Cell-Penetrating Cross-Î² Peptide Assemblies ... - ACS Publications

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Cell-penetrating cross-# peptide assemblies with controlled biodegradable properties Sanghun Han, Mun-kyung Lee, and Yong-beom Lim Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.6b01153 • Publication Date (Web): 06 Dec 2016 Downloaded from http://pubs.acs.org on December 7, 2016

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Cell-penetrating cross-β peptide assemblies with controlled biodegradable properties Sanghun Han,‡ Mun-kyung Lee,†,‡ and Yong-beom Lim* Department of Materials Science & Engineering, Yonsei University, Seoul 03722, Korea KEYWORDS Depsipeptide, self-assembly, controllable degradation

ABSTRACT

Although self-assembled peptide nanostructures (SPNs) have shown potential as promising biomaterials, there is a potential problem associated with the extremely slow hydrolysis rate of amide bonds. Here, we report the development of cell-penetrating cross-β SPNs with a controllable biodegradation rate. The designed self-assembling β-sheet peptide incorporating a hydrolyzable ester bond (self-assembling depsipeptide; SADP) can be assembled into bilayer βsandwich one-dimensional (1D) fibers similarly to conventional β-sheet peptides. The rate of hydrolysis can be controlled by the pH, temperature, and structural characteristics of the ester unit. The 1D fiber of the SADP transforms into vesicle-like 3D structures when the hydrophilic cell-penetrating peptide segment is attached to the SADP segment. Efficient cell internalization

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of the 3D nanostructures was observed, and we verified the intracellular degradation and disassembly of the biodegradable nanostructures. This study illustrates the potential of biodegradable cross-β SPNs and provides a valuable toolkit that can be used with selfassembling peptides.

INTRODUCTION The intracellular delivery of therapeutics such as drugs and genes can be assisted by the use of nanomaterials. In the fields of drug/gene delivery systems, biodegradable polymers have long been used for the controlled release of payloads, the reduction of cytotoxicity, and efficient renal excretion.1,2 Although the peptide bond is considered biodegradable, its hydrolysis is extremely slow under normal conditions in the absence of proteases, with half-lives ranging from several to hundreds of years.3,4 For this reason, the polymers considered to have biodegradable linkages are polyesters, poly(orthoester)s, poly(phosphoester)s, and poly(phosphazene)s.1,2 In particular, polyesters have been extensively used as biodegradable drug and gene delivery carriers. Self-assembled peptide nanostructures (SPNs) have become a topic of considerable research interest because of their potential in various bioapplications. Generally, SPNs are considered biodegradable and biocompatible. As noted above, the term biodegradable for peptide-based materials may need to be reconsidered because of the extremely slow rate of peptide bond hydrolysis. The typical assembly modes of peptide nanostructures include hydrophobic interactions, π−π stacking interactions, α-helical bundle formation, and β-sheet interactions. Among these, the β-sheet interaction is one of the most widely used interaction modes in the development of biomaterials based on SPNs. In fact, nano-assemblies formed by β-sheetforming peptides share a structural similarity to β-amyloid fibrils, which are characterized by


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their cross-β structures.5,6 Despite their advantage of facile aggregation in aqueous solutions, one potential problem with amyloid-like β-sheet fibers is their resistance to proteolytic degradation.7,8 Thus, the degradation of cross-β SPNs may be exacerbated by their slow hydrolysis rate and also by their resistance to proteolysis. Here, we report the development of biodegradable cell delivery materials based on cross-β peptide assemblies. We first designed self-assembling depsipeptides (SADPs) that contain one ester bond in the middle of a β-sheet-forming peptide and compared their self-assembly behavior and degradation kinetics with those of the normal peptide (Figure 1a). A peptide in which one or more of the peptide bonds have been replaced by an ester group is called a depsipeptide.9-11 Next, we designed a self-assembling depsipeptide that has the configuration of a block copolypeptide, in which one block has the function of cross-β assembly and the other block has the cell penetration function. There have been few efforts to assess the possibility of using depsipeptides in β-sheet-mediated self-assembly.12-14 However, because of the absence of hydrogen bonding in the ester bond and the inefficient β-sheet formation propensity of the other residues, the reported depsipeptides display a weak self-association propensity, with working concentrations far higher than those that are biologically relevant.12-14 In this report, the development of bioactive SADPs with the configuration of block copolypeptides was possible because we were able to fabricate a depsipeptide with a strong self-association propensity.

