From the
Department of Molecular Biology and
Genetics and Biophysics Interdepartmental Group, University of Guelph, Guelph,
Ontario N1G 2W1, Canada, ¶Department of
Structural Biology and Biochemistry, Hospital for Sick Children, Toronto,
Ontario M5G 1X8, Canada and Department of Laboratory Medicine and
Pathobiology, University of Toronto, Toronto, Ontario M5G 1L5, Canada, and
||Biophysics Research Institute, Medical College of
Wisconsin, Milwaukee, Wisconsin 53226
Received for publication, March 18, 2003 , and in revised form, May 1, 2003.
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ABSTRACT |
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INTRODUCTION |
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We have previously produced and characterized a recombinant murine 18.5-kDa
MBP (11). Here, we will denote
this protein quasi-C1 (qC1), because it is unmodified post-translationally
(with the exception of an LEH6 tag) and emulates the
least-modified, most basic charge isomer C1. We have also generated by
site-directed mutagenesis a quasi-deiminated form of recombinant murine
18.5-kDa MBP that we call qC8, since it was designed to mimic the less
cationic natural form C8. The recombinant qC8 consists of Arg/Lys Gln
substitutions at the same deimination sites in human MBP that predominate in
chronic multiple sclerosis and has properties similar to those of natural C8
(7). The net charge of qC1 is
+19 at neutral pH, whereas that of qC8 is +13 as for their natural
counterparts.
In this study, we investigated the electrostatic and hydrophobic components
of MBP-lipid interactions by site-directed spin labeling (SDSL) of MBP and
electron paramagnetic resonance (EPR) spectroscopy
(12,
13). The technique of SDSL
involves replacement of residues at selected sites by cysteines, which are
then labeled with a methanethiosulfonate spin label that can be probed by EPR
spectroscopy. This approach enabled us to monitor the electrostatic lipid
interaction profiles of qC1 and qC8 at numerous specific sites. The importance
of hydrophobic interactions was evaluated by spin-labeling sites adjacent to
each of the two Phe-Phe pairs and determining the effects of Phe-Phe
Ala-Ala substitution on spin label mobility and accessibility to lipid-soluble
O2 and water-soluble nickel ethylenediaminediacetic acid (NiEDDA)
as applied by Victor et al.
(14) to the myristoylated
alanine-rich C kinase substrate effector region. The technique of SDSL is
particularly well suited to MBP, because there are no native cysteinyl
residues to be removed prior to mutagenesis and because the EPR spectrum is
not affected by light diffraction and immobilization associated with
MBP-induced lipid vesicle aggregation. Lipid vesicle aggregation is a powerful
mimic of the in vivo function of MBP in the myelin sheath
(15,
16), and SDSL/EPR offers the
advantage of studying residue-specific interactions within this natural
environment.
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EXPERIMENTAL PROCEDURES |
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Phosphatidylcholine (PC), phosphatidylethanolamine, phosphatidylserine (PS), phosphatidylinositol, cholesterol, and sphingomyelin were procured from Avanti Polar Lipids (Alabaster, AL). The depth calibration curve was obtained using the following spin-labeled lipids purchased from Avanti Polar Lipids: 1-palmitoyl-2-stearoyl(n-doxyl)-sn-glycero-3-phosphocholine with n = 5, 7, 10, and 12 (5-doxyl-PC, 7-doxyl-PC, 10-doxyl-PC, and 12-doxyl-PC) and 1,2-dipalmitoyl-sn-glycero-3-phosphotempocholine with TEMPO (1,2-dipalmitoyl-sn-glycero-3-phospho(TEMPO)choline bound to the quaternary ammonium group (TEMPO-PC). All of the lipids were dissolved in chloroform (100%) at concentrations of 510 mg/ml.
Site-directed Mutagenesis of qC1 and qC8 The quasi-deiminated mutant of MBP (qC8) was generated from qC1 by sequential site-directed mutations (first R25Q followed by R33Q, K119Q, R127Q, and R157Q and finally R168Q, murine sequence numbering) using the QuikChange protocol (Stratagene, La Jolla, CA) as described previously (7). A series of matching Cys substitutions in each of qC1 and qC8 was generated for SDSL. In addition, in qC1, each of the two Phe-Phe pairs was separately replaced by Ala-Ala. In summary (Fig. 1a), the following mutations were generated in qC1: S17C, S44C, S67C, H85C, S99C, S129C, S159C, F42A/F43A/S44C, and F86A/F87A/H85C. The following mutations were generated in qC8: S17C, S44C, S67C, H85C, S99C, S129C, and S159C. The introduced cysteinyl residues could then be spin-labeled by MTS-SL (R1) (Fig. 1b).
