Scanning Probe Microscopy - Analytical Chemistry (ACS Publications)

Department of Physics, University of California Santa Barbara, Santa Barbara, California 93106 ... both an M.S. and Ph.D. in mechanical engineering fr...
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Anal. Chem. 2004, 76, 3429-3444

Scanning Probe Microscopy Mark A. Poggi, Elizabeth D. Gadsby, and Lawrence A. Bottomley*

School of Chemistry & Biochemistry, Georgia Institute of Technology, Atlanta, Georgia 30332-0400 William P. King

The George W. Woodruff School of Mechanical Engineering, Georgia Institute of Technology, Atlanta, Georgia 30332-0405 Emin Oroudjev and Helen Hansma

Department of Physics, University of CaliforniasSanta Barbara, Santa Barbara, California 93106 Review Contents Instrumental Innovations Probes Hardware Improvements/New Techniques Theory and Calibration Imaging Applications Scanning Tunneling Microscopy Scanning Electrochemical Microscopy Atomic Force Microscopy Nanofabrication and Nanolithography Thermal Techniques Biological Imaging Sensors Force Spectroscopy Chemical and Polymer Applications Biological Applications Proteins Membrane Proteins Ligand-Receptor Interactions Protein Networks (Mesostructures) Living Cells Nucleic Acids Characterization of Carbon Nanotubes Literature Cited

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Scanning probe microscopy (SPM) comprises a family of techniques that measure surface topography and properties on the atomic scale. There are an ever increasing number of papers devoted to technical advances and applications of SPM. During the period January 1, 2002 through December 31, 2003, more than 11 000 citations were found using the Chemical Abstracts Service that pertained to or applied scanning probe methods. The intent of this review is to highlight innovative contributions to the field published within the aforementioned period. Due to editorial restrictions on the number of citations, the work cited herein illustrates only some of the research avenues currently being explored with SPM. Our selections are, without doubt, subjective.

INSTRUMENTAL INNOVATIONS Probes. Significant advances in the design and application of microcantilever probes have been reported. Manning and co* [email protected]. 10.1021/ac0400818 CCC: $27.50 Published on Web 05/20/2004

© 2004 American Chemical Society

workers fabricated piezoelectric tapping mode cantilevers that have an integreated oscillation drive mechanism on the cantilever (1). Miyahara et al. reported a piezoelectric cantilever capable of simultaneous deflection sensing, cantilever oscillation, and feedback actuation in noncontact mode AFM imaging (2). Rogers and co-workers have fabricated piezoelectric probes capable of imaging in fluid (3). Brook et al. created a unique piezoresistive probe that can carry out both topological and magnetic imaging (4). This probe enables scanning Hall probe imaging of nonconducting or unconnected magnetic samples. Grow et al. presented a simpler method for fabricating silicon nitride cantilevers with oxidation-sharpened tips. The height of the tip is taller than commercially available nitride tips and is especially useful for imaging fragile samples that possess large topological relief (5). Several methods were reported for fabricating nanotube-tipped AFM probes (6-8). Nishino et al. (9) immobilized carbon nanotubes onto a gold-coated STM tip. Images acquired with these showed increased spatial resolution and afforded chemical discrimination of an oxygen functionality present on the substrate surface. Snow and co-workers examined the factors that influence topological imaging with carbon nanotube-tipped probes (10). Bale and Palmer have fabricated tip arrays that are suitable for parallel STM imaging applications (11). Chow and co-workers (12) have addressed approach, alignment, and density issues associated with operating two-dimensional scanning probe arrays. They reported the fabrication and characterization of twodimensional micromachined silicon cantilever arrays with integrated through-wafer electrical interconnects. With these arrays, a substrate domain as large as 3.8 mm × 0.45 mm can be imaged. Several reports of probes specifically designed for the acquisition of optical information about substrates appeared. For example, Aigouy and co-workers attached a fluorescent, rare-earth-doped fluoride glass particle to the end of an AFM tip (13). When this probe was scanned over the surface of a nanostructured sample illuminated by a laser beam, the intensity of fluorescence from the particle was then recorded as a function of the position. This method enabled them to map the location of pinhole defects in opaque films. Crozier et al. (14) fabricated a silicon nitride solid immersion lens onto a cantilever beam. This probe was used for scanning optical microscopy and was capable of achieving optical Analytical Chemistry, Vol. 76, No. 12, June 15, 2004 3429

resolution λ/(2n) where n is the refractive index of the nitride lens. Lee, Ding, and Bard (15, 16) reported the first successful simultaneous topographic and optical imaging of a living unicellular organism using a novel probe tip. The tip, consisting of an optical fiber core, a gold ring, and an insulator, served as an ultramicroelectrode for scanning electrochemical microscopy (SECM) and as a light source for optical microscopy when coupled to a laser. Images were acquired in either constant force or constant current mode. Improved lateral resolution for both optical imaging and SECM imaging was achieved using a constant current while maintaining a fixed distance between the tip and the sample. This technique provides simultaneous electrochemical, optical, and structural information about interfaces. Hughes and Wang created nanoscale cantilevers composed of zinc oxide (17). Extremely small forces could be measured using cantilevers of this size once a means for force/deflection transduction is realized. Similarly, chromium nanocantilevers were fabricated and mechanically tested with an AFM probe (18). Lee and co-workers (19) fabricated a novel probe for integrated AFM-mass spectrometry. The cantilever acts as the force sensor for topological imaging, and the tip acts as a sampler for chemical analysis by time-of-flight mass spectrometry. As the tip in contact with the sample scans across the surface, chemical compounds from the surface adhere to it. At a desired location, the tip is raised via the integrated piezoelectric actuator into a position near an extraction electrode. Application of a potential pulse between the tip and the extraction electrode results in ionization and acceleration of the tip-adherent chemicals into the time-of-flight mass analyzer. The mass spectrum provides identification of the tipadherent molecule(s). Hardware Improvements/New Techniques. Sulchek et al. critically examined ways to increase the scan speeds of both contact and intermittent contact imaging (20). Current limitations on scan speeds can be eliminated by integration of a faster feedback actuator as well as active control of the dynamics of the cantilever. Akiyama et al. (21) devised a fast driving technique that utilizes a tuned filter that boosts the servo signal in proportion to its frequency. An imaging bandwidth of 5 kHz was achieved; images were acquired in constant force mode at tip velocities up to 0.62 mm/s. Stark and Heckl (22) have improved tapping mode imaging by driving cantilever resonance at one frequency and monitoring deflection at a harmonic of this frequency. The result is significant enhancement in image contrast. Zahl and co-workers have created software that can be used to control many different types of SPMs and process images (23). The software is extremely flexible and can be used to control many of the different modes in SPM. The software package is available at no cost to users. Trawick and co-workers used a polynomial mapping method to correct for piezoelectric-induced artifacts in SPM images (24). This correction scheme can reduce the effects of distortion in an AFM image from 5% of the scan width to a single pixel. Kindt et al. (25) developed a real-time method for eliminating the drift component associated with AFM images. Their method incorporates automatic changes in the set point to maintain a set difference in the relative feature richness of two traces taken with different offsets. 3430

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Schaffer has designed an array detector that combines a higher sensitivity and a larger dynamic range (in the z-direction) than conventional two-segment photodiode detectors (26). This detector is less susceptible to nonlinearities during force measurements. Onaran and colleagues (27) utilized radiation pressure generated by a focused acoustic beam to implement tapping mode and elasticity imaging by AFM. Their method enables efficient excitation and spatial mapping of both higher-order flexural and torsional modes of AFM cantilevers in liquids. Arnold et al. (28) presented a new approach for studying friction and stick-slip phenomena analyzing the torsional resonance of the cantilever. Pfeiffer and co-workers (29) proved that lateral forces can be measured even when the tip is not in contact with the surface. Alcarez and co-workers measured and modeled the viscous drag that a cantilever experiences when in proximity with a surface (30). Correction of the drag artifact could lead to an improvement in the scan speed in contact mode imaging of soft samples in liquid and to an increase in the pulling speed range in force spectroscopy measurements. Two groups have examined the cross-talk that occurs during the acquisition of topographical and friction data in AFM (31, 32) and presented methods for detection and correction of this artifact. Buh and Kopanski (33) have looked at the effect of illumination from the laser of an optical beam deflection AFM on a semiconductor sample with a scanning capacitance probe connected in parallel. Significant differences in the capacitance-voltage characteristics were observed and attributed to light spillage over the edges of and transmission through the cantilever. Dubreuil et al. (34) have used an AFM to directly image the air-water interface. Phase and topography images revealed information about the layering of Langmuir-Blodgett films. Burns has presented a unique technique where AFM is coupled in real time with submicrometer confocal fluorescence imaging (35). Hu and co-workers have combined tapping mode atomic force microscopy and fluorescence lifetime imaging microscopy (36). They have demonstrated that spatially mapping the change in fluorescence lifetime and intensity is a promising approach to spectroscopic imaging at the length scales obtainable with AFM. Noy and Huser (37) integrated an AFM into a scanning confocal optical microscope enabling simultaneous acquisition of optical and topographical images of surfaces. Fukushima et al. (38) developed an AFM that mounts in an SEM with easy sample change, optical alignment, and sample positioning capabilities. The viewing angle of the SEM was designed so that the apex of the AFM tip could be observed for most samples. Browne et al. have designed a scanning transmission X-ray microscope for use with synchrotron radiation allowing simultaneous X-ray imaging and topological probing of a surface (39). This technique could provide a plethora of information regarding radiation damage to surfaces or samples and dynamic processes such as specimen corrosion. Bondarenko et al. (40) have developed a scanning magnetic microscope that does not induce appreciable applied forces or magnetic excitations on specimens. This microscope is intended to measure weak magnetic field distributions near the surface at micron and submicrometer scales. Several groups have employed in situ AFM to characterize adsorbates on the active surface of acoustic resonators (41-43).

