Fluorescence Labeling of Carbon Nanotubes and Visualization of a

Mar 11, 2011 - Biological applications of carbon nanotubes have been hampered by the inability to visualize them using conventional optical microscope...
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Fluorescence Labeling of Carbon Nanotubes and Visualization of a Nanotube-Protein Hybrid under Fluorescence Microscope Shige H. Yoshimura,*,†,§ Shahbaz Khan,†,§ Hiroyuki Maruyama,‡ Yoshikazu Nakayama,‡,§ and Kunio Takeyasu† †

Graduate School of Biostudies, Kyoto University, Yoshida-konoe-cho, Sakyo-ku, Kyoto 606-8501, Japan Graduate School of Engineering, Osaka University, Yamadaoka, Suita 565-0871, Japan § Core Research for Evolutional Science and Technology (CREST), Japan Science and Technology Agency (JST), Japan ‡

ABSTRACT: Biological applications of carbon nanotubes have been hampered by the inability to visualize them using conventional optical microscope, which is the most common tool for the observation and measurement of biological processes. Recently, a number of fluorescence labeling methods for biomolecules and various fluorescence probes have been developed and widely utilized in biological fields. Therefore, labeling carbon nanotubes with such fluorophores under physiological conditions will be highly useful in their biological applications. In this Article, we present a method to fluorescently label nanotubes by combining a detergent and a fluorophore commonly used in biological experiments. Fluorophores carrying an amino group (Texas Red hydrazide or BODIPY FL-hydrazide) were covalently attached to the hydroxyl groups of Tween 20 using carbonyldiimidazole. Fluorescence microscopy demonstrated that nanotubes were efficiently solubilized and labeled by this fluorescently labeled detergent. By using this technique, we also demonstrated multicolor fluorescence imaging of a nanotubeprotein hybrid.

’ INTRODUCTION There has been an increasing number of studies using carbon nanotubes for building nanoscale devices to measure various biological processes and reactions1-4 and to generate efficient carriers for biomolecules in drug delivery systems.5-7 In the application of nanotubes in biological systems, visualization of the nanotubes together with biomolecules, living cells, or both is a significant requirement. The visualization of single carbon nanotubes using a conventional optical microscope, which is the most common tool used in biology, has been difficult because of their small diameters. Although an optical microscope can be used to visualize thick multiwalled nanotubes, it cannot be used to observe small single-walled carbon nanotubes (SWNTs), which are more useful in biological applications. Nanotubes can be routinely visualized using electron microscopes and Raman spectroscopy, but these techniques are not suitable for biological samples and are therefore of limited use for biological applications of nanotubes. Considerable progress has been made recently in fluorescence labeling and observation techniques for biological molecules. A number of fluorophores and fluorescent molecules have been developed to label specific biomolecules that can be observed using various types of fluorescence microscopes. Currently, most biomolecules (DNA, proteins, RNA, peptides, and drugs) can be fluorescently labeled, and their localization, dynamics, and in some cases their function, can be monitored under an optical microscope. Combined with progress in observation and detection devices and technologies (CCD cameras, detectors, etc.), the r 2011 American Chemical Society

fluorescence observation of individual biomolecules is now widely utilized to elucidate their behavior, dynamics, and function at the single-molecule level. There is therefore considerable interest in developing a fluorescence labeling method for carbon nanotubes, especially from the viewpoint of constructing new single-molecule nanotube-protein hybrid devices. Previous studies reported that SWNTs emits fluorescence at near-infrared wavelengths (900-1600 nm).8-10 This property has been utilized for the optical chasing and detection of nanotubes following injection into a living body,10 though its application to single-molecule observation has been limited due to the range of fluorescence emission spectra and weak emission signal. Several studies have reported the fluorescent labeling of nanotubes by polymers and hydrophobic probes. For example, poly(vinylpyrrolidone) coupled to fluorophores wraps around the nanotube and makes it visible under a fluorescence microscope.11 Pyrene is a strong fluorophore excited by UV light (∼350 nm) and has been widely used to solubilize nanotubes and functionalize their sidewalls, allowing the immobilization of proteins.12-17 Unfortunately, because UV irradiation can significantly damage biological molecules, and especially nucleic acids and proteins, UV-activatable fluorophores are not suited for biological applications. Received: December 9, 2010 Revised: February 15, 2011 Published: March 11, 2011 1200

