Chemical Information Call-In: Carbon Nanotube Use at ... University respectfully submits the...

STANFORD UNIVERSITY

ENVIRONMENTAL HEALTH & SAFETY

Lawrence '!d. qi66s, CIJf}lssociate'Vice Provost

December 18, 2009

Jeff Wong, PhDChief Scientist, Department of Toxic Substances Control1001 "I" StreetP.O. Box 806Sacramento, California 95812-9806

DIRECTOR'S OFFICEDEPT OF TOXIC SUBSTANCES CONTROL

DEC 23 2009

RECEIVED

Re: Chemical Information Call-In: Carbon Nanotube Use At Stanford University

Dear Dr. Wong,

Stanford University respectfully submits the following information regarding carbon nanotubeuse in research at Stanford University in response to the January 22, 2009 information call-inrequest to Stanford from the Department of Toxic Substances ControL

Stanford University is an institution ofhigher education and academic research. Carbonnanotubes (CNTs) are used regularly on a laboratory scale l in about 16 different laboratoriesin pursuit of various types of basic research on the Stanford University campus. The natureof academic research is dynamic, with researchers frequently changing the direction ofresearch investigation as evidence and data are gathered. Thus, the number oflaboratoriesand faculty investigators that may be involved with research involving CNTs may changeperiodically. Types of research with CNTs are varied and include: medical applicationsinvolving microscopy imaging; electronic devices; energy storage devices; fuel production;and fundamental physics and materials science research. Below is a sampling ofjust some ofthe many publication abstracts from various research activities which illustrate how carbonnanotubes are typically used and applied in academic research at Stanford University.

Circulation and long-term fate of functionalized, biocompatible single-walled carbon nanotnbes inmice probed by Raman spectroscopyZhuang Liu*, Corrine Davist, Weibo Cait, Lina Hef, Xiaoyuan Chen], and Hongjie Dai*§

"Department of Chemistry, Stanford University, Stanford, CA 94305; tDepartment of Comparative Medicine, Stanford UniversitySchool ofMedicine, Stanford, CA 94305; and tThe Molecular Imaging Program at Stanford (MIPS), Department of Radiology and Bio-X Program,Stanford University School of Medicine, Stanford, CA 94305

PNAS Edited by Charles M. Lieber, Harvard University, Cambridge, MA, and approved December 17, 2007 (received for reviewAugust 14,2007)

ABSTRACTCarbon nanotubes are promising new materials for molecular delivery in biological systems. Thelong-term fate ofnanotubes intravenously injected into animals in vivo is currently unknown, an issuecritical to potential clinical applications of these materials. Here, using the intrinsic Raman

1 Per Cal/OSHA 8 CCR 5191, Occupational Exposures to Hazardous Chemicals in Laboratories, laboratory scale refers to thework with substances in which the containers used for reactions, transfers, and other handling of substances are designed to beeasily and safely manipulated by one person. "Laboratory scale" excludes workplaces whose function is to produce commercialquantities of materials.

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spectroscopic signatures of single-walled carbon nanotubes (SWNTs), we measured the bloodcirculation of intravenously injected SWNTs and detect SWNTs in various organs and tissues ofmiceex vivo over a period ofthree months. Functionalization of SWNTs by branched polyethyleneglycol(PEG) chains was developed, enabling thus far the longest SWNT blood circulation up to I day,relatively low uptake in the reticuloendothelial system (RES), and near-complete clearance from themain organs in ~2 months. Raman spectroscopy detected SWNT in the intestine, feces, kidney, andbladder of mice, suggesting excretion and clearance of SWNTs from mice via the biliary and renalpathways. No toxic side effect of SWNTs to mice was observed in necropsy, histology, and bloodchemistry measurements. These findings pave the way to future biomedical applications of carbonnanotubes.

Carbon Nanotubes in Biology and Medicine:In vitro and in vivo Detection, Imaging and Drug DeliveryZhuang Liu, Scott Tabakman, Kevin Welsher, and Hongjie Dai

Department ofChemistry, Stanford University, CA 94305, USA

Received: 23 October 20081 Revised: 1 December 20081 Accepted: 2 December 200&©Tsinghua University Press and Springer-Verlag2009. This article is published with open access at Springerlink.com

ABSTRACTCarbon nanotubes exhibit many unique intrinsic physical and chemical properties andhave been intensively explored for biological and biomedical applications in the past fewyears. In this comprehensive review) we summarize the main results from our and othergroups in this field and clarify that surface functionalization is critical to the behavior ofcarbon nanotubes in biological systems. Ultrasensitive detection of biological specieswith carbon nanotubes can be realized after surface passivation to inhibit the non-specificbinding of biomolecules on the hydrophobic nanotube surface. Electrical nanosensorsbased on nanotubes provide a label-free approach to biological detection. Surfaceenhanced Raman spectroscopy of carbon nanotubes opens up a method of proteinmicroarray with detection sensitivity down to I frnollL. In vitro and in vivo toxicitystudies reveal that highly water soluble and serum stable nanotubes are biocompatible,nontoxic, and potentially useful for biomedical applications. In vivo biodistributions varywith the functionalization and possibly also size of nanotubes, with a tendency toaccumulate in the reticuloendothelial system (RES), including the liver and spleen, afterintravenous administration. If well functionalized, nanotubes may be excreted mainlythrough the biliary pathway in feces. Carbon nanotube-based drug delivery has shownpromise in various In vitro and in vivo experiments including delivery of small interferingRNA (siRNA), paclitaxel and doxorubicin. Moreover, single-walled carbon nanotubeswith various interesting intrinsic optical properties have been used as novelphotoluminescence, Raman, and photoacoustic contrast agents for imaging of cells andanimals. Further multidisciplinary explorations in this field may bring new opportunitiesin the realm of biomedicine.

Highly conductive paper for energy-storage devicesLiangbing Hu, Jang Wook Chci, Yuan Yang, Sangmoo Jeong, Fabio La Mantia, Li-Feng Cui, and Yi Cui

Departments of a)Materials Science and Engineering and b)Electrical Engineering, Stanford University, Stanford, CA 94305

Edited by Charles M. Lieber, Harvard University, Cambridge, MA, and approved October 21, 2009 (received for review August 6,2009)

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ABSTRACTPaper, invented more than 2,000 years ago and widely used today in our everyday lives,is explored in this study as a platform for energy-storage devices by integration with ID

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nanomaterials. Here, we show that commercially available paper can be made highlyconductive with a sheet resistance as low as I ohm per square Usq) by using simplesolution processes to achieve conformal coating of single-walled carbon nanotube (CNT)and silver nanowire films. Compared with plastics, paper substrates can dramaticallyimprove film adhesion, greatly simplify the coating process, and significantly lower thecost. Supercapacitors based on CNT-conductive paper show excellent performance.When only CNT mass is considered, a specific capacitance of200 F/g, a specific energyof 30-47 Watt-hour/kilogram (WbJkg), a specific power of 200,000 W/kg, and a stablecycling life over 40,000 cycles areachieved. These values are much better than those ofdevices on other flat substrates, such as plastics. Even in a case in which the weight of allof the dead components is considered, a specific energy of 7.5 Wh/kg is achieved. Inaddition, this conductive paper can be used as an excellent lightweight current collectorin lithiurnion batteries to replace the existing metallic counterparts. This work suggeststhat our conductive paper can be a highly scalable and low-cost solution for highperformance energy storage devices.

Noninvasive molecular imaging of small living subjects using Raman spectroscopyS. Keren, C. Zavaleta, Z. Cheng, A. de la Zerda, O. Gheysens, and S. S. Gambhir*

MolecularImaging Program at Stanford, Departmentsa/Radiology and Bioengineering, Bio-X Program, Stanford University, /201WelchRoad, Stanford, CA 94305-5484

Edited by Michael E. Phelps, University of California, Los Angeles School of Medicine, Los Angeles, CA, and approved February 5,2008 (received for review November 7, 2007)

ABSTRACTMolecular imaging of living subjects continues to rapidly evolve with bioluminescenceand fluorescence strategies, in particular being frequently used for small-animal models.This article presents noninvasive deep-tissue molecular images in a living subject withthe use of Raman spectroscopy. We describe a strategy for small-animal optical imagingbased on Raman spectroscopy and Raman nanoparticles. Surface-enhanced Ramanscattering nanoparticles and single-wall carbon nanotubes were used to demonstratewhole-body Raman imaging, nanoparticle pharmacokinetics, multiplexing, and in vivotumor targeting, using an imaging system adapted for small-animal Raman imaging. Theimaging modality reported here holds significant potential as a strategy for biomedicalimaging of living subjects.

Self-Sorted Nanotube Networks on Polymer Dielectrics for Low-Voltage Thin-Film TransistorsMark E. Roberts, Melbume C. LeMieux, Anatoliy N. Sokolov, and Zhenan Bao*

Department ofChemical Engineering, Stanford UniVersity, Stauffer Ill,381 North-South Mall, Stanford, California 94305-5025

Nano Letters 2009 Vol. 9, No.7 2525-2531. ReceivedJanuary 27,2009; Revised Manuscript ReceivedMay 8,2009

ABSTRACTRecent exploitations of the superior mechanical and electronic properties of carbonnanotubes (CNTs) have led to exciting opportunities in low-cost, high performance,carbon-based electronics. In this report, low-voltage thin-film transistors with aligned,semiconducting CNT networks are fabricated on a chemically modified polymer gatedielectric using both rigid and flexible substrates. The multifunctional polymer serves asa thin, flexible gate dielectric film, affords low operating voltages, and provides aplatform for chemical functionalizatiou. The introduction of amine functionality to thedielectric surface leads to the adsorption of a network enriched with semiconducting

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CNTs with tunable density from spin coating a bulk solution of unsorted CNTs. Thecomposition of the deposited CNT networks is verified with Raman spectroscopy andelectrical characterization. For transistors at operating biases below 1 V, we observe aneffective device mobility as high as 13.4 cm2lVs, a subthreshold swing as low as 130mY/dec, and typical on-off ratios of greater than 1,000. This demonstration of highperformance CNT thin-film transistors operating at voltages below 1 V and depositedusing solution methods on polymeric and flexible substrates is an lmportant step towardthe realization of low-cost flexible electronics.

Researchers have obtained carbon nanotubes used in such research from either outsidesources or synthesized their own CNTs:(a) Most of the CNTs used at Stanford are acquired from outside vendors (i.e., Hanwha

Nanotech Co., NanoTech Labs, Inc., Unidym).(b) In a very limited number of instances, carbon nanotubes have been synthesized on site at

Stanford via chemical vapor deposition on a laboratory scale, as defmed above, forspecific research purposes. Hence, Stanford University does not engage in "productionor manufacturing of carbon nanotubes for commerce."

2. Sampling, Detection and Measurement Methods Used to Monitor in the Workplace andEnvironmentBased on industrial hygiene workplace exposure evaluation of selected researchersperforming laboratory-scale procedures with carbon nanotubes, it has been determined thatthere is minimal risk ofpersonal exposures and releases to the environment. This is due tothe extremely small laboratory scale quantities of carbon nanotubes used, short durations ofmanipulation, engineering controls employed, work practices followed, and use ofappropriate Personal Protective Equipment, when appropriate. Also, when adhered to asubstrate, such as during deposition and growth, or when used in solution, the potential foraerosolization is minimal. At this time, quantitative sampling and detection have beendeemed not necessary for the risk management program. Stanford researchers do follow arisk based set of control guidelines, which are discussed further in Section 5 below.

Given the fact that the use ofmeaningful quantitative exposure sampling and analyticaltechniques are still under development, Stanford University is exploring a project to haveNIOSH's Nanotechnology field group our visit campus to conduct possible exposureevaluation and monitoring in 2010. We will be happy to provide any findings of this studywith the DTSC, should the project move forward.

The projected presence of carbon nanotubes in the environment could result from incidentalreleases during normal laboratory scale handling and storage, accidental spills, and fromhazardous waste management.

o Routine Laboratory Handling and Storage: Based on a campus-wide survey ofresearch programs, the overall projected use from carbon nanotube research at Stanford



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University is approximately 16 grams per year among the sixteen Principal Investigators.Researchers working with carbon nanotubes endeavor to minimize any product loss dueto its significant expense and research value. Rather, the bulk of carbon nanotubes are insolution, on a substrate, or in a research animal, etc., and ultimately end up as hazardouswaste. Nevertheless, negligible quantities of carbon nanotubes could be released into theenviromnent via laboratory exhaust systems (e.g., fume hoods, glove boxes, ventedfurnaces, general laboratory exhaust) during normal handling and storage. This isnegligible when compared to the amount of existing atmospheric ultrafme particulateswhen one considers the significant amount of nanoparticles that are spewed daily fromdiesel exhaust and other combustion emission sources throughout the state.

o Accidental Spills: Although accidental spills during laboratory procedures are possible,quantities would be relatively small, ranging from 100 micrograms (approximate quantityof raw carbon nanotubes on a spatula) to I gram (if an entire container were spilled).Spill procedures are in place in the event of need to address small or larger spills.

o Nanoparticle Waste: By DTSC regulatory defmition "laboratory waste" is by defaultconsidered regulated hazardous waste unless proven not to meet the waste characteristics.Although we postulate that carbon nanotubes in the amounts used would not meet the testparameters for regulated hazardous waste, Stanford has chosen to manage such materialsas laboratory hazardous waste. Nanoparticle wastes are collected along with otherhazardous wastes from laboratories and sent to the University's hazardous waste vendorsin tightly sealed containers for appropriate hazardous materials management viaincineration at the permitted waste treatment facility.

4. Knowledge of ()c¢qpll.ti()lllli $llfetY.l>ub]jc lItlllltl:lalld.ith.tlEnyiroIlDlclltStanford University monitors the continuing development of new information generated bythe science community to assess the potential health and safety risks associated withengineered nanomaterials. Leading groups in this area of research include work coordinatedby The National Institutes of Health ~NIOSH (http://www.cdc.gov/nioshltopics/nanotechL)and The International Council on Nanotechnology- ICON (http://icon.rice.edulindex.cfm).These organizations collect in one location most of the peer reviewed publications regardingnanomaterial health and safety and these sources, which are regularly monitored for updatedinformation, reflect the current understanding of potential occupational safety, public health,and enviromnental impacts from carbon nanotubes.

Until more is known about the possible risks of nanomaterials, Stanford researchers take aprecautionary, but reasonable approach, and utilize good laboratory safety practices asdescribed in Section 5 below when working with these materials.

Also, it is not clear what DTSC considers to be covered by the CNT call-in. For example,the research animal work done in the Dai laboratory at Stanford (see paper by Liu et al inattachment I) where functionalization of CNTs for use in biological systems appears toindicate such functionalized CNTs are cleared from the body without adverse effect. Thequestion is whether functionalized CNTs are considered to be the same as non-functionalized


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CNTs for purpose of DTSCs information gathering or possible subsequent regulation ofCNTs. It will be very important for DTSC to clearly identify the specific particles that will becovered by any guidance or regulation, as minor changes or addends to the surface of a CNTmay result in significantly different properties.

Stanford University's Chemical Hygiene Plan (CHP), which is required per California'sOccupational Safety and Health Administration (Cal/OSHA) Occupational Exposure toHazardous Chemicals in Laboratories Title 8- California code of Regulations, Section 5191,provides for and supports the procedures, equipment, personal protective equipment, andwork practices for protecting laboratory personnel from potential health hazards of usinghazardous chemicals in the laboratory. The CHP serves as the cornerstone oflaboratoryhealth and safety at Stanford University and is available at:http://chemtoolkit.stanford.edu/docs/Chemical Hygiene Plan.pdf. Stanford University'sLaboratory Chemical Safety Toolkit is a companion to the CHP and is available at:http://ChemToolkit.stanford.edu.

To support the health and safety effortsofresearchers working with nanomaterials a StanfordEH&S guidance document entitled, General Principles and Practices for Working Safelywith Engineered Nanomaterials, has been distributed to faculty identified as working withnanomaterials. The document is also posted on Stanford University's website; seeAttachment 2 herein. The Guide is specific for nanomaterials work in the laboratory and isintended to supplement the requirements of Stanford University's CHP and its companionLaboratory Chemical Safety Toolkit. The Guide provides additional direction on engineeringcontrols, work practices, and personal protective equipment to employ in order to minimizeoccupational exposures to nanomaterials, as well as guidance on proper storage and wastemanagement techniques to avoid releases to the environment. A Standard OperatingProcedure template for working with nanomaterials has also been been made available toresearchers to assist in determining the level and types of controls that may be appropriate forthe projected research activity; see Attachment 3.

6; Waste Handling Practices and Management for Carbon NanotubesNanoparticle wastes, including contaminated lab debris, are managed as hazardous wasteconsistent with federal, state and local requirements. Researchers collect and store wastematerials in closed containers, which include waste labeling (e.g., contains nanosilver waste).

Comment on Future Rule MakingIf rule making regarding the use, fate, and transport of CNTs is pursued in the future, StanfordUniversity would like to continue to partner with the DTSC to provide input, representinginstitutes of higher education. We wish to ensure that any future rule-making considers theunique aspects oflaboratory scale activities utilized in the course of education and academicresearch at universities and colleges throughout California.


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Kindly direct any inquiries regarding this response to Lawrence M. Gibbs, Associate ViceProvost for Environmental Health and Safety, [email protected], (650) 723-7403.

s~'ncerel,

r QlI9d£,L ence M. Gibbs, emAssociate Vice Provost, Environmental Health & Safety

Cc: John Hennessy, President, Stanford University

Attachments:

Attachment I: Sampling of Stanford Faculty Research Articles involving Carbon nanotubes:

• Stanford University Professor Hongjiei DaiZhuang Liu", Corrine Davis'], Weibo Caij, Lina Hef, Xiaoyuan Chenj, and Hongjie Dai'§"Circulation and long-term fate offunctionalized, biocompatible single-walled carbonnanotubes in mice probed by Raman spectroscopy" Proc Natl Acad Sci USA, Dec 2007

• Stanford University Professor Hongjiei DaiLiu, Z; Tabakman, S; Welsher, K; Dai, H. "Carbon Nanotubes in Biology and Medicine: Invitro and in vivo Detection, Imaging and Drug Delivery," Nano Res. 2(85), 120,2009.

• Stanford University Professor Yi CuiHu, L; chor, J; Yang, Y; Jeong, S; La Mantia, F; Cui LF; Cui, Y. (2009) "Highlyconductive paper for energy-storage devices," Proc Natl AcadSci USA, Early Ed.

• Stanford University Professor Sanjiv GambhirKeren, S; Zavaleta, C.; Cbeng, Z.; de la Zerda, A; Gheysens, 0; Gambhir, S. (2008)"Noninvasive molecular imaging of small living subjects using Raman spectroscopy," ProcNat! Acad Sci USA, 105: 5844-5849.

• Stanford University Professor Zhenan BaoRoberts, M; LeMieux, M; Sokolov, A; Bao, Z. "Self-Sorted Nanotube Networks on PolymerDielectrics for Low-Voltage Thin-Film Transistors," Nano Lett. 2009, (3) 2526-2531.

Attachment 2: Stanford University - General Principals and Practices for Working Safely withEngineered Nanomaterials

Attachment 3: Stanford University - Standard Operating Procedure (SOP) Template forWorking with Nanomaterials.

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Attachment la

Edited by Charles M. Ueber, Harvard University. Cambridge, MA, and approved December 17, 2007 (received for review August 14, 2007)

Zhuang Liu*, Corrine Davis", Weibo Cai*, Lina He*, Xiaoyuan Chen*, and Hongjie Dai*§

*Department of Chemistry, Stanford University, Stanford, CA 94305; "Department of Comparative Medicine, Stanford University School of Medicine,Stanford, CA94305; and t-TheMolecular Imaging Program at Stanford (MIPS), Department of Radiology and Blo-XProgram, Stanford UniversitySchool of Medicine, Stanford, CA94305

Author contributions: Z.L., X.C, and H.D.designed research; Z.L., CD., W.C, L.H., and X.Cperformed research; CD. contributed new reagents/analytic tools; Z.L. and CD. analyzeddata; and Z.L. and H.D.wrote the paper.

The authors declare no conflict of interest.

This art1cie is a PNAS Direct Submission.

§Towhom correspondence should be addressed. E-mail:[email protected].

This article contains supporting information online at www.pnas.org/cgilcontentffull/0707654105/0(1.

@2008 by The National Academy of Sciencesof the USA

into animals and are not rapidly excreted (6, 20). Whether or notand how these materials are cleared from the body is unknownin many cases, because of the difficulties in long term in vivotracking and monitoring of the materials. Currently used radiolabels (6, 20) or fluorescent labels for nanomaterials are usefulfor in vivo tracking over short periods of time (a few hours to afew days), but these labels may gradually dissociate from thematerials or decay and lose activity over time. It is thus highlydesirable to detect nanomaterials based on their intrinsic physical properties rather than relying on radiolabels or spectroscopictags for indirect detection/measurement. The direct detectionmethod may lead to a more accurate assessment of how nanomaterials behave in vivo in both short and long terms, i.e., duringthe blood circulation stage and during time periods lastingseveral months.

Here, we show that the intrinsic Raman scattering intensity ofSWNTs (23, 24) does not decay over time while being relativelyinsensitive to the types of noncovalent coatings and solutionenvironments of SWNTs. Unlike the resonance breathing model(RBM) in the SWNT Raman spectrum, the intensity of tangential graphene-like G band is relatively insensitive to the diameterand bundling of nanotubes (25, 26). We then use Ramanspectroscopy to measure the postinjection blood concentrationof SWNTs with different polyethylene-glycol (PEG) coatings inmice and thus glean nanotube blood circulation times. We alsouse Raman spectroscopy and Raman imaging to probe thebiodistribution of SWNTs in various organs of mice ex vivo overa period of several months. We find that the surface chemistry,including the length and branching structure of PEG chains, iscritical to the in vivo behaviors of SWNTs. Longer and morebranched PEG on SWNTs afford longer blood circulation andlower RES uptake. Superior to linear PEG, branched PEGcoating renders SWNTs a significantly prolonged blood circulation, which is desired for future targeted imaging or therapeutic applications. By monitoring the biodistribution of SWNTs inmice organs for months, a relatively slow,but persistent decreasein the SWNT Raman signal is observed, suggesting that theclearance of SWNTs is occurring in the mice. To study theexcretion pathway, nanotubes with the highest possible dose isinjected into mice with their excreta examined by Ramanspectroscopy. Biliary excretion pathway is confirmed by theSWNT Raman signals in the intestine and feces. Althoughthe SWNT concentration in urine is low over a high background,

biodistribution I blood circulation I nanoparticles I excretion I toxicity

Carbon nanotubes are promising new materials for moleculardelivery in biological systems. The long-term fate of nanotubesintravenously injected into animals in vivo is currently unknown.an issue critical to potential clinical applications of these materials.Here. using the intrinsic Raman spectroscopic signatures of singlewalled carbon nanotubes (SWNTs). we measured the blood circulation of intravenously injected SWNTs and detect SWNTs invarious organs and tissues of mice ex vivo over a period of threemonths. Functionalization of SWNTs by branched polyethyleneglycol (PEG) chains was developed. enabling thus far the longestSWNT blood circulation up to 1 day. relatively low uptake in thereticuloendothelial system (RES). and near-complete clearancefrom the main organs in =2 months. Raman spectroscopy detectedSWNT in the intestine. feces, kidney. and bladder of mice. suggesting excretion and clearance of SWNTs from mice via the biliary andrenal pathways. No toxic side effect of SWNTs to mice wasobserved in necropsy. histology. and blood chemistry measurements. These findings pave the way to future biomedical applications of carbon nanotubes.

Circulation and long-term fate of functionalized,biocompatible single-walled carbon nanotubesin mice probed by Raman spectroscopy

The utilization of novel nanomaterials for biological andbiomedical applications has been an active and exciting

direction of research in recent years (1-3). A wide range ofnanomaterials, such as nanopartic1es (4-7), nanorods (8),nanowires (9), and nanotubes (10-12) have been investigated fortheir potential clinical applications in diagnosis and therapeutictreatment of diseases. The interesting structural, chemical, electrical, and optical properties of carbon nanotubes (13, 14), whenused in biological and medical settings, may bring new opportunities to biological detection, imaging, and therapy with highperformance and efficacy. Carbon nanotube-based intercellularmolecular delivery vehicles have been developed for intracellulargene (15-17) and drug delivery in vitro (18, 19). Recently,research began to investigate the behavior of carbon nanotubesin animal bodies in vivo (20, 21), including the finding of lack oftoxicity of well PEGylated single-walled carbon nanotubes(SWNTs) in mice in a pilot study(M. L. Schipper, N. NakayamaRatchford, C.D., N. W. S. Kam, P. Chu, Z.L., X. Sun, L. C. Cork,RD., and S. S. Gambhir, unpublished data).

The biodistribution of intravenously injected carbon nanotubes in mice has been studied by using radiolabeling and isotoperatio mass spectroscopy methods (20, 22). Promising result ofefficient targeted tumor accumulation in vivo has been obtainedby conjugating a ligand peptide to nanotubes to recognizereceptors on the surface of tumor cells (20), suggesting highpotential of nanotube-based drug delivery vehicles for cancertherapy. However, an important unaddressed question for carbon nanotubes and various nanomaterials in general in biomedical applications is their long term fate in vivo. It is known thatmost nanomaterials tend to exhibit high uptake in the reticuloendothelial system (RES) (liver, spleen, etc.) once injected

1410-1415 I PNAS I February 5, 2008 I vel. 105 I no.5 www.pnas.orgjcgijdoij10.1073jpnas.0707654105

concentration in blood or tissue lysates of mice using Ramanspectroscopy. Solutions of same concentration of SWNTs withdifferent PEG coatings (SWNT-I-2kPEG and SWNT-I-5kPEG,SWNT-br-7kPEG) exhibited very similar Raman intensities invarious environments including water, saline, lysis buffer, serum,and liver lysate. These results suggested that the Raman intensityof SWNTs was relatively insensitive to the coatings and solutionenvironments involved in our experiments (SI Fig. 6).

We intravenously injected =200 ,ul of saline solutions ofdifferent PEG functionalized SWNTs at the same nanotubeconcentration (=0.1 mg/ml) into mice and drew blood (=5 ,tI)at different time points postinjection (p.i.) for Raman measurement (Fig. 2 a-e). The measured %lD/g (percentage of injecteddose per gram) in blood vs. time p.i, gave blood circulationbehavior of SWNTs with varions PEGylations (Fig. 2 d and e).We observed that increasing the linear PEG chain length from2 kDa (SWNT-1-2kPEG) to 5 kDa (SWNT-I·5kPEG) significantly extended the blood circulation of SWNTs from =1.2 h to=5 h (Fig. 2 a, b, d, and e). Note that we defined the bloodcirculation time as the time span over which the blood SWNTlevel reduced to 5%ID/g. This result was consistent with ourprevious measurements made with radio-labeled SWNTs (20).However, further increase of linear PEG length to 7 kDa(SWNT-I·7kPEG) and even 12kDa (SWNT-l·12kPEG) showedonly minor effect on the blood circulation time (Fig. 2 d and e).However, SWNT-b,-7kPEG, i.e., SWNTs funetionalized withthree branched PEG chains (Fig. 10), exhibited a remarkableincrease in circulation to =15 h (Fig. 2 C, d, and e, with SWNTsdetected in the blood nearly 1 day p.i.), This finding is importantand suggests that branched PEG structures on SWNTs is highlydesired in affording optimal biological inertness of SWNTs thatresist opsonization or nonspecific binding of proteins in vivo,avoiding rapid RES uptake and thus prolonging circulation inblood. We attribute this improvement to the branched PEGstructure giving better coverage and higher density of hydrophilic PEG groups on SWNTs, thus making nanotubes moreinert and resistant to nonspecific binding and uptake.

To investigate the biodistribution of nanotubes in the mainorgans 1 day p.i. of SWNTs, we killed mice injected withSWNT-1-2kPEG, SWNT-I-5kPEG, and SWNT-b,-7kPEG, respectively. The organs and tissues were homogenized and solubilized in lysis buffers, for measuring SWNT levels in theseorgans and tissues by Raman spectroscopy (Fig. 30 and SI Fig.7). We observed dominant SWNT uptake in the liver and spleenof the RES over other organs and tissues. Clearlyreduced levelsof liver and spleen uptake were seen for SWNT-I-5kPEG andSWNT-br-7kPEG compared with SWNT-I-2kPEG (Fig. 30),suggesting higher degree of surface PEGlyation of SWNTsaffording lower RES uptake. Under the injected dose of 200 J.tIof SWNT at =0.1 mg/ml concentration and detection conditions,no obvious SWNT signals were detected in other main organsexcept for minor kidney signal. Note that the detection limit ofSWNTs was =0.04 p.g/ml in blood and =0.2 p.g/ml in othertissues, corresponding to =O.2%ID/g and =1%ID/g of thenormal injected dose respectively. Therefore, under the injected=200 p.l of =0.1 mg/ml nanotubes, the lack of appreciableSWNT Raman signals in organs other than liver, kidney, andspleen (Fig. 30) does not mean absolutely no SWNT uptake inthose organs. It suggests only that the level of uptake is lowerthan a certain limit (1-2%ID/g). The total amount of SWNTs inthese organs could still be substantial owing to the large mass/weight of the tissues combined.

Indeed, by injecting a highly concentrated SWNT-1-5kPEGsolution (200 p.1 of =05 mg/ml SWNTs in a form of concentratedviscous liquid) into mice, we observed at 1 day p.i, that, besidesdominant liver and spleen uptake, SWNT Raman signalsbecameapparent in the bone (leg bone), kidney, intestine, stomach, andlung of mice with =1%lD/g in the first three organs (Fig. 40).

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~ 20000a:

Fig. 1. NoncovalentlyfunctionalizedSWNTs byvarious PEGylated phospholipids. (a Left) Scheme offunctionalization byvarious phospholipid-PEGs withlinear or branchedPEG chains. (Right) A photo of the 5WNT-br-7kPEG salinesolution atthe concentration (0.1 mg/ml;optical density was 4.6 at808 nm for1 em path) used for injection. (b) A Ramanspectrum of a solution of 5WNT1-2kPEG. The G band peak at 1,590em"! was usedfor 5WNT detection in thiswork. (c) Raman intensity vs. $WNT concentration calibration curve. Lineardependencewasobserved from 0.02 ,u.g/ml to 4,u.g/m1.

the existence of SWNT Raman signals in the kidney and bladderis evident, suggesting renal excretion for nanotubes as wel1.Further, necropsy, histology and blood chemistry studies revealno obvious toxic effect for mice injected with SWNTs, providingfurther supports to the recent finding (M. L. Schipper, N.Nakayama-Ratchford, CD., N. W. S. Kam, P. Chu, Z.L., X. Sun,L. C. Cork, RD., and S. S. Gambhir, unpublished data). Theseresults establish a foundation for further exploration of carbonnanotubes for biomedical applications and may have implications for other nanomaterials.

Results and DiscussionWe used Hipco SWNTs noncovalently functionalized and solubilized (see Materials and Methods) by PEGylated phospholipids(Fig. la) that were stable without aggregation in various biological solutions including serum (15, 17, 20). Our previous study(20) also showed that the phospholipid-PEG coating was stablein vivo without rapid detachment. Centrifugation was used toremove big bundles and impurities, leaving short individual andsmall bundles of tubes in the solution. Atomic force microscopy(AFM) images revealed similar length distributions of differentfunctionalized SWNTs [SWNT-1-2kPEG, 104 ± 49 run; SWNT1-5kPEG, 101 ± 51 run; SWNT-b,-7kPEG, 95 ± 46 nm; supporting information (51) Materials and Methods, and SI Fig. 5].Strong resonance Raman scattering is an intrinsic optical property of SWNTs with sharp peaks and low background in thespectra (Fig. Ib). In this work, the tangential graphite-likephonon mode (G band), the strongest peak in the SWNT Ramanspectrum, was used to detect nanotubes in solution, blood, andtissue lysates. No obvious decay in the Raman signal wasobserved by measuring the Raman spectrum of a SWNT solutionfor np to 3 months (Sl Fig. 6). Raman spectra of SWNT solntionswith known concentrations from 0.02 to 4 ,ug/mlwere taken, andthe G band intensities (integrated peak areas) were plottedagainst SWNT concentrations [measured by their near infrared(NIR) absorptions] as the calibration curve (Fig. Ic). The lineardependence allowed for quantitative measurement of SWNT

Liueral. PNAS I February 5, 2008 I vol. 105 1 no.5 I 1411

25

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clearance of SWNTs from mice organs, with branched PEGfunctionalization giving the most desirable in vivo behavior ofSWNTs. We also carried out Raman spectroscopic mapping andimaging of SWNTs in the liver slices of mice at the 3-month timepoint. Many "hot spots" (locations over the liver slice showinghigh SWNT Raman signals) were observed in the SWNT-l2kPEG treated sample and few in the SWNT-I-5kPEG-treatedsample (Fig. 3d). This observation was consistent with muchlower level of SWNT retention in the liver in the latter case.