EXPERIMENTAL SECTION Materials


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Rink amide MBHA resin LL was purchased from Novabiochem (Germany). All Fmocprotected amino acids were purchased from Novabiochem and Anaspec (USA). Fluorescein-5maleimide was purchased from Anaspec. N-methyl-2-pyrrolidone (NMP) was purchased from Daejung Chemicals & Metals (South Korea). 5(6)-Carboxytetramethylrhodamine and other solvents were purchased from Sigma-Aldrich (USA).

Peptide synthesis and dye-labeling The peptides were synthesized on Rink amide MBHA resin LL using standard Fmoc protocols. Typical amide couplings were performed by the combination of manual and automated peptide synthesis (TributeTM peptide synthesizer; Protein technologies, USA) protocols, and esterification reactions were performed manually. For 25 µmol of resin, glycolic acid (9.5 mg, 5 eq.) or lactic acid (11.3 mg, 5 eq.) was coupled to the peptide using HATU (42.8 mg, 4.5 eq.), HOBt (17.2 mg, 4.5 eq.) and DIPEA (43.5 µL, 10 eq.) in NMP, the mixture was shaken overnight. Then, the hydroxyl-exposed peptide was mixed with Fmoc-Trp(Boc)-OH (65.8 mg, 5 eq.), diisopropylcarbodiimide (DIC) (58 µL, 15 eq.), p-toluenesulfonic acid (PTSA) (4.8 mg, 1 eq.) and 4-dimethylaminopyridine (DMAP) (3 mg, 1 eq.) in DMF and shaken for 2 days for esterification. The subsequent amino acids were coupled to the peptide using standard protocols. The fully synthesized peptides were cleaved using reagent K (trifluoroacetic acid: 1,2ethanedithiol: thioanisole; 95: 2.5: 2.5). For tetramethylrhodamine (TAMRA) labeling, 5(6)carboxytetramethylrhodamine (27.5 mg, 2.56 eq.) was attached to the N-terminal amine of the peptide using HATU (21.9 mg, 2.3 eq.) and DIPEA (26 µL, 6 eq.) in NMP. For fluoresceinlabeling, fluorescein-5-maleimide (10.7 mg, 5 eq.) was mixed with the fully cleaved peptide (5


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µmol) in methanol to be coupled to the thiol group of the C-terminal cysteine via maleimidethiol coupling chemistry. Detailed synthetic schemes are shown in Figures S2 and S3. All peptides were purified by reverse-phase HPLC (water-acetonitrile with 0.1% TFA). Concentrations were determined spectrophotometrically using a molar extinction coefficient of tryptophan (5,500 M-1cm-1) for short β-sheet peptides at 280 nm or TAMRA (80,000 M-1cm-1) at 543 nm for dye-labeled peptides. Yield: 1p (7%), 1e-G (3%), 1e-L (2%), 2p (1%), 2e (0.1%).

Spectroscopic characterization Circular dichroism (CD) spectra were measured using a Chirascan Circular Dichroism Spectrometer equipped with a Peltier temperature controller (Applied Photophysics, Ltd., UK). The CD spectra of the samples were recorded from 260 nm to 190 nm using a 2-mm or 1-mm path-length cuvette. Scans were repeated three or five times and averaged. The mean residue ellipticity (MRE) values were calculated per amino acid residue. FTIR spectra were measured using a Vertex 70 FTIR spectrometer (Bruker, USA). Samples were lyophilized using a lyophilizer (FDU-2100, manufactured by Tokyo Rikakikai Co., Ltd., Japan). Then, the lyophilized samples were dissolved in D2O, and 50 µL of the sample (150 µM, total 7.5 nmol) was placed onto a ZnSe window and dried in a fume hood. DLS was performed with a Zeta sizer Nano ZS instrument (Malvern, UK).

Transmission electron microscopy (TEM) Specimens were observed using a JEOL-TEM 2010 instrument (JEOL, Japan) operating at 120 kV. Two microliters of sample (100 µM) was placed onto a carbon-coated copper grid and


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incubated for 1 min. Then, filter papers were used to wick off the excess liquid, followed by adding 2 µl of 1% (w/v) uranyl acetate solution for 1 min. The excess liquid was again wicked off by filter paper.

