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Formation of size-controlled, denaturation-resistant lipid nanodiscs by an amphiphilic self-polymerizing peptide
Colloids and Surfaces B: Biointerfaces 146 (2016) 423–430
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Formation of size-controlled, denaturation-resistant lipid nanodiscs by an amphiphilic self-polymerizing peptide Hiroaki Kondo, Keisuke Ikeda ∗ , Minoru Nakano Graduate School of Medicine and Pharmaceutical Sciences, University of Toyama, 2630 Sugitani, Toyama 930-0194, Japan
a r t i c l e
i n f o
Article history: Received 16 March 2016 Received in revised form 22 June 2016 Accepted 22 June 2016 Available online 23 June 2016 Keywords: Nanoparticle Phosphatidylcholine Amphiphilic peptide Self-assembly Native chemical ligation
a b s t r a c t Nanodiscs are discoidal particles with a planar phospholipid bilayer enwrapped by proteins such as apolipoprotein A-I. Nanodiscs have been widely used for analyzing structures and functions of membrane proteins by dispersing them in solution. They are expected to be used as drug carriers and therapeutic agents. Amphiphilic peptides are known to form nanodiscs. However, the lipid-peptide nanodiscs are relatively unstable in solution, making them unsuitable for many applications. Here, we report the synthesis of an amphiphilic self-polymerizing peptide termed ASPP1, which polymerizes by intermolecular native chemical ligation reactions. ASPP1 spontaneously formed nanodiscs when added to phospholipid vesicles without using detergents. The diameter of the planar lipid bilayer in the nanodiscs was controlled by the lipid:peptide molar ratio. ASPP1-nanodiscs exhibited greater stability at high temperatures or in the presence of urea than nanodiscs formed by the non-polymerizing amphiphilic peptide or apolipoprotein A–I. Average and maximal degrees of ASPP1 polymerization were 2.4 and 12, respectively. Self-polymerization of the peptide appears to be responsible for stabilization of the nanodiscs. Our results open a new avenue for the development of nanodisc technology. © 2016 Elsevier B.V. All rights reserved.
1. Introduction Lipid nanodiscs are nanometer-sized, discoidal, supramolecular complexes with a patch of planar phospholipid bilayer enwrapped by amphiphilic, ␣-helical polypeptides [1,2]. Nanodiscs dispersed in water are stabilized by hydrophobic interactions, where hydrophobic acyl chains of lipids and nonpolar faces of ␣helices are sequestered from the aqueous solution. Apolipoprotein A–I (apoA–I), the main component of high-density lipoproteins (HDLs), has been reported to form discoidal particles with lipids [3,4]. Cholesterol is incorporated into the discoidal HDLs and metabolized to cholesteryl ester by lectin-cholesterol acyltransferase, resulting in the formation of spherical HDLs which transport excess cholesterol from peripheral tissues to the liver [5]. In addition to the biological significance of discoidal HDLs in cholesterol homeostasis, nanodiscs have attracted considerable interest for their potential use in nanomedicines and drug delivery systems, and as model membrane systems for studying membrane proteins [6–10]. Indeed, a number of membrane proteins, such as bacteriorhodopsin, G-protein coupled receptors, and epidermal growth
∗ Corresponding author. E-mail address: [email protected] (K. Ikeda). http://dx.doi.org/10.1016/j.colsurfb.2016.06.040 0927-7765/© 2016 Elsevier B.V. All rights reserved.
factor receptor, have been assembled into nanodiscs that mimic native lipid bilayer environments. Various biophysical methods and spectroscopic techniques can be used to analyze structures and functions of the solubilized proteins. For a wide range of applications, it is essential to have control over the physicochemical parameters of nanodiscs such as size, stability, lipid composition, and chemical modification. For example, Denisov et al. designed extended and truncated versions of apoA–I, termed membrane scaffold proteins (MSPs), to customize the sizes of the nanodiscs in the range of 9.5–12.8 nm [2]. Hagn et al. designed further truncated versions of MSPs to create smaller nanodiscs with a diameter of ∼6 nm, whose short rotational correlation times were suitable for solution-state NMR studies of integrated membrane proteins [11]. They determined the structure and dynamics of the -barrel protein, OmpX, integrated in the small nanodiscs. Similar to proteins, amphiphilic ␣-helical peptides have also been reported to form nanodiscs in cooperation with lipids [12–14]. Lipid-peptide nanodiscs have several advantages with respect to nanodisc assembly and physicochemical parameters. Nanodiscs can spontaneously form simply by mixing the peptides and lipid suspension (e.g. liposomes) in buffer without the aid of detergents, which may disrupt the structures and functions of membrane proteins. Their sizes can be easily controlled by varying the lipid:peptide ratio when the components are mixed in buffer.
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Table 1 Peptide sequences.
2.2. Peptide synthesis
Peptide
Amino acid sequence
Ac-18A-NH2 ASPP1
Ac-DWLKAFYDKVAEKLKEAF-CONH2 NH2 -CGDWLKAFYDKVAEKLKEAF-COSBna
a
COSBn = benzyl thioester.
Moreover, large-scale synthesis and chemical modifications of the peptides are feasible. Despite these promising properties of lipidpeptide nanodiscs, they are unstable as evidenced by the fact that short amphiphilic peptides of 11 amino acid residues cannot form nanodiscs [15]. In addition, Imura et al. showed that the 22-residue nanodisc-forming peptide derived from human apoA–I (220–241) did not form the assembly below the critical association concentration of ∼10−5 M [13]. These instabilities are probably because relatively short peptides have weaker hydrophobic interactions with the edges of nanodiscs than the linked amphiphilic ␣-helices of apoA–I or MSPs. Stable nanoparticles under physiological conditions are required for in vivo therapeutic applications. Recently, Zhao et al. have reported that branched, multivalent constructs of amphiphilic peptides formed stable nanodiscs that were resistant to enzymatic digestion and had long half-lives in mice [16]. This study raises the possibility that the further chemical ligation of amphiphilic peptides would increase the stability of peptidelipid nanodiscs. In addition, highly stable lipid-peptide nanodiscs are required for studying integrated membrane proteins requiring long experimental period and exposure to high temperatures and denaturing agents. Here, we report synthesis of size-controlled, denaturationresistant lipid nanodiscs using an amphiphilic self-polymerizing peptide termed ASPP1 that can withstand denaturing conditions. The peptide incorporates the apoA–I mimetic sequence 18A, which was originally designed by Anantharamaiah et al. [12]. The sequence of 18A has properties similar to the amphiphilic helices of apoA–I. N- and C-terminal blocked 18A (Ac-18A-NH2 , Table 1) induced a larger increase in ␣-helical content in complex with phosphatidylcholine than unblocked 18A and formed nanodiscs similar to apoA–I nanodiscs [17]. As 18A does not have residues at the junctions of the tandem helices, we attached cysteine-glycine residues at the N-terminus as a flexible linker. We also modified the C-terminus with a benzyl thioester group, which causes a native chemical ligation (NCL) reaction with the free cysteine residue at the N-terminus to form an amide bond [18]. We hypothesized that self-polymerization of ASPP1 would occur, which would facilitate hydrophobic interactions between the interlinked ␣-helices and lipid bilayers, thereby increasing the conformational stability of the nanodiscs. Formation of nanodiscs by ASPP1 with a phospholipid, sizes of the nanodiscs, and intermolecular ligation of ASPP1 were investigated. We also demonstrate the ability of the ASPP1 nanodiscs to resist unfolding of the peptide secondary structures and deformation of the assembly triggered by high temperatures or by denaturant.
