, ramped to 180 mC at
4 mC\min and then to 210 mC at 1.5 mC\min, and finally
held at 210 mC for 5 min. Data were collected on an
SP4770 integrator (Spectra Physics Ltd). In general, fatty
acid composition is expressed as a percentage of total
fatty acids analysed ; however, when it was necessary to
quantify the fatty acids, heneicosanoic acid (C : ) was
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added as an internal standard.
Where necessary, the identity of fatty acids was
confirmed using GC-MS. The separation was carried out
on a Hewlett Packard 5890 Series II gas chromatograph,
and mass spectrometric analysis was performed using a
twin focusing magnetic sector mass spectrometer
(Autospec ; Fisons UK). The same column and temperature programmes were used as described for the
analysis by GC with flame ionization detection.
Injections of 0.5 µl were made in split mode with a split
ratio of 10 : 1. MS analysis was carried out using electronimpact ionization (70 eV) in positive-ion mode and total
ion current monitoring between m\z ratios of 50 and 700
at 1 scan\s.
20 µl injection volume, and 1–2 pmol per injection could
be detected.
Steady-state fluorescence anisotropy
Membrane suspensions equivalent to approx. 100 µg of
protein were incubated at 37 mC for 1 h in the dark with
2 ml of diphenylhexatriene (DPH) (5 µM in PBS). Fluorescent intensities were detected at 37 mC through a
polarizer orientated parallel (I) and perpendicular (IU)
to the direction of polarization of the excitation beam.
All measurements were performed using a PerkinElmer
LS3 fluorescence spectrophotometer, with excitation and
emission wavelengths of 360 nm and 430 nm respectively
and band widths of 10 nm. Fluorescence readings of
DPH in PBS alone were subtracted from the readings
given by DPH plus membranes for each polarizer setting.
RPE membrane suspensions in PBS alone produced
negligible fluorescence.
The polarization of fluorescence was expressed as the
fluorescence anisotropy, rs, where :
rs l (IkIU)\(Ij2IU)
Presentation of results
All results are expressed throughout as either means and
ranges or meanspS.E.M. The significance of differences
between mean values was calculated using Student’s
unpaired t test.
α-Tocopherol
α-Tocopherol was extracted from FBS and medium using
the method of Metcalfe et al. [19]. Extraction from retinal
cells was based on the method of Burton et al. [20] using
SDS to solubilize the membranes. α-Tocopherol concentrations were determined by HPLC with fluorimetric
detection as described previously [19]. Concentrations of
α-tocopherol were determined by direct comparison with
an external standard [5 µM all-rac-α-tocopherol (SigmaAldrich Co. Ltd) in hexane], and were expressed per mg
of protein. The inter- and intra-assay coefficients of
variation were both 5 %. The detection limit was of the
order of 2 pmol. Protein concentrations were determined
using the bicinchoninic acid protein assay [21,21a].
Free MDA
Free MDA was estimated by a modification [22] of the
method of Esterbauer et al. [23] using HPLC. Separations
were carried out on a Spherisorb S5NH column
(25 cmi4.6 mm ; Hichrom UK) using a mobile phase of
0.03 M Trizma (pH 7.0)\acetonitrile (21 : 79, v\v) at a
flow rate of 2.0 ml\min. The free MDA was detected at
267 nm. An MDA standard of known concentration was
prepared from 1,1,3,3-tetramethoxypropane by acid hydrolysis. The assay was linear up to at least 120 pmol\
# 2001 The Biochemical Society and the Medical Research Society
RESULTS
Lipid supplementation
The method of supplementation of cells with phytanic
acid was validated by examining the effects of time and
concentration on the uptake of phytanic acid. Initially,
confluent rat RPE cells were supplemented with 50 µM
phytanic acid over a period of 10 days with two changes
of medium, and the cells were harvested at fixed times.