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Labeling and Purification of Cysteine-containing Mutants of qC1 and qC8 The purification and spin labeling of recombinant murine 18.5-kDa MBPs were done using a modification of a published purification protocol (11). Two types of lysis buffer were used, one containing 20 mM Tris-HCl, pH 8.0, 6 M urea, 5 mM imidazole, and 500 mM NaCl, and the other being identical with the exception of the addition of 1 mM dithiothreitol to prevent the formation of intermolecular disulfide bonds. The cell pellet was lysed using the dithiothreitol-containing buffer, and subsequent loading and washing of the Ni2+-NTA column were done using this buffer. Finally, the column was washed with normal lysis buffer using 25 column volumes to remove the excess dithiothreitol. An additional 25 volumes of spin labeling buffer with urea (20 mM HEPES-NaOH, pH 7.4, 6 M urea, 10 mM NaCl) were then run through the column. The agarose beads were removed from the column and resuspended in 8 ml of spin labeling buffer, and a 10-fold molar excess of MTS-SL was dissolved in 100 µl of Me2SO was added to the bead slurry. The slurry consisting of the protein bound to the Ni2+-NTA beads and MTS-SL was incubated on a nutator overnight in a 15-ml Falcon tube at room temperature. The column was repoured the next day and washed with normal lysis buffer to remove excess MTS-SL, and the bound protein was eluted. The purity of each labeled protein was verified using SDS-PAGE, and the protein-containing fractions were dialyzed against 20 mM HEPES-NaOH, pH 7.4, 10 mM NaCl, and 1 mM EDTA. The labeling efficiency was assessed for one sample using electrospray ionization mass spectroscopy (11, 20) and was found to be complete (data not shown). The protein concentration was estimated by measuring the absorbance at 280 nm using the published extinction coefficients (7). The protein samples were adjusted to 1 mg/ml in the same buffer for EPR spectroscopy.
Preparation of Large Unilamellar Vesicles (LUVs)Aliquots of the chloroform solutions of the lipids were combined in the following molar ratios to form LUVs with a lipid composition similar to that estimated for the cytoplasmic face of the myelin membrane (Cyt-LUVs): 44 mol % cholesterol; 27 mol % phosphatidylethanolamine; 13 mol % PS; 11 mol % PC; 3 mol % sphingomyelin; and 2 mol % phosphatidylinositol (15, 21). The solvent was evaporated under a stream of nitrogen, and the lipid mixture was dried further in a vacuum dessicator overnight. It was hydrated in 20 mM HEPES-NaOH, pH 7.4, 10 mM NaCl, and 1 mM EDTA. The lipid suspension was then passed through a 100-nm polycarbonate membrane 17 times using a syringe extruder (Avanti Polar Lipids). The final lipid concentration after extrusion was verified using a phosphorous assay (22).
EPR SpectroscopyFor signal-averaged EPR spectroscopy, spin-labeled qC1 and qC8 solutions were added to LUVs at a concentration of 100 µg of protein to 2 mg of LUVs, yielding a molar protein:lipid ratio of 1:600. This value is a little less than the MBP:lipid ratio in myelin. Borosilicate glass 50-µl capillary tubes (Fisher Scientific) were used to hold the samples. Aggregation of the vesicles by MBP occurred immediately, and after 10 min, the preparations were spun at 1000 x g to loosely pellet the MBP-LUVs. Most of the supernatant was removed, and the pellets were taken up in capillary tubes. The opposite end of the capillary tube was sealed using a Bunsen burner, and the tubes were centrifuged at 4000 x g for 15 min to give a compact pellet.
The protein-lipid pellet was positioned in the center of the EPR cavity of a Bruker ECS 106 spectrometer (Bruker BioSpin, Milton, Ontario, Canada). The EPR spectra were recorded at room temperature using a microwave power of 10.0 milliwatts and a modulation amplitude of 1.0 G. To estimate the amount of free unbound MBP, the supernatant of the final centrifugation step (see above) was also probed by EPR spectroscopy and no detectable signal was recorded. We conclude that all of the MBP reacted with the lipid vesicles and that there was no unbound protein.