2. Materials and methods
Ac-18A-NH2 was synthesized using Fmoc-based solid-phase synthesis and cleaved from the resin with trifluoroacetic acid/water/triisopropylsilane (95/2.5/2.5) for 3 h. For synthesis of ASPP1 with C-terminal benzyl thioester, the diaminobenzoyl linker-attached Rink amide resin developed by Dawson et al. was employed [20]. Peptide chains were extended on Dbz AM resin (Merck Millipore) by Fmoc-based chemistry. An N-terminal cysteine residue was introduced using Boc-Cys(Trt)-OH/N,N’diisopropylcarbodiimide/1-hydroxybenzotriazole. A solution of p-nitrophenylchloroformate (5 eq) in dichloromethane was added to the resin and the mixture was stirred for 1 h. After washing the resin with DCM and DMF, 0.5 M N,N-diisopropylethylamine in DMF was added and the mixture was stirred for 30 min. After cleavage of peptides from the resin with trifluoroacetic acid/water/triisopropylsilane/1,2-ethanedithiol (92.5/2.5/2.5/2.5), the peptides were dissolved in DMSO containing 620 mM benzylmercaptan and left overnight. The peptides were purified by reverse phase high-performance liquid chromatography (RP-HPLC) using a C18 HPLC column with a gradient of water/acetonitrile containing 0.1% trifluoroacetic acid. Peptides were characterized by HPLC and MALDI-TOF MS (Autoflex-T1, Bruker Daltonics). We determined the purity of ASPP1 to be >90% by HPLC and obtained the expected molecular mass (Fig. S1). Ac-18A-NH2 lyophilized powder was dissolved in water, and ASPP1 lyophilized powder was dissolved in a 1 mM HCl solution and the concentrations were determined by measuring absorbance at 280 nm. 2.3. Liposomes Large unilamellar vesicles (LUVs) were prepared as follows [14]: an aliquot of POPC in chloroform-methanol was transferred to a round-bottom flask and the solvent was removed using a rotary evaporator. Thereafter, the sample was dried overnight under vacuum. Buffer (20 mM MOPS/1 mM EDTA, pH 7.0) was added to the remaining lipid film. The lipid suspension was freeze-thawed 5 times and extruded through a 100 nm pore filter. The size of LUVs was determined using a DelsaMax Core dynamic light scattering (DLS) instrument (Beckman Coulter). The concentration of lipid was determined by an enzymatic assay kit for choline (Wako). 2.4. Nanodiscs Preparation of POPC-apoA–I nanodiscs was based on the sodium cholate dialysis method [21]. Sodium deoxycholate was added to the lipid suspension in MOPS buffer at a POPC:cholate molar ratio of 1:5. Then, apoA–I was added at a molar ratio of POPC:apoA–I of 78:1. After incubation for 24 h at 4 ◦ C, nanodiscs were obtained by dialysis against MOPS buffer for 2 days. The lipid-peptide nanodiscs were obtained by mixing the peptides and POPC LUVs (100 M) at molar ratios of 1:1–1:16 in buffer containing 10 mM 4mercaptphenylacetic acid (MPAA) at 25 ◦ C. Solubilization of LUVs and nanodisc formation was confirmed by right-angle light scattering at a wavelength of 650 nm using an F-4500 fluorescence spectrometer (Hitachi). The sizes of nanodiscs were determined using a DelsaMax Core dynamic light scattering (DLS) instrument (Beckman Coulter) at 20 ◦ C.
2.1. Materials 2.5. Transmission electron microscopy (TEM) 1-Palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) was purchased from Avanti Polar Lipids. 1,6-Diphenyl-1,3,5-hexatriene (DPH) was obtained from Molecular Probes. ApoA–I was isolated from pig plasma using procedures described previously [19]. The purity of apoA–I was determined to be >95% by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE).
A glow-discharged Cu grid coated with thin carbon film (300 mesh) was placed on a droplet of the sample for 5 min. Then, excess sample solution on the grid was removed using filter paper. Next, the grid was placed on a droplet of 2% phosphotungstic acid for 30 s and dried after removal of the excess solution. TEM images
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of nanodiscs were obtained using a JEM-1400 transmission electron microscope (JEOL) operated at 80 keV. Diameters of nanodiscs were measured by PhotoRuler software employing images of 100 particles (Fig. S2). 2.6. SDS-PAGE SDS-PAGE was performed as follows [22]: sample solution was mixed with 3 vol of sample buffer (150 mM Tris/HCl, pH 7.0, 12% SDS (wt/vol), 6% mercaptoethanol (vol/vol), 30% glycerol (wt/vol), 0.05% Coomassie blue G-250) and loaded in 16% polyacrylamide/6 M urea gel. Gels were stained with 0.025% Coomassie blue in 10% acetic acid. The degree of polymerization of ASPP1 was assessed by measuring band densities using ImageJ software. For silver staining, the gel after Coomassie blue staining was washed with 50% methanol, 50 mM ammonium hydrogen carbonate. Silver staining was carried out using the 2D-SILVER STAIN-II kit (Cosmo Bio). 2.7. Circular dichroism (CD) spectroscopy Due to the high absorbance of MPAA, MPAA in solution was removed by dialysis against MOPS-NaOH buffer without MPAA for 48 h. For carrying out measurements in the presence of urea, samples were equilibrated with urea for 72 h at 4 ◦ C. CD spectra were acquired at 25 ◦ C using a J-805 spectrometer (JASCO) with a 1 mm path length quartz cell. Eight scans were averaged for each sample. 2.8. Fluorescence anisotropy A small volume of a methanol solution (final 1% (v/v) methanol) of DPH was added to either LUVs or nanodiscs and incubated at 25 ◦ C for 1 h to incorporate 0.5 mol% DPH into the lipid bilayers. Fluorescence anisotropy of DPH was measured using an F-4500 fluorescence spectrometer with polarizers placed in the excitation and emission paths. Excitation and emission wavelengths were set at 360 and 430 nm, respectively.
Fig. 1. Formation of lipid-peptide nanodiscs at 25 ◦ C. Changes in the right-angle light scattering intensities of POPC LUVs upon addition of Ac-18A-NH2 (A) or ASPP1 (B) were monitored at 650 nm. The POPC:peptide molar ratios were set to 2:1, 4:1 and 8:1 at a POPC concentration of 100 M. Scattering intensity, I(t), was normalized by the intensity of the POPC LUVs before the additions of peptides, I(0).
3. Results 3.1. Formation of lipid-peptide nanodiscs ASPP1 was added to POPC LUVs having diameters of ∼100 nm at 25 ◦ C. We observed a decrease in the right-angle light scattering intensities of the LUVs in the presence of the ASPP1 as well as Ac-18A-NH2 (Fig. 1), suggesting that the vesicles were solubilized into small particles [14]. The decrease in intensities occurred over a period of 1 day, and no further significant changes were observed over a period of 1 week. The formation of small assemblies with hydrodynamic diameters <30 nm was confirmed by DLS measurements of the reaction mixtures (Fig. 2A). Discoidal particles were observed in TEM images of ASPP1- and Ac-18A-NH2 -POPC systems (Figs. 2 B and Fig. S2). Along with circular particles viewed from the direction of bilayer normal, we observed their rouleaux arranged at intervals of ∼5 nm, corresponding to the thickness of lipid bilayers (Fig. 2B, arrowhead). These images are characteristics of nanodiscs or discoidal lipoproteins although populations of these two orientations on a TEM grid are sensitive to experimental procedures of TEM [23]. In addition, the peptides in the particles formed similar ␣-helix-rich structures as judged from their CD spectra (Fig. 4, see below). These results agree with previous studies on nanodiscs formed by Ac-18A-NH2 and apoA–I [12,24], clearly indicating the formation of ASPP1-POPC nanodiscs. Note that we also added MPAA thiol to the reaction mixture to enhance the intermolecular NCL reaction of ASPP1 [25]. The scattering intensity initially decreased upon the addition of ASPP1 to LUVs even in
the absence of MPAA; however, insoluble aggregates were formed over several hours (data not shown). It is possible that transthioesterification of hydrophobic benzyl mercaptan with the hydrophilic MPAA additive increased the solubility of ASPP1 and the nanodiscs. 3.2. Size distributions of nanodiscs DLS size distributions by volume of Ac-18A-NH2 - and ASPP1POPC nanodiscs are shown in Fig. 2A. With both types of nanodiscs, particle sizes increased with the lipid:peptide ratio, indicating that increased numbers of lipid molecules are involved in the formation of each particle. To confirm this, we examined the sizes and morphologies of the nanodiscs using TEM (Figs. 2 B–D and Fig. S2). All electron micrographs showed unimodal size distributions of discoidal particles, whose average diameter increased from 15 nm to 30 nm as a function of the lipid:peptide ratio of 1–16. Thickness of the discoidal particles remained ∼5 nm even at a lipid:peptide ratio of 16 as judged from the rouleaux structures, suggesting a planar bilayer structure (Fig. S2D). These results demonstrate that the diameter of the ASPP1-POPC nanodiscs is controlled by the lipid:peptide ratio, like other lipid-peptide nanodiscs [12–14]. The large nanodiscs may be used to integrate large membrane protein complexes such as cytochrome c oxidase composed of 26 subunits with a full width of ∼16 nm [26]. Note that 18A and ASPP1 exhibited the single DLS peaks at <3 nm without POPC, suggesting that no large complex were formed by these peptides alone under our
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Fig. 2. Size measurements of the lipid-peptide nanodiscs at 25 ◦ C. POPC-peptide nanodiscs were formed by direct solubilization of the lipid vesicles. Nanodiscs were examined 24 h after addition of the peptides to the POPC vesicles (100 M). (A) DLS size distributions by volume of Ac-18A-NH2 -nanodiscs (upper panel) and ASPP1-nanodiscs (lower panel) determined by DLS at lipid:peptide molar ratios of 2:1, 4:1 and 8:1. (B) Negatively stained TEM images (100 × 100 nm2 ) of ASPP1-POPC mixture. (C) Disc diameter distributions of the ASPP1-POPC nanodiscs for every 100 particles in the TEM images. (D) Average disc diameters of the ASPP1-POPC nanodiscs in the TEM images as a function of the lipid:peptide molar ratio. Error bars represent mean ± SD for 100 particles.
