(Received for publication, July 17, 1995; and in revised form, September 5, 1995)
From the
The secondary structure of a 20-amino acid length synthetic peptide corresponding to the N terminus of the second subunit of hemagglutinin (HA2) of influenza virus A/PR8/34 and its interaction with phospholipid bilayers are investigated using ESR, Fourier transform infrared (FTIR), and CD spectroscopy. N-terminal spin labeling of the peptide did not affect the secondary structure of the peptide either in solution or when bound to liposomes as revealed by FTIR and CD spectroscopy. ESR spectra show that the mobility of the labeled peptide is dramatically restricted in the presence of phosphatidylcholine liposomes, suggesting a strong binding to the lipid membranes. The N terminus of the peptide penetrates into the membrane and is located within the hydrophobic core. We find an oblique insertion of the peptide into the lipid bilayer with an angle of about 45° between helix axis and membrane plane using FTIR spectroscopy. No gross changes of the peptide's orientation, motion, and secondary structure were observed between pH 7.4 and pH 5.0. A model of the insertion of the fusion sequence of HA2 into a lipid bilayer is presented taking into account recent investigations on the low pH conformation of HA2 (Bullough, P. A., Hughson, F. M., Skehel, J. J., and Wiley, D. C.(1994) Nature 371, 37-43).
Fusion of influenza viruses with target membranes is mediated by
the major membrane protein of influenza, hemagglutinin (HA). ()HA consists of two disulfide-linked subunits, HA1 and HA2,
which are generated from a precursor form, HA0, by post-translational
cleavage. In the viral membrane, HA is organized as a homotrimer. The
first 20 residues of the N terminus of HA2 are highly conserved within
the influenza virus family (White, 1990) and appear to be crucial for
virus-cell fusion as shown by site-directed mutagenesis (Gething et
al., 1986; Schoch and Blumenthal, 1993). X-ray studies of the
crystallized ectodomain of HA have shown that at neutral pH this
terminus, the so-called ``fusion sequence,'' is buried within
the stem of the trimer 100 Å away from the top of the HA spike
(Wilson et al., 1981; Wiley and Skehel, 1987). The induction
of fusion upon lowering the pH has been ascribed to a structural change
of HA converting it into a fusogenic conformation. Although the low pH
structure of HA is unknown, one consequence of this rearrangement is
the exposure of the fusion sequence (White and Wilson, 1987). However,
very recently, a fragment of the HA2 subunit lacking the fusion peptide
has been crystallized in its low pH conformation (Bullough et
al., 1994). The x-ray crystal structure of this fragment suggests
that the fusion sequence can be delivered at least 100 Å away
from the viral membrane surface enabling its interaction with the
target membrane. This reorientation of the HA2 fragment at low pH is in
line with previous observations on the pH-dependent conformation of a
synthetic peptide sequence comprising 56 residues of this fragment
(Carr and Kim, 1993). The hydrophobic interaction of the HA ectodomain
and its fusion sequence with lipid membranes at low pH have been
demonstrated by hydrophobic photolabeling (Harter et al.,
1988, 1989; Brunner 1989). These studies suggested that the N terminus
interacts with membranes and subsequently adopts an
-helical
structure (Lear and DeGrado, 1987; Wharton et al., 1988;
Rafalski et al., 1991).
In order to elucidate the
particular role of the HA2 N terminus in virus fusion, several studies
have been undertaken to investigate the interaction of synthetic
peptides, corresponding to the fusion sequence, with lipid membranes.
Several features of the peptides which are important for their bilayer
destabilizing and, eventually, fusogenic activity have emerged: (a) the critical pressure of insertion into the lipid phase
(Rafalski et al., 1991), (b) to adopt an -helix
upon binding to the lipid phase (Lear and DeGrado, 1987; Wharton et
al., 1988; Rafalski et al., 1991; Takahashi, 1990), and (c) an asymmetric distribution of hydrophobic and hydrophilic
residues which is also common for fusogenic sequences of proteins of
other viruses (Brasseur et al., 1990). Indeed, the fusion
sequence of the HA2 subunit is characterized by the regular abundance
of glycine and hydrophobic residues (Korte et al., 1992).
Provided the peptide adopts an
-helical conformation, all glycines
are located on one side of the helix, whereas the bulky hydrophobic
residues occupy the opposite side of the helix with respect to the
helix axis (see the Edmundson wheel diagram in Fig. 1A).
