Fluorescent Tau-derived Peptide for Monitoring Microtubules in Living

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Article Cite This: ACS Omega 2019, 4, 11245−11250

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Fluorescent Tau-derived Peptide for Monitoring Microtubules in Living Cells Hiroshi Inaba,*,†,‡ Takahisa Yamamoto,† Takashi Iwasaki,§ Arif Md. Rashedul Kabir,∥ Akira Kakugo,∥,⊥ Kazuki Sada,∥,⊥ and Kazunori Matsuura*,†,‡

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†

Department of Chemistry and Biotechnology, Graduate School of Engineering and ‡Centre for Research on Green Sustainable Chemistry, Tottori University, Koyama-Minami 4-101, Tottori 680-8552, Japan § Department of Bioresources Science, Graduate School of Agricultural Sciences, Tottori University, Koyama-Minami 4-101, Tottori 680-8553, Japan ∥ Faculty of Science and ⊥Graduate School of Chemical Sciences and Engineering, Hokkaido University, Kita 10 Nishi 8, Kita-ku, Sapporo 060-0810, Japan S Supporting Information *

ABSTRACT: Microtubules (MTs) are key cytoskeletal components that modulate various cellular activities with their dynamic structural changes, including polymerization and depolymerization. To monitor the dynamics of MTs in living cells, many drug-based fluorescent probes have been developed; however, these also potentially disturb the polymerization/depolymerization of MTs. Here, we report nondrug, peptide-based fluorescent probes to monitor MTs in living cells. We employed a Tau-derived peptide (TP) that has been shown to bind MTs without inhibiting polymerization/depolymerization in vitro. We show that a tetramethylrhodamine (TMR)-labeled TP (TP−TMR) is internalized into HepG2 cells and binds to intracellular MTs, enabling visualization of MTs as clear, fibrous structures. The binding of TP−TMR shows no apparent effects on polymerization/depolymerization of MTs induced by MT-targeted drugs and temperature change. The main uptake mechanism of TP−TMR was elucidated as endocytosis, and partial endosomal escape resulted in the binding of TP−TMR to MTs. TP−TMR exhibited no cytotoxicity compared with MT-targeted drug scaffolds. These results indicate that TP scaffolds can be exploited as useful MT-targeted tools in living cells, such as in long-term imaging of MTs.

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polymerization of MTs, respectively.13 By conjugating fluorophores to taxol14−22 and colchicine23 scaffolds, intracellular imaging of MTs has been achieved with high efficiency and high resolution. These drug-based probes are useful because of their permeability to cells and high selectivity for MTs. However, these drug scaffolds have several problems, such as low water solubility and complex structures with limited numbers of reactive groups.14−16,18,20,22 Most importantly, since drug-based probes potentially induce cytotoxicity by affecting the polymerization/depolymerization of MTs, optimization of the probe structures is essential for imaging of MTs with minimal cytotoxicity.18,20,22 Thus, a nondrug MTtargeted molecule will be useful to monitor MTs without disturbing the intracellular dynamics of MTs. Peptides are attractive candidates for monitoring MTs because of their biocompatibility, sequence-dependent cell permeability, and molecular recognition capability.24−28 In fact, various MT-binding peptides have been developed but the

INTRODUCTION Microtubules (MTs) are cytoskeletal components that serve important roles in various cellular processes, such as intracellular material transportation, cellular support, and cell division.1−3 MTs are hollow tubular structures consisting of tubulin heterodimers and are constantly extending (polymerizing) and contracting (depolymerizing) in cells. The dynamics of polymerization/depolymerization by MTs are essential to maintain cellular activities, including the cell cycle.1−3 Directional transport of intracellular cargos on MTs by motor proteins (kinesin and dynein) is also important in cells, which have been widely utilized to construct MT-based active materials.2,4−9 Because of the importance of MTs, various methodologies have been developed to monitor MTs in living cells without chemical fixation to understand and modulate the dynamics of MTs.10−12 Expression of fluorescent protein-fused tubulin is a general method to monitor MTs in living cells.11 However, the low efficiency of transfection and low expression levels may be problematic. Alternatively, smallmolecule fluorescent probes have been developed by using MT-binding drugs as scaffolds.12 Taxol and colchicine are wellknown MT-targeted drugs that inhibit depolymerization and © 2019 American Chemical Society