Fig. 3. Self-polymerization of ASPP1. (A) ASPP1- and Ac-18A-NH2 -POPC mixtures with 10 mM MPAA were analyzed by SDS-PAGE. The concentrations of the peptides and POPC were 50 M and 100 M, respectively. (B) Analysis of the population of multimeric ASPP1 after incubations of 1 and 10 days with POPC LUVs. (C) ASPP1-POPC nanodiscs analyzed by silver staining after 3 days incubation. (D) Effect of MPAA concentration on the self-polymerization of ASPP1. ASPP1-POPC nanodiscs were incubated for 3 days with 1, 10, and 100 mM MPAA. (E) Effect on nanodisc formation of ASPP1 polymerization. ASPP1 (50 M) was incubated with or without POPC LUVs (100 M) for 3 days in the presence of 10 mM MPAA. Gels were stained with Coomassie blue except for (C). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
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experimental conditions (data not shown), whereas the previous study showed a formation of Ac-18A-NH2 oligomers in PBS [27].
3.3. Self-polymerization of ASPP1 The degree of polymerization of ASPP1 by NCL reaction was evaluated by SDS-PAGE. We observed a major band corresponding to the monomeric peptide immediately after the addition of ASPP1 to LUVs (Fig. 3A). A second band of lower intensity corresponding to the dimer peptide was also detected. After incubating the nanodiscs for 1 day, we observed the generation of 2–7-mer of ASPP1, whereas no oligomerization of Ac-18A-NH2 was detected, suggesting intermolecular ligations of ASPP1. The population of ASPP1 oligomers was not changed for 10 days (Fig. 3A and B). We could detect a maximum of 12-mer of ASPP1 by silver staining of the gel (Fig. 3C). The population of the multimeric species was not significantly affected by 1, 10, and 100 mM MPAA (Fig. 3D). Moreover, the population was virtually unchanged, regardless of whether ASPP1 was involved in the nanodiscs or not (Fig. 3E).
3.4. Conformational stability of nanodiscs ASPP1 and Ac-18A-NH2 formed ␣-helix-rich structures in the nanodiscs like apoA–I as demonstrated by their CD spectra exhibiting local minima around 208 nm and 222 nm at 25 ◦ C (Fig. 4). We examined the stability of the secondary structures of the polypeptides in the nanodiscs under denaturing conditions such as high temperatures (Fig. 4A–C) and the presence of urea (Fig. 4D–F). Molar ellipticity of Ac-18A-NH2 at 222 nm gradually increased with a rise in temperature from 25 ◦ C to 75 ◦ C, suggesting unfolding of the peptide conformations (Fig. 4A and G). The ellipticity of ASPP1- and apoA–I-POPC nanodiscs also increased, although the shifts were smaller than that of the Ac-18A-NH2 -nanodiscs (Fig. 4B, C, and G). These results suggest that ASPP1 and apoA–I in the nanodiscs have greater tolerance to heat denaturation. Similarly, we evaluated resistance of the peptides and apoA–I to urea denaturation (Fig. 4D–F and H). Surprisingly, no significant changes in the CD spectra for ASPP1 even in the presence of 6 M urea were observed, whereas Ac-18A-NH2 and apoA–I showed increases in ellipticity, indicating ASPP1 is more stable. In addition, the CD spectrum of Ac-18A-NH2 -nanodiscs in the absence of urea at 25 ◦ C (Fig. 4D, purple line) shows smaller helicity than that depicted in Fig. 4A. The difference is due to the 72-h incubation before the measurements shown in Fig. 4D, during which the nanodiscs have partly degraded. However, there was no change in the CD spectra of ASPP1 after 72 h (Figs. 4B and E, purple lines), indicating prolonged stability of the structures. To evaluate the stability of the particles at a temperature range of 25–80 ◦ C, we recorded the right-angle light scattering of the nanodiscs (Fig. 4I). Light scattering intensity increased during the course of heating, which was attributed to a collapse of nanodiscs [28]. ASPP1-POPC nanodiscs collapsed at >70 ◦ C, whereas Ac-18A-NH2 -POPC nanodiscs collapsed at >65 ◦ C. The decrease in the intensities upon cooling suggests the spontaneous re-formation of the peptide nanodiscs. Although apoA–I-POPC nanodiscs showed the highest heat resistance, a further increase in scattering intensity was observed during the cooling process. This could be attributed to the kinetically stable, irreversible deformation of the protein nanodiscs [29,30]. On the other hand, peptide-lipid nanodiscs showed the decrease in scattering intensity during cooling, suggesting the re-formation of the nanodiscs after the deformation, whereas a large hysteresis in the intensity changes implies kinetical stabilization of these nanodiscs.
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3.5. Lipid bilayer environments in the nanodiscs The fluidity of lipid bilayers in the nanodiscs was evaluated by fluorescence anisotropy of DPH (Fig. 5) [31]. The anisotropies in the Ac-18A-NH2 - and ASPP1-POPC nanodiscs at 25 ◦ C were nearly identical to that in POPC LUVs, suggesting similar bilayer fluidities in the liquid crystalline phase. This agrees with previous results showing that acyl chain packing of the Ac-18A-NH2 -POPC nanodiscs were similar to that of LUVs [14]. We previously showed that the acyl chain packing of apoA–I-POPC nanodiscs was dependent on the lipid:protein ratio, whereas Ac-18A-NH2 -POPC nanodiscs exhibited similar packing to POPC vesicles at lipid:peptide ratios of 2 and 4. This discrepancy is caused by the finite length of apoA–I protein. Changes in the number of lipid molecules in the complex induced changes in the shape of the nanodiscs. The planar structures, however, appeared to be maintained in the peptide-lipid nanodiscs by changing the number of peptides fitting to the circumference of the bilayer discs [14].
