LETTER pubs.acs.org/Langmuir
Dynamic Photocontrol of the Gliding Motility of a Microtubule Driven by Kinesin on a Photoisomerizable Monolayer Surface M. K. Abdul Rahim, Tuyoshi Fukaminato, Takashi Kamei, and Nobuyuki Tamaoki* Research Institute for Electronic Science, Hokkaido University, N20, W10, Kita-ku, Sapporo, Hokkaido 001-0020, Japan
bS Supporting Information ABSTRACT: The gliding motility of microtubules driven by kinesin on the surface of an azobenzene monolayer presenting lysine terminal groups is reversibly and repeatedly altered upon photoisomerization of the monolayer.
’ INTRODUCTION The motor proteins in our bodies perform several mechanical functions initiated by chemical reactions; for example, the transformation of chemical energy to mechanical energy as seen in muscle tissue, the transportation of vesicles or organelles in cells, and the synthesis of energy molecules (e.g., adenosine triphosphate (ATP)). Kinesin is one of the most important linear motor proteins; it transports nanoscale objects along microtubules, rails constructed of tubulins, within cells.1 If we could apply such a function to an artificial molecular system, then it might be possible to use it to transport nanoscale objects precisely between desired positions. The realization of such artificial regulation of a motor protein might open up a new field in nanotechnology.2 Several earlier studies have examined the artificial control of kinesin.3 The ideal control would be one that provides on/off switching of the motility function of kinesin at any desired time and any desired position in space. Light is one of the most appropriate signals for switching because it allows regulation with sufficiently high temporal and spatial resolution. The switching of kinesin’s function from the off state to the on state through the action of light was demonstrated quite a while ago using a caged ATP.4 In contrast, reverse switching from the on state to the off state was only recently demonstrated using a caged peptide having the same structure as the kinesin’s tail, which is known to inhibit the motility of kinesin if it does not bear cargo.5 In that study, the authors demonstrated an 80% reduction in the initial gliding velocity of microtubules on the kinesin surface after the photochemical deprotection of the o-nitrobenzyl protecting r 2011 American Chemical Society
group on the caged peptide. Nevertheless, using light to control the motility of kinesin reversibly, as desired for the complete regulation of the linear motor protein both temporally and spatially, has never previously been demonstrated. In this study, we achieved the repeated regulation of the function of kinesin by the reversible photoisomerization of the underlying monolayer using light of two different wavelengths. For the photoresponsive component, we employed a derivative of azobenzene, one of the most studied photochromic compounds.6 Using this approach, the gliding velocity of the microtubules, driven by kinesin immobilized on a monolayer of the lysinedecorated azobenzene, could be controlled repeatedly between fast and slow modes (with a 15% difference in velocity) upon irradiation with UV and visible light, respectively.7
’ RESULTS AND DISCUSSION The chart presents the chemical structure of azobenzene derivative 1 that we applied to prepare a monolayer on the substrate for the motility experiments. The two amino groups on the terminal lysine moiety were protected with tert-Boc groups. Quartz or glass plates were immersed in a solution of 10% 1 and 90% n-dodecyltriethoxysilane 2 to prepare a monolayer through the formation of siloxane linkages between 1 or 2 and the substrate. Subsequent treatment of the plate with trifluoroacetic Received: May 11, 2011 Revised: August 3, 2011 Published: August 06, 2011 10347
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Langmuir acid led to the deprotection of the tert-Boc groups of 1, leaving the azobenzene layer to present free lysine amino groups.8 Figure 1 displays the absorption spectra of a quartz plate featuring an azobenzene monolayer before and after photoirradiation. Prior to irradiation at 366 nm, a strong absorption band appeared at 380 nm, which we assign to the π π* transition of the substituted E-azobenzene chromophore. Assuming the molar extinction coefficient for the oriented azobenzene molecules in the monolayer to be the same as that in solution, the occupied surface area containing one azobenzene molecule is calculated to be 2.0 nm2 by a comparison of the absorbance of the monolayer at 380 nm and that of the solution (ε = 2.20 104 L mol 1 cm 1). It is known that