Dual-fluorescence l -amino acid reports insertion and orientation of melittin peptide in cell membranes

Article abstract of DOI:10.1021/bc400325n

Monitoring insertion and orientation of peptides in situ on cell membranes remains a challenge. To this end, we synthesized an l-amino acid (AFaa) containing a dual-fluorescence dye of the 3-hydroxyflavone family, as a side chain. In contrast to other labeling approaches using a flexible linker, the AFaa fluorophore, introduced by solid phase synthesis into desired position of a peptide, is attached closely to its backbone with well-defined orientation, and, therefore, could reflect its localization in the membrane. This concept was validated by replacing the leucine-9 (L9) and tryptophan-19 (W19) residues by AFaa in melittin, a well-studied membrane-active peptide. Due to high sensitivity of AFaa dual emission to the environment polarity, we detected a much deeper insertion of L9 peptide position into the bilayer, compared to the W19 position. Moreover, using fluorescence microscopy with a polarized light excitation, we found different orientation of AFaa at L9 and W19 positions of melittin in the bilayers of giant vesicles and cellular membranes. These results suggested that in the natural membranes, similarly to the model lipid bilayers, melittin is preferentially oriented parallel to the membrane surface. The developed amino acid and the proposed methodology will be of interest to study other membrane peptides.

Full text of DOI:10.1021/bc400325n

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Article
Dual-Fluorescence L-Amino Acid Reports Insertion
and Orientation of Melittin Peptide in Cell Membranes
Viktoriia Y Postupalenko, Oleksandr M Zamotaiev, Volodymyr V Shvadchak,
Aleksandr V Strizhak, Vasyl G Pivovarenko, Andrey S Klymchenko, and Yves Mély
Bioconjugate Chem., Just Accepted Manuscript • Publication Date (Web): 24 Nov 2013
Downloaded from http://pubs.acs.org on November 28, 2013
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Dual-Fluorescence L-Amino Acid Reports Insertion and
Orientation of Melittin Peptide in Cell Membranes
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Viktoriia Y. Postupalenko,a,‡ Oleksandr M. Zamotaiev,b,‡ Volodymyr V. Shvadchak, a,c Aleksandr V.
Strizhak,b Vasyl G. Pivovarenko, b Andrey S. Klymchenko,*,a Yves Melya
aLaboratoire de Biophotonique et Pharmacologie, UMR 7213 CNRS, Université de Strasbourg, Faculté
de Pharmacie, 74, Route du Rhin, 67401 Illkirch Cedex, France.
bDepartment of Chemistry, Kyiv National Taras Shevchenko University, 01033 Kyiv, Ukraine
cLaboratory of Cellular Dynamics, Max Planck Institute for Biophysical Chemistry, am Fassberg 11,
37077 Göttingen, Germany
KEYWORDS: Fluorescence, amino acid analogue, biomembrane, microscopy, melittin, peptide
orientation, cell membranes
* Address correspondence to andrey.klymchenko@unistra.fr; fax: (+33) 368854313.
Author Contributions
The manuscript was written through contributions of all authors. All authors have given approval to the
final version of the manuscript. ‡These authors contributed equally.
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ABSTRACT
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Monitoring insertion and orientation of peptides in situ on cell membranes remains a challenge. To
this end, we synthesized an L-amino acid (AFaa) containing a dual-fluorescence dye of the 3-
hydroxyflavone family, as a side chain. In contrast to other labeling approaches using a flexible linker,
the AFaa fluorophore, introduced by solid phase synthesis into desired position of a peptide, is attached
closely to its backbone with well-defined orientation, and, therefore, could reflect its localization in the
membrane. This concept was validated by replacing the leucine-9 (L9) and tryptophan-19 (W19)
residues by AFaa in melittin, a well-studied membrane-active peptide. Due to high sensitivity of AFaa
dual emission to the environment polarity, we detected a much deeper insertion of L9 peptide position
into the bilayer, compared to the W19 position. Moreover, using fluorescence microscopy with a
polarized light excitation, we found different orientation of AFaa at L9- and W19 position of melittin in
the bilayers of giant vesicles and cellular membranes. These results suggested that in the natural
membranes, similarly to the model lipid bilayers, melittin is preferentially oriented parallel to the
membrane surface. The developed amino acid and the proposed methodology will be of interest to
study other membrane peptides.
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Introduction
Membrane binding peptides and small proteins are involved in numerous important cellular
processes including membrane fusion, disruption or channel formation. Determination of peptide
insertion depth and orientation in membranes is important for understanding the mechanisms and
developing peptide-based drugs, transfecting reagents, etc.1
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Numerous techniques were developed in vitro to characterize peptide - lipid membrane interactions,
including isothermal titration calorimetry (thermodynamics),2 nuclear magnetic resonance (peptide
structure),3, 4 circular dichroism (conformation),5 electron spin resonance (immersion)6 and
fluorescence (affinity, localization, etc).7 For the last technique, environment-sensitive fluorescent
probes allow measurements at low (physiological) peptide concentrations and provide fast response.8
These probes are essential for experiments with natural cell membranes and constitute versatile tools
for investigating the dynamics of peptide binding.9 Fluorescent probes incorporated into peptides
should combine strong changes of their emission upon membrane binding with relatively small size in
order to minimally perturb the peptide structure and report on the environment as close as possible to
the peptide backbone. The classical bioconjugation strategies for fluorescent labeling of peptides are
based on coupling of sulfhydryl- and amino-reactive dyes with cysteine10 and lysine11 amino acids,
respectively. They were successfully applied to graft environment-sensitive dyes, such as NBD,12
Dansyl13, 14 and Prodan,15 to peptides for site-specific probing of the peptide-membrane interactions.