Received: April 16, 2019 Accepted: June 18, 2019 Published: June 27, 2019 11245

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Figure 1. (a) Tau-derived peptide (TP) designed from Tau protein as a binding motif for microtubules (MTs). (b) Schematic representation of cellular uptake of tetramethylrhodamine (TMR)-labeled TP (TP−TMR) and subsequent binding to MTs.

mammalian cells. Upon incubation of 10 μM TP−TMR with the cells for 6 h at 37 °C, TP−TMR was found to be internalized into the cells and bind to intracellular MTs, observed as clear fibrous structures (Figure 2a and Movie S1).

purpose was construction of MT-targeted drugs, and not livecell imaging of MTs.29−35 Ghosh et al. have developed short (4−9-mer) peptides and peptoids for modulation of intracellular dynamics of MTs by promoting or inhibiting polymerization of MTs.30−35 Although binding of some peptides to MTs in fixed cells has been imaged, no obvious fibrous structures of the peptide-bound MTs were observed.31,32 This may be due to the low affinity of short peptides for tubulin, as their dissociation constants (Kd) are in the millimolar range.31,32 It is anticipated that peptides with high affinity for MTs without disturbing polymerization/ depolymerization of MTs are suitable to monitor MTs in living cells. We previously developed an MT-binding peptide based on an MT-associated protein, Tau (Figure 1a).36 The Tau-derived peptide TP (CGGGKKHVPGGGSVQIVYKPVDL) was designed on the basis of a repeat domain of Tau involved in binding to the inner surface of MTs. From our binding analysis in vitro (not in cellulo), TP binds to a taxol-binding pocket of tubulin located inside MTs with high affinity (Kd = 6.0 μM) by preincubation with tubulin and subsequent polymerization of the peptide−tubulin complex. In contrast, TP binds to the outer surface of MTs when incubated with preformed MTs. Binding of TP does not inhibit the polymerization/ depolymerization of MTs in vitro. In addition, TP has high water solubility at millimolar concentrations. These properties of TP are suitable for an MT-binding motif in living cells. Here, we prepared fluorescently labeled TP and analyzed binding to MTs in living cells. We demonstrated that TP can bind to intracellular MTs via endocytic uptake and subsequent endosomal escape (Figure 1b) without inhibiting polymerization/depolymerization of MTs.

Figure 2. Binding of TP−TMR to MTs in HepG2 cells. Channels for (a) TP−TMR (red), (b) tubulin−GFP (green), and (c, d) TP− TMR, tubulin−GFP, and Hoechst 33342 (cyan) for confocal laser scanning microscopy (CLSM) images. White arrows in image (a) indicate the binding of TP−TMR to MTs at cell edges. (e) Distribution of TP−TMR and tubulin−GFP was quantified by measuring fluorescence intensity in the lined areas of the expanded image (d). All scale bars are 10 μm. See Movie S1 for cross-sectional scans of cells acquired at different heights. Tubulin−GFP was expressed in HepG2 cells; then, 10 μM TP−TMR was incubated with the cells for 6 h at 37 °C under 5% CO2. Cell nuclei were stained with Hoechst 33342.