one alkyl group on the fully covered surface occupies about 0.2 nm2.9 Therefore, the calculated value of 2.0 nm2 corresponds to the surface area of 10 alkylsilane molecules on a fully covered surface. This well matches the azobenzene concentration (1 azobenzene silane in every 10 silanes) applied to monolayer formation, which indicates that the quartz surface is fully covered with the mixture of alkylsilane and azobenzene silane in the expected ratio. After irradiation at 366 nm, the intensity of this absorption band decreased, with the sample reaching its photostationary state (PSS) within 30 s. From the change in the absorbance at 380 nm, it appeared that 40% or more of the azobenzene units had isomerized from E to Z to reach the PSS. Subsequent irradiation at >500 nm led to the recovery of the intensity of the signal at 380 nm; the system reached another PSS within 3 min. We observed similar changes in the absorption spectra of a photoirradiated solution of 1 in CH3CN; these phenomena are typical of the photoisomerization between the E Chart 1. Chemical Structure of 1
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and Z states of azobenzene derivatives.6e,10 Therefore, 1 appears to undergo reversible photoisomerization between its E and Z states in solution and also in the form of a monolayer. For this present study, we used a gliding assay to evaluate the motility of kinesin.11 In this assay, we monitored the gliding of fluorescently labeled microtubules on a substrate decorated with kinesin units using a fluorescence microscope. We prepared flow cells for the motility experiments by fixing a plate presenting a monolayer of 1 to an untreated plate featuring a 100 μm gap using double-sided tape. In most motility experiments, kinesin is typically applied to the substrate in the presence of blocking proteins (e.g., casein) to avoid inactivation.12 In this study, however, we introduced kinesin directly onto the photoresponsive azobenzene monolayer without applying any blocking proteins. First, we confirmed that the microtubules exhibited sufficiently rapid gliding upon kinesin-mediated ATP hydrolysis on the azobenzene monolayer surface in the absence of blocking proteins. Although the length of time that kinesin was catalytically active on the azobenzene monolayer was shorter than that on the corresponding surface featuring blocking proteins, it was long enough to perform all of the motility experiments prior to a serious decrease in activity.13 Next, we investigated the effect of the isomerization of the azobenzene moieties in the monolayer on the motility of the microtubules driven by kinesin-mediated ATP hydrolysis. Just after introducing ATP solution, we irradiated the flow cell alternately with 366 and >500 nm light. Figure 2 shows photographs of microtubules at 0 and 10 s in the motility experiment for the nonirradiated and 366-nm-irradiated flow cell. Under the two irradiation conditions, microtubules move with slightly but statistically significantly different velocities. Figure 3 displays the changes in the gliding velocities of the four states: prior to irradiation (100% E), after irradiation at 366 nm (Z-rich), after subsequent irradiation at >500 nm (E-rich), and after subsequent irradiation at 366 nm (Z-rich). The motility velocity of the microtubule was higher in the Z state of the azobenzene monolayer than in the E state, making it possible to control the velocity repeatedly. The controllable change in the velocity was 13 15% of the initial velocity in the E state, which is confirmed by the experiments with six independent flow cells. In contrast to the results of photoregulation with the deprotected lysine case,
Figure 1. UV vis absorption spectra of the monolayer prepared from 10% compound 1 and 90% n-dodecyltriethoxysilane 2 on a quartz plate after deprotection of the tert-Boc group (left) and in those of 1 in acetonitrile (right). (a) Before irradiation. (b) Photostationary state (PSS) at 366 nm. (c) PSS at >500 nm. The inset shows the absorbance changes at 376 nm after alternating irradiation at 366 and >500 nm over five cycles. Smoothing was done to the original spectra of the monolayer to remove spikelike noise coming from the instrument. 10348
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Langmuir
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Figure 4. Effect of irradiation of the monolayer of 1 and 2 (1:9 molar ratio, after deprotection) on the number of adsorbed microtubules. State 1: Nonirradiated part of the monolayer. State 2: 366-nm-irradiated part (30 s); state 3: after subsequent irradiation at >500 nm for 3 min.