This combined technique facilitated correlation of mass changes with topology. Theory and Calibration. Imaging. Giessibl has written a comprehensive review of common AFM imaging modes and the theory behind them (44). Hofer et al. (45) presented a thorough review of theoretical models that have been used to interpret SPM data. Balantekin and Atalar have devised a model the can elucidate the amount of power that is dissipated into a sample during tapping-mode AFM (46). This model facilitates in the determination of the dampening constant of a sample and could potentially be used to quantify dampening phenomena in composite systems. Boisgard et al. have presented a short overview covering two models that evaluate the loss of energy when a cantilever tip is oscillated in close proximity of a surface (47). Rodriguez and Garcia have provided a thorough interpretation of an AFM when operated under active quality factor control (48). Their theoretical treatment demonstrates that, when using Q-control, the force exerted on the substrate is minimized. Su et al. (49) have correlated residual tip speed before impacting with a surface and the associated tip wear for relatively hard samples. They have shown that a lower set point is not harmful and actually leads to higher resolution images. Stark et al. have presented dynamic experiments where the contact force during a typical tapping mode experiment is determined (50). They’ve shown that, under normal imaging condition, the contact force exceeds 200 nN. Lee and co-workers (51) used nonlinear dynamical systems theory to analyze the oscillatory properties of a cantilever when used in dynamic force microscopy. They brought better understanding of the sudden global changes that occur in the interaction potential at certain gap widths that cause the tip to irregularly tap the sample. Hoffmann has presented simulations of driving a cantilever off-resonance when performing noncontact atomic force microscopy (52), which facilitates a more general route to the reconstruction of the surface force gradients. Couturier et al. have provided a complete analysis of the behavior of a noncontact atomic force microscope (53). They present numerical models for the stability of the cantilever tip when it oscillates close to the sample. Chang and Chu have derived a closed-form expression for the oscillatory behavior of cantilevers with complicated cross-sections (54). Force Spectroscopy. In an effort to model the interaction of a solid colloidal probe with an incompressible liquid drop, Bardos presented a very rigorous theoretical interpretation (55). His models avoid the pitfalls of both perturbation theory and of purely numerical solutions to the Young-Laplace equation. Bedrov and Smith have performed molecular dynamics simulations of the mechanical pulling of poly(ethylene oxide) (PEO) chains in water and n-tridecane to elucidate the mechanism(s) of elastic response of the amphiphilic PEO chain in hydrophilic and hydrophobic environments (56). The simulations quantitatively match AFMbased single-molecule mechanical tests. Biesheuvel et al. have derived models describing the electrostatic repulsion between similar surfaces with ionizable surface groups interacting across aqueous solutions (57). Butt and Volker have theoretically treated the jump to contact point of a cantilever tip through a thin polymer film (58). They have derived a relationship between the force dependence of the activation energy of the point of initial snapdown and the approaching velocity of the tip. Fraxedas and

co-workers have presented a new model based on an equivalent spring constant that takes into account the changes in in-plane interactions during nanoindentation (59). Their model correlates well with experimental data from nanoindentation of several crystalline surfaces. Patrick et al. (60) have used molecular dynamics simulations to provide a detailed description of the adhesive interactions that are probed in chemical force microscopy experiments. Their models take into account atomic-scale motions and distributions of forces. Dean et al. have developed molecularlevel models for electrostatic interactions between polyelectrolyte brushes when explored using chemical force microscopy (61). The relevance of these models to the modeling of native cartilage is discussed in detail. Dudko and co-workers have described a new model that predicts a distribution of forces, the mean rupture force, and the variance during single-molecule pulling experiments (62). The mean rupture force follows a (In V)2/3 dependence on the pulling velocity, V, which differs from earlier predictions, and they have shown that at low pulling velocities a rebinding process can occur that can delay the rupture of the molecule and lead to a bimodal distribution of the observed rupture forces. Leng and Jiang (63) used a hybrid molecular dynamics simulation to investigate adhesion and friction in chemical force microcopy experiments. The hybrid simulation method allows one to simulate force-distance curves (or adhesion) and friction loops (or friction) in the CFM on the experimental time scale for the first time. Friedsam et al. have performed Monte Carlo simulations that demonstrate the severe impact that variable polymer spacer lengths can have on the mean rupture force that is observed during single-molecule mechanical tests (64). Piezoceramic cantilever beams are beginning to play significant roles as sensors, actuators, and resonators. Ballato has theoretically modeled the rotations and deflections of cantilever beams (65). This theoretical treatment identified oscillatory frequencies (both vertical and torsional modes) and derived approximations of the corresponding cantilever displacements at the end of the cantilever beam. The Sader group has investigated the torsional response of a cantilever immersed in a viscous medium and when it is externally driven (66). They have shown experimentally that triangular cantilevers are more susceptible to torque during imaging (67). Gotszalk and co-workers presented a technique for the calibration of Wheatstone bridge cantilevers with an accuracy of better than 5% (68). Jericho and Jericho designed a new instrument that can be used to determine the spring constant of AFM cantilevers with better than 10% uncertainty (69). The device is compact and easy to use. Burnham et al. compared methods for calibrating cantilever spring constants and presented a new method that is easier to perform and comparable in accuracy to the best method available (70).

IMAGING APPLICATIONS Scanning Tunneling Microscopy. STM continues to provide new insight into the balance between intermolecular interactions, molecule-substrate interactions, and thermal energies that govern the diffusion, nucleation, and self-organization of molecules on surfaces. For example, STM images acquired by Barth et al. (71) provided compelling evidence that 4-[trans-2-(pyrid-4-ylvinyl)]Analytical Chemistry, Vol. 76, No. 12, June 15, 2004

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benzoic acid (PVBA) and 4-[(pyrid-4-ylethynyl)]benzoic acid (PEBA) self-assemble into rigid, rodlike molecules. At low temperatures, both PVBA and PEBA form irregular networks of flatlying molecules linked via hydrogen bonding in a diffusion-limited aggregation process. The head-to-tail hydrogen bonding of the related rodlike species PEBA and PVBA stabilizes molecular rows on Ag(111) surfaces. The subtle difference in the molecular geometries is reflected in the lateral ordering: two-dimensional islands are observed with PEBA whereas enantiopure onedimensional nanogratings of supramolecular, chiral H-bonded twin chains are found for PVBA. Berner and colleagues (72) studied the adsorption and twodimensional ordering of chloro(subphthalocyaninato)boron(III) (SubPc) on Ag(111). Sublimation of SubPc onto an Ag(111) results in interesting phase behavior. At 0.2-0.5 monolayer coverage, the molecules assemble into two-dimensional islands composed of a well-ordered honeycomb pattern. At higher coverage, the molecules assemble into a two-dimensional hexagonal close-packed superlattice. Two different orientations of the superstructures with respect to the Ag(111) substrate are observed. Two enantiomorphic superstructures are found for the honeycomb pattern. The arrangement of the molecules in the honeycomb and the hcp pattern were related to the charge distribution of the individual SubPc molecule and the resulting electrostatic interaction with the underlying substrate. They also observed self-assembly of highly ordered, periodic, intermixed monolayers consisting of onedimensional chains or two-dimensional hexagonal patterns on uniform, unreconstructed, atomically clean Ag(111) terraces through competing noncovalent interactions between SubPc and C60 (73). “Nanotube” structures of R-, β-, and γ-cyclodextrins (CyDs), were constructed by Miyake and co-workers (74) using potentialcontrolled adsorption onto Au(111) surfaces from electrolyte solution. At open circuit, CyD molecules adsorbed randomly on this surface and could be desorbed at potentials negative of -0.60 V versus SCE. Ordered molecular arrays were observed for each of the CyDs over specific potential ranges. STM images revealed head-to-head, tail-to-tail, and head-to-tail conformations due to the strength difference between primary-primary and primarysecondary hydrogen bonds of CyDs. Their results demonstrate the utility of controlled potential adsorption in the formation of two-dimensional supramolecular structures on the substrates. De Feyter and De Schryver (75) have published an excellent review of recent progress in the study of two-dimensional supramolecular self-assembly on surfaces probed by STM, with special emphasis on structure, dynamics, and reactivity of hydrogen-bonded systems. Bode and co-workers (76) examined correlations between structural, electronic, and magnetic properties of iron nanowire arrays on stepped W(1 1 0) substrates using spin-polarized STM and STS in ultrahigh vacuum, at low temperatures and under external magnetic fields. They found a strong dependence of the nanomagnetic domain structure on the width of the double-layer iron nanowires as well as atomic-scale domain walls in singlelayer nanowires. These unique properties of low-dimensional magnetic systems are not observed in bulk materials. Several groups focused on the development of novel approaches to provide chemical information while acquiring STM 3432

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images of a surface. For example, Downes and co-workers (77) collected the light emitted from the tip-sample to examine spectroscopically resolved light emission maps of metal surfaces. A rough Au film was shown to exhibit minimal color changes across the sample. With their color mapping approach, the authors were able to distinguish between Ag and Cu clusters with ∼10nm resolution. Ding and co-workers (78) integrated a STM with a time-offlight mass analyzer to locally ionize surface atoms by the combination of an optical laser pulse and tip bias voltage. This approach makes possible the detection of desorbed ions from the nanometer-scale area under the tip. Miyatake and co-workers (79) used a shielded STM tip as a field emission electron source for Auger analysis, enabling elemental analysis of a surface during imaging by Auger electron spectroscopy. During the past few years, spin-polarized STM and STS have proven to be reliable tools for imaging surface magnetic domain structures and for determining the energy-resolved spin polarization of the sample with nanometer resolution. Kubetzka and coworkers (80) addressed the problem of quantifying local surface spin polarization arising from the dependence of tip-sample separation on the magnitude and sign of the spin polarization. Selection of specific stabilization voltages that ensured a non-spinpolarized total current enables imaging under constant separation and results in the disappearance of the signature of domain walls from the topographic image. Hofer (81) reviewed the challenges and errors that accompany interpretation of high-resolution STM images. He encouraged comparison of experimental findings with simulations that predict the influence of chemical interactions between tip and sample to extend the information available about the sample under analysis. In a separate review, he examined the present status of computational modeling of STM imaging applications in studying surface properties, such as adsorption, point defects, spin manipulation, and phonon excitation (45). Voigt and Koch (82) established the theoretical basis for evaluating surface acoustic wave-STM images recorded in constant current imaging mode. They examined the surface motion induced by a Rayleigh wave on LiNbO3 to quantitatively determine the geometry of the atomic oscillation ellipse given by its eccentricity and the amplitude of the transverse displacement. They report that their system is capable of generating and detecting surface acoustic waves with transverse displacement amplitudes as small as 0.001 Å with spatial resolution for acoustic measurements in the nanometer range. Okawa and Aono (83) fabricated and imaged electrically conductive organic nanowires using STM. A monolayer of diacetylene compounds chemisorbed onto graphite self-assembled into parallel lines on the substrate. They then used a STM tip to initiate linearly propagating chain polymerization of the diacetylenes at any predetermined point and terminate it at another predetermined point with nanometer spatial precision. Scanning Electrochemical Microscopy. Keung and coworkers (84) reported the simultaneous imaging of topography and electrochemical activity using integrated SECM-AFM cantilevers operated in Tapping Mode. The current response at the microring electrode was equivalent to that observed in contact mode. The potential utility of these bifunctional cantilevers for

