Carbon Nanotubes: Methods and Protocols (Methods in Molecular Biology, 625)

Methods

in

Molecular Biology™

Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK



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Carbon Nanotubes Methods and Protocols

Edited by

Kannan Balasubramanian Max-Planck-Institut für Festkörperforschung, Stuttgart, Germany

Marko Burghard Max-Planck-Institut für Festkörperforschung, Stuttgart, Germany

Editors Kannan Balasubramanian, Ph.D. Max-Planck-Institut für Festkörperforschung Stuttgart Germany [email protected]

Marko Burghard, Ph.D. Max-Planck-Institut für Festkörperforschung Stuttgart Germany [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-60761-577-4 e-ISBN 978-1-60761-579-8 DOI 10.1007/978-1-60761-579-8 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2010920245 © Humana Press, a part of Springer Science+Business Media, LLC 2010 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or ­dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, ­neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Humana Press is a part of Springer Science+Business Media (www.springer.com)

Preface Since their discovery in 1991, carbon nanotubes (CNTs) have had an enormous impact in materials science. More recently, CNTs have successfully entered the fields of molecular biology, biomedicine, and bioanalytical chemistry. Much of the increasing interest in CNTs is owed to their rare combination of high chemical stability and exceptional optical and electrical properties. Another major factor that has promoted the utilization of CNTs in biological sciences is their unique structure. In fact, their high surface-to-volume ratio and high aspect ratio render them close-to-ideal candidates as active components of biosensors, or as “nanosyringes” enabling the injection of drugs or biological markers into living cells. Over the last couple of years, a wide variety of high-quality CNTs have become commercially available, a fact that has strongly stimulated the recent development of biologyrelated CNT applications. However, the obtainable material often differs in purity, agglomeration state, as well as the length and diameter distribution of the tubes, all of which have a profound influence on important parameters like the tubes’ dispersability and surface properties. It is hence highly desirable to make reliable protocols, which include as many details on the used nanotubes as possible, available to a wide range of readers coming from different fields. We strongly hope that the present collection of protocols will contribute to the achievement of common standards and help to avoid discrepancies in future biology-related CNT studies. This book is organized into five parts. The first part focuses on CNT chemical functionalization approaches, which are required to tackle a major obstacle for using CNTs in biology and medicinal chemistry, namely their inherent hydrophobic character and the resulting lack of solubility in most solvents compatible with the biological milieu. CNT functionalization based upon covalent or non-covalent schemes has proven to be highly effective for enhancing the water solubility of the nanotubes and thus transforming their biocompatibility profile. At the same time, non-covalent functionalization often serves as the basis for further purification of the tubes via centrifugation or chromatography. The second part is devoted to toxicity studies of CNTs. In the meanwhile, it is well established that various types of functionalized CNTs exhibit a capacity to be taken up by a wide range of cells and are able to traffic through different cellular barriers. Recent studies have demonstrated that the cellular uptake of CNTs is largely independent of the nature and density of the appended functional groups, which paves the way for nanotube-tube delivery of a broad range of agents, including proteins, DNA, and synthetic polymers. The intracellular traffic of functionalized CNTs will be the topic of the third part, encompassing three different methodologies. Part 4 deals with modified CNT networks as scaffolds for cell growth, an area that attracts increasing attention due to the future perspective of designing therapies for CNS regeneration or the development of neurochip devices. Finally, Part 5 provides protocols related to CNT-based biosensors, with emphasis on amperometric detection principles. One central topic here is the use of tube coatings to enhance the selectivity of the sensor response toward specific analytes. Stuttgart, Germany

Kannan Balasubramanian Marko Burghard

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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Part I  Functionalization   1 Non-covalent Attachment of Proteins to Single-Walled Carbon Nanotubes . . . . . Luís F.F. Neves, Ta-Wei Tsai, Naveen R. Palwai, David E. Martyn, Yongqiang Tan, David W. Schmidtke, Daniel E. Resasco, and Roger G. Harrison   2 Covalent Conjugation of Multi-walled Carbon Nanotubes with Proteins . . . . . . . Changqing Yi, Suijian Qi, Dawei Zhang, and Mengsu Yang   3 Covalently Linked Deoxyribonucleic Acid with Multi-walled Carbon Nanotubes: Synthesis and Characterization . . . . . . . . . . . . . . . . . . . . . . . Weiwei Chen, Changqing Yi, Tzang Chi-Hung, Shuit-Tong Lee, and Mengsu Yang   4 Temperature and pH-Responsive “Smart” Carbon Nanotube Dispersions . . . . . . Dan Wang and Liwei Chen

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Part II Toxicity   5 Effects of Carbon Nanotubes on the Proliferation and Differentiation of Primary Osteoblasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41 Dawei Zhang, Changqing Yi, Suijian Qi, Xinsheng Yao, and Mengsu Yang   6 Carbon Nanotube Uptake and Toxicity in the Brain . . . . . . . . . . . . . . . . . . . . . . . 55 Leying Zhang, Darya Alizadeh, and Behnam Badie   7 In Vitro and In Vivo Biocompatibility Testing of Functionalized Carbon Nanotubes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 Gianni Ciofani, Vittoria Raffa, Orazio Vittorio, Alfred Cuschieri, Tommaso Pizzorusso, Mario Costa, and Giuseppe Bardi   8 Real-Time Monitoring of Cellular Responses to Carbon Nanotubes . . . . . . . . . . . 85 Qingxin Mu, Shumei Zhai, and Bing Yan   9 Reducing Nanotube Cytotoxicity Using a Nano-Combinatorial Library Approach . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Qiu Zhang, Hongyu Zhou, and Bing Yan 10 DNA Damage by Carbon Nanotubes Using the Single Cell Gel Electrophoresis Technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Olga Zeni and Maria Rosaria Scarfì

Part III Trafficking 11 Assessment of Cellular Uptake and Cytotoxicity of Carbon Nanotubes Using Flow Cytometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123 Khuloud T. Al-Jamal and Kostas Kostarelos

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12 Cell Trafficking of Carbon Nanotubes Based on Fluorescence Detection . . . . . . . 135 Monica H. Lamm and Pu Chun Ke 13 Carbon Nanotubes as Intracellular Carriers for Multidrug Resistant Cells Studied by Capillary Electrophoresis–Laser-Induced Fluorescence . . . . . . . . 153 Ruibin Li, Hanfa Zou, Hua Xiao, and Renan Wu

Part IV  Scaffolds 14 Carbon Nanotube-Based Neurochips . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171 Moshe David-Pur, Mark Shein, and Yael Hanein 15 Effect of Carbon Nanotubes on HepG2 Adhesion and Spreading . . . . . . . . . . . . . 179 Suijian Qi, Changqing Yi, Dawei Zhang, and Mengsu Yang

Part V  Biosensors 16 Enzymatic Detection Based on Carbon Nanotubes . . . . . . . . . . . . . . . . . . . . . . . Martin Pumera 17 Carbon Nanotube Biosensors Based on Electrochemical Detection . . . . . . . . . . . Martin Pumera 18 Biosensors Based on Carbon Nanotube-Network Field-Effect Transistors . . . . . . . Cristina C. Cid, Jordi Riu, Alicia Maroto, and F. Xavier Rius 19 Detection of Biomarkers with Carbon Nanotube-Based Immunosensors . . . . . . . Samuel Sánchez, Esteve Fàbregas, and Martin Pumera 20 Carbon Nanotube Biosensors with Aptamers as Molecular Recognition Elements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hye-Mi So, Dong-Won Park, Hyunju Chang, and Jeong-O Lee Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors Khuloud T. Al-Jamal  •  Nanomedicine Laboratory, Centre for Drug Delivery Research, The School of Pharmacy, University of London, London, UK Darya Alizadeh  •  Division of Neurosurgery, City of Hope National Cancer Center, Duarte, CA, USA Behnam Badie  •  Division of Neurosurgery, City of Hope National Cancer Center, Duarte, CA, USA Giuseppe Bardi  • Institute of Neurosciences, CNR, Via Moruzzi, Pisa, Italy Hyunju Chang  • NanoBio Fusion Research Center, Korea Research Institute of Chemical Technology, Jang-dong 100, Eusung-gu, Daejeon 305–343, Korea Liwei Chen  • Department of Chemistry and Biochemistry, Ohio University, Athens, OH, USA; Suzhou Institute of Nano Tech and Nano Bionics, Chinese Academy of Science, Suzhou, Jiangsu, P. R. China Weiwei Chen  • Department of Physics & Materials Science, City University of Hong Kong, KLT, Hong Kong Tzang Chi-Hung  • Department of Biology and Chemistry, City University of Hong Kong, KLT, Hong Kong Cristina C. Cid  • Department of Analytical and Organic Chemistry, Universitat Rovira i Virgili, Tarragona, Catalonia, Spain Gianni Ciofani  • Scuola Superiore Sant’Anna, Pisa, Italy Mario Costa  • Institute of Neurosciences, CNR, Via Moruzzi, Pisa, Italy Alfred Cuschieri  • Scuola Superiore Sant’Anna, Pisa, Italy Moshe David-Pur  • School of Electrical Engineering, Tel-Aviv University, Tel-Aviv, Israel Esteve Fàbregas  • Sensors and Biosensors Group, Analytical Chemistry Department, UAB.Edifici Cn, Bellaterra, Spain Yael Hanein  • School of Electrical Engineering, Tel-Aviv University, Tel-Aviv, Israel Roger G. Harrison  • School of Chemical, Biological and Materials Engineering, and Carbon Nanotube Technology Center, University of Oklahoma, Norman, OK, USA Pu Chun Ke  • Department of Physics and Astronomy, Clemson University, Clemson, SC, USA Kostas Kostarelos  • Nanomedicine Laboratory, Centre for Drug Delivery Research, The School of Pharmacy, University of London, London, UK Monica H. Lamm  • Department of Chemical and Biological Engineering, Iowa State University, Ames, IA, USA Jeong-O Lee  • NanoBio Fusion Research Center, Korea Research Institute of Chemical Technology, Jang-dong 100, Eusung-gu, Daejeon 305–343, Korea Shuit-Tong Lee  • Physics & Materials Science, City University of Hong Kong, KLT, Hong Kong Ruibin Li  • National Chromatographic R & A Center, Dalian Institute of Chemical Physics, Chinese Academy of Sciences, Dalian, P.R. China