Figure 1: Schematic structure of the influenza fusion peptide and the spin label. A, schematic wheel diagram of the 20-residue fusion peptide illustrating the amphipathic structure of the helix. The hydrophobic residues (highlighted) are located on the lower left of the wheel opposite the glycine-rich region (hatched). The N and C terminus are indicated by N and C, respectively. B, spin label 2,2,5,5-tetramethyl-3-pyrroline-1-oxyl-3-carboxylate attached to the peptide.
Despite numerous efforts, it is still unclear how the fusion peptide corresponding to the HA2 N-terminal segment destabilizes lipid membranes. To elucidate this mechanism, information is required not only on the structure of the peptide bound to a membrane, but also on its orientation within the membrane. Very recently it has been suggested from FTIR measurements on the fusion sequence of the transmembrane protein gp32 of simian immunodeficiency virus, that a prerequisite of the fusogenicity of fusion sequences is their oblique insertion into the membrane increasing the negative curvature of the bilayer (Martin et al., 1994; Epand et al., 1994). However, the orientation of the fusion segment of the HA2 N terminus within the membrane is unknown. Ishiguro et al. (1993) found for water-soluble peptides, which share typical features of the sequence of the influenza fusion peptide, a rather parallel orientation of the peptide with respect to the membrane plane of lipid multibilayers by using attenuated total reflection infrared (ATR-FTIR) spectroscopy. Recently, we have studied the tryptophan fluorescence from the 20-residue synthetic peptide corresponding to the HA2 N terminus of the ``B'' strain of influenza virus by stationary and time-resolved fluorescence. We calculated the distance of the tryptophan residue from the bilayer center, suggesting that the tryptophan residues are at or near the hydrocarbon-polar interface (Clague et al., 1991). No significant change in position was observed between two pH values (pH 5.0 and 7.4). However, those studies do not allow judgment of the membrane orientation of the entire peptide.
In order to gain information on the membrane orientation of
the whole fusion peptide, we examine here the interaction of a
synthetic peptide of the influenza fusion sequence with liposomes by
using both electron spin resonance (ESR) spectroscopy and ATR-FTIR
spectroscopy. The primary structure of the peptide
(Gly-Leu-Phe-Gly-Ala-Ile-Ala-Gly-Phe-Ile-Glu-GlyGly-Trp-Thr-Gly-Met-Ile-Asp-Gly)
represents the first 20 residues of the HA2 subunit of the strain
A/PR8/34. For the ESR measurements, the peptide was covalently
spin-labeled at the N terminus. Labeling did not significantly affect
the secondary structure as well as the membrane-destabilizing effect of
the peptide. We could show that the fusion sequence adopts an
-helical structure upon binding to the membrane and most
importantly penetrates into a hydrophobic environment at an oblique
angle independent of the pH value.
The N-terminal labeled fusion peptide was
dissolved in MeSO to obtain a stock solution of 1
mmol/liter. One µl of this stock solution was added to a 100-µl
vesicle suspension under rapid vortexing. All samples contained less
than 1% Me
SO to avoid any disturbance of the membrane
bilayer. The molar lipid to peptide ratio (L/P (mol/mol)) was varied by
diluting the vesicle suspension before adding the peptide to ensure a
constant spin label concentration in all samples. Depending on L/P, the
measured spectra exhibit a superposition of two components arising from
membrane-associated peptide as well as from free tumbling peptide in
buffer. To obtain the membrane spectrum, the spectrum of the labeled
peptide in buffer obtained by separate experiments was subtracted. The
intensity was calculated by double-integrating the spectra. The
relative amount of membrane-bound peptide was determined by the ratio
of the membrane signal intensity to the intensity of the highest L/P,
when all peptide is bound to the membrane. In some experiments, the
vesicle-peptide suspension was applied to a Sephadex G-50 gel
filtration column (Pharmacia Biotech Inc., 10
50 mm)
equilibrated with buffer pH 7.4 and 5.0, respectively, in order to
remove unbound peptide. Turbid, lipid-containing fractions were
collected and tested for tryptophan fluorescence. The fractions with
the highest tryptophan fluorescence were pooled and concentrated with
Centricon 30 microcentrifugation units (Amicon) prior to use in further
experiments. To characterize the insertion of the spin-labeled peptide,
the accessibility of the aqueous quencher
K
[Fe(CN)
] to the NO group was
investigated. For that purpose, aliquots of freshly prepared
K
[Fe(CN)
] stock solution were added
to the liposome-peptide suspension to a final concentration of 80
mmol/liter.
where H
is equal to the central line
width (in gauss) and h
and h
are the amplitudes of the central and the high field line,
respectively. For comparison, we have calculated the correlation time
of the spectra of the membrane-bound peptide from also,
but, due to the restricted motion of the peptide, these values
represent an apparent correlation time
.
