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Exp. Eye Res. (1996) 62, 47–53
Lipid–Protein Interactions in Human and Bovine Lens Membranes
by Fourier Transform Raman and Infrared Spectroscopies
HIDETOSHI SATOa,b, DOUGLAS BORCHMANa*, YUKIHIRO OZAKIb, OM P. LAMBAa,
W. CRAIG BYRDWELLc, M. C. YAPPERTc C. A. PATERSONa
aDepartment of Ophthalmology and Visual Sciences, and cDepartment of Chemistry, University of
Louisville, Louisville, KY 40292, U.S.A. and bDepartment of Chemistry,
Kwansei Gakuin University 1-1-155, Uegahara, Nishinomiya 662, Japan
(Received Rochester 15 June 1995 and accepted in revised form 23 August 1995)
In other systems, proteins have been shown to alter the molecular structures of lipids in the cellmembrane bilayer. We wished to determine if proteins altered the structure of lens lipids. The structureof lipid hydrocarbon chains in urea purified human lens membrane vesicles containing intrinsic,hydrophobically bound proteins was compared to the structure of lipids in vesicles without protein.Fourier transform Raman spectroscopy was used to characterize lipid and protein structure. To study lipidinteractions with extrinsic, surface bound proteins, the lipid structure was compared in bovine lipidvesicles with and without α-crystallin bound to the surface of the membrane. Lipid structure was studiedusing Fourier transform infrared spectroscopy. No change in lipid structure was detected even atprotein}lipid weight ratios of two to one. Human lens intrinsic proteins contained a high amount ofa helical structure (60%), but did not alter hydrocarbon chain interactions.
# 1996 Academic Press LimitedKey words : lens ; spectroscopy; protein ; lipid structure.
1. Introduction
Mateu et al. (1978) have shown that proteins alter
lipid structure. Extrinsic proteins, proteins that bind
to the surface of membranes, were found to disorder
the hydrocarbon chains of lipids. Intrinsic proteins,
proteins that span the bilayer or are deeply imbedded
in the bilayer, ordered the hydrophobic layer of lipid
membranes. Changes in lipid–protein interactions in
cataractous lenses (Takemoto and Rintoul, 1983) may
affect the structure of protein bound lipids and thus
could impact upon lens membrane passive perme-
ability properties and}or cation pump activity. About
50% of the weight of lens membranes is attributable to
intrinsic proteins (Takemoto and Rintoul, 1983; Roy
et al., 1982; Broekhuyse and Kuhlmann, 1978;
Broekhuyse and Kuhlmann, 1979). These proteins
could contact as much as 40% of the lipid hydrocarbon
chains and could affect the structure of the membrane.
α-Crystallin has been shown to bind to lens membranes
(lfeani and Takemoto, 1989; Mulders et al., 1985,
1989; Liang and Li, 1992; Ifeani and Takemoto,
1990, 1991a, 1991b; Zhang and Augusteyn, 1994),
and to immobilize the lipids (Liang and Li, 1992;
Puskin and Wiese, 1982), evidence that structural
alterations could exist. We have determined that
hydrocarbon chain structural order increases in
human lenses with age (Borchman et al., 1994b) and
cataract (Borchman et al., 1993) in the cortex and
* For correspondence at : Department of Ophthalmology andVisual Sciences, 301 E Muhammad Ali Blvd., Louisville, KY 40292,U.S.A.
nucleus. Dynamic order has also been shown to
increase (Takemoto and Rintoul, 1983; Puskin and
Wiese, 1982; Liang et al., 1989). The increase in lipid
order with age has been correlated with an increase in
sphingomyelin and decrease in phosphatidylcholine
(Borchman et al., 1994a, 1994b). Our previous lipid
structural studies were conducted without protein
being present.
In this study we examined protein–lipid inter-
actions by extrinsic and intrinsic proteins in bovine
and human lens membranes, respectively.