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Alicia Maroto  • Department of Analytical and Organic Chemistry, Universitat Rovira i Virgili, Tarragona, Catalonia, Spain David E. Martyn  • School of Chemical, Biological and Materials Engineering, and Carbon Nanotube Technology Center, University of Oklahoma, Norman, OK, USA Qingxin Mu  • School of Chemistry and Chemical Engineering, Shandong University, Jinan, P.R. China; St. Jude Children’s Research Hospital, Memphis, TN, USA Luís F.F. Neves  • School of Chemical, Biological and Materials Engineering, and Carbon Nanotube Technology Center, University of Oklahoma, Norman, OK, USA Naveen R. Palwai  • School of Chemical, Biological and Materials Engineering, and Carbon Nanotube Technology Center, University of Oklahoma, Norman, OK, USA Dong-Won Park  • NanoBio Fusion Research Center, Korea Research Institute of Chemical Technology, Jang-dong 100, Eusung-gu, Daejeon 305–343, Korea Tommaso Pizzorusso  • Institute of Neurosciences, CNR, Via Moruzzi, Pisa, Italy Martin Pumera  • Biomaterial Systems Group, Biomaterials Center and International Center for Materials Nanoarchitectonics (MANA), Tsukuba, Ibaraki, Japan Suijian Qi  • Department of Biology and Chemistry, City University of Hong Kong, KLT, Hong Kong Vittoria Raffa  • Scuola Superiore Sant’Anna, Pisa, Italy Daniel E. Resasco  • School of Chemical, Biological and Materials Engineering, and Carbon Nanotube Technology Center, University of Oklahoma, Norman, OK, USA Jordi Riu  • Department of Analytical and Organic Chemistry, Universitat Rovira i Virgili, Tarragona, Catalonia, Spain F. Xavier Rius  • Department of Analytical and Organic Chemistry, Universitat Rovira i Virgili, Tarragona, Catalonia, Spain Samuel Sánchez  • Sensors and Biosensors Group, Analytical Chemistry Department, UAB. Edifici Cn, Bellaterra, Spain Maria Rosaria Scarfì  • CNR-Institute for Electromagnetic Sensing of Environment (IREA), Naples, Italy David W. Schmidtke  • School of Chemical, Biological and Materials Engineering, and Carbon Nanotube Technology Center, University of Oklahoma, Norman, OK, USA Mark Shein  • School of Electrical Engineering, Tel-Aviv University, Tel-Aviv, Israel Hye-Mi So  • NEMS Bio Team, National NanoFab Center, 355 Gwahangno, Yuseong-gu Daejeon 304–806, Korea Yongqiang Tan  • School of Chemical, Biological and Materials Engineering, and Carbon Nanotube Technology Center, University of Oklahoma, Norman, OK, USA Ta-Wei Tsai  • School of Chemical, Biological and Materials Engineering, and Carbon Nanotube Technology Center, University of Oklahoma, Norman, OK, USA Orazio Vittorio  • Scuola Superiore Sant’Anna, Pisa, Italy; Department of Oncology, Transplantation and Advanced Technologies in Medicine, University of Pisa, Pisa, Italy Dan Wang  • Department of Chemistry and Biochemistry, Ohio University, Athens, OH, USA

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Renan Wu  • National Chromatographic R & A Center, Dalian Institute of Chemical Physics, Chinese Academy of Sciences, Dalian, P.R. China Hua Xiao  • National Chromatographic R & A Center, Dalian Institute of Chemical Physics, Chinese Academy of Sciences, Dalian, P.R. China Bing Yan  • School of Chemistry and Chemical Engineering, Shandong University, Jinan, P.R. China; St. Jude Children’s Research Hospital, Memphis, TN, USA Mengsu Yang  • Department of Biology and Chemistry, City University of Hong Kong, KLT, Hong Kong Xinsheng Yao  • Department of Natural Products Chemistry, Shenyang Pharmaceutical University, Shenyang, China Changqing Yi  •  Biotechnology & Health Centre, Shenzhen Virtual University Park, Shenzhen, China, City University of Hong Kong, KLT, Hong Kong Olga Zeni  • CNR-Institute for Electromagnetic Sensing of Environment (IREA), Naples, Italy Shumei Zhai  • School of Chemistry and Chemical Engineering, Shandong University, Jinan, P.R. China Dawei Zhang  • Department of Biology and Chemistry, City University of Hong Kong, KLT, Hong Kong Leying Zhang  • Division of Neurosurgery, City of Hope National Cancer Center, Duarte, CA, USA Qiu Zhang  • School of Chemistry and Chemical Engineering, Shandong University, Jinan, Shandong, P. R. China Hongyu Zhou  • School of Chemistry and Chemical Engineering, Shandong University, Jinan, P.R. China; St. Jude Children’s Research Hospital, Memphis, TN, USA Hanfa Zou  • National Chromatographic R & A Center, Dalian Institute of Chemical Physics, Chinese Academy of Sciences, Dalian, P.R. China

Part I Functionalization

Chapter 1 Non-covalent Attachment of Proteins to Single-Walled Carbon Nanotubes Luís F.F. Neves, Ta-Wei Tsai, Naveen R. Palwai, David E. Martyn, Yongqiang Tan, David W. Schmidtke, Daniel E. Resasco, and Roger G. Harrison Abstract A method for the non-covalent attachment of proteins to single-walled carbon nanotubes (SWNTs) is described. In this method, the protein is adsorbed to SWNTs that are suspended using sodium cholate, a surfactant and bile salt. The sodium cholate is then removed by dialysis with retention of the protein on the SWNTs. This method has resulted in good protein loadings and good retention of protein activity. Key words: Non-covalent attachment, Adsorption, Dialysis, Proteins, Single-walled carbon nanotubes, Sodium cholate, Surfactant

1. Introduction Single-walled carbon nanotubes (SWNTs) have important optical, thermal, mechanical, and electronic properties (1), and are being developed for applications in various biological systems (2). For some of these biological applications, it is necessary to attach proteins to the SWNTs, for example, to target the SWNTs to specific cells or to construct a biosensor. In this chapter, we provide detailed information on a method that we have used successfully to attach proteins to SWNTs by adsorption, a non-covalent form of attachment (3). This method is advantageous in that important features in the UV-vis-NIR adsorption spectra of the SWNTs are preserved, which is critical in applications where it is desired, for example, that the SWNTs strongly absorb energy when NIR radiation is applied. One problem with direct covalent