Therefore, we have estimated
also according
to the formalism given by Freed(1976) assuming a Brownian rotational
diffusion model:
The outer hyperfine splitting A was obtained
from the membrane spectra of the peptide. The values a and b, 5.4
10
s and -1.36,
respectively, and A
= 33.6 gauss were
derived from spectral simulations for slow isotropic rotational motion,
assuming a line width parameter of 3 gauss (Freed, 1976).
The
isotropic hyperfine splitting constant a was
calculated according to Marsh (1981):
The outer (A) and inner (A
) hyperfine splittings were determined (in
gauss) from the measured spectra after smoothing the noisy signals
using the software provided by Bruker (Fourier transformation).
Figure 2:
ESR spectra of the fusion peptide in
solution. A, spectrum of the spin label used for the labeling
procedure and the labeled peptide (B) both dissolved in
MeSO. The spectra were signal-averaged over 8 consecutively
measured spectra with a modulation amplitude of 0.5 gauss and with a
scan range of 50 gauss C, spectra are of the labeled fusion
peptide C
and of a mixture of labeled and unlabeled peptide
(1:10, w/w) (D). The peptides were dissolved in
Me
SO and added to the buffer (Me
SO/sodium
acetate buffer, pH 7.4 (1:100 v/v)). The arrows indicate the
broad component. Spectra C and D were signal-averaged
over 16 consecutively measured spectra with a modulation amplitude of 4
gauss and with a scan range of 50 gauss. Note that the sharp signals
are overmodulated and therefore could not be compared directly with
spectra A and B.
To investigate
peptide-membrane interactions, appropriate aliquots of a peptide stock
solution in MeSO were added to the vesicle suspension (for
details see ``Materials and Methods''). The spectrum of the
labeled peptide becomes anisotropic in the presence of vesicles
indicating a strong interaction with the membrane. In Fig. 3,
spectra of the spin-labeled peptide in the presence of egg PC LUV at
various L/P are shown for pH 7.4 and pH 5.0 (room temperature). Nearly
all peptide is bound to liposomal membranes at L/P greater than 300.
Lowering L/P, two phenomena could be observed: (i) a sharp component
appears in the ESR spectrum which is best seen in the high field line
and (ii) the total intensity of the spectra decreases.
Figure 3: ESR spectra of spin-labeled fusion peptide bound to egg PC LUV in sodium acetate buffer, pH 7.4 (A-C, left) and pH 5.0 (D-E, right), at room temperature. The spectra were signal-averaged over 16 consecutively measured spectra with a scan range of 100 gauss and a modulation amplitude of 4 gauss. The probes contain different molar lipid to peptide ratios (L/P) as denoted in the figure. The arrows indicate the appearance of a second mobile component.
The narrow signal arises most likely from unbound peptide. Indeed, by subtracting the spectrum of the free tumbling peptide in buffer (Fig. 2C) from the original spectrum (Fig. 3), an anisotropic spectrum without any narrow lines was obtained. Therefore, we contribute this spectrum to a peptide population which does not insert into the bilayer. Excess free spin label would cause a similar narrow spectrum. Since the free spin label and the labeled peptide can be well separated by HPLC, it is very unlikely that samples contain free spin label.
The decreasing signal intensity upon lowering L/P at constant concentration of labeled peptide certainly is a consequence of peptide aggregation in buffer accompanied by spin-spin interaction as described above.