Because no significant SWNT signals were detected in themain organs of mice other than in the RES over the severalmonth period, we suggest that the clearance of SWNTs from theRES was due to the excretion of SWNTs out of the body, ratherthan nanotubes migrating to other organs of mice. The excretionoccurred faster for SWNTs with higher degree of PEGylation,hydrophilicity, and biological inertness. To find out the detailedexcretion pathway, we collected the urine and feces of mice atdifferent time points after injection of high doses of SWNTs andcarried out Raman measurement. We observed appreciableSWNT Raman signal in the dry feces sample of mice aftersubtracting a high background (Fig. 4c). The obvious presence ofnanotubes in the feces and intestine (Fig. 4b) clearly revealsSWNT excretion via the biliary pathway in the feces. Althoughno significant SWNT Raman signal was observed in the urine

1-7kPEG

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Fig. 2. Blood circulation behavior of SWNTs probed by Raman spectroscopy. (e-c) Raman spectra of blood samples drawn from BALBk mice at various timepoints after injection with SWNT-I-2kpEG (a) SWNT-'-5kPEG (b), and SWNT·br·7kPEG (c) solutions, respectively. Note that spectrum baselines were subtractedin a-Co (el)Blood circulation data probed by the Raman method for 5WNT-I-2kPEG. 5WNT-I-5kPEG,SWNT-I-7kPEG, SWNT-I-12kPEG, arid SWNT-br-7kPEG.lnjectedinto BALBk mice (Inset). The SWNTlevels in blood were determined aspercentage of injected SWNTamount per gram of blood (%ID/g in blood). SWNT-I-5kPEG,5WNT-I-7kPEG, and SWNT-I-12kpEGshowed similar blood circulation time, significantly longer than that of SWNT-'-2kPEG.The longest blood circulation wasobserved for SWNT-br-7kPEG.The error bars are based on four mice in each group. (e) Blood circulation time of 5WNTswith different PEGylations. The bloodcirculation time was defined as the time duration through which the blood 5WNT level reduced to 5%ID/g.

Because bone marrow is part of the RES, uptake of SWNTs inthe bone is understandable and consistent with the previousbiodistribution study for other nanomaterials (27). The existenceof SWNTs in the kidney and intestine of mice is interesting,suggesting possible urinary and biliary excretion routes forSWNTs, as discussed in the following paragraphs.

To glean the long term fate of SWNTs in vivo, injected micewere killed at 1, 2, and 3 months p.i, for biodistribution measurements with three to four animals per group at each timepoint. We found that the concentration of SWNTs remained verylow in most of the organs except for the liver and spleen. In thesetwo organs, we did observe SWNT levels steadily decreasing overa 3-month period, with the concentration of retained SWNTsfollowing the trend of SWNT-I-2kPEG > SWNT-I-5kPEG >SWNT-br-7kPEG at later time points (Fig. 3 b and e). In the caseof SWNT-I-2kPEG, appreciable amounts of SWNTs remained inthe liver and spleen, with a concentration of =7%ID/g at even3 months p.i. In contrast, very low levels (=2%ID/g) of SWNT1-5kPEG were retained in the RES of mice at 3 months p.i. (Fig.3 band c). The least retention of nanotubes in the RES wasobserved for SWNT-br-7kPEG, with <2%ID/g retention at 2months p.i. These results-suggest that, in addition to the advantages of longer blood circulation and lower initial RES uptake,higher degree of PEGylation of SWNTs affords more rapid

1412 I www.pnas.orgjcgijdoi/l0.l073jpnas.0707654105 Uu etal.

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stable radiolabels on SWNTs or the intrinsic Raman, photoluminescence or isotope properties of the nanotube themselves.The slowexcretion contradicts with the report bySingh et al (36),who suggested fast urinal excretion of carbon nanotubes basedon radiolabel signals, which could be due to detachment ofradiolabels from nanotubes or free unconjugated excess radiolabels in the injected samples. Such problems are avoided indirect detection methods using the intrinsic physical or chemicalproperties of nanotubes.

Neither mortality nor significant loss of body weight wasobserved with any of the mice (>30) iujected with 0.1 mg/mlSWNTs during the 3-month period of the study. To investigateany potential toxic effect of SWNTs, necropsy, histology andblood chemistry examinations were performed on our SWNTtreated mice at 3 months p.i, No obvious toxicity of nanotubeswas noticed in this small-scale toxicity study (SI Fig. 8 and SITable 1). This observation is consistent with a recent pilottoxicity study carried out with four to five mice per groupinjected with similarlyfunctionalized nanotubes (M. L. Schipper,

40

Fig. 4. Biodistribution measured under ultra-high dose SWNT (0.5 mg/ml)injection. (a) Biodistribution of SWNT-I-5kPEG in various organs of micereceived 200 p..1 of 0.5 mg/ml SWNT-I-SkPEG at 24 h p.i. After increasing of theinjected dose, SWNTs became detectable by Raman inseveral organs includingbone, lung, kidney, stomach, and intestine, with slightly over 1%IDIgin bone,kidney and intestine. Error bars were based on three mice. (b and c) Ramanspectra of intestine lysate at 24 h p.i. (b) and dry feces sample collected 8 h poi.(c). Because of high background in the feces sample, the spectrum was takenby 600 s long time collection (5 s for all of the other samples). An appreciableamount of nanotubes was noticed in the intestine and feces, providing clearevidence forthe biliary excretion pathway of intravenously injected SWNTs, (dand e) Raman spectra of kidney (eI)and bladder (e) Iysates from mice at 24 hp.i. The existence of SWNTs in the kidney and bladder indicate that a smallportion of nanotubes is excreted via the kidney-urine pathway. Note thatbackground has been subtracted in the spectra b-e.

.!1'e 20

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Fig. 3. SWNTs in mice tissues probed by ex vivo Raman spectroscopy afterinjection into mice. (a) Biodistribution of SWNT-f-2kPEG, 5WNT-f-5kPEG, andSWNT-br-7kPEG, respectively, at 1 day p.i. measured by Raman spectroscopy.The SWNTconcentrations in most organs are below detection limit. (b and c)Evolution of the concentrations of SWNTsretained in the liver and spleen ofmice over a period of 3 months. Compared with SWNT-I-2kPEG. much lowerconcentrations of retained SWNTs in the liver and spleen were observed forSWNT-I-5kPEG and SWNT-br-7kPEG. (d) Raman mapping images of liver slicesfrom mice treated with SWNT-I-2kPEG (Left), SWNT-I-5kPEG (Center), andSWNT-br-7kEG (Right)at 3 months p.i. More SWNT signals were observed inthe SWNT-I-2kPEG-treated mouse sample than the other two samples underthe same Raman imaging conditions (laser power, beam size, etc). The errorbars in a-cwere based on three to four mice per group. Note that the injectedSWNT solutions had a concentration of 0.1 mg/ml (optical density was 4.6 at808 nm for 1 cm path).

samples because of high background and dilution by large urinevolume, we did observe clear SWNT signals in the kidney andbladder of mice at 24 h p.i. (Fig. 4 d and c). Further, for miceinjected with SWNTs at 3 days p.i., SWNT signals in the kidneywere much lower and almost nonexistent in the bladder (data notshown). This finding snggests that SWNTs were also excretedthrough the renal pathway. Considering the average lengths ofSWNTs (mean =100 run, SI Fig. 5) exceeding the renal excretionthreshold (28) and the fact that the majority of nanotubesaccumulates in the liver,we suggest that urinary excretion occursfor a small percentage of nanotnbes with very short length «50nm in length, diameter 1-2 nm) in our sample at early time pointsafter injection. Because of the existence of nanotubes in feces,the nonbiodegradable nature of Sp2 carbon SWNT structure andthe fact that foreign nondegradable substances (including various nanoparticles) in the liver are known to be excreted throughthe biliary pathway from the liver to the bile duct, intestine, andfeces (29-35), we propose that excretion of onr SWNTs frommice occurs over time mainly via the biliary pathway and endsup in feces.

Notably, our results here including the dominant RES uptakeof SWNTs and the relatively slow excretion are consistent withseveral previous reports by ns (20) and other groups (21,22) whoused detection methods based on strongly anchored serum-

Liu et al. PNAS I February 5, 2008 I vel. 105 I no. 5 I 1413

[SWNT]tissue lysate X VUssue lysate%ID/g ~ [SWNT] V .. hinjected X injected SWNT X tissue weig t

x 100%.

, [SWNT] hlood lysate X Vhlood lysate%ID/g ~ .

[SWNT]injected X VinjectedSWNT X blood weight

Fuctionalization and Characterization of PEGylated SWNTs. Raw Hipco SWNTs(0.2 mg/ml) were sonicated in a 0.2-mM solution of PL-PEG for 1 h followed bycentrifugation at 24,000 X 9 for 6 h, yielding a suspension of SWNTs withnoncovalent PL-PEG coating in the supernatant (15,20,40). Excess surfactantand unreacted PEG molecules in the caseofsynthesized PL-PEG were removedby filtration through a 100-kOaMWCOfilter(Millipore), typically 1day beforein vivo experiments. The filtration efficiency was confirmed by fluorescentlabeled PEG, showing completed removal of PEG molecules after 6 timesfiltration and washing (data not shown). Right before injection, the solutionwas centrifuged again at 24,000 x 9 for 6 h to remove any potential aggregates. UV-VIS-NIR absorption spectrum of the SWNTsolution was acquired bya Cary 60001 UV-visible-NIR spectrometer. Atomic force microscopy (AFM)images were taken by depositing SWNTs from solution onto Si02 substrates.The length of SWNTs was measured to be =100 nm averaged over 100tubesimaged by AFM. The concentration of a SWNTsolution was determined byRaman spectroscopy and by optical absorbance at 808 nm with a weightconcentration basedextinction coefficient of 46 L'g-t'cm-t or a molar extinction coefficient of 3.9 X 106 M-l'cm-t for typical -e too-nm-ronc tubes (11).

SWNT i.v. Injection and Raman Spectroscopy Method for Measuring BloodCirculation and Biodistribution in Mice. Six-week-old BALBIcmice were used inour study. Two hundred microliters of =0.1 mg/ml SWNT(optical density was4.6 at 808 nm for 1 cm path) saline solution was intravenously (l.v.) injectedinto the tail vein of each mouse. Before injection of the SWNT solution, aRamanspectrum was recorded and used to calculate the SWNTconcentrationbasedon the calibration curve described above. At various time points postinjectlon (p.f.), =5 p.1 of blood was collected from the tail vein (using a differentvein from the injected one) and dissolved in 5 p.lof lysisbuffer (1% SOS, 1%Triton X-100, 40 mM Trls acetate, 10 mM EOTA, 10 mM Orr) for detectingSWNTs in the blood by Raman measurement. The Raman G band peak areaswere measured to calculate the SWNT concentrations in the blood. Thepercent injected dose per gram (%IO/g) of blood was calculated by thefollowing equation:

x 100%.

RamanMeasurement of SWNT Solutions for Calibration Curve of SWNT RamanIntensity vs.Concentration. SWNTsolutions of various concentrations in capillary glass tubes (Fisher) were measured by using a Renishaw microRamaninstrument (laser excitation wavelength = 785 nm). A glass capillary tubefilled with a SWNTsolution was placed under the objective (X20) ofthe Ramanmicroscope. As low as2 p.lof solution sample was required for each measurement. After focusing at the center of the capillary, we recorded the Ramanspectrum of the solution (100 mW power with laserspot sizeof =25 p.m 2, 10-scollection time). At least four spectra were taken for each sample for averaging. For a given concentration of SWNT solution, the Raman intensity wasobtained by integrating the SWNTG-band peak area from 1,570cm-1to 1,620em"! and averaged over several spectra. Note that SWNTs with the sameconcentration in water, buffer, serum, and tissue lysate were found to exhibitsimilar Raman intensities (see 51 Fig. 6).

For biodistribution study performed at ultra-high SWNTdose, 4 ml of SWNT1-5kPEG solution with normal concentration (0.1 mg/ml) was concentrateddown to 0.8 ml of (0.5 mg/ml) by filtration through a 100"kDa filter. Thesolution was a black liquid with an optical density >20 at 808 nm (t-ern pathlength).

For biodistribution study, mice were killed at 1,30, 60, and 90 daysp.i., andthe organsltissues were collected, weighed, and solubilized in the lysisbufferusing a homogenizer (strong stirring and sonication, 1 min for each sample).After heating at 700e for =2 h, clear homogenate tissue solutions wereobtained for Ramanmeasurement. Acontrol experiment wasdone to confirmthat this treatment did not affect the Raman intensity of an SWNTsolution(data not shown). The biodistribution of SWNTs in various organs of mice wasthen calculated and plotted in unit of %IO/g basedon the following equation,

N. Nakayama-Ratchford, CD., N. W. S. Kam, P. Chu, Z.L., X.Sun, L. C. Cork, RD., and S. S. Gambhir, unpublished data).These two independent studies combined provide a strongindication of the lack of toxicity of well functionalized SWNTsin mice before clearance from the body. In contrast to a previousstudy of nonfunctionalized pristine carbon nanotubes causingfiber toxicity to mice (37), our well functionalized SWNTs arehighly biocompatibility for in vivo applications. The functionalization chemistry of nanotubes is critical to the in vivobehaviorsof nanotubes, including blood circulation, retention, excretion,and toxicity.

ConclusionWe have shown in the current work that the Raman spectroscopycan be used to detect carbon nanotubes in animals to glean theblood circulation behavior and biodistribution in main organs,especially in the RES. The robust Raman scattering property ofSWNTs allows us to track them for a long period with highfidelity, without the concern of labels falling off or decay overtime. It is found that the surface chemistry of nanotubes iscritical to their in vivo behavior, a conclusion that will likely applyto most nanomaterials, if not all. This result is expected becausepristine carbon nanotubes have very hydrophobic surfaces andare highly nonspecific in binding to biological species (38, 39).Recently, it has been shown that intravenously injected pristineSWNTs are highly rich in the lung as well as RES and remain inmice indefinitely (22). This hydrophobicity has to be blocked byproper chemical functionalization such as the PEG coatingsdescribed here, which enables biologically inert SWNTs withlong blood circulation, low RES uptake, and relatively fastclearance from organs and excretion from the body. The degreeof PEGylation of SWNTs is important to the in vivo behaviorsof nanotubes. Longer PEG chains, especially those withbranched structures, are excellent in affording SWNTs with themost desirable characteristics for in vivo applications. This resultshould also be applicable to functionalization of various othernanomaterials (nanocrystals, particles, etc.) for in vivo research.SWNTs detected in the feces of mice clearly reveal the biliaryexcretion pathway. A small percentage of nanotubes seems to beexcreted via the renal pathway. Last, no obvious toxic effect isfound in the necropsy, histology, and blood chemistry studies,which warrants the safety of properly functionalized carbonnanotubes for future in vivo biomedical applications.

Materials and MethodsVarious PEGylated Phospholipids (PL-PEGs) Used for Noncovalent Functionalization of Nanotubes.SeveralPL-PEGswithlinear PEG structure and one PL-PEGwith branched PEG structure were used in this study (Fig. la) includingcommercially available DSPE-PEG(2000)Amine (denoted as"L-I-2kPEG"whereI stands for linear, Avanti) and SUNBRIGHT OSPE-050PA (PL-I-5kPEG, NOFcooperation). PL-I-7kPEG was synthesized by mixing 1 eq of OSPE-050PA with2 eq of NHS-mPEG2000 (Nektar) in methylene chloride overnightfolJowed byaddition of 2 eq of N,N'-dicyclohexylcarbodiimide (Oee, Aldrich). Dee wasused to reactivate any potentially hydrolyzed NHS group on PEG duringstorage. The solvent was evaporated after another 24 hreaction. Water wasadded, and the insoluble solid (unreacted oeq was removed by vacuumfiltration. Thefinal productwasa clearwater solution and was stored at -20°Cfor future usage. PL-f-12kPEG and PL-br-7kPEG (where br stands for"branched") were synthesized by similar methods except for the startingmaterials. One eq of OSPE-PEG(2000)Amine and 2 eq of NHs-mPEG10000(Nektar) were usedto synthesize PL-I-12kpEG whereas 1eq of OSPE-050PA and1.5 eq of (Methyl-PEOt2h-PE04-NHS Ester (Pierce) were used to make PL-br7kPEG. Thefinal products were characterized and confirmed (data not shown)by MALOI (matrix-assisted laser desorption/ionization) massspectrometry inStanford PAN facility, showing no existence of starting PL-I-5kPEG or PL-I2kPEG materials. No further purification was performed because the excesshydrophilic NHS-PEG molecules were confirmed to exhibit no binding affinityto the hydrophobic nanotube surface. The excess NHS-PEGs were removedduring the nanotube filtration step.

1414 I www.pnas.org/cgijdoijl0.1073jpnas.0707654105 Liu et a/.

We used three to four mice per group at each time point poi. to obtain theaverage value and standard deviation for both blood circulation and biodlstribution measurements.

We also used a microRaman technique (15) to carry out Raman imaging ofSWNTs in liver slices.To obtain the Raman mapping image of liver slices (formice killed at 90 days p.i.), 5-Mm-thick paraffin-embedded liver slices weremounted on Si02 substrate and mapped under a Renishaw microRamanmicroscope with a line-scan model (100 mW laser power, 40 urn x 2 um laserspot size, 20 pixels each line, 2-s collection time, X20 objective). The SWNT

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PNAS I February 5, 2008 I vol. 105 I no.5 I 1415

Attachment Ib

Nano Res (2009) 2: 85-120DOl 10.1007/s12274-009-9009-8

~

Carbon Nanotubes in Biology and Medicine:In vitro and in vivo Detection, Imaging and Drug Delivery

ABSTRACTCarbon nanotubes exhibit many unique intrinsic physical and chemical properties and have been intensively

explored for biological and biomedical applications in the past few years. In this comprehensive review, we

summarize the main results from our and other groups in this field and clarify that surface functionalization

is critical to the behavior of carbon nanotubes in biological systems. Ultrasensitive detection of biological

species with carbon nanotubes can be realized after surface passivation to inhibit the non-specific binding

of biomolecules on the hydrophobic nanotube surface. Electrical nanosensors based on nanotubes provide

a label-free approach to biological detection. Surface-enhanced Raman spectroscopy of carbon nanotubes

opens up a method of protein microarray with detection sensitivity down to 1 fmol/L. In vitro and in vivo

toxicity studies reveal that highly water soluble and serum stable nanotubes are biocompatible, nontoxic,

and potentially useful for biomedical applications. In vivo biodistributions vary with the functionalization

and possibly also size of nanotubes, with a tendency to accumulate in the reticuloendothelial system (RES),

including the liver and spleen, after intravenous administration. If well functionalized, nanotubes may be

excreted mainly through the biliary pathway in feces. Carbon nanotube-based drug delivery has shown

promise in various In vitro and in vivo experiments including delivery of small interfering RNA (siRNA),

paclitaxel and doxorubicin. Moreover, single-walled carbon nanotubes with various interesting intrinsic

optical properties have been used as novel photoluminescence, Raman, and photoacoustic contrast agents for

imaging of cells and animals. Further multidisciplinary explorations in this field may bring new opportunities

in the realm of biomedicine.

KEYWORDSCarbon nanotubes, biomedical applications, surface functionalization, biosensor, drug delivery, biomedical

imaging

Introduction

Nanomaterials have sizes ranging from about one

nanometer up to several hundred nanometers,

comparable to many biological macromolecules

Address correspondence to hdaicsstanford.edu

such as enzymes, antibodies, and DNA plasmids.Materials in this size range exhibit interesting

physical properties, distinct from both the molecular

and bulk scales, presenting new opportunities forbiomedical research and applications in various

__________________lIilllTl _.'g~ ~:t ~, ~ •\ ..." j _ Springer

86

areas including biology and medicine. The emerging

field of nanobiotechnology bridges the physical

sciences with biological sciences via chemicalmethods in developing novel tools and platforms

for understanding biological systems and disease

diagnosis and treatment [1-3].

Carbon nanotubes (CNTs) are rolled up seamless

cylinders of graphene sheets, exhibiting unparalleled

physical, mechanical, and chemical properties which

have attracted tremendous interest in the past decade

[4-8]. Depending on the number of graphene layers

from which a single nanotube is composed, CNTs

are classified as single-walled carbon nanotubes

(SWNTs) or multi-walled carbon nanotubes

(MWNTs). Applications of CNTs span many fields

and applications, including composite materials [9],

nanoelectronics [10, 11], field-effect emitters [12],

and hydrogen storage [13]. In recent years, efforts

have also been devoted to exploring the potential

biological applications of CNTs, motivated by their

interesting size, shape, and structure, as well asattractive and unique physical properties [14-17].

With diameters of 1-2 nm, and lengths ranging

from as short as 50 nm up to 1 em, SWNTs are

one-dimensional (l-D) nanomaterials which may

behave distinctly from spherical nanoparticles in

biological environments, offering new opportunitiesin biomedical research. The flexible 1-D nanotube

may bend to facilitate multiple binding sites of a

functionalized nanotube to one cell, leading to a

multi-valence effect, and improved binding affinity

of nanotubes conjugated with targeting ligands.

With all atoms exposed on the surface, SWNTs have

ultrahigh surface area (theoretically 1300 m'/g) that

permits efficient loading of multiple molecules

along the length of the nanotube sidewall. Moreover,

supramolecular binding of aromatic molecules can be

easily achieved by rrvrt stacking of those molecules

onto the polyaromatic surface of nanotubes [18].

SWNTs are quasi 1-D quantum wires with sharp

densities of electronic states (electronic DOS) at the

van Hove singularities (Fig. l(a», which impart

distinctive optical properties to SWNTs [19]. SWNTs

are highly absorbing materials with strong optical

absorption in the near-infrared (NIR) range due to

Ell optical transitions (Figs. l(a) and l(b», and thus

Nano Res (2009) 2: 85-120

can be utilized for photothermal therapy [20, 21] and

photoacoustic imaging [22]. Semiconducting SWNTs

with small band gaps on the order of 1 eV exhibit

photoluminescence in the NIR range. The emission

range of SWNTs is 800-2000 nm [17, 23, 24], which

covers the biological tissue transparency window,

and is therefore suitable for biological imaging.

SWNTs also have distinctive resonance-enhanced

Raman signatures for Raman detection/ imaging,

with large scattering cross-sections for single tubes

[25, 26]. The intrinsic physical properties of SWNTs

can be utilized for multimodality imaging and

therapy.

In contrast to SWNTs, MWNTs are formed by

multiple layers of graphene and have much largerdiameters (1Q--100 nm), Although MWNTs exhibit less

rich and attractive optical properties than SWNTs,

their use in biological systems could be different from

that of SWNTs due to their larger sizes, which could

offer different platforms for different purposes, such

as delivery of large biomolecules including DNAplasmids into cells [27-30].

Motivated by various properties of CNTs,

research towards applying carbon nanotubes for

biomedical applications has been progressing rapidly.

CNT-based sensors have been developed to detect

biological species including proteins and DNA [14,

31, 32J. Relying on their optical properties, SWNTs

can be utilized as optical tags or contrast agents for

various biological imaging techniques [17, 22, 24,

26]. Our group and others have shown that properly

functionalized CNTs are able to enter cells without

toxicity, shuttling various biological molecular

cargoes into cells [15, 16, 29, 33-36J. Our latest study

has shown the promise of using CNTs for in vivocancer treatment in a mouse model [37J. However,

despite these exciting findings, researchers have

reported the negative sides of CNTs, showing that

nonfunctionalized nanotubes are toxic to cells andanimals [38-43]. The biodistribution and long-term

fate of CNTs have been explored by us and several

different groups, with different results obtainedusing different methods and materials [44-50]. These

controversial findings require clarification in order to

avoid confusion to the public.In this review, we first summarize the various

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Nano Res (2009) 2: 85-120 87

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IBm 2M ~ Springer

88 Nano Res (2009) 2: 85-120

routes used to functionalize carbon nanotubes

including covalent and noncovalent methods. Carbon

nanotube-based electronic and optical biosensors

are then discussed. Surveying our and others

results, we next describe work which has shown

that while nonfunctionalized, hydrophobic CNTs

have shown toxicity [38-43], those with carefully

designed biocompatible coatings are harmless to

cells In vitro [17, 18, 20, 51-57] and in vivo, at least

to mice within tested dose ranges [45, 58]. In terms

of biodistribution, although direct comparison

between different studies may not be valid because

of the different CNT materials used, tracking SWNTs

themselves by their intrinsic physical properties

(Raman scattering, photoluminescence, 13C isotope

mass spectrum) shows that, similar to other

nanoparticles in vivo, after systemic administration

SWNTs are predominantly localized in thereticuloendothelial system (RES) including the liver

and spleen [45, 48, 50]. We also review the current

progress in the use of carbon nanotubes for In vitrodrug delivery studies as well as pioneering efforts

towards in vivo cancer treatment. Lastly, SWNT

based biomedical imaging In vitro and in vivo are

discussed.

1. Functionalization of carbon nanotubes forbiological applications

As grown, raw carbon nanotubes have highly

hydrophobic surfaces, and are not soluble in

aqueous solutions. For biomedical applications,

surface chemistry or functionalization is requiredto solubilize CNTs,' and to render biocompatibility

and low toxicity. Surface functionalization of carbon

nanotubes may be covalent or noncovalent. Chemical

reactions forming bonds with nanotube sidewalls

are carried out in the covalent functionalization

case, while noncovalent functionalization exploits

favorable interactions between the hydrophobic

domain of an amphiphilic molecule and the CNT

surface, affording aqueous nanotubes wrapped by

surfactant.

1.1 Covalent functionalization of carbon nanotubes

Various covalent reactions have been developed

to functionalize carbon nanotubes, oxidation

being one of the most common. CNT oxidation

is carried out with oxidizing agents such as nitric

acid [59, 60]. During the process, carboxyl groups

are formed at the ends of tubes as well as at the

defects on the sidewalls. Zeng et al. observed

sp" carbon atoms on SWNTs after oxidation and

further covalent conjugation with amino acids

[61]. However, although oxidized CNTs are soluble

in water, they aggregate in the presence of salts

due to charge screening effects, and thus cannot

be directly used for biological applications due to

the high salt content of most biological solutions.

Further modification can be achieved by attaching

hydrophilic polymers such as poly(ethylene glycol)

(PEG) to oxidized CNTs, yielding CNT-polymer

conjugates stable in biological environments (Fig.

2(a» [18,58, 62]. We have used covalently PEGylated

SWNTs synthesized by this strategy for both In vitroand in vivo applications [18, 58].

Another widely used type of covalent reaction

to functionalize CNTs is the cylcoaddition reaction,

which occurs on the aromatic sidewalls, instead

of nanotube ends and defects as in the oxidation

case. [2+1] Cycloadditions can be conducted by

photochemical reaction of CNTs with azides (Fig.

2(b» [63, 64] or carbene generating compounds via

the Bingel reaction (Fig. 2(c)) [65, 66]. A 1,3-dipolar

cycloaddition reaction on CNTs developed by Prato

et al. is now a commonly used reaction (Fig. 2(d» [67,

68]. An azomethine-ylide generated by condensation

of an a-amino acid and an aldehyde is added to the

graphitic surface, forming a pyrrolidine ring coupled

to the CNT sidewall. Functional groups (e.g., amino

terminated PEG) introduced via a modified a-amino

acid can be used for further conjugation of biological

molecules such as peptides or drugs [36, 69].

Despite the robustness of the covalent func

tionalization method, the intrinsic physical properties

of CNTs such as photoluminescence and Raman

scattering are often destroyed after chemical reactions

due to the disrupted nanotube structure. The

intensities of Raman scattering and photoluminescence

of SWNTs are drastically decreased after covalent

modification, reducing the potential of optical

applications of these materials.

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Nano Res (2009) 2: 85--120 89

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Figure 2 Schemes of covalent functionalization of carbon nanotubes: (a) CNTs are oxidized and then conjugated with hydrophilicpolymers (e.q., PEG) or other functional moieties; (b) photoinduced [1,2] addition of azide compounds with CNTs; (c) Bingel reactionon CNTs; (d) l,3-dipolar cylcoaddition on CNTs. For biological applications, "R" in the figure is normally a hydrophilic domain which

renders CNTs water soluble. Further conjugationof bioactive molecules can be applied based on such functionalizations

1.2 NoncovaIent functionaIization of carbon nanotnbes

In contrast to covalent functionalization, noncovalent

functionalization of CNTs can be carried out bycoating CNTs with amphiphilic surfactant moleculesor polymers. Since the chemical structure of theIT-network of carbon nanotubes is not disrupted,

except for shortening of length due to the sonication

employed in the functionalization process, the

physical properties of CNTs are essentially preserved

by the noncovalent approach. Consequently, aqueous

solutions of CNTs, especially SWNTs, engineered

by noncovalent functionalization are promising for

~ Springer

90 Nano Res (2009)2: 85-120

(c)

Figure 3 Schemes of noncovalent functionalization of carbon nanotubes. (a)

Proteins areanchored on the SWNT surface via pyrene n-n stacked on a nanotubesurface. Right: A transmission electron microscope (TEM) image of an SWNT

conjugated with proteins. Copyright 2001 American Chemical Society [70]. (b)An SWNT coated bya single-stranded DNA via IT-n stacking. Copyright 2005 theNational Academy of Sciences [20]. (c) An SWNT functionalized with PEGylatedphospholipids. Both linear PEG (I-PEG) or branched PEG (br-PEG) can be used in thismethod. Copyright 2005 the National Academy of Sciences [45]

of SWNT-based biosensors [14]. Pluronic tri-block

polymer was used by Cherukuri et al. to solubilize

SWNTs for in vivo experiments [50]. However, the

Pluronic coating is not sufficiently stable and is

(a)

multiple biomedical applications including imaging.

The polyaromatic graphitic surface of a carbon

nanotube is accessible to the binding of aromaticmolecules via rrvrr stacking [70, 71]. Taking advantage

of the rr-rr interaction between pyrene and

the nanotube surface, we and others have

used pyrene derivatives to noncovalently

functionalize carbon nanotubes (Fig.

3(a)) [70, 72]. Chen et al. showed that

proteins can be immobilized on SWNTs

functionalized by an amine-reactive pyrene

derivative [70]. A recent study conducted

by Wu et al. also used pyrene conjugated

glycodendrimers to solublize carbonnanotubes [72]. Beside pyrene derivatives,

single-stranded DNA molecules have been

widely used to solubilize SWNTs by virtue

of the rr-rr stacking between aromatic DNA

base units and the nanotube surface (Fig.

3(b)) [20, 73, 74]. However, a recent report

by Moon et al. showed that DNA molecules

coated on SWNTs could be cleaved by

nucleases in the serum, suggesting that

DNA functionalization of SWNTs might

not be stable in biological environments

containing nucleases [75]. We also haveshown that fluorescein (FITC) terminated

PEG chains are able to solubilize SWNTswith the aromatic FITC domain rr-rt stacked

on the nanotube surface, yielding SWNTs

having visible fluorescence which are useful

for biological detection and imaging [76].

Furthermore, other aromatic molecules

such as porphyrin derivatives have also

been used for noncovalent functionalization

of CNTs [77].

Various amphiphiles have been used

to suspend carbon nanotubes in aqueous

solutions, with hydrophobic domains

attached to the nanotube surface via van

der Waals forces and hydrophobic effects,

and polar heads for water solubility [78].

We used Tween-20 and a Pluronic triblock

copolymer to noncovalently functionalize

nanotube surfaces to reduce the non

specific binding of proteins in the case

fi ill-----------------------N ana Research

Nano Res (2009) 2: 85-120

quickly replaced by serum proteins once SWNTs are

intravenously injected. Other traditional surfactants

including sodium dodecyl sulfate (SDS) and Triton

X-lOa have also been used to suspend CNTs in

water [79]. Carbon nanotubes solubilized by those

amphiphiles with relatively high critical micelle

concentrations (CMC) are typically not stable

without an excess of surfactant molecules in the

solution. Large amounts of surfactants may lyse cell

membranes and denature proteins, and are therefore

not useful in biological environments.