The percentage of phytanic acid in the total lipid from the
crude membrane extracts increased rapidly from an initial
value of 0.5 % to 5.7 % after 2 days and then more slowly,
reaching 7.7 % after 10 days. Cells were also exposed to a
range of phytanic acid concentrations over 7 days with
one change of medium. The percentage of phytanic acid
increased linearly with concentration, from 0.6 % with
no supplementation to 32.8 % at 200 µM phytanic acid.
Similar results were obtained with Y79 cells. Based on
these results, RPE and Y79 cells were routinely supplemented with up to 200 µM phytanic acid for 7 days.
The method of supplementation of rat RPE and Y79
cells with α-tocopherol was validated by examining
uptake with increasing time and concentration of
α-tocopherol. Confluent rat RPE cells were supplemented
Phytanic acid and retinal cell membranes
with α-tocopherol (mean concentration of 92 µM) over a
period of 13 days. The medium was replaced on alternate
days and cells were harvested at various times and stored
under N in PBS at k20 mC until analysed. α-Tocopherol
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concentrations increased steadily with time ; from being
initially undetectable, they rose to 9 nmol\mg of protein
after 1 day, and to 30 and 46 nmol\mg after 7 and 13 days
respectively. RPE cells were also supplemented with a
range of α-tocopherol concentrations over 7 days.
Concentrations of α-tocopherol increased linearly from
just detectable with unsupplemented medium to a mean
concentration of 29.5 nmol\mg of protein with 85 µM
α-tocopherol. Thereafter the cellular α-tocopherol
concentration began to plateau.
Dose-dependent uptake of α-tocopherol by Y79 cells
was also observed. The increase in uptake was linear
between 0 and 75 µM supplementation, increasing from
undetectable to 8.4 nmol\mg of protein. At supplementation of 100 µM there was a sharp increase in the
α-tocopherol concentration to 33.5 nmol\mg, and
thereafter the concentration continued to rise linearly
to 46.3 nmol\mg at 150 µM supplementation.
Uptake of phytanic acid into cellular lipid
fractions
In order to investigate the fate of phytanic acid taken up
by the cells, the distribution of phytanic acid in the
different lipid fractions was determined using rat RPE
cells supplemented with 100 µM phytanic acid for 7 days.
The mean distribution of phytanic acid in two separate
experiments (as a percentage of phytanic acid in the
whole-cell lipid extract) was : triacylglycerol 48.3 %,
phospholipid 18.5 %, non-esterified fatty acid 8.1 % and
cholesterol ester 7.9 %. Similar results were obtained
with Y79 cells, with 60.3 % of the phytanic acid in the
triacylglycerol and 20.4 % in the phospholipid fraction.
These results indicated that phytanic acid taken up by the
two retinal cell lines was handled in a similar manner,
with a significant proportion incorporated into phospholipids, but with the major portion stored in the triacylglycerol fraction. Subsequent histochemical studies after
staining cells with Oil Red O indicated that neutral lipid
droplets accumulated in the cytoplasm of phytanic acidsupplemented cells.
The uptake of phytanic acid into the phospholipid
fraction of RPE cells was used as a marker of its uptake
into cellular membranes. A dose–response study was
carried out in which confluent RPE cells were supplemented with 0–200 µM phytanic acid for 7 days. The
percentage of phytanic acid in the phospholipid fraction
increased linearly from 0.3 % (unsupplemented) to
15.3 % (200 µM phytanic acid).
The fatty acid profiles of the RPE cell total lipid and
phospholipid fractions following supplementation with
0, 100 and 200 µM phytanic acid for 7 days are shown in
Figure 2 Fatty acid profiles of (a) total lipid and (b)
phospholipid fractions of rat RPE cells after supplementation
with various concentrations of phytanic acid for 7 days
phyt, phytanic acid.
Figures 2(a) and 2(b). The changes were similar for these
two lipid fractions. An increase in the relative amount of
phytanic acid was accompanied by decreases in the
percentages of all the fatty acids analysed (C : , C : ,
"' " ") !