After subtracting the narrow signal from the measured spectrum, the ratio of bound to unbound peptide could be estimated by double integration of the spectra (see ``Materials and Methods''). Above L/P of approximately 300, more than 90% of the peptide is bound to the membranes. Below this L/P, the relative amount of peptide bound to vesicles decreases, indicating a saturation of binding to the bilayer. The data can be described approximately by a Langmuir binding isotherm with a limiting stoichiometry of 32 lipids/peptide for pH 7.4 at room temperature. At L/P = 60, the ratio used for FTIR measurements (see below), approximately 60% (pH 7.4) and 50% (pH 5.0) of the peptide is bound to vesicles. Only marginal differences of the bound fraction were observed upon variation of the lipid composition (egg PC or DPPC) or size of the vesicles (data not shown).
In order to separate the unbound peptide at low L/P from the vesicles, we apply the LUV suspension to a gel filtration column as described under ``Materials and Methods.'' The lipid-containing fractions were collected and tested for ESR activity. We obtain a nearly pure membrane spectrum overlapped with a negligible narrow signal contributing to less than 2% of the total measured intensity (data not shown) ensuring that almost all unbound peptide was removed by gel filtration.
To compare the restricted motion of
labeled peptide upon binding to the membrane with the free tumbling of
the peptide in solvent on a quantitative basis, we calculated the
apparent correlation time according to .
We found for egg PC LUV at both pH 7.4 and pH 5.0 nearly the same value
of 5.4 ± 0.2 ns and 5.3 ± 0.2 ns, respectively. The
rotational correlation time
`
estimated according to
the approach of Freed(1976) () was in the same order:
`
= 3.0 ± 0.2 ns and 3.0 ± 0.3 ns
for pH 7.4 and pH 5.0, respectively. Similarly, we did not observe a
significant difference of the outer hyperfine splitting A
between pH 7.4 and 5.0.
In
order to give an estimate of the insertion depth of the spin label, we
have measured a of the spin-labeled fatty acids
I(m,n) in relation to the position of the NO moiety.
For that purpose, 1 mol % of I(m,n) was incorporated
into egg PC and DPPC vesicles. The isotropic hyperfine splitting a
as a function of the immersion depth of the NO
moiety is shown in Fig. 4. The position of the NO group was
taken from our previous work (Clague et al., 1991). The
carboxyl group of the fatty acids corresponds to the depth 0 (membrane
surface). By comparing a
of the labeled peptide
with this plot, we conclude that the spin label is located near the
hydrocarbon-polar interface of the bilayer.
Figure 4:
Isotropic hyperfine splitting a of fatty acids labeled with the doxyl spin label
at different sites (see formula at the right-hand side) versus the insertion depth of the NO group. The carboxyl group
of the fatty acids corresponds to the depth 0 nm (membrane surface).
The fatty acids were incorporated in DPPC SUV
(
-
) and in egg PC SUV (
- -
-
); both vesicle types were prepared in PBS, pH 7.4. The
value of a
for the labeled peptide, bound to DPPC
SUV (solid line) and to egg PC SUV (dashed line) is
indicated; both vesicle types were prepared in sodium acetate buffer,
pH 7.4.
To sustain this
observation, we have investigated the accessibility of the hydrophilic
quencher K[Fe(CN)
] to the nitroxy
group. In the presence of 80 mmol/liter
K
[Fe(CN)
] about 20 to 30% of the
membrane spectrum is quenched, while the spin label or the labeled
peptide alone in buffer is almost completely quenched. In Table 1, the ratio of the signal intensity in the presence of
K
[Fe(CN)
] (A
) to the intensity of the membrane spectrum (A
) in the absence of the quencher is given at
pH 7.4 and pH 5.0. Intensities were obtained by double integrating the
spectra. Taking into account the error of double integration, no
substantial difference between pH 7.4 and pH 5.0 could be observed. The
portion of the quenched signal does not change, within the experimental
error, by variation of L/P in the chosen concentration range (L/P from
1350 to 135). The narrow spectrum vanishes totally as it is expected
for a signal corresponding to unbound peptide in aqueous solution (data
not shown). In conclusion, the spin label attached to the peptide is
partly accessible to the quencher. This confirms our previous results
(see above) that the label attached to the N terminus of the peptide is
located in the hydrophobic part of the lipid phase close to the
hydrocarbon-polar head group interface.
Figure 5:
ATR
spectra of the fusion peptide (solid line) dissolved in
MeSO (A) (4.4 mg/ml) and in a mixture of
trifluoroethanol/PBS, 2:1 (0.44 mg/ml) at pH 7.4 (B) and pH
5.0 (C), respectively, at room temperature. The dotted
lines represent the results of the deconvolution procedure with a
resolution enhancement factor of K = 1.86 (A)
and 1.8 (B and C), respectively (Kauppinen et
al., 1981).