2. Materials and Methods
All aspects of this work adhered to the recom-
mendations from the Declaration of Helsinki, Finland.
Extrinsic Protein–Lipid Interactions Using Bovine Lipids
and Proteins
Bovine lens lipid extraction. Bovine eyes were
obtained fresh from a slaughterhouse. Lenses were
removed and cortical and nuclear tissues were
separated and lipid was extracted from the cortical
tissue. A monophasic methanolic extraction
(Borchman et al., 1994) followed by a hexane–
isopropanol purification was used to extract lipid from
136 g of pooled cortical material. After the methanol
extraction (Borchman et al., 1994), the methanol was
evaporated in an atmosphere of argon in a rotary
evaporator, RE121 (Buchi, Switzerland) at 55°C. The
lipid was redissolved in hexane–isopropanol (1:1, v}v)
and centrifuged. The purpose of this purification was
0014-4835}96}01004707 $12.00}0 # 1996 Academic Press Limited
48 H. SATO ET AL
to eliminate a non-protein impurity (or impurities)
(which is not soluble in hexane–isopropanol) that
absorbs at 270 nm. The elimination of the impurity
from our extracted lipid was confirmed by the absence
of any absorbing band at 270 nm. We found that this
monophasic extraction}purification procedure was
the simplest, quickest and most efficient method for
the extraction of lens lipids and is much better than
the standard Folch procedure (Folch et al., 1957). To
be certain that all of the lipid was extracted from the
lens tissue, the pellet from the first methanol extraction
step, which may contain trace amounts of lipid, was
extracted by the standard Folch (1957) procedure and
then by diethyl ether extraction. The lipids from all of
the extractions were pooled and stored in an at-
mosphere of nitrogen at ®70°C. The amounts
obtained after the Folch (1957) extraction wash and
diethyl ether extractions were negligible which indi-
cates that all the lipid was extracted in the methanol
step. The yield of lipid was 0±4 g per 136 g of lens
tissue (wet weight), measured gravimetrically. The
$"P-nuclear magnetic resonance (NMR) spectrum, (see
Borchman et al., 1994, for methods) Fig. 1, shows
qualitatively no phospholipid compositional abnor-
malities. Because a fast pulsing sequence was used to
acquire the $"-NMR spectrum in Fig. 1, only a
qualitative estimation of phospholipid composition
can be made as the relaxation of the phospholipids was
not complete.
α-Crystallin–lipid binding protocol. Bovine lipid was
dispersed by sonication in Hepes buffer (pH 7±4, 5 m)
at 4 mg lipid ml−". α-crystallin (Sigma Chemical Co.,
St. Louis, MO, U.S.A., 12% other crystallins) was
dissolved at a concentration of 1±2 mg ml−" in Hepes
buffer (pH 7±4, 5 m). The concentration was verified
by a modified Lowry assay (Peterson, 1977) designed
for membrane protein samples. The secondary struc-
ture of the α-crystallin was determined by infrared
spectroscopy (Lamba et al., 1993). Six solutions
containing 4 mg lipid each were incubated overnight
at 36°C with increasing amounts of protein (0–1±2mg) in a total volume of 2 ml. Each sample was then
spun at 170000 g for 1 hr. The supernatant was
decanted and assayed for protein. Infrared spectra of
some of the pellets containing the highest levels of
protein were measured to assess lipid structural order.
Spectroscopic instrumentation. Infrared spectra
were acquired with a Mattson model 5000 FTIR
spectrometer equipped with a TGS detector. Approxi-
mately 250 interferograms were recorded, coadded,
and apodized with a Happ–Genzel function prior to
Fourier transform, yielding an effective resolution of
1±0 cm−". Temperature was monitored to ³0±4°C by a
copper-constantan type T thermocouple and LFE
Instruments (Chesterland, OH, U.S.A.), model 3000
controller. Temperature was maintained at 36 °Cusing a Specac Limited (Fairfield, CT, U.S.A.) variable
temperature cell (model 21500). All procedures of
spectral routines have been described in our recent
publications (Borchman et al., 1993; Lamba et al.,
1993). Signal averaging, data smoothing using the
Savitsky–Golay procedure, baseline correction, buffer
subtraction, and related spectral routines were per-
formed with the Grams}386 software (Salem, NH,
U.S.A.).