4. Discussion To stabilize the lipid-peptide nanodiscs, we employed a modified peptide that can self-polymerize by the NCL reaction. This approach has been previously used for the synthesis of collagenlike polymers, resulting in the formation of fibrils that mimic natural collagen with high thermal stabilities [32]. We designed amphiphilic self-polymerizing peptide, ASPP1, which solubilized POPC vesicles over a period of 1 day as well as the parent peptide, Ac-18A-NH2 . In addition, particle sizes, polypeptide secondary structures, and lipid bilayer environments of ASPP1-POPC complex were similar to those of Ac-18A-NH2 -POPC nanodiscs, suggesting that the N-terminal cysteine-glycine residues and C-terminal thioester group have little effects on interactions with POPC. ASPP1 polymerized to generate >10-mer with a mean degree of polymerization of 2.4. The reaction was completed within 1 day and further ligation was not observed. This is probably because of hydrolysis of the thioester group at the C-terminus of ASPP1, which prevented further polymerization of the peptides [25]. Formation of cyclic polypeptides by the intramolecular NCL reaction between the Nand C-terminus may also inhibit polymerization [33]. In addition, the NCL reaction of ASPP1 was not influenced by the lipid bilayers or the formation of nanodiscs in spite of the increased local concentration of peptide and the reduced dimensionality. We have previously shown that the NCL reaction rate of amphiphilic peptides in the presence of POPC vesicles at a lipid:peptide ratio of 4 was approximately 3-fold faster than that in the absence of the lipid vesicles without thiol additives [15]. An excess amount of MPAA additive has been reported to enhance the NCL reaction rate >18-fold faster than without thiols [25]. Therefore, the lipid bilayers may not significantly contribute to the reaction rates under our conditions in the presence of MPAA. Polymerization of amphiphilic peptides in nanodiscs might be achieved by the cross-linking of the peptides without introducing any additional amino acid residues or chemical modifications at the N- and C-terminals of the 18A sequence. ASPP1 has the advantage of neither disrupting the amphiphilic structure of the ␣-helix nor affecting cationic lysine residues, a main target of cross-linking reagents. ASPP1 can also selectively polymerize even in the presence of other molecules such as integrated membrane proteins and chemical groups reactive to cross-linkers. In addition, cysteine side chain can be capped by thiol-reactive compounds like iodoacetamide after formation of ASPP-POPC nanodiscs and ASPP1 polymerization, preventing an undesired disulfide bond formation between ASPP1 and other molecules [16]. ASPP1 was able to polymerize to a maximum of 12-mer, and ASPP1-POPC nanodiscs exhibited increased stability at high
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Fig. 4. Stability of the nanodiscs. CD spectra of Ac-18A-NH2 - (A,D), ASPP1- (B,E) and apoA–I-POPC nanodiscs (C,F) measured while increasing temperature from 25 ◦ C to 75 ◦ C (A–C) or in the presence of urea ranging from 0 M to 6 M at 25 ◦ C (D–F). Changes in the molar ellipticity at 222 nm are plotted as functions of temperature (G) and concentration of urea (H). (I) Right-angle light scattering intensity of 18A-, ASPP1- and apoA–I-nanodiscs monitored at 650 nm during heating (solid lines) and cooling (dotted lines). The lipid:peptide molar ratio was 2:1 for Ac-18A-NH2 - and ASPP1-POPC nanodiscs and 78:1 for apoA–I-POPC nanodiscs. apoA–I nanodiscs were prepared by sodium cholate dialysis, whereas peptide nanodiscs were formed by direct solubilization of the lipid vesicles. (For interpretation of the references to colour in the text, the reader is referred to the web version of this article.)
temperatures or in the presence of denaturant as compared to Ac18A-NH2 - and ApoA–I-POPC nanodiscs. In addition, ASPP1-POPC nanodiscs had longer half-lives. These results are consistent with previous studies. A number of studies investigating apoA–I mimetic peptides revealed that the cooperativity of ␣-helical domains influenced lipid vesicle solubilization and cholesterol efflux from living cells [34]. Anantharamaiah et al. demonstrated that a prolinelinked dimer of 18A, 37pA exhibited higher lipid affinity than 18A [12]. Zhao et al. also reported that multivalent peptide constructs containing 2–4 copies of amphiphilic peptide showed increased half-lives in mice [16]. In addition, most recently Kariyazono et al. showed that proline-linked amphiphilic peptide dimers can stabilize nanodisc structures at high temperatures [35]. The increase in stability appears to be caused by hydrophobic interactions between the amphiphilic helices and lipid bilayers that are enhanced by the ligation of the peptides. In addition, the 18A sequence in ASPP1 has a high membrane affinity and nanodisc-forming activity. We estimated the approximate free energy change of the peptide for
partitioning into the bilayer by forming ␣-helical structures using the following equation [36]: G = −4.62 (±0.15) − 0.46 (±0.03) H (kcal/mol). Here, H is a hydrophobic moment of the amphiphilic peptide [37]. Substituting the H values of 0.467 for ASPP1 and 0.495 for 18A gives G values of −4.83 and −4.85 kcal/mol, respectively. We estimated the mean free energy change of the ligated ASPP1 with a mean degree of polymerization of 2.4 to be −11.6 kcal/mol. In these calculations, we assumed the transition from the unstructured amphiphilic peptides in solution to the ␣-helical structures at the interface of the nanodiscs. In contrast, apoA–I is partially folded in the lipid-free state to sequester the hydrophobic surfaces from the aqueous phase. Therefore, the free energy change of nanodisc formation of apoA–I is expected to be smaller than that of unfolded polypeptides. Indeed, the free energy change of apoA–I for forming nanodiscs with POPC was −8.6 kcal/mol per protein at 25 ◦ C as determined by extrapolation of the experimental Van’t Hoff plot [38]. These free energies might explain why nan-
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Acknowledgement This work was supported in part by Grants-in-Aid for Scientific Research from the Japanese Ministry of Education, Culture, Sports, Science and Technology (nos. 26287098 and 26860020). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.colsurfb.2016.06. 040. Fig. 5. Fluorescence anisotropy of DPH in POPC LUVs, Ac-18A-NH2 - and ASPP1-POPC nanodiscs at 25 ◦ C. The lipid:peptide molar ratio was 2:1 for the nanodiscs. Error bars represent mean + SD (n = 2).
odiscs formed by polymerized ASPP1 are more stable than apoA–I nanodiscs, whereas the Ac-18A-NH2 nanodiscs are less stable. In addition, the mixture of ASPP1 oligomers with different degrees of polymerization may play a role in the stabilization of the nanodiscs. We have previously shown synergistic effects in formation of nanodiscs by amphiphilic peptides with different lengths [15]. Short amphiphilic peptides of 11 amino acid residues did not form nanodiscs with POPC. However, these peptides were integrated into nanodiscs formed by 22-residue amphiphilic peptides, resulting in an increase in the number of the particles. Short peptides may also bind to the hydrophobic patches of the nanodiscs of long peptides to increase formation and stabilization of the nanodiscs. Peptide-lipid complexation also depends on amino acid sequences of amphiphilic ApoA–I mimetic peptides, which have been developed for a potential agent of treating atherosclerosis [34]. For example, an Ac-18A-NH2 mutant peptide 4F, in which two leucine residues of 18A were replaced by phenylalanine, exhibited the enhanced insertion into lipid monolayer, vesicle solubilization, and biological activities [27]. In addition to design of amino acid sequence, our self-polymerizing peptide strategy also provides a potential way for further stabilization of apoA–I mimetic peptidelipid complexes.
5. Conclusion In the present study, we demonstrated that ASPP1 formed nanodiscs with POPC, similar to those formed by 18A peptide or apoA–I protein. The nanodiscs had lipid bilayers in the liquid crystalline phase like those in liposomes. In addition, discoidal structures were spontaneously formed without adding detergent and subsequent dialysis, which is a typical feature of peptide nanodiscs. Our findings indicate that ASPP1-nanodiscs are useful for a variety of applications such as media for solubilizing membrane proteins and as therapeutic agents. The detergent-free formation of nanodiscs was also observed in styrene-maleic acid copolymers [39]. Recently, Dörr et al. reported formation of native nanodiscs by adding the copolymer to E. coli membranes without using detergents [40]. They isolated membrane proteins of the cell membranes with the surrounding lipid molecules. The peptide-based strategy using ASPP1 also has the potential to allow a detergent-free isolation of membrane proteins from living cells. We also showed that the sizes of the ASPP1-POPC nanodiscs were controlled by the number of peptides and lipids in the individual particles. The selfpolymerization of ASPP1 by NCL reaction increased the thermal stability of the assembly. These features may improve the pharmacokinetics of the nanodiscs.