To investigate the structure of the peptide bound to
the membrane, we added the peptide from a MeSO stock
solution to a suspension of egg PC LUV to a final L/P of 60 at pH 7.4
and 5.0, respectively (see the corresponding FTIR spectra in Fig. 6, A and C). The analysis of the amide I
region reveals that at both pH values the peptide exhibits mainly
-sheet and random coil structures while only approximately 20% of
the peptide is in an
-helical conformation. However, the pH
dependence of the ratio
-sheet/
-helical structures is similar
to that found for the peptide in trifluoroethanol/PBS: the contribution
of
-sheet structures rises with decreasing pH. We applied the
vesicle-peptide mixtures at pH 7.4 as well as at pH 5.0 on a gel
filtration column, as described under ``Material and
Methods.'' In Fig. 6, B and D spectra of the amide I band
at both pH values are given. Due to the elimination of the unbound
peptide, L/P increases from 60 to 150 and 95 for pH 7.4 and 5.0,
respectively. The spectra show an almost complete disappearance of
-sheet structures. The remaining peptide adopts
-helical and
random coil structures with small components of
-turns. This
proves unambiguously that the membrane-bound peptide is mostly in an
-helical and random coil conformation while the
-sheet
structure corresponds to the unbound peptide.
Figure 6:
ATR spectra at room temperature of the
fusion peptide (solid line) bound to egg PC LUV in PBS, pH 7.4 (left) and pH 5.0 (right), representing the amide I
region. The cut peak at approximately 1738 cm corresponds to the lipids
(C=O) vibration. Spectra
were recorded before (A and C, top) and
after (B and D, bottom) G-50 gel filtration
of the lipid-peptide suspension for removal of unbound peptide. The
probes were deuterated for at least 1 h before recording the spectra.
The dotted lines represent the results of the deconvolution
procedure with a resolution enhancement factor of K =
1.87 (Kauppinen et al. 1981).
To verify that the
spin label group at the N terminus does not change the secondary
structure of the peptide, we recorded IR spectra of the labeled
peptide, too. The results are shown in Table 2. The labeled
peptide adopts the same structure as the unlabeled when it is dissolved
in MeSO (70%
-sheet). Only small differences in
comparison to the unlabeled peptide were found in the presence of LUV
(see Table 2). The secondary structure of the labeled peptide
associated with the membrane was almost identical with that of the
unlabeled peptide. When we applied the vesicle peptide suspension onto
a G-50 column, the peptide fraction which adopted
-sheet
conformation was separated from the vesicles, and the remaining bound
peptide took up predominantly
-helical and random coil structures.
From the ESR spectra subsequently measured, we can ensure that almost
all remaining peptide is bound to membranes.
Figure 7:
ATR spectra of the fusion peptide inserted
into egg PC LUV (in PBS, pH 5.0) before (A, left) and
after (B, right) G-50 gel filtration, recorded using
90° and 0° polarization. The dichroism spectra obtained by
subtracting the (90°-0°) recorded spectra are plotted at the bottom of the figure, expanded 3-fold in the ordinate
direction. The hatched areas represent the amide I band,
whereas the phospholipid (CH
) at 1468 cm
and the
(CH
) at 1200 cm
are
indicated by arrows.
The orientation of the secondary structures was
determined from the 90°-0° difference spectra (Fig. 7, hatched areas). No deviation for the -helical structure
could be revealed, suggesting that the
-helix is neither parallel
nor perpendicular to a normal of the ATR element surface. The
curve-fitting applied to the polarized spectra in the amide I band
allows evaluation of the dichroic ratio (A
/A
) for the
-helical structure and calculation of the angle
between the
long axis of the
-helix and the germanium plate (Cabiaux et
al., 1989). For the helical structure associated with the
unlabeled peptide, A
/A
of 1.48 corresponds to an angle of 45° ± 5°. For
the labeled peptide we found A
/A
= 1.45
and
= 40° ± 5°. This
-helical
orientation is not significantly modified after passage of the sample
on the G-50 column, suggesting that the
-helix corresponds to a
structure that is inserted into the lipid bilayer. No alteration of
within the experimental error was found at pH 7.4 and 5.0,
respectively.