Intrinsic Protein–Lipid Interactions Using Human Lens
Membranes
Human lens tissue. Clear human lenses were
obtained within 8 hr of death through the Kentucky
Lions Eye Bank. The epithelium, cortex and nucleus
were dissected. Three regions were pooled according
to age as follows: pool I, 0 to 15 years old (n¯16);
pool II, 16 to 30 (n¯34); pool IV, 46 to 60 (n¯42);
pool V, 61 to 75 (n¯91); and pool VI, 76 and older
(n¯44).
Urea treatment of membrane. The protocol of
Broekhuyse and Kuhlman (1978) was followed to
prepare urea-insoluble membranes enriched in in-
trinsic proteins. All reagents were purchased from the
Sigma Chemical Company. Lens tissue was homo-
genized, using a Teflon douncer in 0±1 Tris
(hydroxymethyl)aminomethane (Tris–HCL) at pH¯7±4 and 5 m dithiothreitol (DDT). One microliter of
buffer per lens cortical tissue was used. The homo-
genate was spun at 10000 g for 30 min at 10°C. The
pellet was resuspended in 7 urea, 0±1 M Tris–HCl,
pH¯7±4 with 5 m DTT using the same volume as
for the initial homogenization. The homogenate was
diluted seven-fold with 0±1 Tris–HCl, pH¯7±4,
5 m DTT and spun at 80000 g for 30 min. The
supernatant was decanted and the pellet resuspended
in 0±1 Tris–HCl, pH¯7.4, 5 m DTT by homo-
genization with a Teflon douncer and spun at 80000
g for 30 min. The pellet was resuspended and spun
three more times. The final pellet was placed in a
capillary tube for the FT-Raman structural study.
FT-Raman spectroscopy of urea-treated membrane. The
FT-Raman (FTR) system was a Jeol JRS-FT 6500N
spectrometer equipped with an InGaAs detector. The
excitation wavelength at 1064 nm was provided by a
cw Nd:YAG laser (Spectron SL301), and the laser
power at the sample position was typically 500 mW.
Raman spectra were obtained with a spectral res-
olution of 4 cm−", and 500 scans were accumulated to
obtain an acceptable signal-to-noise ratio.
All numerical manipulations and data treatment
were performed with Grams}386 software (Salem, NH,
U.S.A.). Spectra were 11-point smoothed using the
Savitsky–Golay procedure.
CH#
stretching band and lipid order. Lens material
for this study came from the same pool as that used in
LENS LIPID–PROTEIN INTERACTIONS 49
F. 1. $"P-NMR spectrum of bovine lipid extracted from the lens cortical region. U¯unidentified; PEplas¯phosphatidylethanolamine plasmalogen; PE¯ phosphatidylethanolamine; PC¯ phosphatidylcholine; PS¯ phosphatidyl-serine ; SM¯ sphingomyelin ; PG¯ phosphatidylglycerol.
Raman shift (cm–1)
2900 2850 2800
γ
F. 2. FT-Raman spectra of the CH#
stretching regionof (------) dipalmitoylphosphatidylcholine solid, which iscompletely ordered, (——) lens cortical lipid completelydisordered in chloroform.