An empirical motion parameter 0 was determined from the
first derivatives of the absorption spectra using
Equation 1,
![]() | (Eq. 1) |
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Depth Measurements via Continuous Wave (CW) Power
Saturation The depths of penetration of spin-labeled sites of all
of the proteins into Cyt-LUVs were determined from CW power saturation.
Experiments and data analysis were performed essentially as described
previously (25,
26) using a Varian E102
Century Series spectrometer (Varian Associates, Palo Alto, CA) equipped with a
loop-gap resonator (Medical Advances, Milwaukee, WI). Because of the increased
sensitivity of this resonator, the amount of sample used for CW saturation was
halved (1 mg of LUVs, 50 µg of protein). After spinning down the
lipid-protein aggregate, 5 µl of it was loaded into a gas-permeable TPX
capillary (27). The field
modulation was 1.25 G, and the incident microwave power was varied from 0.1 to
81 milliwatts. The P value was determined by
fitting the amplitude A of the mI = 0 peak to
Equation 2,
![]() | (Eq. 2) |
![]() | (Eq. 3) |
This parameter describes the relative depth of the spin label in the bilayer due to the gradient of NiEDDA and O2 along the bilayer normal.
The dependence of on distances from the membrane surface was
determined using head group-labeled TEMPO-PC and various PCs with doxyl
nitroxides along the acyl chain at positions 5, 7, 10, and 12. The
nitroxide-labeled PCs were mixed with the lipids used for Cyt-LUVs at a molar
ratio of 1:500 in chloroform, and LUVs were prepared as described above. The
calibration was done using Cyt-LUVs without protein as well as two
concentrations of unlabeled qC1. The first concentration was a molar ratio of
lipid to protein of 600:1 (identical to that used for the depth measurements),
and the second concentration was a saturating amount of unlabeled qC1 (150:1
lipid to protein). The resultant
values were used in conjunction with
the known depth (distance of the nitroxide nitrogen from the phosphate) of
spin-labeled PC doxyl nitroxides
(29) to generate a standard
curve for determining the absolute depth of the MTS-SL labels in the protein
when bound to the bilayer. The distance of the nitrogen to the phosphate in
TEMPO-PC was estimated to be 5 Å based on molecular models by
Farahbakhsh et al.
(28).
Previous results have demonstrated that the linear dependence of the depth
parameter breaks down with increasing distance into the aqueous phase and
behaves according to Equation 4
(30),
![]() | (Eq. 4) |
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RESULTS |
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The qC8 Spin Labels Are Less Immobilized than the qC1 Spin Labels in the BilayerThe interactions with Cyt-LUVs of qC1 and qC8 spin-labeled at sites S17C, S44C, S67C, H85C, S99C, S129C, and S159C were monitored from the EPR spectra. Upon interaction of these proteins with Cyt-LUVs, the motion became more anisotropic and the sharp hyperfine lines were broadened, indicating a decrease in mobility (Figs. 2b and 3, a and b). In some cases, especially H85R1, S129R1, and S159R1 in qC1, a second more immobilized component was observed as indicated by an arrow on the low field side in Fig. 3a. There was no exchange broadening indicative of MBP oligomerization in any of the spectra.
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The spectra in Fig. 3, a and
b, showed several differences between qC1 and qC8. The
mobility of most labeled residues in qC8 was greater as can be seen especially
from the sharper high field peak at mI = 1. There
was also a decrease in the more immobilized component seen for qC1 at H85R1
and S129R1. To quantify the degree of mobility, the widths of the center-lines
(H) were measured and compared as 1/
H
(Fig. 4). Generally, a higher
value of 1/
H means that the relative mobility is greater. The
spin labels at sites S17C, S67C, S129C, and S159C in qC8 were more mobile than
at the same sites in qC1. Overall, the difference between the spin-label
environments in qC1 and qC8 was more pronounced the closer the label was to an
Arg/Lys
Gln substitution. However, H85R1 was more mobile in qC8 than in
qC1 (seen especially from the loss of immobilized component from the spectrum
in Fig. 3b, rather
than from the change in
H in
Fig. 4) even though there was
no Arg/Lys
Gln substitution in the vicinity. This result indicates that
this site might be more sensitive to its environment or that quasi-deimination
and reduced lipid binding effected a local structural perturbation.