K. Balasubramanian and M. Burghard (eds.), Carbon Nanotubes: Methods and Protocols, Methods in Molecular Biology, vol. 625, DOI 10.1007/978-1-60761-579-8_1, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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attachment of molecules to the SWNTs is that this method results in the complete elimination of the UV-vis-NIR adsorption bands, which is thought to be due to the SWNT’s p system being disrupted (4, 5). The adsorption method we will describe was first used for proteins by Graff et  al. (6). The first step in this method is to completely suspend the SWNTs in an aqueous solution of sodium cholate, which is a surfactant and a bile salt. After centrifugation, the protein is added, and the suspension is dialyzed using a dialysis membrane that will retain the protein but allow the sodium cholate to pass through. The suspension is centrifuged again, and the supernatant containing the suspended SWNTs with protein adsorbed is retained. This method for us has given good protein loading and good retention of biological activity for horseradish peroxidase (HRP, MW = 40  kDa) (3) and glucose oxidase (MW =  160 kDa) (unpublished data): HRP loading of 2 mg protein/mg SWNTs and 98% retention of native enzyme activity; glucose oxidase loading of 22 mg protein/mg SWNTs and 87% retention of native enzyme activity. Atomic force microscopy (AFM) analysis was performed on SWNTs with HRP adsorbed, and it is possible to visualize the HRP adsorbed on the nanotubes (Fig. 1) (3). In addition, for SWNTs with HRP adsorbed, there was retention of a substantial fraction of the NIR absorption at 980 nm (Fig. 2) (3). Other methods that have been used to adsorb proteins on SWNTs are the organic solvent displacement method (7) and the

Fig. 1. AFM images showing SWNT with sodium cholate and HRP protein. (a) SWNT in sodium cholate after sonication and centrifugation. Arrows indicate solid sodium cholate with a height of 1.0–1.5 nm (does not include 0.8 nm SWNT height). (b) SWNT/protein after dialysis and centrifugation. Arrows indicate protein associated with SWNT. The height is 3.8–6.0 nm (does not include 0.8 nm SWNT height). Reproduced from ref. (3) with permission from IOP Publishing Ltd.

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Fig. 2. UV-vis-NIR absorption spectra of the pristine SWNTs suspended in sodium cholate and the SWNT/HRP complex after the final centrifugation. Reproduced from ref. (3) with permission from IOP Publishing Ltd.

aqueous sonication method (8); however, these methods gave problems with complete dispersion of SWNTs and with the maintenance of protein activity or structure.

2. Materials 2.1. SWNT Preparation and Suspension

1. Purified and freeze dried CoMoCAT SG65 SWNTs, rich in (6,5) type with an average diameter of 0.75  nm, were provided by Southwest Nanotechnologies, Inc. (Norman, OK). 2. Sodium cholate (Sigma-Aldrich) was used as a dispersant. 3. A horn sonicator equipped with a microtip of 3.2  mm in diameter was used (CPX750, Cole Parmer). 4. An ultracentrifuge was used to centrifuge the SWNT suspension (Optima XL series preparative ultracentrifuge, Beckman Coulter).

2.2. Adsorption of Proteins on SWNTs

1. Sodium phosphate (20 mM, pH 7.4) for buffering. 2. Dialysis membranes (10  kDa and 100  kDa, Spectrum Laboratories). 3. Bradford protein assay kit (Quick Start, Bio-Rad).

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3. Methods 3.1. SWNT Preparation and Suspension

1. Add 3.0 mg of SWNTs to 7 mL of a 2 wt % sodium cholate solution and sonicate at a power of 7 W for 30 min. 2. Centrifuge the resulting suspension at 29,600 × g for 30 min (see Note 1).

3.2. Adsorption of Proteins on SWNTs

1. Mix sodium phosphate buffer with the SWNT suspension. 2. Add 20 mg of protein to the suspension at 4°C. 3. Dialyze the solution at 4°C for 12 h against 2 L of sodium phosphate buffer using a 10 kDa dialysis membrane to remove unadsorbed sodium cholate (see Note 2). 4. Transfer the solution to a 100  kDa dialysis membrane and then dialyze at 4°C against 2 L of sodium phosphate buffer, with a change of the 2 L of buffer at 2, 4, 16, and 24 h from the start of dialysis (see Note 3). Carry out the final dialysis for 4 h. 5. Centrifuge the solution at 29,600 × g for 1 h at 4°C. 6. Save the supernatant (see Note 4). 7. Perform the Bradford protein assay in order to quantify the protein concentration in the solution (see Note 5). Also, measure the protein concentration in the dialysate (the solution on the outside of the dialysis membrane). 8. Measure the absorbance at 800 nm of the initial suspension (SWNTs dispersed using sodium cholate) and the final suspension (SWNT/protein complex) for determination of the SWNT concentration (see Note 6).

4. Notes 1. The centrifugation step that is performed during the production of the suspension is important, since it eliminates nanotubes aggregates. The presence of aggregates in suspension is undesirable; the existence of aggregates can contribute to significant changes in the properties of the suspension. During the centrifugation step, it is important to use at least 29,600 × g centrifugal force when using single-walled carbon nanotubes, in order to assure a good nanotube dispersion of single-walled nanotubes, and to remove any aggregated nanotubes. 2. The reason for the use of a 10  kDa dialysis membrane is because the sodium cholate (MW = 431  Da) will be able to

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pass through the membrane pores freely and therefore will permit the removal of unadsorbed sodium cholate. Proteins with molecular weights higher than 10 kDa will be retained inside the membrane. 3. A membrane with a larger pore size (100  kDa molecular weight cutoff) is used in the second dialysis, in order to remove unadsorbed protein through the membrane. Extensive dialysis is used to ensure complete removal of unadsorbed protein; less dialysis can possibly be used for some proteins, especially low molecular weight proteins. The removal of unadsorbed protein can be evaluated by measuring the protein concentration in the dialysate. For low protein concentrations, it may be necessary to use a micro protein assay (for example, the micro BCA protein assay from Pierce). It is important to note that the membrane pore size should always be larger than the protein molecular size. Different membrane pore sizes are commercially available for the dialysis of proteins with a range between 10 and 300  kDa. It is also important to note that higher than a 100  kDa molecular weight cutoff of the membrane can lead to some loss of nanotubes through the membrane. 4. The initial suspension (SWNT dispersed using sodium cholate) and the final suspension (SWNT/protein complex) can be analyzed by atomic force microscopy (see the AFM images in Fig.  1 for the adsorption of HRP using this procedure). 5. The amount of protein used gives a relatively low coverage of the SWNT surface area. For example, using the weights of SWNTs and protein in Subheading 3 and assuming the protein is HRP, the ratio of the number of protein molecules to the number of six-carbon groups in the SWNTs is calculated to be 0.012. Assuming the protein is globular, the diameter of the protein is approximately 7.5 times the diameter of a SWNT. These calculations are verified by the AFM image of HRP adsorbed on SWNTs shown in Fig.  1.1, where the coverage by the protein on the SWNTs is relatively sparse and the protein is much larger than a SWNT. Seven and a half times the SWNT average height of 0.75 nm is 5.6 nm, which falls within the 3.8–6.0 nm height of the HRP protein measured. 6. A wavelength of 800 nm is used to measure the concentration of the SWNTs, since bands are not present on the absorption spectra at this wavelength (see Fig.  2 for SWNTs with HRP adsorbed). A calibration curve can be made from a plot of absorbance at 800 nm versus SWNT concentration.

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Acknowledgments This work was supported by the U.S. Department of Energy-Basic Energy Sciences (DE-FG02-06ER64239), the U.S. Department of Defense Breast Cancer Research Program (W81XWH07-1-0536), and the Foundation for Science and Technology of Portugal (Luis Neves).

References 1. Han J (2005) In: Meyyappan M (ed) Structures and properties of carbon nanotubes. CRC, Moffett Field, CA, pp 2–24 2. Lin Y, Taylor S, Li H, Fernando KAS, Qu L, Wang W, Gu L, Zhou B, Sun YP (2004) Advances toward bioapplications of carbon nanotubes. J Mater Chem 14:527–541 3. Palwai NR, Martyn DE, Neves LFF, Tan Y, Resasco DE, Harrison RG (2007) Retention of biological activity and near-infrared absorbance upon adsorption of horseradish peroxidase on single-walled carbon nanotubes. Nanotechnology 18:235601/1-235601/5 4. Buffa F, Hu H, Resasco DE (2005) Side-wall functionalization of single-walled carbon nanotubes with 4-hydroxymethylaniline followed by polymerization of e-caprolactone. Macromolecules 38:8258–8263

5. Bahr JL, Yang J, Kosynkin DV, Bronikowski MJ, Smalley RE, Tour JM (2001) Functionalization of carbon nanotubes by electrochemical reduction of aryl diazonium salts: a bucky paper electrode. J Am Chem Soc 123:6536–6542 6. Graff RA, Swanson JP, Barone PW, Baik S, Heller DA, Strano MS (2005) Achieving individual-nanotube dispersion at high loading in single-walled carbon nanotube composites. Adv Mater 17:980–984 7. Karajanagi SS, Vertegel AA, Kane RS, Dordick JS (2004) Structure and function of enzymes adsorbed onto single-walled carbon nanotubes. Langmuir 20:11594–11599 8. Matsuura K, Saito T, Okazaki T, Ohshima S, Yumura M, Iijima S (2006) Selectivity of watersoluble proteins in single-walled carbon nanotube dispersions. Chem Phys Lett 429:497–502