C : , C : ,n− , C : ,n− , C : ,n− , C : ,n− and
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C : ,n− ), except for C : in the total lipid extract, and
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"% !
C : and C : in the phospholipid fraction. The changes
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in C : were not consistent in the total lipid extract. A
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similar fatty acid profile of the total lipid and phospholipid fractions was seen in Y79 cells following supplementation with 100 µM phytanic acid (results not
shown).
Membrane fluidity
Confluent RPE cells were supplemented with or without
150 µM phytanic acid for 7 days, and the anisotropy
values of membrane suspensions from eight separate
flasks per group were determined. The phytanic acidsupplemented cells had a significantly lower anisotropy
value (meanspS.E.M. : supplemented, 0.15p0.01 ;
unsupplemented, 0.18p0.01 ; P 0.001).
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Figure 3 Uptake of phytanic acid by rat RPE cells in the
presence of 100 µmol/l phytanic acid and increasing concentrations of α-tocopherol for 7 days
n l 3 for each point ; bars indicate ranges. Significance of differences : *P
0.001 compared with no added α-tocopherol.
Competition between phytanic acid and
α-tocopherol
Exposure of RPE cells to 50 µM α-tocopherol and
increasing concentrations of phytanic acid (0–200 µM)
did not have a significant effect on the α-tocopherol
concentration, which remained at approx. 20 nmol\mg
of protein despite the percentage phytanic acid content
increasing in the crude RPE membranes from 0 to 36 %.
In contrast, when confluent RPE cells were exposed to
100 µM phytanic acid and between 0 and 170 µM
α-tocopherol for 7 days, the mean percentage of phytanic
acid increased from 13.8 to 21.6 % (P 0.001) when the
α-tocopherol concentration of the medium was increased
from 0 to 100 µM (Figure 3). Over this concentration
range the α-tocopherol concentration in the RPE cells
increased from undetectable to 45 nmol\mg of protein.
This increase in percentage of phytanic acid in RPE cells
was a consistent finding. In one study the distribution of
phytanic acid within the various lipid fractions following
supplementation with increasing concentrations of
α-tocopherol was also investigated. The percentage of
phytanic acid in the phospholipid fraction increased from
5.8 to 8.4 % with an increase in the α-tocopherol
concentration in the medium from 0 to 100 µM. There
were no consistent changes in the percentage phytanic
acid content of the triacylglycerol, cholesterol ester or
non-esterified fatty acid fractions.
Another study was carried out to determine whether
phytanic acid supplementation would displace α-tocopherol already present in rat RPE cells after presupplemention with the vitamin. Confluent rat RPE cells
were supplemented with 150 µM α-tocopherol for
7 days, after which the α-tocopherol concentration in
crude membrane preparations was determined (day 0).
The cells were then supplemented with or without 150 µM
phytanic acid with no added α-tocopherol over the
subsequent 7 days, and cells from the two groups were
# 2001 The Biochemical Society and the Medical Research Society
Figure 4 Effects on the nk3 fatty acid composition of Y79
cells of supplementation with increasing concentrations of
linolenic acid (C18 : 3,n−3) for 7 days
n l 3 at linolenic acid concentrations of 0, 30 and 90 µmol/l ; otherwise n l 1.
Visible bars indicate ranges ; otherwise ranges fall within symbols. Significance of
differences : *P 0.001 compared with no added C18 : 3,n−3 ; jP 0.001
compared with 30 µM C18 : 3,n−3.
harvested on days 3 and 7. The mean (n l 3 for all
groups) α-tocopherol concentration on day 0 was
34.8 nmol\mg of protein. The addition of phytanic acid
did not significantly affect the mean α-tocopherol concentration at either time point. On day 3 the mean
α-tocopherol concentrations were 23.3 and 25.1 nmol\mg
with and without phytanic acid respectively, and on day
7 the values were 16.2 and 15.9 nmol\mg respectively.