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Colloids and Surfaces B: Biointerfaces journal homepage: www.elsevier.com/locate/colsurfb
Formation of size-controlled, denaturation-resistant lipid nanodiscs by an amphiphilic self-polymerizing peptide Hiroaki Kondo, Keisuke Ikeda ∗ , Minoru Nakano Graduate School of Medicine and Pharmaceutical Sciences, University of Toyama, 2630 Sugitani, Toyama 930-0194, Japan
a r t i c l e
i n f o
Article history: Received 16 March 2016 Received in revised form 22 June 2016 Accepted 22 June 2016 Available online 23 June 2016 Keywords: Nanoparticle Phosphatidylcholine Amphiphilic peptide Self-assembly Native chemical ligation
a b s t r a c t Nanodiscs are discoidal particles with a planar phospholipid bilayer enwrapped by proteins such as apolipoprotein A-I. Nanodiscs have been widely used for analyzing structures and functions of membrane proteins by dispersing them in solution. They are expected to be used as drug carriers and therapeutic agents. Amphiphilic peptides are known to form nanodiscs. However, the lipid-peptide nanodiscs are relatively unstable in solution, making them unsuitable for many applications. Here, we report the synthesis of an amphiphilic self-polymerizing peptide termed ASPP1, which polymerizes by intermolecular native chemical ligation reactions. ASPP1 spontaneously formed nanodiscs when added to phospholipid vesicles without using detergents. The diameter of the planar lipid bilayer in the nanodiscs was controlled by the lipid:peptide molar ratio. ASPP1-nanodiscs exhibited greater stability at high temperatures or in the presence of urea than nanodiscs formed by the non-polymerizing amphiphilic peptide or apolipoprotein A–I. Average and maximal degrees of ASPP1 polymerization were 2.4 and 12, respectively. Self-polymerization of the peptide appears to be responsible for stabilization of the nanodiscs. Our results open a new avenue for the development of nanodisc technology. © 2016 Elsevier B.V. All rights reserved.
1. Introduction Lipid nanodiscs are nanometer-sized, discoidal, supramolecular complexes with a patch of planar phospholipid bilayer enwrapped by amphiphilic, ␣-helical polypeptides [1,2]. Nanodiscs dispersed in water are stabilized by hydrophobic interactions, where hydrophobic acyl chains of lipids and nonpolar faces of ␣helices are sequestered from the aqueous solution. Apolipoprotein A–I (apoA–I), the main component of high-density lipoproteins (HDLs), has been reported to form discoidal particles with lipids [3,4]. Cholesterol is incorporated into the discoidal HDLs and metabolized to cholesteryl ester by lectin-cholesterol acyltransferase, resulting in the formation of spherical HDLs which transport excess cholesterol from peripheral tissues to the liver [5]. In addition to the biological significance of discoidal HDLs in cholesterol homeostasis, nanodiscs have attracted considerable interest for their potential use in nanomedicines and drug delivery systems, and as model membrane systems for studying membrane proteins [6–10]. Indeed, a number of membrane proteins, such as bacteriorhodopsin, G-protein coupled receptors, and epidermal growth
∗ Corresponding author. E-mail address: [email protected] (K. Ikeda). http://dx.doi.org/10.1016/j.colsurfb.2016.06.040 0927-7765/© 2016 Elsevier B.V. All rights reserved.
factor receptor, have been assembled into nanodiscs that mimic native lipid bilayer environments. Various biophysical methods and spectroscopic techniques can be used to analyze structures and functions of the solubilized proteins. For a wide range of applications, it is essential to have control over the physicochemical parameters of nanodiscs such as size, stability, lipid composition, and chemical modification. For example, Denisov et al. designed extended and truncated versions of apoA–I, termed membrane scaffold proteins (MSPs), to customize the sizes of the nanodiscs in the range of 9.5–12.8 nm [2]. Hagn et al. designed further truncated versions of MSPs to create smaller nanodiscs with a diameter of ∼6 nm, whose short rotational correlation times were suitable for solution-state NMR studies of integrated membrane proteins [11]. They determined the structure and dynamics of the -barrel protein, OmpX, integrated in the small nanodiscs. Similar to proteins, amphiphilic ␣-helical peptides have also been reported to form nanodiscs in cooperation with lipids [12–14]. Lipid-peptide nanodiscs have several advantages with respect to nanodisc assembly and physicochemical parameters. Nanodiscs can spontaneously form simply by mixing the peptides and lipid suspension (e.g. liposomes) in buffer without the aid of detergents, which may disrupt the structures and functions of membrane proteins. Their sizes can be easily controlled by varying the lipid:peptide ratio when the components are mixed in buffer.
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Table 1 Peptide sequences.
2.2. Peptide synthesis
Peptide
Amino acid sequence
Ac-18A-NH2 ASPP1
Ac-DWLKAFYDKVAEKLKEAF-CONH2 NH2 -CGDWLKAFYDKVAEKLKEAF-COSBna
a
COSBn = benzyl thioester.
Moreover, large-scale synthesis and chemical modifications of the peptides are feasible. Despite these promising properties of lipidpeptide nanodiscs, they are unstable as evidenced by the fact that short amphiphilic peptides of 11 amino acid residues cannot form nanodiscs [15]. In addition, Imura et al. showed that the 22-residue nanodisc-forming peptide derived from human apoA–I (220–241) did not form the assembly below the critical association concentration of ∼10−5 M [13]. These instabilities are probably because relatively short peptides have weaker hydrophobic interactions with the edges of nanodiscs than the linked amphiphilic ␣-helices of apoA–I or MSPs. Stable nanoparticles under physiological conditions are required for in vivo therapeutic applications. Recently, Zhao et al. have reported that branched, multivalent constructs of amphiphilic peptides formed stable nanodiscs that were resistant to enzymatic digestion and had long half-lives in mice [16]. This study raises the possibility that the further chemical ligation of amphiphilic peptides would increase the stability of peptidelipid nanodiscs. In addition, highly stable lipid-peptide nanodiscs are required for studying integrated membrane proteins requiring long experimental period and exposure to high temperatures and denaturing agents. Here, we report synthesis of size-controlled, denaturationresistant lipid nanodiscs using an amphiphilic self-polymerizing peptide termed ASPP1 that can withstand denaturing conditions. The peptide incorporates the apoA–I mimetic sequence 18A, which was originally designed by Anantharamaiah et al. [12]. The sequence of 18A has properties similar to the amphiphilic helices of apoA–I. N- and C-terminal blocked 18A (Ac-18A-NH2 , Table 1) induced a larger increase in ␣-helical content in complex with phosphatidylcholine than unblocked 18A and formed nanodiscs similar to apoA–I nanodiscs [17]. As 18A does not have residues at the junctions of the tandem helices, we attached cysteine-glycine residues at the N-terminus as a flexible linker. We also modified the C-terminus with a benzyl thioester group, which causes a native chemical ligation (NCL) reaction with the free cysteine residue at the N-terminus to form an amide bond [18]. We hypothesized that self-polymerization of ASPP1 would occur, which would facilitate hydrophobic interactions between the interlinked ␣-helices and lipid bilayers, thereby increasing the conformational stability of the nanodiscs. Formation of nanodiscs by ASPP1 with a phospholipid, sizes of the nanodiscs, and intermolecular ligation of ASPP1 were investigated. We also demonstrate the ability of the ASPP1 nanodiscs to resist unfolding of the peptide secondary structures and deformation of the assembly triggered by high temperatures or by denaturant.