imaging of soft samples was illustrated with a patterned substrate composed of glucose oxidase confined in a polymer matrix. A new method for measuring local interfacial impedance properties with high lateral resolution was developed by Katemann and co-workers (85). They coupled electrochemical impedance spectroscopy with SECM to visualize microscopic domains of different conditions and electrochemical activities on solid-liquid interfaces immersed into an electrolyte. The performance of the method was illustrated by imaging an array of Pt microelectrodes in the absence of a redox mediator. Zhang and Unwin (86) used a proton feedback approach to study lateral proton diffusion processes at stearic acid monolayers at the air-water interface by SECM. The method afforded quantitation of the rate of lateral proton diffusion under steadystate conditions while the monolayer was maintained under welldefned surface pressure control. Their findings demonstrate that the state of the monolayer has a significant influence on lateral proton diffusion. Liljeroth and co-workers (87) examined the potential of ringdisk ultramicroelectrodes as SECM probes both theoretically and experimentally. Theoretical approach curves to both insulating and conducting substrates for disk generation/ring collection mode of operation were calculated by numerical methods. Their calculations predicted that the current response as a function of distance depends on the ring radius but is relatively insensitive to ring thickness and overall tip radius. They experimentally verified their calculations by tracking the partitioning of iodine across an immiscible liquid-liquid interface with a ring-disk SECM probe. Treutler and Wittstock (88) developed a combined EC-STM/ SECM instrument enabling the collection of spatially correlated topographic and reactivity data on conductive surfaces. Their approach involved bringing a combined ECSTM/SECM probe into tunneling contact with a dodecanethiolate-covered gold surface and then retracting it by defined distances (5-30 nm) to perform electrochemical experiments outside the tunneling gap. The combination of the two imaging modes together with the use of nanometer-sized electrodes allowed tremendous improvement of lateral resolution for SECM imaging. Numerical simulations of the complex nonsymmetrical 3D systems encountered in SECM experiments were performed by Sklyar and Wittstock (89). Steady-state amperometric SECM responses were simulated in three dimensions with the boundary element method using an exterior Laplace formulation. The new formulation afforded significant reduction in the computational errors arising from the incorrect positioning of the bulk boundary. Simulations were undertaken to analyze the influence of SECM probe tilt and distance control modes on the spatial resolution of images acquired over conductive and insulating substrates. Zoski et al. (90) examined the effect of shielding of an ultramicroelectrode by a larger substrate electrode in SECM for reversible and quasi-reversible reactions. Shielding occurs when the potential of the substrate and the tip are the same. Feedback occurs when the electrode is within a few micrometers of the conductive substrate. For reversible reactions, the shielding effect causes the tip current to decrease below that expected for an insulator. For quasi-reversible reactions, the effect of shielding depends on the standard rate constant k°, the transfer coefficient,

and the dimensionless potential parameter. For large values of k°, the tip current versus distance curves approaches that expected for a conductor as for reversible kinetics, while for very small k°, insulator behavior is slowly approached. Thus, shielding provides a way of measuring fast kinetics at a substrate surface. This approach holds promise as a means of mapping the electrontransfer activity of a surface by SECM.

ATOMIC FORCE MICROSCOPY There is a continuing interest in scanned probe-based examination of self-assembled monolayer properties. Beebe et al. have studied the contact resistance of aliphatic self-assembled monolayers (91). They have measured the effect of the surface linking group as well as the effect of the work function of the metal substrate on the bulk measurement. Cui et al. (92) have measured the conductance of alkanethiol monolayers, correlating conductance with the length of the alkyl chain. Zhao and Davis used a similar approach to investigate the force-dependent conduction of blue copper metalloprotein (93). Fan and co-workers used a tuning fork-based atomic force microscopy (AFM) technique to measure the conductance of key components of molecular wires (94). Bockrath and co-workers have devised a scanned probe technique based on electrostatic force microscopy capable of probing the conductance of samples without requiring attached leads (95). The technique was successfully used to probe the conductance of DNA. Jiang et al. (96) elucidated lamellar branching during the formation processes of poly(bisphenol A-co-decane) spherulites with variable-temperature AFM. They found irregular lamellar branches at 35 °C and straight lamellas formed at higher temperatures. Magonov and Yerina (97) used high-temperature AFM to investigate the surface morphology of ultrathin films of C60H122. At temperatures nearing 140 °C, there was an observed spontaneous reorientation of alkane lamellas. Schwarz et al. have reviewed the use of low-temperature AFM and the progress that has been made to date in the thrust to achieve atomic-scale resolution (98). Barner et al. have used the AFM to monitor and manipulate two individual polymer chains on a surface (99). The two polymer molecules were topologically characterized prior to and after photochemically driven covalent attachment. Hembacher et al. (100) discussed why only every other atom is visible when resolving the atoms on the graphite surface. They have recently resolved the “hidden atom” in graphite using low-temperature AFM. Calleja et al. have studied the voltage-driven dependence of water bridge formation between an AFM tip and a substrate when the two are electrically biased (101). The results are important in understanding AFM-based nanolithography experiments. Nanofabrication and Nanolithography. The use of scanning probes for surface modification has recently made great advances, contributing to advancements in data storage, nanolithography, and templating techniques for nanomanufacturing. Dip pen nanolithography (DPN) is a technique where a chemically coated atomic force microscope cantilever tip is coated with a mobile molecule and then “inks” the surface over which it scans, transferring the molecules from the cantilever to the substrate. Analytical Chemistry, Vol. 76, No. 12, June 15, 2004

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Originally demonstrated and typically used as a contact modeonly technique, Agrawal (102) performed DPN in tapping mode, where they modulated deposition of a peptide through control of the tapping amplitude. Tapping mode DPN could allow deposition on soft surfaces. Many new molecules and surfaces have been shown to be accessible with the DPN technique, including deposition of proteins (103, 104), sol-gel precursors for magnetic materials (105), oligonucleotides on metals and insulators (106), highly reactive alkoxysilanes on glass (107), and optically active inks (108). The versatility of the DPN technique has not yet been bounded, and in the future, DPN could emerge as one of the most important applications of SPM. Advances in microfabrication have benefited DPN; for example, two-dimensional arrays of cantilevers have been used to pattern a surface in a parallel fashion (109). Another advance has been the ability to affix a soft elastomeric tip to the cantilever such that highly local contact printing can be performed (110). At present, the technique has shown a resolution of 500 nm but this is by no means the lower limit. In thermomechanical data storage, a heated AFM cantilever tip forms an ultrasmall indentation into a polymer. This technology remains a high-profile application of atomic force microscopy, with several advances recently reported (111). In particular, advances in understanding tip-polymer interaction during the formation of nanometer-scale contact have aided single-indent erasing. Thermomechanical data storage using cantilevers with carbon nanotube tips has been shown to modify surfaces, achieving indentation sizes of 22 nm in diameter (112). The heat transport mechanisms during writing and reading have been closely investigated, resulting in improvements in cantilever design for enhanced reading (113) and for high-speed and low-power writing (114). New cantilevers have also been developed that include a heater for writing and a piezoresistive element for detecting deflections (115). More established SPM-based nanolithography approaches, such as SPM-enhanced local oxidation and surface scratching have made advances, particularly for application to real device fabrication. For example, one group showed the oxidation fabrication of 20-nm SiGe devices (116), and another showed the oxidation fabrication of sub-10-nm silicon wire-based devices (117). AFMbased lithography also allowed precise placement of AlGaAs/GaAS quantum dots and wires with nanometer precision (118). SPM-based electrostatic nanolithography was shown to pattern nanometer-scale dots and lines in a polymer through resistive heating of the polymer in contact with the tip (119). The technique can produce both raised and depressed regions, by operating the cantilever tip in either an attractive or repulsive contact mode. Cavallini et al. (120) imaged amide-based rotaxane films that were grown/deposited on mica or HOPG. When imaging in contact mode at loading rates less than 2 nN, the films look rather homogeneous. If the vertical load during scanning is increased above 2nN, the film becomes mechanically perturbed, and during repetitive scanning of a single line along the film, results in a series of dots along the imaging axis. This pressure-sensitive film (and others like it) could potentially be used as SPM-based data storage bins. Crook and co-workers have demonstrated a new SPM-based lithography technique called erasable electrostatic lithography 3434

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(121). This technique can generate patterns of charge on a surface with a negatively biased scanning probe tip. The proof of concept involved drawing and erasing quantum antidots. Thermal Techniques. There is at present great interest in the measurement of thermal-physical properties and temperature at extremely small length scales, for which scanning probe techniques are lending important measurements. Thermal measurements can be categorized as techniques that use standard probe microscopy measurements on a heated sample or techniques that use heat-generating or temperature-sensing cantilevers. For measurements made on heated samples, the AFM can be a sensitive tool for measuring the temperature dependence of physical properties. Phase-change phenomena in thin organic layers can have strong length-scale dependence, and AFM performed on hot plate stages can make in situ and highly local measurements of crystallization and melting. For example, hot stage AFM measurements of the phase change properties of poly(ethlyene oxide) (122) and poly(bisphenol A-co-decane) (123) showed several types of lamellar microstructure, depending upon annealing time and annealing rate. It is possible to make in situ observation of the temperature dependence of materials deposition, for example, with lithium on nickel (124), where the lithium formed tightly packed islands of diameters near 100-200 nm at temperatures ranging from 60 to 80 °C. Another group (125) observed copper growth on atomically flat MgO surfaces. The temperature dependence of near atomically thin copper island formation, diffusion, and coalescence was observed between room temperature and 350 °C. The temperature dependence of mechanical properties can be of importance for many engineering applications. AFM was used to measure the deformation response of nanofabricated silicon beams with diameters in the range of 200-800 nm (126) at temperatures up to 300 °C. The internal stress required to plastically deform the silicon was in the range of 4-7 GPa, which is 10 times higher than found in bulk samples. There have also been advances in measurements of temperature dependence of biochemical phenomena. One group (127) observed the denaturation of DNA molecules on mica. The DNA was seen to degrade from double-stranded to single-stranded between 50 and 80 °C, with full molecular decomposition at 100 °C. Another group (128) performed pull-off measurements to investigate the temperature dependence of the force required to rupture a biotin-avidin bond, finding a 50% decrease in rupture force for a heating temperature of 37 °C compared to the roomtemperature measurement. While capable of reaching a temperature as high as 500 °C, hot stage AFM is typically slow as the hot stage is quite large and likely induces thermomechanical expansion that can be difficult to mitigate. Trawick and co-workers (129) microfabricated a heater and a thermometer on the same centimeter-scale chip, reducing the overall heat load on an AFM system, reducing the heating time, and offering improvements in calibration through the integrated temperature sensor. In scanning thermal microscopy (SThM), an atomic force microscope cantilever has a temperature sensor integrated into the tip. It is possible to use SThM to perform high-resolution thermometry, for example, of microfabricated integrated circuits