The percentage of phytanic acid after 3 and 7 days of
supplementation was 17.8 and 19.1 % respectively, compared with 0.2 % at both time points in the cells without
added phytanic acid.
Lipid peroxidation
Studies were conducted to determine whether added
phytanic acid might influence the antioxidant properties
of α-tocopherol and thus result in increased lipid peroxidation of membrane polyunsaturated fatty acids
(PUFA). The Y79 cell line was used, as it is possible to
increase its content of long-chain PUFA by supplementation with linolenic acid (C : ,n− ) [24]. The changes in
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the percentages of C : ,n− and of the major nk3 PUFA
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after supplementing Y79 cells with 0–160 µM C : ,n−
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for 7 days are shown in Figure 4. Prior to supplementation, C : ,n− was undetectable, but its content
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increased to 5.1 % at 100 µM and to 26.4 % at 160 µM
C : ,n− . There was a significant increase in the per") $ $
centages of both C : ,n− and C : ,n− with increasing
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C : ,n− supplementation, with C : ,n− reaching a
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maximum of 8.3 % at 75 µM and C : ,n− a maximum of
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Phytanic acid and retinal cell membranes
MDA concentration after oxidative stress at all phytanic
acid concentrations.
DISCUSSION
Figure 5
cells
Free MDA concentrations in cell sonicates of Y79
Y79 cells were supplemented for 5 days with 100 µmol/l linolenic acid and
increasing concentrations of phytanic acid, in the presence or absence of
α-tocopherol (0.25 µmol/l), and with or without oxidative stress. Oxidative stress
was achieved by exposing cell sonicates to 10 mmol/l H2O2 and 0.2 mmol/l CuSO4.
, Oxidative stress alone ; $, oxidative stressjα-tocopherol ; , no
oxidative stress ; >, no oxidative stress, but with α-tocopherol. n l 3 at
phytanic acid concentrations of 0, 50 and 100 µmol/l ; otherwise n l 1. Visible
bars indicate ranges ; otherwise ranges fall within symbols. Significance of
differences : *P 0.05 compared with no added phytanic acid.
3.3 % at 95 µM supplementation. The percentage of
C : ,n− increased significantly (P 0.001) on sup## ' $
plementation with 30 µM C : ,n− , and reached a peak of
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12.3 % at 75 µM. Thereafter the percentage declined
markedly ; at 160 µM C : ,n− the percentage of C : ,n−
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was 2.3 %, which was lower than that observed without
any supplementation (4.6 %). The percentages of C : ,
"% !
C : , C : ,n− , C : and C : ,n− remained relatively
"' ! ") # ' #! !
#! % '
constant, but the percentages of C : and C : decreased
") "
"' "
with increasing C : ,n− supplementation (results not
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shown).
The effect of phytanic acid on the susceptibility of
C : ,n− -supplemented Y79 cells to in vitro oxidative
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stress was investigated by modulating the phytanic acid
and α-tocopherol concentrations, and measuring the
concentration of free MDA with and without exposure
to oxidative stress. Y79 cells were supplemented with
100 µM C : ,n− , with or without 0.25 µM α-tocopherol
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and 0–100 µM phytanic acid, for 5 days in the presence or
absence of oxidative stress (H O \CuSO ). The results
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are shown in Figure 5. In the cells that were not exposed
to oxidative stress, increasing the phytanic acid concentration, with or without the addition of α-tocopherol,
had little effect on free MDA concentrations. The free
MDA produced after oxidative stress both in the absence
and in the presence of α-tocopherol decreased with
increasing concentrations of phytanic acid supplementation, reaching significance at 50 µM phytanic acid.