Chapter 2 Covalent Conjugation of Multi-walled Carbon Nanotubes with Proteins Changqing Yi, Suijian Qi, Dawei Zhang, and Mengsu Yang Abstract  Linkage of proteins to carbon nanotubes (CNTs) is fundamentally important for applications of CNTs in medicinal and biological fields, as well as in biosensor or chemically modulated nanoelectronic devices. In this contribution, we provide a detailed protocol for the synthesis and characterization of covalent CNT-protein adducts. Functionalization of multiwalled carbon nanotubes (MWCNTs) with proteins has been achieved by the initial carboxylation of MWCNTs followed by amidation with the desired proteins. Attenuated total reflection Fourier transform infrared (ATR-FTIR) and X-ray photoelectron spectroscopy (XPS) measurements validated the presence of a covalent linkage between MWCNTs and proteins. The visualization of proteins on the surface of MWCNTs was furthermore achieved using atomic force microscopy (AFM). The protein-conjugated nanocomposites can also be assembled into multidimensional addressable heterostructures through highly specific biomolecular recognition system (e.g., antibody–antigen). Keywords: Carbon nanotubes, Covalent conjugation, Protein, Antibody, ATR-FTIR, XPS, AFM

1. Introduction The unique properties of nanomaterials (NMs) in combination with the biorecognition abilities of proteins offer particularly exciting opportunities in molecular imaging (1–3), therapy (4, 5), biomolecule delivery (6, 7), and design of functional nanodevices (8–12). The loss or retention of the native structure of proteins upon their conjugation onto NMs provides an additional variable for controlling NM assembly – intercomponent spacing (13). Though biomolecules, such as DNA and proteins, can be linked to nanotubes via noncovalent interactions (14, 15), the use of covalent chemistry is expected to provide better stability, accessibility, and selectivity (16, 17).

K. Balasubramanian and M. Burghard (eds.), Carbon Nanotubes: Methods and Protocols, Methods in Molecular Biology, vol. 625, DOI 10.1007/978-1-60761-579-8_2, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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Herein, we report a protocol for covalent conjugation of proteins to carbon nanotube (CNT) surface, which has been achieved by the initial carboxylation of CNTs followed by amidation with the desired proteins. This two-step chemical method employs mild conditions and results in tethering of the organic functionality through a covalent bond. This protocol can also be employed to functionalize CNTs with various proteins and amines, including primary amines and secondary amines. The proteinconjugated nanocomposites can be further assembled into multidimensional addressable heterostructures through highly specific biomolecular recognition system, such as antibody– antigen interation.

2. Materials 2.1. Carboxylation of MWCNTs

  1. Pristine MWCNTs, prepared by the chemical vaporization deposition (CVD) method (Nanotech Port, China). 2. 10-µm pore size PTFE filter paper (Advantec MFS, Inc.). 3. Ultrasonic cleaning bath (Electron Microscopy Sciences).

2.2. Amidation

  1. Mouse monoclonal IgG (Santa Cruz Biotechnology). 2. 1-ethyl-3-(3-dimethyl-aminopropyl) carbodiimide (EDC) (Sigma-Aldrich). 3. N-hydroxysuccinimide (NHS) (Sigma-Aldrich). 4. Phosphate buffer solution (PBS): 0.1 mol/L, pH 7.4.

2.3. Equipment

   1. Transmission electron microscope (TEM), Tecnai 12 (Philips). 2. Copper grids with formvar film (Electron Microscopy Sciences). 3. ULVAC-PHI 5802 XPS system (Kanagawa, Japan). 4. FTIR spectrometer Spectrum One (Perkin Elmer). 5. FTIR microscope equipped with a HgCdTe detector cooled with liquid nitrogen (i-Series). 6. Multimode atomic force microscope (Veeco Instruments).

3. Methods (See Notes 1 and 2) 3.1. Carboxylation

  1. Pristine MWCNT powder was oxidized in a 3:1 mixture of concentrated H2SO4 (98%) and HNO3 (69%) at 70°C for 4 h (18–20) and filtered through a 10-µm pore size PTFE filter paper. Or

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Pristine MWCNTs were refluxed in 4  M HNO3 for 24  h and filtered through a 10-µm pore size PTFE filter paper. 2. After filtration, the refluxed MWCNTs were exposed to 1 M HCl and sonicated for about 30 min. 3. The carboxylated MWCNTs were filtered, and washed with deionized water and dried in air. 3.2. Amidation

    1. MWCNT-COOH and mouse monoclonal IgG was mixed in a tube at the ratio 1:10. 2. A mixture solution of 0.40 M 1-ethyl-3-(3-dimethyl-aminopropyl) carbodiimide (EDC) and 0.10  M N-hydroxy succinimide (NHS) was added to initiate cross-linking reactions between carboxyl groups on MWCNTs and amine groups in IgG. 3. The reaction tubes were rotated at room temperature for 1 h. 4. After centrifugation and decanting supernate, mouse IgG functionalized MWCNTs were re-suspended in PBS buffer.

3.3. TEM Characterization

    1. 5 mL of dilute aqueous sample was spotted onto a grid and left for 90 s. 2. Lightly touch one edge of the grid with filter paper to wipe off moisture. 3. Grids were then allowed to air dry prior to analysis. 4. TEM: Observe the samples on grids in electron microscopy with an accelerating voltage of 120 kV. Figure 1 shows the results obtained.

3.4. XPS Measurements (See Notes 3 and 4)

    1. Parameter setting: (a) Pressure ranges are as follows: 2 × 10−6 mbar (fast entry chamber), 4 × 10−8 mbar (preparation chamber), and 4 × 10−9 mbar (sample analysis chamber). (b) High transmission FAT mode, 14.12 keV, 25 mA, Al Ka (1,486.7  eV), was used for the analysis at 90° electron take off angle for normal non-charging samples (45° for the charging samples). (c) The analyzer slit width was set for 0.8 mm, and the resulting overall energy resolution was 0.35 eV. 2. The SCIENTA software was used for data acquisition and data analysis. 3. The binding energy of the C1s of graphite, 284.5  eV (±0.35 eV energy resolution of the spectrometer at the settings employed) was taken as the reference. 4. Prior to individual elemental scans, a survey scan was taken for all the samples in order to detect the elements present.

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Fig. 1. (a) and (b) TEM images of the carboxylated MWCNTs

5. The obtained XPS spectra of CNT-protein adduct are listed in Fig. 2. The C1s XPS of the oxidized nanotubes in Fig. 2a shows a large peak at 284.4 eV from the nanotubes, a smaller peak at 286.4 eV, and a well-separated peak at 288.8 eV. The peak at 288.8 eV is attributed to the carbonyl group in the carboxylic acid group. The peak at 286.4, 2 eV higher than the main peak, is attributed to C atoms in ether-like linkages. The corresponding N(1s) spectrum shows no signal above the detection limit of the instrument, even with extensive signal averaging. After conjugation with proteins, the modified tubes were characterized by XPS after briefly warming to 75°C in an ultrahigh vacuum to remove any residual physically adsorbed proteins. Compared to the main bulk C1s at 284.4  eV, the resulting C1s photoelectron spectrum shows some narrowing of the bulk peak (Fig. 2b). We notice that there is no significant intensity near 288.8 eV. The absence of intensity at 288.8 eV is important because the C1s binding energy of carboxylic groups is expected to decrease significantly when a carboxylic acid group is converted to a carbonyl amide, which is at 288.45  eV. Carbon atoms in carboxylic acid groups and in carbonyl amide groups typically have C1s binding energies ~4.0  eV and ~3.0  eV higher, respectively, than C atoms in alkanes (21). Thus, the changes we observe in the C1s spectrum support the formation of a carbonyl amide linkage to the nanotubes. The N1s spectrum shows a peak

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Fig. 2. XPS of chemically modified CNTs. C1s spectrum of CNT-COOH (a); C1s spectrum of protein functionalized CNTs (b), showing elimination of the carboxylic peak and increased peak from the amide group. N1s spectrum of protein functionalized MWCNTs (c)

with a binding energy of 399.7 eV (Fig. 2c). Previous studies have shown that amides have binding energies in the range of ~399.5–400.2 eV (21). Therefore, the peak energies are consistent with the formation of the amide bonds. 3.5. ATR-FTIR Measurements (See Notes 5 and 6)

  1. Sample preparation: MWCNT-protein adducts were grinded to a fine powder. 2. Sample loading: The ZnSe through top-plate of the horizontal ATR was covered by a fine layer of the grinded sample, avoiding air bubble formation on the crystal surface. 3. Parameters setting: A detector gain of 1 and a speed of the moving mirror of 0.6 cm−1 were employed.