Atomic force microscopy (AFM) Specimens were observed using a Nanoscope IV instrument (Digital Instruments, USA) and a NX-10 (Park Systems, South Korea). The peptides were dissolved in H2O or 15 mM potassium phosphate buffer with 150 mM KF, and the samples were incubated at least overnight at room temperature. Five microliters of sample (50~70 µM) was placed on a mica surface and dried in air.

Wide-angle X-ray scattering (WAXS) and small-angle X-ray scattering (SAXS) X-ray scattering experiments were performed at the 4C SAXS II beamline of the Pohang Accelerator Laboratory (PAL). A light source from an In-vacuum Undulator 20 (IVU 20: 1.4 m length, 20 mm period) of the Pohang Light Source II storage ring was focused with a vertical focusing toroidal mirror coated with rhodium and monochromatized with a Si (111) doublecrystal monochromator (DCM), yielding an X-ray beam wavelength of 0.675 Å. Samples were mounted in solution sample cells with a mica window that had a thickness of 10 µm, a volume of 50 µL, and an X-ray beam path length of 0.8 mm and irradiated for 30 s of exposure time at room temperature. Scattered radiation was acquired using a two-dimensional (2D) chargecoupled detector (Mar USA, Inc.) positioned 1 m (0.04 Å–1 < q < 0.50 Å–1) and 0.2 m (0.21 Å–1 < q < 2.83 Å–1) away from the sample for WAXS and SAXS, respectively. The SAXS data were collected in five successive frames of 0.1 min each to minimize radiation damage. Each 2D


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SAXS pattern was circularly averaged from the beam center and normalized to the transmitted X-ray beam intensity, which was monitored with a scintillation counter placed behind the sample. The scattering of water was used as the experimental background. To determine structural information from the SAXS experimental curves, the pair distance distribution function p(r) was obtained with the GNOM program using an indirect Fourier transform as follows: 1 pr = sin 2

where q is the momentum transfer (q = (4π/λ) sin θ, where 2θ is the scattering angle and λ is the wavelength of the X-ray), and r is the distance between the paired scattering elements. Using this approach, the maximum diameter of a given macromolecule (Dmax) could be acquired from the distance at which p(r) converges to zero, and the radius of gyration (Rg,p(r)) could be calculated by the following equation: ,

= 2

The low-resolution ab initio shape of the peptide in water was reconstructed using the DAMMIF program without any symmetry restriction. The surface rendering of the structural models was performed using Discovery Studio 3.5 (Accelrys, Inc.).

Degradation study For the degradation study of the peptides, 1p or 1e (-G or L) was dissolved in 10 mM phosphate buffer (pH 7.4). For the determination of the degradation rate of each peptide, the samples were analyzed using reverse-phase HPLC (water-acetonitrile with 0.1% TFA) at


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different time points. Peaks indicating degraded or non-degraded peptide were determined on an HPLC chromatogram, and then the sizes of the peaks, indicating the amplitude and area of each peak, were measured. The calculated areas were normalized and plotted, and all curves were fitted by a pseudo first-order kinetic model. For proteolytic degradation study, trypsin (0.01 to 0.1 mg/mL) or chymotrypsin (0.1 mg/mL) was added to the peptide sample in water. Aliquots taken at appropriate time intervals were analyzed with MALDI-TOF MS.

Cell culture, confocal microscopy, and fluorescence-activated cell sorting (FACS) For the microscopic observation of the intracellular delivery of the fluorescent dye containing peptides, HeLa cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% FBS and 1% penicillin and incubated at 37 °C/ 5% CO2. Then, the cells (2 × 104) were seeded on an 8-well Lab-Tek ll chambered cover-glass system in DMEM. The next day, the cells were washed with Dulbecco’s phosphate-buffered saline (DPBS) and treated with the dye-labeled peptides (2e or 2p, total 20 pmol in media) containing Opti-MEM media. The peptide-treated cells were observed by confocal microscopy using a Carl Zeiss (Germany) LSM 880 with a 63x oil immersion lens at different time points. The fluorescence signal of TAMRA was collected using excitation with a 561 nm laser and a 568-690 nm band-pass emission filter. The fluorescence signal of TAMRA by fluorescence resonance energy transfer (FRET) (the carboxyfluorescein (FAM)-TAMRA pair) was also collected using excitation with a 488 nm laser and a 568-690 nm band-pass emission filter. To compare the two signals (excitation with 488 nm and 561 nm), the emission using 561 nm was visualized in red and that using 488 nm, that is, the emission by FRET, was visualized in green.