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RESULTS AND DISCUSSION Red fluorescent tetramethylrhodamine (TMR)-5-maleimide was conjugated to the N-terminal cysteine of TP as a fluorophore according to our reported procedure.36 The TMR-labeled TP (TP−TMR) and 5(6)-carboxytetramethylrhodamine (TMR−COOH) showed similar fluorescence spectra (Figure S1), indicating that TP−TMR can be used for imaging without quenching TMR. Cellular uptake of TP− TMR into human hepatoma HepG2 cells was analyzed by confocal laser scanning microscopy (CLSM). Before incubation with TP−TMR, the cells were transfected with green fluorescent protein-fused tubulin (tubulin−GFP) using BacMam technology (Thermo Fischer Scientific), which uses baculovirus as a carrier to deliver and express genes in

The fibers in the TP−TMR channel were essentially colocalized with the fibers observed in the tubulin−GFP channel (Figure 2b,c). Quantitative analysis of the distribution of fluorescent intensity also showed colocalization of TP− TMR and tubulin−GFP fiber structures (Figure 2d,e), indicating the binding of TP−TMR to intracellular MTs. In particular, binding of TP−TMR to MTs at the edges of cells was clearly observed (white arrows in Figure 2a), probably because of the high density of MTs at cell edges to exert 11246

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cellular support and cell migration.37,38 Distribution of TP− TMR-bound MTs throughout cells can be seen in Movie S1, which shows cross-sectional scans of cells acquired at different heights. On the other hand, the dotlike structures observed in the TP−TMR channel (Figure 2a) were not colocalized with tubulin−GFP (Figure 2b,c). The dots were expected to be endosomes, as investigated below. Even by using lower concentration of TP−TMR and shorter incubation time compared with the above condition (10 μM TP−TMR, 6 h incubation), the binding of TP−TMR to intracellular MTs was still observed; however, the fluorescence intensity of TP− TMR was low (Figures S2 and S3). Thus, we used the above condition (10 μM TP−TMR, 6 h of incubation) in the following experiments. In contrast with TP−TMR, entry of TMR−COOH into HepG2 cells was not observed even after 6 h of incubation (Figure S4), indicating that the TP moiety is required for cellular uptake. These results show that TP−TMR can penetrate into cells and bind to intracellular MTs. In addition, even after incubation of the cells for 24 h upon uptake of TP−TMR, the binding of TP−TMR to intracellular MTs was still observed, indicating that TP−TMR can be used for long-term imaging of MTs (Figure S5). The effects of MT-binding drugs taxol and colchicine on the binding of TP−TMR to intracellular MTs were evaluated. Even after coincubation of 10 μM TP−TMR and 10 μM taxol (a MT-stabilizing agent) with HepG2 cells, fibrous structures were observed in the TP−TMR channel (Figure 3a). The fibers were essentially colocalized with the fibers in the tubulin−GFP channel (Figure 3b,c), indicating the binding of TP−TMR to intracellular MTs even in the presence of taxol. We have previously shown that TP−TMR has two binding sites in MTs: (1) a taxol-binding pocket of tubulin when incubating TP−TMR with tubulin and subsequent polymerization of the peptide−tubulin complex and (2) the outer surface of MTs when incubating TP−TMR with preformed MTs.36 Because coincubation with taxol showed little effect on the fibrous structures in the TP−TMR channel (Figure 3a), it is possible that TP−TMR mainly binds to the outer surface of preformed MTs. When 10 μM TP−TMR was incubated with HepG2 cells in the presence of 10 μM colchicine (an MTdestabilizing agent), cell morphology changed to being round and only dotlike structures were observed but no fiber structures were observed in the TP−TMR channel (Figure 3d). Also, tubulin−GFP was homogeneously distributed in the cytoplasm and did not form fibrous MTs (Figure 3e). These results clearly indicate that the polymerization of MTs was inhibited by colchicine and TP−TMR-bound MTs were not formed under the conditions employed. To elucidate whether TP−TMR-bound MTs depolymerize/polymerize in cells, TP−TMR-treated cells were cooled and heated as depolymerization and polymerization conditions, respectively. The fibrous structures in the TP−TMR channel observed before cooling (Figure 3g) disappeared after cooling (Figure 3h), showing the depolymerization of TP−TMR-bound MTs. Upon heating the cells, the polymerization occurred to form TP−TMR-bound MTs (Figure 3i). Thus, binding of TP− TMR exhibits no apparent effects on depolymerization/ polymerization of MTs in living cells, indicating that TP− TMR may be useful in monitoring MT dynamics. The cellular uptake mechanism of TP−TMR was elucidated by cooling the cells during uptake to inhibit an energydependent uptake process.39 Uptake of TP−TMR in HepG2 cells was significantly inhibited at 4 °C (Figure S6). This result