published lipid structural and compositional studies
(Borchman et al., 1994a, 1994b, 1995). In this study
the lipid hydrocarbon order of membranes with
intrinsic protein is compared to that of lipid mem-
branes without protein to determine if the membrane
proteins alter lipid structure. The Raman spectral
features used to characterize lipid order include the CH
stretching band near 2850 cm−", which is the largest
band in the Raman spectra of all lipids, both native
and synthetic. Figure 2 depicts the CH stretching
region for a typical lens lipid disordered in chloroform
and for dipalmitoylphospatidylcholine that is com-
pletely ordered. The abscissa unit is wavenumbers
and, in general terms, is proportional to the frequency
of a vibration, in this instance the C–H stretching
vibration. The ordinate shows the relative number of
molecules vibrating at a given frequency. Note that
when lipids become more disordered, the band
frequency expressed in wavenumbers increases. The
frequency changes in this band have been used in our
FTIR and FTR studies to estimate lipid hydrocarbon
chain order (Borchman et al., 1993, 1994, 1995).
In addition to a shift to higher wavenumbers when
lipid acyl chains become more disordered, the intensity
of the band at 2880 cm−" decreases. The 2880 cm−"
Raman band is a Fermi resonant band, sensitive to
intra- as well as interchain interactions (Lamba et al.,
1993). The intensity of this band decreases as intra-
and intermolecular chain disorder increases. The peak
height intensity ratio I#))!
}I#)&!
, has been used as an
order parameter to determine the relative hydrocarbon
chain ‘fluidity ’ of a membrane (Gaber and Peticolas,
1977).
3. Results
Extrinsic Protein–Lipid Interactions Using Bovine Lipids
and Proteins
α-Crystallin–lipid binding model. As a model for the
study of structural changes caused by extrinsic
proteins binding to lens lipids, α-crystallin was bound
to lens lipids. Figure 3 shows the maximum weight
ratio of α-crystallin to lipid in this study was 0±3. This
corresponds to 164 g protein per mole of lipid
(phospholipid}cholesterol, 1 :1 mole ratio). In a pre-
vious study, Ifeanyi et al. (1991) demonstrated that
phosphatidylcholine vesicles bound a maximum of
31±3 g of protein per mole of phospholipid. Bovine
lipids in this study were capable of binding five times
more α-crystallin, perhaps because of differences in
technique or the differences in the composition of the
vesicles. After centrifugation, all of the α-crystallin
bound to the vesicles leaving no α-crystallin in the
supernatant [Fig. 3(b)]. Centrifugation alone could not
bring the α-crystallin out of solution (Fig. 3). Fifteen
hours were required to complete the binding process.
50 H. SATO ET AL
0.4
–0.10.0
(B) Protein/lipid (w/w)
O.D
. (75
0 n
m)
0.3
0.2
0.1
0.0
0.1 0.2 0.3 0.4
0 0.4 0.8 1.2 0.4
(A) [Protein] (mg ml–1)
(a)
(b)
F. 3. Optical density of supernatant after centrifugationof sample ; (A) aqueous bovine α-crystallin ; (B) aqueousα-crystallin at the same concentration as (A), with 4 mgbovine lens cortical lipid.
Raman shift (cm–1)
3100 2900 28003000
2924
2852
(B)
(A)
F. 4. Fourier transform infrared spectra of (A) pellet fromcentrifugation of α-crystallin plus bovine lens cortical lipidsolution. Pellet contained 0±3 protein}lipid (w}w); (B) pelletfrom centrifugation of bovine lens cortical lipid.
To determine whether lipid structure was changed
due to protein binding, infrared spectra of the vesicle
pellets containing protein were acquired. The bands in
the CH stretching region, 3100 to 2800 cm−", are very
sensitive to lipid order and have been used to probe the
lipid order of human, and rabbit lens lipids (see
Methods for details). Hydrocarbon chain lipid order is
defined structurally in terms of the freedom of rotation
about the carbon–carbon hydrocarbon bonds. Figure
4 shows the infrared spectra of vesicle pellets with [Fig.