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4. Spectra measurements: The spectra were obtained in the absorbance mode from 4,000 to 700 cm−1 by accumulating 100 scans, working with a spectral resolution of 2 cm−1. 5. Control: Absorbance spectra were corrected versus a spectrum of distilled water, obtained in the same instrumental conditions. 6. Figure 2.3 shows the typical ATR-FTIR spectra of MWCNTCOOH before and after functionalization by IgG. The absorption peak at 1,569 cm−1 indicates the presence of amide bond which comes from the covalent linkage between CNTs and IgG through the functional groups. 3.6. AFM Measurements

1. MWCNT-protein adducts suspension: Suspend MWCNTprotein adducts in PBS to a concentration of 1 mg/mL. This concentration provides convenient coverage for AFM imaging and may be used for a variety of similar size samples. 2. Prepare mica: Cleave a fresh mica surface by first pressing some adhesive tape against the top mica surface, then peeling off the tape. Glue mica to a small puck (e.g., using epoxy). 3. Deposit sample solution on mica: Deposit 50 mL of protein solution on the freshly cleaved mica. 4. Sample to bind to substrates: Allow 20–30 min for the sample to bind to the mica substrate. Binding time may vary with different samples (it can be up to 24 h). 5. Rinse unbound sample: Rinse the sample with a large quantity of buffer to remove unbound protein. 6. AFM measurements: AFM measurements were performed under ambient conditions using a NanoScope V Controller

Fig.  3. ATR-FTIR spectra of CNTs. The absorption peak at 1,569  cm−1 indicates the presence of amide linkage between MWCNTs and IgG

Covalent Conjugation of Multi-walled Carbon Nanotubes with Proteins

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with a multimode atomic force microscope in tapping mode with standard 125 mm single-crystal silicon cantilevers. 7. Figure 4 shows the herein obtained AFM images: MWCNTs appear as bright lines in the image and the circle particles represent bound proteins. Proteins are connected to the sidewall of nanotubes, indicating that oxidization took place in the defect sites of sidewalls. From the observations of several samples, we conclude that protein attachment mainly occurred at nanotube sidewalls, because chemical functionalization occurred primarily at the sidewalls. This AFM image is similar to that of cytochrome c-functionalized purified SWCNTs which was taken by Davis and co-workers (14).

Fig.  4. (a) and (b) AFM (tapping mode) images of protein-MWCNT adducts. MWCNTs appear as bright lines and the circle particles represent bound proteins

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4 Notes 1. Unless stated otherwise, water used in this protocol was Milli-Q deionized water. 2. Unless stated otherwise, the CNT suspensions should be prepared in D.I water and followed by ultrasonication for 30 min. 3. Use only polyethylene gloves. Other gloves may contain silicones that can contaminate the surface. 4. Make sure everything, used to handle or store your samples, is clean (tweezers, etc.). It is recommended to have a dedicated set of clean tools for handling your samples. In particular, take care to avoid grease, oils, and silicone contaminants around your tools and work area. A general cleaning protocol that often works is to clean the utensils that will handle samples with the following solvents (in this order): Hexanes, Methylene chloride, Methanol, and Acetone. 5. The sample must be in direct contact with the ATR crystal, because the evanescent wave or bubble only extends beyond the crystal 0.5–5 mm. 6. The refractive index of the crystal must be significantly greater than that of the sample or else internal reflectance will not occur – the light will be transmitted rather than internally reflected in the crystal.

Acknowledgments The financial support of Key Laboratory Funding Scheme of Shenzhen Municipal Government, BTC operation fund (CityU project No. 9683001) and City University of Hong Kong (Project No. 7002100) are gratefully acknowledged.

References 1. Loo C, Hirsch L, Lee MH, Chang E, West J, Halas N, Drezek R (2005) Gold nanoshell bioconjugates for molecular imaging in living cells. Opt Lett 30:1012–1014 2. Medintz IL, Uyeda HT, Goldman ER, Mattoussi H (2005) Quantum dot bioconjugates for imaging, labelling and sensing. Nat Mater 4:435–446

3. Lewis JD, Destito G, Zijlstra A, Gonzalez MJ, Quigley JP, Manchester M, Stuhlmann H (2006) Viral nanoparticles as tools for intravital vascular imaging. Nat Med 12:354–360 4. Loo C, Lowery A, Halas N, West J, Drezek R (2005) Immunotargeted nanoshells for integrated cancer imaging and therapy. Nano Lett 5:709–711

Covalent Conjugation of Multi-walled Carbon Nanotubes with Proteins 5. Kam NWS, O’Connell M, Wisdom JA, Dai HJ (2005) Carbon nanotubes as multifunctional biological transporters and near-infrared agents for selective cancer cell destruction. Proc Natl Acad Sci USA 102:11600–11605 6. Pantarotto D, Briand JP, Prato M, Bianco A (2004) Translocation of bioactive peptides across cell membranes by carbon nanotubes. Chem Commun 16–17 7. Kam NWS, Jessop TC, Wender PA, Dai HJ (2004) Nanotube molecular transporters: internalization of carbon nanotube-protein conjugates into mammalian cells. J Am Chem Soc 126:6850–6851 8. Connolly S, Fitzmaurice D (1999) Programmed assembly of gold nanocrystals in aqueous solution. Adv Mater 11:1202–1205 9. Caswell KK, Wilson JN, Bunz UHF, Murphy CJ (2003) Preferential end-to-end assembly of gold nanorods by biotin-streptavidin connectors. J Am Chem Soc 125:13914–13915 10. Lee JA, Govorov AO, Dulka J, Kotov NA (2004) Bioconjugates of CdTe nanowires and Au nanoparticles: Plasmon-exciton interactions, luminescence enhancement, and collective effects. Nano Lett 4:2323–2330 11. Lee J, Govorov AO, Kotov NA (2005) Bioconjugated superstructures of CdTe nanowires and nanoparticles: multistep cascade Forster resonance energy transfer and energy channeling. Nano Lett 5:2063–2069 12. Wang S, Mamedova N, Kotov NA, Chen W, Studer J (2002) Antigen/antibody immunocomplex from CdTe nanoparticle bioconjugates. Nano Lett 2:817–822 13. Srivastava S, Verma A, Frankamp BL, Rotello VM (2005) Controlled assembly of protein-

14.

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18.

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nanoparticle composites through protein surface recognition. Adv Mater 17:617–621 Azamian BR, Davis JJ, Coleman KS, Bagshaw CB, Green MLH (2002) Bioelectrochemical single-walled carbon nanotubes. J Am Chem Soc 124:12664–12665 Shim M, Kan NWS, Chen RJ, Dai HJ (2002) Functionalization of carbon nanotubes for biocompatibility and biomolecular recognition. Nano Lett 2:285–288 Chen WW, Tzang CH, Tang JX, Yang MS, Lee ST (2005) Covalently linked deoxyribonucleic acid with multiwall carbon nanotubes: synthesis and characterization. Appl Phys Lett 86:103114 Sarah EB, Cai W, Lasseter TL, Weidkamp KP, Hamers RJ (2002) Covalently bonded adducts of deoxyribonucleic acid (DNA) oligonucleotides with single-wall carbon nanotubes: synthesis and hybridization. Nano Lett 2:1413–1417 Yi CQ, Fong CC, Zhang Q, Lee ST, Yang MS (2008) The structure and function of ribonuclease A upon interacting with carbon nanotubes. Nanotechnology 19:095102 Yi CQ, Fong CC, Chen WW, Qi SJ, Tzang CH, Lee ST, Yang MS (2007) Interactions between carbon nanotubes and DNA polymerase and restriction endonucleases. Nano technology 18:025102 Zhang DW, Yi CQ, Zhang JC, Chen Y, Yao XS, Yang MS (2007) Effects of carbon nanotubes on the proliferation and differentiation of primary osteoblasts. Nanotechnology 18: 475102 Moulder JF, Stickle WF, Sobol PE, Bomben KD (1992) Handbook of X-ray photoelectron spectroscopy. Perkin-Elmer, Eden Prairie, MN