Supplementation with α-tocopherol lowered the free
The aim of this study was to develop an in vitro model of
ARD in retinal cells, in order to investigate the molecular
distortion and anti-metabolite hypotheses for the pathogenesis of ARD. Methods were established for the
modulation of fatty acid and α-tocopherol profiles in two
mammalian cell lines of retinal origin (i.e. rat RPE and
Y79 retinoblastoma cells). The results demonstrated that
the two cell lines were able to reproducibly take up
phytanic acid, linolenic acid and α-tocopherol in a doseand time-dependent manner. The cells were typically
incubated with up to 200 µM phytanic acid. This is higher
than the plasma phytanic acid concentrations observed in
control individuals ( 16 µM), but not as high as concentrations in untreated ARD patients, which may reach
3200 µM. ARD patients treated by dietary restriction
may have plasma phytanic acid concentrations of 300–
1000 µM [7,8]. The concentration of phytanic acid
incorporated into the total cellular lipids of rat RPE cells
reached 32.8 % of the total fatty acid concentration after
supplementation with 200 µM phytanic over 7 days. This
concentration was comparable with the values of 35.4 %
and 9.6 % reported in the retinas of two ARD patients
[5].
When the two cell lines were supplemented with
100 µM phytanic acid, approx. 50 % of the phytanic acid
was incorporated into triacylglycerol ; this was most
probably stored within the neutral lipid droplets that
accumulated in the cytoplasm. Accumulation of triacylglycerol by cells supplemented with fatty acids in the
culture medium has also been reported by others [25],
and appears to be a normal response to this form of
treatment. This response may be relevant to the in vivo
situation, as fat deposits have been found in the RPE of a
patient with ARD [26] as well as in other neural tissues
[8]. The next largest proportion of phytanic acid (approx.
20 %) was found in the phospholipid fraction, indicating
that a substantial proportion of the phytanic acid was
incorporated into cellular membranes. The increase in
phytanic acid uptake into the phospholipid fraction of
both cell types was accompanied by decreases in the
percentages of virtually all the other fatty acids, with the
exception of some of the saturated fatty acids. The
increases in these saturated fatty acids may represent an
active response by the supplemented cells to compensate
for the increase in phytanic acid, which, as shown by our
results, increases membrane fluidity. It is thought that
one way in which cells regulate membrane fluidity is by
altering the unsaturation status of phospholipid acyl
chains through a process known as homeoviscous
# 2001 The Biochemical Society and the Medical Research Society
703
704
S. P. Young, A. W. Johnson and D. P. R. Muller
adaptation [25]. The effects of phytanic acid incorporation
into cellular membranes on the physical and functional
properties of the membranes will depend on whether it is
present as the free acid or bound covalently to phospholipids. However, less than 2 % of the phytanic acid in a
crude membrane lipid extract of RPE cells supplemented
with 100 µM phytanic acid was incorporated into the
non-esterified fatty acid fraction (results not shown).
Supplementation of RPE cells with 150 µM phytanic
acid was shown to decrease the steady-state anisotropy of
DPH in crude membrane suspensions, indicating an
increase in membrane fluidity. The methyl groups along
the phytanyl chain would be expected to decrease
hydrophobic interactions and increase the distance between neighbouring fatty acyl groups, thus reducing the
packing order of phospholipids. It has been reported
previously that incorporation of phytanic acid into the
phospholipids of a strain of the fungus Neurospora crassa
was associated with a lowering of the phase transition
temperature of its membranes [27]. The results of other
studies investigating the effects of free phytanic acid on
the physical properties of liposomes [28,29] and liver
mitochondrial membranes [30] have provided evidence
that free phytanic acid also decreases phospholipid
interactions, but to an extent similar to unbranched nonesterified fatty acids [30]. The results from our studies on
the fate of phytanic acid taken up by cell lines suggest that
the increase in membrane fluidity results from an
alteration of the profile of the fatty acyl chains incorporated into the phospholipid fraction of the membrane lipids, rather than from the direct incorporation of
the free unesterified phytanic acid. Various studies have
investigated the effects of altering the lipid composition
of membranes on membrane physical parameters and the
activity of membrane proteins [25,31]. Correlations
between the physical state of membranes and the activity
of membrane enzymes, receptors and transport processes
have been shown. For instance, Brown [32] provided
evidence that the transition of metarhodopsin I into
metarhodopsin II was influenced by the overall physical
properties of the bilayer. Thus the decrease in anisotropy
of phytanic acid-supplemented membranes, shown in the
present study, may well have significance for the function
of certain membrane proteins, and is compatible with the
molecular distortion hypothesis.