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Role of the Phe-Phe Sites in Anchoring qC1 and qC8 in the BilayerThe spin labels at sites S44C and H85C next to two Phe-Phe pairs at positions Phe-42/Phe-43 and Phe-86/Phe-87 were the most immobilized in qC1 and qC8 (Figs. 3b and 4), suggesting that they anchor qC1 and qC8 in the bilayer. The two Phe-Phe pairs were expected to provide a hydrophobic component to the free energy of binding of MBP to lipid bilayers.
To estimate the relative importance of these Phe-Phe sites, both
Phe-42/Phe-43 and Phe-86/Phe-87 were substituted by Ala-42/Ala-43 and
Ala-86/Ala-87 in qC1-S44C and qC1-H85C, respectively. The Phe-Phe
Ala-Ala substitutions caused significant differences in the environments of
adjacent probes and hence mobilities and spectral line shapes, especially for
qC1-H85R1 (Fig. 3c,
arrows). In fact, qC1-F86A/F87A/H85R1 was the most mobile of all of
the sites probed as can be seen from the
H values in
Fig. 4. In contrast,
qC1-F42A/F43A/S44R1 was motionally restricted even with the Phe-Phe
Ala-Ala substitutions.
Depth Measurements via CW Power SaturationThe EPR spectral
line shapes (and hence values of H and
0)
could depend not only on the depth of penetration of the spin label into the
lipid bilayer but also on secondary structural changes that can be induced in
MBP by its interaction with lipids. For this reason, direct measurements of
depth of penetration of each spin label into the membrane were performed using
power saturation approaches
(25,
26).
Table I summarizes the
and
values calculated for all of the proteins when bound to Cyt-LUVs.
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Due to the unique combination of lipids present in the Cyt-LUVs, proper
calibration of the value using the spin-labeled lipids was essential.
The NiEDDA accessibility of the lipid nitroxide was significantly higher in
the cholesterol-containing Cyt-LUVs than in the PC or PC/PS lipid mixtures
usually used for these measurements
(18,
30,
32). Subczynski et
al. (32,
33) have shown that
cholesterol increases the water permeability and reduces oxygen permeability
of the bilayer up to C7 or C9 of the acyl chain of saturated and unsaturated
lipids, respectively. Consequently, NiEDDA would actually penetrate Cyt-LUVs
more easily up to a depth of 10 Å, which means that all of the nitroxide
spin labels in MBP would be more available for collisional quenching than they
would be in a bilayer lacking cholesterol. The experimentally determined depth
of the spin labels (29) was
plotted versus the
values
(Fig. 5a), and a curve
was then fit to the data using the hyperbolic tangent function described in
Equation 4. This function was
first reported by Frazier et al.
(30) and provides a
description of the limiting behavior of
that can be applied to sites on
the aqueous side of the bilayer interface.
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To determine the lipid spin label accessibility in the presence of MBP bound to the surface of the membrane, the calibration curve was also determined with the addition of 50 µg of unlabeled qC1 (as used for spin-labeled MBP) and with 180 µg of unlabeled qC1 to saturate the acidic phospholipid head groups. The presence of MBP increased the depth parameter (Fig. 5a), and the curve was fit to data for the samples containing 180 µg of qC1. Despite the differences in lipid composition from that used by Frazier et al. (30), Equation 4 resulted in an extremely good fit of the data (r2 = 0.9998) (Fig. 5a).
The distance data for the various spin-labeled MBPs were obtained by
solving for the distance using experimentally determined values
(Fig. 5b). The
calibration curve changes very little below a
value of 2, making
it impossible to solve for the relative depths of some of the more exposed
spin label sites. To calculate a distance for
values below 2, we
used linear regression to fit data for the head group PC and 5- and
7-doxyl-PCs (these values served for comparative purposes only).
The depth values for qC1 showed that with the exception of S17R1, spin-labeled residues in the N-terminal half were more deeply embedded in the bilayer than those in the C-terminal half, whereas those in the midsection were located near the level of the nitrogen of the PC head groups (Fig. 5b). The spin label S44R1 next to Phe-42/Phe-43 was the most deeply penetrating one, but H85R1 next to Phe-86/Phe-87 penetrated only a little below the nitrogen of the PC head group, even though it was the most immobilized residue.
Comparison of the depth values of qC1 and qC8 revealed dramatic differences at sites in the C terminus, which relocated to the aqueous phase in qC8. In contrast to qC8, S129R1 and S159R1 of qC1 were found at the interface and at 3.5 Å below the phosphates, respectively. The relocation of residues in the C-terminal domain of qC8 to the aqueous phase accounts for the greater mobility of these residues.