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For FACS analysis, Hela cells (1 × 105) were seeded on 24-well plates and incubated overnight in DMEM supplemented with 10% FBS and 1% Pen/Strep at 37 °C/ 5% CO2. Then the cells were washed DPBS and the medium was replaced with Opti-MEM containing the peptides. After overnight incubation, the cells were washed with DPBS and trypsinized, followed by resuspension in DPBS supplemented with 10% cell dissociation buffer and 1% FBS. The cells were analyzed by FACSCalibur (BD biosciences, USA) using CELLQuest software.

Cytotoxicity assay Hela cells (2 × 104) were seeded on 96-well plates and cultured in DMEM supplemented with 10% FBS and 1% Pen/Strep at 37 °C/ 5% CO2 overnight. The next day, the cells were washed DPBS and the medium was replaced with Opti-MEM containing the peptides. After 4h incubation at 37 °C/ 5% CO2, the cells were washed with DPBS, and the medium was replaced with DMEM supplemented with 10% FBS and 1% Pen/Strep without phenol red. The cells were incubated at 37 °C/ 5% CO2 overnight. MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2Htetrazolium bromide) solution (5 mg/mL in DPBS) was added to each wells and incubated at 37 °C/ 5% CO2 for 2 h. Then medium was changed to DMSO and incubated at 37 °C/ 5% CO2 for 1 h. The OD at 490 nm was measured using a Victor X5 Multilabel plate reader (PerkinElmer, USA).


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RESULTS AND DISCUSSION

Figure 1. Self-assembly of depsipeptides and their intracellular delivery followed by degradation. (a) Structures of SADPs. (b) Self-assembly of 1e-G/1e-L into 1D fibers and their


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hydrolytic degradation. (c) Self-assembly of 2e into 3D vesicles, intracellular delivery, and biodegradation.

The designed SADP 1e-G consists of alternating aromatic/hydrophobic and positively or negatively charged amino acids (Figure 1a). Glycolic acid was used for the ester bond formation, which was conducted by the carbodiimide esterification method (Figures S2 and S3).15 The presence of aromatic tryptophan is likely to increase the strength of the β-sheet association via π−π stacking interactions. The conformation of the peptides was investigated using CD spectroscopy. A control peptide (1p) with all backbone linkages composed of peptide bonds exhibited the typical characteristics of a β-sheet in an aqueous solution, with strong negative and positive bands at 214 nm and 198 nm, respectively (Figure 2a).16 The SADP 1e-G was similarly shown to form β-sheets, as indicated by the strong negative and positive bands at 215 nm and 197 nm, respectively (Figure 2b). The value of molecular ellipticity at 215 nm ([θ]215), the signature of the β-sheet in the CD spectrum,17 indicated that the [θ]215 of 1e-G was 60% that of 1p. Thus, albeit reduced, 1e-G retained significant β-sheet formation propensity even after the incorporation of the ester linkage. In our study, we observed strong β-sheet formation in 1e-G at a concentration of 5 µM in phosphate-buffered saline (PBS). The strong [θ]215 at 5 µM, combined with the presence of β-sheet signature at lower concentration (Figure S6), suggests that the critical aggregation concentration (cac) should be lower than this concentration. Therefore, 1e-G can maintain the cross-β assembled state at biologically relevant concentrations. Fourier transform infrared spectroscopy (FTIR) revealed strong bands at 1624 cm-1 and 1627 cm1

for 1p and 1e-G, respectively, further confirming the presence of β-sheets in the SADP


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assembly (Figure 2c, d). Additionally, the presence of bands at 1692 cm-1 and 1690 cm-1 for 1p and 1e-G, respectively, indicates that the β-sheets in both peptides have an antiparallel orientation.18-21

Figure 2. Spectroscopic characterization of SADP assembly in solution. CD spectra of (a) 1p and (b) 1e-G. FTIR spectra of (c) 1p and (d) 1e-G. Original spectra (red) and after Fourier selfdeconvolution (black).