4(A)] and without [Fig. 4(B)] protein bound to the
CH2 stretch
(i)
(ii)
(iii)
2800 2700 2600 2500
Wavenumber (cm–1)
(i)
(ii)
(iii)1700 1600 1400 1200
Am
ide
1
Tyr
osin
e
Lipidamide
Tryptophan Random
Hel
ix Tyr
osin
e
(iii)
Sheet
Helix
1700 1650
(A)
(B)
(C)
F. 5. Fourier transform infrared spectra of : i, urea-purified membrane; ii, aqueous dispersion of lipid extractedfrom urea purified membrane; iii, spectrum i®spectrum ii.(A) CH stretching region, (B) fingerprint region, (C) amide Iband.
pellet. Careful analysis of the spectra revealed no
differences in the CH#
symmetric stretching band
frequency, which indicates there was no change in
lipid order, even at high lipid to protein ratios of 164
g α-crystallin per mole lipid (cholesterol}phospholipid,
1:1 mole ratio). α-Crystallin has no affect on the
conformation of the lipid chains.
Intrinsic Protein–Lipid Interactions Using Human Lens
Membranes
The CH#
symmetric stretching band was used to
compare the lipid order of lens lipid membranes with-
out protein, with urea-purified membranes enriched
with intrinsic proteins (see Materials and Methods).
Lipid contributes about 80% the CH stretching
band intensity in the Raman spectra of membranes,
Fig. 5(A). Fig. 5(A)i shows the spectral region
LENS LIPID–PROTEIN INTERACTIONS 51
100
2851
28490
Age (years)
Pea
k ce
nte
r of
gra
vity
(w
ave
nu
mbe
rs)
20 40 60 80
2850
Moreordered
Morefluid
F. 6. The center of gravity was measured for the CH#
symmetric stretching band and is proportional to theconformational order of the lipid hydrocarbon chains. At2854±5 cm−" the lipid is completely disordered. At 2849cm−" the lipid is completely ordered. (D) human lens corticallipid ; (E) urea-purified human lens membranes.
corresponding to the CH#stretching Raman bands for
urea purified cortical membranes in aqueous buffer.
The frequency corresponding to the center of gravity
of the CH#
symmetric stretching band, measured in
wave numbers, was shown to decrease with increasing
age for the cortical lipids (no protein) in aqueous
buffer, Fig. 6. There was no statistical difference
between lipid order measured with or without intrinsic
membrane protein present (Fig. 6). Four of the six
urea-treated membrane samples exhibited lipid orders
within the dotted 95% confidence limits (Fig. 6) and
the order of these membranes is therefore indis-
tinguishable from those measured for lipids without
protein. In terms of lipid order, completely ordered
lipids have a CH#symmetric band frequency of 2847±2
cm−". Lens lipids completely disordered in CHCl$have
a CH#symmetric band frequency of 2854 cm−". Using
these numbers to estimate the percentage lipid order,
the maximum difference between lens membranes
with and without protein was, at most, 12%. The
averaged difference was 4±0%, comparable to the
standard deviation of the percentage order estimation
for the six samples measured. Lipid order may range
from 0 to 100%, so a difference of 4% order is small
and because of the experimental deviation, this
difference can not be considered significant.
The infrared spectral region corresponding to the
amide I band is shown in Fig. 5(B). This band arises
from the protein backbone amide moiety. Protein
secondary structure can be estimated by mathematical
treatment of the amide I band (Lamba et al., 1993).
We determined the secondary structure of bovine
crystallins using infrared spectroscopy and found that
the crystallin proteins were predominantly β-sheet
and contained less than 10% α-helix (Lamba et al.,
1993). Using a similar approach, we estimated the
average secondary structure of the intrinsic membrane
proteins in our samples. The estimation of secondary
structure of membrane proteins is more complicated
than that of soluble proteins because the lipid amide
band from sphingolipids interferes with the protein
amide I band. By curve fitting the amide I band of the
spectra with lipid and protein, we estimate that
41%³14 (.., n¯6) of the membrane amide I band
intensity is due to sphingolipids. Comparison of the
spectrum in Fig. 5(B)i with that in Fig. 5(B)ii, one may
see that the spectrum in Fig. 5(B)i contains a
considerable number of protein bands which indicates
a large mass of the sample is presumably due to
membrane intrinsic protein. To remove the con-
tribution of the lipid from the spectrum of lipid and
protein, Fig. 5(B)i, the spectrum of pure lipid, Fig.