Figure 2. Photographs of rhodamine-labeled microtubules driven by kinesin-mediated ATP hydrolysis on the deprotected lysine azobenzene monolayer. (Top) Prior to irradiation at 0 s (left) and 10 s (right) in the motility experiment. (Bottom) After 366 nm irradiation at 0 s (left) and 10 s (right) in the motility experiment. Red arrows point to the front position of a microtubule at the indicated time. The dotted arrows on the right photographs point to the front positions of the same microtubules at 0 s. Field of view: 82 μm 82 μm.
Figure 3. Changes in microtubule motility upon photoirradiation. The velocity was calculated with the distance of the trace that a terminal of a microtubule passed during 10 s of motility. State 1: Prior to irradiation. State 2: After irradiation at 366 nm for 30 s. State 3: After subsequent irradiation at >500 nm for 3 min. State 4: After subsequent irradiation at 366 nm for 30 s. The number of microtubules measured was 50, 53, 55, and 28 for states 1 4, respectively. The error bars indicate the standard error of the mean.
substantially no change in the velocity of the microtubules upon photoirradiation was detected for the tert-Boc-protected lysine azobenzene, which is confirmed with five independent flow cells (Supporting Information). To understand the origin of motility control through photoisomerization of the azobenzene monolayer, we investigated the affinity of the monolayer surface toward the microtubules in a flow cell, in the absence of kinesin, by using a fluorescence microscope to count the number of attached microtubules. Figure 4 displays the difference in the number of microtubules attached to the monolayer surface with and without irradiation after equilibration. Fewer microtubules were adsorbed on the Z-rich surface after irradiation at 366 nm than were on the 100%
E surface prior to irradiation. After subsequent irradiation at >500 nm, the surface recovered its ability to bind to a greater number of microtubules.14 These results suggest that the Z form of the lysine-presenting azobenzene monolayer surface had a lower affinity for the microtubules than did its E form. Exploiting this difference in affinity made it possible to provide the sticky and less-sticky surfaces for microtubules repeatedly, merely by irradiating it with visible and UV light. There are two possible mechanisms explaining the change in motility of the kinesin/microtubule system upon the photoisomerization of our monolayer surface. One is that photoisomerization induced a change in the interaction between the monolayer surface and the microtubules. The surface of a microtubule is rich in carboxylate groups, providing a net negative charge; the surface of our monolayer presented lysine residues with two free amino groups of positive charge in the buffer solution.15 It is therefore likely that electrostatic attraction occurred between the microtubules and the monolayer surface. We found that the affinity between the microtubules and the monolayer surface decreased upon the E-to-Z photoisomerization of the azobenzene moieties in the monolayer. Although the exact structure of the monolayer remains unclear to us at present, it is possible that isomerization to the Z form of the azobenzene units caused the lysine groups to dive into the monolayer and shield their amino groups from the surface. On such a surface exhibiting affinity toward the microtubules, competition between the driving by kinesin and the fixation by the surface will occur, with slower motility expected for a surface with higher affinity. (For a theoretical analysis of this effect on the motility, see the Supporting Information.) The other possibility is that the photoresponsive monolayer affects the activity of kinesin. It is known that the activity of kinesin in vivo is changed depending on the nature of the surface of the substrate; in extreme cases, kinesin is completely deactivated on some substrates in the absence of a blocking protein.12c Martin et al. reported the reversible control of kinesin activity depending on the doping states of a conduction polymer equipped underneath. They insisted that the ion-pair interaction between the polymer and kinesin was switched by doping and dedoping.3i Our photoresponsive surface in this present study changed its property as observed for the affinity to microtubules upon irradiation; this change in the surface property might have affected the structure of kinesin, resulting in a change in its motility activity. Further experiments will be necessary to elucidate the mechanisms underlying the photoregulated motility of microtubules on this surface. 10349
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Langmuir In summary, the gliding motility of microtubules driven by kinesin on the surface of an azobenzene monolayer presenting lysine terminal groups is reversibly and repeatedly altered upon photoisomerization of the monolayer. The findings can provide a new methodology in nanotechnology to regulate the transportation of nanoobjects precisely in time and space. We expect that some modifications in the chemical structure, such as the use of other amino acids at the terminal, of the photoresponsive molecule with respect to the monolayer would induce a substantial increment in the magnitude of the motility change, which is necessary for actual applications.