2. Materials and methods
Ac-18A-NH2 was synthesized using Fmoc-based solid-phase synthesis and cleaved from the resin with trifluoroacetic acid/water/triisopropylsilane (95/2.5/2.5) for 3 h. For synthesis of ASPP1 with C-terminal benzyl thioester, the diaminobenzoyl linker-attached Rink amide resin developed by Dawson et al. was employed [20]. Peptide chains were extended on Dbz AM resin (Merck Millipore) by Fmoc-based chemistry. An N-terminal cysteine residue was introduced using Boc-Cys(Trt)-OH/N,N’diisopropylcarbodiimide/1-hydroxybenzotriazole. A solution of p-nitrophenylchloroformate (5 eq) in dichloromethane was added to the resin and the mixture was stirred for 1 h. After washing the resin with DCM and DMF, 0.5 M N,N-diisopropylethylamine in DMF was added and the mixture was stirred for 30 min. After cleavage of peptides from the resin with trifluoroacetic acid/water/triisopropylsilane/1,2-ethanedithiol (92.5/2.5/2.5/2.5), the peptides were dissolved in DMSO containing 620 mM benzylmercaptan and left overnight. The peptides were purified by reverse phase high-performance liquid chromatography (RP-HPLC) using a C18 HPLC column with a gradient of water/acetonitrile containing 0.1% trifluoroacetic acid. Peptides were characterized by HPLC and MALDI-TOF MS (Autoflex-T1, Bruker Daltonics). We determined the purity of ASPP1 to be >90% by HPLC and obtained the expected molecular mass (Fig. S1). Ac-18A-NH2 lyophilized powder was dissolved in water, and ASPP1 lyophilized powder was dissolved in a 1 mM HCl solution and the concentrations were determined by measuring absorbance at 280 nm. 2.3. Liposomes Large unilamellar vesicles (LUVs) were prepared as follows [14]: an aliquot of POPC in chloroform-methanol was transferred to a round-bottom flask and the solvent was removed using a rotary evaporator. Thereafter, the sample was dried overnight under vacuum. Buffer (20 mM MOPS/1 mM EDTA, pH 7.0) was added to the remaining lipid film. The lipid suspension was freeze-thawed 5 times and extruded through a 100 nm pore filter. The size of LUVs was determined using a DelsaMax Core dynamic light scattering (DLS) instrument (Beckman Coulter). The concentration of lipid was determined by an enzymatic assay kit for choline (Wako). 2.4. Nanodiscs Preparation of POPC-apoA–I nanodiscs was based on the sodium cholate dialysis method [21]. Sodium deoxycholate was added to the lipid suspension in MOPS buffer at a POPC:cholate molar ratio of 1:5. Then, apoA–I was added at a molar ratio of POPC:apoA–I of 78:1. After incubation for 24 h at 4 ◦ C, nanodiscs were obtained by dialysis against MOPS buffer for 2 days. The lipid-peptide nanodiscs were obtained by mixing the peptides and POPC LUVs (100 M) at molar ratios of 1:1–1:16 in buffer containing 10 mM 4mercaptphenylacetic acid (MPAA) at 25 ◦ C. Solubilization of LUVs and nanodisc formation was confirmed by right-angle light scattering at a wavelength of 650 nm using an F-4500 fluorescence spectrometer (Hitachi). The sizes of nanodiscs were determined using a DelsaMax Core dynamic light scattering (DLS) instrument (Beckman Coulter) at 20 ◦ C.
2.1. Materials 2.5. Transmission electron microscopy (TEM) 1-Palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) was purchased from Avanti Polar Lipids. 1,6-Diphenyl-1,3,5-hexatriene (DPH) was obtained from Molecular Probes. ApoA–I was isolated from pig plasma using procedures described previously [19]. The purity of apoA–I was determined to be >95% by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE).
A glow-discharged Cu grid coated with thin carbon film (300 mesh) was placed on a droplet of the sample for 5 min. Then, excess sample solution on the grid was removed using filter paper. Next, the grid was placed on a droplet of 2% phosphotungstic acid for 30 s and dried after removal of the excess solution. TEM images
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of nanodiscs were obtained using a JEM-1400 transmission electron microscope (JEOL) operated at 80 keV. Diameters of nanodiscs were measured by PhotoRuler software employing images of 100 particles (Fig. S2). 2.6. SDS-PAGE SDS-PAGE was performed as follows [22]: sample solution was mixed with 3 vol of sample buffer (150 mM Tris/HCl, pH 7.0, 12% SDS (wt/vol), 6% mercaptoethanol (vol/vol), 30% glycerol (wt/vol), 0.05% Coomassie blue G-250) and loaded in 16% polyacrylamide/6 M urea gel. Gels were stained with 0.025% Coomassie blue in 10% acetic acid. The degree of polymerization of ASPP1 was assessed by measuring band densities using ImageJ software. For silver staining, the gel after Coomassie blue staining was washed with 50% methanol, 50 mM ammonium hydrogen carbonate. Silver staining was carried out using the 2D-SILVER STAIN-II kit (Cosmo Bio). 2.7. Circular dichroism (CD) spectroscopy Due to the high absorbance of MPAA, MPAA in solution was removed by dialysis against MOPS-NaOH buffer without MPAA for 48 h. For carrying out measurements in the presence of urea, samples were equilibrated with urea for 72 h at 4 ◦ C. CD spectra were acquired at 25 ◦ C using a J-805 spectrometer (JASCO) with a 1 mm path length quartz cell. Eight scans were averaged for each sample. 2.8. Fluorescence anisotropy A small volume of a methanol solution (final 1% (v/v) methanol) of DPH was added to either LUVs or nanodiscs and incubated at 25 ◦ C for 1 h to incorporate 0.5 mol% DPH into the lipid bilayers. Fluorescence anisotropy of DPH was measured using an F-4500 fluorescence spectrometer with polarizers placed in the excitation and emission paths. Excitation and emission wavelengths were set at 360 and 430 nm, respectively.
Fig. 1. Formation of lipid-peptide nanodiscs at 25 ◦ C. Changes in the right-angle light scattering intensities of POPC LUVs upon addition of Ac-18A-NH2 (A) or ASPP1 (B) were monitored at 650 nm. The POPC:peptide molar ratios were set to 2:1, 4:1 and 8:1 at a POPC concentration of 100 M. Scattering intensity, I(t), was normalized by the intensity of the POPC LUVs before the additions of peptides, I(0).
3. Results 3.1. Formation of lipid-peptide nanodiscs ASPP1 was added to POPC LUVs having diameters of ∼100 nm at 25 ◦ C. We observed a decrease in the right-angle light scattering intensities of the LUVs in the presence of the ASPP1 as well as Ac-18A-NH2 (Fig. 1), suggesting that the vesicles were solubilized into small particles [14]. The decrease in intensities occurred over a period of 1 day, and no further significant changes were observed over a period of 1 week. The formation of small assemblies with hydrodynamic diameters <30 nm was confirmed by DLS measurements of the reaction mixtures (Fig. 2A). Discoidal particles were observed in TEM images of ASPP1- and Ac-18A-NH2 -POPC systems (Figs. 2 B and Fig. S2). Along with circular particles viewed from the direction of bilayer normal, we observed their rouleaux arranged at intervals of ∼5 nm, corresponding to the thickness of lipid bilayers (Fig. 2B, arrowhead). These images are characteristics of nanodiscs or discoidal lipoproteins although populations of these two orientations on a TEM grid are sensitive to experimental procedures of TEM [23]. In addition, the peptides in the particles formed similar ␣-helix-rich structures as judged from their CD spectra (Fig. 4, see below). These results agree with previous studies on nanodiscs formed by Ac-18A-NH2 and apoA–I [12,24], clearly indicating the formation of ASPP1-POPC nanodiscs. Note that we also added MPAA thiol to the reaction mixture to enhance the intermolecular NCL reaction of ASPP1 [25]. The scattering intensity initially decreased upon the addition of ASPP1 to LUVs even in
the absence of MPAA; however, insoluble aggregates were formed over several hours (data not shown). It is possible that transthioesterification of hydrophobic benzyl mercaptan with the hydrophilic MPAA additive increased the solubility of ASPP1 and the nanodiscs. 3.2. Size distributions of nanodiscs DLS size distributions by volume of Ac-18A-NH2 - and ASPP1POPC nanodiscs are shown in Fig. 2A. With both types of nanodiscs, particle sizes increased with the lipid:peptide ratio, indicating that increased numbers of lipid molecules are involved in the formation of each particle. To confirm this, we examined the sizes and morphologies of the nanodiscs using TEM (Figs. 2 B–D and Fig. S2). All electron micrographs showed unimodal size distributions of discoidal particles, whose average diameter increased from 15 nm to 30 nm as a function of the lipid:peptide ratio of 1–16. Thickness of the discoidal particles remained ∼5 nm even at a lipid:peptide ratio of 16 as judged from the rouleaux structures, suggesting a planar bilayer structure (Fig. S2D). These results demonstrate that the diameter of the ASPP1-POPC nanodiscs is controlled by the lipid:peptide ratio, like other lipid-peptide nanodiscs [12–14]. The large nanodiscs may be used to integrate large membrane protein complexes such as cytochrome c oxidase composed of 26 subunits with a full width of ∼16 nm [26]. Note that 18A and ASPP1 exhibited the single DLS peaks at <3 nm without POPC, suggesting that no large complex were formed by these peptides alone under our
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Fig. 2. Size measurements of the lipid-peptide nanodiscs at 25 ◦ C. POPC-peptide nanodiscs were formed by direct solubilization of the lipid vesicles. Nanodiscs were examined 24 h after addition of the peptides to the POPC vesicles (100 M). (A) DLS size distributions by volume of Ac-18A-NH2 -nanodiscs (upper panel) and ASPP1-nanodiscs (lower panel) determined by DLS at lipid:peptide molar ratios of 2:1, 4:1 and 8:1. (B) Negatively stained TEM images (100 × 100 nm2 ) of ASPP1-POPC mixture. (C) Disc diameter distributions of the ASPP1-POPC nanodiscs for every 100 particles in the TEM images. (D) Average disc diameters of the ASPP1-POPC nanodiscs in the TEM images as a function of the lipid:peptide molar ratio. Error bars represent mean ± SD for 100 particles.