An ideal noncovalent functionlization coating

on CNTs for biological applications should have

the following characteristics. First, the coating

molecules should be biocompatible and nontoxic.

Second, the coating should be sufficiently stable

to resist detachment from the nanotube surface in

biological solutions, especially in serum having high

salt and protein contents. The amphiphilic coating

molecules should have very low CMC values so

that the nanotube coating is stable after removal of

most of the excess coating molecules from the CNT

suspension. Lastly, the coating molecules should

have functional groups which are available for

bioconjugation with antibodies or other molecules

to create various functional CNT conjugates for

different biological applications.

Noncovalent functionalization of SWNTs by

PEGylated phospholipids (PL-PEG) was developed

by our group to meet the above requirements,

including high water solubility of nanotubes and

versatile functionalities (Fig. 3(c» [18, 20, 35, 37,

44]. Phospholipids are the major component of

cell membranes, and are safe to use in biological

systems. The two hydrocarbon chains of the lipid

strongly anchor onto the nanotube surface with

the hydrophilic PEG chain extending into the

aqueous phase, imparting water solubility and

biocompatibility. Unlike nanotubes suspended by

typical surfactants, PEGy1ated SWNTs prepared by

this method are highly stable in various biological

solutions including serum, and even under harsh

conditions without requiring the presence of excess

PL-PEG (e.g., they are stable without coating

detachment upon heating in phosphate buffered

91

saline at 70 'C for weeks). PL-PEGs with different

PEG lengths and structures (linear vs, branched)

can be used to obtain specific PEGylated SWNTs

for desired applications. Conjugation of biological

molecules can be effected by using the functional

group (e.g., amine) at the PEG terminal. Relying on

this functionalization strategy, we have succeeded in

using SWNTs for a range of biomedical applications

including biological sensing, imaging and drug

delivery In vitro with cells or in vivo with animals [18,

20,24,35,37,44,45,51].

2. Selective protein-protein interactions onCNTs and biosensor applications

Functionalization strategies as presented above

suggest that both single-walled and multi-walled

carbon nanotubes may present scaffolds for

biomolecule immobilization, allowing subsequent

applications in biosensing, utilizing the intrinsic

electronic or optical properties of CNTs for signal

transduction. The remarkable physical properties

of carbon nanotubes, including high surface area,

semiconducting behavior, band gap fluorescence, and

strong Raman scattering spectra, lend themselves

well to measuring or detecting proximal or adsorbed

biomolecule interactions along the carbon nanotube

sidewall, at functionalized cap regions [80], and

even within the nanotube shell [81]. Proximity of

reasonably charged or polarized biomolecules yields

gating effects on isolated semiconducting carbon

nanotubes, or net semiconducting networks of

CNTs, thus yielding field -effect transistors (FETs)

capable of quantifying the degree of specific or non

specific binding of biomolecules [14]. Moreover, the

photoluminescent and Raman scattering properties

of single-walled carbon nanotubes may be applied

to biosensing, by specific conjugation of targeting

ligands to SWNT tags, coupled with sufficient

sidewall passivation in order to prevent non-specific

binding (NSB). Owing to their length scale and

distinctive structure, carbon nanotubes are of great

interest for the development of highly sensitive and

multiplexed biosensors for applications ranging from

the laboratory to the clinic.

l"il ~.r ~~ z, •\- ..... J _ Springer

92

2.1 Non-specific and specific protein-nanotube

interactions

In 1999, Balavoine et al. reported the crystallization

of streptavidin, a biotin-binding protein expressed in

Streptomyces avidinii, on the hydrophobic surface

of MWNTs. While stochastic binding of the ~70

kDa protein to the nanotube sidewall was the major

observation, helical packing of a streptavidin monolayerwas observed on occasion [82]. A similar observation

was found for the HupR protein, derived from

Rhodobacter capsulatus, suggesting that non-specific

interactions between proteins and carbon nanotubes arecommon for such water-soluble biomolecules.

Our group was the first to report that the interaction

between proteins and the CNTs sidewall is general,

and may be ascribed to hydrophobic interactions

between the exterior fullerene surface and regions of

high hydrophobic residue density within the protein

tertiary structure. As such, non-specific adsorption of

proteins does not appear to be dependent upon protein

isoelectric point (pl) and occurs in a variety of buffers

and solvents [14, 70, 83, 84]. To date, non-specific

adsorption of a variety of proteins, ranging in size and

pl, has been observed on both single- and multi-walled

carbon nanotubes (Fig. 4).

Ca)

(b)

Figure 4 Tapping mode AFM images of the non-specific adsorption

characteristics of streptavidin (SA) onto (a) bare single-walled carbonnanotubes (SWNTs) on SiO/Si, (b) Adsorption of SA onto SWNTs

and SiO/Si following application of Triton X-1QO. Copyright 2002

American Chemical Society [83]

Nano Res (2009) 2: 85--120

The mechanism by which water-soluble proteins

interact with bare carbon nanotubes was elucidated,

both by the discovery of peptides binding directly

to fullerene surfaces [85, 86], and by discovering

methodologies to prevent protein adsorption. Our

group demonstrated that amphiphilic coatings,

containing PEG components, such as Tween-20

and Pluronics, were able to effectively decrease the

hydrophobicity of the CNTs sidewall, thus reducing

and often eliminating the non-specific adsorption

of proteins [32, 83, 87]. This result suggested that

non-specific protein-CNT interactions are highly

dependent upon hydrophobic interactions [88], which

are thermodynamically unstable in the presence of

surfactants.

A variety of polymers and amphiphiles have

been demonstrated for CNT passivation, including

neutral molecules such as PL-PEGs [32], Triton

X-IOO [83], Tween-20, and Pluronics [14], all of which

contain PEG components. Resistance to protein

adsorption appears to scale with both the mass of

the hydrophilic PEG block, as well as the number ofPEG branches. Surfactant-coated carbon nanotubes

immobilized on substrates demonstrate degrees of

non-specific protein adsorption, ranging from partial

inhibition of NSB [14] imparted by Triton X-I00(containing linear PEG Mn~400 Da) to full inhibition

by Tween-20 (containing branched PEG, Mn~900 Da)

[14, 83]. Carbon nanotubes that are individualized

and suspended in the aqueous phase present

significantly higher sterically available surface area

(by mass) for non-specific protein interactions,

and thus require greater amounts of PEGylation

for sufficient passivation. It was experimentally

observed [32] that PL-PEG amphiphiles with PEG

having Mn~2000 Da did not efficiently prevent non

specific protein adsorption; however, longer linearPEG (Mn~5000 Da) or branched PEG (Mn~8000 Da)

prevented most interactions.

While carbon nanotube-protein interactions can

be observed via electron and force microscopies,

utilization of such supramolecular systems for

biosensing requires that protein activity and

specificity remain intact. A comparative study of theenzymes soybean peroxidase and a-chymotrypsin

non-specifically adsorbed onto SWNTs revealed

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~anoRes(2009)2:85-120

varying degrees of specific activity loss, seemingly

proportional to protein melting temperature andloading density [89]. Loss of specific enzyme activity

was correlated with inverse proportional changes

in the number of a-helices (increased) and B-sheets

(decreased) upon adsorption onto hydrophobic

SW~Ts.

For applications of molecular recognition

and readout based on carbon nanotubes, ligand

target affinity must be maintained. Thus, the non

specific immobilization of analyte proteins onto the

hydrophobic nanotube sidewall is not desirable,

owing to loss of function and lack of chemical control.

As described above, several methods for specific

nanotube functionalization exist; however, covalentmodifications to the nanotube sidewall damage the

interesting electronic and optical properties expected

to be utilized as readout mechanisms in biomolecular

detection.

In order to impart chemical functionality to

swms immobilized on substrates or suspended in

aqueous media (a requirement for most biosensing

applications) without hindering electrical or optical

readout, noncovalent approaches are critical.

Amphiphiles have been employed to impart

robust functionality to C~Ts as noted above, and

ideal candidates for biosensing applications must

simultaneously repel non-specific interactions and

biofouling by including an inert hydrophilic spacer

such as PEG.

Specific protein-nanotube conjugates, synthesized

via supramolecular chemistry, have been reported

in both surface-immobilized and solution phase

methodologies. A proof of principle study demon

strated the chemically directed conjugation of

streptavidin to bare SW~Ts on Si/ SiO" while non

specific adsorption was prevented by a combination

of Triton XI00 and PEG [83]. Additionally, thesurfactant Tween-20, adsorbed onto substrate

bound nanotubes, was covalently conjugated via

carbonyldiimidazole to biotin, Staphylococcal protein

A (SpA), and UIA antigen [14, 70J, which imparted

specific binding of streptavidin, immunoglobulin

G, and the monoclonal mouse antibody 10E6,

respectively, while preventing non-specific

adsorption of non-target proteins.

93

2.2 Electrical detection of protein-nanotube

interactions: Applications and mechanism

Following the first demonstration of SW~Ts acting

as molecular sensors via field-effect doping by the

gaseous species ~H3 and ~02 [90], our lab amongst

others set out to utilize field-effect transistors based

on carbon nanotubes for label-free, highly specific

biomolecule detection. As noted above, conjugation

of biotin, Staphylococcal protein A (SpA), and UIA

antigen to SW~Ts [14, 70], amongst other ligands

with specific binding characteristics, has been

reported to impart specific binding of streptavidin,

immunoglobulin G, and the monoclonal mouse

antibody 10E6, respectively, allowing in situ direct

detection of these analytes in the nmol/L range via

crrr FET devices (Fig. 5).

Surprisingly, in all cases of C~T FET-based

detection, conductance across a semiconducting

network of C~Ts was reduced following specific

analyte protein binding, regardless of analyte pI

[14, 87]. Further studies revealed the importance

of Schottky barrier modulation as a result of

biomolecule adsorption/binding at nanotube rmetal

electrode contacts in detecting biomolecules with

low net charge [87], thus elucidating the unexpected

gating behaviors observed in FET biomolecule

detection.

Understanding the Schottky barrier modulation

mechanism of carbon nanotube FETs in biomolecule

detection allowed for further improvements to

be made in device architecture, thus allowing

greater sensitivity and lower limits of detection

to be coupled with specific binding, imparted

by supramolecular ligand conjugations. Angular

shadow mask deposition of Au/ Cr metal contacts

onto semiconducting SW~T networks increased the

contact area and reduced the contact thickness, raising

hopes of increasing the Schottky barrier modulation

component in FET conductance measurement of

biomolecules (Fig. 6). As hypothesized, increased

Schottky barrier area was correlated with improved

sensor device characteristics, including a reduction in

specific detection limit by four orders of magnitude,

over a -1 umolj L to 1 pmol/L range of analyte

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94 Nano Res (2009) 2: 85-120

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Figure 5 Specific chemical conjugation of biomolecules to SWNTs coated in Tween-20 surfactant, which prevents non-specificinteractions, allows both quartz crystal microbalance (QeM) and FET measurementof target analyte binding. SWNTs functionalizedby biotinylated tween-zc (a)demonstrateQeMshifts dependentupon analyte, streptavidin, concentration (b), and analyte mediatedconductance changes (c)asmeasured across a mat of SWNTs. (d) SWNTs mayalsobe selectively coupledto Staphylococcal protein A(SpA) for selective binding of murine IgGasevidenced by QCM (e)and FET (f) measurements. Copyright2003 the National Academyof Sciences [14J

BSA 1 pmollLI

200 400 600Time (s)

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400 800 1200 1600Time (s)

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~

Figure 6 SWNTnetwork device architecture for increased Schottky contact area affords biosensors of increased sensitivity:(a) Shadow mask deposition of Au/Cr contacts at a 23° angle onto a semiconducting SWNT network increases Schottky contactarea; (b) FET devices fabricated from SWNT networks with high Schottkycontact area demonstrate enhanced sensitivities and limitsof detection in specific biomolecule detection by four orders of magnitude.Copyright 2006 American Chemical Society [91]

iLi4NanD Research l!IittliittliiOliittliiOliiOlii1llll1llll1llll1!22!!OIiiOlii!illll!illll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1llll1lllll

Nano Res (2009) 2: 85-120

protein concentration [91].

A variety of carbon nanotube FETs and

electrochemical devices have been demonstrated for

selective detection of oxidase and dehydrogenase

activity, as well as for other enzymes and

biomolecules of interest, in label-free fashion [92,

93]. However, the sensitivity of such devices in

performing detection of biomolecules is limited by

charge screening of both CNT-bound ligands and ions

common in physiological buffers. Schottky barrier

modulation and chemical gating effects of FETs are

distance-dependent processes, thus reducing the

utility of CNT FETs for complex (indirect) bioassays.

Especially, troublesome for FET applications in

detection of biomolecules is the incompatibility

of common aqueous buffers with conductance

modulated readout. The high ionic strength of

physiological buffers, often required to retain protein

structure and function, shields FET devices from

charge effects of analyte biomolecules, thus reducing

the simplicity and utility of this detection strategy.

As a result, the detection of analyte concentrations in

the nmol/L to pmol/L range has yielded poor signalto-noise ratios with these devices.

2.3 Photoluminescent detection of proteins-based

on semiconducting SWNTs

Exploiting the interesting optical properties of

semiconducting SWNTs, rather than monitoring

their cond uctance in transistor devices, opensanother route for sensitive and selective biomolecule

detection using these extraordinary materials. SWNT

band gap fluorescence [23] has been explored as a

methodology for NIR-imaging of both In vitro and

in vivo biological systems [17, 24, 26, 50, 94, 95J, and

holds promise for In vitro biomolecule detection

assays in both direct and sandwich-assay formats [96

-98J. SWNT NIR fluorescence does not photobleach

under high excitation powers and benefits from

the negligible auto-fluorescence contributions of

other assay components in the NIR range. SWNT

fluorescence also demonstrates a large Stokes shift

compared with traditional fluorophores, and allows a

range of excitation energies to be used.

As described above, aqueous phase processing of

SWNTs is made possible by PEGylated amphiphiles,

95

which reduce biofouling and provide sites of

functionality for ligand conjugation to the SWNT

tags. Recently, our group has demonstrated specificand sensitive detection of biomolecules in sandwich

assay format, using the band gap photoluminescence

of SWNTs as NIR fluorophores, with sensitivity from

the micromolar to the picomolar range (unpublished

results). SWNT NIR fluorescence may be multiplexed

via microarray printing technology, and unique

SWNT fluorophores may be obtained by chirality

separation [74]. NIR fluorescence detection via

SWNT fluorophores may facilitate high throughput

bioassays with low background contributions

and thus improved sensitivity over conventional

techniques.

In addition to direct measurement of SWNT

fluorophore emission in immunoassay formats,

Strano and coworkers have sought to use band

gap modulation and charge transfer effects

via photoluminescence for transduction andquantification of biomolecules such as DNA and

glucose. Noncovalent functionalization of SWNTs

with 24-mer ssDNA and subsequent hybridization

of eDNA in proximity to the SWNT surface alters the

dielectric constant at the SWNT surface, and produces

a 2 meV increase in band gap energy [97], observed

as a blue shift in emission. Such methodology yields

a theoretical detection limit of 6 nmol/L for 24-mer

DNA. Moreover, utilizing a similar noncovalent

modification strategy, the same group demonstrated

signal transduction via fluorescence quenching for

measuring glucose concentrations at physiologically

relevant conditions, from the micromolar to

millimolar range [98J. Direct detection of protein

binding events by relief of band gap fluorescence

quenching has also been demonstrated. Satishkumar

et al. demonstrated that small molecule quenchers

may be removed from SWNT surfaces by avidin

and albumin in a specific and non-specific manner,

respectively, with detection limits in the micromolar

range [99].

In addition to quenching of the inherent SWNT

band gap fluorescence for sensor applications, specific

detection of biomolecules by quenching may also be

realized by applying the carbon nanotube as both a

scaffold for recognition ligands and as a quencher of

I~ ~:f ~, " •\. "... J _ Springer

96 Nano Res (2009) 2: 85-120

small molecule fluorophores. An excellent example of

such quenching assays is the use of molecular beacons in

real-time polymerase chain reaction (PCR) applications.

In comparison with commonly used molecular beacons,

SWNTs-noncovalently functionalized by FAM

labeled ssDNA-demonstrate superior quenching in

the unhybridized state, and thus improved signal

to-noise ratios when hybridized to a complementary

strand [100]. Additionally, this new class of beacon

affords improved thermal stability and is general to a

wide variety of fluorophores.

While SWNT NIR band gap photoluminescence is

promising for biomolecule detection, several problems

remain to be solved. While only semi-conducting

SWNTs demonstrate band gap photoluminescence,

separation, and isolation of these fluorophores from

other nonfluorescing SWNT isomers is challenging.

Moreover, the quantum yield of those SWNTs that do

fluoresce in the NlR is dependent upon their chemical

environment, and processing is required to avoid

quenching and maximize quantum yield [101]. Signal

transduction via band gap modulation and quenching

suffers from the limits of spectral resolution, as well

as photoluminescence intensity, which restricts the

utility of these methods to analytes at relatively high

concentration. Applications of SWNTs as quenchers

are promising for detection of ssDNA hybridization,

however additional work is necessary to monitor

protein interactions by this method. Future

elucidation and optimization of SWNT fluorescence in

the NIR range, as well as isolation of semiconducting

SWNTs [74, 102] may improve the photoluminescent

detection limit of SWNT fluorophores in protein

assays by many orders of magnitude.

2.4 Surface-enhanced SWNT-Raman tags for highly

sensitive detection of proteins

To avoid the issues plaguing SWNT photo

luminescence-based detection of proteins Invitro, our

lab sought to utilize the intense Raman scattering

cross section of SWNTs in immunoassay format.

While methodologically similar to fluorescence-based

sandwich assays [103-105] the application of SWNT

Raman tags in lieu of traditional or nontraditional

[106] fluorophores offers many potential benefits.

In addition to scattering efficiencies that rival the

quantum yield of organic fluorophores [26], the

Raman scattering spectra of SWNTs are simple,

with strong, well-defined Lorentzian peaks of

interest, demonstrating FWHM of 1-3 nm. As such,

the Raman scattering spectra of SWNTs are easily

distinguishable from noise, and no "auto-scattering"

is observed for conventional assay surfaces or

reagents [32]. Photobleaching of SWNT Raman tags

is not observed even under extraordinarily high laser

powers, a reflection of the stability of the SWNT sp'

carbon lattice.

Surface-enhanced Raman scattering (SERS) [106]

is a technique that may be applied to vastly increase

the intensity of Raman active molecules in proximity

to appropriately tuned surface plasmons, usually

associated with gold, silver, or copper nanostructures

[107]. Coupling the intense resonance enhancement

of 1-D SWNT Raman tags with SERS presents the

opportunity to extend the limit of detection of

traditional fluorescence assays from approximately

1 pmol/L [108] to the femtomolar level or below. By

fabricating a gold-coated assay substrate via electron

beam evaporation and roughening the gold surface

via annealing at 400°C in H, following analyte and

tag binding, our group was able to demonstrate

quantitative SERS over a large area. While such

treatment would destroy most small, organic

Raman active molecules, SWNTs are robust and are

undamaged by the process. This strategy for SERS

yielded a nearly 100-fold increase in SWNT Raman

scattering intensity [32] (Fig. 7).

As previously discussed, noncovalent surfactant

wrapping may be used to both functionalize and

passivate SWNTs. Recently, our group employed

SWNTs, suspended by linear PL-PEG-NH, (M, ~5000

Da) and branched PL-mPEG (M, ~8000 Da) (Fig. 7),

coupled to goat anti-mouse immunoglobulin G (GaM

- IgG) for specific detection in immunoassays. To

test the selective binding and non-specific behavior

of such conjugates, a variety of analyte proteins

were immobilized on SERS-active substrates, and,

following incubation with GaM-IgG-SWNTs,

selective detection of mouse IgGs was observed (Fig.

8). Extending this methodology to concentration

dependent sandwich immunoassays, we have been

able to push limits of detection to 1 fmol/L of model

dEN ana Research lilliEiii:1IIi~_1IIi~:fJli:0?Jif.gMmIlli1.,"!IIIIIIIIIIIIIIIIIIIIIIIIIii 1llllll1IIIIii1llllll1llllllll1llllll1llllll1llllll1llllllll1llllllll1llllllll1llllll1llllllll1llllllll1llllllll1llllll1llllllll1llllllll1llllllll1llllllll1llllllll1llllllll1llllllll1llllllll1llllllll

Nano Res (2009) 2: 85-120 97

120 G+- BeforeSERS

.i;' 90 --After SERS~--:-$ ~

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CO00 ~E~ G-oo 30

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200 240 280Raman shift (em-')

17001650160015500~~E::::::;:::==:;::=::;=:===~=:;::==;1500

Raman shift (cnr')

(a)

DSPE-PEG-NH, (5 kDa)

11 ~jOvJ;NH--<,>r,"'-_",~.'l?~O"" 0

''r,

1 :1 DSPE-PEG R t/ mixture '"<amen ag

Cb)

Figure7 SWNT-antibody conjugates may be utilized in conjunction with SERS asbright Ramantags for sandwich assay protein detection: (a) preparation of a spatially uniform SERS substrate

for SWNTRaman tag detection of biomolecules significantly increases Raman scattering intensity,thus improving signal-ta-noise and reducing assay time; (b) SWNT-antibody conjugates wereformulated by first suspending SWNTs in aqueous mediavia PEGylated surfactants, which provide

functionality and prevent non-specific interactions to the hydrophobic 5WNT surface, andsubsequently coupledto antibodiesvia bifunctional cross-linkers. Such antibody-Raman tags havebeenusedin direct and indirect immunoassays. Copyright2008 NaturePublishing Group [32]

100

10

I

200

Ca) (b)

Figure 8 SWNT-Raman tags have been applied to immunoassays demonstrating both high selectivity and sensitivity: (a) SWNT-anti-mouseIgG conjugates were applied to direct detection of mouse IgGs, demonstrating excellent signal-to-noise and minimal cross-reactivity; (b) 5WNTanti-mouse IgGconjugates were used as Raman tags for the indirect detection of mouse anti-human serum albumin (aHSA) IgGcaptured ontoan SERS active substrate by HSA. A limit of detection of 1 fmol/L analyte was reproducibly observed (two separate trials are shown). The dataarewell fitted by a logistic regression (solid red curve, fit to data shown in green)allowing accurate quantitation of ana!yte over eight orders ofmagnitude.Copyright 2008 Nature Publishing Group [32]

~ Springer

98 Nano Res (2009) 2: 85-120

analyte, approximately three orders of magnitude

better than common fluorescence methods.

Ultrasensitive protein detection by SWNT Raman

tags has been demonstrated not only for model

ana1ytes, but also for true biomarkers of human

autoimmune disease [32] and cancer (unpublished

results), with dynamic ranges over 6-8 orders of

magnitude (Fig. 8).Biomolecule detection by SWNT Raman tags

appears to be generalizable to systems other

than high affinity antigen-antibody interactions,

including biotin-streptavidin binding, protein

A/G-IgG interaction and DNA hybridization

[32]. By coupling the characteristic intense

Raman scattering efficiency of 1-D SWNTs with

quantitative SERS substrates and noncovalent

strategies for both specific antibody conjugation

and non-specific binding passivation, highly

sensitive biosensors have been developed and

demonstrated. Moreover, by taking advantage of

highly multiplexable microarray technology, and

employing isotopically labeled SWNTs (composedof pure 12e and 13e, respectively) our group has

demonstrated multicolor detection of multiple

analytes simultaneously, utilizing only a single

excitation source (Fig. 9). The application and

utility of such sensitive, multiplexed biosensors

remains to be fully explored, though with recent

advances in biomarker discovery [109] there is

much promise for detection and monitoring of

early stage cancer.

(a)

Raman shift (cm-')

lJC-SWNT

r1650

"C-$WNT

rAnti-mouse

IgG

160015501500

10

£ 8~-:-Q) ~ 6E a;.~ 0"'~ 4Ec'"no 2

0

Cb) (c)

Figure9 SWNT tags made from pure 12e and Be feedstocks may be used for multicolor Raman-based protein detection:(a)Schematic diagram of a protein rnicroarray employing multi-color SWNT tags for high throughput, multiplexed detection;(b) 12C and 13e SWNTs demonstrateeasily resolvable c-band Raman scattering spectra; (c) direct Raman detection of mouseIgG (red) and human IgG (green) by "C and BC SWNT tags. respectively. Copyright 2008 Nature Publishing Group [321

z: ._lIIiIilllil__,!ili! _

N ana Research

Nano Res (2009) 2: 85-120

3. Toxicity of carbon nanotubes

Safety is the first requirement of any material used

in medicine. A large number of studies have been

performed in the past several years to explore the

potential toxic effects of carbon nanotubes. The

conclusions of these reports varied drastically,

showing a large dependence on the type of nanotube

materials as well as functionalization approaches.

Cell culture experiments and in vivo pilot studies

conducted by various groups observed no obvious

toxicity of properly functionalized carbon nanotubes

[15, 53, 58, 72]. On the other hand, raw carbon

nanotubes were shown to be toxic to mice after

inhalation into the lung [39, 40, 110, 111]. Recent

research showed that unfunctionalized, long MWNTs

may pose a carcinogenic risk in mice [43]. As a result

of the wide variety of reports, both the public and

research communities are currently concerned about

using carbon nanotubes for biomedical applications.

It is thus critical and urgent to clarify the toxicity

issue of carbon nanotubes. The current status is that

toxicity appears to be dependent on the material

preparation, especially geometry and surface

functionalization. Well functionalized CNTs with

biocompatible surface coatings have been shown to

be nontoxic In vitro to cells and in vivo in mice.

3.1 In vitro toxicity of carbon nanotubes

Even in cell culture experiments, the issue of

toxicity of carbon nanotubes is still controversial.

While inhibition of HEK 293 cell proliferation

after exposure to SWNTs was reported by Cui et

al. [38], Ding et al. observed that MWNTs induce

cell cycle arrest and increase apoptosis/ necrosisof human skin fibroblasts [41]. However, neither

of these studies used functionalized carbon

nanotubes. Apoptosis of T lymphocytes induced

by oxidized MWNTs was observed by Bottini et

al. [42]. However, simple oxidation is not enough

to render carbon nanotubes soluble and stable in

saline and cell media, and thus does not represent a

biocompatible functionalization. Sayes et al. further

reported that the toxicity of CNTs was dependent

on the density of functionalization, with minimal

toxicity for those heavily functionalized with

99

the highest density of phenyl-St.i.X groups [112].

These results are understandable because CNTs

without proper functionalization have a highly

hydrophobic surface, and thus may aggregate in

the cell culture and interact with cells by binding to

various biological species, including proteins, via

hydrophobic interactions, and induce certain cell

responses such as cell toxicity.

Other factors may also contribute to the observed

toxicity of CNT samples In vitro. Surfactants, present

in excess in the CNT suspensions, are known to be

highly toxic to cells [113]. The metal catalyst content

in CNTs should also be considered when the toxicity

of carbon nanotubes is investigated [114]. Moreover,

proper assays must be employed in toxicity tests

to avoid interference of carbon nanotubes with

the assay reagents [115, 116]. For these reasons, In

vitro toxicity assays of carbon nanotubes should be

carefully designed and performed with well prepared

and characterized materials, as well as suitable assay

methods.

We and many other groups have successfully

used well functionalized, serum stable carbon

nanotubes for In vitro cellular uptake experiments

without observing apparent toxicity [17, 18, 20, 51

57]. Cells exposed to SWNTs, PEGylated by various

PL-PEG amphiphiles, used in our work exhibited

neither enhanced apoptosis/neurosis, nor reduced

proliferation of various cell lines In vitro [18, 20,

51]. Carbon nanotubes covalently functionalized

by 1,3-dipolar cycloaddition developed by Prato et

al. also appeared to be safe to the tested cell lines,

including primary immune cells [52, 53]. Carbon

nanotubes with a biomimetic coating engineered

by Bertozzi et al. were also nontoxic to cells [54, 55].

Several other independent groups also reported that

CNTs coated by DNA, amphiphilic helical peptides

and serum proteins were not toxic to cells [26, 56,

57]. In very recent work, Jin et al. discovered that

SWNTs taken up by cells via endocytosis exited cells

through exocytosis without affecting the viability

of cells [95]. It appears that raw CNTs and CNTs

without serum-stable functionalization show toxicity

to cells at moderate dosage, while serum-stable,

functionalized CNTs show little toxicity even at high

dosages.

l~' bo. S .\~~ ~ pnnger

100

3.2 In vivo toxicity of carbon nanotubes

To address the possible side effects of CNTs on

human health and our environment, researchershave investigated the toxicology of CNTs in

animal models. Unfunctionalized raw CNTs have

been intratracheally (IT) instilled into animals,

showing obvious pulmonary toxicity including

unusual inflammation and fibrotic reactions due

to the aggregation of hydrophobic raw CNTs in

the lung airways [39, 40, 110, 111]. Those results

suggest that aerosol exposure of raw CNTs in the

workplace should be avoided to protect humanhealth. Nevertheless, toxicities observed by

intratracheal instillation of large amounts of raw

CNTs may have little relevance to the toxicology

profile of functionalized soluble CNTs for biomedical

applications, especially when they are administered

through other routes such as intraperitoneal (IP) and

intravenous (IV) injections, by which lung airways

are not exposed to CNTs.

In a recent pilot study, Poland et al. noticed

asbestos-like pathogenic behaviors such as

mesothelioma associated with exposing the

mesothelial lining of the body cavity of mice to

large MWNTs (length 10-50 urn, diameter 80

160 nm) following intraperitoneal injection [43].

Despite the importance of this finding for potentiai

negative effects of CNTs to human health, it

should be noted that the MWNT materials used in

this study were simply sonicated in 0.5% bovine

serum albumin (BSA) solutions without careful

surface functionalization and hence are not directly

meaningful as far as the functionalized CNTs with

biocompatible coatings recommended for biomedical

applications are concerned. Furthermore, length

dependent pathogenicity was observed, as no

obvious toxic effect was observed for shorter and

smaller MWNTs (length 1-20 I'm, diameter 10-14

nm), indicating that the toxicology profiles of CNTs

may significantly differ between CNTs of various

sizes (diameter and length). It is worth noting that

functionalized SWNTs used in typical biomedical

research have length 50-300 nm and diameter 1-2

nm, which are entirely different from the geometry

of MWNTs used by Poland et al.

Nano Res (2009)2: 85-120

The first reported in vivo toxicity study of

functionalized SWNTs was conducted by the

Gambhir group and our group [58]. Both covalently

and noncovalently PEGylated SWNTs were used

in this study. Mice intravenously injected with

PEGylated SWNTs (-3 mg/kg) were monitored over

four months, with systolic blood pressure, complete

blood counts and serum chemistry recorded every

month. Careful necropsy and tissue histology

examinations were performed at the end of four

months. Normal blood chemistries and histological

observations were observed in this study, suggesting

that functionalized biocompatible SWNTs may be

safe for in vivo biological applications. Another

separate study by our group showed similar

results, suggesting that PEGylated SWNTs are

slowly excreted from the body following systemic

distribution in mouse models, without exhibiting

obvious toxicity in the process [45]. Recently, Yang

et al. showed in a three-month toxicity study that

SWNTs suspended by Tween-80, which is probably

not an ideal coating molecule, exhibited low toxicities

to the tested mice at a very high dose (-40 mg/kg)

following IV administration. Such toxicity may

be due to the oxidative stress induced by SWNTs

accumulated in the liver and lungs [117]. The toxicity

observed was dose-dependent, and appeared to be

less obvious at lower doses (2 mg/kg and 16 mg/kg).

Another recent report by the same group showed

that their covalently PEGylated SWNTs, with much

higher aqueous stabilities and biocompatibilities,

exhibited an ultralong blood circulation half-life in

mice [118]. Although the long-term toxicology of such

"improved" SWNTs has yet to be determined, no

acute toxicity has been reported even at a high dose

(24mg/kg).