Figure 8: Circular dichroism spectra at room temperature of both labeled (dashed line) and unlabeled (solid line) fusion peptide dissolved in a mixture of trifluoroethanol/PBS (4:3, v/v), pH 7.4. The concentrations were 0.25 mg/ml and 0.17 mg/ml for the unlabeled and labeled peptide, respectively.
In the present study we show that the parallel approach of ESR and FTIR measurements allows to examine both the secondary structure of membrane active peptides and their localization and orientation within the lipid bilayer. While the FTIR method can provide information on the structure and orientation of the whole peptide in the membrane, selective spin labeling of the peptide allows us to determine the insertion depth of a given sequence as well as the dynamics of peptide motion within the membrane.
Of course, the
general disadvantage of the covalent (spin) labeling approach lies in
the influence of the reporter group on the structure and on the
membrane interaction (including the orientation) of the peptide
sequence. However, we have given several lines of evidence that
labeling does not affect the peptide in its structure and properties
relevant for membrane interaction. (a) Investigation of the
secondary structure in MeSO by using FTIR revealed no
significant differences in the relative content of
-helix,
-sheet and random coil structures between the unlabeled and
labeled peptide. This finding is strongly supported by the CD spectra
recorded in trifluoroethanol/H
O. (b) Similar to
the nonlabeled peptide, we observed a significant enhancement of
-helical structures when labeled peptide associates with lipid
membranes. (c) As shown by a lipid mixing assay, the fusogenic
activity of the peptide is not modified by covalent spin
labeling.
As expected, the secondary structure of the
peptide is sensitive to the polarity of its environment. In
MeSO, mainly
-sheet structures were found. In a
mixture of trifluoroethanol and aqueous buffer, both
- and
-structures, were present, with a higher portion of
-structure at acidic pH. A similar behavior was reported for a
peptide with a related sequence (Takahashi, 1990). The
-helical
portion of the peptide increases dramatically upon binding to the
membrane as shown previously for fusion sequences of the influenza
strains B/Lee (Lear and DeGrado, 1987) and X31 (Rafalski et
al., 1991). The membrane-bound sequence of A/PR8/34 adopts mainly
-helical and random coil structures, at similar extents. The
contribution of
-sheets was
10%. This suggests that the
-sheet is not the typical structure of membrane-bound fusion
peptide. Presumably, the
-sheet peptide is not able to penetrate
or even insert into the bilayer. This supports the model where the
peptide upon binding to the lipid phase adopts an almost helical
conformation combined with unordered structures at the ends of the
helix. The last 3 residues at the C terminus of the sequence cannot
adopt an
-helical structure in principle, since hydrogen bridges
cannot form. This is in line with recent observations that both ends of
an
-helical peptide exhibit typically more motional dynamics
(Miick et al., 1993). An homogeneous peptide population is
supported by our ESR data, since we did not find any indication for
different peptide classes.
As judged from the isotropic ESR signals in Fig. 3, at L/P > 300 nearly all the peptide present binds to the lipid bilayer. Upon lowering L/P, the bound peptide portion decreases, indicating a saturation effect of the membrane. We found a lipid to peptide stoichiometry of 32 at pH 7.4 which is similar to the value reported for the 20-residue fusion peptide of influenza B Lee (Lear and DeGrado, 1987).
The spin-labeled peptide is highly mobile
in solution as deduced from ESR spectra: equals 350
ps when the peptide is dissolved in Me
SO. In this solvent,
we found no indication for oligomerization of the peptide in even at
high concentrations. In contrast, the solubility of the peptide in
aqueous buffer is so low that only a small portion of the peptide is
monomeric while most of the peptide aggregates. The correlation time of
the monomeric peptide in Me
SO is lower than expected for a
molecule with a molecular mass of approximately 2000 Da, which is known
to be >800 ps (Miick et al., 1993). This indicates that the
spin label group undergoes additional motion relative to the global
mobility of the peptide.