Figure 3. Effects of (a−c) taxol and (d−f) colchicine on the binding of TP−TMR to MTs. Channels for (a, d) TP−TMR, (b, e) tubulin− GFP, and (c, f) TP−TMR, tubulin−GFP, and Hoechst 33342 in CLSM images. Tubulin−GFP was expressed in HepG2 cells, then 10 μM TP−TMR with 10 μM taxol or 10 μM colchicine was incubated with the cells for 6 h at 37 °C under 5% CO2. Cell nuclei were stained by Hoechst 33342. (g−i) Temperature-dependent depolymerization/ polymerization of TP−TMR-bound MTs in HepG2 cells. CLSM images were acquired (g) immediately, (h) upon cooling at ∼4 °C for 40 min, and (i) upon heating at ∼40 °C for 5 min. White arrows indicate TP−TMR-bound MTs. All scale bars are 10 μm.

indicates that uptake of TP−TMR involves an endocytic pathway. Since dotlike structures along with fibrous structures were observed in the TP−TMR channel when incubated at 37 °C (Figure 2a), it is possible that TP−TMR is partially released from endosomes, distributed in cytoplasm, and then bound to MTs. To investigate this hypothesis, colocalization of TP−TMR and a typical endosome marker, fluorescein isothiocyanate-labeled transferrin (FITC−transferrin), was identified.40 As a result, the dotlike structures in the TP− TMR channel were colocalized with FITC−transferrin, whereas the fibrous structures in the TP−TMR channel were not observed in the FITC−transferrin channel (Figure S7). Thus, the dotlike structures in the TP−TMR channel are attributed to endosomes and partial endosomal escape of TP− TMR results in the binding to MTs observed as fibrous structures in the TP−TMR channel. When human glioma U251 cells were used instead of HepG2 cells, only dotlike 11247

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structures were observed but no fiber structures of MTs were observed in the TP−TMR channel (Figure S8), indicating that TP−TMR was internalized into U251 cells via an endocytic pathway without subsequent endosomal escape. We checked the bioactivity of TP−TMR using the watersoluble tetrazolium salt (WST) assay, a method to measure cell viability by evaluating mitochondrial activity in living cells. TP−TMR over a range of 1−20 μM showed no cytotoxicity even after incubation with HepG2 cells for 24 h (Figure 4).