To fully address the toxicity concern of CNTs,

further investigations-including animal models

other than mice-and on larger scales, are still

required. Moreover, the interactions betweenadministered CNTs and the immune complement

system, whose activation is an important first line ofdefense against foreign species, especially microbes,

require more attention [119]. Moreover, increased

efforts are needed not only from the chemical aspect

i.e., further ?ptimizing CNT surface chemistry and

Nano Research 1lIIII1lIIII1lIIII~~~iII.I11IIII11IIII11III__&I11IIII11IIII11III.I11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11IIII11III

Nano Res (2009) 2: 85-120

geometry for improved biocompatibility, but also

from those with biological expertise, to systematically

study the complete CNT toxicology profile in

different animal models with different routes of

administration.

4. In vitro delivery of biomolecules bycarbon nanotubes

The work using carbon nanotubes for drug delivery

in our group was triggered by an unexpected finding

that functionalized CNTs are able to enter cells by

themselves without obvious toxicity [15]. Similar

results were published by the Prato group around the

same time [36]. The CNT cellular uptake mechanism

may differ depending on the functionalization and

size of the CNTs, including endocytosis as reported

by us and several other groups [15, 26, 34, 56,

95], or passive diffusion as observed by the Prato

group when CNTs are functionalized by 1,3-dipolar

cycloaddition [16, 120]. CNTs have been used to

efficiently shuttle various biological cargoes, ranging

from small drug molecules to biomacromolecules,

such as proteins and DNA(RNA, into different types

of cells. Once taken up by cells via endocytosis,

SWNTs are able to exit cells through exocytosis [95].

4.1 Delivery of small drug molecules by carbon

nanotubes

Small drug molecules can be covalently conjugated to

CNTs for In vitro delivery. Fluorescent dyes and drug

cargoes were simultaneously linked to 1,3-dipolar

cycloaddition functionalized CNTs via amide bonds

for the delivery of an anti-cancer drug [69] or an

antifungal drug [52] into cells. In collaboration with

our group, Feazell and Lippard used noncovalently

PEGylated SWNTs (using PL-PEG, Mn-2000 Da)

as a longboat delivery system to internalize a

platinum(IV) complex, a prodrug of the cytotoxicplatinum( II), into cancer cells [121]. The inert

platinum(IV) prodrug compounds developed by the

Lippard group are activated only after being reduced

to the active platinum( II) form. SWNTs tethered

with the platinum(IV) complexes through peptide

linkages are taken into cancer cells by endocytosis

and reside in cell endosomes, where reduced pH

101

induces reductive release of the platinum( 11) core

complex, thus killing the cancer cells. The cytotoxicity

of the platinum(IV) complex increases over 100-fold

after attachment to SWNTs. We have also conjugated

paclitaxel, a commonly used anti-cancer drug, to

branched PEG-coated SWNTs via a cleavable ester

bond [37]. The SWNT-PTX conjugate was tested both

In vitro and in vivo.

Beside covalent conjugation, novel noncovalent

supramolecular chemistry for loading aromatic

drug molecules onto functionalized SWNTs by IT

-IT stacking has been uncovered in our lab (Figs.

10 (a) and 10(b)) [18]. Doxorubicin, a commonly

used cancer chemotherapy drug, can be loaded on

the surface of PEGylated SWNTs with remarkably

high loading, up to 4 g of drug per 1 g of nanotube,

owing to the ultrahigh surface area of SWNTs. The

loading(binding is pH dependent and favorable for

drug release in endosomes and lysosomes, as well as

in tumor micro-environments with acidic pH (Fig.

10(c)). Similar drug loading behaviors have been

reported for MWNTs [122], single-walled carbon

nanohorns [123] and nano-graphene oxide [124, 125J.

The supramolecular approach of drug loading on

CNTs opens new opportunities for drug delivery.Targeting ligands including folic acid [20, 126],

peptides [18, 44] and antibodies [24, 127-129] have

been used to target CNTs to specific types of cellsIn vitro or to tumors in vivo. Targeted drug delivery

with CNTs requires conjugation of both targeting

molecules and drug molecules to the same nanotube,

and thus requires carefully designed strategies [18,

126]. In the work reported by Dhar et al., folic acid

(FA) was linked to a Pt(lV) prodrug compound,

and then conjugated to PEGylated SWNTs [126],

yielding an SWNT-Pt(IV)-FA conjugate that showed

enhanced toxicity to folate receptor (FR) positive

cells but not to FR negative cells as the result of FA

targeted delivery. For the delivery of aromatic drugs

such as doxorubicin, which are directly loaded on

the nanotube surface via IT-IT stacking, the functional

groups on the SWNT coating molecules (e.g., PL-PEG

-amine) can be conjugated with targeting molecules

such as Arg-Cly-Asp (RGD) peptide for targeted

delivery (Fig. 10(a)) [18].

Besides drug conjugation and loading on the

~~.~ ~§ Ill' . .\ ..... J _ Springer

102 Nano Res (2009) 2: 85-120

to

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o

Figure 10 Supramolecular chemistry of functionalized SWNTs for efficient drug loading and delivery. (a) Schematic of doxorubicin (DOX)

n-stackinq onto a nanotube pre-functionaiized by Pl-PEG. Targeting ligands such as RGD peptide can be conjugated on the PEG termini for

targeted drug delivery. Inset: AFM images of SWNT before (top) and after (bottom) DOX loading. The height of SWNTs increased after DOX

loading. (b) UV-vis-NIR absorbance spectra of solutions of free doxorubicin (green), plain SWNTs (black), and DOX loaded 5WNTs. (c) Releasecurves of DOX from SWNTs at different pH. DOX loaded on SWNTs is stable at basic and neutral pH with very slow release but exhibits faster

release in acidic environments. Copyright 2007 American Chemical Society [18]

external surfaces of nanotubes, the hollow structure

of CNTs may allow the encapsulation of drugmolecules inside nanotubes for drug delivery.Fullerene balls [130], metal ions [131J, smallcompounds such as metallocenes [132], and evenDNA molecules [133] have been encapsulated insideCNTs. Although a number of theoretical modelingstudies predicted the insertion of biomoleculesincluding chemotherapy drugs [134, 135] into CNTs,drug delivery by encapsulation of drugs insideCNTs has been rarely reported. Further experimentalstudies are still needed to examine the possibility ofutilizing the encapsulation strategy in CNT-baseddrug delivery.

4.2 Delivery of biomacromoiecuies by carbon

nanotubes

Unlike various small drug molecules which are

able to diffuse into cells, biomacromolecules

including proteins, DNA, and RNA rarely cross cell

membranes by themselves. Intracellular delivery

is thus required in order to use these molecules

for therapeutic applications. Proteins can be either

conjugated or noncovalently absorbed on nanotubes

for intracellular delivery [15, 33]. The hydrophobic

surface of partially functionalized SWNTs (e.g.,

oxidized SWNTs) allows non-specific binding of

proteins. After being translocated into cells by

2££ 2IlIIllIlllllIlllllWl~WI~ilj1IIII1IIII1IIII1lIIll1lllll.1IIII1IIII.1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIII1IIIIN ano Research

Nano Res (2009) 2: 85-120 103

160

120[

80

40

o

200

201510fJi'tl

5

5

o

10

Ca)

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(d)

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2015

o..oj-o,. ..

, o'

oo·~O·,..i'j:~'.·:.'

o-

10fJnl

5

favorable for siRNA delivery into cells (Fig. l1(d)). We

propose that our SWNTs functionalized with short

PEG (2 kDa) groups retain a certain hydrophobicity

5

o

20

15

E::> 10

Figure 11 siRNA delivery bycarbon nanotubes: (a) a scheme of 5WNT

siRNA conjugation via disulfide linkage; (b)confocal images of untreated cells(left) and SWNT-siRNAcxCR4 treated cells (right) after PE-anti CXCR4 staining.Scale bars: 40 IJm; (c) CXCR4 expression levels on (EM cells three daysafter various treatments, including four types of liposomes (Upo1-4) and

luciferase (Luc) siRNA control; (d) G-mode Raman intensity maps of single(EM cells afterincubation for 1 d in SWNTs functionalized by PL~PEG2000 (left)and PL-PEGS400 (right) chains respectively. Inset: optical microscope images ofCEM cells. Copyright 2007 Wiley-VCH [511

nanotubes, proteins can become bioactive once they

are released from endosomes [33].

CNTs can be modified with positive charges to

bind DNA plasmids for gene transfection [27-

30]. Pantarotto et al, and Singh et al, used amine

terminated SWNTs and MWNTs functionalized

by 1,3-dipolar cycloaddition to bind DNA

plasrnids, and have achieved reasonable

transfection efficiency [27, 29]. In the work of Gao

et al., amine groups were introduced to oxidized

MWNTs for DNA binding and transfection,

successfully expressing green fluorescence

protein (GFP) in mammalian cells. Although the

MWNT-based method was less efficient than

commercial gene transfection agents, such as

lipofectamine 2000, the MWNTs exhibited much

lower toxicity [30]. In another study carried out

by Liu et al., polyethylenimine (PEI) grafted

MWNTs were used for DNA attachment and

delivery, which afforded comparable efficacy to

the standard PEl transfection method with the

benefit of reduced cytotoxicity [28].

Small interfering RNA (siRNA) is able

to silence specific gene expression via RNA

interference (RNAi) and has generated a great

deal of interest in both basic and applied biology

[136J. Although the viral-based siRNA delivery

method has shown promise in animal models

as well as clinical trials, the safety concern of

viral vectors is significant. It is thus important to

develop nonviral vectors for siRNA delivery [137,

138]. With a cleavable disulfide bond linkage

between siRNA and SWNTs, we successfully

delivered siRNA into cells by nanotubes and

observed a gene silencing effect (Fig. ll(a))

[35]. We further showed that our SWNT-based

siRNA delivery was applicable to those hard

to-transfect human T cells and primary cells,

which were resistant to delivery by conventional

cationic liposome-based transfection agents (Figs.

11 (b) and l1(c» [51]. Surface functionalization

dependent cell uptake of SWNTs was observed.

Compared with SWNTs coated with long PEG

(5.4 kDa) groups, shorter PEG (2 kDa) coated

SWNTs with more exposed hydrophobic surface

showed higher cellular uptake, which was

ailll lilli,

104

(due to incomplete coverage of nanotube sidewalls),

which can cause binding and association with

cells, resulting from hydrophobic interactions with

hydrophobic cell membrane domains. Cell binding

of SWNTs is an important first step for cellular entry

via endocytosis. Our results suggest that balanced

chemical functionalization schemes that impart

sufficient aqueous solubility and biocompatibility

to nanotubes, and also retain the ability of the

nanotube to bind with cell surfaces are important for

intracellular delivery of biomacromolecules by CNTs.

Beside our work, other CNT-based siRNA delivery

has also been reported, showing efficacy In vitro andeven in vivo [139].

5. In vivo biodistribution and long termfate of carbon nanotubes

Encouraged by the success in using CNTs for In vitro

sensing, drug delivery, and imaging, research in these

fields has moved to in vivo animal research. The first

critical questions to address are the biodistribution

profile of CNTs after systemic administration

into animals and whether there is any toxicity.

In the past few years, in vivo biodistribution and

pharmacokinetic studies have been carried out by

a number of groups using different CNT materials,

different surface functionalizations, and different

tracking methodologies, thus obtaining varying and

sometimes controversial results.

Radiolabeled C"In-DTPA) SWNTs and MWNTs

functionalized by 1,3-dipolar cycloaddition were

used by Singh et al. [46] and Lacerda et al. [47]

to determine biodistribution. Surprisingly, after

intravenous injection of CNTs into mice, they

observed fast urinal clearance of CNTs, with the

majority (>95%) cleared out within 3 h, and no

uptake in RES organs such as the liver and spleen.

Those phenomena were similar to the in vivo behavior

of small molecules, but drastically differed from that

expected of most nanoparticles with sizes exceeding

the glomerular filtration threshold. To explain their

results, the researchers proposed that despite their

lengths, the small diameters of CNTs allowed for fast

urinal excretion of CNTs. However, this conclusionis debatable considering well-characterized size-

Nano Res (2009) 2: 85-120

dependent protein biodistribution and excretion

behavior (as shown in Table 1, proteins larger than

7-9 nm start showing high RES uptake and limited

renal excretion) and also contradictory findings

in a study reported by Choi et al. using quantum

dots (QDs) [140]. It is found that the maximum size

allowing fast urinal excretion of spherical QDs is

~6 nm, including coating molecules [140], which is

indeed larger than the diameter of individual SWNTs

(1-2 nm), However, the QDs were much smaller

than the diameter of SWNT bundles (10-40 nm) [46]

or MWNTs (20-30 nm) [47] used in these two CNT

biodistribution studies. The reported fast CNT urinal

excretion requires confirmation.

Several other labs have also studied the bio

distribution of radiolabeled CNTs in mice. Wang

et aJ. reported relatively slow urinal excretion

and low RES uptake in their first study [141].

Later reports by the same group using 14C-taurine

functionalized CNTs, however, revealed dominant

and persistent liver accumulation of CNTs after

intravenous injection [49,142]. Another independent

study by McDevitt et al. using antibody conjugated

radiolabeled CNTs functionalized by 1,3-dipolar

cycloaddition also showed high CNT uptake in the

liver and spleen with slow urinal excretion [129]. A

significant amount of CNTs remained in the body

even after 15 days. We have also investigated the

biodistribution of radiolabeled, PEGylated SWNTs,

observing dominant SWNT uptake in RES organs,

including liver and spleen, without rapid clearance

[44]. Although the radiolabel method is a convenient

way to examine the biodistribution of a substance,

excess free radioisotopes in the radiolabeled CNT

samples, if not completely removed, may lead

to false results-especially for excretion-as free

radioisotopes are small molecules that are rapidly

excreted through urine after intravenous injection.

Also, radiolabels could be gradually released from

CNTs in vivo, and be slowly excreted in the free

form. As a consequence, radiolabeling is not an ideal

strategy to study the excretion and long term fate of

CNTs.

Without using radiolabels, researchers have

investigated the in vivo behavior of SWNTs relying on

their intrinsic properties. Individual semiconducting

Nano Research

Nano Res (2009) 2: 85-120106

60

50

-o 400:§.5 30

~;f!. 20

10

0

0 5 10

80 -r------------,

-- SWNT-/-2kPEG

~ SWNT-/-5kPEG

""7r""- SWNT-/-7kPEG-- SWNT-f-12kPEG-- SWNT-br-7kPEG

20 25

:§ 60o~! 40

.5

!z 20

~

o

-- SWNT-/-2kPEG

-- SWNT-f-5kPEG

SWNT-br-7kPEG

50Time (Day)

Cb)

100

__ SWNT-f-2kPEG

-- SWNT-/-5kPEG

SWNT-br-7kPEG

50Time (Day)

(c)

100

Cd)

Figure 12 In vivo pharmacokinetics and long-term biodistribution of SWNTs with different PEG coatings (Fig. 2(c» measured by Ramanspectroscopy. (a) Blood circulation curves of different PEGylated SWNTs after intravenous injection to mice. Long and branched PEG coating onSWNTs prolongs blood circulation half-lives of nanotubes. SWNT levels in liver (b) and spleen (c)over time. SWNTs areslowly excreted from RESorgans in a few months, with faster clearance rate for those with good PEG coating. Notethat SWNTs have much lower uptakes in other organs(less then 2% 10/g). (d) Raman images of mouse liver slices three months after SWNT injection. Much lower SWNT residues are observed in theliver of mice injected with nanotubes with more hydrophilic surface coating. Copyright 2008 the National Academy of Sciences {45]

intravenously injected into glioblastoma U87MG

tumor-bearing mice, which were monitored by

micro-positron emission tomography (micro-PET)

over time (Fig. 13). RGD-conjugated SWNTs with along PEG coating (SWNT-PEG54oo-RGD) exhibited

a high tumor uptake of -13% of injected dose per

gram tissue (%ID / g), compared with 4%-5% ID/ g

obtained with plain SWNTs without RGD (SWNT

PEG54(0) . Interestingly, we found that efficient tumor

targeting could only be realized when SWNTs were

Nano Research

Nano Res (2009) 2: 85-120 105

Table 1 Biodistributionand pharmacokinetics of proteinswith different sizes. Small proteinswith diameter below 6~7 nm are quickly excretedthrough urinal clearance with very short blood circulation and whole body retention half-lives. Biggerproteins in contrast show high uptake inRES organsand much longer blood and whole body half-lives. Theyare unableto undergo fast urinal excretion because of their largesizes

Biodistribution

Protein molecule MW(kOa) d(nm) (%(lO/g)at 4 h) Bloodhalf-life Whole body(min) half-life (h)

Liver Spleen Kidney

SeFv [1431 30 5.3 [140] 1.3 0.9 0.8 <10 -3.8

Fab' [1431 50 6.0 [144] 1.3 1.8 21.5 -30 -6.7

Se(Fv), [143,1451 60 7.0 [1401 2.0 2.8 2.9 78 -10

F(ab'), [1431 100 7.8 7.1 9.8 -200 -20

[Se(Fv),], [145] 120 9.3 [140] 7.0 6.2 3.6 170

IgG [143, 1451 152 11 [144] 18.7 18.0 4.7 330 -80

SWNTs exhibit NIR photoluminescence, which hasbeen used by Cherukuri et al. to track nanotubesin rabbits [50]. Without obtaining detailed biodistribution data, the authors observed SWNT

photoluminescence signals in the liver, but not inother organs such as the kidney. In another study,Yang et al. used isotope ratio mass spectroscopyto examine the biodistribution of 13C enriched

unfunctionalized SWNTs over a month, showing highnanotube uptake in lung, liver, and spleen withoutapparent excretion within 28 days [48]. Taking

advantage of the bright Raman scattering signaturesof SWNTs, we used Raman spectroscopy to studythe long-term fate of nanotubes in mice [45]. It wasfound that our PEGylated biocompatible SWNTs were

predominantly accumulated in the liver and spleenafter intravenous administration, but slowly excretedwithin months, probably via the biliary pathway intothe feces. There could be a small portion of SWNTswith very short lengths that were excreted throughurinal elimination, as indicated by the weak SWNT

Raman signal observed in the mouse kidney andbladder.

Most importantly, we have systematically studiedthe relationship between PEG coating and the in vivobehavior of SWNTs. Prolonged blood circulationtime is generally desired for drug delivery andtumor targeting. We have concluded that long PEGcoatings on SWNTs generally confer a prolongedblood circulation half-life, reduced RES uptake andaccelerated excretion (Fig. 12). Moreover, PEGs

having a branched structure offer a more efficientcoating on an SWNT surface than linear PEGs, andallow a longer SWNT blood circulation half-life,namely, -5 h for branched PEG compared with -2

h for linear PEG of the same molecular weight (7kDa) (Fig. 12) [45]. Another recent study by Yang et

al. obtained an even longer blood circulation halflife (22 h) using covalently PEGylated SWNTs [118].

These results are thus important guides to futureresearch into the use of carbon nanotubes for in vivobiomedical applications.

6. In vivo tumor targeting and preliminaryefforts towards in vivo cancer therapy

In order to use CNTs for potential cancer treatment

and / or imaging, targeting nanotubes to tumors ishighly desirable. Both passive targeting, relying onthe enhanced permeability and retention (EPR) effectof cancerous tumors, and active targeting guided bytumor targeting ligands, have been employed forvarious nanoparticle-based drug delivery systems.Thus far, there are two published papers reporting

in vivo tumor targeting by CNTs conjugated withtargeting ligands. We showed that efficient tumortargeting was achieved by conjugating a RGD peptide

-which recognizes integrin a"[33' known to be upregulated on various solid tumor cells and tumorvasculatures-to PEGylated SWNTs [44]. SWNTswith two different PEG coatings conjugated withboth RGD peptide and radiolabels ("Cu-DOTA) were

2 .an. l~' f":\ S .\~~ ~ pnnger

Nano Res (2009)2: 85--120

PEG2000-RGD

SWNT-PEGzOQO

-RGD

PEG5400-OOTA-64Cu ~

0-Tyr (y)y NH,

RGO = '-..Pi Lys (K)PEG54oo-RGD OyNH HN.

HOOC-/. 0 Arg (R)

Asp (0) o-e~~~ NH01('

Gly (G) NHSWNT -PEG"oo-RGO

Ca)

107

0.5 h

6h

24 h

SWNT-PEG"oo-RGO

U87MG

SWNT -PEG"oo-RGO

U87MG

SWNT-PEG"",-RGOw/RGOblocking

U87MG

Cb)

SWNT-PEG"",-RGO

HT-29

Figure 13 In vivo tumor targeting with 5WNTs. (a)Scheme of PEGylated SWNTs with RGD conjugation and radiolabeling. (b) Micro-PET imagesof mice. Arrows point to the tumors. High tumor uptake (~13 %ID/g) of SWNT-PEGS400-RGD is observed in the U87MG tumor (2nd column),

in contrast to the low tumor uptake (t'' column) of SWNT-PEG1ooo-RGD. The 3'd column is a control experiment showing blocking of SWNT

PEG S400-RGD tumor uptake by co-injection of free c(RGOyK). The4th column isa control experiment showing low uptake of SWNT-PEGS400-RGD

in integrin av~3-negative HT-29 tumor. Copyright 2007 Nature Publishing Group [44]

coated with long PEG (SWNT-PEG54oo-RGD) but not

with short PEG (SWNT-PEG,ooo-RGD). The latterhad short blood circulation time, and thus lowerprobability of being trapped in tumors or to bindthe tumor receptors. Our data suggest that surfacefunctionalization of SWNTs is also important for

tumor targeting in vivo. Another study carried outby McDevitt et al. [129] showed tumor targeting of

CNTs by antibody conjugation.The first in vivo cancer treatment study with CNTs

was reported by Zhang et al. using positively chargedSWNTs to delivery therapeutic siRNA into cancercells [139]. However, this was a proof-of-conceptstudy, with SWNT-siRNA complexes directly injectedinto tumors, instead of systemic administration.

Our recent work showed that paclitaxel (PTX),

;z m ~ Springer

108 Nano Res (2009) 2: 85-120

****** ***

** **

Individual semiconducting SWNTs have small

band gaps on the order of -1 eV, depending on the

7.1 In vitro photoluminescence imaging

*

--SWNT-PTX

-PlainSWNT

---Taxol®

-+- Untreated

---PEG-PTX•.",--DSPE-PEG-PTX

2

4

6

10

12.,.------------------;---,

0-lr---,.......,~---r-_r_-_._-_,~.,_----1

7. Biological imaging usingcarbon nanotubes

a commonly used chemotherapy drug, may be

conjugated to branched PEG functionalized SWNTs

via a cleavable ester bond (Fig. 14(a)) [37]. The SWNT

PTX conjugate was tested in a 4Tl murine breast

cancer model in mice, exhibiting

improved treatment efficacy over

the clinical Cremophor-based

PTX formulation, Taxol® (Fig.

14(b». Pharmacokinetics and

biodistribution studies revealed

longer blood circulation half

life and higher tumor uptake of

SWNT-PTX than those of simple

PEGylated PTX and Taxol®,

consistent with the observed

efficacies of different PTX

formulations. The high passive

tumor uptake of SWNT-PTX is

likely due to the EPR effect. In

addition, PTX molecules carried to

RES organs (e.g., liver and spleen)

by SWNTs were rapidly dissociated

from nanotubes and excreted,

diminishing the RES toxicity of this

SWNT-based PTX formulation.

Our work is the first to show that

carbon nanotubes can be used for

in vivo drug delivery for cancer

therapy by systemic administration

[37].

Figure 14 In vivo drug delivery with carbon nanotubes for cancer treatment. (a) Schematic

illustration of paclitaxel conjugation to SWNT functionalized by phospholipids with branchedPEG chains. The PTX molecules are reacted with succinic anhydride (at the circled OH site) to

form cleavable ester bonds and linked to the termini of branched PEG, via amide bonds. This

allows for release of PTX from nanotubes by ester cleavage in vivo. (b) Tumor growth curves

of 4T1 tumor-bearing mice receiving the different treatments indicated. The same PTX dose

(5 mg/kg) was injected (on days 0, 6, 12, and 18, marked by arrows) for Taxol®, PEG-PTX,

DSEr-PEG-PTX and SWNT-PTX. p Values (Taxol® vs SWNT-PTX): * p<0.05, ** p<0.01, ***p<O.001. Inset A photo of representative tumors taken out of an untreated mouse, a Taxol®

treated mouse and an SWNT-PTX treated mouse at the end of the treatments. Copyright 2008

American Association for Cancer Research [37]

In addition to applications for

drug delivery and treatment,

the intrinsic optical properties

of SWNTs make them useful

as optical probes. Owing to

their quasi I-D nature, SWNTs

exhibit strong resonance Raman

scattering, high optical absorption

and photoluminescence in the

NIR range, all of which have been

utilized for imaging in biological

systems In vitro and in vivo.

10

Time (day)

(b)

15 20 25

N ano Research ----,.. ----------------

Nano Res (2009) 2: 85-120 109

1000

900

800

700

600

500

400

300

200

100

o

'No emission'

(e)

(a)

147]. This seems to contradict studies on single

SWNTs with much higher quantum yield estimates

of up to 0.07 [148]. Many factors contribute to the

quantum yield including exciton quenching by

bundles [149], sidewall defects [150], and length

[151]. Crochet et al. showed that the quantum yield

of bulk SWNT suspensions could be increased

by an order of magnitude by centrifugation in an

iodixanol gradient to remove bundled nanotubes

[152]. Careful preparation techniques, such as these,

need to be applied in making suspensions of pristine,

unbundled SWNTs in bio-inert coatings, such as

PL-PEG, in order to fully realize their potential as

(b)

Emission

900-1600 om

~

Figure 15 SWNT as NIR fluorescent labels. (a) Schematic of targeting cells with 5WNT

Herceptin conjugates to 8T-474 (HER2/neu positive) and MCF-7 (HER2/neu negative) cells. (b)NIR photoluminescence image of BT-474 cells treated with the 5WNT-herceptin conjugate. Thehigh level of fluorescence signal indicates surface binding of the SWNT-Herceptin conjugate.(c) NIR photoluminescence image of MCF-7 cells. Very little NIR signal is seen owing to thelow non-specific binding of the SWNTHHerceptin conjugate and low cellular autofluorescence.NJR photoluminescence images were taken using a homebuilt scanning confocal microscopewith InGaAs detector, Photoluminescence was collected from 900--1600 nm. Copyright 2008American Chemical Society [24]

diameter and chirality of a given nanotube. This

band gap allows for photoluminescence in the NIR

range (900-1600 nm) which is useful for biological

imaging, due to the high optical transparency of

biological tissue near 800-1000 nm and the inherently

low autofluorescence from tissue in the NIR range

[146]. SWNTs have a further advantage due to the

large separation between the excitation (550-850 nm)

and emission bands (900-1600 nm). This spacing

further reduces background from autofluorescence

and Raman scattering.

O'Connell et al. [23] first demonstrated NIR

photoluminescence from micelle-encapsulated

SWNTs, yielding an estimatedquantum efficiency of about 10-'.

Relying on this intrinsic NIR

photoluminescence, Cherukuri et

al. imaged nonspecific uptake of

SWNTs in phagocytic cells [17]. In

more recent work, [in et al. usedNIR photoluminescence to track

endocytosis and exocytosis of

SWNTs in NIH-3T3 cells in real

time [95]. In our lab, we developed

bio-inert PEGylated SWNTs

conjugated with antibodies as

NIR fluorescent tags for selective

probing of cell surface receptors

[24]. Figure 15 shows NIR fluore

scence images of (b) BT-474,

which is HER2/ neu positive, and(c) MCF-7, which is HER2/ neu

negative, treated with an SWNT"

herceptin conjugate. We observed

ultralow NIR autofluorescence

between different cell lines,

demonstrating the advantage of

SWNTs as an NIR fluorophore,There is still work to be done

in the application of SWNTs

as effective fluorophores. Bio

inert samples must be made

with optimized quantum yields.

Initial estimates of the quantum

yield of SWNT suspensions wereon the order of 10-3 or less [23,

r 3E ~ I"t; ~~ ~~ " .\ ..... J _ Springer

110

sensitive NIR fluorophores.

7.2 In vitro Raman imaging

Owing to their quasi 1-D nature, SWNTs exhibit

strong resonance Raman scattering due to their

sharp electronic density of states at the van Hove

singularities. SWNTs have several distinctive Raman

scattering features including the radial breathing

mode (REM) and tangential mode (G-band) [25],

which are sharp and strong peaks that can be easily

distinguished from fluorescence backgrounds, and

are thus suitable for optical imaging. We and Heller

et al. have used Raman microscopy to image SWNTs

in liver cells, as well as tissue slices, using either the

REM peak or G-band peak of SWNTs [26, 45, 51, 58].

Our latest work showed that SWNTs with different

isotope compositions exhibited shifted G band peaks,

Nano Res (2009) 2: 85-120

and thus could be used as multicolor contrast agents

for multiplexed Raman imaging [127]. Cancer cells

with different receptor expression profiles were

selectively labeled with three isotopically unique

formulations of "colored" SWNTs, conjugated

with various targeting ligands including Herceptin

(anti-Her2), Erbitux (anti-Her'l ), and RGD peptide,

allowing for multicolor confocal Raman imaging

of cells in a multiplexed manner using a singleexcitation (Figs. 16(c) and 16(d)). Raman signals of

SWNTs are highly robust against photobleaching,

allowing long term imaging and tracking [26, 45].

With narrow peak features, SWNT Raman signals

are easily differentiated from the auto-fluorescence

background. The SWNT Raman excitation and

scattering photons are in the NIR region, which is

the most transparent optical window for biological

(b)

15000

12000

~8 9000

c~ 6000

i2.3000

~-_. 12C-SWNT-anti~her2

-- "C/"C-SWNT-RGD

-- 13C-SWNT-anti-her1

1500 1550 1606

Raman shift (em-')

(b) (e)

Figure 16 Multi-color Raman imaging with isotopically modified SWNTs. (a) Schematic illustration of SWNTs with three different isotopecompositions (13C-SWNT, llC12rC-SWNT, 12C-5WNT) conjugated with different targeting ligands. (b) Solution phase Raman spectra of the

three SWNT conjugates under 785 nm laser excitation. Different G-band peak positions were observed. (c) A deconvoluted confocal Raman

spectroscopy image of a mixture of three cell lines with different receptor expressions incubated with the three-color SWNT mixture. Scalebar =

100 urn. Copyright 2008 American Chemical Society [127J

Nano Research__._III1!l1fK 1IIII

Nano Res (2009) 2: 85-120

systems In vitro and in vivo. Thus, SWNTs are novel

Raman tags which show promise for multiplexed

biological detection and imaging.

7.3 SWNTs for in vivo animal imaging

SWNT-based biomedical imaging has also been

conducted in animal models. The first ever imaging

of nanotubes inside a living animal was achieved

by the Weisman group in 2007 [94]. In this work,

Drosophila larvae were fed by food containing

SWNTs and imaged by NIR fluorescence microscopy.The biodistribution of SWNTs in live larvae was

monitored by the nanotube fluorescence signals.

Recently, the Gambhir group successfully used RGD

conjugated PEGylated SWNTs provided by us as

Raman probes for in vivo tumor imaging in live mice

(Fig. 17) [153, 154]. Intravenous injection of targeted

SWNTs to living mice bearing a tumor xenograft,

showed strong SWNT Raman signals in the tumor,

while little signal was observed in the tumor after

injection of non-targeted SWNTs. This is the first

success of in vivo tumor imaging via carbon nanotube

labels.

SWNTs have strong optical absorption in the

visible and NIR range. We and Chakravarty et

111

al. have shown that SWNTs can be utilized as

photothermal therapeutic agents to kill cancer

cells [20, 128]. NIR laser irradiation was used in

both cases to generate heat, causing destruction of

cancer cells with specific SWNT internalization.

Beside its potential applications in therapy, the high

optical absorption of SWNTs can also be utilized

in photoacoustic imaging. Photoacoustic imaging,

in which sounds are generated as a result of local

heating by the absorption of laser light, has higher

spatial resolution than traditional ultrasound, and

deeper tissue penetration than fluorescence imaging[155]. de la Zerda et al. used our RGD-conjugated

SWNTs as the contrast agent for photoacoustic

molecular imaging of cancer in a mouse tumor model

(Fig. 17) [22]. This work opens up new opportunities

for in vivo biological imaging with SWNTs.