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Biomacromolecules The goal of this study is to establish the method to label nanotubes with fluorophore, which are commonly used in biology, and handle them under a physiological condition together with biomolecules such as proteins. The previous study successfully labeled nanotubes by fluorophores via noncovalent hydrophobic interaction and visualized them in water under an optical microscope.18 However, handling nanotubes in physiological solution (containing ∼130 mM salt) causes a strong association/aggregation of nanotubes as well as nonspecific adsorption of proteins to the surface of the nanotubes because hydrophobic interaction generally becomes stronger as the salt concentration increases. Therefore, it is desirable to develop a procedure to solubilize and label the nanotube in a physiological buffer. In this report, we present a method for simultaneously solubilizing and labeling carbon nanotubes by using a detergent covalently coupled with a variety of fluorophores commonly used in biology.Because of their stability under physiological conditions and their varied fluorescence properties, fluorescently labeled nanotubes can be easily utilized in combination with biomolecules such as proteins.

’ EXPERIMENTAL SECTION Fluorescence Labeling of Nanotubes by FluorophoreLinked Tween 20. Polyoxyethylene sorbitan monolaurate (Tween 20, Sigma-Aldrich) and carbonyldiimidazole (CDI, Thermo Fisher Scientific) were dissolved in 25 mL of dehydrated dimethyl sulfoxide (DMSO, Nacalai Tesque) and incubated for 2 h at 40 °C with constant stirring (∼100 rpm). Excess CDI was removed by extraction with diethylether. The precipitate was redissolved in DMSO (∼1 mL) and extracted with diethylether (∼10 mL) for four times to ensure the complete removal of free CDI. SWNTs synthesized by chemical vapor deposition (CVD) method (Nanocyl s.a.) were dispersed in ultrapure water by ultrasonication. CDI-activated Tween 20 was then added to a final concentration of 10%. The nanotubes were washed with 50 mM sodium borate (pH 7.5) to remove excess Tween 20 and then incubated with fluorophore carrying an amino group (Texas Red hydrazide or BODIPY FL-hydrazide, Invitrogen) at 25 °C for 4 h. Excess fluorophore was removed either by passing the solution through a gel filtration column (Micro Bio-Spin 6, Bio-Rad Laboratories) or by repeating centrifugations and redissolution. Alternatively, CDI-activated Tween 20 was first incubated with fluorophore in 50 mM sodium borate (pH 7.5) at 25 °C for 4 h. The nanotubes were then mixed with fluorophorelinked Tween20. Microscopy. Fluorescence images of the nanotubes were obtained using an inverted fluorescence microscope (IX70, Olympus) with a filter set indicated. The images were captured by a CCD camera (EMCCD, Hamamatsu) and were processed by the software AquaCosmos (Hamamatsu). Transmission electron microscopy (TEM) images of nanotubes were obtained by JEM-2500SE (JEOL) with an accelerating voltage at 90 kV. Preparation of Fluorescent Protein. The coding region of ECFP in pECFP-C1 (Invitrogen) was amplified by PCR to create 50 NcoI and 30 XhoI sites and then inserted into pET29a(þ) vector (Merck Bioscience) to create a hexahistidine-tagged fusion protein, as previously reported.19 The fusion protein was expressed in E. coli cells (BL21CodonPlus RIL, Agilent Technologies, CA) and purified using nickelagarose beads (Ni-NTA, Qiagen) according to the manufacturer’s protocol. In brief, the cells were suspended in lysis buffer (50 mM NaPO4, pH 8.0, 300 mM NaCl, 10 mM imidazole) and disrupted by sonication at 4 °C for 5 min. The solubilized protein was separated from cell debris by centrifugation (10 000g, 30 min, 4 °C), and incubated with Ni-NTA resin with gentle agitation (1 h, 4 °C). The beads were then

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washed with wash buffer (50 mM NaPO4, pH 8.0, 300 mM NaCl, 20 mM imidazole) and histidine-tagged protein was eluted by a discontinuous imidazole gradient from 100 to 500 mM in lysis buffer. The yield and the purity of the purified protein were examined by polyacrylamide gel electrophoresis (SDS-PAGE) and subsequent staining by coomasie brilliant blue.