The anti-metabolite hypothesis was investigated by
studying the effects of exposure to 100–200 µM phytanic
acid on (a) concentrations of α-tocopherol in crude
membranes of rat RPE cells and (b) the susceptibility of
phytanic acid-supplemented Y79 cells to oxidative stress
in vitro. These high concentrations of phytanic acid did
not significantly influence the concentration of αtocopherol in crude membranes of rat RPE cells or its
displacement from these membranes. An unexpected
finding was the significantly enhanced uptake of phytanic
acid into the phospholipid fraction of RPE cells when
# 2001 The Biochemical Society and the Medical Research Society
co-supplemented with α-tocopherol. α-Tocopherol is
thought to influence the physical properties of membranes, not only via its capacity as an antioxidant,
protecting unsaturated acyl chains, but also by increasing
the order of phospholipids in the bilayer. Our results
suggest that α-tocopherol stabilizes membranes containing phospholipids with phytanoyl side chains, which
we have shown to have a disruptive effect on membrane
order. A modulatory effect of α-tocopherol on the
phospholipid acyl composition of cultured cells has been
shown previously by Giasuddin and Diplock [33].
Supplementation of Y79 cells with linolenic acid
(C : ,n− ) increased the percentage of nk3 PUFA to a
") $ $
similar extent to that reported by Hyman and Spector
[24], and provided an increase in the potential substrate
for lipid peroxidation. C : ,n− was the major nk3
## ' $
PUFA in the Y79 cell fatty acid profile up to a linolenic
acid concentration of 75 µM. At higher C : ,n−
") $ $
concentrations there was a rapid decrease in the percentage of C : ,n− , which suggested that, at these
## ' $
concentrations, there was inhibition of its synthesis from
C : ,n− . Using Y79 cells with increased concentrations
") $ $
of nk3 PUFA, there was no evidence that supplementation with phytanic acid interfered with the antioxidant
properties of α-tocopherol in these membranes. There
was, however, a consistent and significant decrease in the
concentration of free MDA with increasing phytanic acid
concentrations in the cells exposed to oxidative stress
with or without the addition of α-tocopherol. This most
probably resulted from reduced incorporation of nk3
PUFA into Y79 cell membranes with increasing concentrations of phytanic acid and, therefore, to a decrease in
the substrate available for lipid peroxidation. In summary, no evidence was found to support the antimetabolite hypothesis for the pathogenesis of ARD.
The use of transformed cell lines in these studies had
the advantage that a sufficient number of cells could be
cultured for the necessary experiments. However, the
relative simplicity of these in vitro tissue culture models
can only approximate to the much more complex
situation occurring in vivo. Creation of a knock-out
animal model of the disease is possible, now that the gene
encoding the phytanoyl-CoA α-hydroxylase has been
identified. This approach would overcome the problems
that have previously hampered the creation of a successful
animal model [8,34], and is likely to provide insights into
the precise mechanism(s) involved in the pathogenesis of
ARD.
ACKNOWLEDGMENTS
S. P. Y. thanks the Child Health Research Appeal Trust
(CHRAT) and the Szeben Peto Foundation for providing
financial support. We are grateful to Dr J. Greenwood
(The Institute of Ophthalmology, London, U.K.) for
Phytanic acid and retinal cell membranes
kindly supplying the immortalized rat RPE cell line, and
Mr G. Lynes (Great Ormond Street Hospital NHS
Trust, London, U.K.) for assistance with the synthesis of
phytanic acid.