was carried out using a FluoView FV10i (Olympus). The reagents were purchased from Watanabe Chemical Ind., Ltd., Tokyo Chemical Industry Co., Dojindo Laboratories Co., Ltd. and Wako Pure Chemical Industries. All chemicals were used without further purification. Cell Culture. Human hepatoma HepG2 cells and human glioma U251 cells were cultured in Dulbecco’s modified Eagle’s medium. All medium contained 10% fetal bovine serum (v/v), 100 μg/mL streptomycin, 100 units/mL penicillin, 1 mM sodium pyruvate, and 1% MEM nonessential amino acids (v/v, Sigma M7145). Cells were maintained at 37 °C in a 5% CO2 incubator, and a subculture was performed every 3−4 days. CLSM Measurements. HepG2 cells and U251 cells were seeded onto a single-well glass bottom dish at 2.0 × 104 cells/ well in a final volume of 100 μL and incubated for ∼24 h at 37 °C, 5% CO2. CellLight tubulin−GFP, BacMam 2.0 (Thermo Fisher Scientific) was added to the cells (1 μL per well) and incubated for further 24 h, at 37 °C, 5% CO2. After removal of the medium, 10 μM TP−TMR in the medium was added to the cells and incubated for a designed period. For evaluation of the effects of MT-binding drugs, 10 μM taxol or 10 μM colchicine was coincubated with 10 μM TP−TMR at the same time. For evaluation of the endosomal escape, 0.1 mg/mL FITC−transferrin was coincubated with 10 μM TP−TMR at the same time. After washing with PBS, the cells were incubated with 5 μg/mL Hoechst 33342 for 10 min at 37 °C, 5% CO2. After washing with PBS, the medium was added to the cells and fluorescence images were acquired. TP−TMR was excited with 550 nm and observed through a 574 nm emission band-pass filter (red). Tubulin−GFP was excited with 489 nm and observed through a 510 nm emission bandpass filter (green). FITC−transferrin was excited with 495 nm and observed through a 519 nm emission band-pass filter (green). Hoechst 33342 was excited with 352 nm and observed through a 455 nm emission band-pass filter (cyan). Cell Viability Assay (WST Assay). The cultured HepG2 cells were plated in 96-well plates at 1.0 × 104 cells/well in a final volume of 100 μL and incubated for 24 h at 37 °C, 5% CO2. The culture medium was replaced with medium containing 0−20 μM TP−TMR, taxol, or colchicine and incubated for 24 or 72 h. The culture medium was replaced with fresh medium, and 10 μL of the Cell Counting Kit-8 reaction solution (Dojindo, Japan) was added to each well. After incubation for 4 h, the absorbance at 450 nm was measured. Cell viability (%) was calculated by setting the absorbance of the untreated control cells as 100%.

Figure 4. Cytotoxicity of TP−TMR against HepG2 cells. HepG2 cells were incubated with 0−20 μM TP−TMR (red), taxol (black), or colchicine (blue) for 24 h at 37 °C under 5% CO2. Cell viability was evaluated by the WST assay. The error bars represent standard deviation.

Even after longer incubation time (72 h), TP−TMR showed no cytotoxicity (Figure S9). In contrast, MT-binding drugs taxol and colchicine showed clear cytotoxicity in a concentration-dependent manner (Figure 4). The cytotoxicity of taxol and colchicine is due to inhibition of depolymerization and polymerization of MTs, respectively.13 Thus, TP−TMR showed no cytotoxicity because binding of TP−TMR to MTs exhibits minimal effects on depolymerization/polymerization of MTs, as described above. These results indicate that TP is useful as a scaffold for MT binding for long-term usage without disturbing cellular activity.

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CONCLUSIONS In conclusion, we have demonstrated that a fluorescently labeled Tau-derived peptide, TP−TMR, can be successfully used for imaging of MTs in living cells. Binding of TP−TMR to MTs showed no apparent effects on polymerization/ depolymerization of MTs and no cytotoxicity in contrast with the known MT-binding drugs taxol and colchicine. Thus, TP− TMR can be used for imaging of MTs without disturbing cellular activities. Especially, time-lapse imaging of MTs during cell division is important to analyze the functions of MTs in living cells. We also envision that the TP scaffold may be applicable for MT-targeted modulation of cellular activities. A notable example is optical modulation of MT dynamics by designing photoswitchable MT-destabilizing agents, enabling control of mitosis and cell death.41 In such a way, design of TP to control the dynamics of MTs is in progress.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsomega.9b01089. Fluorescence spectra of TP−TMR and TMR−COOH; CLSM images; cell viability assay (PDF) Movie S1: Cross-sectional scans of cells acquired at different heights (MPG)

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METHODS Equipment and Materials. TP−TMR was prepared as described in our previous work.36 High-performance liquid chromatography was performed to purify TP−TMR using a Shimadzu LC-6AD liquid chromatograph with GL Science Inertsil WP300 C18 columns (20 × 250 mm2). Fluorescence measurements were performed using a Jasco FP-8200. Confocal laser scanning microscopy (CLSM) measurement