8. Summary and perspective

In this article, we have comprehensively reviewed

the current research regarding the use of carbon

nanotubes for biomedical applications. Various

covalent and noncovalent chemistries have been

developed to functionalize CNTs for biomedical

Plain SWNT

SWNT-RGD

Photoacoustic

invivo imagingRaman

invivo imaging

Figure 17 In vivo tumor imaging with 5WNTs. In vivo photoacoustic (strong lightabsorption of SWNTs generates anacoustic signal) and Raman images of U87MG tumors in live mice injected with plain'SWNT or RGD-conjugated SWNT.Strong tumor contrast induced by 5WNT-RGD was observed in both imaging techniques. Copyright 2008 AmericanChemical Society 11 54!. 2008 Nature Publishing Group[22!. and 2008 the National Academy of Sciences [1531

'7 Z= (~1 ~ Springer

112

research. Relying on their electric or optical

properties, functionalized CNTs have been used

for ultrasensitive detection of biological species.

Surveying the literatures, we clarify that In vitro and

in vivo toxicities of CNTs are highly dependent on

CNT functionalization. Appropriately functionalized

CNTs with biocompatible coatings are stable in

biological solutions, and nontoxic In vitro to cells

and in vivo to mice at the tested doses. Various

reports have shown that CNTs are able to shuttle

biological molecules including small drug molecules

and biomacromolecules including proteins,

plasmid DNA, and siRNA into cells In vitro via an

endocytosis pathway. In vivo behaviors, including

blood circulation, biodistribution, and long term fate

of CNTs, have been studied in the past two years,

showing dominant uptake of CNTs in RES organs,

similar to most nanomaterials tested in vivo. CNTs

are able to target tumors by both passive targeting

relying on the EPR effect and active targeting

guided by targeting ligands, showing promise for

in vivo cancer treatment. Moreover, SWNTs exhibit

unparalleled intrinsic optical properties and have

been used for biological imaging In vitro and in vivo.

Surface functionalization chemistry is the rr:ost

essential and fundamental factor as far as biomedical

applications of CNTs are concerned. Highly

hydrophilic coatings, such as long and branched

PEG on CNTs, impart "inertness" in biological

environments, minimizing the toxicity of CNTs and

reducing NSB to biological species, such as serum

proteins and cell surface proteins. Minimal NSB is

critical since it leads to enhanced detection sensitivity

in CNT-based biosenors and imaging probes.

Decreased NSB reduces the non-specific endocytosis

of CNTs, which favors targeted drug delivery to

specific cell types. Similarly, the in vivo behaviors of

CNTs are highly dependent on their surface coating.

Prolonged blood circulation and reduced RES

uptake can be achieved by using CNTs with highly

hydrophilic coatings. Further efforts are required to

optimize the surface chemistry of SWNTs, to further

enhance biocompatibility. With improved surface

coating, further improvements in biological sensing

and imaging, better tumor targeting attributed

to prolonged blood circulation and reduced RES

Nano Res (2009)2: 85-120

uptake, and accelerated excretion may be realized.

By minimizing the non-specific protein binding of

nanotubes via improved surface functionaIization,complement activation may also be reduced for

SWNTs.

CNTs, especially SWNTs are highly promisingin biomedicine for several reasons. CNTs are

composed purely of carbon, while many inorganic

nanomaterials (e.g., quantum dots) contain more

hazardous elements, such as heavy metals. The

distinctive 1-D structure and tunable length of CNTs

provide an ideal platform to investigate size and

shape effects in vivo. Lastly, unlike conventional

organic drug carriers, the intrinsic physical

properties of SWNTs including resonance Raman

scattering, photoluminescence, and strong NIR

optical absorption can provide valuable means of

tracking, detecting, and imaging. Taken together,

CNTs may serve as a unique platform for potential

multimodality cancer therapy and imaging.

Although numerous encouraging results using

CNTs in biomedicine have been published in the

past several years, much more work is still needed

before CNTs can enter the clinic. The most important

issue to be addressed is still the concern of long

term toxicity. Although we have shown that well

functionalized carbon nanotubes are not toxic In vitro

to cells and in vivo to mice at our tested doses, further

systematic investigations using different animal

models on larger scales with various doses are

required. Special attention should be paid to CNTs

with surface functionalization optimized for such

applications, with greater chances of minimizing

toxic side effects.

Although we have succeeded in using carbon

nanotubes for In vitro siRNA delivery, in vivo delivery

of this type of biological macromolecules remains a

challenge. A paradox exists because of the need for

high In vitro cellular uptake and the requirement

of favorable in vivo behavior such as long blood

circulation and low RES uptake of SWNTs. The

former suggests reduced PEGylation of nanotubes,

while the latter requires dense biocompatible

surface coatings. Conjugation of targeting ligands

on appropriately coated SWNTs may help to solve

this problem, allowing enhanced cellular uptake

Nano Research IliILllillillillillilli;_II.i!aIl·;,II·IIIIIIIIIIIIIIIIII••••1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII1IIIII11111

Nano Res (2009) 2: 85-120

via receptor mediated endocytosis, without loss

of optimal SWNT in vivo characteristics. Further

development of suitable bioconjugation chemistry

on nanotubes may create versatile SWNT-based

bioconjugates for actively targeted in vivo drug andgene delivery.

Although not specifically discussed in this review,

the fabrication of CNTs may also play an important

role in their future biomedical applications. Most of

the CNT samples used in the published biomedical

studies are heterogeneous mixtures of nanotubes

with different lengths, diameters, and chiralities. The

lengths of CNTs may affect In vitro cellular uptake as

well as in vivo pharmacokinetics of nanotubes. It is

thus important to obtain and test nanotube samples

with narrow length distributions. A density column

based length separation method has been developed

in our group [156]. Further studies will uncover

potential length dependent effects on the behavior of

CNTs in biological systems. The electronic structures

of SWNTs are determined by their chiralities (Fig. 1).

Achieving SWNTs with single diameter and chirality

may bring a revolution to the semiconductor industry

and has been one of the ultimate goals of nanotube

research for a decade. Progress has been made by

selective synthesis of SWNTs under special conditions

[157], removal of SWNTs with undesired diameters

by etching [102], and chirality separation based on

chromatography [73, 74]. Different semiconducting

SWNTs with single chirality compositions can serve

as different colors in NIR photoluminescence imaging

(Fig. 1). One the other hand, Raman scattering of

SWNTs with single chirality will be largely enhanced

because all nanotubes can be in resonance with

a selected excitation wavelength. The diameter

dependent Raman RBM band of SWNTs may also be

utilized in multi-color Raman imaging [25, 158]. CNT

samples with homogenous length, diameter, and

chirality distributions are the ideal candidates for

further studies in biomedicine.

Lastly, biomedical imaging, based on the inherent

physical properties of SWNTs, may be combined

with drug delivery for multimodality cancer

diagnosis and therapy. Phototherapy relying on the

strong NIR light absorption ability of SWNTs may be

conducted simultaneously, along with chemotherapy

113

delivered by SWNTs, to enhance treatment efficacy

in vivo. Despite challenges on the way towards the

clinic, carbon nanotubes exhibit great potential

for biomedicine, and may bring unprecedented

opportunities for the future of cancer diagnosis and

therapy.

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Nano Research :

Attachment Ic

Liangbing Hu··', Jang Wook Choi··'. Yuan Yang·''. Sangmoo Jeongb, Fabio La Mantia·, Li-Feng Cui·, and Yi Cui··2

Departments of »Materlals Science and Engineering and bElectrical Engineering, Stanford University, Stanford, CA 94305

Edited by Charles M. Ueber. Harvard University, Cambridge, MAt and approved October 21, 2009 (received for review August 6. 2009)

Author contributions: L.H., 1.W.C.,Y.Y., and Y.c. designed research; L.H., l.W.C.,Y.Y., 5.1.,F.L.M., and L.-F.C. performed research; L.H., 1.W.C., Y.Y., and Y.c. analyzed data; and L.H.,1.W.C., Y.Y., and Y.c. wrote the paper.

The authors declare no conflict of interest.

Thisarticle is a PNAS Direct Submission.

lL.H.,1.W.C., and Y.Y. contributed equally to this work.

2Towhom correspondence should be addressed. E-mail:[email protected].

This article contains supporting information online at www.pnas.orglcgifcontentlfullf0908858106/DCSupplemental.

hierarchical porous fiber structures, surface charges, and functional groups in paper, which are under fine control today (9).Paper has continued to expand its applications beyond information recording, and recently, novel applications such as microfluidic and electronic devices have been demonstrated. Whitesidesand colleagues fabricated 3D microfluidic devices by stackingpaper and adhesive tape (10) and, in a separate study, portablebioassays on patterned paper substrates (11). Researchers in theflexible electronics community have also explored paper assubstrates for organic photodiodes (12), organic thin-film transistors (13, 14), circuits (15), and active matrix displays (16). Inthis study, we demonstrated that the application of paper can beexpanded even further to important energy-storage devices byintegrating with single-walled eNTs and metal nanowires bysolution-based processes. The coated ID nanomaterial filmsshow high conductivity, high porosity, and robust chemical andmechanical stability, which lead to high-performance supercapacitors (SCS) and lithium-ion (Li-ion) batteries.

Results and DiscussionAqueous CNT ink with sodium dodecylbenzenesulfonate(SDBS) as a surfactant was used in this study (17), where SDBSand CNT were 10 and 1-5 mg/mL in concentration, respectively.Once CNT ink was applied onto paper by the simple Meyer rodcoating method (Fig. 1A), the paper was transformed into highlyconductive paper with a low sheet resistance around 10 fl/sq(Fig. 1B), which is lower than previous reports by several ordersof magnitude because of the ink formulation and the choice ofsubstrates (18, 19). Fig. 1 C and D shows the conformal coatingof CNTs on the fiber structure of the paper, which contributesto high film conductivity (see Figs. SI and S2 for more details).One important reason for this conformal coating might be theporous structure of paper, which leads to large capillary force forthe ink. The strong capillary force enables high contactingsurface area between flexible nanotubes and paper after thesolvent is absorbed and dried out. We also applied the samemethod to produce conductive paper based on ink of othernanoscale materials, by using Ag NWs as an example (Fig. IEand Table 51; see Materials and Methods for detailed procedures). The sheet resistances at different effective film thicknesses for CNTs and Ag NWs are plotted in Fig. IF. Benefittingfrom the conformal coating, the sheet resistances reached a lowlevel of 1 flIsq for Ag NWs at the effective film thickness of 500nm. As film thickness increased, the scaling of the resistancechanged from percolation-like to linear behavior, which is similarto CNT networks on flat substrates. The cross-over from percolation to linear region was found to be =20-30 nm on other flatsubstrates, which is close to our value, = 10 nm (Fig. IF), and thedifference is likely due to the length differences of CNTs (20, 21).

conformal coating! carbon nanotubes I nanomater!al I solution process

Printable solution processing has been exploited to depositvarious nanomaterials, such as fullerene, carbon nanotubes

(CNTs), nanocrystals, and nanowires for large-scale applications, including thin-film transistors (1-3), solar cells (4, 5), andenergy-storage devices (6, 7), because the process is low-costwhile maintaining the unique properties of the nanomaterials. Inthese processes, flat substrates, such as glass, metallic films, Siwafers, and plastics, have been used to hold nanostructure films.Nanostructured materials are usually first capped with surfactant molecules so that they can be well-dispersed as separatedparticles in a solvent to form "ink." The ink is then depositedonto the flat substrates, followed by surfactant removal andsolvent evaporation. To produce high-quality films, significantefforts have been spent on ink formulation and rheology adjustment. Moreover, because the surfactants are normally insulating, and thus limit the charge transfer between the nanomaterials, their removal is particularly critical. However, this stepinvolves extensive washing and chemical displacement, whichoften cause mechanical detachment of the film from the flatsubstrate. Polymer binders or adhesives have been used toimprove the binding of nanomaterials to substrates, but these canalso cause an undesirable decrease in the film conductivity.These additional procedures increase the complexity of solutionprocessing and result in high cost and low throughput. Here, weexploit paper substrates used in daily life to solve these issues anddevelop a simple, low-cost, high-throughput, and printable process for achieving superior device performance.

Throughout human history, paper has been the most important medium to express and propagate information and knowledge. The Prisse Papyrus, the oldest existing piece of writing onpaper, dates back to 2,000 n.c. (8). The critical feature thatenables paper to record information in such an enduring manneris the strong adhesion of ink onto paper, resulting from the 3D

Highly conductive paper for energy-storage devices

Paper. invented more than 2,000 years 8g0 and widely used todayin our everyday lives. is explored in this study as a platform forenergy-storage devices by integration with 10 nanomaterials.Here. we show that commercially available paper can be madehighly conductive with a sheet resistance as low as,1 ohm persquare (!l/sq) by using simple solution processes to achieve conformal coating of single-walled carbon nanotube (CNn and silvernanowlre films. Compared with plastics. paper substrates candramatically improve film adhesion. greatly simplify the coatingprocess, and significantly lower the cost. Supercapacitors based on(NT-conductive paper show excellent performance. When onlyCNT mass is considered. a specific capacitance of 200 F/g, a specificenergy of 30-47 Watt-hour/kilogram (Wh/kg), a specific power of200,000 W/kg. and a stable cycling life over 40.000 cycles areachieved. These values are much better than those of devices onother flat substrates. such as plastics. Even in a case in which theweight of all of the dead components is considered. a specificenergy of 7.5 Wh/kg is achieved. In addition, this conductive papercan be used as an excellent lightweight current collector in Iithiumion batteries to replace the existing metallic counterparts. Thiswork suggests that our conductive paper can be a highly scalableand low-cost solution for high-performance energy storagedevices.

www.pnas.org/cgijdoijl0.1073/pnas.0908858106 PNA5 Early Edition I 1 of 5

D••

Paper

_ Before Washing",,"wAfterWashing

E

4 6 6 10Bending Radius (mm)

PET

C1.0 •0.8

r£oc 0.6

0.4

0.2

0.02

Fig. 2. Various performance tests of conductive paper. (A) Sheet resistancechange of CNTs on paper and PETbefore and after the process of washing toremove surfactants. (8) The comparison of film peeling after washing surfactants. CNTfilms on PETare easily peeled off, whereas CNTs on paperstill stickwell. (Q Sheet resistance changes after bending conductive paper into different radii. (D) Film adhesion test with Scotch tape: CNTs on paper remainbound, whereas CNTs on PETare peeled off. (E) Direct writing of CNTink onthe paper with Chinese calligraphy.

necessary in the case of conductive paper, and thus its fabricationprocess becomes greatly simplified.

The conductive paper also has excellent mechanical properties.The conductive paper with eNT thicknesses from 100 run to 5 MIDcan be bent down to a 2-mm radins (Fig. 2C) or folded withont anymeasurable change in electrical conductivity. Fatigue tests showthat the conductive paper can be bent to a 2-mm radius for 100 timeswith resistance increase less than 5%. These mechanical behaviorsare likely dne to the combined effect of the flexibility of individnalCNTs, the strong binding of the CNTs with the paper fibers, and theporons morphology ofthe paper, which can relax the bending strain.Such flexibility could satisfy the requirement in SCS and batteries.In comparison, conductive paper with a 50~nm gold layer evaporated on Xerox paper showed R, of 7 fl/sq. The sheet resistanceincreased by 50% after folding the condnctive gold paper threetimes. Moreover, the strong adhesion of the CNTs to paper lead tohigh film stability against damage, such as scratching and peelingoff. This is demonstrated via a Scotch tape test showing the clearsuperiority of nsing paper over the PET snbstrate (Fig. W). TheScotch tape did not peel off any CNTs on paper, and the sheetresistance remained the same, =10 fl/sq. As in the conformalcoating, the exceptionally strong binding is attributed to the largecapillary force and maximized contact area and, subsequently, Vander Waals force between the CNTs and paper (23). The superiorflexibility and high stability make conductive paper promising forvarious rolled-up devices. It is noteworthy that the conductive paperdescribed here is completely distinct from previously reported bulkypapers (24) or other condnctive paper (25) in several perspectives.(i) We nsed widely available commercial paper, not paper-like filmsproduced by more complicated processes. Therefore, we could

,0"

'0·'1..,,"'0'""'~-''''O''''--,"'0"',--,"'0Effective Thickness (nm)

Because paper absorbs solvents easily and binds with CNTsstrongly, the fabrication process for the conductive paper is muchsimpler than that for other flat substrates, such as glass orplastics. First, contrary to other substrates, ink rheology forpaper is not strict at all. In glass and plastics, the ink surfaceenergy needs to match with that of substrates, and the viscositymust be high enough to avoid surface tension-driven defects,such as rings and dewetting in the coating and drying processes(22). Therefore, various additives are incorporated in the ink totune the rheology properties. These insulating additives decreasethe conductivity of the final film. In contrast, our CNT ink doesnot need any additives to adjust the rheology, which simplifiesthe process and leads to high film conductivity. Second, thepaper does not require surfactant washing processes to achievehigh film conductivity, which is necessary for other substrates. Asshown in Fig. 2 A and B, the sheet resistance of the CNT paperwas already as low as 30 fl/sq before washing, and there was nofilm delamination and sheet resistance change after washing. Incontrast, washing of CNTs on Polyethylene terephthalate (PET)substrates resulted in significant film cracking and peel-off (Fig.2B). The local resistance of CNT film on PET decreasedsignificantly (Fig. S3A); however, the global resistance became=1,000 times higher (Fig. 2A). Althongh the detailed mechanism behind this unusual phenomenon is unclear, it is likely thatwhile solvent is sucked into paper by capillary force, surfactantsbecome rearranged, perhaps toward porous fiber networks, suchthat surfactants do not hinder charge transport within eNT filmsas much as they do with flat substrates. This unusual property ofpaper renders the typical surfactant washing requirement un-

Fig. 1. Conformal coating of CNTs or Ag NWs on commercial paper. (A)Meyer rod coating of (NT or Ag NW ink on commercial Xerox paper. (8)Conductive Xerox paper after (NT coating with sheet resistance of =10 flIsq.SEM images of (Q surface morphology of Xerox paper, (D) conformal (NTcoating along fibers in Xerox paper, and (E) conformal Ag NW coating onXerox paper. (A Sheet resistances of conductive paper based on CNTs and AgNWswith various thicknesses.

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both aqueous and organic electrolytes by using galvanostatic (Fig.38) and cyc1icvoltammetric (Fig. S5) methods. Detailed proceduresfor the device preparation and performance characterization aredescribed in Materials and Methods and Sf Materials and 'Methods.As shown in Fig. 3C, the specific capacitances of all-paper SCS atvarious currents are superior to the previously reported values withpure CNT electrodes on flat substrates (6, 26-33), and are evenclose to those of pseudocapacitors based on the polymer/CNTcomposites (34, 35). A high specific capacitance of 200 Fig wasachieved for devices in sulfuric acid electrolyte (Fig. 3C). Furthermore, our devices can maintain excellent specific capacitance evenunder high-current operations. Even at =40 A/g, capacitanceslarger than 70 F/g were maintained in both aqueous and organicphases. Such high capacitances at large currents are attributed tothe excellent ion accessibility from both sides of the CNT fihn andintimate electrolyte-CNT wetting that originates from the porousnature of paper. For comparison, PET-based SCSprepared in thesame way only showed capacity less than 50 Fig, (Fig. 3C), whichindicates the importance of the porous nature of paper for betterSC performance. This is also confirmed by another control experiment with Au conductive paper. Devices with the same amount ofCNTs on Au-coated Xerox paper (50 nm, 7 U/sq) showed 36 Fig at10 Ng, which is 4 = 5-fold lower than our CNT conductive paperSCS(=160 Fig at the same current density). This is likely becausethe Au film blocks pores in paper and impedes the ion access fromthe paper side. When operated at 3 V in organic solvent, the specificenergy and power reached 47 Wh/kg and 200,000 W/kg, respectively,which exceed previously reported data (Fig. 3D) (6, 26, 27,30, 31). Our data in this plot are calculated based on CNT mass onlyto comparewith other data in the references that were also acquiredin the same way. Furthermore, even in a case when CNT massloading increased to 1.7 mgcm'', the device performance was stillsuperior to others reported (Fig. 3D). The superior capacitancesmay be a result of the higher ion accessibility in the paper due tothe strong absorption of solvent by the paper. The CNT film onpaper is also thinner, with the same mass loading as a result of thelarger surface area of the rough paper. Also, to compare withcommercial devices (36) and see the significance of mass saving byreplacing metal current collectors with conductive paper, we plotted another Ragone plot (Fig. 3E). In this plot, the mass of deadcomponents (electrolyte, paper, separator = 1.1,3.3, 1.6 mg/cm2,

respectively) was considered in addition to active materials(CNTs ~ 1.7 mg/cnr'), This Ragone plot shows that our data arebetter in both energy/power densities compared with commercialdevices as-well as reported data in the literature (33) that are alsobased on the mass of all of the device components.

Cycle life, one of the most critical parameters in the SC operations, turns out to be excellent for 40,000cycles (Fig. 3F).Only 3%and 0.6% capacitance losses were observed in sulfuric acid andorganic electrolyte, respectively. Moreover, the conductive paperwas mechanically robust and did not show any cracks or evidenceof breakage for 60 days. CNT films as thick as ~14 urn (1.33mgfcm2) were also tested in sulfuric acid, and capacitances as highas =122 Fig were achieved (Fig. S68). The devices with 1.33and 1.7mg/cm? CNTs were compared with the reported densest CNTassembly data (Fig. S6C) (28). Our devices show larger capacitancesper area compared with their films, although their fihns are 10 timesthicker. Furthermore, their devices also leave plenty of unusedspace, and therefore might not be scalable. More data on cyclicvoltammetry (CV; Fig. S5) and thickness dependence of thecapacitance (Fig. S6A) are presented in the Sf Materials andMethods.

Rechargeable batteries are another type of energy-storagedevice with high energy density, but they are still too heavy forapplications such as vehicle electrification. In this work, conductive paper was used to replace the heavy metallic currentcollectors, which could reduce the weight of batteries up to 20%(Table 82; see Sf Materials and Methods for details) with

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benefit from the well-established paper technology. (ii) Our conductive paper takes advances of intrinsic properties of paper, whichlargely simplifies tbe fabrication process. (iii) Our fabrication processis scalable, with roll-to-roll fashion. (iv) Other painting methodscan also be applied to fabricate conductive paper. Chinese calligraphy (Fig. 2E) and pen writing (Fig. S38) are demonstrated asexamples.

Because of the high conductivity and the large surface area, theconductive paper was studied in SC applications as active electrodesand current collectors. CNTs deposited on porous paper are moreaccessible to ions in the electrolyte than those on flat substrates (6,26-33), which can result in high power density. In addition, thepaper itself can function well as a separator. Therefore, all-paperscs have been realized by simple fabrication processes (Fig. 3A).We fabricated such all-paper SCS and tested their performance in

Fig. 3. Conductive paper as SC electrodes. (A) Schematic illustration ofall-paper SCs based on (NT conductive paper. Zoomed-in schematic illustratesthat ion accessibility is enhanced by the strong solvent absorption. (B) Galvanostatic charging/discharging curves taken from 3 V with organic electrolyte and 1 Vwith sulfuric acid. (Q Gravimetriccapacitances at various currentsmeasured in aqueous and organic electrolytes. Data from CNTs on PET areplotted together for comparison. (D) A Ragone plot showing that the allpaper SCs outperform the PET SC devicesand reported data in literature (6, 26,27, 30, 31). The color coding (black, red, green) of the traces isthe same asinC. Data in B-D aswell as data from references mentioned in 0 are calculatedbasedon a massloading ofCNTs only. Typical massCNTloadings in Cand 0 are72 JLg!cm2, and data from a large mass loading of 1.7 mg!cm2 are alsopresented (D, blue plot). (E) A Ragone plot based on the massof differentcombinations of dead components. A plot from ref. 33 based on the massofentire device components is also presented. The trapezoid showing the performance of commercial devices is from ref. 36. (F) Capacitance retentionmeasured in different electrolytes. After 40,000 cycles, 97% and 99.4% ofinitial capacitances are maintained for sulfuric acid and organic electrolytes,respectively.

Hu et al. PNAS EarlyEdition I 3 of 5

CycIO NurrlJor

Fig.5. Tests on the stability of paper in the organic electrolyte. The side withCNTs (A) and without CNTs (8) of the conductive paper soaked in organicelectrolyte (1 M UPF6 in EClDEC). From left to right: as-coated conductivepaper, paper soaked for 2 months at 30 QC, and paper soaked for 1 month at50 QC. (0 Atotal of500 cyclesof li4Tis012nanopowderswith conductive paperas the current collector. The mass loading of li 4Ti s0 12 is =2 mg/cm2.The totaltime is about 3.5 months.

component and device levels. After soaking the CNT conductivepaper in organic electrolyte [1 M LiPF6 in ethylene carbonate/diethyl carbonate (EC/DEC)] at 30 °C for 2 months or 50°C for1.5 months, there was no detectable disintegration of paper andCNTs (Fig. 5 A and B). Moreover, the sheet resistance of theconductive paper decreased from =60 ,Q./sq down to 9 il;sq aftersoaking in the organic electrolyte, which might be due to thedissolution of surfactant or hole doping of CNTs (44). On thedevice level, batteries with CNT conductive paper as currentcollectors were cycled 500 times, as shown in Fig. 5C. Thecapacity retention was 95% after 280 cycles at C/3. Moreover,during the following 220 cycles at the rate of C/2, the capacitydecay was less than 0.01% per cycle. The total time for the 500cycles is =3.5 months. In addition, after stopping cycling for aweek, the charge capacity only increased by 3.3%, indicating thatthere is not much self-discharge for batteries with paper-basedbatteries. These experiments demonstrate that paper is stable inelectrolyte for at least months, and the stable period of paper inorganic electrolyte could further extend to more than 1 year,which is close to the shelf life requirement for some applications.

ConclusionIn conclusion, we have made highly conductive CNT paper byconformal coating of CNTs onto commercial paper, whoseconductivity can be further enhanced by incorporating metalnanowire strips as global current collectors for large-scale energy-storage devices (Figs. S4D). The intrinsic properties ofpaper, such as high solvent absorption and strong binding withnanomaterials, allow easy and scalable coating procedures.Taking advantage of the mature paper technology, low cost, lightand high-performance energy-storage devices are realized byusing conductive paper as current collectors and electrodes. Theconcept of using paper as a novel substrate together withsolution-processed nanoscale materials could bring in new opportunities for advanced applications in energy storage andconversion. By combining our paper-based energy storage withother types of devices developed, such as bioassays or displays onpaper, full paper electronics could be realized in the future.

Materials and MethodsPreparation of Inks and Conductive Paper. To form a CNT ink, CNTs grown bylaser ablation and SDBS (Sigma-Aldrich) were dispersed in deionized water.Their concentrations were 10 and 1-5 mg/mL. respectively. After bath sonication for 5 min, the CNT dispersion was probe-sonicated for 30 min at 200 Wfl!C505; Sonics)to form an ink. Meyer rods (Rdspecialties) were used to coatthe CNT ink onto Xerox paper. The sheet resistance of conductive paper wasmeasured by using the four-point probe technique (EDTM). To make silvernanowire (Ag NW)ink, Ag NWswere produced insolution phasefollowingthemethod of Xia and colleagues (45). In the first step, a mixture of 0.668 g ofpolyvinylpyrrolidone (PVP) and 20 mLof ethylene glycol (EG) was heated in aflask at 170 QC. Once the temperature was stabilized, 0.05~ g of silver chloride

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reasonable internal resistance (Fig. S7),even considering the weightof all components in a battery. Hence, higher gravimetric energydensity can be achieved in such paper batteries. Previously, Pushparaj et aI. (25) fabricated "paper batteries" with CNTs themselvesas the active electrode material for Li-ion batteries. However, as anactive material, CNTs suffer from issues of poor initial Coulombefficiency, unsuitable voltage profiles, and fast-capacity decay (25,37). Instead of using the CNTs to store lithium ions, we used themto function as lightweight current collectors to achieve practicalbatteries with a long cycle life. Fig. 4A shows the structure ofconductive paper-based battery. LiMn204 nanorods (38) andLi,Ti5012 nanopowders (=200 nm; Slid Chemie) or Si/C (39)nanowires were coated onto conductive paper to act as the cathodeand anode, respectively. Fig. 4B displaysthe initial charge/dischargecurves of half cellsconsisting of the LiMnZ04nanorods or Li4Tis012

nanopowders coated on conductive paper as working electrodesand lithium foil as counter electrodes. Voltage profiles were closeto those with metal current collectors, according to previous work,and no apparent voltage drop was observed (38, 40-42). Thecycling performance of these conductive paper-supported electrodes is shown in Fig. 4C. The LiMnz04 nanorod and Li4Tis0 12

nanopowder electrodes achieved initial discharge capacities of 110mAhlg and 149 mAhlg, and capacity retentions of 93% and 96%after 50 cycles at 03, respectively. These values are comparablewith metal collector-based batteries (38, 40-42). In our devices, thecoulomb efficiency was generally =98.5% for LiMnZ04 and over99.5% for LL,Ti,012. To demonstrate a practical paper battery, asern?full cell with conductive paper acting as the current collectorin both cathode and anode was used to repeatedly light a blue LED,as shown in Fig.4D. In this demonstration, the cathode is LiMnz04,whereas the anode is carbon/silicon core/shell nanowires.

The chemical stability of conductive paper in the electrolytemay be a main concern for practical uses. Paper has been usedas the separator in aluminum electrolytic capacitors with aqueous solution (43), and our tests also show that paper is stable inaqueous electrolyte (1 M H2S04) for 2 months. Regardingorganic electrolyte, the stability of paper is tested under both

Fig. 4. Conductive paper as the current collector for li-ion batteries. (A)Schematic illustration of the conductive paper battery configuration. Themetal leads contact only CNTs but not active materials. Lithium foil is used asthe counter electrode in half-cell tests. (B)Galvanostaticchargingfdischargingcurves of UMnZ04 nanorod cathode (3.5-4.3 V) and li4Tis012 nanopowderanode (1.3~1.7 V) half-cells with conductive paper current collectors. Thecurrent rate isCIS. (C)Cyclingperformance of liMnz04 nanorod (Cl3,49 rnA/g)and li4Tis0 12. nanopowder (03, 58 rnA/g) half-cells. (D) A 5 cm2 paper battery(a full cell with liMn204 nanorod cathode, aSi core/shell NW anode, andconductive paper current collectors) used to repeatedly light up a blue LED.

4 of 5 I www.pnas.org/cgi/doi!10.1073/pnas.0908858106 Hu et el.

(AgCI)was ground finely and added to the flask for initial nucleation. After 3minutes, 0.22 g of silver nitrate (AgN03) was titrated for 10 minutes. Then, theflask was kept at the same temperature for another 30 minutes. After thereaction was completed, the solution was cooled down and centrifuged threetimes to remove solvent, PVP, and other impurities.

Cell Preparation and Measurements of SCs. For aqueous electrolyte devices,two pieces of CNT conductive paper were first attached on glass slides. CNTfilms were used as both electrodes and current collectors. At the end of theCNT paper, a small piece of platinum was clipped onto the CNT conductivepaper by a toothless alligator clip to connect to a battery analyzer (Maccor4300). Both glass slides were assembled with a separator (Whatman 8-p.m

filter paper) sandwiched in between. The paper assembly was wrapped withparafilm and then dipped in the electrolyte solution. The active area overlapped by both CNT conductive papers was 1 cm2. For organic electrolytedevices, cells were assembled by inserting the same separator soaked with thestandard battery electrolyte (1 M UPF6 in ethylene carbonate:diethylenecarbonate> 1:1 vol/vol; Ferro) between two CNTconductive paper substrates.The active area overlapped by both CNTconductive paper substrates was also'1 cm''.Then, the entire assembly was sealed in a polybag (Sigma-Aldrich). Asin the aqueous cells, small pieces of platinum were attached to the end of CNTconductive paper for a good electrical contact. The current collectors came outthrough the sealed edges of polybags and then were connected to the batteryanalyzer. All steps in the cell preparation were done in an argon-filled glovebox (oxygen and water contents below 1 and 0.1 ppm, respectively). Typicalmass loadings for data shown in the main text Fig. 3 C and 0 are 72 ~ 270p.glcmz. Larger mass loadings up to 1.7 mglcmZwere also tested (Fig. 3 0 and£, and Fig. S6C), and the capacitances are plotted in Fig. 56. Capacitance,energy density, and power density are all characterized by galvanostaticmeasurements. A total of 0.02 = 20 mAlcm z were applied to cells whilepotentials between both electrodes swept between cutoff values (0 ::0; V ::0;0.85 = 1 Vin aqueous phase, 0::0; V ::0;2.3 "'"3 Vin organic phase). Voltages wererecorded every 0.01 "'" 0.2 seconds. For the cycling test in both phases ofelectrolyte, =5 Alg was applied. The cutoff potentialsforthe sulfuric acid andorganic electrolyte were 0.85 and 2.3 V, respectively.