REFERENCES
1 ten Brink, H. J., Schor, D. S., Kok, R. M., Poll-The,
B. T., Wanders, R. J. and Jakobs, C. (1992) Phytanic acid
alpha-oxidation : accumulation of 2-hydroxyphytanic acid
and absence of 2-oxophytanic acid in plasma from
patients with peroxisomal disorders. J. Lipid Res. 33,
1449–1457
2 Pahan, K. and Singh, I. (1993) Intraorganellar localization
of CoASH-independent phytanic acid oxidation in
human liver peroxisomes. FEBS Lett. 333, 154–158
3 Jansen, G. A., Ofman, R., Ferdinandusse, S. et al. (1997)
Refsum disease is caused by mutations in the phytanoylCoA hydroxylase gene. Nat. Genet. 17, 190–193
4 Mihalik, S. J., Morrell, J. C., Kim, D., Sacksteder, K. A.,
Watkins, P. A. and Gould, S. J. (1997) Identification of
PAHX, a Refsum disease gene. Nat. Genet. 17, 185–189
5 Levy, I. S. (1970) Refsum’s syndrome. Trans.
Ophthalmol. Soc. U.K. 90, 181–186
6 Steinberg, D. (1993) Refsum disease. In The Molecular
and Genetic Basis of Neurological Disease (Rosenberg,
R., Prusiner, S., DiMauro, S., Barchi, R, and Kunkel, L.,
eds), pp. 389–398, Butterworths, London
7 Steinberg, D. (1995) Refsum Disease. In The Metabolic
and Molecular Bases of Inherited Disease (Scriver, C.,
Beaudet, A., Sly, W. and Valle, D., eds), pp. 2351–2369,
McGraw-Hill Inc., London
8 Refsum, S., Stokke, O., Eldjarn, L. and Fardeau, M.
(1984) Heredopathica atactica polyneuritiformis (Refsum
disease). In Peripheral Neuropathy (Dyck, P., Thomas, P.
and Lambert, E., eds), pp. 1680–1703, Saunders,
Philadelphia
9 Muller, D. P. R., Lloyd, J. K. and Wolff, O. H. (1983)
Vitamin E and neurological function Lancet i, 225–228
10 Burton, G., Joyce, A. and Ingold, K. U. (1983) Is vitamin
E the only lipid-soluble, chain-breaking antioxidant in
human blood plasma and erythrocyte membranes ? Arch.
Biochem. Biophys. 221, 281–290
11 Greenwood, J., Pryce, G., Devine, L. et al. (1996) SV40
large T immortalised cell lines of the rat blood-brain and
blood-retinal barriers retain their phenotypic and
immunological characteristics. J. Neuroimmunol. 71,
51–63
12 Reid, T. W., Albert, D. M., Rabson, A. S. et al. (1974)
Characteristics of an established cell line of
retinoblastoma. J. Natl. Cancer Inst. 53, 347–360
13 Calder, P. C., Bond, J. A., Harvey, D. J., Gordon, S. and
Newsholme, E. A. (1990) Uptake and incorporation of
saturated and unsaturated fatty acids into macrophage
lipids and their effect upon macrophage adhesion and
phagocytosis. Biochem. J. 269, 807–814
14 Tran, K. and Chan, A. C. (1992) Comparative uptake of
alpha- and gamma-tocopherol by human endothelial cells.