However, with both bioconjugation strategies these dyes are attached to a peptide through a flexible
linker that is at least 0.5 nm long, so that they are not positioned precisely at the site of labeling.16 In
order to locate the probes closer to the peptide backbone 17 and site-specifically monitor the peptide
environment, another strategy is to prepare fluorescent amino acid analogs,18-21 based on
solvatochromic fluorophores like prodan22 or aminonaphtalimide.8, 23
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Though effective enough in the mentioned studies, these amino acids show limited spectroscopic
properties and sensitivity to binding events. To overcome these drawbacks, recently we designed the
first chiral L-amino acid based on a 2-furyl-3-hydroxychromone fluorophore and applied it to probe
site-specifically peptide-nucleic acid interactions.24 Due to an excited state intramolecular proton
transfer (ESIPT) reaction,25 3-hydroxychromone (3HC) and 3-hydroxyflavone (3HF) dyes show dual
fluorescence with superior sensitivity to environment parameters such as polarity and hydration, since
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they combine solvatochromic shifts of normal (N*) and tautomer (T*) emission bands with changes of
the ratio of these bands.26 Such a multiparametric sensing allows fine discrimination of environment of
different types of membranes27-30 as well as monitoring peptide-DNA24, 31 and protein-protein
interactions.32
In the present study, we focused on the 4’-(dimethylamino)-3-hydroxyflavone fluorophore, as its
environment-sensitivity is adapted to membrane research and its spectral properties are suited for
microscopy studies. An L-amino acid AFaa bearing this fluorophore (Fig. 1A) was constructed from
natural tyrosine to allow close location of the fluorophore to the peptide backbone. The fluorophore
moiety of AFaa is similarly lipophilic as the indole unit of tryptophan and should thus mimic its
localization and orientation in the lipid membrane.
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Figure 1. Fluorescent amino acid and labeling of melittin. A) Structure of the 3-hydroxyflavone-based
amino acid (AFaa) and label for amino groups (MFL). The blue arrow shows the direction of the
transition dipole moment.33 B) Excited state intramolecular proton transfer (ESIPT) reaction in
3-hydroxyflavones. C) Structure of melittin based on NMR data (pdb 3QRX).34 Lipophilic amino acids
are shown in cyan; lysines are in blue. Labeling sites L9 and W19 are in pink and orange, respectively.
Five unstructured residues at the basic C-terminus are not shown. The structure of AFaa in red is
overlaid with W19 for size comparison. D) Sequence of melittin. The arrows show the two alternative
positions where the natural amino acids were replaced by the AFaa amino acid.28 Residues forming
membrane-bound α-helix are underlined; cationic amino acids are in blue.
As a model peptide for testing the new fluorescent amino acid, we selected melittin, a well-described
cationic membrane-binding peptide, corresponding to the main toxic component of European honey
bee venom.35 It is composed of 26 amino acids that are predominantly hydrophobic, but contains a
stretch of positively charged amino acids at the C-terminal domain (Fig. 1). Melittin is soluble in water
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and adopts predominantly a random coil conformation, while it self-associates into α-helical tetramers
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at high ionic strength or high peptide concentration.35
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Melittin spontaneously binds to natural and artificial membranes, adopting an α-helical conformation.36
Interestingly, depending on the membrane properties and lipid-to-protein ratio, melittin was reported to
be transmembrane37 or to orient parallel to membrane surface.38, 39 Melittin as an active component of
honey bee venom induces the leakage of cellular content. Some authors suggest that this process is the
result of formation of pores with transmembrane orientation of the peptide.40 The others show that the
relatively high peptide to lipid ratios (>1:200) are needed for its membrane permeabilization activity,
which is more consistent with its binding parallel to the membrane surface.41 In our previous work, we
labeled the N-terminus of melittin with 4’-dimethylamino-3-hydroxyflavone label (MFL, Fig. 1A),
which allowed us monitoring its binding and insertion into model membranes. However, these studies
were limited to the probing of the environment of the peptide N-terminus. Moreover, the spacer used to
attach the probe made the analysis of the peptide orientation in the bilayer imprecise.28
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In the present work, based on the same fluorophore, we synthesized the fluorescent amino acid AFaa
and substituted it for the hydrophobic residues leucine-9 (L9) and tryptophan-19 (W19), middle and
end of α-helical domain of melittin, respectively. This allowed us to determine the insertion depth and
orientation of melittin in model and cell membranes.
Materials and methods
Instrumentation. Proton NMR spectra were recorded on a 300 MHz Bruker spectrometer, and mass
spectra were recorded on a Mariner System 5155 mass spectrometer using the electrospray ionization
(ESI) method. All chemicals and solvents for synthesis and spectroscopic measurements were from
Sigma-Aldrich.