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Figure 3. Nanostructural characteristics of SADP assemblies in solution. AFM images of (a) 1p and (b) 1e-G. (c) SAXS data and SAXS-derived models. The models were reconstituted using an ab initio shape determination program, DAMMIN. (d) WAXS data. Black (1p); red (1e-G).

The morphologies of the 1p and 1e-G assemblies were then investigated. AFM observation revealed that both 1p and 1e-G formed fibrous 1D objects (Figure 3a, b). In terms of the nanostructural length, the 1e-G assemblies (~ 400-800 ± 300 nm) were longer than the 1p


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assemblies (~ 200-500 ± 100 nm). Because the samples are in the dried state during the AFM measurement and placed on a 2D surface, the real morphology and size of the objects in solution may not be correctly represented with this technique. To accurately define the structures in solution and to gain further structural insight, we performed synchrotron SAXS and WAXS measurements in solution.22,23 Structural ab initio models reconstructed from the experimental SAXS data corroborated the formation of 1D objects in solution (Figures 3c and S7). Additionally, the calculation of the radius of gyration, Rg, and the maximum dimension, Dmax, further confirmed that the length of the 1e-G assemblies is longer than that of the 1p assemblies. The self-assembled nanostructures of 1e-G were then characterized using WAXS in an aqueous solution. We found reflections corresponding to d-spacings of 2.87, 4.80, and 7.57 Å. The peak at 2.87 Å is a characteristic of the π−π stacking distance between tryptophan residues, and 4.80 Å corresponds to the β-sheet interstrand distance.24 The d-spacing of 7.57 Å corresponds to the β-sheet intersheet distance, indicating that the 1D fiber of the SADP is a bilayer β-sandwich structure.25,26 The controlled degradation and release of payloads should be an important parameter for biodegradable materials intended for use in intracellular delivery applications. We first asked the question of whether the SADP can be degraded by ester hydrolysis (Figure 1b). To address this question, NaOH was added to the solution of 1e-G at a final concentration of 5 mM, and the sample was subjected to CD measurement at several time intervals. As shown in Figure 4a, the signature of the β-sheet at approximately 215 nm decreased over time. This disassembly behavior implies that the hydrolytic cleavage of the ester weakens the strength of the cooperative β-sheet assembly. To corroborate the hydrolysis of the ester, we investigated the changes in the molecular weight of the sample by matrix-assisted laser desorption/ionization time-of-flight mass


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spectrometry (MALDI-TOF MS). Indeed, a MALDI spectrum following the incubation of 1e-G in a basic condition revealed peaks corresponding to the molecular weights of the hydrolyzed peptides (Figure 4b). By contrast, the hydrolytic cleavage of 1p could not be observed under similar basic conditions. When 1e-G was incubated under acidic conditions, the hydrolytic event was barely observed (Figure 4c). Thus, the study illustrated that the rate of hydrolysis is dependent on the solution pH, and the hydrolysis of the SADP ester bond is followed by the disassembly of the molecular assemblies.

Figure 4. Degradation of SADP. (a) CD spectra of 1e-G in 5 mM NaOH. t0 = immediately after NaOH addition. [1e-G] = 30 µM. MALDI-TOF MS spectra of 1e-G after (b) 24 h of incubation in 50 mM NaOH and (c) 2 weeks of incubation in 50 mM HCl. The m/z for [M+H]+: (i) calc'd


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(834.95), found (835.74); (ii) calc'd (892.00), found (892.81); (iii) calc'd (1707.93), found (1707.77).

Figure 5. Time-course study of degradation using HPLC. (a) 0 h. (b) After 1 day, (c) 4 days, and (d) 6 days. [1e-G] = 30 µM in 10 mM phosphate buffer, pH 7.4. (e) Plot of relative amount of


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intact SADP as a function of time. 1e-G at 25 °C (closed circles), 1e-G at 37 °C (closed triangles), 1e-L at 25 °C (open circles), 1e-L at 37 °C (open triangles).