Battery Fabrication and Test. The cathode materials LiMnz04 nanorods weresynthesized according to our previous work, with modification (38). Typically,

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micropatterning processfor paper-based microfluidics. Anal Chem 81:7091-7095.12. Lamprecht8, et al. (2005) Organicphotodiodes on newspaper. PhysicaStatus Solidi A

202:R50-R52.13. Kaihovirta NWC, Makela T, Wilen C, Osterbacka R (2009) Self-supported ion

conductive membrane-based transistors. Adv Mater 21:2520-2523.14. EderF,et al. (2004) Organicelectronics on paper. Appl Phys Lett 84:2673-2675.15. Kim DHK, et at. (2009) Ultrathin siliconcircuitswith strain-isolation layersand mesh

layouts for high-performance electronics on fabric, vinyl, leather, and paper. AdvMater 21:3703-3707.

16. Anderson P,et al. (2002)Activematrix displaysbased on all-organic electrochemicalsmart printed on paper.Adv Mater 14:1460-1464.

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18. Kordas K, et at. (2006) Inkjet printing of electricallyconductive patterns of carbonnanotubes. SmaJ/2:1021-1025.

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Hu et al.

8 mmol MnS04'HZO and 8 mmol (NH4)zSZOS were dissolved in 20 ml ofdeionized water, and the solution was transferred to a 45-ml, Teflon-linedstainless steel vessel (Parr). The vessel was sealed and heated at 150 "Cfor 12 hto obtain I3~Mn02 nanorods. The as-synthesized Mn02 nanorods were mixedand ground with lithium acetate (Aldrich) at a molar ratio of 2:1. A total of 1mL of methanol was added to make a uniform slurry mixture. Then, themixture was slntered at 700 "Cfor 10 h under airto obtain LiMnz04 nanorods.The carbon/silicon core/shell nanowires were synthesized by CVD method.Carbon nanofibers (Sigma-Aldrich) were loaded into a tube furnace andheated to 500 "c. Then, silane gas was introduced and decomposed ontocarbon nanofibers. The weight ratio of silicon shell to carbon core wastypically=2:1. Li4Tis012 powder was used as received from SOd Chemie.

Electrodes for electrochemical studies of LiMn204 and Li4Tis012 were prepared by making slurry of 70 wt % active materials, 20 wt % Super P Carbon,and 1Owt % PVDFbinder in N-methyl-2~pyrrolidone (NMP) as the solvent. Theslurry was coated onto a piece of conductive CNT paper by an applicator andthen dried at 100 "C in a vacuum oven overnight. For ClSicore/shell nanowires,the as-synthesized nanowlres were dropped onto a CNT paper and dried toform the anode.

The half-cell tests of both cathode (UMnZ04) and anode (Li4TisOd werecarried out inside a coffee bag (pouch) cell assembled in an argon-filledglovebox (oxygen and water contents below 1 and 0.1 ppm, respectively).Lithium metal foil (Alfa Aesar) was used asthe counter electrode in eech case.A 1 M solution of LiPF6 in EOOEC(1:1 vollvol; Ferro) was used asthe electrolyte,with separators from Asahi Kasel. The charge/discharge cycles were performed at different rates at room temperature, where 1C was 148 mAlg forLiMnz04 and 175 mAlg for Li4Tis012, respectively. The voltage range was3.5-4.3 V for LiMn204 and 1.3-1.7 V for li4Tis012. Tests were performed byeither Bio-Loqic VMP3 battery testers or MTIbattery analyzers. To fabricate afull cell with high voltage to light a blue LED; slliconlcarbon core/shell nanowires and LiMn204 nanorods were used as anode and cathode, respectively.Then, the two electrodes were assembled to make a 5 cm2 pouch cell asdescribed above, and it was used to repeatedly light the blue LED.

ACKNOWLEDGMENTS. This work was supported byThe Korea Foundation forAdvanced Studies (S.J.) and King Abdullah University of Science and Technology Investigator Award KUS-11-001-12 (to v.cj.

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PNAS Early Edition I 5 of 5

Attachment ld

S. Keren, C. Zavaleta, Z. Cheng, A. de la Zerda, O. Gheysens, and S. S. Gambhir*

Molecular Imaging Program at Stanford, Departments of Radiology and Bioengineering, Blo-XProgram, Stanford University, 1201 Welch Road, Stanford.CA 94305-5484

Edited by Michael E. Phelps, University of California, los Angeles School of Medicine, los Angeles, CA, and approved February 5, 2008 (received for reviewNovember 7, 2007)

Author contributions: S.K. and C.Z.contributed equally to this work; S.K.,C.Z.,and 5.5.G.designed research; S.K.,C.Z.,z.c., A.d.l.Z., and O.G. performed research; c.z. analyzed data;and S.K.,eLZ., and 5.5.G. wrote the paper.

The authors declare no conflict of Interest.

This article is a PNASDirect Submission.

Freely available online through the PNASopen access option.

"To whom correspondence should be addressed. E-mail:[email protected].

This article contains supporting information online at www.pnas.org/cgilcontentifuIV0710575105/DCSuppiementa I.

@2008 by The National Academy of Sciences of the USA

nitude of the Raman effect (=1 photon is inelastically scatteredfor every 107 elastically scattered photons) limits the sensitivityand, as a result, the biomedical applications of Raman spectroscopy. The discovery of the surface enhanced Raman scattering(SERS) phenomenon offers an excitingopportunity to overcomethis lack of sensitivity and introduce Raman spectroscopy intonew fields (8). SERS is a plasmonic effect where moleculesadsorbed onto nano-roughened noble metal surfaces experiencea dramatic increase in the incident electromagnetic field, resulting in high Raman intensities comparable to fluorescence (6).

Single-walled carbon nanotubes (SWNTS) also show an intense Raman peak produced by the strong electron-phononcoupling that causes efficient excitation of tangential vibrationin the nanotubes quasi one-dimensional structure upon lightexposure (9). Recent demonstrations of tumor targeting, usingradiolabeled SWNTS (10) combined with lowtoxicity effects andrapid renal excretion (11), suggest that carbon nanotubes mayalso become promising molecular imaging agents for livingsubjects. Here, we report for the first time, to our knowledge,noninvasive imaging of SERS active nanoparticles [namely,Nanoplex Biotags (Oxonica) were used in this study and henceforth are referred to as SERS-biotags or biotags] and SWNTnanoparticles in small living subjects by using Raman spectroscopy. Instrument adaptation to small-animal imaging, quantification of reproducibility and sensitivity, pharmacokinetics ofnanoparticies, demonstration of multiplexing, in vivo tumortargeting after tail-vein injection, and whole-body deep-tissueimaging are all established in this work.

ResultsOptimization of Raman Imaging Instrumentation. Optical microscopes designed for surface imaging through transparent mediaface significant losses of light when imaging through a diffusivetissue because of light scattering. We used a Raman imagingsetup based on a Renishaw InVia Raman microscope equippedwith an excitation laser at 785 nm, which is typically used forsurface imaging. Several measures were taken to optimize thissetup specifically for small-animal imaging. A high numericalaperture (NA) objective, commonly used on this microscopesetup, wasfound to be less than ideal when imaging small animalsfor both illumination and light collection. A small spot sizeillumination obtained by a high NA objective limits the permissible laser power that can be used before tissue damage. Moreover, high NA objectivescollect light efficiently onlyfrom a small

Molecular imaging of living subjects continues to rapidly evolvewith bioluminescence and fluorescence strategies. in particularbeing frequently used for small-animal models. This article presents noninvasive deep-tissue molecular images in a living subjectwith the use of Raman spectroscopy. We describe a strategy forsmall-animal optical imaging based on Raman spectroscopy andRaman nanoparticles. Surface-enhanced Raman scattering nanoparticles and single-wall carbon nanotubes were used to demonstrate whole-body Raman imaging, nanoparticle pharmacokinetics, multiplexing. and in vivo tumor targeting. using an imagingsystem adapted for small-animal Raman imaging. The imagingmodality reported here holds significant potential as a strategy forbiomedical imaging of living subjects.

M olecular imaging of living subjects provides the ability tostudy cellular and molecular processes that have the

potential to impact many facets of biomedical research andclinical patient management (1-4). Imaging of small-animalmodels is currently possible by using positron emission tomography (PET). single photon emission computed tomography.magnetic resonance imaging, computed tomography, opticalbioluminescence and fluorescence, high frequency ultrasound,and several other emerging modalities. However, no singlemodality currently meets the needs of high sensitivity, highspatial and temporal resolution, high multiplexing capacity, lowcost, and high-throughput.

Fluorescence imaging, in particular, has significant potentialfor in vivostudies but is limited by several factors (5, 6), includinga limited number of fluorescent molecular imaging agentsavailable in the near infra-red (NIR) window with large spectraloverlap between them, which restricts the ability to interrogatemultiple targets simultaneously (multiplexing). In addition,background autofluorescence emanating from superficial tissuelayers restricts the sensitivity and the depth to which fluorescence imaging can be used. Moreover, rapid photobleaching offluorescent molecules limits their useful lifetime and preventsstudies of prolonged duration. Therefore, We have attempted todevelop new strategies that may solve some of the limitations offluorescence imaging in living subjects.

Raman spectroscopy can differentiate the spectral fingerprintof many molecules, resulting in very high multiplexing capabilities. Narrow spectral features are easily separated from thebroadband autofluorescence, because Raman is a scatteringphenomenon as opposed to absorption/emission in fluorescence,and Raman active molecules are more photostable comparedwith fluorophores, which are rapidly photobleached. Unfortunately, the precise mechanism for photobleaching is not wellunderstood. However, it has been linked to a transition from theexcited singlet state to the excited triplet state. Photobleachingis significantlyreduced for single molecules adsorbed onto metalparticles because of the rapid quenching of excited electrons bythe metal surface, thus preventing excited-state reactions andhence photobleaching (7). However, the inherently weak mag-

nanotubes ! SERS nanoparticles

Noninvasive molecular imaging of small livingsubjects using Raman spectroscopy

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spot size; a highly diffusive propagation of light photons thronghtissue results in only a very small number of scattered photonsemanating from the same small spot size. For these reasons, wefound that a X 12 open field lens with a defocused beam,resultingin a spot size afZO X 200 p.m,workedwell. The spectralresolutionwas adjustedto allowhigh sensitivitywhile being ableto multiplex different SERS nanoparticles by opening the monochromatorslit (lOO-,u,lll width) to obtain a spectralresolution of10 cm'<'. A computer controlled stage was also added to themicroscope setup to allow automated mappingof large surfaces.Raman images were obtained by using a Raman point-mappingmethod. The computer-controlled x-y translation stage performed a raster scan of a region of interest over the mouse, theRaman spectrum was measured at every point, and softwarealgorithms were used to calculate the SERS nanoparticle concentration and generate a two-dimensional mapping image ofthe SERS nanoparticle distribution. Finally, a heated bed and ananesthesia inhalation unit were attached to the microscope stageto allow a long, motionless scan time at a stabilized bodytemperature. A photograph of the imaging setup is shown insupporting information (Sf) Fig. Sla.

Raman Nanoparticles. The ideal properties of a nanoparticle usedfor small-animal Raman spectroscopy would include small dimensions, simple conjugation methods, no toxicity effects, andintense and unique Raman spectra. Recently developed SERSactive nanoparticles, called Nanoplex Biotags (Oxonica), composed of a gold core, Raman-active molecular layer (see Methods), and silica coating, schematically shown in Fig. SIb, holdsignificant potential for in vivo imaging applications (12). Theglass coating of these SERS nanoparticles guarantees physicalrobustness, insensitivity to environmental conditions, and simplebiofunctionalization of the well studied silica surface chemistry.The SERS-nanoparticles were designed to maximize the Ramansignal, and the NIR excitation and emission profiles are ideal forminimizing light absorption by tissue. SERS nanoparticles usedin this study had a mean diameter of 120 nm, as seen in thetransmission electron microscopy image in Fig. S2.

SWNTS exhibit a strong Raman peak at 1,593 cm", whichallows high sensitivity detection (Fig. S3). Unlike the SERSnanoparticles, these SWNTs are inherently Raman active and donot use a metal surface enhancer to increase Raman detection.The high aspect ratio of the carbon structure of SWNTs is idealfor bioconjugation, and recent reports have successfully shownspecific tumor targeting in vivo within various tumor models,using various functionalized SWNTs (10, 13, 14). SWNTs havea very small diameter of =3 nm and a length of 200 nm, asschematically shown in Fig. S1c and on the atomic force microscopy image in Fig. S2b. SERS and SWNT nanoparticles wereboth tested for stability in mouse serum and incubated at 37°Cover a 5-day period with no degradation of Raman signaldetected (data not shown).

Imaging SystemCharacterization. We demonstrated the in vitro andin vivo reproducibility of the measured and processed Ramanspectra. Five samples (5-iLlvolume), each with 6.6 fmol of SERSnanoparticles, were measured on a piece of parafilm, using theRaman imaging setup. The coefficient of variance (COY) of thecalculated SERS nanoparticle concentration, using the quantitative analysis method described in Methods, showed a reproducibility of 1.9%. Reproducibility was also evaluated in a nudemouse with four separate s.c. injections of 16 fmol of SERSnanoparticle (10-f..d volume) mixed with a gelatinous proteinmixture known as matrigel, which resembles the complex extracellular environment found in many tissues (10-pJ volume). Thematrigel was used to keep the SERS nanoparticles from diffusingquickly out of the skin and showed no inherent Raman spectraas expected (Fig. S4). The SERS nanoparticle component con-

Keren et al.

centration calculated for different injection sites showed a COYof 3.1%. Deep tissue reproducibility was also determined byinjecting 260 fmol of SERS nanoparticles (200-iLI volume) via thetail-vein into three nude mice. Nanoparticles of various types arenaturally taken up by the reticuloendothelial system and thus canbe found in the Kupffer cells of the liver (15). The liver of eachmouse was imaged =30 min after tail-vein injection, and component analysis revealed a COY of 16.7% between the threemice. Naturally, the higher variability could be attributed to anumber of factors, such as positioning, injection technique, andindividual animal pharmacokinetics. Reproducibility in the liverof a single mouse imaged repeatedly (eight times) showed a COYof 2.8% (data not shown).

Sensitivity was evaluated by s.c. injecting three mice withdecreasing concentrations of SERS nanoparticles in a volume of20 n.l. The data revealed a highly linear relationship betweencalculated and injected concentration of nanopartic1es with anR2 ~ 0.99. The smallest amount of SERS nanoparticles detectedin a 20-pJ s.c. injection was 8.125 pM. Ex vivo measurementsshowed high linearity with R2 = 0.997 and a detection limit as lowas 600 particles (Fig. S5).

The maximum depth of penetration for our Raman microscope was evaluated by using a tissue mimicking phantom, wherea maximum depth of 2 mm was observed by using 6 nM SWNTsand 5.5mm, using 1.3 nM SERS nanoparticles (for more details,see Sf Text). However, deeper penetration could be achieved byincreasing administered nanoparticle concentration or slightlyincreasing the laser power while stilI remaining below exposurelimits.

The effect of changing the working distance between the lensand the subject was also evaluated ex vivo, by changing the stageheight that contained a 5-iLl sample of 6.6 fmol of SERSnanopartic1es and in vivo with a mouse that had been injected viatail vein, with a 200-iLI sample of 260 fmol of SERS nanoparticles. The component concentration was found to increaseexponentially as the sample per mouse on the stage was movedcloser to the lens of the microscope (data not shown). However,calculated concentration only weakly depends on variations inobjective-subject distance when detected through the diffusivemouse tissue.

Demonstration of SERS NanopartideMultiplexing in Mice. Each typeof the SERS nanoparticles (Nanoplex biotags) contains a differentRaman-active material with its own unique spectral fingerprintallowing detection and quantification of multiple tags simultaneously within the same animal. Two types, SERS 421 and SERS440, with different Raman signatureswere s.c. injected into a mouseto demonstrate the multiplexing capabilities in living subjects. Thefirst s.c. injection consisted of 5 p.l (6.6 fmol) of S421 and 15 iLl ofmatrigel (Fig. 1a). The second s.c. injection consisted of 5-iLl (6.6fmol) of SERS 440 and 15 iLl of matrigel (Fig. Ib). The third s.c.injection consisted of an equal mix of SERS 421 (5 iLl),SERS 440(5 iLl), and 10 p.l of matrigel (Fig. Ie). Based on their differentRaman spectra, the concentration of each SERS nanopartic1e couldbe calculated by using the component analysis method describedlater (see Methods). Fig. I shows the concentration mapping of theinjection sites. The area of the three s.c. injectionswas mapped witha step size of 500 ,LLm and an integration time of 1 s. The map showsthe three individual injection sites; each assigned a color by theNanoplex software based on their corresponding spectra, with theconcentration magnitude portrayed as pixel brightness. Note thatthe third injection site was calculated to have a yellow color thatcorrelates with an equal mix of the SERS 421 (red) and SERS 440(green) SERS nanoparticles. Ex vivo experiments verifying thesystem's multiplexing capability are shown in Fig. 56.

In addition, we have been able to successfully multiplex fourtypes of SERS Raman nanoparticles ofvarying concentrations ina living mouse. The first four s.c. injections consisted of 5 ,LLI (6.6

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Fig. 1. Evaluation of multiplexing experiment. (a) Ramanspectrum acquired from first S.c. injectionof 5421SERS nanoparticles. Thesoftware hasassigned thecolorred forthis particularRamanspectrum. (b) Ramanspectrum acquiredfrom second s.c. injectionof 5440SERS nanopartkles.Thesoftware hasassigned thecolor green for this particular Ramanspectrum. (c) Ramanspectrum acquiredfrom the third s.c. injection of an equal mix of 5421 and 5440. Notice how thisspectrum represents an equal mix of both individualspectrums asif they had been overlaid. Asa result,the coloryellow iscalculated by the analysis softwareto represent an equal mix of the red (5421) and green (5440) SERS nanoparticles in the map at Right.

WholewBody Noninvasive Imaging of SERS Nanoparticles. For wholebody mapping, mice were tail-vein injected with 260 fmol ofSERS nanoparticles in a 200-ILI volume. A raster scan wasacquired 2 h after injection over a large portion of the mousebody with 1 mm step size and 3 s integration time. Quantitativeanalysis software calculated the concentration of SERS nanoparticles and generated an image showing accumulation of theSERS nanoparticles in the liver (Fig. 3a). A finer mapping withstep size of 750 ,urn over the liver region reveals a detailed imageof the liver. A slight distinction between liver lobes can be seenin this higher resolution image (Fig. 3b). SERS nanoparticleswere visualized in the liver region out to 24 days after tail-veininjection, and continued to produce a recognizable spectrumwith sustainable intensity for 24 days. Killed animals underwentex vivo tissue analysis, and we confirmed the presence of SERSnanoparticles in the liver of mice (data not shown).

different SERS nanoparticles, which nearly simultaneously accumulated in the liver (Fig. S9). The first group of mice (n ~ 3)that received an equal mix of 5 kDa of PEG and nonpegylatedSERS nanoparticles showed no difference in liver accumulationbetween each of the nanoparticles and at 2 min after injection(Fig. 2a). The second group of mice (n ~ 3), which received anequal mixof 20 kDa PEG and nonpegylated SERS nanoparticles,also showed no difference in liver accumulation between each ofthe nanoparticles and plateaued at 4 min after injection (Fig. 2b).As with many nanoparticles administered intravenously, evadingthe macrophages of the reticuloendothelial system remains aconstant problem, where in some in vivo cases pegylation isinsufficient, as seen in our data and reported by Moghimi et at.The pharmacokinetics of SWNTs were also evaluated over aperiod of 90 min in 4 mice to reveal a fluctuation in nanotubedistribution in the liver over the first 10 min followed by acontinuous increase out to 90 min (Fig. 2c).

Whole-Body Noninvasive Imaging and Tumor Targeting of SWNTs.SWNTs were also imaged in nude mice after tail-veininjection of=60 pmol in 200I-'!. An intense peak at 1593cm" makes nanotubeseasily detected with Raman spectroscopy. A Raman image wasacquired with a raster scan 2 h after injectionwith a step size of 1

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liver Pharmacokinetics of SERS Nanoparticles and 5WNTs. Takingadvantage of the multiplexing capabilities ofSERS nanoparticlesdetected by Raman spectroscopy, we studied the circulationproperties of both pegylated and nonpegylated SERS nanoparticles simultaneously in living mice. An equal mixture of pegylated (PEG) and nonpegylated SERS nanoparticles (260 fmol ofin a 200-JLl volume), each with different Raman signatures, weretail-vein injected to evaluate their accumulation in the liver as afunction of time. The laser was positioned over the mouse's liverbefore injection, and a Raman spectrum was acquired with a 10-sintegration time over a period of 90 min. Quantitative analysiswas used to calculate the relative concentration of the two

fmol) of each SERS particle and 5 1-'1 of matrigel as shown in Fig.S7a as follows: SERS 482 (green), SERS 420 (red), SERS 481(yellow), and SERS 421 (blue). The fiftb s.c, injection (purple)at the far right consisted of a mixture of these four SERSparticles with varying concentrations to determine multiplexingin vivo (Fig. S7a). The mixture contained 41-'1 of SERS 420 (5.28fmol), 3 1-'1 of SERS 421 (3.96 fmol), 2 1-'1 of SERS 481 (2.64fmol), 11-'1 of SERS 482 (1.32fmol), and 10 1-'1 of matrige!. Basedon their different Raman spectra, the concentration of eachSERS nanoparticle could be calculated by using the componentanalysis method. Fig. S7 b-e show the four SIlRS componentsand the correlating intensity of the fifth injection site corresponding to the concentration of that particularcomponent. Thedifferent intensities of the fifth injection site represented in eachof the maps (Fig. S7 b-e) qualitatively correlates with thedifferent concentrations of each SERS nanoparticle mixed.Notice how SERS 420 has the most intense pixel brightness in thefifth injection site (Fig. S7c), followed by SERS 421 (Fig. S7e)with the second most intense, then SERS 481with the third mostinteuse (Fig. S7d), and finally SERS 482 with the least intense atthe fifth injection site (Fig. S7b). The area of the five s.c.injections was mapped with a step size of 750 JLm and anintegration time of 1 s. The map shows the five individualinjection sites; each is assigned a color by the software based ontheir corresponding spectra, with the concentration magnitudeportrayed as pixel brightness. The unique Raman spectra ofthese four different SERS nanoparticle are depicted in Fig. S8.

5846 ! www.pnas.org/cgi/doi/l0.l073/pnas.0710575105 Keren et al.

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Fig. 3. Raster-scan image of mouse liver, using Raman spectroscopy inconjunction with SERS nanoparticles. (a) Whole-body map (1-mm steps) ofnude mouse 2 h after tail-vein injection ofSERS nencpertldes. Note how mostof the SERS particles accumulated in the liver (L) (arrow), resulting in a welldefined image. (b) Map of liver (750-tLm steps), showing higher definition ofliver (arrow) and slight distinction between the two liver lobes.

Fig.4. Accumulation of RGD SWNTsand plain nontargeted SWNTswithin anlnteqrln-posltlve U87MG tumor model at 24 h after Lv. injection. (a) Digitalphotograph of mouse depicting tumor area (black square) and correspondingRaman images acquired 24 h after SWNTinjection by raster scan with 750-tLmsteps. Noticethe accumulation of RGD SWNTsin the tumor area as opposed tothe plain nontargeted SWNTsthat show littleto no accumulation inthetumorarea. (b) Raman spectral analysis of RGD nanotubes and plain nontargetednanotubes within the tumor at 24 h after SWNTinjection. The graphed datashow a significant increase (-. P < 0.05) in Raman signal in mice (n = 3 pergroup) injected with RGDnanotubes as opposed to mice injected with plainnanotubes, thus indicating accumulation of RGD SWNTto tumor site. Thisquantitative data supports the Raman images displayed in a.

of RGD conjugated SWNTs of =60 pmol in 200 iLL The secondgroup (control) received an i.v. injection of plain nontargetedSWNTs of the same concentration. Raster scans were acquired at24 h after SWNT injection over the tumor area with 750-iLm stepsto generate Raman images (Fig. 4a). The images revealed anintense accumulation of RGD conjugated SWNTs in the tumorarea; however, little to no accumulation of plain SWNTs wasobserved in the tumor area at 24 h after injection. Raman spectrawere taken in living mice at 24 h after injection and revealed asignificant increase (P < 0.05) in SWNT accumulation in the tumorareas of experimental mice receiving RGD SWNTs (SWNT concentration: 0.0204 ± 0.0087) compared with the control groupreceiving plain SWNTs (SWNT concentration: 0.0016 ± 0.0005)(Fig.4b).

DiscussionIn summary, we adapted a Raman microscope to demonstrateRaman imaging of small living subjects while using two differenttypes of Raman nanoparticles, SERS active nanoparticles (Nano-

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Fig.2. Pharmacokinetics of SERS nanoparticles and SWNTsin the liver [dataacquisition starts at 10 s, zero accumulation at time point zero (data notshown)]. (a) Accumulation of nonpegylated SERS versus 5 kDa of PEG SERSnenoperttcles in liver of nude mouse. The graph depicts the mean normalizedconcentration ofSERSnenopartlcles in three mice::!: SEM.(b) Accumulation ofnonpegylated SERS versus 20 kDa of PEGSERS nanoparticle in liver of nudemouse. The graph depicts the mean normalized concentration of SERS nanoparticles in three mice :t SEM. (c) Pharmacokinetics of nanotubes in liverevaluated >90 min after tail-vein injection. The graph depicts the meannormalized concentration of nanotubes in four mfce c SEM.Note the gradualincrease of SWNTaccumulation in liver after 30 min after injection.

mm and an integration time of 3 s. The map revealed accumulationof nanotubes in the liver and a random distribution faintly dispersedacross the peritoneal cavity (Fig. SlOa). The SWNTs were alsoevaluated daily for liver accumulation. and showed an increase inRaman intensity several days after tail-vein injection. Because ofthis continuous rise in Raman signal in the liver, another map of thesame area, using the same image acquisition parameters, was takenat 72 h after injection to reveal a better defined liver with betterdelineation of the liver than the previous 2 h image of the same area(Fig. SlOb). Daily evalnation of the liver continued to show Ramansignal in the liver until 12 days after injection, at which time theanimals were killed. Killed animals underwent ex vivo tissue analysis, and we confirmed the presence of nanotubes in the liver ofmice (data not shown).

Furthermore, preliminary data demonstrated the ability of ourmodified Raman microscope to detect targeting of SWNTs conjugated with arginine-glycine-aspartate (RGD) peptide in an integrin positive U87MG tumor model in living mice. This RGDpeptide binds to cxvf33 integrin, which is overexpressed in angiogenicvessels and various tumor cells (16). Tumor targeting of RGDSWNTs was described by Liu et al. (10), using microPET imagingand exvivo Raman imaging of tissues. Sixmice, s.c. inoculated with2 X 10' U87MG cellsnear the right shoulder were divided into twogroups. The first group (experimental) received a tail-vein injection

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plex Biotags), and SWNTs. We have shown relatively high signalreprodncibility both in vitro and in living subjects and the ability toproduce an imageof nanoparticlesfrom both s.c.locationsand fromdeeper tissues (e.g., liver) in living mice. Furthermore, we havedemonstrated the ability to follow the arrival of nanoparticles inliver tissue to create time-activity curves. A minimum detectionsensitivity of 8.125pM was observed in a livingmouse while usingSERS nanoparticles. The ability to multiplex with four SERSnanopartic1es presenting different Raman spectra was also demonstrated with the rapid and straightforward distinction betweenthese nanoparticles in livingmice. These initial results are encouraging and demonstrate the potential robustness of a Raman-basedimaging strategy for small living subjects.

In the current proof-of-principle work, we demonstrated detection of Raman nanopartic1es in both superficial and deep tissuesalong with an initial evaluation of its potential to detect tumortargeting with SWNTs conjugated to RGD peptide in an U87MGanimal model. The primary limitation to Raman imaging of largersubjects will be those also faced by other opticaltechniqnes and islimited by NIR light penetration beyond a few centimeters of tissue(17). The key advantages of the current Raman imaging strategyover fluorescence is the very high multiplexing capability and lackof confounding background signal from autofluorescence. A studyby Souza et 01. (18) was able to collect a Raman spectra from thesurface of a mouse (without forming any images) but had to useintratumoral injection of nanoparticles and also a very large concentration (3.8 I-'M) of Raman nanoparticles in a relatively largevolume of 3001-'1 (18). Auother group has applied multiple Ramansignals to follow the distribution of cholesterol in a rat eye whileshowing the phenotyping of lymphocytes by using a single nonenbanced Raman-labeled (filipin) polystyrene bead to monitor cholesterol in a rat eye in conjunction with the inherent Raman peakassociated with proteins to follow the distribution of cholesterol(19). Furthermore, it was showu that these Raman-labeled polystyrene spheres could be successfully used in combination with afluorescent label. Using an additional Raman label, can overcomethe many limitations that arise with fluorescence such as photobleaching, autofluorescence, and a limited number of fluorescentlabels. Although we demonstrated multiplexing with only fourSERS nanoparticles, we should be able to easily image many moresimultaneously injected"SERS nanoparticles, and as many as 10spectrally distinct SERS nanoparticles are already available (e.g.,Oxonica).

Glass-encapsulated SERS active nanoparticles are only onekind of Raman particle recently developed that can be used ascontrast agents for molecular imaging. Roughened surface noblemetal nanoparticles labeled with binding affinity molecules havealso been shown to dramatically increase the Raman signal oftheir complementary molecules once attached through theSERS mechanism (12, 20, 21). Conjugation of monoclonalantibodies with SERS nanoparticles for targeting and imaging ofspecific cancer markers in live cultured cells was also recentlyinvestigated (5). Another study presents the first in vivo application of SERS for glucose measurements in a rat (22) by s.c.implantation of functionalized SERS nanospheres.

Recent publications have reviewed the rapid increase ofRaman spectroscopy in many biomedical applications (23-25).One report has demonstrated the ability of Raman spectroscopy to diagnose benign and malignant excised breast tissuewith high sensitivity aud specificity (26). Another articlediscusses the potential of Raman spectroscopy to analyze themorphologic composition of atherosclerotic coronary arterylesions and assess plaque instability and disease progression invivo (27). The recent development of a highly efficient Ramanoptical fiber probe has now bridged the gap between Ramanspectroscopy and the clinical setting. The optical fiber probehas successfully demonstrated its usefulness in assessing human tissue models for disease and thus has great potential as

5848 I www.pnas.org/cgijdoij10.1073/pnas.0710575105

a clinically practical technique (28, 29). In particular, thisRaman optical fiber probe has the potential of being anextremely useful tool in an intraoperative setting, for instancein surgical debulking of cancers. Its ultra-high sensitivity wouldbe useful in detecting even the smallest presence of malignanttissues in the human body for excision. By using targeted SERSnanoparticles and noninvasive in vivo imaging, Raman spectroscopy can become an important clinical diagnostic tool, asdemonstrated in this article.