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Amino acid synthesis. The synthesis of the 3-acetyl-L-tyrosine hydrochloride (1) and 3-acetyl-N-
(tert-butoxycarbonyl)-L-tyrosine (2) was performed as described previously.42, 43
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N-(tert-butoxycarbonyl)-3-(3-[4-(dimethylamino)phenyl]prop-2-enoyl)-L-tyrosine (3). 5.00 g (15.48
mmol) of 3-acetyl-N-Boc-L-tyrosine 2 were dissolved in 50 mL of ethanol upon stirring under Ar-
atmosphere. Then, 50 mL of degassed 25% solution of sodium hydroxide were slowly added to the
reaction mixture (cold water bath). After that, 4.00 g of 4-dimethylaminobenzaldehyde (26.81 mmol,
1.73 eq.) were added and the mixture was stirred for 12h at room temperature under Ar. Additional
0.52 g of aldehyde were added and the mixture was stirred overnight. Reaction was monitored by TLC
(CH2Cl2: MeOH, 7: 3 v/v). For that, small aliquots were carefully acidified with HCl and extracted
with ethyl acetate. Then, the orange mixture was carefully acidified with HCl and diluted twice with
water. The obtained oil was separated, dissolved in CH2Cl2, washed with water, dried with MgSO4 and
the solvent was evaporated in vacuo. The obtained oil was crystallized upon standing overnight under
petroleum ether. Orange crystals were recrystallized from toluene/petroleum ether mixture. Yield 5.48
g (78%) of 3 as an orange powder. Mass spectroscopy, ion polarity – negative, m/z: 453.2 [M–H]–; ion
polarity – positive, m/z: 455.2 [M+H]+. 1H-NMR (300MHz, DMSO-d6): δ 1.25 (s, 9H, Boc), 3.00 (s,
6H, N(CH3)2), 3.07 - 3.17 (m, 2H, CH2), 4.09 (m, 1H, CH), 6.78 (d, 2H, J=9Hz, ArH), 7.19 (m, 1H,
ArH), 7.45 (d, 2H, J=9Hz, ArH), 7.53 (d, 1H, J=15Hz, CH), 7.57 (br. s, 1H, Ph-OH), 7.62 (m, 2H,
ArH), 7.91 (d, 2H, J=15Hz, CH), 9.05 (br. s, 1H, NH), 12.78 (br. s, 1H, COOH).
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N-(tert-butoxycarbonyl)-3-(2-[4-(dimethylamino)phenyl]-3-hydroxy-4-oxo-4H-chromen-6-yl)-L-
alanine (4) 5.00 g (11.00 mmol) of chalcone 3 was dissolved in 60 mL of ethanol, cooled in ice bath
and 44 mL of 1 M solution of sodium hydroxide was carefully added with stirring. After that, 2.9 mL
of 30% hydrogen peroxide was added carefully and the mixture was stirred in ice bath for
approximately 6h under Ar-atmosphere. The reaction was monitored by TLC (CH2Cl2:MeOH (8:2 v/v).
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Then the reaction mixture was diluted with 200 mL of ether, acidified with conc. HCl to pH 2 and the
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obtained orange precipitate was filtered and washed with water. Yield of the crude 3-hydroxychromone
4 was 3.55 g. It was purified by crystallization from methanol/water (4:1) mixture. Yield 1.7 g (33%)
of yellow crystals. Mass spectroscopy, ion polarity – negative, m/z: 467.0 [M–H]–; ion polarity –
positive. m/z: 469.0 [M+H]+.1H-NMR (300MHz, DMSO-d6): δ 1.28 (s, 9H, Boc), 3.01 (s, 6H,
N(CH3)2), 3.14 (m, 2H, CH2), 4.15 (m, 1H, J=3 Hz, CH), 6.83 (d, 2H, J=9Hz, ArH), 7.19 (d, 1H,
J=9Hz, ArH), 7.64 (m, 2H, ArH), 7.95 (br. s, 1H, OH), 8.11 (d, 2H, J=9Hz, ArH), 9.14 (br. s, 1H, NH),
12.66 (br. s, 1H, COOH).
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3-(2-[4-(dimethylamino)phenyl]-3-hydroxy-4-oxo-4H-chromen-6-yl)-L-alanine hydrochloride (5).
1.14 g (2.43 mmol) of chromone 5 was dissolved in 14 mL of dioxane and after that, 14 mL of conc.
HCl was added with cooling the reaction mixture in an ice water bath. After 10 min, the bath was
removed and the mixture was stirred for 2-3 h under Ar-atmosphere. Then, it was evaporated under
reduced pressure. Yield 0.896 g (91%) of dark yellow hygroscopic powder. Mass spectroscopy, ion
polarity – negative, m/z: 367.0 [M–H]–; ion polarity – positive, m/z: 369.0 [M+H]+. Corresponds to the
free base, M= 368,4.1H-NMR (300MHz, DMSO-d6): δ 3.04 (s, 6H, N(CH3)2), 3.29 (dd, 2H, J=4.2 Hz,
J=3Hz, CH2), 4.26 (q, 1H, J=3 Hz, CH), 6.91 (d, 2H, J=8Hz, ArH), 7.69 (dd, 1H, J=6Hz, J=3Hz, ArH),
+
7.73 (d, 1H, J=6Hz, ArH), 8.01 (d, 1H, J=3Hz, ArH), 8.14 (d, 2H, J=8Hz, ArH), 8.44 (br. s, 3H, NH3 ).