Coupling of Recombinant Fluorescent Protein to the End of the Nanotubes. Single-walled carbon nanotubes (Nanocyl s.a.) were dispersed on a silicon wafer in isopropanol and then oxidized in air at 400 °C for 1 h. Nanotubes were then collected, dispersed in water by ultrasonication, and incubated with TxRed-coupled Tween 20. After excess Tween 20 was removed, the labeled nanotubes were incubated with 2.5 mg/mL 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC, Sigma-Aldrich) and 2.5 mg/mL sulfo N-hydroxysuccinimide (sulfo-NHS, Thermo Fisher Scientific) in 50 mM 2-(N-morpholino)ethanesulfonic acid (pH 7.0) at 4 °C for 1 h. Following removal of free sulfo-NHS and EDC using a gel filtration column (Micro Bio-Spin 6, Bio-Rad Laboratories), purified ECFP was added and incubated at room temperature for 10 min. After free protein was removed by centrifugation, the nanotubes were observed under a fluorescence microscope (IX70, Olympus).

’ RESULTS AND DISCUSSION Fluorescence Labeling of Nanotubes in Aqueous Solution. Biological applications of nanotubes require solubilization of the nanotubes because hydrophobic surfaces such as glass, silicon, and nanotubes tend to adsorb proteins through hydrophobic interactions and van der Waals forces. Previous studies have reported the solubilization of nanotubes using various surfactants20-24 and direct sidewall modifications,25 coupled to vigorous stirring or ultrasonication. We chose the surfactant, polyoxyethylene sorbitan monolaurate (Tween 20), which is often used as detergent in a wide variety of biochemical experiments, to reduce nonspecific interactions between proteins and between proteins and surfaces without affecting protein function. The procedure for coupling the hydroxyl groups of Tween 20 to a fluorophore carrying an amino group is presented in Figure 1. Tween 20 carries hydroxyl groups at the ends of polyoxyethylene chain and therefore can be directly cross-linked to other functional groups by an appropriate cross-linker, such as CDI. TEM images of nanotubes are shown in Figure 2a. The length and the diameter of the nanotubes ranged from 3 to 6 μm and 2 to 3 nm, respectively. Most of the nanotubes appeared to be single- or double-walled. When the nanotubes were dispersed in 10% Tween 20 solution and sonicated, they were efficiently solubilized into the aqueous phase. The same effect could be seen with CDI-activated Tween 20, indicating that activated Tween 20 retains potent surfactant properties, although a small fraction of nanotubes remained unsolubilized. It should be noted that once the nanotubes were solubilized in water by Tween 20, they remained soluble for several days even after the detergent was removed from the solution, enabling a long-term storage of the solubilized nanotubes. Nanotubes covered with CDI-activated Tween 20 in aqueous phase were reacted with fluorophores carrying an amino group. Two commercially available fluorophores with different emission spectra were tested: Texas Red hydrazide (ex: 596 nm, em: 620 nm) and BODIPY FL-hydrazide (ex: 488 nm, em: 520 nm) (Figure 3). Both fluorophore are often used to label various biomolecules. After removing excess fluorophore, the nanotubes were observed under a conventional fluorescence microscope. 1201

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Figure 3. Chemical structures of the fluorophores utilized in this study: BODIPY FL Hydrazide (ex: 488 nm, em: 520 nm) and Texas Red Hydrazide (ex: 596 nm, em: 620 nm).

Figure 4. Fluorescence images of nanotubes. Fluorescence images of nanotubes labeled with (a) Texas Red and (b) BODIPY FL. Images were captured using an inverted microscope equipped with an EM-CCD camera. For observation of Texas Red and BIDIPY FL, the filter sets used were WMIY2 (excitation: BP545-580 nm, dichroic mirror: 600 nm, emission: LP610 nm) and WMBV2 (excitation: BP400-440 nm, dichroic mirror: 455 nm, emission: LP 475 nm), respectively. Scale bars: 5 μm. Figure 1. Activation and coupling of Tween 20 to amino-carrying compounds. Tween 20 was first treated with carbonyldiimidazole to activate the hydroxyl groups at the ends of polyoxyethylene chain. A compound carrying an amino group was then reacted to generate covalent linkage. The mass spectrometry analysis confirmed that the average number of total oxyethylene groups (n = w þ x þ y þ z) is 20 (data not shown).