At the N terminus, S17R1 was slightly more exposed in qC1 as opposed to
qC8, despite the fact that the mobility was significantly higher in qC8.
Accessibility of S44R1 and S99R1 was similar in qC8 and qC1, consistent with
the lack of effect of Arg/Lys Gln substitution on mobility of these
residues. In addition, S99R1 was found in the aqueous region consistent with
its high mobility, whereas the location of S44R1 in the bilayer was consistent
with its low mobility.
The Arg/Lys Gln substitution in qC8 caused H85R1 to move further out
of the bilayer, consistent with an increase in mobility. However, the location
of this residue in the head group region in qC1 suggests that the low mobility
of this residue may be due to secondary structure in this region rather than
the result of deep bilayer penetration. There was a significant increase in
the aqueous accessibility of H85R1 after the F86A/F87A substitution, but no
change for S44R1 with the F42A/F43A mutation. This result was somewhat
surprising considering the depth of S44R1; however, the increase in mobility
observed at this site was not nearly as extreme as it was for H85R1 upon
substitution of Ala-Ala for Phe-Phe near these residues.
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DISCUSSION |
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Electrostatic Interactions of MBP with Lipid BilayersWe used a fully charged version of recombinant murine MBP (qC1, net charge +19 at pH 7.0) as well as a less cationic form (qC8, net charge +13 at pH 7.0) to study how the overall charge affected lipid-protein interactions at various sites. Most spin-labeled sites were found to be more mobile in qC8 than in qC1, although the accessibility measurements indicated that the C terminus in qC8 was the most accessible to the aqueous phase, indicating a potentially decreased interaction of these qC8 sites with the lipid bilayer.
Other studies have shown that MBP/C1 can bind to actin filaments while bound to Cyt-LUVs and that the protein dissociates from the membrane when calmodulin is added (43). This dissociation would be predicted to be faster with qC8. Thus, not only does the modified protein aggregate lipid to a lesser extent, but the greater mobility of spin-labeled sites and the increased accessibility at the C terminus indicate that it is less embedded in the bilayer, which would expose it to potential ligands. In summary, long range Coulombic forces appear to play an important role in attracting MBP to the surface of a membrane and in determining how deeply it is embedded in the bilayer.
The high mobility and accessibility of S99R1 in qC1 and qC8 indicated that this region was exposed to the aqueous environment. This segment of 18.5-kDa MBP is of interest because of its proximity to the PRTPPPS motif (the triproline repeat region characteristic of all of the known mammalian MBPs) (Fig. 1a). The sequence of PRTP represents a PXXP motif that can bind to an SH3 domain (Src homology 3) containing protein such as non-receptor tyrosine kinases (44). Moreover, the threonyl residue within this motif is a mitogen-activated protein kinase target (45). Our results suggest that this segment of MBP is naturally exposed and available for modification and recognition.
The Two Phe-Phe Pairs in MBP Are Not EquivalentThe
importance of aromatic residues in the two dual Phe-Phe sites was studied by
substituting these residues for Ala-Ala. The spin-labeled residues immediately
adjacent to the Phe-Phe pairs, S44R1 and H85R1 in both qC1 and qC8,
demonstrated significant motional restriction compared with other sites. After
the Phe-Phe Ala-Ala substitutions, the mobility of each of these sites
increased, especially H85R1.
When the regions surrounding Phe-42/Phe-43 and Phe-86/Phe-87 are portrayed
as helical wheels (Fig. 6),
there are distinct disparities. Segment mMBP
(3849)
has neither a hydrophobic nor a hydrophilic face indicative of an amphipathic
-helix. In contrast, segment mMBP-(8293) is strongly amphipathic
with very hydrophobic amino acids (Val, Phe, and Ile) on one side and polar
amino acids (Lys, Thr, His, and Asn) on the other side of the
-helix (a
similar prediction has been noted previously
(46,
47)). The segment
mMBP-(8293) has a hydrophobic-hydrophilic residue ratio of 13:5, which
in peptides has been shown to result in immersion of their hydrophobic regions
into lipid bilayers (48). In
fact, this type of
-helix had the largest hydrophobic face in that
study and had the strongest binding to bilayers. The mean helical hydrophobic
moment (µH) of the putative
-helix mMBP-(8293) was
calculated using the sum of the hydrophobicities of the side chains
(49) and found to be 0.332,
indicating that the
-helix is amphiphilic perpendicular to its axis.