Table 1. Half-life for SADP assemblies

a

SADPs

25 °C

37 °C

1e-G

2.8a

0.54

1e-L

4.0

0.92

Half-life in days. We then quantitatively measured the degradation kinetics of SADPs in a physiological buffer

(pH 7.4) using high-performance liquid chromatography (HPLC). Aliquots taken from the solution of 1e-G were subjected to HPLC analysis (Figure 5a-d). When 1e-G was incubated for an appropriate time, the formation of new two peaks that corresponded to the hydrolyzed peptides could be observed (Figure 5b). Plotting the integral values for the areas under the peaks as a function of time allowed a quantitative representation of the SADP degradation kinetics (Figure 5e). The hydrolysis is the reaction between water and the SADP. Because of the large excess of water, the concentration of water remained constant during hydrolysis. Thus, assuming pseudo-first-order reaction kinetics, the degradation data were fitted using the rate equation

[SADP] = [SADP] ! "#$

where [SADP]0 is the initial concentration, k is the rate constant, and t is the incubation time. Then, the half-life (τ) for the hydrolysis reaction was calculated by


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τ=

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ln 2 '

The calculated half-lives at 25 °C and 37 °C were 2.8 and 0.54 d, respectively (Table 1). Thus, the rate of hydrolysis for the SADPs increased with increasing temperature. We then questioned whether further control of the hydrolysis rate is possible by modifying the SADP molecular structure. We hypothesized that the use of lactic acid that has an additional methyl group might extend the half-life compared to glycolic acid. Because of the hydrophobic nature of the methyl group, the water accessibility to the ester is likely to be decreased. In the field of drug delivery systems, the copolymerization of lactic acid with glycolic acid has been a popular method to control the degradation kinetics of biodegradable polymers.1,27,28 As expected, the determination of the degradation rate for the SADP incorporating lactic acid (1e-L) revealed an increase in the half-life compared with 1e-G (Figure 5e and Table 1). Taken together, the rate of hydrolytic degradation of SADPs can be controlled by the pH, the temperature, and the structure of the ester unit.


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Figure 6. MALDI-TOF MS spectra from proteolytic degradation study. a) and b) trypsin-treated samples. c) and d) chymotrypsin-treated samples. Samples were treated with trypsin (0.01 mg/mL) or chymotrypsin (0.1 mg/mL) for 24 h and 5 h, respectively. Curly lines (red) indicate the proteolytic and/or hydrolytic cleavage sites. Ga: glycolic acid.

Degradation of peptide bonds can be facilitated by the presence of cellular proteases. We compared the proteolytic degradation rates of the normal peptide (1p) and an SADP (1e-G). First, the peptides were incubated with trypsin and aliquots taken at appropriate time intervals were analyzed using MALDI-TOF MS. Representative mass spectra in Figure 6a and 6b show the


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significant degradation of 1e-G, while 1p remaining nearly intact after the prolonged incubation with the protease. Chymotrypsin treatment also induced faster degradation of 1e-G than 1p (Figure 6c,d). Thus, the data suggest that, combined with the hydrolytic degradation data, in vivo degradation rate of SADP assemblies is likely to be faster than that of normal peptide assemblies. We further compared the proteolytic degradation rate of differently structured SADPs. As shown in Figure 7, we observed faster proteolytic degradation of 1e-G than that of 1e-L, which is in line with the hydrolytic degradation data (Figure 5).


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Figure 7. MALDI-TOF MS spectra from proteolytic degradation study. a) and b) trypsin-treated samples. c) and d) chymotrypsin-treated samples. Samples were treated with trypsin (0.01 mg/mL) or chymotrypsin (0.1 mg/mL) for 16 h and 24 h, respectively. Curly lines (red) indicate the proteolytic and/or hydrolytic cleavage sites. La: lactic acid.

Next, we sought to address the cell delivery possibility and the question of the intracellular degradation of SADP assemblies. We designed 2e, which has a cell-penetrating peptide (CPP) Rev segment and SADP segment.29 The N- and C-termini of 2e were fluorescently labeled with FAM and TAMRA for intracellular tracking. 2p is a control in which all the bonds are composed of amides. A CD investigation showed the coexistence of unstructured and β-sheet populations in both the 2p and 2e assemblies (Figure 8a). The bands for unstructured peptides at approximately 203 nm likely resulted from the cell penetration segments. An AFM investigation of the SPNs revealed that both 2p and 2e assemble into vesicle-like structures (Figures 8b, c). Dynamic light scattering (DLS) experiments were conducted to obtain ensemble-averaged hydrodynamic diameters and the results showed that both assemblies had similar sizes (Figure S8). We interpret that the premature termination of the β-sheet growth because of the steric interference among flexible CPP segments30 and the increased volume fraction of the hydrophilic segment are the primary driving forces in the formation of the vesicle-like structures.