Figure 2. TEM images of carbon nanotubes. Nanotubes were observed by TEM (a) before and (b) after oxidation in air. Most of the nanotubes are single- or double-walled. Scale bars: 10 nm.

As shown in Figure 4, clear fluorescence images of the nanotubes could be seen. There was no clear difference in labeling efficiency between the two fluorophores tested: each provided uniform

fluorescence labeling along the nanotubes with the expected emission wavelengths (Figure 4). However, in several cases (7 out of 79), a spiral pattern of fluorescence signal was observed along the nanotube axis, possibly due to a helical alignment of Tween 20 molecules along the nanotube sidewall. The average length of observed fluorescent nanotubes was 5 μm, showing good agreement with observations by electron microscopy before labeling (Figure 2a). A few thick bundles of multiple nanotubes were observed, indicating that each nanotube is welldispersed in the aqueous phase by Tween 20. Alternatively, CDI-activated Tween 20 was reacted with fluorophore before mixing with the nanotubes. In this case, a slight reduction in detergent activity was observed when it was mixed with nanotubes; that is, insoluble particles of nanotubes were visible at the bottom of the tube. This is probably because the replacement of hydroxyl groups with fluorophore reduces the hydrophilic properties of Tween 20. After removing insoluble nanotubes by centrifugation, we removed the supernatant under a fluorescence microscope. The fluorescence image of the nanotubes was very similar to the image described above (data not shown), indicating that the fluorophore can be coupled to Tween 20 either before or after incubation with the nanotubes. Characterization of Labeled Nanotubes in Physiological Conditions and Visualization of Nanotube-Protein Hybrid. The fluorescence properties of the labeled nanotubes were compared in different buffer conditions commonly used in biological experiments. After labeling, the nanotubes were transferred 1202

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based on the TEM image, the diameter of the nanotube was approximately 2 to 3 nm, which allows binding of several proteins simultaneously.

Figure 5. Protein attachment and multicolor imaging of the nanotubeprotein hybrid. Purified ECFP was incubated with Texas Red-labeled oxidized nanotubes in the presence of cross-linker (EDC and sulfoNHS). The fluorescence signal was observed using a conventional fluorescence microscope with two different filter sets: WMIY2 (excitation: BP545-580 nm, dichroic mirror: 600 nm, emission: LP610 nm) for Texas Red and WMBV2 (excitation: BP400-440 nm, dichroic mirror: 455 nm, emission: LP 475 nm) for ECFP. Under our experimental condition, the amount of purified ECFP against nanotubes was kept low to reduce the number of free ECFP signals in the fluorescence image. Under such a condition, ECFP signal was found at only one end of the nanotube and not both ends. Although the addition of a large amount of ECFP increased the probability of colocalization between the nanotube signal and the ECFP signal, it became more difficult to judge whether ECFP is covalently coupled to the nanotube or it simply associates on the nanotube surface. Scale bar: 5 μm.

to 100 mM sodium phosphate (pH 7.4) or 50 mM sodium borate (pH 5.2 and 8.0). The fluorescence signal from the nanotubes observed under a fluorescence microscope did not drastically change in these different conditions, indicating that the fluorophore covalently attached to the surfactant is not affected by these various environments. The labeled nanotubes were also incubated with proteins such as bovine serum albumin (BSA, 1 mg/mL). The fluorescence signal was not affected (data not shown), indicating that these labeled nanotubes are stable in a biological environment. We then conjugated fluorescent protein to the end of labeled nanotubes and simultaneously imaged this nanotube-protein hybrid in two different colors under a fluorescence microscope. The ends of the nanotubes are desirable sites for various modifications, and the attachment of a protein to the end(s) is an important technique for the development of nanotube devices. We have previously reported that the oxidation of a nanotube generates carboxyl groups at its ends, which can be cross-linked to the amino group of a protein using a heterobifunctional crosslinker, carbodiimide (EDC).26 The oxidized nanotubes (Figure 2b) were fluorescently labeled with Tween 20-TexasRed as described above, and purified enhanced cyan fluorescent protein (ECFP, ex: 433 and 453 nm, em: 475 and 501 nm) was attached to the end of the nanotubes using EDC. As shown in Figure 5, ECFP signals were observed at the end of the nanotubes (20 out of 165) using a fluorescence microscope but not on their sidewalls (0 out of 165). Quantification of ECFP signal intensity suggested that, in some cases, multiple ECFP molecules attached to the end. This is probably due to the size of the nanotube tip;