The correlation time, , of
the rotational diffusion of such rather small spin labels as used here
in the membrane is about 300 ps. (
)This value is one order
of magnitude lower than the value measured for the spin-labeled peptide
bound to the membrane. Although the covalent attachment of the label to
the peptide reduces significantly its rotational motion, we can surmise
that the membrane spectrum of the labeled peptide does not reflect
solely the global motion of the peptide but, presumably, also a
``local segment'' motion within the peptide. (
)This result may also account for the absence of
significant differences between spectra recorded in the presence of egg
PC and DPPC, which is rather surprising, since at room temperature the
DPPC membrane is in the crystalline-gel state, while the membranes
consisting of egg PC are in a fluid-crystalline-like state. Most
likely, the local motions within the peptide structure are not
sensitive to the membrane fluidity.
As has been shown recently, the
depth of the NO group covalently bound to a residue within the sequence
does not necessarily reflect the average insertion depth of the peptide
backbone (Yu et al., 1994). Here, we have estimated that the
distance between the NO moiety of a spin label (bound to the peptide
terminus) and the -carbon of the first residue is in the order of
6 Å. Both the partial quenching by
K
[Fe(CN)
] and the hyperfine coupling
constant a
of the spin-labeled peptide bound to
the membrane clearly show that the spin label inserts into the bilayer
close to the hydrocarbon-polar interface independent of the pH. Thus,
the N terminus of the peptide as well is localized near the interface
or even deeper in the hydrophobic core. The concluded immersion depth
of the N terminus is in accordance with our previous investigation on
the localization of the tryptophan of the fusion sequence of influenza
B/Lee. We found that this residue is at or near the hydrocarbon-polar
interface with no gross positional change when switching the pH between
7.4 and 5.0 (Clague et al., 1991).
Since recent data have
suggested that the hydrophobicity of the N-terminal viral fusion
peptide as well as the -helix structure are not the only
parameters required for fusion to occur and that the orientation of the
peptide at the lipid-water interface plays a crucial role during the
fusion event (Horth et al., 1991; Martin et al.,
1993a, 1994), we have investigated the orientation of influenza fusion
peptide when it is inserted in the lipid bilayer.
Our polarized FTIR
results indicate that influenza peptide is obliquely inserted in the
vesicle membranes (see Fig. 7). As has been pointed out
previously (Martin et al., 1991), FTIR spectroscopy does not
discriminate between a fixed uniaxial orientation and the motional
averaging of different orientations resulting in an average oblique
orientation. However, as deduced from our ESR measurements, the strong
motional restriction of the peptide bound to the membrane makes it very
unlikely that the FTIR spectra reflect the mean of an ensemble of
different orientations. The oblique mode of insertion has been
predicted by computer analysis for the N-terminal fusogenic domains of
a series of enveloped viral proteins (Brasseur et al., 1990)
and has been experimentally demonstrated for the simian
immunodeficiency virus fusion peptide (Martin et al., 1994) as
well as for the human immunodeficiency virus fusion peptide (Martin et al., 1993b). We note that the angle between the
helix long axis and the bilayer plane, which we determine to be about
45°, differs significantly from the angle (
about 20°)
which was observed for water-soluble peptides related to this fusion
peptide (Ishiguro et al., 1993). The oblique membrane
insertion of viral fusogenic peptides seems to be an essential
requirement for membrane fusion. It is likely that this orientation
markedly alters the rather parallel alignment of the phospholipid acyl
chains. This alteration of membrane organization does not occur in the
case of amphipathic or transmembrane peptides and could prefigure a
more dramatic change in lipid order, giving rise to new lipid phases
which are thought to be associated with the initial events of membrane
fusion. Moreover, the existence of a transitional nonbilayer lipidic
structure has been demonstrated recently by
P NMR in
presence of the influenza fusion peptide (Epand et al., 1994).
In a recent investigation, a fragment of the HA2 subunit was
crystallized in its low pH conformation. The x-ray structure analysis
suggested that the N terminus of the HA2 subunit could be delivered
approximately 100 Å toward the target membrane when a coiled coil
ensemble is formed of the HA2 subunits of the HA trimer at low pH
(Bullough et al., 1994). This fragment lacks the fusion
peptide and two antiparallel -strands at its N terminus. In Fig. 9A, we present a model of the membrane topology of
the three fusion peptides of such a coiled coil structure. According to
our results, the fusion peptides are inserted at an oblique angle with
the N terminus located in the hydrophobic core. The C terminus with its
two charged residues is located in the polar head group region. The
arrangement of the three peptides shown in Fig. 9B (view along the long axis of the coiled coil) reflects the C3
symmetry of the coiled coil. The wedge shape of the peptide due to its
asymmetric distribution of small and bulky moieties (see introduction
to the text) supports a concave local bilayer curvature which may be
involved in the local alteration of the lipid phase.