Biodistribution of these Raman nanoparticles are currently beingevaluated in our lab with the use of radiolabels (e.g., with positronemitters) so that quantification of exact nanoparticle concentrationcan be explored as we have recently done with radiolabeledqnantum dots (30). Additioual work with tissue targeting of theRaman nanoparticles (SERS and SWNTS) should also help tofurther expand the eventual utility of the strategies developed in thiswork. Fnrther studies with both SERS and SWNTS will still beneeded to understand any potential limitations, including deliverydue to nanoparticle size, optimal injected dose, and potential fortoxicity.Just as quantum dots are finding increasing applications forimaging of small-animal models (31-33), so should Raman nanoparticles lead to increasing applications without the limitationsfaced by conventional fluorescence imaging. Furthermore, the highsensitivity associated with our modified Raman microscope inconjunction with SERS nanoparticles (8.125 pM) represents animportant advantage over the sensitivity of conventional fluoroscopy imaging devices in conjunction with quantum dots [=11 nM,using IVIS (Xenogen) and Maestro (CRi, Inc.) imaging systems(unpnblished data)]. Additional comparisons between quantumdots and Raman nanoparticles should help to forther demonstratethe trne advantages of each technique. Although no single molecular imaging strategy (including Raman) isoptimal for allbiologicalmodels, the use of the current Raman imaging strategy along withexisting optical and nonoptical strategies should help expand theavailable toolbox for the field of molecular imaging. In addition, weare developing new Raman spectroscopy instrumentation for dedicated imaging of small living subjects, which should lead to fasterimage acquisition times; the potential to estimate signal depth; and,eventually, tomographic imaging. Further optimized instrumentation and more studies with various Raman nanoparticles shouldhelp to markedly expand the use of Raman imaging of small livingsubjects and should also impact newer clinical imaging strategies.The current work sets the foundation for future studies andsupports continued investigation of Raman imaging of livingsubjects.

MethodsRaman Imaging Setup. The Renishaw InViaRaman microscope. shown in Fig.51,consisted of a semiconductor diode near-infrared laser that operates at 785 nmand delivers 60 mWto the sample. Light was guided through a collimator ontoa series of mirrors that focused the light through an open field 12X microscopelens.The mouse was illuminated with the laser beam. Light from the illuminatedspot was collected with a lens and sent through a monochromator. Rayleighscattering close to the laser line was filtered through an edge filter. The remaining inelastic (Raman) scattered light was then focused through a slit (100-,umwidth) and dispersed by a diffraction grating (600 grooves per millimeter) ontoa ((D detector (deep depletion, Pt-cooled to -70°(, with a size of 576 by 384pixels; each pixel size is 22 x 22 ,urn), which then sends the detected Ramanspectra to a workstation for further processing.

SERS and SWNT Nanoparticles. SERS active nanoparticles (Nanoplex Biotags;Oxonica) consisted of gold nanoparticles covered with a layer of Raman-activematerial and coated with glass. For more details on the structures of these5ERS nanoparticles, please see Fig. 511 and 51Text. The SWNTS were providedby H. Oai (Stanford University, Stanford, CA) and had dimensions of a fewnanometers in diameter and =200 nm in length (Fig. 51c).

Animal Experiments. Female a-week-old nude mice (Charles RiverLaboratories)were used for all Raman spectroscope studies. All procedures performed on theanimals were approved bythe Stanford UniversityInstitutional Animal Care and

Keren et al.

Use Committee and were within the guidelines of humane care of laboratoryanimals.

Animal Injections. Four mice were s.c. injected with 13 fmol of SERS nanopartlcles in a 20-,u1 volume, using a 26 gauge needle. Four separate mice wereinjected via tail-vein with 260 fmol of SERS nanopertfcles in a 200-,u1 volume,using a 26-gauge needle. Four additional mice were injected via tail-vein with60 pmol of SWNTs in a 200-,u1 volume, using a 26 gauge needle.

Mouse Tumor Model. U87MG human glioblastoma (American Type CultureCollection) were cultured under standard conditions. The U87MG tumor modelswere generated by a s.c.injection of 20 x 106 cells in 200 J.t>l of PBS near the rightshoulder of the mice. Six female nude mice were inoculated with a s.c.injectionnear the right shoulder. When the tumor volume reached =200 mm', three micewere tail-vein injected with RGD SWNTs (experimental group), and three micewere injected with plain SWNTs (control group). In vivo Raman spectroscopicmeasurements were taken ofthe tumor site in living mice 24 h after IV injectionto evaluate accumulation of RGDversus plain SWNTsin the tumor.

Preparation and Conjugation of Pegylated SWNTs with RGD. Single-wall nanotubes were prepared as described by Liu et al. A detailed description-of RGDchemistry conjugation is provided in ref. 10. The SWNTwere provided by H. Dai

and have dimensions of a few nanometers in diameter and =200 nm in lengthyielding a concentration of 300 nM and a molecular massof 170 kDa.

1. Blasberg RG(2003) Molecular imaging and cancer. Mo! Cancer Ther 2:335-343.2. Gambhir 55(2002) Molecular imaging of cancer with positron emission tomography. Nat

RevCancer2:683-693.3. Massoud TF, Gambhir 55 (2003) Molecular imaging in liVingsubjects: Seeing funda

mental biological processes in a new light. GenesDev 17:545-580.4. Weissleder R(2006) Molecular imaging in cancer. Science312:1168-1171.5. Lee 5, eta/. (2007) Biological imaging of HEK293cells expressing PLCgamma1 using

surface-enhanced Raman microscopy. Anal Chern 79:916-922.6. Faulds K, Barbagallo RP,Keer Jf", Smith WE,Graham D(2004) 5ERRS as a more sensitive

technique for the detection of labelled oligonucleotides compared to fluorescence.Analyst 129:567-568.

7. Nie 5, Emory 5R (1997) Probing single molecules and single nancpartkles by surfaceenhanced raman scattering. Science275:1102-1106.

8. Fleischmann M, Hendra PJ,Mcquillan AJ (1974) Raman spectra of pyridine adsorbed ata sliver electrode. Chern PhysLett 26:163-166.

9. Jorio A, Saito R, Dresselhaus G, Dresselhaus MS(2004) Determination of nanotubesproperties by Raman spectroscopy. Phi/os Trans Roy Soc London Ser A 362:23112336.

10. Liu Z, et et. (2007) In vivo blodlstrtbutton and highly efficient tumour targeting ofcarbon nanotubes in mice. Nature Nanotech 2:47-52.

11. Lacerda L,Bianco A, Prato M, Kostarelos K(2006) Carbon nanotubes as nanomedicines:From toXicology to pharmacology. Adv Drug DeHvRev 58:1460-1470.

12. Mulvaney SP, Musick MD, Keating CD, Natan MJ (2003) Glass-coated, analyte-taggednanoparticles: A new tagging system based on detection with surface-enhancedRaman scattering. Langmuir 19:4784-4790.

13. Yang R, et al. (2006) Single-wailed carbon nanotubes-medtated in vivo and in vitrodelivery of siRNAinto antigen-presenting cells. Gene Ther 13:1714-1723.

14. McDevitt MR, et al. (2007) Tumor targeting with antibody-functionalized, radiolabeled carbon nanotubes. J Nucl Med 48:1180-1189.

15. Popielarski SR, Hu-LJeskovan5, French 5W, Triche TJ, Davis ME(2005) A nanopartlclebased model delivery system to guide the rational design of gene delivery to the liver.2. In vitro and in vivo uptake results. Bioconjugate chemistry 16:1071-1080.

16. Meyer A, Auernheimer J, Modlinger A, Kessler H (2006) Targeting RGD recognizinglnteqrlns: Drug development, biomaterial research, tumor imaging and targeting. CurrPharm Des 12:2723-2747.

17. Troy T,Jekic-McMullen D, Sambucetti L,Rice B(2004) Quantitative comparison of thesensitivity of detection offiuorescent and bioluminescent reporters in animal models.Molecular imaging 3:9-23.

Keren et el.

Raman Spectroscopic Imaging in Living Mice. Raman measurements wereperformed with a Renishaw microscope system. A semiconductor diode nearinfrared laser operating at A '" 785 nm was used asthe excitation source with alaser power of 60 mW measured at the surface of the mouse's skin (exposurelimits in 51 Text). Raman images were obtained by using a Raman point-mappingmethod. Acomputer-controlledx-ytranslation stage was used to raster-scanthemouse, creating a spectral image by measuring the Raman spectrum of eachindividual pixel in the area of interest with 500-pm, 750-Mm, or 1-mm step sizes.Integration times of3 sper step were usedto acquire our Raman maps (see51 Textfor more details). The objective lens usedwas a 12x open field in a dimly lit room.

Quantitative Spectral Analysis. The direct classical least squares method, alsocalled linear un-mixing and K-matrix methods, was used in this work toperform a quantitative analysis of Raman spectroscopy (34, 35). More detailsare provided in 51 Text.

ACKNOWLEDGMENTS. We thank Drs.Bill Doering, Glenn Davis, Ian Walton andDavid Guagliardo at Oxonica for their assistance with Nanoplex biotags; Dr.Zhenhuan Chi (Rensihaw) and Dr. Timothy Doyle (Stanford Small Anlmallmaqing) for their support. This work was funded in part by National Cancer InstituteCenter of Cancer Nanotechnology Excellence Grant U54 CA119367, NationalInstitute of Biomedical Imaging and BioEngineering Bioengineering ResearchPartnership Grant 5-R01-EBB000312, and ICMIC P50 CA114747 (to S5.G.) andNational Institutes of Health Training Grant 132 CA09695-15 Advanced Tech-niques for Cancer Imaging (to C.Z.). .

18. SOUZij GR, et el. (2006) In vivo detection of gold-imidazole self-assembly complexes:NIR-5ER5 signal reporters. Anal Chern 78:6232-6237.

19. 5itjtsema NM,Duindam JJ, Puppels GJ, Otto C,Greve JM (1996) Imaging with extrinsicRaman labels. App! Spec 50:545-551.

20. Moskovits M (2005) Surface-enhanced Raman spectroscopy: A brief retrospective. JRaman Spectrosc 36:485-496.

21. Driskell JD, et al. (2005) Low-level detection of viral pathogens by a surface-enhancedRaman scattering based immunoassay. Anal Chern 77:6147-6154.

22. Stuart DA, et al. (2006) In vivo glucose measurement by surface-enhanced Ramanspectroscopy. Anal Chern 78:7211-7215.

23. Choo-Smith LP,eta!. (2002) Medical applications of Raman spectroscopy: From proofof principle to clinical implementation. Biopolymers 67:1-9.

24. Elkje N5,Aizawa K, Ozaki Y (2005) Vibrational spectroscopy for molecular characterisation and diagnosisof benign, premalignant and malignant skin tumours. BiotechnolAnnu Rev 11:191-225.

25. Hanlon EB, et al. (2000) Prospects for in vivo Raman spectroscopy. Phys Med Bio!45:Rl-R59.

26. Haka AS,etal. (2005) Diagnosing breast cancer by using Raman spectroscopy. ProcNatlAcad Sci USA 102:12371-12376.

27. Buschman HP, et et. (2001) Diagnosis of human coronary atherosclerosis by morphology-based Raman spectroscopy. Cardiovasc Patho/1 0:59-68.

28. MotzJT, etal. (2004) Optical fiber probe for biomedical Raman spectroscopy.ApplOpt43:542-554.

29. Motz Jr, et al. (2005) Real-time Raman system for in vivo disease diagnosis. J BiomedOpt 10:031113.

30. Schipper Ml, et al. (2007) microPET-based biodistribution of quantum dots in livingmice. J Nucl Med 48:1511-1518.

31. Gao X, Chung LW, Nie 5 (2007) Quantum dots for in vivo molecular and cellularimaging. Methods Mol Bioi 374:135-146.

32. So MK,Xu C, Loening AM, Gambhir SS,Rao J (2006) Self-illuminating quantum dotconjugates for in vivo imaging. Nat Biotechno/24:339-343.

33. Tada H, Higuchi H,Wanatabe TM, Ohuchi N(2007) In vivo real-time tracking of singlequantum dots conjugated with monoclonal anti-HER2 antibody in tumors of mice.Cancer Res67:1138-1144.

34. Haaland OM, Easterling RG (1980) Improved sensitivity of infrared spectroscopy by theapplication of least squares methods. Appl Spec 34:539-548.

35. Pelletier MJ (2003) Quantitative analysis using Raman spectroscopy. Appl Sped57:20A-42A.

PNAS I April 15, 2008 I vol. lOS I no. 15 I 5849

Attachment Le

Self-Sorted Nanotube Networks onPolymer Dielectrics for Low-VoltageThin-Film TransistorsMark E. Roberts, Melburne C. LeMieux, Anatoliy N. Sokolov, and Zhenan 8ao*

Department of Chemical Engineering, Stanford University, Stauffer III,381 North-South Mall, Stanford, California 94305-5025

Received January 27, 2009; Revised Manuscript Received May 8, 2009

ABSTRACT

NANDLETTERS

2009Vol. 9, No.72526-2531

Recent exploitations of the superior mechanical and electronic properties of carbon nanotubes (CNTs) have led to exciting opportunities inlow-cost, high performance, carbon-based electronics. Inthis report, low-voltage thin-film transistors withaligned, semiconducting CNT networksare fabricated on a chemically modified polymer gate dielectric using both rigid and flexible substrates. The multifunctional polymer servesasa thin, flexible gate dielectric film, affords lowoperating voltages, and provides a platform for chemical functionalization. The introductionof amine functionality to thedielectric surface leads to theadsorption of a network enriched with semiconducting CNTs withtunable densityfrom spincoating a bulksolution of unsorted CNTs. The composition of thedeposited CNT networks is verified withRaman spectroscopy andelectrical characterization, For transistors at operating biases below 1 V, we observe an effective device mobility as highas 13.4 cm'Ns, asubthreshold swing as low as 130 mV/dec, and typical on-off ratios of greater than 1,000. This demonstration of high performance CNTthin-film transistors operating atvoltages below 1 Vand deposited using solution methods onpolymeric and flexible substrates is animportantstep toward the realization of low-cost flexible electronics.

Single-wall carbon nanotubes (SWNTs) are ideal componentsfor flexible, low-power electronic devices owing to theirsuperior electrical and mechanical properties, as well asenvironmental stability.l-' Their excellent charge-transportproperties are particularly snitable for applications relatingto display drivers, photovoltaics, and circuits."? Additionally,their extreme environmental sensitivity, intrinsic of a onedimensional electronic material consisting of only surfaceatoms, makes SWNTs ideal active components for biologicaland chemical sensorss" as charge transport properties aredirectly influenced by molecular adsorbates. To be costeffective, new macroelectronic applications including disposable sensors, electronic skins, and bendable large displaysrequire nanoscale, thin semiconducting films uniformlydeposited over large areas in processes compatible withflexible substrates, such as comparatively rough polymericplatforms.

Despite rapid progress in the field, directed, reproducibleassembly and orientation of SWNTs via solution depositionover large areas remain an unsolved issue. While isolated(single-tube) SWNT devices have been demonstrated as theactive component for transistors," this is not practical forlarge-scale integration. Recently developed random nanotubenetworks.t''!' easily deposited from solution, offer an attractive alternative. Still, the random nature of SWNTs in terms

*To whom correspondence should be addressed. E-mail: [email protected]. Phone: (650) 723-2419.

1O.1021/n1900287p CCC: $40.75 © 2009 American Chemical SocietyPublished on Web 06/0512009

of chirality, and variations in tube density, alignment, andbundling of SWNT networks results in poor device performance and reproducibility, inhibiting fundamental structurelproperty relationship studies. To date, alignment/densitycontrol has been addressed through dielectrophoresis,'? gasflow," and absorption (or evaporation) onto affinity pat,terns,13.14 while chirality purification has been achievedthrough simple metallic "burnoff"," ultracentrifugation.l'v'?and microfluidics combined with dielectrophoresis methods"among others. However, these are either bulk (solution) phaseseparations involving several steps or device level postdeposilion techniques similarly not suitable for direct integrationinto devices.

We have recently reported on a process that can achievechirality selection and alignment in a one-step solutiondeposition process through controlling substrate surfacechemistry.19 This self-sorting method results in a moderatelyaligned SWNT submonolayer network enriched with semiconducting, unbundled nanotubes of reproducible topology(density, alignment, and chirality) from device to device.Although demonstrated on silicon with a SiO, dielectric,'?our envisioned low-cost flexible electronics will requireoperation at lower voltages and a dielectric material thatexhibits a high degree of bendability.i" Polymeric materialsare an excellent cboice as they provide tunable surfacechemistry, morphology, can undergo high elastic deformation, and can planarize or smooth rough substrates. Impor-

A Spin-coatPVPfitms

Figure 1. Schematic of the TFT fabrication with self-sorted SWNT networks on thin polymer dielectric layers and the resulting topologyof sortedandunsorted networks. (A) The SWNT solution depositionprocess, which involves spin coating 10 .ug/mL in NMP at 3600-4000rpm on the cross-linked PVP layer; (B) AFM image (15 x 15 fl-m2 , z-scale = 5 urn) of a typical sortedSWNT networkwith a maximumsurface coverage area of ,....,10% on APTES + PVP surfaces showing the radial alignment and uniformity; (C) AFM image (2 x 2 jim2

,

z-scale= 5 nrn) of the sortedSWNT network in which densitywas controlled at depositionto be at thepercolation threshold ("-'5% surfacecoverage); (D) molecular structure of the cross-linkedPVP withIIDA; (E) AFM image at 10 x 10 Jlm2 (z-scale = 20 urn) of an unsortedSWNT networkwith as surfacecoverage of .......20%;and(F) a similaranalysis as in panel E, butat 5 x 5 p.m2 (z-scale = 20 nm) highlightingthe excessive bundling in the unsorted surfaces.

tantly, recent work has shown these materials offer highcapacitance, desirable of a gate dielectric material,2!,22combined with suitable hydrophobicity that allows operationin ambient and aqueous environments,22,23 a key for autonomous sensing and other applications.

The formation of well-ordered SWNT arrays, precedingactual working SWNT thin-film transistors (SWNT TFTs),on polymer surfaces is not a trivial task. Previously, this hasbeen accomplished with a postdeposition transfer printingstep of highly aligned CVD grown SWNT films onto plasticsheets, resulting in on/off ratios of approximately 103 usingspecial channel geometry.V" The forces involved in thismethod may result in cracks within the polymer film therebydiminishing the dielectric properties.' Single-step fabricationof SWNT TFTs directly from solution on polymer substrateshas been achieved by Snow' and Gruner, II yielding on/offratios as high as 100 with no alignment or density control.

In this report, we demonstrate high performance TFTs withsemiconducting SWNT (scSWNT) networks on a chemicallymodified polymer gate dielectric layer with operating biasesbelow I V and excellent cycling stability in water. Thedielectric layers are deposited on both a rigid Si, as well asa transparent, flexible ITOIPET substrate. Specifically, thesingle-step construction of well-ordered SWNT networktransistors with tunable topology on polymer dielectricsurfaces has been accomplished. Our solution depositionmethod is compatible with polymeric dielectric layers andrepresents a major step toward enabling SWNT networkintegration with flexible, low power electronics. Importantly,we show that the overall SWNT network topology (alignment, density), controlled by the surface modification and

Nano Lett., Vol. 9, No.7, 2009

the SWNT deposition conditions, along with chiralitypurification in the SWNT network, has a critical impact onthe transistor characteristics.

SWNT-based TFrs were fabricated on insulating poly(4vinylphenol) (PVP) films cross-linked with 4,4' -Ihexafluoroisopropylidene)diphthalic anhydride (HDA).22 Briefly, thepolymer dielectrics were deposited via spin coating at 7000rpm (Headway Research, Inc.) from a solution of 20 mg!mL PVP, 2 mg/mL HDA, and I ,uLlmL triethylamine inpropylene glycol monomethyl ether acetate (pGMEA). Thelayers were deposited and characterized with a thickness of25 nm on rigid silicon substrates or 35 nm on flexiblesubstrates (indium tin oxide on polyethylene terephthalate(ITOIPET» (Figure I). The resulting capacitance of the 25and 35 nm films is 165 and 115 nF/cm2, respectively. Acomplete description of the experimental details is availablein the Supporting Information,

To achieve the desired enrichment of semiconductingSWNTs (scSWNTs), the PVP surface was modified withamine functionality prior to SWNT deposition. While theresidual hydroxyl groups on the polymer surface may be usedto introduce amine groups, further oxidation of the polymersurface through a gentle UV-ozone exposure (3 min) leadsto increased grafted amine density. Ozone treatment resultedin a decrease in contact angle to <10°, much lower than fororiginal PVP (62.4 ± 1.1) and similar to that of treated Si02.The substrates were placed in a solution of aminopropyltriethoxysilane (APTES) (Gelest, silanes were distilled priorto use) in anhydrous toluene for 1 h inside a dry N2 glovebox,followed by rinsing in anhydrous toluene. The samples weresubsequently annealed in a vacuum oven at 80°C for a period

2527

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·1 Raman ~ft em·!

14150

1500 15'50 1600Ra~n Shift em·1

1650 17

Figure 2. Raman spectroscopy using 1.96 eV excitation energy, which is resonant with metSWNT (EMl1 ) and scSWNT (£833) in a nearly

50/50 ratio for the CNTs used in this study. (A) RBM analysis from averaged spectra (orange curve) from 12-point mapping (see SupportingInformation) on sorted surfaces showing predominantly semiconducting tubes, which exhibit strong £833 (red shaded region) deconvolutedpeaks (red curves) at 170 and 155 cm", and a smaller one at 146 cm". (B) RBM region of the random unsorted SWNT network showinga broader overall average spectrum (black curve) and several additional modes (blue lines) than was observed from the sorted networks,including four additional EMU peaks (blue shaded region). (C) Comparison of the tangential mode spectra of the sorted (orange) andunsorted (black) CNT networks.

of 10 min. Following the chemical modification with APTES,we observed an increase in thickness from 25.1 ± 0.2 to26.8 ± 2.2 nm and a change in contact angle to 43.5 ± 1.0°with no appreciable change in atomic force microscopy(AFM) measured surface roughness (Supporting Information,Table SI-1). We found that the optimal UV-ozone treatmenttime of 3 min was critical, as longer times resulted in roughersurfaces with reduced SWNT adsorption and shorter timesled to a poor quality amine layer (Supporting Information,Figure SI-1). However, the ozone and subsequent aminetreatment had no observable effect on the PVP capacitanceand surface roughness with gate leakage remaining low (seebelow). This is important for several reasons. A major keyfor the deposition of a self-sorted, debundled SWNT networkis a smooth surface,'? and partial delamination or pinholeformation in the polymer dielectric would adversely affectdevice performance and/or charge transport in the overlayingsemiconductor layer.26,27

Sorted SWNT networks are deposited via spin-coating ontop of functionalized PVP as discussed in SupportingInformation. Briefly, a solution of 10 flg/rnL of purifiednanotubes in I-methyl-2-pyrrolidone (NMP) (50 flL per ern'substrate)was deposited on a substrate rotating at 3600-4000rpm prior to solution addition. Atomic force micrographs ofthe sorted and aligned nanotube networks on PVP-HDAsurfaces are shown in Figure 1, revealing no damage ordelamination of the underlying polymer layer. The SWNTnetwork deposited on the APTES/PVP-HDNSiO, substrateshowed high uniformity from AFM analysis at differentlength scales. The polymer film roughness of 0.3 nm iscomparable to that of the underlying silicon oxide. Becauseof the increased surface roughness of the flexible ITOIPETsubstrates, the SWNT networks exhibited a reduction indensity and alignment (Supporting Information, Figure SI1) when deposited according to the conditions optimized forrigid silicon substrates. As a control, unsorted SWNTnetworks were also deposited on the APTES-modified PVPsubstrates; however, the solution was allowed to settle on

2528

the surface for an extended period of time (60 s) before thesubstrate was spun dry. Figure IE shows the topology ofthe unsorted networks on the APTESIPVP-HDNSiO, substrates, while Figure IF highlights the excessive bundlingfound on these networks. In contrast, Figure IB shows thelarge-scale topology of the sorted SWNT networks tohighlight the degree of alignment and Figure 1C shows anenhanced magnification to illustrate the clear lack of bundlingwithin the networks.

To verify the enrichment of scSWNTs through the selfsorting process on amine-modified PVP, the chirality of theadsorbed SWNT network was studied using Raman spectroscopy.fl-Rarnan mappingwasconductedover largeareasof eachsamplesurfacewithinthe transistorchannelsto identify the typeof adsorbed tubes" using 1.96 eV excitation energy. Theobserved resonant radial breathing modes (RBMs) were correlatedwith chiralityusing a diameterlRBMrelationdevelopedfor isolated SWNTs on SiO,.29 In particular, the 1.96 eV lineis resonant with metSWNT (£,'"11) and scSWNT (F!'33) in anearly 50/50 ratio for the arc-discharge tubes used in this study(mean diameter 1.4 ± 0.6 nm from AFM) and is best suitedfor separation analysis. RBM analysis (Figure 2A,B) ofaveraged spectra taken from sorted SWNT networks revealspredominantly scSWNTs exhibiting strong F!'33 peaks at 170and 155 ern-I, a smaller peak at 146 cm", and perhaps verysmall EM

11 peaks at 195 ern"! and 200 cm" that were notsignificant from noise, and thus were not fitted.

On the unsorted samples, there is a much stronger metalliccontribution and an overall higher number of RBM peaks(Figure 2B). At least four peaks (183, 192, 200, and 210cm") are attributed to metSWNTs, with a much largerintegrated area of the RBM region in the [!MIl wavenumbers,as compared with the sorted networks. The enhancedBreit-Wigner-Fano (BWF) line shape broadening of theG- tangential band observed in the unsorted SWNT networkis also indicative of a larger fraction of metSWNTs whencompared to the sorted SWNT networks (Figure 2C). Ingeneral, the RBM region is characterized by a much broader

Nano ten., Vol. 9, No.7, 2009

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Figure 4. SWNT TFT operation in aqueous conditions. (A) Ios- Vos of a TFT with a sorted SWNT network on 25 nm PVP-HDA in water;(B) los-Va for 1Ff in panel A with a Pcff = 0.52 cm2Ns, VT = 0.38 V, and an onloff ratio of 102; and (C) IDs- Va for TIT in panel Aafter removing the device from the water and drying under a stream of dry, compressed air. (The dashed black line represents the measurementin ambient before exposure to water.)

Figure 3. Electrical characteristics of CNT networks on APTESIPVP-HDA substrates. (A) 1D8 - VDS of a TFT with a sorted SWNT networkon "25 nrn PVP-HDA on silicon; (B) IDS-VO for the TFT in panel A with a #eff = 13.4 cm2Ns, V T = 0.67 V and an on/off ratio of 1.6 x1(}3; (C) histogram of device mobilities for sorted SWNT networks on 25 run PVP-IIDA on Si substrates; (D) histogram of on/off ratiosfor devices shown in panel C; (E) IDs- Vos for a TFf with an unsorted SWNT network; (F) 10s-Va for TIT in E; (G) 10s-Vos of a TFTwith a sorted SWNT network on ITOIPET substrates with 35 nm PVP-HDA; and (H) los-Va on the TFT in G with a Peff= 1.0 cm2Ns,

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with the electrode, and thus represents a lower limit of deviceperformance. The devices exhibited an average thresholdvoltage (VT ) of 0.31 V and an average on/off ratio of 1.75x 103 with subthreshold slopes as low as 130 mV/dec andgate leakage comparable to those previously reported.F-"In some cases, individual devices showed an effectivemobility as high as 13.4 cm2Ns. Histograms of the devicemobility and on/off ratio are shown in Figure 3C,D. Thespread in performance can be attributed to slight variationsin alignment and density resulting from subtle variations inbatch-to-batch (~20 batches) device fabrication as well assome minimal on-chip variations owing to a radial densitygradient associated with our assembly process as describedearlier. [9 It is important to consider the subthreshold valuewhen characterizing plastic dielectrics to maintain highperformance at low operating biases, and this value (~130

mV/dec)is comparable with the best values reported for CNTnctworks.>? or organic transistors."

The device characteristics on the ITO/PET substrateswere inferior to those on rigid silicon owing to the lowerSWNT density. The lower density may be attributed tothe increased roughness of the ITOIPET surface (5-10nm) when compared to the PVPISi (0.5-1 nm). This

2529

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peak distribution compared to sorted networks with sharppeaks. This is attributed to both the presence of a largernumber of tube chiralities and increased bundling.30,31 Indeed,this result combined with AFM analysis demonstrates thatthe amine surface on PVP is effective at sorting chiralityand controlling bundling of the SWNTs, and that partiallyaligned nanotube films can be efficiently deposited onpolymer surfaces.

Top-contact thin-film transistors with a channel length (L)= 50 usn, channel width (W) = I mm were completed bythermally evaporating gold electrodes" on top of the SWNTnetworks. The transfer characteristics (source-drain current,IDS, vs gate voltage, VG) ofTFTs with sorted (71 devices onSi and 10 devices on ITOIPET) and unsorted (lOon Si)SWNT networks were measured with a draio voltage, VDS,

of -I V and a gate voltage sweep from 0.5 V (or I V) to-I V with a VG step of 0.02 V (Keithley 4200-SCSSemiconductor Parameter Analyzer). From the transfercurves of the TFTs on Si substrates using the electrode WIL= 20, we determined an average effective device mobility(fl'ff) of 2.65 cm2Ns~ It should be noted that this effectivemobility is calculated from the WIL of the electrodes withoutcorrecting for the W based on the number of CNTs in contact

NanD Lett., Vol. 9, No.7, 2009

results in an effective device mobility of 0.73 cm'Ns, athreshold voltage of 0.30 V, and an on/off ratio greaterthan 3 x 10' with a subthreshold swing of 390 mV/decThese values still represent the best performance for CNTnetworks assembled directly from solution on flexibleplastic surfaces. The electrical characteristics of the sortedSWNT TFTs on silicon and ITO/PET substrates are shownin Figure 3G,H. As suggested by the Raman analysis, asignificantly higher degree of metSWNTs is adsorbed inunsorted networks compared to sorted networks. Thisobservation is confirmed with electrical characterization,showing that the SWNT films with unsorted, randomnetworks are dominated by metallic pathways and aregenerally conductive with much lower on-off ratios «2)and higher los (~1O-5 A). The output (Ios- Vos) and transfercharacteristics (Ios-Vol for the TFTs with unsorted networksare also shown in Figure 3E,F.

The robustness and durability of our CNT-based TFTs isdemonstrated through operation under aqueous conditions.During the electrical characterization in water, the Vos andVo were reduced to -0.6 V to keep the parallel ionic currentthrough the solution to a minimum." The 1FT characteristicsin water were calculated for Vos = Va = -0.6 V resultingin afl.,ff, VT , and an on/off ratio of 0.52 cm2Ns, 0.38 V, and102, respectively (compared to 1.2 cm2Ns, 0.24 V, and 1.1x 103 for the same device in ambient). The output andtransfer characteristics of a typical sorted SWNT networktransistor under aqueous conditions are shown in Figure4A,B, respectively. Notably, the decrease in on/off ratio isa result of both a higher off-current, owing to ionicconduction through the solution, and the doping effect ofwater on semiconducting nanotubes." Although limited bya low on/off ratio of around 102

, the SWNT TFTs were stablein water for over 400 electrical cycles of Va = 0.6 to -0.6V with Vee = -0.6 V. The water-stability is likely to beeven higher as we have demonstrated an extended stabilityof 104 cycles for less stable OTFTs in water." Importantly,upon removal from water, the performance of the SWNTTFTs returned to the previous values measured in air,indicating the absence of degradation of the SWNTs and thedielectric layer (Figure 4C). The ability to operate in wateris critical for futnre applications to electronic sensors capableof detection in aqueous or biologically complex media.

In summary, recent advances in SWNT processing haveled to the ability to deposit CNTs in a self-sorting, alignedmanner with the potential for large-scale manufacturing. Inthis report, we have expanded upon these achievements byreducing the power requirements while maintaining theelectronic performance and demonstrating compatibility witha flexible substrate. These robust networks also demonstratedstable performance within aqueous media with no signs ofdegradation. We showed that using a controlled solutiondeposition method on amine-modified polymer dielectricsleads to an aligned, uniform network of carbon nanotubescontrasting the random networks generated through standarddeposition methods. fl.-Raman spectroscopy was used to showenrichment in semiconducting nanotubes within the aligned,sorted networks, corroborating the superior transistor char-

2530

acteristics. The fact that the self-sorting approach can be. applied to polymer dielectrics through a room temperature,

solution deposition method represents a major step towardthe realization of inexpensive SWNT electronic devices ontransparent, flexible substrates. Their particular stability inambient and aqueous conditions is especially attractive forfuture sensor applications based on SWNT TFTs (i.e.,autonomous underwater detection systems).

Acknowledgment. M.E.R. acknowledges partial supportfrom the NASA GSRP fellowship. M.C.L. acknowledgessupport from the Intelligence Community Postdoctoral Fellowship Program. Z.B. acknowledges financial support fromNSF-EXP sensor program, the Stanford Center for PolymericInterfaces and Macromolecular Assemblies (NSF-CenterMRSEC), and the Sloan Research Fellowship.