Fig. 3. Self-polymerization of ASPP1. (A) ASPP1- and Ac-18A-NH2 -POPC mixtures with 10 mM MPAA were analyzed by SDS-PAGE. The concentrations of the peptides and POPC were 50 M and 100 M, respectively. (B) Analysis of the population of multimeric ASPP1 after incubations of 1 and 10 days with POPC LUVs. (C) ASPP1-POPC nanodiscs analyzed by silver staining after 3 days incubation. (D) Effect of MPAA concentration on the self-polymerization of ASPP1. ASPP1-POPC nanodiscs were incubated for 3 days with 1, 10, and 100 mM MPAA. (E) Effect on nanodisc formation of ASPP1 polymerization. ASPP1 (50 M) was incubated with or without POPC LUVs (100 M) for 3 days in the presence of 10 mM MPAA. Gels were stained with Coomassie blue except for (C). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
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experimental conditions (data not shown), whereas the previous study showed a formation of Ac-18A-NH2 oligomers in PBS [27].
3.3. Self-polymerization of ASPP1 The degree of polymerization of ASPP1 by NCL reaction was evaluated by SDS-PAGE. We observed a major band corresponding to the monomeric peptide immediately after the addition of ASPP1 to LUVs (Fig. 3A). A second band of lower intensity corresponding to the dimer peptide was also detected. After incubating the nanodiscs for 1 day, we observed the generation of 2–7-mer of ASPP1, whereas no oligomerization of Ac-18A-NH2 was detected, suggesting intermolecular ligations of ASPP1. The population of ASPP1 oligomers was not changed for 10 days (Fig. 3A and B). We could detect a maximum of 12-mer of ASPP1 by silver staining of the gel (Fig. 3C). The population of the multimeric species was not significantly affected by 1, 10, and 100 mM MPAA (Fig. 3D). Moreover, the population was virtually unchanged, regardless of whether ASPP1 was involved in the nanodiscs or not (Fig. 3E).
3.4. Conformational stability of nanodiscs ASPP1 and Ac-18A-NH2 formed ␣-helix-rich structures in the nanodiscs like apoA–I as demonstrated by their CD spectra exhibiting local minima around 208 nm and 222 nm at 25 ◦ C (Fig. 4). We examined the stability of the secondary structures of the polypeptides in the nanodiscs under denaturing conditions such as high temperatures (Fig. 4A–C) and the presence of urea (Fig. 4D–F). Molar ellipticity of Ac-18A-NH2 at 222 nm gradually increased with a rise in temperature from 25 ◦ C to 75 ◦ C, suggesting unfolding of the peptide conformations (Fig. 4A and G). The ellipticity of ASPP1- and apoA–I-POPC nanodiscs also increased, although the shifts were smaller than that of the Ac-18A-NH2 -nanodiscs (Fig. 4B, C, and G). These results suggest that ASPP1 and apoA–I in the nanodiscs have greater tolerance to heat denaturation. Similarly, we evaluated resistance of the peptides and apoA–I to urea denaturation (Fig. 4D–F and H). Surprisingly, no significant changes in the CD spectra for ASPP1 even in the presence of 6 M urea were observed, whereas Ac-18A-NH2 and apoA–I showed increases in ellipticity, indicating ASPP1 is more stable. In addition, the CD spectrum of Ac-18A-NH2 -nanodiscs in the absence of urea at 25 ◦ C (Fig. 4D, purple line) shows smaller helicity than that depicted in Fig. 4A. The difference is due to the 72-h incubation before the measurements shown in Fig. 4D, during which the nanodiscs have partly degraded. However, there was no change in the CD spectra of ASPP1 after 72 h (Figs. 4B and E, purple lines), indicating prolonged stability of the structures. To evaluate the stability of the particles at a temperature range of 25–80 ◦ C, we recorded the right-angle light scattering of the nanodiscs (Fig. 4I). Light scattering intensity increased during the course of heating, which was attributed to a collapse of nanodiscs [28]. ASPP1-POPC nanodiscs collapsed at >70 ◦ C, whereas Ac-18A-NH2 -POPC nanodiscs collapsed at >65 ◦ C. The decrease in the intensities upon cooling suggests the spontaneous re-formation of the peptide nanodiscs. Although apoA–I-POPC nanodiscs showed the highest heat resistance, a further increase in scattering intensity was observed during the cooling process. This could be attributed to the kinetically stable, irreversible deformation of the protein nanodiscs [29,30]. On the other hand, peptide-lipid nanodiscs showed the decrease in scattering intensity during cooling, suggesting the re-formation of the nanodiscs after the deformation, whereas a large hysteresis in the intensity changes implies kinetical stabilization of these nanodiscs.
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3.5. Lipid bilayer environments in the nanodiscs The fluidity of lipid bilayers in the nanodiscs was evaluated by fluorescence anisotropy of DPH (Fig. 5) [31]. The anisotropies in the Ac-18A-NH2 - and ASPP1-POPC nanodiscs at 25 ◦ C were nearly identical to that in POPC LUVs, suggesting similar bilayer fluidities in the liquid crystalline phase. This agrees with previous results showing that acyl chain packing of the Ac-18A-NH2 -POPC nanodiscs were similar to that of LUVs [14]. We previously showed that the acyl chain packing of apoA–I-POPC nanodiscs was dependent on the lipid:protein ratio, whereas Ac-18A-NH2 -POPC nanodiscs exhibited similar packing to POPC vesicles at lipid:peptide ratios of 2 and 4. This discrepancy is caused by the finite length of apoA–I protein. Changes in the number of lipid molecules in the complex induced changes in the shape of the nanodiscs. The planar structures, however, appeared to be maintained in the peptide-lipid nanodiscs by changing the number of peptides fitting to the circumference of the bilayer discs [14].