The hydrophobic moment of this section of MBP places it directly in the domain
of known surface-seeking
-helices
(48). Moreover, this
-helix may even be stable in solution as indicated by digestion of MBP
with cathepsin D, which cleaves at Phe-Phe linkages. The Phe-42/Phe-43 pair is
cleaved far more quickly than Phe-86/Phe-87
(7,
50). The CW power saturation
accessibility of H85R1 indicates that this site is above the plane of the
lipid phosphate groups. Since H85R1 is located on the polar side of the
putative amphipathic
-helix (Fig.
6), the conclusion that it is positioned at the interface is
sound.
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The CW power saturation data positions H85R1 in the Ala-Ala-containing
mutant in the bulk aqueous phase several angstroms from the membrane location
of H85R1 in the Phe-Phe-containing protein. It might have been expected that
this site would have been even more exposed based on the considerable change
in mobility that was observed. Another factor to consider is the fact that the
spectrum of the spin-labeled residue is sensitive to -helix motion
(51,
52). The membrane-anchoring
effect of the Phe-Phe pair might prevent rocking motions of the
-helix.
Rocking motion would become much more pronounced when the Phe-Phe was
converted to Ala-Ala.
Another interesting feature of segment mMBP-(8293) is the terminal
prolyl residues at positions Pro-82 and Pro-93. Prolines are commonly found in
kink or bend points within proteins and disrupt -helices, especially in
membrane proteins (53). These
two residues could represent natural extents of the putative amphipathic
-helix, and the points at which this section of MBP would dip into the
membrane. This idea is supported by the observation that both prolyl residues
line up in an orientation where they could connect with the rest of the
protein (Fig. 6), whereas the
amphipathic
-helix penetrates the membrane.
The Phe-42/Phe-43 Ala-42/Ala-43 substitutions had less of an effect
on the mobility of S44R1. The CW power saturation indicated that the depth of
this residue was unchanged by this substitution, even though it was the most
deeply penetrating site. From this result, it does not appear that the
presence of the Phe-42/Phe-43 pair alone is vital for membrane anchoring of
this region of the protein. Thus, Phe-42/Phe-43 and Phe-86/Phe-87 may play
different roles in terms of the association of MBP with the membrane. The
reason for the Phe-Phe-independent deep penetration of S44R1 and for the deep
penetration of the N-terminal half of MBP is not known. However, it is
consistent with greater labeling of the N-terminal half of MBP by the
hydrophobic photolabel relative to the C-terminal half
(40). In myristoylated
alanine-rich C kinase substrate, a highly basic 25-residue lipid-effector
region interacts with the cell membrane by a combination of electrostatic
interactions and partial insertion of hydrophobic phenylalanines into the
lipid bilayer (14,
54). Our results show that
this highly effective design for strong membrane association is only partially
shared by MBP with significant differences in behavior (and thus role) of one
of its Phe-Phe pairs.
Another amphipathic -helix was predicted earlier for the C terminus
of MBP (55), suggesting that
this region might bind acidic lipids with high affinity and/or contain a
calmodulin-binding site. The more immobilized component in the spectrum of
S159R1 of qC1 (not present in the spectra of S17R1, S67R1, and S99R1) and its
deeper penetration at a depth of 3.5 Å below the lipid phosphates are
consistent with the localization of S159R1 on the hydrophobic side of this
putative amphipathic
-helix.
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CONCLUSIONS |
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FOOTNOTES |
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Recipient of a Multiple Sclerosis Society of Canada Studentship.
** To whom correspondence should be addressed: Dept. of Molecular Biology and Genetics, University of Guelph, 50 Stone Rd., E., Guelph, Ontario N1G 2W1, Canada. Tel.: 519-824-4120 (ext. 52535); Fax: 519-837-2075; E-mail: gharauz{at}uoguelph.ca.
1 The abbreviations used are: MBP, myelin basic protein; q, quasi; SDSL,
site-directed spin labeling; EPR, electron paramagnetic resonance; NiEDDA,
nickel ethylenediaminediacetic acid; MTS-SL,
[1-oxyl-2,2,5,5-tetramethyl-D-pyrroline-3-methyl]methanethiosulfonate;
NTA, nitrilotriacetic acid; PC, phosphatidylcholine; PS, phosphatidylserine;
LUV, large unilamellar vesicle; CW, continuous wave.
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ACKNOWLEDGMENTS |
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REFERENCES |
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