Supporting Information Available: Materials, methods,and additional film properties. This material is available freeof charge via the Internet at http://pubs.acs.org.

References(1) Baughman, R. H.; Zakhidov, A. A.; de Heer, W. A. Science 2002,

297 (5582).787-792.(2) O'Connell, M. Carbon nanotubes: properties and applications; CRC

Taylor & Francis: Boca Raton, FL, 2006; p 319.(3) Avouris, P.; Chen, Z.; Perebeinos, V. Nat. Nanotechnol. 2007, 2 (10),

605-615.(4) Rowell, M. W.; Topinka, M. A; McGehee, M. D.; Prall, H.-J.; Dennler,

G.; Sariciftci, N. S.; Hu, L.; Gruner, G. Appl. Phys. Lett. 2006, 88(23), 233506.

(5) Cao, Q.; Kim, H.-s.; Pimparkar, N.; Kulkarni, J. P.; Wang, c, Shim,M.; Roy, K.; Alam, M. A; Rogers, J. A Nature 2008, 454 (7203),495-500.

(6) Allen, B. L.; Kichambare, P, D.; Star, A. Adv. Mater. 2007, 19 (11),1439-1451.

(7) Snow, E. S.; Perkins, F. K.; Houser, E. J.; Badescu, S. c., Reinecke,T. L. Science 2005, 307 (5717), 1942-1945.

(8) Kauffman, D. R.; Star, A Chern. Soc. Rev. 2008,37 (6), 1197-1206,(9) Auvray, S.; Derycke, V.; Goffman, M.; Filoramo, A.; Jost, 0.;

Bourgoin, J.-P. Nano Lett. 2005, 5 (3), 451--455.(10) Lay, M. D.; Novak, J. P.; Snow, E. S. Nano Lett. 2004,4 (4), 603

606.(11) Artukovic, E.; Kaempgen, M.; Hecht, D. S.; Roth, S.; Gruner, G. Nano

Lett. 2005,5 (4), 757-760.(12) Krupke, R.; Hennrich, F.; Lohneysen, H. V.; Kappes, M. M. Science

2003,301 (5631), 344-347.(13) Sharma, R.; Lee, C. Y.; Choi, J. H.; Chen, K.; Strano, M. S. Nano

Lett. 2007, 7 (9), 2693-2700.(14) Wang, Y.; Maspoch, D.; Zou, S.; Schatz, G. c.. Smalley, R. E.; Mirkin,

C. A Proc. Natl, Acad. Sci. U,S.A. 2006, 103 (7), 2026-2031.(15) Collins, P, G.; Arnold, M. S.; Avouris, P, Science 2001, 292 (5517),

706-709.(16) Arnold, M. S.; Green, A. A.; Hulvat, J. F.; Stupp, S. I.; Hersam, M. C.

Nat. Nanotechnol. 2006, 1, 60.(17) Engel, M.; Small, J. P.; Steiner, M.; Freitag, M.; Green, A A; Hersam,

M. c, Avouris, P. ACS Nano 2008, 2 (12), 2445-2452,(18) Shin, D. H.; Kim, J.~E.; Shim, H. C.; Song, J.-W.; Yoon, J.-H.; Kim,

J.; Jeong, S.; Kang, J.; Baile, S.; Han, C.-S. Nano Lett. 2008, 8 (12),4380-4385.

(19) LeMieux, M. C.; Roberts, M.; Bannan, S.; Jin, Y. W.; Kim, J. M.;Bao, Z. Science 2008, 321 (5885), 101-104.

(20) Forrest, S. R. Nature 2004, 428 (6986), 911-918.(21) Kim, c.. Wang, Z.; Choi, H.-J.; Ha, Y.-G.; Facchetti, A; Marks, T. J.

J. Am. Chem. Soc. 2008, 130 (21), 6867-6878.(22) Roberts, M. E.; Mannsfeld, S, C. B.; Queralto, N.; Reese, C.; Locklin,

J.; Knoll, W.; Baa, Z. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 12134,(23) Roberts, M. E.; Queralto, N.; Mannsfeld, S. C. B.; Reinecke, B, N.;

Knoll, W.; Baa, Z. Chern. Mater. [Online early access]. DOl: lO.102l!cm900637p.

(24) Cao, Q.; Hur, S. H.; Zhu, Z. T.; Sun, Y. G.; Wang, C. J.; Meitl, M. A.;Shim, M.; Rogers, J. A Adv. Mater. 2006, 18 (3), 304-309.

NanD Len., Vol. 9, No.7, 2009

(25) Detailed experimental details are available in Supporting Information.(26) Kim, C.; Facchetti, A.; Marks, T. J. Science 2007, 3]8 (5847), 76

80.(27) Tello, M.; Chiesa, M.; Duffy, C. M.; Sirringhaus, H. Adv. Funct. Mater.

2008,18 (24), 3907-3913.(28) Iorio, A; Saito, R; Hafner, J. R; Lieber, C. M.; Hunter, M.; McClure,

T.; Dresselhaus, G.; Dresselhaus, M. S. Phys. Rev. Lett. 2001, 86 (6),I1l8.

(29) Jorio, A.; Fantini, c.: Pimenta, M. A.; Heller, D. A; Strano, M. S.;Dresselhaus, M. S.; Oyama, Y.; Jiang, J.; Saito, R Appl. Phys. Lett.2006, 88 (2), 023109.

Nano Leff., Vol. 9, No.7, 2009

(30) Rao, A M.; Chen, J.; Richter, E.; Schlecht, D.; Eklund, P. c.,Haddon,R. c., Venkateswaran, D. D.; Kwon, Y. K.; Tomanek, D. Phys. Rev.Lett. 2001, 86 (17),3895.

(31) Heller, D. A; Barone, P. W.; Swanson.J. P.; Mayrhofer, R. M.; Strano,M. S. J. Phys. Chern B 2004, 108 (22), 6905-<5909.

(32) Fll, Q.; Liu, J. Langmuir2005, 21 (4), ll62-1165.(33) Someya, T.; Kim, P.; Nuckolls, C. Appl. Phys. Lett. 2003,82 (14),

2338-2340.

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Attachment 2

What are Nanomaterials?

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What are Nanomaterials?

Nanomaterials are defined as materials with at least one external dimension in the size range from approximately 1-100nanometers. Nanoparticles are objects with all three external dimensions at the nanoscate-. Nanoparticles that arenaturally occurring (e.g., volcanic ash, soot from forest fires) or are the incidental byproducts of combustion processes(e.g., welding, diesel engines) are usually physically and chemically heterogeneous and often termed ultrafine particles.Engineered nanoparticles are intentionally produced and designed with very specific properties related to shape, size,surface properties and chemistry. These properties are reflected in aerosols, colloids, or powders. Often, the behavior ofnanomaterials may depend more on surface area than particle composition itself. Relative-surface area is one of theprincipal factors that enhance its reactivity, strength and electrical properties.

Engineered nanoparticles may be bought from commercial vendors or generated via experimental procedures byresearchers in the laboratory (e.g., CNTs can be produced by, laser ablation, HiPCO (high-pressure carbon monoxide, arcdischarge, and chemical vapor deposition (CVD)). Examples of engineered nanomaterials include: carbon buckeyballs orfullerenes; carbon nanotubes; metal or metal oxide nanoparticles (e.g., gold, titanium dioxide); quantum dots, amongmany others.

1 National Institute for Occupational Safety and Health (NIOSH) Approaches to Safe Nanotechnology: Managing the Health and Safety

Concerns Associated with Engineered Nanomaterials (March 2009). http://www.cdc.gov/niosh/docs/2009-125/

Back to General Principles and Practices for Working Safely with Engineered Nanomaterials

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General Principles and Practices for Working Safely with Engineered Nanomaterials Page 1 of 1

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General Principles and Practices for Working Safely with Engineered Nanomaterials

• Puroos.eo Action Items for Principal Investigators

• What are Nanomaterials?• Current Occupational Health and Safety Concerns• Guidelines for Working with Nanomaterials

a Quick GUide: Exposure Risks and Control Measures for Common Laboratory Operations InvolvingNanomaterials

• Spill Response

• Disposal• Additional Information and References• Standard Operating Procedure Template for Work with Nanomaterials

Back to Research and Laboratory Safety

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Purpose

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Purpose

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GENERAL RESEARCH a RENOVATION & j(IN~ ~

HEALHI LA80RATORY CONSTRUCTION ENVIRONMENTAL r \"ABOUT US a SAFETY SAFETY SAFETY PROGRAMS h0111e

This document provides environmental, health and safety guidance to researchers working with engineered nanomaterialsin Stanford University laboratories. It is intended to supplement the requirements of Stanford University's ChemicalHygiene Plan and its companion Laboratory Safety Toolkit which are available at:

As the scientific community continues to gather data to assess the potential health and safety risks associated withengineered nanomaterials, these gUidelines may be updated as information becomes available. Until more is known aboutthe possible risks of nanomaterials, it is prudent and appropriate to take a precautionary approach and utilize goodlaboratory safety practices when working with these materials.

Action Items for Faculty Working With or Creating Engineered Nanomaterials

1. Review this document - Stanford University's General Principles and Practices for Working Safely withEngineered Nanomaterials.

2. Instruct all research personnel in your lab to follow the work practices described in this document.3. Create Standard Operating Procedures (SOPs) for processes and experiments involving nanomaterials using

the template contained in this document. Priority for SOP development should be given to those operationswhere there is higher risk of exposure (e.g., manipulation of nanoparticles in gas stream, work with drydispersible nanoparticles). A SOP template for work with nanornaterials is available here.

Back to General Principles and practices for Working Safely with Engineered Nanomaterials

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Current Occupational Health and Safety Concerns Page 1 of 1

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Current Occupational Health and Safety Concerns

Exposure to nanomaterials may occur through inhalation, dermal contact, or ingestion depending on how personnel useand handle them. The full health effects of exposures to nanomaterials are not fully understood at this time. For example,a peer-reviewed toxicity study on carbon nanotubes (CNTs) indicated that the toxicity of nanoparticles depends on specificphysiochemical and environmental factors and thus the toxic potential of each nanoparticle needs to be evaluatedseparately [Helland et al., 2007]. Results of existing studies in animals or humans provide some basis for preliminaryestimates of areas of concern. According to the National Institute for Occupational Safety and Health (NIOSH)2 , studies todate have indicated:

• Increased toxicity of ultrafine particles or nanoparticles as compared to larger particles of similar composition.Chemical composition and other particle properties can also influence toxicity. [Oberdorester et aI., 1992, 1994a,b,2005a; uson et al., 1997; Tran et al., 1999, 2000; Brown et al., 2001; Duffin et al., 2002; Barlow et al. 2005;Maynard and Kuempel 2005; Donaldson et al. 2006].

• A greater proportion of inhaled nanoparticles will deposit in the respiratory tract as compared to larger particles.[ICRP 1994; Jaques and Kim 2000; Daigle et al. 2003; Kim and Jaques 2004.]

• Nanoparticles can cross cell membranes and interact with sub cellular structures where they have been shown tocause oxidative damage and impair function of cells in culture. [Moller et al. 2002, 2005; Li et al. 2003; Geiser et al.2005].

• Nanoparticles may be capable of penetrating healthy intact skin and translocating to other organ systems followingpenetration. [Tankenaka et at. 2001; Kreyline et al 2002; Oberd6rester et al. 2002, 2004; Semmler et at. 2004;Geiser et al. 2005.]

• Catalytic effects and fire or explosion are other hazards to consider. [Pritchard 2004].

2 National Institute for Occupational Safety and Health (NIOSH) Approaches to Safe Nanotechnology: Managing the Health and Safety

Concerns Associated with Engineered Nanomaterials (March 2009) htto:/Iwww.cdc.gov/niosh/docS/2009-125/


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Lab Safety Guidelines for Handling Nanomaterials Page I of2

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Lab Safety Guidelines for Handling Nanomaterials

Exposure standards have not been established for engineered nanoparticles in the United States or internatinally [SafeNanotechnology 2008.] Until more definitive findings are made regarding the potential health risks of handlingnanomaterials, researchers planning to work with nanomaterials must implement a combination of engineering controls,work practices, and personal protective equipment to minimize potential exposures to themselves and others. For a quickguide to the exposure risks and prudent control measures to be used for common laboratory operations involvingnanomaterials, refer to the table. It is important to consider if the nanoparticles are in an agglomerated or aggregatedform, functionallzed, suspended in liquid, or bound, as these conditions may affect the exposure potential.

1. Engineering Controls:o Use glove bags, glove boxes, fume hoods, or other containment or exhausted enclosures when there is a

potential for aerosolization, such as: handling powders; creating nanoparticles in gas phase; pouring ormixing liquid media which involves a high degree of agitation. (DO NOT use horizontal laminar flow hoods(clean benches), as these devices direct the air flow towards the worker.) Consult with EH&S if engineeringcontrols are not feasible.

o Use fume hoods or other local exhaust devices to exhaust tube furnaces and or chemical reaction vessels.

o Perform any maintenance activities, such as repair to equipment used to create nanomaterials orcleaning/replacement of dust collection systems, in fume hoods or under appropriate local exhaust.

2. Work Practices:o Selection of Nanomaterials:

• Whenever possible, handle nanomaterials in solutions or attached to substrates to minimize airbornerelease.

• Consult the Material Safety Data Sheet (MSDS), if available, or other appropriate references prior tousing a chemical or nanomaterial with which you are unfamiliar. Note: Information contained in someMSDSs may not be fully accurate and/or may be more relevant to the properties of the bulk materialrather than the nano-stze particles.

o Safety Equipment:

• Know the location and proper use of emergency equipment, such as safety showers, fire extinguishers,and fire alarms.

o Hygiene:• Do not consume or store food and beverages, or apply cosmetics where chemicals or nanomaterials are

used or stored since this practice increases the likelihood of exposure by ingestion.• Do not use mouth suction for pipetting or siphoning.• Wash hands frequently to minimize potential chemical or nanoparticle exposure through ingestion and

dermal contact.• Remove gloves when leaving the laboratory, so as not to contaminate doorknobs, or when handling

.common use objects such as phones, multiuser computers, etc.a Labeling and Signage:

• Store in a well-sealed container, preferable one that can be opened with minimal agitation of thecontents.

• Label all chemical containers with the identity of the contents (avoid abbreviations/ acronyms); includeterm "nano" in descriptor (e.q., "nano-zinc oxide particles" rather than just "zinc oxide." Hazardwarning and chemical concentration information should -also be included, if known.

• Use cautious judgment when leaving operations unattended: i) Post signs to communicate appropriatewarnings and precautions, Ii) Anticipate potential equipment and facility failures, and iii) Provideappropriate containment for accidental release of hazardous chemicals.

o Cleaning:• Wet wipe and or HEPA-vacuum work surfaces regularly.

a Transporting:• Use sealed, double-contained container when transporting nanomaterials inside or outside of the

building.

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Lab Safety Guidelines for Handling Nanomaterials Page 2 of2

o Buddy System:

• Communicate with others in the building when working alone in the laboratory; let them know whenyou arrive and leave. Avoid working alone in the laboratory when performing high-risk operations.

3. Personal Protective Equipment:o Wear gloves, lab coats, safety goggles, long pants, closed-toe shoes, and face shields, as appropriate

dependent on the nature of the materials and procedure.o If work cannot be conducted inside a fume hood or other ventilated enclosure, consult with EH&SlS

Occupational Health and Safety Program (723-0448) regarding the need for respiratory protection or otheralternative controls.

4. Training:o Ensure that researchers have both general safety training and lab-specific training relevant to the

nanomaterials and associated hazardous chemicals used in the process/experiment. See SU's LaboratoryChemical Safety Toolkit (SU Toolkit) for quidance on training(htlp:llchemtoolkit.stanford.eduIChemSafetyTralning)

0 Lab-specific training can include a review of this safety fact sheet, the relevant Material Safety Data Sheets (ifavailable), and the lab's Standard Operating Procedure for the experiment.

5. Standard Operating Procedures:o Prepare a Standard Operating Procedure (SOP) for operations Involvtnq nanomaterials. A general use

Standard Operating Procedure for working with nanomaterials is available here. The SOP should be tailored tobe specific to the proposed experimental procedure.

o Consider the hazards of the precursor materials in evaluating the process.o Special consideration should be given to the high reactivity of some nanopowders with regard to potential fire

and explosion. [Pritchard 2004].

6. Consultation:Consult with your Principal Investigator prior to procuring or working with nanomaterials. See SU's Toolkit forinformation regarding consultation and prior approvals (http://chemtoolkit.stanford.edu/PriorApprovals). Foradditional assistance, contact EH&S's Occupational Health & Safety Program at 723-0448.


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Quick Guide: Exposure Risks and Control Measures for Common Laboratory Operations ... Page 1 of 1

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"ATERIALSENGI~

Quick Guide: Exposure Risks and Control Measures for Common Laboratory Operations InvolvingNanomaterials

Activity types, by Risk of Exposure Primary Control Measures

Low Probability:Disposable nitrile or PVC gloves. Do not reuse•

Non-destructive handling of solid nanoparticlegloves.

• • Wet cleaning procedures and/or HEPA vacuumcomposites or nanoparticles permanently bonded to a for surfaces/equipment.substrate

Medium / High Probability:

• Conduct task within a fully enclosed system• Working wi nanomaterials in liquid media during pouring (e.g., glovebox), or fume hood

or mixing, or where a high degree of agitation is • Disposable gloves appropriate for the solvent ininvolved (e.g., sonication) which the particles are suspended. Do not reuse

• Handling nanostructured powders* gloves.• High speed abrading/grinding nano-composite materials • Safety eyewear (+ face shield if splash potential• Maintenance on equipment used to produce exists)

nanomaterials • Wet cleaning procedures for surfaces/equipment• Cleaning of dust collection systems used to capture

nanoparticles

High Probability:

• Work in enclosed systems only (e.q., glovebox,• Generating nanoparticles in the gas phase or in aerosol glovebag, or sealed chamber).

(spill or liquid)

• Manipulation of nanoparticles in gas stream

* EH&S recognizes that low-density nanomaterials (e.q., carbon-based) become aerosolized by even the slightest airmovement and may not be practical when weighed or handled in laboratory fume hoods. Consult with EH&S on alternativesets of controls.


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Spill Response

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Spill Response

In addition to following EH&S general spill response directions (http://chemtoalkit.stanford.edu/emeraencies)/ integratethe additional measures for spills involving nanomaterials:

• Use wet clean up methods or vacuum cleaners equipped with HEPA-filters.• Do not dry sweep or use conventional vacuum cleaners.• Collect spill cleanup materials in a tightly closed container• Manage spill cleanup debris as hazardous waste.


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Disposal

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Disposal

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• As a prudent measure, manage nanoparticle wastes, including contaminated lab debris, as a part of your normallaboratory hazardous waste stream. See http://ChemToolkit.stanford.edu/ChemWasteDisDosa[ for additionalinformation on hazardous waste management.

• Collect and store waste materials in a tightly closed container. Include information describing the nanoparticulatenature of the materials on the waste tag (e.g., contains nanosilver material).


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Additional Information and References Page 1 of2

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Additionailliformation and References

The field of nanotechnology is rapidly evolving. The following entities provide additional information regarding the researchefforts underway by governmental agencies and other institutions to fill in knowledge gaps.

• National Institute for Occupational Safety and Health (NIOSH) - Nanotechnologywww.cdc.gov/niosh/topics!nanotech/

• National Institute of Occupational Safety and Health - Approaches to Safe Nanotechnology: Managing the Health andSafety Concerns Associated with Engineered Nanomaterials (March 2009)http://www.cdc.gov/niosh/docs/2009-125/

• United States Department of Laborhttp://www.osha.gov/dsg/nanotechnology/nanotechnology.html

• American Chemical Society: Nanotechnology Safety Resourceshttp://membership.acs.org/C/CCS/nano.htm

• Environmental Protection Agency - Nanotechnologyhttp://es.epa.gove/ncer/nano/index.html

• National Nanotechnology Initiativehttp://www.nano.gov

• GoodNanoGuidehttp://goodnanoguide.org/tiki-index.php?page-HomePage

References:

Barlow PG, Clouter-Baker AC, Donaldson K, MacCallum J, Stone V [2005]. Carbon black nanoparticles induce type II epithelial cells to releasechemotaxins for alveolar macrophages. Particle and Fiber Toxicol 2, 14 pp [open access].

Brown DM, Wilson MR, MacNee W, Stone V, Donaldson K [2001]. Size-dependent proinflammatory effects of ultrafine polystyrene particles: A rolefor surface area and oxidative stress in the enhanced activity of ultrafines. Toxicology and Applied Pharmacology 175(3): 191-199.

Daigle CC, Chalupa DC, Gibb FRMorrow PE, Oberdorster G, Utell MJ, Frampton MW [2003]. Ultrafine particle deposition in humans during rest andexercise. Inhalation Toxieol 15(6):539-552.

Donaldson K, Aitken R, Tran L, Stone V, Duffin R, Forrest G, Alexander A. [2006] Carbon Nanotubes: a Review of Their Properties in Relation toPulmonaryToxieology and Workplace Safety. Toxieol Sci. 92(1): 5-22.

Duffin R, Tran Cl, Clouter A, Brown OM, MacNee W, Stone V, Donaldson K [2002]. The importance of surface area and specific reactivity in theacute pulmonary inflammatory response to particles. Ann Occup Hyg 46:242-245.

Duffin R, Tran L, Brown D, Stone V, Donaldson K [2007}. Proinflammogenic effects of low-toxicity and metal nanoparticles in vivo and in vitro:highlighting the role of particle surface area and surface reactivity. Inhal Toxicol 19(10):849-856.

Helland A, Wick P, Koehler A, Schmid K, Som C [2007]. Reviewing the Environmental and Human Health Knowledge Base of Carbon Nanotubes.Environ Health Perspectives 115(8): 1125-1131.

Geiser M, Rothen-Rutishauser B, Kapp N, Schurch S, Kreyling W, Schulz H, Semmler M, Im Hof V, Heyder J, Gehr P [2005]. Ultrafine particles crosscellular membranes by nonphagocytic mechanisms in lungs and in cultured cells. Environ Health Perspectives 113(11):1555-1560.

ICRP [1994]. Human respiratory tract model for radiological protection. Oxford, England: Pergamon, Elsevier Science Ltd., InternationalCommission on Radiological Protection Publication No. 66.

Jaques PAl Kim CS [2000]. Measurement of total lung deposition of inhaled ultrafine particles in healthy men and women. Inhal Toxico! 12(8):715731.

Kim CS, Jaques PA [2004]. Analysis of total respiratory deposition of inhaled ultrafine particles in adult subjects at various breathing patterns.Aerosol Sci Technol 38:525-540.

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Additional Information and References Page 2 of2

Kreyling WG, Semmler M, Erbe F, Mayer P, Takenaka 5, Schulz H, Oberdorster G, Ziesenis A [2002]. Translocation of ultrafine insoluble iridiumparticles from lung epithelium to extrapulmonary organs is size dependent but very low. J Toxlcol Environ Health 65(20): 1513-1530.

lison, D., C. Lardot, F. Huaux, G, Zanetti, Fubini B [1997]. Influence of particle surface area on the toxicity of insoluble manganese dioxide dusts.Arch. Toxlcol. 71(12): 725-729.

Maynard AM, Kuempel ED [2005J. Airborne nanostructured particles and occupational health. J Nanoparticle Research 7(6):587-614.

Moller W, Hofer T, Ziesenis A, Karg E, Heyder J [2002]. Ultrafine particles cause cytoskeletal dysfunctions in mecrophaqes. Toxieol Appl Pharmacal182(3): 197-207.

Moller W, Brown OM, Kreyling WG, Stone V [2005]. Ultrafine particles cause cytosketeta! dysfunctions in macrophages: role of intracellular calcium.Part Fibre Toxtcot. 2:7, 12pp.

Oberdorster G, Fer-in J, Gelein R, Soderholm SC, Finkelstein J [1992]. Role of the alveolar macrophage in lung injury-studies with ultrafineparticles. Environ Health Perspect 97:193-199.

Oberdorster G, Ferin J, Lehnert BE [1994a]. Correlation between particle-size, in-vivo particle persistence, and lung injury. Environ Health Perspect102(SS}: 173-179.

Oberdorster G, Ferin J, Soderholm S Gelein R, Cox C, Baggs R, Morrow PE [1994b]. Increased pulmonary toxicity of inhaled ultrafine particles: dueto lung overload alone? Ann. Occup. Hyg. 38(Suppl. 1): 295-302.

Oberdcrster G, Sharp Z, Atudorei V, Elder A, Gelein R, lunts A, Kreyling W, Cox C [2002]. Extrapulmonary translocation of ultrertne carbon particlesfollowing whole-body Inhalation exposure of rats. J Toxicol Environ Health 65 Part A(20): 1531-1543.

Oberdcrster G, Sharp Z, Atudorei V, Elder A, Gelein R, Kreyling W, Cox C [2004]. Translocation of inhaled ultrafine particles to the brain. InhalToxlcol 16(6-7):437-445.

Oberdorster G, Oberdorster E, Oberdorster ] [2005a]. Nanotoxicology: an emerging discipline evolving from studies of ultrafine particles. EnvironHealth Perspect. 113(7):823-839.

Pritchard OK [2004]. Literature review-explosion hazards associated with nanopowders. United Kingdom: Health and Safety Laboratory,HSL/2004/12.

"Safe Nanotechnology in the Workplace - An Introduction for Employers, Managers, and Safety and Health Professionals" [Feb. 2008]. NationalInstitutes of Health. DHHS (NIOSH) Publication No. 2008-112.

Semmier M, Seitz J, Erbe F, Mayer P, Heyder J, Oberdorster G, Kreyling WG [2004]. Long-term clearance kinetics of inhaled ultrafine insolubleIridium particles from the rat lung, including transient translocation into secondary organs. Inhal Toxicol 16(6-7): 453-459.

Takenaka S, Karg OJ Roth CJ Schulz H, Ziesenis A, Heinzmann U, Chramel P, Heyder J [2001]. Pulmonary and systemic distribution of inhaledultrafine silver particles in rats. Environ Health Perspect 109(suppl. 4):547-551.

Tran Cl, Cullen RT, Buchanan 0, Jones AD, Miller BG, Searl A, Davis JMG, Donaldson K [1999]. Investigation and prediction of pulmonary responsesto dust. Part II. In: Investigations into the pulmonary effects of low toxicity dusts. Contract Research Report 216/1999 Suffolk, UK: Health andSafety Executive.


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Attachment 3

Standard Operating Procedure Template for theLaboratory Use of Nanomaterials

Process or Experiment Overview: Description of the process or experiment that the SOP#2 covers. This process may be described in general terms, such as cleaning and purification

of single walled carbon nanotubes.

D one time D daily D weekly D monthly

Risk Assessment: Identify potential chemical and safety hazards. Special consideration#3 should be given to the high reactivity of some nanopowders with regard to potential fire and

explosion. Consider the hazards of the precursor materials in evaluating the process.

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#4Controls: (Check those that apply to the risks and procedures for the workthat will be done)

...~"I:I"'~~' "~I:I'.~" "~ ...."

EI'I!lil'leetil'lg~ol'ltt()ls ..... ,.,;;,.,'W~~ yf·...~·'·~~

D General laboratory • Non-destructive handling of solid nanoparticle

ventilationcomposites or nanoparticles permanently bonded to asubstrate

D Laboratory fume hood; or • Working wi nanomaterials in liquid media duringpouring or mixing, or where a high degree of agitation

D Exhausted enclosure is involved (e.g., sonication)

• Handling nanostructured powders*

• Maintenance on equipment used to producenanomaterials

• Cleaning of dust collection systems used to capturenanoparticles

D Glove box; or • Generating nanoparticles in the gas phase or inaerosol

D Other sealed enclosure • Manipulation of nanoparticles in gas stream

.

*If work cannot be conducted with appropriate engineering controls, consult with EH&S, 723-0448.

-r- ~ •. ro' ..~....0 gloves; indicate type:

0 safety goggles o face shield o lab coats 0 other:

0 appropriate street clothingRespiratory protection is generally not required for lab research, provided the appropriate engineeringcontrols are employed. For additional guidance on respiratory protection, consult with EH&S, 723-0448.

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#4c Location of Nearest Emergency Safety Equipment:

Item: I - , ' -

-,

First Aid Kit

_,i",,~ Spill

Fire .._.- ,

.....u ..~

~ire_'•.' ....'a.......a"

Step-By-Step Operating Procedure: Provide a sequential description of work, including#5 details such as chemical concentrations and when special safety equipment is to be utilized.

Clearly label areas where nanomaterials are used.

#6 Decontamination:

• Upon leaving the nanomaterial work area, remove any personal protective equipment wornand wash hands, forearms, face, and neck.

• After each use (or day), wipe down and or HEPA vacuum the immediate work area andequipment to prevent accumulation of nanoparticles.

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#7 I Spill and Accident Procedures:

Health Threatening Emergencies (ex. Fire Explosion, Serious Injury, or other ImmediateDanger)

1. CALL 9-911 (286 in School of Medicine) for the Fire Department.2. Alert people in the vicinity, activate local alarm systems.3. Evacuate the area.4. REMAIN NEARBY TO ADVISE EMERGENCY RESPONDERS.5. Once personal safety is established, call EH&S at 725-9999 (or in the School of Medicine,

x286) and proceed with local notifications, below.

If Personnel Exposed:1. Remove exposed/contaminated individual(s) from area, unless unsafe to do so because of (a)

medical condition of victim(s), or (b) potential hazard to rescuer(s).2. If immediate medical attention is required, notify SU Emergency 9-911 (or 286 at the School

of Medicine).3. Notify EH&S to report the potential exposure by calling 5-9999 (or in the School of Medicine,

x286).4. Administer First Aid as appropriate.5. Flush contamination from eyes/skin using the nearest emergency eyewash /shower for a

minimum of 15 minutes. Remove any contaminated clothing.6. Take copy of MSDS(s) of chemical(s) to hospital with victim.7. Contact EH&S's Occupational Health Center at (650) 725-5308 for follow-up.

Non-Health Threatening Emergencies (ex. spills requiring cleanup assistance)In the event of a spill or release, which mayor has impacted the environment (storm drain, soil, airoutside the building), or spill or release that cannot be cleaned up by local personnel:

1. Notify Stanford Responders: Call Notify Stanford Responders: Call 725-9999 (286 in theSchool of Medicine) (24 hours/day, 7 days/week).

2. Provide local notifications to your supervisor.

Small Spills/Local Cleanup:In the event of a minor spill or release that can be cleaned up by local personnel using readilyavailable equipment (absorbent, available from EH&S in Small Spill Kit):

1. Notify personnel in the area and restrict access. Eliminate all sources of ignition.2. Review the MSDS for the spilled material, or use your knowledge of the hazards of the

material to determine the appropriate level of protection.3. Wearing appropriate Personal Protective Equipment, clean up spill involving

nanomaterials using wet methods and/or HEPA vacuum. DO NOT SWEEP SPILLEDOR DRIED NANOMATERIALS. Collect spill cleanup materials in a tightly closedcontainer. Manage spill cleanup debris as Hazardous Waste.

4. If greater than 30 ml, or if it will take longer than 15 minutes for you to clean-up, immediatelycall EH&S at 725-9999 (or in the School of Medicine, x286) to report the spill, and notify yoursupervisor.

5. Submit on-line waste pickup request to EH&S.

#8 I Waste Disposal:

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• Manage nanoparticle wastes, including contaminated lab debris, as a part of your normallaboratory Hazardous Waste stream.

• Collect and store waste materials in a tightly ciosed container. Include informationdescribing the base nanoparticie materials on the waste tag.

#9 Training Requirements:

General Training (check all that apply):

DGeneral Safety & Emergency Preparedness (EHS-4200)

DChemical Safety for Laboratories (EHS-1900)

. DOther:

I~~~~ti~~~~~t~!~~~~~~~ I IMaintained:

Laboratory-specific training (check all that apply):

DReview of MSDS for Nanomaterial (if available)

DReview of MSDS for other chemicals involved in process/experiment

DReview of this SOP

DOther:

1;~;~!i~~j~~i~~~~~g~80~ I IMaintained:

Approval Required: Identify any tasks that require prior approval by the PI/ Laboratory

#10Supervisor (e.g., use of Restricted Chemical and other higher hazard chemicals, andrunning of higher hazard operations). Refer tohttp://chemtoolkit.stanford.edu/RestrictedChem for more information.

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