4. Discussion To stabilize the lipid-peptide nanodiscs, we employed a modified peptide that can self-polymerize by the NCL reaction. This approach has been previously used for the synthesis of collagenlike polymers, resulting in the formation of fibrils that mimic natural collagen with high thermal stabilities [32]. We designed amphiphilic self-polymerizing peptide, ASPP1, which solubilized POPC vesicles over a period of 1 day as well as the parent peptide, Ac-18A-NH2 . In addition, particle sizes, polypeptide secondary structures, and lipid bilayer environments of ASPP1-POPC complex were similar to those of Ac-18A-NH2 -POPC nanodiscs, suggesting that the N-terminal cysteine-glycine residues and C-terminal thioester group have little effects on interactions with POPC. ASPP1 polymerized to generate >10-mer with a mean degree of polymerization of 2.4. The reaction was completed within 1 day and further ligation was not observed. This is probably because of hydrolysis of the thioester group at the C-terminus of ASPP1, which prevented further polymerization of the peptides [25]. Formation of cyclic polypeptides by the intramolecular NCL reaction between the Nand C-terminus may also inhibit polymerization [33]. In addition, the NCL reaction of ASPP1 was not influenced by the lipid bilayers or the formation of nanodiscs in spite of the increased local concentration of peptide and the reduced dimensionality. We have previously shown that the NCL reaction rate of amphiphilic peptides in the presence of POPC vesicles at a lipid:peptide ratio of 4 was approximately 3-fold faster than that in the absence of the lipid vesicles without thiol additives [15]. An excess amount of MPAA additive has been reported to enhance the NCL reaction rate >18-fold faster than without thiols [25]. Therefore, the lipid bilayers may not significantly contribute to the reaction rates under our conditions in the presence of MPAA. Polymerization of amphiphilic peptides in nanodiscs might be achieved by the cross-linking of the peptides without introducing any additional amino acid residues or chemical modifications at the N- and C-terminals of the 18A sequence. ASPP1 has the advantage of neither disrupting the amphiphilic structure of the ␣-helix nor affecting cationic lysine residues, a main target of cross-linking reagents. ASPP1 can also selectively polymerize even in the presence of other molecules such as integrated membrane proteins and chemical groups reactive to cross-linkers. In addition, cysteine side chain can be capped by thiol-reactive compounds like iodoacetamide after formation of ASPP-POPC nanodiscs and ASPP1 polymerization, preventing an undesired disulfide bond formation between ASPP1 and other molecules [16]. ASPP1 was able to polymerize to a maximum of 12-mer, and ASPP1-POPC nanodiscs exhibited increased stability at high
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Fig. 4. Stability of the nanodiscs. CD spectra of Ac-18A-NH2 - (A,D), ASPP1- (B,E) and apoA–I-POPC nanodiscs (C,F) measured while increasing temperature from 25 ◦ C to 75 ◦ C (A–C) or in the presence of urea ranging from 0 M to 6 M at 25 ◦ C (D–F). Changes in the molar ellipticity at 222 nm are plotted as functions of temperature (G) and concentration of urea (H). (I) Right-angle light scattering intensity of 18A-, ASPP1- and apoA–I-nanodiscs monitored at 650 nm during heating (solid lines) and cooling (dotted lines). The lipid:peptide molar ratio was 2:1 for Ac-18A-NH2 - and ASPP1-POPC nanodiscs and 78:1 for apoA–I-POPC nanodiscs. apoA–I nanodiscs were prepared by sodium cholate dialysis, whereas peptide nanodiscs were formed by direct solubilization of the lipid vesicles. (For interpretation of the references to colour in the text, the reader is referred to the web version of this article.)
temperatures or in the presence of denaturant as compared to Ac18A-NH2 - and ApoA–I-POPC nanodiscs. In addition, ASPP1-POPC nanodiscs had longer half-lives. These results are consistent with previous studies. A number of studies investigating apoA–I mimetic peptides revealed that the cooperativity of ␣-helical domains influenced lipid vesicle solubilization and cholesterol efflux from living cells [34]. Anantharamaiah et al. demonstrated that a prolinelinked dimer of 18A, 37pA exhibited higher lipid affinity than 18A [12]. Zhao et al. also reported that multivalent peptide constructs containing 2–4 copies of amphiphilic peptide showed increased half-lives in mice [16]. In addition, most recently Kariyazono et al. showed that proline-linked amphiphilic peptide dimers can stabilize nanodisc structures at high temperatures [35]. The increase in stability appears to be caused by hydrophobic interactions between the amphiphilic helices and lipid bilayers that are enhanced by the ligation of the peptides. In addition, the 18A sequence in ASPP1 has a high membrane affinity and nanodisc-forming activity. We estimated the approximate free energy change of the peptide for
partitioning into the bilayer by forming ␣-helical structures using the following equation [36]: G = −4.62 (±0.15) − 0.46 (±0.03) H (kcal/mol). Here, H is a hydrophobic moment of the amphiphilic peptide [37]. Substituting the H values of 0.467 for ASPP1 and 0.495 for 18A gives G values of −4.83 and −4.85 kcal/mol, respectively. We estimated the mean free energy change of the ligated ASPP1 with a mean degree of polymerization of 2.4 to be −11.6 kcal/mol. In these calculations, we assumed the transition from the unstructured amphiphilic peptides in solution to the ␣-helical structures at the interface of the nanodiscs. In contrast, apoA–I is partially folded in the lipid-free state to sequester the hydrophobic surfaces from the aqueous phase. Therefore, the free energy change of nanodisc formation of apoA–I is expected to be smaller than that of unfolded polypeptides. Indeed, the free energy change of apoA–I for forming nanodiscs with POPC was −8.6 kcal/mol per protein at 25 ◦ C as determined by extrapolation of the experimental Van’t Hoff plot [38]. These free energies might explain why nan-
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Acknowledgement This work was supported in part by Grants-in-Aid for Scientific Research from the Japanese Ministry of Education, Culture, Sports, Science and Technology (nos. 26287098 and 26860020). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.colsurfb.2016.06. 040. Fig. 5. Fluorescence anisotropy of DPH in POPC LUVs, Ac-18A-NH2 - and ASPP1-POPC nanodiscs at 25 ◦ C. The lipid:peptide molar ratio was 2:1 for the nanodiscs. Error bars represent mean + SD (n = 2).
odiscs formed by polymerized ASPP1 are more stable than apoA–I nanodiscs, whereas the Ac-18A-NH2 nanodiscs are less stable. In addition, the mixture of ASPP1 oligomers with different degrees of polymerization may play a role in the stabilization of the nanodiscs. We have previously shown synergistic effects in formation of nanodiscs by amphiphilic peptides with different lengths [15]. Short amphiphilic peptides of 11 amino acid residues did not form nanodiscs with POPC. However, these peptides were integrated into nanodiscs formed by 22-residue amphiphilic peptides, resulting in an increase in the number of the particles. Short peptides may also bind to the hydrophobic patches of the nanodiscs of long peptides to increase formation and stabilization of the nanodiscs. Peptide-lipid complexation also depends on amino acid sequences of amphiphilic ApoA–I mimetic peptides, which have been developed for a potential agent of treating atherosclerosis [34]. For example, an Ac-18A-NH2 mutant peptide 4F, in which two leucine residues of 18A were replaced by phenylalanine, exhibited the enhanced insertion into lipid monolayer, vesicle solubilization, and biological activities [27]. In addition to design of amino acid sequence, our self-polymerizing peptide strategy also provides a potential way for further stabilization of apoA–I mimetic peptidelipid complexes.
5. Conclusion In the present study, we demonstrated that ASPP1 formed nanodiscs with POPC, similar to those formed by 18A peptide or apoA–I protein. The nanodiscs had lipid bilayers in the liquid crystalline phase like those in liposomes. In addition, discoidal structures were spontaneously formed without adding detergent and subsequent dialysis, which is a typical feature of peptide nanodiscs. Our findings indicate that ASPP1-nanodiscs are useful for a variety of applications such as media for solubilizing membrane proteins and as therapeutic agents. The detergent-free formation of nanodiscs was also observed in styrene-maleic acid copolymers [39]. Recently, Dörr et al. reported formation of native nanodiscs by adding the copolymer to E. coli membranes without using detergents [40]. They isolated membrane proteins of the cell membranes with the surrounding lipid molecules. The peptide-based strategy using ASPP1 also has the potential to allow a detergent-free isolation of membrane proteins from living cells. We also showed that the sizes of the ASPP1-POPC nanodiscs were controlled by the number of peptides and lipids in the individual particles. The selfpolymerization of ASPP1 by NCL reaction increased the thermal stability of the assembly. These features may improve the pharmacokinetics of the nanodiscs.
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