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected] (H.I.). *E-mail: [email protected] (K.M.). 11248

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ORCID

W.; Johnsson, K. Fluorogenic probes for live-cell imaging of the cytoskeleton. Nat. Methods 2014, 11, 731−733. (19) Lukinavičius, G.; Reymond, L.; Umezawa, K.; Sallin, O.; D’Este, E.; Göttfert, F.; Ta, H.; Hell, S. W.; Urano, Y.; Johnsson, K. Fluorogenic Probes for Multicolor Imaging in Living Cells. J. Am. Chem. Soc. 2016, 138, 9365−9368. (20) Lee, M. M.; Gao, Z.; Peterson, B. R. Synthesis of a Fluorescent Analogue of Paclitaxel That Selectively Binds Microtubules and Sensitively Detects Efflux by P-Glycoprotein. Angew. Chem., Int. Ed. 2017, 56, 6927−6931. (21) Butkevich, A. N.; Ta, H.; Ratz, M.; Stoldt, S.; Jakobs, S.; Belov, V. N.; Hell, S. W. Two-Color 810 nm STED Nanoscopy of Living Cells with Endogenous SNAP-Tagged Fusion Proteins. ACS Chem. Biol. 2018, 13, 475−480. (22) Lukinavičius, G.; Mitronova, G. Y.; Schnorrenberg, S.; Butkevich, A. N.; Barthel, H.; Belov, V. N.; Hell, S. W. Fluorescent dyes and probes for super-resolution microscopy of microtubules and tracheoles in living cells and tissues. Chem. Sci. 2018, 9, 3324−3334. (23) Zhang, H.; Wang, C.; Jiang, T.; Guo, H.; Wang, G.; Cai, X.; Yang, L.; Zhang, Y.; Yu, H.; Wang, H.; Jiang, K. Microtubuletargetable fluorescent probe: site-specific detection and superresolution imaging of ultratrace tubulin in microtubules of living cancer cells. Anal. Chem. 2015, 87, 5216−5222. (24) Hamley, I. W. Small Bioactive Peptides for Biomaterials Design and Therapeutics. Chem. Rev. 2017, 117, 14015−14041. (25) Zhang, P.; Cui, Y.; Anderson, C. F.; Zhang, C.; Li, Y.; Wang, R.; Cui, H. Peptide-based nanoprobes for molecular imaging and disease diagnostics. Chem. Soc. Rev. 2018, 47, 3490−3529. (26) Qi, G.-B.; Gao, Y.-J.; Wang, L.; Wang, H. Self-Assembled Peptide-Based Nanomaterials for Biomedical Imaging and Therapy. Adv. Mater. 2018, 30, 1703444. (27) Tsutsumi, H.; Mihara, H. Soft materials based on designed selfassembling peptides: from design to application. Mol. BioSyst. 2013, 9, 609−617. (28) Inaba, H.; Matsuura, K. Peptide Nanomaterials Designed from Natural Supramolecular Systems. Chem. Rec. 2019, 19, 843−858. (29) Pieraccini, S.; Saladino, G.; Cappelletti, G.; Cartelli, D.; Francescato, P.; Speranza, G.; Manitto, P.; Sironi, M. In silico design of tubulin-targeted antimitotic peptides. Nat. Chem. 2009, 1, 642− 648. (30) Biswas, A.; Kurkute, P.; Jana, B.; Laskar, A.; Ghosh, S. An amyloid inhibitor octapeptide forms amyloid type fibrous aggregates and affects microtubule motility. Chem. Commun. 2014, 50, 2604− 2607. (31) Biswas, A.; Kurkute, P.; Saleem, S.; Jana, B.; Mohapatra, S.; Mondal, P.; Adak, A.; Ghosh, S.; Saha, A.; Bhunia, D.; Biswas, S. C.; Ghosh, S. Novel hexapeptide interacts with tubulin and microtubules, inhibits Aβ fibrillation, and shows significant neuroprotection. ACS Chem. Neurosci. 2015, 6, 1309−1316. (32) Jana, B.; Mondal, P.; Saha, A.; Adak, A.; Das, G.; Mohapatra, S.; Kurkute, P.; Ghosh, S. Designed Tetrapeptide Interacts with Tubulin and Microtubule. Langmuir 2018, 34, 1123−1132. (33) Mondal, P.; Das, G.; Khan, J.; Pradhan, K.; Ghosh, S. Crafting of Neuroprotective Octapeptide from Taxol-Binding Pocket of βTubulin. ACS Chem. Neurosci. 2018, 9, 615−625. (34) Pradhan, K.; Das, G.; Gupta, V.; Mondal, P.; Barman, S.; Khan, J.; Ghosh, S. Discovery of Neuroregenerative Peptoid from Amphibian Neuropeptide That Inhibits Amyloid-β Toxicity and Crosses Blood-Brain Barrier. ACS Chem. Neurosci. 2019, 10, 1355− 1368. (35) Mondal, P.; Das, G.; Khan, J.; Pradhan, K.; Mallesh, R.; Saha, A.; Jana, B.; Ghosh, S. Potential Neuroprotective Peptide Emerged from Dual Neurotherapeutic Targets: A Fusion Approach for the Development of Anti-Alzheimer’s Lead. ACS Chem. Neurosci. 2019, 10, 2609−2620. (36) Inaba, H.; Yamamoto, T.; Kabir, A. M. R.; Kakugo, A.; Sada, K.; Matsuura, K. Molecular Encapsulation Inside Microtubules Based on Tau-Derived Peptides. Chem. - Eur. J. 2018, 24, 14958−14967.