or sensors (130, 131), and it is also possible to make local measurements of thermal conductivity (132, 133). One of the significant open questions in SThM is the quality of calibration techniques, limited at present by a lack of models and measurements of thermal conduction mechanisms in SThM. Shi and Majumdar (134) have made the most significant progress on this to date, making measurements of thermal transport across a 50-nm-diameter contact between a thermocouple tip and a heated surface. This study found that the surface topography can significantly affect the temperature signal in SThM, illustrating the need for extremely detailed calibration for high-precision temperature measurements with AFM. A study by Thurber (135) used magnetic resonance force microscopy to measure the temperature at the end of a cantilever tip and found that the thermal resistance of the cantilever and cantilever tip was sufficiently high such that the temperature at the end of the tip could be significantly different from the temperature in the cantilever. Thus, for SThM approaches that do not have a temperature sensor at the very end of the cantilever tip, SThM measurement error can be higher than previously thought. Finally, Lefevre (136) developed a technique for the calibration of AFM tips that have integrated heater-thermometers. Previously, only transient techniques had been available, offering the ability to measure thermal diffusivity but not thermal conductivity. Lefevre’s contribution was to offer a dc-calibration technique, which allows for steady-state temperature measurement and measurement of thermal conductivity. Biological Imaging. Today’s research on biological AFM imaging differs from earlier research in that the AFM images are much more likely to be simply an adjunct to exciting research, instead of being an end in themselves. A few of the many reviews of biological AFM include Vol. 68 of Methods in Cell Biology (137) and ref 138. Scanning Probe Evolution in Biology reviews the first 20 years of this field, in an article that presents beautiful images of the detail seen in membrane protein arrays of aquaporin and fusion pores (139). Nucleic acids are reviewed in Chromatin Fibers, One at a Time (140) and ref 141. Proteins, including membrane proteins, are reviewed in refs 142 and 143. The sections that follow are only a small selection of the many recent highlights in biological imaging by SPM. Membrane Proteins. Rhodopsin dimers in paracrystalline arrays are seen in native disk membranes of rod outer segments from the mouse retina (144). Native membranes are not readily imaged at such a high resolution by techniques other than probe microscopy. Rhodopsin is a G-protein-coupled receptor and is therefore a member of a large and important class of proteins. Prion Proteins. Prions are currently one of the most exciting classes of proteins. A single type of prion protein can polymerize into amyloid fibers with different morphologies, and these different fiber morphologies are “inherited” in the sense that seeds of one fiber type will promote the formation of fibers of the same type. An ingenious method uses AFM for distinguishing the locations and rates of fiber growth. Two prion variants were prepared. One variant contained an antibody recognition site, which gave broad fibers by AFM after antibody labeling (145). In this way, initial fiber growth using one prion variant was distinguished from subsequent fiber growth using the other prion variant. In another

prion investigation, 20-nm bands were seen along the amyloid fibrils by AFM (146). In a merger of two hot research areas, nanowires have been built by self-assembly of amyloid fibers and selective gold coating (147). This is a “bottom-up” approach for fabricating a nanostructured material. Amyloid fibrils are a good biomaterial for bottomup synthesis of nanomaterials because the amyloid fibrils, being composed of prion proteins, are unusually resistant to harsh treatments that would destroy most biomaterials. Prion proteins containing accessible cysteine residues are used for these nanostructures. The cysteine residues are covalently attached to colloidal gold particles, which are then further metallized by treatment with gold and silver salts. Lengths of the gold nanowires could be roughly controlled from the growth conditions for the amyloid fibers, to produce nanowires 60 nm to hundreds of micrometers long. Nucleic Acids. DNA papers outnumber RNA papers by about 10 to 1 in the literature on SPM of nucleic acids. It is increasingly difficult to select from among these papers, since the overall quality of the AFM research has risen so much over the past several years. “Designer” nucleic acids are being used to create “nanofabrics” with amazing patterns resembling stripes and plaids. Some of the newest patterns made with designer nucleic acids are known as “waffles” and “bar codes” (148, 149). The father of this field, Ned Seeman, has most recently developed a DNA rotary device, in collaboration with a new leader in the field, Hao Yan, and other colleagues (150). A clever sequence-specific method for coating DNA with gold uses RecA-coated single-stranded (ss) DNA complementary to a sequence on double-stranded (ds) DNA (151). Homologous recombination produces dsDNA interrupted by the bound RecAssDNA complex. The dsDNA was pretreated with glutaraldehyde to create aldehyde-derivatized DNA, which was stretched on a passivated silicon substrate. In an AgNO3 solution, silver aggregates form on the aldehydes of the unprotected dsDNA. The silver aggregates serve as catalysts for gold deposition, creating linear devices consisting of discontinuous gold nanowires. A variation of this technique is used to build a field-effect transistor, by attaching a single-walled carbon nanotube (SWNT) to the RecAcoated region of the DNA strand, using a streptavidin-functionalized SWNT on a “sandwich filling” of anti-RecA antibody and a biotinlylated secondary antibody (152). In the “device” category is a “nondevice”. Noncontact AFM scans of DNA and carbon nanotubes in contact with a gold electrode show that the carbon nanotube is a conductor, while the DNA is not a conductor and, therefore, cannot be used as a molecular wire (153). Sequence Recognition and Sequence-Dependent Structures. The inorganic mineral surface of mica shows a highly selective preference for one side of intrinsically curved DNA containing A-tracts. In DNA containing short A-tracts, the A’s tend to be positioned on one side, and the T’s tend to be positioned on the other side of the DNA molecule. This DNA preferentially binds to mica with the T-rich side facing down (154). Sequence-dependent condensation in Ni(II)-containing solutions was observed for GC-repeating and AT-repeating dsDNA. Poly(dG-dC)‚(dG-dC) (GC-DNA) condensed extensively, forming Analytical Chemistry, Vol. 76, No. 12, June 15, 2004

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toroids and condensed networks. These condensates were absent in poly(dA-dT)‚(dA-dT) (AT-DNA). The clue to understanding this work came from Kornyshev’s and Leiken’s electrostatic zipper theory of DNA condensation, coupled to the specificity of Ni(II) for binding to the N7 of guanine. (155). GC-DNA and AT-DNA are specialized examples of DNA sequence characterized by alternating pyrimidines and purines (Pyr/Pur). Long Pyr/Pur sequences occur with an unusually high frequency in eukaryotic genomes. Mirror image Pyr/Pur sequences can form triple-helical structures, which appear as thickened regions in the AFM (156). Abnormal DNA repeats are a common feature in many genetic diseases. The structures of these repeatssopen DNA loops and condensed DNA segmentsshave been visualized by AFM and used to propose a mechanism by which the abnormal repeats might cause aberrant replication (157). Sensors. Microcantilever-based sensor technologies have grown rapidly over the last several years in an attempt to develop sensitive and selective detection of biological and chemical substances and environmental factors. Much work has been done to broaden the range of substances detected from neurotoxins (158) to explosives (159) and cardiac proteins (160) to enantiomers (161). Major progress has also been made in sensor system improvements. The general scheme of microcantilever sensors is based on detecting the bending (static mode) or shifting resonance frequency (dynamic mode) of an SFM cantilever upon binding of a given substance or due to some other force on the cantilever. Microcantilever sensors do not require labeling of proteins or antibodies, which saves time and avoids adulteration of interactions (160). They have greater sensitivity to low analyte concentrations than surface acoustic wave and quartz crystal microbalance sensors (161). It has become apparent that microarrays are the preferred design because of their greater reliability through the inclusion of a reference to account for factors such as thermal drift, viscosity, nonspecific adsorption, nonspecific chemical reactions, and solution flow dynamics. In addition, multiple targets can be detected simultaneously leading to high-throughput measurements and producing distinct recognition patterns from complex mixtures, as was demonstrated by Gerber and colleagues with their “electronic nose” system (162). System advances have been made for improved accuracy and sensitivity. For example, the mass change from molecular adsorption that induces resonance frequency shift is often complicated by consequent changes in the spring constant. Cherian and Thundat have corrected for the spring constant change by concurrent measurement of resonant frequency and cantilever bending (163). Dufour and Fadel have modeled the response of microcantilevers when used as resonant-based chemical sensors (164). They have provided criteria for the optimization of cantilever-based sensors. Vidic et al. developed a system that maintains the resonance frequency of the cantilever through a closed feedback loop. The constant resonance frequency resulted in a 1000-fold increase in the quality factor and was applicable in liquid and gas phases (165). Naik and co-workers evaluated the dynamic response of a cantilever in liquid near a solid wall. The ratio of cantilever width to the height of the beam above a surface 3436