3-(2-[4-(dimethylamino)phenyl]-3-hy droxy-4-oxo-4H-chromen-6-yl)-N-[(9H-fluoren-9-
ylmethoxy)carbonyl]-L-alanine (6, Fmoc-AFaa). 600 mg (1.48 mmol) of chlorohydrate of amino acid
6 was dissolved in a solution of 500 mg (5.95 mmol) of sodium bicarbonate in 20 mL of water.
Subsequently, 30 mL of DMF was added. Then, 500 mg (1 eq.) of FmocOSu was added and the
mixture was stirred for 24 h under Ar-atmosphere. After that, the mixture was diluted with 20 mL of
water and the precipitate of sodium salt of the crude product was filtered. Finally, it was purified by
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dissolution of the sodium salt in hot water (about 100 mL per 1 g) and further acidification with HCl.
Yield 640 g (73%), orange powder. Mass spectroscopy, ion polarity – positive, m/z: 591.0 [M+H]+. 1H-
NMR (300MHz, DMSO-d6): δ 3.01 (s, 6H, N(CH3)2), 3.25 (m, 2H, CH2), 4.17 (m, 2H, CH2), 4.26 (m,
1H, CH), 6.85 (d, 2H, J=9Hz, ArH), 7.25 (m, 2H, ArH), 7.36 (m, 3H, ArH), 7.58 (m, 2H, ArH), 7.65
(m, 2H, ArH), 7.79 (s, 1H, ArH), 7.84 (d, 2H, J=8Hz, ArH), 8.01 (br. s, 1H, OH), 8.11 (d, 2H, J=9Hz,
ArH), 9.13 (br. s, 1H, NH), 12.79 (br. s, 1H, COOH).
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Peptide synthesis, labeling, and purification. N-terminally labeled MFL-melittin was synthesized
as described previously.28 Melittin peptides, where L9 and W19 residues were replaced by AFaa, were
synthesized by solid phase peptide synthesis on a 433A synthesizer (ABI, Foster City, CA). The
synthesis was performed at a 0.1 mmol scale using standard side-chain protected
fluorenylmethoxycarbonyl (Fmoc)-amino acids and HBTU/HOBt coupling protocol. LL-amide resin
(54 mmol/g reactive group concentrations) was used as a solid support.
Coupling of the fluorescent AFaa amino acid was performed manually, in 10 mL flask using the
same coupling solution as for automated synthesis. In our experiments, we used only a 2-fold excess of
the AFaa over the peptidyl resin (comparing with 10-fold for other amino acids). Therefore, we
increased the coupling time to 2 h (comparing with 20 min) and added the labeled amino acid in two
steps (“double coupling”) to minimize formation of peptide lacking the labeled residue. To ensure that
no deletion product could form, we capped the unreacted N-terminal amino groups with acetic
anhydride. At the end of the synthesis, the peptidyl resin was isolated and washed twice by NMP.
Cleavage and deprotection of the peptidyl resin were performed for 2 h using a 10 mL trifluoroacetic
acid (TFA) solution containing water (5%, v/v), TIS (iPr3SiH, 2.5%, v/v), thioanisole (5%, v/v) and
ethanedithiol (2.5%, v/v). During the deprotection step, common scavengers used to protect Lys or
Arg (such as iPr3SiH) residues were added to prevent the AFaa fluorophore to undergo ring opening
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reactions. The solution was concentrated in vacuo and the peptide was precipitated by ice-cold diethyl
ether and then, pelleted by centrifugation. The pellet was washed with diethyl ether and dried. The
peptides were solubilized with aqueous TFA (0.05 %, v/v). HPLC purification was carried out on a C8
column (uptisphere 300A, 5µm; 250Х10, Interchim, France) in a water/acetonitrile mixture containing
0.05% TFA with linear gradients depending on the peptide: for L9 - 30-60% acetonitrile in 40 min and
for W19 - 40-60% acetonitrile in 30 min. Molecular masses obtained by ion spray mass spectrometry
were: 3082,79 for L9; 3009,79 for W19, in agreement with the expected theoretical masses. Prior to
use, peptides were dissolved in distilled water, aliquoted and stored at -20°C. Concentrations of the
labeled peptides were determined using the label absorbance at 400 nm ( = 33000 M-1cm-1).
Fluorescence measurements. Absorption spectra were measured on a Cary 4 spectrophotometer
(Varian) and fluorescence spectra on a FluoroLog (JobinYvon, Horiba) spectrofluorometer.
Fluorescence emission spectra were recorded at 400 nm excitation wavelength in 20 mM phosphate
buffer containing 150 mM NaCl (pH = 7.4) at 20°C. All the spectra were corrected for Raman
scattering and background fluorescence. Fluorescence quantum yields were determined using
4’-(diethylamino)-3-hydroxyflavone in ethanol (quantum yield 0.52)44 as a reference. Titration
experiments were performed by mixing of 300 nM melittin with 3 to 300 μM lipids, and incubation for
1 h prior to measurements. Each data point was performed independently. Deconvolution of the
fluorescence spectra of the labelled peptides into three bands, corresponding to the normal (N*), H-
bonded normal (H-N*) and tautomer (T*) forms, was performed using the Siano software kindly
provided by Dr. A.O. Doroshenko (Kharkov, Ukraine), as previously described.28, 45 The individual
emission bands were approximated by a log-normal function accounting for several parameters:
maximal amplitude, Imax, spectral maximum position, νmax, and position of half-maximum amplitudes,
ν1 and ν2, for the blue and red parts of the band, respectively. These parameters determine the shape
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parameters of the log-normal function, namely the full width at the half-maximum, FWHM=ν1−ν2, and
the band asymmetry, P=(ν1−νmax)/(νmax−ν2). For the iteration process, the FWHM of the two short-
wavelength bands (N* and H-N*) were fixed at 3000 cm-1. For the H-N* band, the asymmetry and the
band position were fixed at 0.9 and 18400 cm-1, respectively. The other parameters were allowed to
vary in the iteration process. The resulting fluorescence intensities of the separated N* and T* bands
were used to calculate the N*/T* ratio.