Lipids 27, 38–41
15 Folch, J., Lees, M. and Sloane, S. G. M. (1957) A simple
method for the isolation and purification of total lipids
from animal tissues. J. Biol. Chem. 226, 497–507
16 Jellum, E., Eldjarn, L. and Try, K. (1966) Conversion of
phytol into dihydrophytol and phytanic acid. Acta
Chem. Scand. 20, 2535–2538
17 Lepage, G. and Roy, C. C. (1986) Direct
transesterification of all classes of lipids in a one-step
reaction. J. Lipid Res. 27, 114–120
18 Liebich, H. M., Wirth, C. and Jakober, B. (1991) Analysis
of polyunsaturated fatty acids in blood serum after fish
oil administration. J. Chromatogr. 572, 1–9
19 Metcalfe, T., Bowen, D. M. and Muller, D. P. R. (1989)
Vitamin E concentrations in human brain of patients with
Alzheimer’s disease, fetuses with Down’s syndrome,
centenarians and controls. Neurochem. Res. 14,
1209–1212
20 Burton, G. W., Webb, A. and Ingold, K. U. (1985) A
mild, rapid, and efficient method of lipid extraction for
use in determining vitamin E\lipid ratios. Lipids 20,
29–39
21 Smith, P. K., Krohn, R. I., Hermanson, G. T. et al. (1985)
Measurement of protein using bicinchoninic acid. Anal.
Biochem. 150, 76–85
21a Smith, P. K., Krohn, R. I., Hermanson, G. T. et al. (1987)
Erratum. Anal. Biochem. 163, 279
22 MacEvilly, C. J. and Muller, D. P. R. (1996) Lipid
peroxidation in neural tissues and fractions from vitamin
E-deficient rats. Free Radical Biol. Med. 20, 639–648
23 Esterbauer, H., Lang, J. J., Zaddravec, S. and Slater, T. F.
(1984) Detection of malonaldehyde by high-performance
liquid chromatography. Methods Enzymol. 105, 319–328
24 Hyman, B. T. and Spector, A. A. (1981) Accumulation of
N-3 polyunsaturated fatty acids in cultured human Y79
retinoblastoma cells. J. Neurochem. 37, 60–69
25 Spector, A. A. and Yorek, M. A. (1985) Membrane lipid
composition and cellular function. J. Lipid Res. 26,
1015–1035
26 Toussaint, D. and Danis, P. (1971) An ocular pathologic
study of Refsum’s syndrome. Am. J. Ophthalmol. 72,
342–347
27 Friedman, K. J. and Glick, D. (1982) Role of lipids in the
Neurospora crassa membrane : IV. Biochemical and
electrophysiological changes caused by growth on
phytanic acid. J. Membr. Biol. 64, 1–9
28 Yue, J., Thewalt, J. L. and Cushley, R. J. (1988)
Deuterium nuclear magnetic resonance study of the
interaction of branched chain compounds (phytanic acid,
phytol) with a phospholipid model membrane. Chem.
Phys. Lipids 49, 205–213
29 Fukuzawa, K., Chida, H. and Suzuki, A. (1980)
Fluorescence depolarization studies of phase transition
and fluidity in lecithin liposomes containing alphatocopherol. J. Nutr. Sci. Vitaminol. Tokyo 26, 427–434
30 Schonfeld, P. and Struy, H. (1999) Refsum disease
diagnostic marker phytanic acid alters the physical state
of membrane proteins of liver mitochondria. FEBS Lett.
457, 179–183
31 Viret, J., Daveloose, D. and Letterrier, F. (1990)
Modulation of the activity of functional membrane
proteins by the lipid bilayer fluidity. In Membrane
Transport and Information Storage (Aloia, R. C., Curtain,
C. C. and Gordon, L. M., eds.), pp. 239–253, Alan R. Liss
Inc., New York
32 Brown, M. F. (1994) Modulation of rhodopsin function
by properties of the membrane bilayer. Chem. Phys.
Lipids 73, 159–180
33 Giasuddin, A. S. and Diplock, A. T. (1981) The influence
of vitamin E on membrane lipids of mouse fibroblasts in
culture. Arch. Biochem. Biophys. 210, 348–362
34 Steinberg, D., Avigan, J., Mize, C. E. et al. (1966) Effects
of dietary phytol and phytanic acid in animals. J. Lipid
Res. 7, 684–691
Received 7 March 2001/25 July 2001; accepted 28 August 2001
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