Hiroshi Inaba: 0000-0002-7658-7827 Akira Kakugo: 0000-0002-1591-867X Kazunori Matsuura: 0000-0001-5472-7860 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by KAKENHI (Nos. 17K14517 and 19K15699 for H.I.) from the Japan Society for the Promotion of Science (JSPS), the Inamori Foundation, and Konica Minolta Science and Technology Foundation for Konica Minolta Imaging Science Encouragement Award.

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REFERENCES

(1) Fletcher, D. A.; Mullins, R. D. Cell mechanics and the cytoskeleton. Nature 2010, 463, 485−492. (2) Hess, H.; Ross, J. L. Non-equilibrium assembly of microtubules: from molecules to autonomous chemical robots. Chem. Soc. Rev. 2017, 46, 5570−5587. (3) Nogales, E. Structural insights into microtubule function. Annu. Rev. Biochem. 2000, 69, 277−302. (4) Kabir, A. M. R.; Kakugo, A. Study of active self-assembly using biomolecular motors. Polym. J. 2018, 50, 1139−1148. (5) Bachand, G. D.; Spoerke, E. D.; Stevens, M. J. Microtubulebased nanomaterials: Exploiting nature’s dynamic biopolymers. Biotechnol. Bioeng. 2015, 112, 1065−1073. (6) Song, W.; Mö hwald, H.; Li, J. Movement of polymer microcarriers using a biomolecular motor. Biomaterials 2010, 31, 1287−1292. (7) Song, W.; Zhu, J.; Kong, W.; Möhwald, H.; Li, J. Different Microtubule Structures Assembled by Kinesin Motors. Langmuir 2018, 34, 9768−9773. (8) Keya, J. J.; Suzuki, R.; Kabir, A. M. R.; Inoue, D.; Asanuma, H.; Sada, K.; Hess, H.; Kuzuya, A.; Kakugo, A. DNA-assisted swarm control in a biomolecular motor system. Nat. Commun. 2018, 9, 453. (9) Matsuda, K.; Kabir, A. M. R.; Akamatsu, N.; Saito, A.; Ishikawa, S.; Matsuyama, T.; Ditzer, O.; Islam, M. S.; Ohya, Y.; Sada, K.; Konagaya, A.; Kuzuya, A.; Kakugo, A. Artificial Smooth Muscle Model Composed of Hierarchically Ordered Microtubule Asters Mediated by DNA Origami Nanostructures. Nano Lett. 2019, 19, 3933−3938. (10) Sahl, S. J.; Hell, S. W.; Jakobs, S. Fluorescence nanoscopy in cell biology. Nat. Rev. Mol. Cell Biol. 2017, 18, 685−701. (11) Zwetsloot, A. J.; Tut, G.; Straube, A. Measuring microtubule dynamics. Essays Biochem. 