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Large unilamellar vesicles. Large unilamellar vesicles (LUVs) of ~ 100 nm diameter (according to
dynamic light scattering, DLS) were obtained by the classical extrusion method as previously described
45: evaporation of chloroform solution of lipids, hydration of the resulted film in buffer and ten
extrusions of the obtained solution through membrane with 100 nm pores. Small unilamellar vesicles
(SUVs) for CD spectroscopy were prepared by ultrasonication (~40 min, 20°C) of the lipid suspension
with size distribution control by DLS (45±7 nm).
Giant unilamellar vesicles. Giant unilamellar vesicles (GUVs) were generated by electroformation
in a home-built liquid cell, using previously described procedures.46, 47 1 mM solution of lipids in
chloroform was deposited on the platinum wires of the chamber, and the solvent was evaporated under
vacuum for 30 min. The chamber was filled with a 300 mM sucrose solution, and a 2-V, 10-Hz
alternating electric current was applied to this capacitor-like configuration for ca. 1.5 h. Then, a 50 μL
aliquot of the obtained stock solution of GUVs in sucrose was added to 200 μL of 300 mM glucose
solution to give the final suspension of GUVs used in microscopy experiments. Staining of GUVs was
performed by addition of an aliquot of the peptide solution to obtain a 0.2 μM final concentration.
Cell preparation and Staining. HeLa cells were cultured in Dulbecco's modified Eagle medium
(DMEM, high glucose, Gibco-Invitrogen) supplemented with 10% (v/v) fetal bovine serum (FBS,
Lonza), 1% antibiotic solution (penicillin-streptomycin, Gibco-Invitrogen) in a humidified incubator
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with 5% CO2 atmosphere at 37°C. A cell concentration of 5-10104 cells/mL was maintained by
removal of a portion of the culture and replacement with fresh medium, three times per week. For
microscopy studies, cells were seeded onto a chambered coverglass (IBiDi) at a density of 5×104
cells/IBiDi. After washing the cells with Opti-MEM, a solution of labeled melittin diluted in 1 mL of
Opti-MEM was added to the cells to a final concentration of 1 µM and immediately measured.
Microscopy. Fluorescence microscopy experiments were performed by using a home-built two-
photon laser scanning setup based on an Olympus IX70 inverted microscope with an Olympus 60x
1.2NA water immersion objective.48 Two-photon excitation at 830 nm was provided by a titanium-
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sapphire laser (Tsunami, Spectra Physics). A density filter of 0.2 was used on the red channel. Images
corresponding to the blue and red channels were recorded simultaneously using a dichroic mirror at
555 nm and two APDs. The images were processed with LabView-based (for GUVs) or ImageJ-based
(for cells) programs that generate a ratiometric image by dividing the image of the blue channel by that
of the red channel.47, 49 For each pixel, a pseudocolor scale is used for coding the ratio, while the
intensity is defined by the integrated intensity recorded for both channels at the corresponding pixel.
Results and Discussion
Design and synthesis. The Fmoc-protected fluorescent L-amino acid Fmoc-AFaa was designed by
extending the side chain of L-tyrosine into a 3-hydroxyflavone group bearing an electron-donating
dimethylamino group (Scheme 1). The lipophilicity of AFaa is close to the one of tryptophan
(calculated logP values are equal to 0.48 and 0.11, respectively). Moreover, the side chain of the AFaa
amino acid is only ~0.5 nm longer than the tryptophan indole ring, and the distance from the
fluorophore to the peptide backbone is 0.2-0.7 nm (Fig. S1), which is less than the diameter of α-helix.
All these features ensure a better site-specific probing of the environment than the cysteine labeling
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approach (>1 nm distance from the backbone). Similarly to natural aromatic amino acids, the rotation
of AFaa fluorophore with respect to the peptide backbone is limited to C-Cβ and Cβ-C6 bonds. Thus,
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the rotational freedom around C-Cβ bond should be as for tyrosine and tryptophan in native peptides50
with the preference for trans-conformation of the aromatic residue relative to the carbonyl in their -
helix. A key feature is that the orientation of the transition dipole of the AFaa fluorophore is parallel to
its rotation axis (Cβ-C6 bond, Fig 1), which decreases the uncertainty of the dye dipole orientation in
respect to the peptide and thus, allows the determination of the peptide orientation by the orientation of
the AFaa fluorophore.