5(B)ii was subtracted. The results of this subtraction
are shown in Fig. 5(B)iii.
By curvefitting the amide band as illustrated in Fig.
5(C), we estimate that the proteins are predominately
α-helical (72³15%), 18³11% β-sheet, 6³7% turns
and 4³4% other structure. Bands near 1680, 1655,
1634 and 1601 cm−" were assigned to turn, α-helix,
β-sheet and amino acid residues, respectively. The
standard deviation of the mean was calculated from
six different samples. No age-related correlations were
observed. There was no difference in the secondary
structure determined by analysis of the original amide
I band or of the Fourier self deconvolved amide bands.
The intensity at 1274 cm−" in the amide III region
[Fig. 5(B)iii] confirms the large α-helical content as
calculated from the amide I band (Warren et al.,
1976). The small amount of β-sheet and large α-
helical content indicate that the proteins in the
membrane preparation must be mostly intrinsic, since
all of the crystallins are predominately β-sheet (Lamba
et al., 1993). The 18% β sheet component may come
from crystallins shown to be present in urea mem-
branes (Russel et al., 1981). Since extrinsic proteins do
not influence lipid structure, they do not interfere with
our assessment of the effect of intrinsic proteins on
lipid structure.
4. Discussion
Weight ratios of about 1 protein to 1 phospholipid
have been reported for membrane preparations such
as the one used in this study (Takemoto and Rintoul,
1983; Roy et al., 1982; Broekhuyse and Kuhlmann,
1978, 1979). Assuming that a 100000 molecular
weight protein may contact 30 annular (the lipids in
contact with the protein) lipids (Warren et al., 1976)
(the main intrinsic protein 26 contacts eight lipids) we
calculate that 25% of the lens membrane lipids would
be in contact with the intrinsic protein. From a static
structural perspective, lipid hydrocarbon order was
not perturbed by either extrinsic or intrinsic proteins.
52 H. SATO ET AL
Even at a high 1:1 weight ratio of extrinsic protein to
lipid, no statistical differences in lipid hydrocarbon
chain order were detected. Our results are unlike those
reported by Mateau et al. (1978) which show that
extrinsic proteins disorder membranes and intrinsic
proteins order membranes. We define lipid order in a
structural sense and should not be confused with
other definitions of lipid order based on lipid mobility
or wobble. Liang et al. (1989) have shown that lens
membrane proteins do immobilize lens lipids. This has
been shown for many proteins (Devaux and
Seigneuret, 1983).
We have shown that lipid order increases with age
and cataract. These studies were performed on lipid
vesicles without protein present. It was important to
determine if proteins influence lipid order and if the
changes in lipid order reported with age and cataract
were to compensate for structural perturbations
caused by increases in membrane associated protein.
It is well known that the association of proteins with
the membrane insoluble fraction increases with
increasing age and cataract. We learn from this study
that protein, at concentrations found in human lenses,
has no influence on lipid hydrocarbon chain structure.
The increase in lipid structural order with age may be
associated with phosphatidylcholine and sphingo-
myelin compositional changes (Borchman et al.,
1994a, 1994b). The structure of lipid hydrocarbon
chains in lens lipid vesicles containing no protein is
likely to be similar to the structure in vivo where
proteins are present.
Acknowledgements
Supported by Public Health Service research grantsEY07975 and EY06916 (Bethesda, MD, U.S.A.) and theKentucky Lions Eye Foundation (Louisville, KY, U.S.A.), andan unrestricted grant from Research to Prevent Blindness,Inc. Christopher A. Paterson, Ph.D., D.Sc. is a Research toPrevent Blindness Senior Scientific Investigator.
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