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influences measurement of added mass and viscosity damping coefficients. They recommended calibration to account for nonuniform gap height (166). Headrick et al. demonstrated enhanced sensitivity of disordered nanostructure surfaces compared to smooth microcantilever coatings (167). Modifications to cantilever coatings, such as hydrogels (168) and zeolites (169), have been used to improve selectivity. Another variation of the microcantilever sensor assesses the detachment of an agarose bead from the tip resulting from a displacement reaction (158). Domanski et al. (170) fabricated a piezoresistive cantilever with a porous silicon element and measured its response to changes in humidity. The large increase in active surface area afforded by the porous silicon structure dramatically improved the sensitivity of this sensor. Sensors have been developed to detect a wide variety of analytes for numerous applications, only a few of which can be highlighted here. Detection of trace explosives is of interest for national security. Microcantilever arrays have been designed to detect explosive vapors using a voltage pulse to instigate a miniature deflagration that is unique to the type of explosive compound (159). Microcantilever arrays have also found potential application as early and rapid medical diagnostic devices. Two cardiac biomarker proteins important for diagnosis of acute myocardial infarction have been measured using antibody-coated cantilevers (160). Chiral evaluation of R-amino acids has been achieved using smooth and nanostructured cantilever arrays modified with stereoselective antibodies. This type of sensor allows rapid quantification of enantiomeric impurities (161). Microcantilever sensor use has been extended to reactions using thermal cycling, such as polymerase chain reaction. The system quickly stabilized during temperature plateaus and remained reproducible through several cycles (171). The thermal response of the cantilever during enzymatic binding was analyzed and shown to be independent of heat generated by the enzymatic reaction (172). Miniaturization of the system onto a single chip incorporating a resonant cantilever and integrated transducers and driving circuitry was accomplished by Lange et al. by fabrication of complementary metal oxide semiconductors (CMOS) (173). This system was shown to be effective in detecting volatiles with sensitivity similar to other acoustic gas sensors and was able to detect the thickness of the polymer deposition on the cantilever array to monitor the coating procedure. Additional miniaturization of the cantilever to dimensions of 40 µm × 500 nm × 500 nm (length, width, thickness) was achieved by Davis et al. to improve both mass and spatial resolution (174). The quality factor was significantly improved, especially in a vacuum, where both the fundamental and harmonic modes were excited. The nanocantilever was fabricated using nanolithography and integrated into a CMOS circuit for resonance frequency tracking. Microcantilever arrays have also been integrated into microfluidic systems (175). Significantly more power efficient systems have been developed by Adams and colleagues (176). Microcantilever sensor systems will continue to evolve with improvements in sensitivity and specificity, as well as smaller size and cost, for introduction into fields including medical diagnostics, environmental monitoring, food quality, and chemical detection.

FORCE SPECTROSCOPY Chemical and Polymer Applications. Humphris et al. have presented a new force spectroscopic technique called transverse dynamic force microscopy (177). This approach allows one to monitor shifts in the cantilever drive frequency, phase, and amplitude during a single pulling event. Lim and co-workers used sample modulation force spectroscopy to study the effects of tip roughness and geometry in the AFM measurements of solvation (178). The sample modulation response curves allow one to make a direct measurement of the interaction stiffness (force gradient) as a function of tip-sample distance, and this technique is capable of measuring both repulsive and attractive solvation potentials in a single approach. Kappl and Butt (179) have presented a thorough review of colloidal probe force spectroscopic measurements. Zhang and Zhang have written a comprehensive review covering many of the force spectroscopic techniques that have been used to probe the local mechanical properties of polymers (180). Akhremitchev et al. present a thorough review of the use of scanning probe microscopy in characterizing the function and aging of textured, minimally adhesive polymer surfaces (181). Chen and co-workers have claimed to be the first to successfully measure the lateral Casimir force (182). A full understanding of the Casimir force could play a large part in applying/utilizing this force in microscale devices. Kudera et al. used single-molecule force spectroscopy to investigate the mechanical properties of individual bis(terpyridine)ruthenium(II) complexes (183). Marszalek et al. observed the chair-boat transitions of a single polysaccharide molecule using force spectroscopy (184). Zapotoczny and co-workers measured the rupture forces of individual β-cyclodextrin-ferrocene hostguest complexes in an aqueous medium (185). The observed rupture force for the host-guest conjugate was 55 ( 10 pN. Tivanski et al. have measured both conduction and adhesion forces simultaneously of a polythiophene (186). During chemical force microscopy measurements between selfassembled layers of methoxytri(ethylene glycol) and methylterminated alkanethiols, Dicke and Haehner have shown that water is crucial for the stability of the surface charge that is associated with organic films and acts as a template for hydroxyl adsorption (187). Zepeda et al. (188) have determined the energy barriers of alternative interfaces at variable temperatures. This exhaustive endeavor reveals an important role of solvation in many of the systems probed using chemical force microscopy. Green has reviewed the recent progresses of using a new inverted AFM technique (189), which can carry out combinatorial atomic force microscopy (190). Connell et al. used adhesion mapping to locate nanometerscale oil droplets existing on a polystyrene surface (191). Force curve mapping was used to gently probe the surface of the fluid droplets, and through automated analysis of the force curves, the true topography and microscopic contact angle of the droplets were determined. Eaton and co-workers utilized topographic imaging, adhesion force mapping, and indentation mapping to investigate the surface of an elastomeric filled silicone coating. Topographic observations revealed randomly distributed protruding features, which lead to a source of error when assessing the nanoscale stiction (192). The source of peak broadening in the

adhesion force histograms acquired in typical chemical force microscopy experiments was further characterized by Sato and co-workers on homogeneous and nonhomogeneous surfaces (193). To aid in the design of nanotube-based composite systems, Bottomley et al. have examined adhesive interactions between an alkanethiol-modified gold-coated tip and the sidewall of a singlewalled carbon nanotube (194). Akabori and co-workers have monitored the mechanical properties of a polystyrene film using lateral force microcopy while the film was subjected to a temperature ramp (195). The authors were able to observe a thinning-induced relaxation process, called surface β-relaxation. Bliznyuk et al. investigated the surface glass transition temperature of several molecular weight films of polystyrene by studying the hysteresis in the loading-unloading cycles of force-distance curves (196). Harmon and co-workers have measured the change in elastic modulus of hydrogel films as the films were gradually heated past their transition point (197). They also performed the inverse of the previous experiment where a hydrogel sphere was affixed to the cantilever tip and its compressibility measured during its phase transition. Hodges has presented a very thorough review of AFM-based mechanical testing of polymer films in liquid environments (198). Bunker et al. used an interfacial force microscope (IFM) to study the surface chemistry changes that occur during the photoactivated opening and closing of rings in tethered spiropyran monolayers (199). This study demonstrated the ability of the IFM to probe differences in surface polarity, providing insight into the impact of surface charge on electrokinetic flow in microfluidic systems. Houston and Kim Hyun reviewed the use of the IFM to characterize the adhesion, friction, and mechanical properties of self-assembled systems on gold surfaces (200). Hugel et al. performed force spectroscopic measurements on a polymer composed of bistable photosensitive azobenzenes (201). The polymers were exposed to ultraviolet light that induced lengthening and contraction of individual polymers through photoinduced switching of the azo groups between their trans and cis configurations. The polymer was found to contract against an external force acting along the polymer backbone, thus delivering mechanical work. Rabe and co-workers used atomic force acoustic microscopy (202) and monitored the amplitude and phase of the cantilever vibration as well as the shift of the cantilever resonance frequencies to interpret the local tip-sample contact stiffness and used this information to generate elastic maps of a surface. Benmouna and co-workers (203) have developed a unique imaging mode that is capable of measuring the thermal resonance of the cantilever as it travels above a latex surface. This approach provides topologically defined “maps” of the cantilever’s resonance as it travels above a substrate. They have also used a similar setup to monitor the mechanical pulling of acrylic fibers (204). Biological Applications. Reviews of biological force spectroscopy include two edited books. One is a volume of Methods in Cell Biology (137). The other is reprinted from the Journal of Muscle Research and Cell Motility (205); it is well illustrated with useful figures and is good for both novices and specialists in the field. Current Opinion reviews include well-referenced articles about force spectroscopy of DNA and RNA (206), in microbiology (207), single-molecule folding (208), and a dramatic image of AFM Analytical Chemistry, Vol. 76, No. 12, June 15, 2004

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cantilevers with a human hair (209). A leader in the field has written a detailed review (210). Another review, comparing AFM and FRET, concludes that “the route to fast unfolding, whether chemical or mechanical, is well-defined and straightforward, whereas the many possible conformational routes to folding result in very slow dynamics” (211). Proteins. Titin. Single-molecule force spectroscopy was pioneered on titin, a multidomain protein from vertebrate striated and cardiac muscle cells. This protein plays a major role in generating the elasticity and passive tension in muscle cell sarcomeres. The I-band part of titin is responsible for its elastic properties and consists of tandem repeats of immunoglobulin (Ig)like domains at distal and proximal parts, while the central part is occupied by linker domains (PEVK and some others) with structures that are not well-understood. Force spectroscopy on polyproteins, assemblies of identical protein domains placed in series, leads to force spectra with repetitive features that permit the identification and, thus, the investigation of mechanical properties of individual protein domains. Thus, numerous recombinant polyproteins, containing tandems of one or a few types of Ig domains from titin, were produced by genetic engineering in order to mimic native titin but in a simplified version. Force spectroscopy has shown that the Ig domains from the proximal part of titin could be unfolded by stretching forces in the150200-pN range with all Ig domains demonstrating similar mechanical strength (212). In contrast, the Ig domains from the distal part of titin required stronger stretching forces (150-350 pN) and have shown a strong hierarchy in their mechanical strength. Similar data on the recombinant polyproteins with Ig domains interspersed with one or a few linker domains (PEVK or N2B) demonstrated that these linker domains behave as entropic springs without any unfolding pattern. They are easily unfolded with only small forces. The overall mechanical extension of titin under physiological conditions was reconstituted from experimental data for its individual domains and was described as an extension of linker domains, followed by unraveling of a few proximal Ig domains. The feasibility of such a projection of individual Ig-domain properties onto the whole native protein was confirmed in recent studies on the force-induced unfolding kinetics of Ig domains in relation to their neighborhood (213). This research demonstrated that Ig domains I28, I29, I30, I31, and I32 show significant differences in unfolding kinetics and that these differences were influenced little, or not at all, by the surrounding environment. The authors also proposed that observed differences in kinetics for Ig domains could be important in preventing the misfolding of titin during muscle relaxation. Fibronectin. Fibronectin (FN) is one of the naturally occurring polyproteins that consists of numerous individually folded domains belonging to three distinct types (FNI, FNII, FNIII). Cells produce this protein as a part of their extracellular matrix (ECM). Mechanical deformations of some of the FNIII domains inside FN can trigger many physiological processes inside cells and can regulate ECM assembly and its interactions with the cells. By using AFM force spectroscopy, sequential increases in unfolding forces were observed during mechanical unfolding of native FN molecules (214). Forces required to unfold native FN ranged from 80 pN for the weakest domain to 200 pN for the strongest one. 3438