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Scheme 1. Synthesis of Fmoc-AFaa. i) AcCl, AlCl3, PhNO2; ii) Boc2O; iii)
4-dimethylaminobenzaldehyde, NaOH, EtOH/H2O; iv) H2O2, NaOH, EtOH; v) HCl, dioxane; vi)
FmocOSu, NaHCO3, DMF/H2O.
Fmoc-protected fluorescent L-aminoacid Fmoc-AFaa was prepared from L-tyrosine according to the
strategy we described earlier for other fluorescent amino acid.24 Synthesis of Fmoc-AFaa was done
through tyrosine C-acylation, Boc-protection and condensation of the resulted product (2) with
aldehyde into N-Boc-protected 2'-hydroxychalcone (3). Then, oxidative cyclization in basic conditions
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into N-Boc-protected-3-hydroxyflavone (4) was performed, followed by the change of the protective
group to Fmoc (Scheme 1).
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In the preparation of AFaa, none of the reaction steps include the asymmetry center conversion, but the
steps iii and iv require application of alkali. Therefore to check the optical purity of the obtained amino
acid, we synthetized the amide derivatives of (5) with (R)- and (S)-α-phenyl-ethylamine (Fig. S2). The
obtained diastereoisomers show differences in 1H-NMR-spectra (Figure S3 and discussion in SI) and
comparison of peak intensities of the unpurified (S,R)- and (S,S)-amides shows that racemization is
less than 10%.
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Synthesis and Structure of Melittin Fluorescent Variants. Melittin variants labeled by AFaa amino
acids at positions L9 and W19 were synthesized according to typical Fmoc SPPS protocols. Coupling
of Fmoc AFaa amino acid was done using a 2-fold excess and 2h reaction time. The 3-OH group of
AFaa was not unprotected, because being acylated by amino acid or acetic anhydride it was easily
recovered at the conditions of peptide cleavage from the resin. We saw no signs of AFaa
decomposition on HPLC profile of the crude peptides. Melittin peptide labeled at the N-terminus with
MFL label was prepared as described previously.28
To check the influence of the introduction of the fluorescent amino acid on the affinity of melittin to
lipid membranes, we titrated the labeled melittin peptides with POPC liposomes and monitored the
increase in α-helix content by CD spectroscopy at 222 nm (Fig. S4). The labeled peptides show binding
profiles similar to the one reported for unmodified melittin (Fig. S4),50, 51 indicating that substitution of
either L9 or W19 by AFaa does not significantly perturb the peptide conformation and its membrane
binding affinity.
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Fluorescent properties of AFaa and the labeled peptides in solution. The AFaa amino acid shows a
strong dependence of its dual emission on solvent polarity and its fluorescence is strongly quenched in
aqueous media. These properties are similar those of the MFL label28 and can thus be used to report on
the local environment of the labeled sites of melittin.
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Being diluted in buffer solution to a final concentration 0.3 µM, all labeled peptides show complex
time dependent behavior, as their fluorescence intensity strongly decreases with time and the spectral
shape changes. Indeed, for the N-terminal labeled melittin (Fig. 2A) and W19-substituted peptide (Fig.
2B), the single band, which can be assigned to the N* emission shifts to the red, while for L9-
substituted melittin the dual emission degrades into single red shifted N* band (Fig. 2C). The
spectroscopic changes observed for all three peptides clearly suggest the strong increase in the polarity
and hydration of the probe environment, which can only be explained by the dissociation of the melittin
aggregates after their dilution in the buffer from the concentrated aqueous stock solution of ~100 µM.
Indeed, in the concentrated solution melittin molecules are reported to be in the form of tetramers,35
while after dilution in the buffer to micromolar concentration they dissociate giving monomeric
species, presenting much higher exposure of the 3HF dye to bulk water. It should be noted, in
oligomeric melittin (spectra recorded immediately after dilution) the label at position W19 and N-
terminus show a single band emission typical for highly hydrated media, while at position L9 it shows
two well separated emission bands, reporting nonpolar aprotic environment, in agreement with the
previously described structure of tetramer51 where L9 segment of peptide is screened from water, while
the other labeled positions are more exposed.
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24h
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Figure 2. Changes of fluorescence spectra upon dissociation of melittin oligomers. A) N-terminal
labeled melittin, B) and C) W19 and L9 labeled variants. Time after preparation: 2 min. (blue lines) or
24 h. (red lines). Peptide concentration was 300 nM, phosphate buffer pH 7.2, 150 mM NaCl, 25°C.
Probing Peptide-Membrane Interactions. Melittin efficiently binds to natural and model membranes.
Interaction of the labeled peptides with DOPC vesicles results in the appearance of two emission bands
typical for nonpolar media and a strong (up to 20-fold) increase in their fluorescence intensity (Fig. 3a),
in line with an immersion of the label in the hydrophobic viscous medium of the membrane. Due to
these strong changes in the fluorescence intensity, the fluorescence of free peptide is negligibly small,
so that only bound form contributes for the fluorescence spectra in the presence of vesicles.