’ ASSOCIATED CONTENT
bS
Supporting Information. Experimental details including synthesis, the preparation of a monolayer, optical measurements and photoisomerization of monolayers, the structure of a flow cell, and an adhesion and motility assay. This material is available free of charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION Corresponding Author
*Fax: +81 11 706 9357. Tel: +81 11 706 9356. E-mail: tamaoki@ es.hokudai.ac.jp.
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’ ACKNOWLEDGMENT This work was supported by a grant-in-aid for science research in a priority area “New Frontiers in Photochromism (no. 471)” from the Ministry of Education, Culture, Sports, Science, and Technology (MEXT), Japan. M.K.A.R. acknowledges The Ushio Foundation for a scholarship. ’ REFERENCES (1) (a) Brady, S. T. Nature 1985, 317, 73–75. (b) Vale, R. D.; Reese, S. T.; Sheetz, M. P. Cell 1985, 42, 39–50. (2) (a) Agarwal, A.; Hess, H. Prog. Polym. Sci. 2010, 35, 252–277. (b) Goel, A.; Vogel, V. Nat. Nanotechnol. 2008, 3, 465–475. (c) van den Heuvel, M. G. L.; Dekker, C. Science 2007, 317, 333–336. (d) Fischer, T.; Hess, H. J. Mater. Chem. 2007, 17, 943–951. (3) (a) Stracke, R.; B€ohm, K. J.; Wollweber, L.; Tuszynski, J. A.; Unger, E. Biochem. Biophys. Res. Commun. 2002, 293, 602–609. (b) van den Heuvel, M. G. L.; Butcher, C. T.; Lemay, S. G.; Diez, S.; Dekker, C. Nano Lett. 2005, 5, 235–241. (c) Platt, M.; Muthukrishnan, G.; Hancock, W. O.; Williams, M. E. J. Am. Chem. Soc. 2005, 127, 15686–15687. (d) Hiratsuka, Y.; Tada, T.; Oiwa, K.; Kanayama, T.; Uyeda, T. Q. P. Biophys. J. 2001, 81, 1555–1561. (e) van den Heuvel, M. G. L.; Butcher, C. T.; Smeets, R. M. M.; Diez, S.; Dekker, C. Nano Lett. 2005, 5, 1117–1122. (f) Limberis, L.; Magda, J. J.; Stewart, R. J. Nano Lett. 2001, 1, 277–280. (g) Du, Y. Z.; Hiratsuka, Y.; Taira, S.; Eguchi, M.; Uyeda, T. Q. P.; Yumoto, N.; Kodaka, M. Chem. Commun. 2005, 2080–2082. (h) Ionov, L.; Stamm, M.; Diez, S. Nano Lett. 2006, 6, 1982–1987. (i) Martin, B. D.; Vlea, L. M.; Soto, C. M.; Whitaker, C. M.; Gaber, B. P.; Ratna, B. Nanotechnology 2007, 18, 055103. (j) Tucker, R.; Katira, P.; Hess, H. Nano Lett. 2008, 8, 221–226. (4) (a) Higuchi, H.; Muto, E.; Inoue, Y.; Yanagida, T. Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 4395–4400. (b) Hess, H.; Clemmens, J.; Qin, D.; Howard, J.; Vogel, V. Nano Lett. 2001, 1, 235–239. (5) Nomura, A.; Uyeda, T. Q. P.; Yumoto, N.; Tatsu, Y. Chem. Commun. 2006, 3588–3590. (6) (a) Ercole, F.; Davis, T. P.; Evans, R. A. Polym. Chem. 2010, 1, 37–54. (b) Tamaoki, N.; Kamei, T. J. Photochem. Photobiol., C 2010, 11, 47–61. (c) Seki, T.; Nagano, S. Chem. Lett. 2008, 37, 484–489. 10350
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