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The mechanical stability of some individual FNIII domains was elucidated by force spectroscopy on engineered polyproteins, containing multiple copies of the corresponding FNIII domain, alternating with identical I27 domains from titin. It was found that domains in position 10 and 13 (10FNIII and 13FNIII), which contain cell-binding signals, are mechanically the weakest domains. In contrast, domains 1FNIII and 2FNIII proved to be mechanically the strongest among all FNIII domains. Interestingly, 1FNIII domain (the domain that has the first position inside FN) was mechanically unstable due to an unfolded or misfolded state when alternating with identical I27 domains in the synthetic polyprotein. However, the mechanical stability of 1FNIII domain was restored upon substitution of I27 domains by 2FNIII domains (native neighbor of 1FNIII domain in FN), suggesting a chaperone-like activity of 2FNIII domain toward 1FNIII domain. Ubiquitin. By using engineered polyproteins assembled from individual ubiquitin proteins, researchers were able to directly observe the folding of individual ubiquitin proteins (215, 216). Force-clamp spectroscopy was used to directly observe folding trajectories of ubiquitin under a constant force load (216). At least four folding stages could be resolved by this method after the initial unfolding of a ubiquitin polyprotein under a high stretching force in the 100-120 pN range. First, immediately upon relaxing from high stretching forces down to low quenching forces in the 15-50 pN range (to facilitate refolding of polyprotein domains), the protein backbone spontaneously collapsed as a result of its elastic recoil. The following two stages have shown a continuation of polyprotein backbone collapse (folding) but at a much slower speed (in the range of a few seconds). Folding stage number three was different from stage number two in that it had a faster collapse rate. The authors note that it was problematic to discriminate between these two stages on some of the experimental curves. The last, the fourth stage, pictured a fast collapse of semifolded protein into its fully folded state. The duration of the whole folding event was dependent on the contour length of the polyprotein and on the level of quenching force. Interestingly, all four folding stages had relatively large fluctuations (at least a few nanometers) in the end-to-end length of the protein. These fluctuations start only after unfolding of the protein has happened, and they calm down immediately after the folding process is finished. Carrion-Vazquez and coauthors found (215) that the forces required to unfold ubiquitin domains in Lys48-C-linked ubiquitin chains are more than two times smaller than the forces required for N-C-linked ubiquitin chains. These findings indicate that the force spectra of mechanical unfolding of this protein are strongly dependent on the place and direction of the applied mechanical stretching forces. Membrane Proteins. Force spectroscopy was used to study stability and unfolding pathways of bacteriorhodopsin (BR) molecules in the native purple membranes of Halobacterium salinarum. Sequential unfolding of transmembrane domains in BR can be achieved by this method, and the contour length of each unfolded domain can be resolved with relatively high precision (217). The authors studied pH effects, temperature, and site mutations on the stability of individual transmembrane domains. Extracellular loops connecting transmembrane domains were shown to contribute significantly to the stability of BR molecules and to serve as energy barriers in the mechanical unfolding of

this protein (217, 218). Interestingly, removal of the retinal group from the protein backbone resulted in the formation of an additional unfolding barrier at that place in the protein. Ligand-Receptor Interactions. The sensitivity of AFM instruments allows measurement of the forces between single pairs of ligand and receptor molecules. Nevo and coauthors used AFM-based force spectroscopy to study interactions between small GTPase Ran and its nuclear receptor protein importin β1 (219). They have shown that interactions between importin β1 and Ran protein in the GTP-bound state results in the formation of a ligand-receptor complex that exists as a population of two energetically distinct forms with, respectively, higher and lower stretching strengths. Interestingly, the application of an external force shifted equilibrium toward a higher strength form of the complex. In contrast, the ligand-receptor complex between importin β1 and Ran protein in the GDP-bound state had only a single energetic state. Protein Networks (Mesostructures). Although the majority of force spectroscopy research is focused on experiments at the single-molecule level, more complex structures with numerous molecules organized into networks or mesostructures can also be successfully studied by this technique. Force spectroscopy, together with AFM imaging, gave new insight into the organization of spider dragline silk fibers (220, 221). The design of the recombinant protein used in this study was based on highly repetitive sequences from a native spider dragline silk protein. This recombinant protein had 16 identical repetitive modules, and each module was predicted to have a length of 14 nm when extended. AFM images showed that this protein spontaneously formed segmented nanofibers on a mica surface. Force spectroscopy on these nanofibers revealed repetitive spectra with multiple ruptures of “sacrificial bonds”. These rupture events had similar rupture forces and rupture lengths in multiples of 14 nm. The authors proposed a model based on these results and described the possible folding of individual spider dragline silk proteins as well as their organization into silk nanofibers. Force spectroscopy data from native spider capture silk fibers demonstrated an exponential increase in force upon stretching (222) with multiple ruptures of “sacrificial bonds”. A remarkable self-healing ability of capture silk fibers was demonstrated in these experiments. The authors proposed a model that describes capture silk fibers as a cascade of springs with restricted extendable length and with identical or similar spring constants. Living Cells. Mechanical changes on the cell surface can be studied with force spectroscopy under physiological conditions. Changes in the rigidity of the cell surfaces of Pseudomonas putida KT2442 cells (223, 224) were dependent on the ionic strength of the surrounding environment and were attributed to changes in the flexibility of cell surface biopolymers. Another study measured the mechanical forces behind the adhesion of bacterial cells Escherichia coli JM109 to the surface (225). These adhesion forces were related to the content and the electrochemical status of lipopolysaccharides in the cell walls. Selectin-glycoprotein complexes from cell walls are responsible for the attachment and rolling of leukocytes on vascular surfaces. AFM-based force measurements made possible direct detection, measurement, and discrimination between the catchand slip-types of bonds in selectin-ligand complex (226). The

dual character of the forces in selectin-ligand complex seems to be critical for regulating cell adhesion under variable mechanical stress. Nucleic Acids. Force changes in single-nucleic acid molecules have been successfully studied by AFM instruments with resolution as high as 5 pN (206). The forced extension of a single dsDNA molecule gives a force versus extension curve with characteristic features. The large force plateau at 65 pN describes the progressive loss of double-helical structure of dsDNA due to the loss of base-stacking interactions under applied forces. The second force plateau near 150 pN describes the progressive melting of dsDNA into ssDNA. The interactions of dsDNA with small-molecule drugs can influence the corresponding dsDNA force spectra (227). Force spectra for dsDNA differed from force spectra for drug-bound dsDNA, with differences that were characteristic for the particular drug-DNA interaction. The authors demonstrate that these changes occur in a concentration-dependent manner. The results indicate that AFM-based force spectroscopy can be effectively used to study basic mechanisms of DNA-drug interactions, as well as being utilized in massive screening, sensing applications, or both. Conventional AFM-based rupture force spectroscopy has been extended (228) to the "differential force” test, in which a reference nucleic acid duplex is placed in series with the nucleic acid duplex of interest. When force is applied to this system, the weaker bond will have higher chance of rupturing. By determining which nucleic acid duplex survived during the extension cycle, one should be able to directly compare the rupture forces for the different DNA structures. The sensitivity of this method was high enough to detect a single base pair mismatch in a 20-bp duplex, as well as to distinguish between shear and unzipping ruptures between analogous DNA duplexes. dsRNA molecules demonstrate force spectra that are similar, but not identical, to dsDNA (229). The first and the only force plateau on dsRNA force versus extension spectra describes the transition of A′-form RNA into S-form. Average transition forces at this plateau were 20% higher for dsRNA molecules when compared to dsDNA with similar base compositions. This increase in transition forces can be due to the significantly higher energy of base-stacking interactions in A′-form dsRNA when compared to B-form dsDNA. Interestingly, the distribution of the plateau forces was also much broader among individual dsRNA molecules. The melting plateau was not detected in dsRNA force spectra, suggesting that melting forces for dsRNA probably exceed 200 pNsthe typical disengagement force between nucleic acid molecules and the AFM tip. CHARACTERIZATION OF CARBON NANOTUBES Scanning probe techniques are invaluable tools in the characterization and manipulation of carbon nanotubes. The utility of STM in probing the topological and electrical properties of carbon nanotubes has been reviewed (230). It has been demonstrated that an AFM tip can be used to cut either multiwalled (231) or single-walled (232) nanotubes, affording a straightforward method for removing defect-laden portions or creating nanotube-fragment quantum dots. Woodside and McEuen used the AFM to study single-electron motion in nanotube quantum dots (233). They remind us that although scanned probe techniques have great Analytical Chemistry, Vol. 76, No. 12, June 15, 2004

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sensitivity, these techniques alter the native properties of the system under measurement. Measurement of the impact of mechanical stress on the electrical properties of carbon nanotubes has been a research focus. Bozovic et al. (234) studied the effect of strain on the electrical properties of single-walled carbon nanotubes. Using scanning gate microscopy, they showed that defect sites, created by mechanical deformation, cause a significant decrease in electron transport along the tube. Kim et al. (235) elucidated the electrical properties of a MWNT while it was being mechanically deformed using nanomanipulators, Thelander and Samuelson (236) made electrical contact to both ends of a nanotube and then used an AFM tip to manipulate the nanotube on the surface. Tip-induced bending of the nanotube caused measurable decreases in the tube conductance. Minot and co-workers (237) have shown that the band structure of a carbon nanotube can be dramatically altered by mechanical strain. They suspended a nanotube over a trench and clamped both ends with electrical contacts. An AFM tip was brought into contact with the tube and used to vary the strain on the nanotube. Nanotube conductance was monitored as a function of strain. Their results demonstrate that strain can open a band gap in a metallic nanotube and modify the band gap in a semiconducting nanotube. Williams and co-workers present a unique AFM-based technique for measuring the torsional stiffness of multiwalled nanotubes (238). They found that the torsional stiffness of a multiwalled nanotube becomes larger during repeated torsional strains. Chen et al. bridged a carbon nanocoil between two cantilever tips (239). The nanocoil was mechanically pulled, and the tensile properties of the coil were evaluated. The nanocoil behaved like an elastic spring with a spring constant of 0.12 N/m in the low-strain region. Nanotubes filled with spherical molecules are known as “nanopeapods”. STM can be used to visualize the location of the inclusion compounds in the peapod (240). Conductance along the backbone of the nanopeapod was measured; a drop in band gap energy was found at locations where fullerenes resided in the tube (241, 242). Theoretical treatment of the conduction in these nanostructures has been presented ((243). Carbon nanotubes are thermally and electrically conductive materials with high tensile strength. Their incorporation into polymer composites portends of next-generation lightweight, highstrength materials and motivates research in this area. Technical challenges that presently inhibit realization of new materials with these properties include control of the orientation and dispersion of the nanotube within the polymer matrix. Characterization of the defect density of nanotubes in polymer composites is important for the development of next-generation composites. Techniques enabling visualization of nanotube dispersion into polymeric matrixes include current sensing AFM and magnetic force microscopy (244, 245). Wagner’s group has measured the force required to detach a nanotube from a polymer by laterally scratching the nanotube out of the polymer using an AFM tip (246). More recently, they embedded a nanotube-tipped cantilever in a polymer and then measured the force required to pull the nanotube out (247). After removal, images of the polymer surface revealed the location where the nanotube was previously embedded. 3440 Analytical Chemistry, Vol. 76, No. 12, June 15, 2004