The spectral profiles of the three peptide variants differ greatly in DOPC vesicles, indicating that the
insertion of the 3HF fluorophore in lipid membranes depends on its position in melittin. Using
previously established deconvolution of the spectra into three bands (N*, hydrated H-N*, and T*), we
estimated the ratio N*/T* intensity ratio, the parameter that correlate directly with the dielectric
constant of the environment.28 AFaa at position L9 shows the lowest N*/T* band ratio (0.27), which
corresponds to the dielectric constant   5. According to the dielectric model of lipid bilayer suggested
by Griffith et al,52 this very low value of polarity indicates the deep immersion into membrane in the
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region of alkyl chains of fatty acids (Fig. 4). For W19 variant, the N*/T* ratio is much larger (1.14) and
corresponds to   20, indicating that W19 position is much more water exposed, close to the
lipid/water interface. The N-terminal label connected through a flexible linker (MFL) shows
intermediate N*/T* ratio value (0.70) with   10, which in line with our previous report28 suggests the
dye localization in the region of the ester groups of phospholipids. Our data are consistent with the
current models of melittin in the lipid bilayer, since for both vertical and horizontal orientation of the α-
helix, the L9 residue is assumed to protrude into the bilayer, while the W19 residue, placed next to
charged amino acids, is expected to localize at the interface. Such interfacial localization of Trp has
been reported for many other membrane binding peptides.53 Thus, both L9- and W19-labeled peptides
show similar insertion as the native one, which is probably because in both cases the hydrophobic
amino acids (L and W) were replaced with a hydrophobic AFaa.
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Remarkably, addition of cholesterol to the lipid bilayers induces only minor changes in the dual
emission of L9- and W19-labelled peptides (Fig. 3B). Deconvolution analysis shows that the N*/T*
ratio is nearly the same for L9 (0.26) and only marginally increased for W19 peptide variant (1.34). In
contrast, the N-terminal label exhibits 2-fold decrease in the N*/T* ratio (to 0.36), indicating a strong
decrease in the environment polarity of the label. As in the N-terminal labeled melittin the 3HF dye is
connected to the peptide through a flexible linker, we speculate that cholesterol directs the probe deeper
into the bilayer, independently from the peptide. Therefore, fluorescent amino acid AFaa (in L9 and
W19 positions) is far better than the N-terminal probe for studying insertion of the peptide into the
membranes due to rigid connection to the backbone.
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Figure 3. Probing peptide-membrane interactions, using labeled melittin peptides. A) Fluorescence
spectra of N-terminal, L9- and W19-labeled melittin peptides in buffer (dash) and DOPC vesicles
(solid lines) at lipid/peptide (L/P) ratio = 250/1. B) Normalized emission spectra of peptides in DOPC
(solid lines) and DOPC:Cholesterol (6:4) (dashed lines) vesicles. Concentration of melittin and lipids
was 300 nM and 15 μM, respectively. C) Normalized emission spectra of peptides bound to live cells.
300 nM of melittin variants were added to HeLa cell suspensions and recorded after 2 min incubation.
Finally, we recorded the fluorescence spectra of the labeled melittin peptides added to suspensions of
live HeLa cells. The spectra show the same trends as for model membranes (Fig. 3C), suggesting that
the insertion of melittin into cell plasma membranes should be similar to the model systems.
Determination of peptide orientation in membrane. The amphiphilic helix of melittin could lay
parallel to the membrane surface on the water/lipid interface or form pores with transmembrane
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orientation perpendicular to the surface. These two possible conformations could be discriminated
using AFaa-labeled melittins and fluorescence microscopy with a polarized light source, to excite
selectively fluorophores that are oriented parallel to the polarization plane of the light.47, 54 Since the
AFaa amino acid is structurally close to a natural aromatic amino acid, its fluorophore should be
oriented almost perpendicular to the α-helix. Moreover, according to NMR data,34 the W19 and L9
lateral chains in the -helix of melittin are oriented almost perpendicular to each other (Fig. 4).
Therefore, if melittin α-helix is oriented perpendicular to the membrane plane, both L9 and W19 labels
will be oriented parallel to the membrane plane. In contrast, a parallel orientation of the melittin α-helix
to the membrane plane will result in opposite orientations of W19 and L9 with respect to the membrane
plane (Fig. 4).
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Figure 4. Model of melittin insertion and orientation in the lipid membrane (front and side views). The
conformation of the peptide is reported elsewhere.34 The curved double arrows show the expected
orientational freedom for the AFaa fluorophore, where the hydrated species (dotted arrows) are
shallower in the bilayer than the non-hydrated ones (solid arrows). We assume that the orientation of
transition dipole moment in AFaa fluorophore is the same for these two species.