Chapter 3 Covalently Linked Deoxyribonucleic Acid with Multi-walled Carbon Nanotubes: Synthesis and Characterization Weiwei Chen, Changqing Yi, Tzang Chi-Hung, Shuit-Tong Lee, and Mengsu Yang Abstract In this chapter, a multi-step protocol for covalently linking functionalized multi-walled carbon nanotubes (MWCNT) to deoxyribonucleic acid (DNA) oligonucleotides is provided. X-ray photoelectron spectroscopy (XPS) is used to characterize the initially formed amine-terminated MWCNTs, to which DNA is covalently anchored. Atomic force microscopy (AFM) investigation of the DNA–MWCNT conjugates reveals that the chemical functionalization occurs at both the ends and sidewalls of the nanotubes. The described methodology represents an important step toward the realization of DNA-guided self-assembly for carbon nanotubes. Key words: Carbon nanotubes, Covalent conjugation, DNA, ATR-FTIR, XPS, AFM

1. Introduction The discovery of carbon nanotubes in 1991 (1) and their subsequent production in bulk quantities (2) have paved the way to the exploration of the physical, chemical, and biological properties of single-walled carbon nanotubes (SWCNTs) and multiwalled carbon nanotubes (MWCNTs). Due to their unique electronic, chemical, and mechanical properties, carbon nanotubes (CNTs) have shown great promises in biosensing, tissue engineering, and biomedical applications (3–5). Many of the interesting and unique properties of nanoscale materials are realized when they are integrated into more complex assemblies (6, 7). In biology, the highly selective binding between complementary sequences of deoxyribonucleic acid (DNA) plays the central role in genetic replication. This selectivity can, in principle, K. Balasubramanian and M. Burghard (eds.), Carbon Nanotubes: Methods and Protocols, Methods in Molecular Biology, vol. 625, DOI 10.1007/978-1-60761-579-8_3, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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be used to assemble a wide range of materials, by forming adducts between DNA and the materials of interest (8). While biomolecules like DNA can be linked to nanotubes via noncovalent interactions (9, 10), the use of covalent chemistry is expected to provide better stability, accessibility, and selectivity during competitive hybridization (11). In this chapter, a multi-step protocol to covalently link functionalized MWCNTs to deoxyribonucleic acid (DNA) oligonucleotides is demonstrated. And we combine the use of X-ray photoelectron spectroscopy (XPS) and atomic force microscopy (AFM) to achieve the chemical analysis of nanotube functionalization, as well as the direct visualization of the DNA–MWCNT adducts that were obtained from our developed multi-step method.

2. Materials 2.1. Carboxylation of MWCNTs

1. Pristine MWCNTs, prepared by the chemical vaporization deposition (CVD) method (Nanotech Port, China). 2. 10-µm pore size PTFE filter paper (Advantec MFS, Inc.). 3. Ultrasonic cleaning bath (Electron Microscopy Sciences). 4. Milli-Q system (Millipore).

2.2. CNT Functionalization with Poly-l-lysine

1. Poly-l-lysine (Sigma-Aldrich). 2. 1-[3-(dimethyl aminopropyl)]-3-ethylcarbodiimide hydrochloride (Sigma-Aldrich). 3. N-hydroxysuccinimide (NHS) (Sigma-Aldrich). 4. EtONH4 (Sigma-Aldrich). 5. Phosphate buffer solution (PBS): 0.1 M, pH 7.4.

2.3. Linkage of DNA Strands to MWCNTs

1. A pair of primers, HBB-1 (5¢ Amine-AGGGTTGGCCAAT­ CTACTCC-3¢) and HBB-2 (5¢ Amine-TCTCCCCTTCCTA­ TGACATGA-3¢) (Invitrogen). 2. PCR reaction solution containing genomic DNA, the pair of primers, reaction buffer, MgCl2, dNTPs, and Taq DNA polymerase (Roche, Basel Switzerland). 3. Biophotometer (Eppendorf, Germany).

2.4. Equipment

1. ULVAC-PHI 5802 XPS system (Kanagawa, Japan). 2. Multimode atomic force microscope (Veeco Instruments). 3. Mica substrates (Ted Pella, Inc.).

Covalently Linked Deoxyribonucleic Acid with Multi-walled Carbon Nanotubes

3. Methods (See Notes 1 and 2 and Fig. 1) 3.1. Carboxylation of MWCNTs

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1. Pristine MWCNT powder was oxidized in a 3:1 mixture of concentrated H2SO4 (98%) and HNO3 (69%) at 70°C for 4 h and filtered through a 10-µm pore size PTFE filter paper. Or Pristine MWCNTs were refluxed in 4 M HNO3 for 24 h and filtered through a 10-µm pore size PTFE filter paper. 2. After filtration, the refluxed MWCNTs were exposed to 1 M HCl and sonicated for about 30 min. 3. The carboxylated MWCNTs were filtered, washed with deionized (D.I) water, and dried in air.

3.2. Functionalization of CNTs with Poly-l-lysine

1. The purified, oxidized MWCNTs were suspended in 0.4  mL 1-[3-(dimethyl aminopropyl)]-3-ethylcarbodiimide hydrochloride and 0.1 mL N-hydroxysuccinimide in an ultrasonic bath. 2. After centrifugation, 300 µL poly-l-lysine (0.1%) was added and the mixture was stirred for 1 h at 25°C. 3. This dispersion was centrifuged and washed two times with EtONH4 (pH = 8.5) to remove excess poly-l-lysine. 4. After rinsing with D.I water for three times, the mixture was placed in a 55°C oven for 15 min to form the amine-terminated nanotubes.

Fig. 1. Scheme showing the steps involved in the fabrication of covalently linked DNAnanotube adducts. Reprinted with the permission from ref. (14), copyright 2005 American Institute of Physics

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3.3. Linkage of DNA Strands to MWCNTs

1. Linking DNA strands to the nanotube requires specially prepared DNA strands. An amine-terminal labeled 684 base pairs DNA was amplified from human b-hemoglobin (HBB) gene by polymerase chain reaction (PCR). 2. A pair of primer, HBB-1 (5¢ Amine-AGGGTTGGC­ CAATCTACTCC-3¢) and HBB-2 (5¢ Amine-TCTCCC­ CTTCCTATGACATGA-3¢), was designed to amplify the 5¢ amine labeled 684 base pairs PCR product from the template of exon 1 + 2 of the HBB gene. 3. Each 50 µL PCR reaction solution contained 100–200 ng of genomic DNA, 0.2–0.4 µM of the pair of primers, 1× reaction buffer, 1.5 mM MgCl2, 200 µM dNTPs, and 5.0 U Taq DNA polymerase. 4. After the reaction mixture was set up in a thin-walled PCR tube, the thermal cycling was carried out as follows: 95°C for 2 min, followed by 30 cycles of denaturation at 94°C for 30 s, annealing at 60°C for 30 s and extension at 72°C for 30 s, and a last extension step of 5 min at 72°C. 5. The PCR products were precipitated with 0.2 × volume of 3 M sodium acetate, pH = 4.5, and 2 × volume of 100% ethanol, and washed with 75% ethanol. 6. The purified PCR products were quantified by a photometer. 7. Poly-l-lysine-treated nanotubes were then mixed with 30 µL amine labeled DNA (100  ng/µL), followed by shaking at 26°C for 30 min. 8. After baking at 80°C, the product was rehydrated through water vapor for 30 s. 9. The sample was immersed into a mixture of methanol and acetic acid (3:1) for 5 min and baked again for 30 min. 10. Steps 7 and 8 were repeated twice to ensure that the incorporated DNA was removed from the DNA–MWCNTs adduct.

3.4. XPS Analysis (See Notes 3 and 4)

1. Parameter setting: (a) Pressure ranges are as follows: 2 × 10−6 mbar (fast entry chamber), 4 × 10−8 mbar (preparation chamber), and 4 × 10−9 mbar (sample analysis chamber). (b) High transmission FAT mode, 14.12 keV, 25 mA, Al Ka (1,486.7  eV) was used for the analysis at 90° electron take off angle for normal non-charging samples (45° for the charging samples). (c) The analyzer slit width was set for 0.8 mm and the resulting overall energy resolution was 0.35 eV. 2. The SCIENTA software was used for data acquisition and data analysis.