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Figure 8. Self-assembly of 2p and 2e. (a) CD spectra of 2p (left) and 2e (right). Arrows indicate the signature of the β-sheet at approximately 215 nm. AFM images of (b) 2p and (c) 2e SPNs. Left (height images), right (phase images).


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First, we compared the cytotoxicity of 2p and 2e in mammalian cells, HeLa cells. Interestingly, the biodegradable 2e showed significantly reduced toxicity compared to 2p (Figure S10). To visualize the intracellular biodegradation, the cells were treated with the SPNs of 2p and 2e and incubated for 48 h to allow an appreciable amount of biodegradation. Laser scanning confocal microscopy (LSCM) was performed with excitation at 543 nm and the emission bandpass filter set between 568–690 nm for red fluorescence detection. In this condition, only the red fluorescence from TAMRA is observed. In principle, the fluorescence intensity from TAMRA would remain the same regardless of the degradation status. As shown in the middle images of Figure 9, the cells treated with 2p and 2e exhibited similar levels of TAMRA fluorescence. Then, we observed Fluorescence resonance energy transfer (FRET) between FAM and TAMRA by irradiating the sample at 488 nm and setting the emission bandpass filter between 568–690 nm. Because the FRET efficiency exponentially decreases as the distance between the two probes increases, decreased FRET emission is expected following the degradation and disassembly of the SADP assemblies. As shown in the bottom images of Figure 9, we observed a decrease in FRET emission in the cells treated with the 2e assemblies. To further confirmed the faster degradation of 2e in the cellular context, we conducted flow cytometry-based FRET assay (Figure S9). Following the treatment with the peptides, cells were trypsinized and the FACS analysis was performed with the excitation of FAM at 488 nm and the acquisition at 530 nm (FL1 channel) and 585 nm (FL2 channel) for the detection of emissions from FAM and TAMRA, respectively. The ratios of fluorescence intensities at 585 nm and 530 nm were 12 ± 1.3 and 8 ± 0.8 for 2p and 2e, respectively (mean ± s.d. n=4). The less efficient FRET signal indicates the faster degradation of 2e than 2p within the cell. Therefore, the


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microscopy and FACS data verified that the SADP assemblies can be biodegraded following internalization into the cells.

Figure 9. Cell delivery and intracellular degradation of SADP assemblies. HeLa cells treated with (a) 2p and (b) 2e. [2p] = [2e] = 0.67 µM. Merged bright-field and fluorescence images (top


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images). Fluorescence images with excitation at 543 nm and emission from 568 to 690 nm (middle images). Fluorescence images with excitation at 488 nm and emission from 568 to 690 nm (bottom images). In the bottom images, the original red fluorescence was pseudo-colored to green for optimal visual contrast. Bar = 20 µm.

CONCLUSIONS In conclusion, we have designed cell-penetrating cross-β assemblies with controlled biodegradable properties. The self-assembling β-sheet peptides incorporating an ester bond in the middle of the backbone formed 1D fibers with β-sandwich structures. The degradation rate of the SADP assemblies could be controlled as a function of the pH, the temperature, and the structure of the ester unit. The hydrolysis of the ester bond induced the disassembly of the self-assembled nanostructures. SADP assemblies also displayed faster proteolytic degradation kinetics than peptide assemblies composed of all peptide bonds. A block copolypeptide of the CPP and SADP mediated efficient cellular internalization and eventual intracellular degradation. We envision that self-assembled nanostructures based on SADPs can solve the potential problems associated with the negligible degradability of cross-β assemblies.

ASSOCIATED CONTENT Supporting Information. The supporting information is available free of charge on the ACS Publications website at DOI:


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Peptide chemical structures, reaction schemes, MALDI spectra, HPLC chromatograms, CD spectra, SAXS and DLS data, FACS and cytotoxicity data (PDF).

AUTHOR INFORMATION Corresponding Author *Tel: +82-2-2123-5836; E-mail: [email protected]

Present Addresses † Samsung Bioepis Co., Ltd., Incheon, Korea

Author Contributions ‡These authors contributed equally.

ACKNOWLEDGMENT This work was supported by grants from the National Research Foundation (NRF) of Korea (2014R1A2A1A11050359, 2014M3A7B4051594) and Yonsei University Future-leading Research Initiative.


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