’ CONCLUSIONS We developed a one-step procedure to solubilize and label simultaneously carbon nanotubes by a variety of amine-carrying fluorophores. The application of this procedure is not restricted by the property of the fluorophore (excitation and emission wavelengths) nor by the solution used (buffer, salt, protein, etc.). Because the fluorophore does not directly contact the surface of the nanotube, the fluorescence property of the fluorophore does not seem to be affected by the nanotube surface. This will enable the generation of many combinations of nanotubes with fluorescently labeled biomolecules. We further demonstrated that the ends of these solubilized nanotubes remained active and could react with protein molecules. Nanotubes have been utilized in various biological applications such as biological sensor devices1-4 and scanning probe microscopes.27-31 The fluorescence labeling of nanotubes will allow a wider range of applications, especially in optics-based single-molecule devices. ’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. Tel/Fax: þ81-75-753-7906.

’ ACKNOWLEDGMENT We would like to thank Japan Science and Technology Agency (JST) (Core Research for Evolutional Science and Technology (CREST)) for financial support. We thank Prof. Ohno at Gifu University for providing plasmid encoding the histidine-tagged fusion protein. We thank Mr. Senga (Osaka University) for his help in taking TEM images presented in Figure 2. ’ REFERENCES (1) Star, A.; Gabriel, J. C.; Bradley, K.; Grner, G. Nano Lett. 2003, 3, 459–463. (2) Star, A.; Tu, E.; Niemann, J.; Gabriel, J. C.; Joiner, C. S.; Valcke, C. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 921–926. (3) Besteman, K.; Lee, J.; Wiertz, F. G. M.; Heering, H. A.; Dekker, C. Nano Lett. 2003, 3, 727–730. (4) Chen, R. J.; Choi, H. C.; Bangsaruntip, S.; Yenilmez, E.; Tang, X.; Wang, Q.; Chang, Y. L.; Dai, H. J. Am. Chem. Soc. 2004, 126, 1563–1568. (5) Kam, N. W.; Liu, Z.; Dai, H. J. Am. Chem. Soc. 2005, 127, 12492–12493. (6) Singh, R.; Pantarotto, D.; McCarthy, D.; Chaloin, O.; Hoebeke, J.; Partidos, C. D.; Briand, J. P.; Prato, M.; Bianco, A.; Kostarelos, K. J. Am. Chem. Soc. 2005, 127, 4388–4396. (7) Feazell, R. P.; Nakayama-Ratchford, N.; Dai, H.; Lippard, S. J. J. Am. Chem. Soc. 2007, 129, 8438–8439. (8) O’Connell, M. J.; Bachilo, S. M.; Huffman, C. B.; Moore, V. C.; Strano, M. S.; Haroz, E. H.; Rialon, K. L.; Boul, P. J.; Noon, W. H.; Kittrell, C.; Ma, J.; Hauge, R. H.; Weisman, R. B.; Smalley, R. E. Science 2002, 297, 593–596. (9) Bachilo, S. M.; Strano, M. S.; Kittrell, C.; Hauge, R. H.; Smalley, R. E.; Weisman, R. B. Science 2002, 298, 2361–2366. (10) Cherukuri, P.; Gannon, C. J.; Leeuw, T. K.; Schmidt, H. K.; Smalley, R. E.; Curley, S. A.; Weisman, R. B. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 18882–18886. (11) Didenko, V. V.; Moore, V. C.; Baskin, D. S.; Smalley, R. E. Nano Lett. 2005, 5, 1563–1567. 1203

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