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3. The binding energy of the C1s of graphite, 284.5 eV (±0.35 eV energy resolution of the spectrometer at the settings employed) was taken as the reference. 4. Prior to individual elemental scans, a survey scan was taken for all the samples in order to detect the elements present. 5. The obtained XPS spectra of oxidized nanotubes are listed in Fig. 2. The spectrum shows a large peak at 284.4 and 285 eV

Fig. 2. XPS of chemically modified nanotubes. (a) C (1s) spectrum of oxidized nanotubes (upper ), along with fit to three peaks. C (1s) spectrum of oxidized nanotubes functionalized with poly-l-lysine (lower ), showing elimination of the carboxylic peak and increased peak from the amide group. (b) N(1s) spectrum of oxidized nanotubes functionalized with poly-l-lysine (upper ); N (1s) spectrum of oxidized nanotubes (lower ), showing no detectable N (1s) signal. Reprinted with the permission from ref. (14), copyright 2005 American Institute of Physics

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from the nanotubes, a smaller peak at 286.4  eV, and a well-separated peak at 288.6  eV. The peak at 288.6  eV is attributed to the carbonyl group in the carboxylic acid group. The smaller peak at 285 eV is attributed to the satellite peaks arising from excitation of electronic transitions within the nanotubes. The peak at 286.4, 2 eV higher than the main peak, is attributed to C atoms in ether like linkages. The corresponding N(1s) spectrum (Fig. 2b) shows no signal above the detection limit of the instrument. 6. The amine-terminated nanotubes were characterized by XPS after briefly warming to ~75°C in an ultrahigh vacuum to remove any residual physically adsorbed amines. Compared to the main bulk C(1s) at 284.4 eV (as in Fig. 3.2a upper figure), the resulting C(1s) photoelectron spectrum (Fig. 3.2a lower one) shows some narrowing of the bulk peak and three shoulders at 285.05, 286.3, and 288.15 eV. There is no significant intensity near 288.6 eV. The absence of intensity at 288.6 eV is important because the C(1s) binding energy of carboxylic groups is expected to decrease significantly when a carboxylic acid group is converted to a carbonyl amide, which is at 288.15 eV. The shoulders at 285.05 and 286.3 eV are both attributed to the C atoms in alkanes; the higher 286.3 eV is due to electron donation from the adjacent N atom. Carbon atoms in carboxylic acid groups and in carbonyl amide groups typically have C(1s) binding energies, ~4.0  eV and ~3.0  eV higher, respectively, than C atoms in alkanes (12). Thus, the changes observed in the C(1s) spectrum support the formation of a carbonyl amide linkage to the nanotubes. The N(1s) spectrum (Fig. 3.2b lower figure) shows a peak with a binding energy of 399.8 eV. Previous studies have shown that amides have binding energies in the range of ~399.5–400.2 eV (12, 13). Therefore, the peak energies are consistent with the formation of the product depicted in Fig. 3.1a lower figure. 3.5. AFM Characterization

1. DNA–MWCNT adducts suspension: Suspend DNA–MWCNT adducts in PBS to a concentration of 1 mg/mL. This concentration provides convenient coverage for AFM imaging and may be used for a variety of similar size samples. 2. Prepare mica: Cleave a fresh mica surface by first pressing some adhesive tape against the top mica surface, then peeling off the tape. Glue mica to a small puck (e.g., using epoxy). 3. Deposit sample solution on mica: Deposit 50  mL of DNA– MWCNT adducts solution on the freshly cleaved mica with 10 mM MgCl2. 4. Sample to bind to substrates: Allow 20–30 min for the sample to bind to the mica substrate. Binding time may vary with different samples (it can be up to 24 h).

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Fig. 3. (a) and (b) AFM (tapping mode) images of DNA–MWCNT adducts. MWCNTs appear as bright lines and the paler strands represent bound DNA. Reprinted with the permission from ref. (14), copyright 2005 American Institute of Physics

5. Rinse unbound sample: Rinse the sample with a large quantity of buffer to remove unbound protein. 6. AFM measurements: AFM measurements were performed under ambient conditions in tapping mode with standard 125 mm single-crystal silicon cantilevers. 7. As shown in Fig. 3a, at the tip site and the middle site of the carbon nanotube, there are two features, which can be attributed to DNA strands. Carbon nanotubes appear as bright lines in the image. While in Fig. 3b, DNA strands are connected to the sidewall of nanotubes, indicating that oxidization took place in the defect sites of sidewalls. From the observations of several samples, we conclude that DNA attachment occurred at nanotube ends and sidewalls, because chemical functionalization occurred primarily at the ends and sidewalls.

4 Notes 1. Unless stated otherwise, water used in this protocol was D.I water. 2. Unless stated otherwise, the CNT suspensions should be prepared in D.I water and followed by ultrasonication for 30 min. 3. Use only polyethylene gloves. Other gloves may contain silicones that can contaminate the surface. 4. Make sure everything used to handle or store your samples is clean (tweezers, etc.). It is recommended to have a dedicated

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set of clean tools for handling your samples. In particular, take care to avoid grease, oils, and silicone contaminants around your tools and work area. A general cleaning protocol that often works is to clean the utensils that will handle samples with the following solvents (in this order): Hexanes, Methylene chloride, Methanol, Acetone.

Acknowledgments The financial support of Key Laboratory Funding Scheme of Shenzhen Municipal Government, BTC operation fund (CityU Project No. 9683001) and City University of Hong Kong (Project No. 7002100) are gratefully acknowledged. References 1. Iijima S (1991) Helical microtubules of graphitic carbon. Nature 354:56–58 2. Ebbesen TW, Ajayan PM (1992) Large-scale synthesis of carbon nanotubes. Nature 358: 220–222 3. Chen RJ, Zhang YG, Wang DW, Dai HJ (2001) Noncovalent sidewall functionalization of single-walled carbon nanotubes for protein immobilization. J Am Chem Soc 123:3838–3839 4. Heller DA, Baik S, Eurell TE, Strano MS (2005) Single-walled carbon nanotube spectroscopy in live cells: towards long-term labels and optical sensors. Adv Mater 17: 2793–2799 5. Gao LZ, Nie L, Wang TH, Qin YJ, Guo ZX, Yang DL, Yan XY (2006) Carbon nanotube delivery of the GFP gene into mammalian cells. Chembiochem 7:239–242 6. Elghanian R, Storhoff JJ, Mucic RC, Letsinger RL, Mirkin CA (1997) Selective colorimetric detection of polynucleotides based on the distance-dependent optical properties of gold nanoparticles. Science 277:1078–1081 7. Perez JM, O’Loughin T, Simeone FJ, Weissleder R, Josephson L (2002) DNAbased magnetic nanoparticle assembly acts as a magnetic relaxation nanoswitch allowing screening of DNA-cleaving agents. J Am Chem Soc 124:2856–2857

8. Soto CM, Srinivasan A, Ratna BR (2002) Controlled assembly of mesoscale structures using DNA as molecular bridges. J Am Chem Soc 124:8508–8509 9. Guo Z, Sadler PJ, Tsang SC (1998) Immobilization and visualization of DNA and proteins on carbon nanotubes. Adv Mater 10:701–703 10. Shim M, Kan NWS, Chen RJ, Dai HJ (2002) Functionalization of carbon nanotubes for biocompatibility and biomolecular recognition. Nano Lett 2:285–288 11. Sarah EB, Cai W, Lasseter TL, Weidkamp KP, Hamers RJ (2002) Covalently bonded adducts of deoxyribonucleic acid (DNA) oligonucleotides with single-wall carbon nanotubes: synthesis and hybridization. Nano Lett 2: 1413–1417 12. Moulder JF, Stickle WF, Sobol PE, Bomben KD (1992) Handbook of X-ray photoelectron spectroscopy. Perkin-Elmer, Eden Prairie, MN 13. Lin Z, Strother T, Cai W, Cao X, Smith LM, Hamers RJ (2002) DNA attachment and hybridization at the silicon (100) surface. Langmuir 18:788–796 14. Chen WW, Tzang CH, Tang JX, Yang MS, Lee ST (2005) Covalently linked deoxyribonucleic acid with multiwall carbon nanotubes: synthesis and characterization. Appl Phys Lett 86:103114

Chapter 4 Temperature and pH-Responsive “Smart” Carbon Nanotube Dispersions Dan Wang and Liwei Chen Abstract Carbon nanotubes (CNTs) are a family of all-carbon quasi one-dimensional nanomaterials that are highly hydrophobic and typically aggregated in bundles. Recent accomplishments in dispersing CNTs in aqueous solutions open possibilities for their new applications in biomedicine. In many occasions, biological and biomedical applications demand an actuation mechanism; thus, it is highly desirable to control the dispersion and aggregation of CNTs in aqueous solutions with external stimuli. Here, we report two “smart” single-walled CNT (SWNT) aqueous dispersions that respond to temperature and pH changes through environment-responsive polymers, poly (N-isopropylacrylamide) (PNIPAAm) and Poly-l-lysine (PLL). Key words: Single-walled carbon nanotube, Dispersion, Aggregation, Temperature-responsive, pH-responsive, poly (N-isopropylacrylamide), Poly-l-lysine, Atomic force microscopy, Fluorescence