Ding et al. (248) have observed polymer sheathing in a carbon nanotube-polycarbonate composite. Contact of the polymer sheath with an AFM tip perturbs the polymer multilayer structure, and the polymer sheath rolls into a ball. These observations suggest the importance of both nanotube-polymer and polymerpolymer interactions in enhancing the performance of nanotubepolymer composites. In summary, SPM has revolutionized the way we perceive atoms on surfaces and molecules adherent to them. With proximal probes, scientists and engineers can manipulate matter on the atomic and molecular scale, measure intermolecular forces, initiate chemical reactions, and determine the mechanical properties of single molecules. SPM is the cornerstone upon which nanotechnology is based. Future innovations and advances in nanotechnology will educe from the diligence, ingenuity, and creativity of practitioners of scanning probe microscopy. Mark A. Poggi is a graduate student completing his doctoral degree in analytical chemistry at the Georgia Institute of Technology. He obtained his baccalaureate degree in chemistry at Michigan State University. His current research interests include microcantilever array-based sensors and the mechanical and interfacial properties of carbon nanotubes. Elizabeth D. Gadsby is a research program leader at Kimberly-Clark Corporation in Roswell, GA. She obtained her baccalaureate degree in microbiology at the University of Florida and will shortly complete her doctoral degree in biochemistry at the Georgia Institute of Technology. Her dissertation research focused on AFM of drug-DNA interactions. Lawrence A. Bottomley is a professor of chemistry at the Georgia Institute of Technology. He obtained his B.S. in chemistry at California State University, Fullerton, and his Ph.D. in analytical chemistry at the University of Houston. His current research interests include the biological and nanotechnological applications of scanning probe microscopy and electrochemistry. William P. King is an Assistant Professor of Mechanical Engineering at the Georgia Institute of Technology. He obtained his B.M.E. at the University of Dayton and both an M.S. and Ph.D. in mechanical engineering from Stanford University. Dr. King spent eighteen months with IBM Research in Zurich, Switzerland, prior to joining the faculty at Georgia Tech. Dr. King’s research interests focus on transport physics at small scales, nanoscale thermal processing, novel manufacturing of soft materials for MEMS, and micro/biofluidics. Emin Oroudjev is a researcher at the University of California Santa Barbara. He obtained his master’s degree in physiology at Moscow State University, Moscow, USSR, and his doctoral degree at the Institute of Plant Physiology, Moscow, Russia. His current research interests include the biological and nanotechnological applications of scanning probe microscopy. Helen Hansma is a Research Biophysicist at the University of California in Santa Barbara. She obtained her B.A., M.A., and Ph.D. at Earlham College, UC Berkeley, and UC Santa Barbara, respectively. Her current research interests include new applications for probe microscopy of biomaterials.

LITERATURE CITED (1) Manning, L.; Rogers, B.; Jones, M.; Adams, J. D.; Fuste, J. L.; Minne, S. C. Rev. Sci. Instrum. 2003, 74, 4220-4222. (2) Miyahara, Y.; Deschler, M.; Fujii, T.; Watanabe, S.; Bleuler, H. Appl. Surface Sci. 2002, 188, 450-455. (3) Rogers, B.; York, D.; Whisman, N.; Jones, M.; Murray, K.; Adams, J. D.; Sulchek, T.; Minne, S. C. Rev. Sci. Instrum. 2002, 73, 32423244. (4) Brook, A. J.; Bending, S. J.; Pinto, J.; Oral, A.; Ritchie, D.; Beere, H.; Henini, M.; Springthorpe, A. Appl. Phys. Lett. 2003, 82, 35383540. (5) Grow, R. J.; Minne, S. C.; Manalis, S. R.; Quate, C. F. J. Microelectromechan. Syst. 2002, 11, 317-321. (6) Delzeit, L.; Nguyen, C. V.; Stevens, R. M.; Han, J.; Meyyappan, M. Nanotechnology 2002, 13, 280-284. (7) Yenilmez, E.; Wang, Q.; Chen, R. J.; Wang, D.; Dai, H. Appl. Phys. Lett. 2002, 80, 2225-2227. (8) Yang, Y.; Zhang, J.; Nan, X.; Liu, Z. J. Phys. Chem. B 2002, 106, 4139-4144. (9) Nishino, T.; Ito, T.; Umezawa, Y. Anal. Chem. 2002, 74, 42754278.

(10) Snow, E. S.; Campbell, P. M.; Novak, J. P. J. Vac. Sci. Technol., B 2002, 20, 822-827. (11) Bale, M.; Palmer, R. E. J. Vac. Sci. Technol., B 2002, 20, 364369. (12) Chow, E. M.; Yaralioglu, G. G.; Quate, C. F.; Kenny, T. W. Appl. Phys. Lett. 2002, 80, 664-666. (13) Aigouy, L.; De Wilde, Y.; Mortier, M. Appl. Phys. Lett. 2003, 83, 147-149. (14) Crozier, K. B.; Fletcher, D. A.; Kino, G. S.; Quate, C. F. J. Microelectromech. Syst. 2002, 11, 470-478. (15) Lee, Y.; Ding, Z.; Bard, A. J. Anal. Chem. 2002, 74, 3634-3643. (16) Lee, Y.; Bard, A. J. Anal. Chem. 2002, 74, 3626-3633. (17) Hughes, W. L.; Wang, Z. L. Appl. Phys. Lett. 2003, 82, 28862888. (18) Nilsson, S. G.; Sarwe, E. L.; Montelius, L. Appl. Phys. Lett. 2003, 83, 990-992. (19) Lee, D. W.; Despont, M.; Drechsler, U.; Gerber, C.; Vettiger, P.; Wetzel, A.; Bennewitz, R.; Meyer, E. J. Microelectron. Eng. 2003, 67-68, 635-643. (20) Sulchek, T.; Yaralioglu, G. G.; Quate, C. F.; Minne, S. C. Rev. Sci. Instrum. 2002, 73, 2928-2936. (21) Akiyama, T.; Staufer, U.; de Rooij, N. F. Rev. Sci. Instrum. 2002, 73, 2643-2646. (22) Stark, R. W.; Heckl, W. M. Rev. Sci. Instrum. 2003, 74, 51115114. (23) Zahl, P.; Bierkandt, M.; Schroder, S.; Klust, A. Rev. Sci. Instrum. 2003, 74, 1222-1227. (24) Trawick, M. L.; Megens, M.; Harrison, C.; Angelescu, D. E.; Vega, D. A.; Chaikin, P. M.; Register, R. A.; Adamson, D. H. Scanning 2003, 25, 25-33. (25) Kindt, J. H.; Thompson, J. B.; Viani, M. B.; Hansma, P. K. Rev. Sci. Instrum. 2002, 73, 2305-2307. (26) Schaffer, T. E. J. Appl. Phys. 2002, 91, 4739-4746. (27) Onaran, A. G.; Degertekin, F. L.; Hadimioglu, B. Appl. Phys. Lett. 2002, 80, 4063-4065. (28) Arnold, W.; Hirsekorn, S.; Kopycinska, M.; Rabe, U.; Reinstaedtler, M.; Scherer, V. Proc. SPIE Int. Soc. Opt. Eng. 2002, 4703, 5364. (29) Pfeiffer, O.; Bennewitz, R.; Baratoff, A.; Meyer, E.; Grutter, P. Phys. Rev. B: Condens. Matter 2002, 65, 161403/161401161403/161404. (30) Alcaraz, J.; Buscemi, L.; Puig-de-Morales, M.; Colchero, J.; Baro, A.; Navajas, D. Langmuir 2002, 18, 716-721. (31) Piner, R.; Ruoff, R. S. Rev. Sci. Instrum. 2002, 73, 3392-3394. (32) Fujisawa, S.; Ogiso, H. Rev. Sci. Instrum. 2003, 74, 5115-5117. (33) Buh, G. H.; Kopanski, J. J. Appl. Phys. Lett. 2003, 83, 2486-2488. (34) Dubreuil, F.; Daillant, J.; Guenoun, P. Langmuir 2003, 19, 84098415. (35) Burns, A. R. Langmuir 2003, 19, 8358-8363. (36) Hu, D.; Micic, M.; Klymyshyn, N.; Suh, Y. D.; Lu, H. P. Rev. Sci. Instrum. 2003, 74, 3347-3355. (37) Noy, A.; Huser, T. R. Rev. Sci. Instrum. 2003, 74, 1217-1221. (38) Fukushima, K.; Kawai, S.; Saya, D.; Kawakatsu, H. Rev. Sci. Instrum. 2002, 73, 2647-2650. (39) Browne, M. T.; Charalambous, P.; Burge, R. E.; Yuan, X. C. Ultramicroscopy 2002, 92, 221-232. (40) Bondarenko, S. I.; Nakagawa, N.; Shablo, A. A.; Pavlov, P. P. Physica B (Amsterdam) 2003, 329-333, 1512-1513. (41) Choi, K. H.; Friedt, J. M.; Frederix, F.; Campitelli, A.; Borghs, G. Appl. Phys. Lett. 2002, 81, 1335-1337. (42) Hayden, O.; Bindeus, R.; Dickert, F. L. Meas. Sci. Technol. 2003, 14, 1876-1881. (43) Friedt, J. M.; Francis, L.; Choi, K. H.; Frederix, F.; Campitelli, A. J. Vac. Sci. Technol., A 2003, 21, 1500-1505. (44) Giessibl, F. J. Rev. Mod. Phys. 2003, 75, 949-983. (45) Hofer, W. A.; Foster, A. S.; Shluger, A. L. Rev. Mod. Phys. 2003, 75, 1287-1331. (46) Balantekin, M.; Atalar, A. Phys. Rev. B: Condens. Matter 2003, 67, 193404. (47) Boisgard, R.; Aime, J. P.; Couturier, G. Appl. Surf. Sci. 2002, 188, 363-371. (48) Rodriguez, T. R.; Garcia, R. Appl. Phys. Lett. 2003, 82, 48214823. (49) Su, C.; Huang, L.; Kjoller, K.; Babcock, K. Ultramicroscopy 2003, 97, 135-144. (50) Stark, M.; Stark, R. W.; Heckl, W. M.; Guckenberger, R. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 8473-8478. (51) Lee, S. I.; Howell, S. W.; Raman, A.; Reifenberger, R. Ultramicroscopy 2003, 97, 185-198. (52) Hoffmann, P. M. Appl. Surf. Sci. 2003, 210, 140-145. (53) Couturier, G.; Boisgard, R.; Nony, L.; Aime, J. P. Rev. Sci. Instrum. 2003, 74, 2726-2734. (54) Chang, W.-J.; Chu, S.-S. Phys. Lett. A 2003, 309, 133-137. (55) Bardos, D. C. Surf. Sci. 2002, 517, 157-176. (56) Bedrov, D.; Smith, G. D. J. Chem. Phys. 2003, 118, 6656-6663. (57) Biesheuvel, P. M. Langmuir 2002, 18, 5566-5571. (58) Butt, H.-J.; Franz, V. Phys. Rev. E: Statistical, Nonlinear, Soft Matter Phys. 2002, 66, 031601. (59) Fraxedas, J.; Garcia-Manyes, S.; Gorostiza, P.; Sanz, F. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 5228-5232. (60) Patrick, D. L.; Flanagan, J. F.; Kohl, P.; Lynden-Bell, R. M. J. Am. Chem. Soc. 2003, 125, 6762-6773.