Figure 9:
Model of the insertion of the fusion
peptides of a HA trimer in the coiled coil form at low pH into a
phospholipid membrane. A, view in the plane of a lipid
bilayer. The three cylinders at the top symbolize the end of
the coiled coil (for details, see Bullough et al.(1994)) The
cylinders to the left and right represent the
-helical fusion peptide inserted at an oblique angle into the
outer layer of a bilayer. The third peptide, oriented toward the
viewer, is dashed. The thick line, linking the fusion
peptide with the coiled coil, represents a protein segment of unknown
low pH structure, which consists at neutral pH of an antiparallel
-sheet (see ``Discussion''). B, view along the
coiled coil (three hatched circles in the center) illustrating
the proposed C3 symmetry of the inserted
peptides.
An important
result of our study is that we did not detect any significant influence
of the pH either on the secondary structure or on the orientation of
the peptide and the insertion depth of the N terminus in the limit of
the accuracy of our methods. Due to the presence of negatively charged
carboxyl groups in the sequence of the influenza fusion peptide one
would anticipate, for example, that acidification would favor a deeper
insertion of the peptide into the membrane. However, assuming that the
pK of 3.86 and 4.24 for isolated aspartic acid and
glutamic acid (Abraham and Leo, 1987), respectively, is not changed
dramatically within the fusion sequence, it is obvious that a
significant amount of theses residues is still in the deprotonated form
at pH 5.0. We have previously shown by a hydrophobicity plot of the N
terminus of the HA2 subunit of A/PR8/34 (including pH-dependent
protonation of carboxyl groups) that the hydrophobicity profile does
not change dramatically when shifting the pH from 7.4 to 5.0 (Korte et al., 1992). Using the scale values of Abraham and
Leo(1987), the average hydrophobicity of the first 20 residues of this
sequence changes only from 0.64 (pH 7.4) to 0.90 (pH 5.0). For
comparison, the average hydrophobicity values of the membrane-spanning
part of glycophorin and HA2 are significantly higher, 1.51 and 1.75,
respectively (Clague et al., 1991). This suggests that, even
at pH 5.0, the fusion peptide is not able to penetrate through the very
hydrophobic core (midplane) of the bilayer. A similar conclusion can be
drawn from the average hydrophobic moment which is indicative for the
formation of peptide helices with amphipathic properties (Eisenberg,
1984). Assuming an
-helical structure, the average hydrophobic
moment of the fusion sequence is 0.76 and 0.58 at pH 7.4 and 5.0,
respectively. The value for the membrane-spanning part of HA2 is 0.19.
Thus, even the hydrophobic moment of the fusion peptide at acidic pH
suggests amphipathic properties. From this, it can be concluded that
the peptide will remain near the hydrocarbon-polar interface after
acidification, which is in agreement with our experimental results here
and those presented recently (Clague et al., 1991).
We cannot rule out that an alteration of the oblique angle of peptide orientation occurs upon lowering the pH within the range of the experimental error (±5°). A rough estimate shows that even a peptide reorientation of about 10° may account for the observed pH dependence of tryptophan fluorescence (see Clague et al.(1991)).
In summary, our approach confirms previous results
that the fusion peptide of influenza virus adopts an -helical
structure upon binding to phospholipid bilayers and extends these
studies in the respect that for the first time the orientation of the
peptide within the membrane has been elucidated. We could show that the
peptide inserts with an oblique angle into the bilayer with the N
terminus localized within the hydrophobic core of the bilayer. This
topology of the influenza fusion peptide may promote intermediate lipid
structures which may resemble a major step in merging of the virus with
the target membrane.
with D (Perrin, 1936; John and
Jähnig, 1988)
where a and b refer to the half-axes of the ellipsoid. To
approximate the dimension of the peptide, we used a =
1.5 nm and b = 0.6 nm. Taking the viscosity of the
membrane, , with 0.1 pascal, we obtain D
= 1.6
10
s
and
= 6.3
10
s. Even if we assume an
uncertainty of a factor 10, the estimated rotational relaxation time is
still one order of magnitude higher than the measured
.