1. Introduction The unique all carbon quasi one-dimensional structure of SWNTs has intrigued much fundamental research on their mechanical, thermal, and electrical properties as well as potential applications in nanocomposites, field emission displays, and molecular electronics (1–5). Recent introduction of individually dispersed SWNTs to aqueous and biology compatible media has opened new frontiers of carbon nanotubes in biology and nanomedicine (6–8). Generally speaking, the surface of as-produced SWNTs is highly hydrophobic; and thus SWNTs exist in aggregated bundles. SWNT dispersions in organic or aqueous solvents can be obtained by either covalent functionalization or non-covalent solublization, with the latter being more favored for the preservation of nanotube structure and properties. Since non-covalent

K. Balasubramanian and M. Burghard (eds.), Carbon Nanotubes: Methods and Protocols, Methods in Molecular Biology, vol. 625, DOI 10.1007/978-1-60761-579-8_4, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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interactions with SWNT sidewalls (p–p stacking, van-der-Waals interactions, and hydrophobic interactions) do not require specificity and directionality, a surprisingly large variety of small molecule surfactants and water-soluble amphiphilic polymers are suitable for this purpose (9–17). Multi-functional polymer surfactants are also exploited for the separation (14, 16–18), alignment (19, 20), and hierarchical assembly of SWNTs (21), as well as for attaching fluorescent chromophore and biologically active cargos to SWNTs (22, 23). Many biomedical applications involve active processes, for example, drug and/or gene delivery involves active release of load at target locations; photothermal and photodynamic therapies involve light-induced actuation: heating and photochemistry, respectively. Utilization of amphiphilic polymer surfactants in SWNT dispersion enables potential actuation mechanisms via selected multi-functional polymers (24). Here, we discuss two “smart” SWNT dispersions that respond to temperature and pH changes using poly (N-isopropylacrylamide) (PNIPAAm) and poly-l-lysine (PLL). The action in response to temperature or pH stimuli is the aggregation of SWNTs in the dispersion. Controlled aggregation of SWNT may help clogging local blood vessel, “deliver” SWNT along with attached load by precipitating from circulation, or enhance photothermal effects. Therefore, the two environmentally responsive SWNT dispersions presented here are pioneering examples of SWNT actuation in biocompatible environments, and they may lead to more sophisticated biotechnologies in the future. PNIPAAm has long been known as a temperature responsive polymer (25). The structure of PNIPAAm contains balanced hydrophilicity from amide bonds and hydrophobicity from hydrocarbon main chain and isopropyl groups on side chains. At temperatures lower than the lower critical solution temperature (LCST, ~33°C for PNIPAAm), PNIPAAm assumes an extended chain conformation in which N and O atoms in amid bonds form H-bonding with surrounding water molecules. When the temperature is higher than the LCST, water molecules are released to maximize entropy while amid groups form intra-chain H-bonding among themselves; thus, the polymer chain takes a coiled and compact conformation (26–28). The extended conformation of PNIPAAm at low temperature allows the amphiphilic polymer chain to cover a large area of SWNT surface in dispersion. Raising temperature causes the chain conformation to shrink and thus exposes hydrophobic SWNT surface to water. This provides a driving force for SWNTs, which are dispersed at low temperatures, to aggregate into small bundles. PLL is an amphiphilic polyelectrolyte that contains both hydrophobic hydrocarbon moieties and partially protonated primary amine groups. SWNTs interact with PLL in aqueous

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environment because of two interactions: one is hydrophobic interaction between PLL hydrocarbon linker moieties (–C4H8–) and SWNT sidewalls; the other is cation-p interaction between protonated amine groups and the p-electron system of SWNTs (26). The cation-p interaction is completely turned off at pH value equal to or higher than the isoelectric point of lysine (9.7) due to the deprotonation of –NH3+ groups. The hydrophobic interaction between –C4H8– linker and SWNT is also affected by pH via changes in the secondary structure of PLL. PLL chain adopts the a-helix conformation at high pH but changes to uncoiled conformations in acidic or neutral pH due to the electrostatic repulsion among side chain cations (29, 30). Since both interaction mechanisms between PLL and SWNT respond to the pH change of the solution, the PLL–SWNT dispersion is expected and indeed observed to be pH sensitive.

2. Materials 1. HiPCO SWNT powders, purified grade (Carbon Nano­ technologies, Inc., Houston, TX). 2. Poly (N-isopropylacrylamide), molecular weight 20,000– 25,000 (Aldrich). 3. 0.1 % (w/v) Poly-l-lysine solution in H2O (Sigma). 4. Sonicator VCX 130 (Sonix, Newtown, CT). 5. Atomic force microscopy (AFM) microscope MFP3D (Asylum Research, Santa Barbara, CA). 6. AFM probes NSC15/AlBS (MikroMasch USA, Wilsonville, OR) with resonance frequency around 325 KHz. 7. Fluorescence NS1 NanoSpectralyzer (Applied NanoFluo­ rescence, LLC, Houston, TX). 8. Spectropolarimeter Jasco-715 (Jasco Inc., Easton, MD).

3. Methods 3.1. SWNT Dispersion in PNIPAAm 3.1.1. Preparation of PNIPAAm–SWNT Dispersion

1. 10  mg/ml of PNIPAAm solution is prepared by dissolving 30 mg PNIPAAm in 2.4 ml water and then adding 0.6 ml of 0.1 M NaOH solution (see Note 1). 2. Add ~1 mg of HiPCO SWNT powder to 3 ml of 10 mg/ml PNIPAAm solution. This mixture is sonicated for 90 min in an ice-water bath at a power level of 130 W (Fig. 1).

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Fig. 1. Experimental setup for SWNT sonication: (1) sonication probe; (2) polymer and SWNTs; (3) ice-water bath

Fig.  2. Absorption spectrum of PNIPAAm–SWNT dispersion. Inset: the structure of PNIPAAm

3. The solution is centrifuged for 5 min at 800 × g and 3 min at 2,300 × g to yield a PNIPAAm–SWNT dispersion in the supernatant (see Note 2). 3.1.2. Spectroscopic Characterization of PNIPAAm–SWNT Dispersion

The absorption spectrum (Fig. 2) showed resolved peaks in spectral ranges for the first interband transitions for metallic SWNTs (M11), the first (S11) and second (S22) interband transitions for semiconducting SWNTs (31, 32). The Raman spectrum (Fig. 3) showed features of SWNTs including the radial breathing mode (RBM), the tangential G-band, the disorder induced D-band,

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Fig. 3. Raman spectrum of PNIPAAm–SWNT dispersion

and its second-order harmonic, the G¢-band (33). The van Hove peaks in absorption spectrum and the low intensity of D-band in Raman spectrum indicate that PNIPAAm molecules are noncovalently physisorbed on the sidewall of SWNTs (26). 3.1.3. Temperature Response of PNIPAAm– SWNT Complexes

To test the temperature response of the SWNT dispersion in PNIPAAm, we use atomic force microscopy to characterize the morphology of individual PNIPAAm–SWNT complexes and fluorescence spectra to characterize the response at the ensemble level. The PNIPAAm–SWNT complexes were first measured at room temperature, then heated in a 40°C water bath for 2 min and measured after cooling to room temperature, finally re-dispersed by 2  min sonication at 0°C and measured at room temperature. Photographs (Fig.  4) showed the solution becomes turbid after heating due to the aggregation of free PNIPAAm. When the solution is cooled down to room temperature, it became clear again. 1. AFM sample preparation: 10 µl of 0.1% w/v PLL solution was spin-coated onto a freshly cleaved mica and then 10 µl of the PNIPAAm–SWNT dispersion was spin coated on this substrate. The samples were then rinsed with deionized water and dried with argon gas. All AFM images were taken at room temperature and ambient conditions in AC mode. The images show that SWNTs aggregate into larger bundles (4–20  nm diameter) after heating (Fig. 5) (see Note 3). However, after 2  min sonication in 0°C ice-water bath, SWNTs are re-dispersed individually or in small bundles similar to those from the original dispersion ( MWNTs. CNTs were found to inhibit the formation of mineralized nodules greatly and dose-dependently during the final stage of osteoblast differentiation, causing a 50% decrease in the formation of mineralized nodules at the concentration of 50  mg/mL. The expression of important proteins such as Runx-2 and Col-I in osteoblasts was also greatly inhibited by the CNTs. TEM results revealed that the effects on cellular behavior may be exerted by the CNTs from in- and outside of the cells. Key words: Carbon nanotubes, Primary osteoblast, Alkaline phosphatase, Adipocyte, TEM

1. Introduction Carbon nanotubes (CNTs) have potential applications in biosensors, tissue engineering, and biomedical devices, because of their unique electronic, chemical, and mechanical properties (1–4). Recent studies also suggested that carbon-based nanomaterials may be present in the atmospheric environment (5). Experimental evidence over the past few years showed that ultrafine nanomaterials with mean diameters