We performed two-photon excitation microscopy of melittin bound to DOPC giant unilamellar vesicles
(GUVs) (Fig. 5a). The polarized two-photon light source allows photo-selection of the fluorophores
when their transition dipole is parallel to the direction of the light polarization 47, 54. The emission of the
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N* and T* species of AFaa was recorded in the blue (480-565 nm) and red (>565 nm) channels,
respectively. According to our previous studies, the T* emission is directly linked to the non-hydrated
species of the dye, while the N* emission corresponds to a mixture of non-hydrated and hydrated
species.45 The hydrated species usually exhibit a shallower location and therefore a somewhat different
orientation with respect to the non-hydrated species.47 Our results show that for the L9-labeled melittin,
the effect of selective excitation by the polarized light is observed mostly for the blue channel, as
shown by the bright emission of the vesicle at its equator region (Fig. 5 A1). As a result, the higher
values of the blue/red ratios in the ratiometric image correspond to the equator region of GUVs (Fig. 5
A3). Assuming the same orientation of the transition dipole moments in AFaa fluorophore for hydrated
and non-hydrated species, the observed differences in the photo-selection profiles suggest that these
two species are oriented differently in the bilayer. We can conclude that the orientation of AFaa
fluorophore in L9-position varies from titled for non-hydrated species till horizontal closer to the
bilayer surface for the hydrated species (Fig. 4). In contrast, the W19 melittin analogue shows exactly
the opposite behavior, since the effect of light polarization is observed only for the red channel, with a
clear photo-selection for the equator region of the GUVs (Fig. 5 B2). As a consequence, in the ratio
images, the color contrast appears the opposite to that of L9 melittin (Fig. 5 B3). Thus, for the W19
melittin, the orientation of AFaa fluorophore varies from horizontal for the non-hydrated species to
tilted towards the bilayer surface for the hydrated species (Fig. 4). Consequently, the L9 and W19
positions of melittin differ drastically not only by the insertion of the 3HF fluorophore, but also by its
orientation in the lipid bilayer. The fact that L9 and W19 derivatives show different behaviors strongly
suggests that their orientation in the bilayer is imposed by the melittin peptide. Taken together, these
data support a parallel orientation of the melittin to the membrane surface,55 where the L9 and W19
residues present different orientations relatively to the surface (Fig. 4). Our results are consistent with
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recent EPR38 and oriented circular dichroism studies56 showing that most of the melittin -helix stays
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Finally, the data of the N-terminal labeled melittin suggest that orientation of 3HF fluorophore varies
from vertical for the non-hydrated species to tilted towards the membrane surface for the hydrated
species, in line with our previous studies.28 However, fluorescence of the N-terminal labeled melittin
does not provide a clear information about the peptide orientation, as its fluorophore is connected to the
peptide through a flexible linker. As a consequence, its non-hydrated species tends to orient vertically
in the bilayer, as it is observed for membrane probes based on this fluorophore.45, 47
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Figure 5. Two-photon fluorescence microscopy images of L9- (A), W19- (B), and N-terminal (C) –
labeled melittin bound to DOPC GUVs. The images were recorded at the blue channel (A1, B1 and C1)
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and red channel (A2, B2 and C2). Then, the blue/red ratio images were calculated (A3, B3 and C3).
The color of each pixel of these ratio images represents the Iblue/Ired intensity ratio, while the pixel
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intensity corresponds to the total intensity at both channels. Two-photon excitation wavelength was at
830 nm. Double headed arrows indicate the orientation of light polarization, while other arrows
indicate the photo-selection areas. Scale bar is 10 μm. Concentration of melittin was 0.2 μM.
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Next, the same approach was applied to determine the melittin orientation in cell membranes. Since
melittin is a membrane active peptide, it leads even at 1 μM concentration to the disruption of the
plasma membrane of HeLa cells and formation of micrometer-scale vesicles (Fig. 6), which originate
from the plasma membranes. Microscopy images of these vesicles clearly show the effects of light
polarization, which are the same as those observed for GUVs (Fig. 5). Thus, the orientation of AFaa
dye at L9 and W19 positions of melittin in the plasma membranes is similar to that in the model lipid
bilayers. Such orientation of the labels can be realized only when melittin -helix binds parallel to the
cell membrane surface and does not adopt transmembrane orientation. Thus, using the new amino acid,
for the first time to our knowledge, we were able to register the orientation of melittin in natural
membranes, and show that it is the same as in model membranes.
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Figure 6. Two-photon fluorescence microscopy images of Hela cells stained with L9- (A), W19- (B)
and N-terminal- (C) labeled melittin. The upper panels (A1-C1) represent typical cells generating
vesicles in the presence of melittin, while the lower panels (A2-C2) show selected images of the
produced vesicles. The color of each pixel represents the value of the intensity ratio Iblue/Ired, while the
pixel intensity corresponds to the total intensity at both channels. Double-headed arrows indicate the
orientation of the light polarization. Two-photon excitation wavelength was at 830 nm. Sizes of the
images were 70 × 70 μm. Peptide concentration was 1μM.
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Conclusion
A new fluorescent L-amino acid (AFaa) bearing an environment-sensitive 3-hydroxychromone
fluorophore as a side chain was synthesized and introduced into the melittin peptide at two alternative
positions by solid-phase synthesis. The structure of AFaa is derived from the L-tyrosine amino acid,
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which confers a well-defined orientation of the fluorophore in respect to the peptide α-helix and
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approaches through a flexible linker. These attractive features were successfully used to determine the
insertion and orientation of the labeled melittin peptide in lipid bilayers. Our spectroscopy and
microscopy data suggested that melittin is preferentially oriented parallel to the bilayer surface in
model and natural (cellular) membranes. The developed amino acid and proposed methodology will be
of interest to study other membrane peptides.
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Acknowledgements
V.Y.P., O.M.Z, V.G.P. and this work were supported by ANR Fluoaant, ANR Femtostack, ANR
Fluometadn, ANR Cellmemprobes, CNRS, and Université de Strasbourg. O.V.S. was supported by
ARCUS Alsace. We thank Cyril Antheaume and Patrick Wehrung for NMR and mass measurements,
respectively.
Supporting Information Available
Additional characterization data on the fluorescent amino acid and melittin peptides are presented. This
material is available free of charge via the Internet at http://pubs.acs.org.
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