2018, 62, 725−735. (12) Wang, L.; Frei, M. S.; Salim, A.; Johnsson, K. Small-Molecule Fluorescent Probes for Live-Cell Super-Resolution Microscopy. J. Am. Chem. Soc. 2019, 141, 2770−2781. (13) Jordan, M. A.; Wilson, L. Microtubules as a target for anticancer drugs. Nat. Rev. Cancer 2004, 4, 253−265. (14) Souto, A. A.; Acuña, A. U.; Andreu, J. M.; Barasoain, I.; Abal, M.; Guerri, F. A. New Fluorescent Water-Soluble Taxol Derivatives. Angew. Chem., Int. Ed. 1996, 34, 2710−2712. (15) Guy, R.; Scott, Z.; Sloboda, R.; Nicolaou, K. Fluorescent taxoids. Chem. Biol. 1996, 3, 1021−1031. (16) Evangelio, J. A.; Abal, M.; Barasoain, I.; Souto, A. A.; Lillo, M. P.; Acuña, A. U.; Amat-Guerri, F.; Andreu, J. M. Fluorescent taxoids as probes of the microtubule cytoskeleton. Cell Motil. Cytoskeleton 1998, 39, 73−90. (17) Devaraj, N. K.; Hilderbrand, S.; Upadhyay, R.; Mazitschek, R.; Weissleder, R. Bioorthogonal Turn-On Probes for Imaging Small Molecules inside Living Cells. Angew. Chem., Int. Ed. 2010, 49, 2869− 2872. (18) Lukinavičius, G.; Reymond, L.; D’Este, E.; Masharina, A.; Göttfert, F.; Ta, H.; Güther, A.; Fournier, M.; Rizzo, S.; Waldmann, H.; Blaukopf, C.; Sommer, C.; Gerlich, D. W.; Arndt, H.-D.; Hell, S. 11249

DOI: 10.1021/acsomega.9b01089 ACS Omega 2019, 4, 11245−11250

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(37) Ingber, D. E. Tensegrity I. Cell structure and hierarchical systems biology. J. Cell Sci. 2003, 116, 1157−1173. (38) Ganguly, A.; Yang, H.; Sharma, R.; Patel, K. D.; Cabral, F. The role of microtubules and their dynamics in cell migration. J. Biol. Chem. 2012, 287, 43359−43369. (39) Richard, J. P.; Melikov, K.; Vives, E.; Ramos, C.; Verbeure, B.; Gait, M. J.; Chernomordik, L. V.; Lebleu, B. Cell-penetrating peptides. A reevaluation of the mechanism of cellular uptake. J. Biol. Chem. 2003, 278, 585−590. (40) Mukherjee, S.; Ghosh, R. N.; Maxfield, F. R. Endocytosis. Physiol. Rev. 1997, 77, 759−803. (41) Borowiak, M.; Nahaboo, W.; Reynders, M.; Nekolla, K.; Jalinot, P.; Hasserodt, J.; Rehberg, M.; Delattre, M.; Zahler, S.; Vollmar, A.; Trauner, D.; Thorn-Seshold, O. Photoswitchable Inhibitors of Microtubule Dynamics Optically Control Mitosis and Cell Death. Cell 2015, 162, 403−411.

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