(61) Dean, D.; Seog, J.; Ortiz, C.; Grodzinsky, A. J. Langmuir 2003, 19, 5526-5539. (62) Dudko, O. K.; Filippov, A. E.; Klafter, J.; Urbakh, M. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 11378-11381. (63) Leng, Y.; Jiang, S. J. Am. Chem. Soc. 2002, 124, 11764-11770. (64) Friedsam, C.; Wehle, A. K.; Kuehner, F.; Gaub, H. E. J. Phys.: Condens. Matter 2003, 15, S1709-S1723. (65) Ballato, A. Ceram. Trans. 2003, 136, 533-554. (66) Green, C. P.; Sader, J. E. J. Appl. Phys. 2002, 92, 6262-6274. (67) Sader, J. E.; Sader, R. C. Appl. Phys. Lett. 2003, 83, 3195-3197. (68) Gotszalk, T.; Grabiec, P.; Rangelow Ivo, W. Ultramicroscopy 2003, 97, 385-389. (69) Jericho, S. K.; Jericho, M. H. Rev. Sci. Instrum. 2002, 73, 24832485. (70) Burnham, N. A.; Chen, X.; Hodges, C. S.; Matei, G. A.; Thoreson, E. J.; Roberts, C. J.; Davies, M. C.; Tendler, S. J. B. Nanotechnology 2003, 14, 1-6. (71) Barth, J. V.; Weckesser, J.; Trimarchi, G.; Vladimirova, M.; De Vita, A.; Cai, C.; Brune, H.; Guenter, P.; Kern, K. J. Am. Chem. Soc. 2002, 124, 7991-8000. (72) Berner, S.; De Wild, M.; Ramoino, L.; Ivan, S.; Baratoff, A.; Guentherodt, H. J.; Suzuki, H.; Schlettwein, D.; Jung, T. A. Phys. Rev. B: Condens. Matter 2003, 68, 115410. (73) De Wild, M.; Berner, S.; Suzuki, H.; Yanagi, H.; Schlettwein, D.; Ivan, S.; Baratoff, A.; Guentherodt, H.-J.; Jung, T. A. ChemPhysChem 2002, 3, 881-885. (74) Miyake, K.; Yasuda, S.; Harada, A.; Sumaoka, J.; Komiyama, M.; Shigekawa, H. J. Am. Chem. Soc. 2003, 125, 5080-5085. (75) De Feyter, S.; De Schryver, F. C. Chem. Soc. Rev. 2003, 32, 139150. (76) Bode, M.; Kubetzka, A.; Pietzsch, O.; Wiesendanger, R. Surf. Sci. 2002, 514, 135-144. (77) Downes, A.; Guaino, P.; Dumas, P. Appl. Phys. Lett. 2002, 80, 380-382. (78) Ding, Y.; Micheletto, R.; Hanada, H.; Nagamura, T.; Okazaki, S. Rev. Sci. Instrum. 2002, 73, 3227-3231. (79) Miyatake, Y.; Nagamura, T.; Hattori, K.; Kanemitsu, Y.; Daimon, H. Jpn. J. Appl. Phys., Part 1 2003, 42, 4848-4851. (80) Kubetzka, A.; Pietzsch, O.; Bode, M.; Wiesendanger, R. Appl. Phys. A: Mater. Sci. Process. 2003, 76, 873-877. (81) Hofer, W. A. Prog. Surf. Sci. 2003, 71, 147-183. (82) Voigt, P. U.; Koch, R. J. Appl. Phys. 2002, 92, 7160-7167. (83) Okawa, Y.; Aono, M. Surf. Sci. 2002, 514, 41-47. (84) Kueng, A.; Kranz, C.; Lugstein, A.; Bertagnolli, E.; Mizaikoff, B. Angew. Chem., Int. Ed. 2003, 42, 3238-3240. (85) Katemann, B. B.; Schulte, A.; Calvo, E. J.; Koudelka-Hep, M.; Schuhmann, W. Electrochem. Commun. 2002, 4, 134-138. (86) Zhang, J.; Unwin, P. R. Phys. Chem. Chem. Phys. 2002, 4, 38143819. (87) Liljeroth, P.; Johans, C.; Slevin, C. J.; Quinn, B. M.; Kontturi, K. Anal. Chem. 2002, 74, 1972-1978. (88) Treutler, T. H.; Wittstock, G. Electrochim. Acta 2003, 48, 29232932. (89) Sklyar, O.; Wittstock, G. J. Phys. Chem. B 2002, 106, 7499-7508. (90) Zoski, C. G.; Aguilar, J. C.; Bard, A. J. Anal. Chem. 2003, 75, 2959-2966. (91) Beebe, J. M.; Engelkes, V. B.; Miller, L. L.; Frisbie, C. D. J. Am. Chem. Soc. 2002, 124, 11268-11269. (92) Cui, X. D.; Primak, A.; Zarate, X.; Tomfohr, J.; Sankey, O. F.; Moore, A. L.; Moore, T. A.; Gust, D.; Nagahara, L. A.; Lindsay, S. M. J. Phys. Chem. B 2002, 106, 8609-8614. (93) Zhao, J.; Davis, J. J. Nanotechnology 2003, 14, 1023-1028. (94) Fan, F.-R. F.; Yang, J.; Cai, L.; Price, D. W., Jr.; Dirk, S. M.; Kosynkin, D. V.; Yao, Y.; Rawlett, A. M.; Tour, J. M.; Bard, A. J. J. Am. Chem. Soc. 2002, 124, 5550-5560. (95) Bockrath, M.; Markovic, N.; Shepard, A.; Tinkham, M.; Gurevich, L.; Kouwenhoven, L. P.; Wu, M. W.; Sohn, L. L. Nano Lett. 2002, 2, 187-190. (96) Jiang, Y.; Yan, D.-D.; Gao, X.; Han, C. C.; Jin, X.-G.; Li, L.; Wang, Y.; Chan, C.-M. Macromolecules 2003, 36, 3652-3655. (97) Magonov, S. N.; Yerina, N. A. Langmuir 2003, 19, 500-504. (98) Schwarz, A.; Schwarz, U. D.; Langkat, S.; Holscher, H.; Allers, W.; Wiesendanger, R. Appl. Surf. Sci. 2002, 188, 245-251. (99) Barner, J.; Mallwitz, F.; Shu, L.; Schluter, A. D.; Rabe, J. P. Angew. Chem., Int. Ed. 2003, 42, 1932-1935. (100) Hembacher, S.; Giessibl, F. J.; Mannhart, J.; Quate, C. F. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 12539-12542. (101) Calleja, M.; Tello, M.; Garcia, R. J. Appl. Phys. 2002, 92, 55395542. (102) Agarwal, G.; Sowards, L. A.; Naik, R. R.; Stone, M. O. J. Am. Chem. Soc. 2003, 125, 580-583. (103) Lee, K.-B.; Lim, J.-H.; Mirkin, C. A. J. Am. Chem. Soc. 2003, 125, 5588-5589. (104) Lim, J.-H.; Ginger, D. S.; Lee, K.-B.; Heo, J.; Nam, J.-M.; Mirkin, C. A. Angew. Chem., Int. Ed. 2003, 42, 2309-2312. (105) Fu, L.; Liu, X.; Zhang, Y.; Dravid, V. P.; Mirkin, C. A. Nano Lett. 2003, 3, 757-760. (106) Demers, L. M.; Ginger, D. S.; Park, S. J.; Li, Z.; Chung, S. W.; Mirkin, C. A. Science 2002, 296, 1836-1838. (107) Jung, H.; Kulkarni, R.; Collier, C. P. J. Am. Chem. Soc. 2003, 125, 12096-12097. (108) Su, M.; Dravid, V. P. Appl. Phys. Lett. 2002, 80, 4434-4436.

Analytical Chemistry, Vol. 76, No. 12, June 15, 2004

3441

(109) Zhang, M.; Bullen, D.; Chung, S.-W.; Hong, S.; Ryu, K. S.; Fan, Z.; Mirkin, C. A.; Liu, C. Nanotechnology 2002, 13, 212-217. (110) Wang, X.; Ryu, K. S.; Bullen, D. A.; Zou, J.; Zhang, H.; Mirkin, C. A.; Liu, C. Langmuir 2003, 19, 8951-8955. (111) Vettiger, P.; Cross, G.; Despont, M.; Drechsler, U.; Durig, U.; Gotsmann, B.; Haberle, W.; Lantz, M.; Rothuizen, H.; Stutz, R.; Binnig, G. IEEE Trans. Nanotechnol. 2002, 39-55. (112) Lantz, M.; Gotsmann, B.; Durig, U.; Vettiger, P.; Nakayama, Y.; Shimizu, T.; Tokumoto, H. Appl. Phys. Lett. 2003, 83, 12661268. (113) King, W. P.; Kenny, T. W.; Goodson, K. E.; Cross, G. L. W.; Despont, M.; Durig, U. T.; Rothuizen, H.; Binnig, G.; Vettiger, P. J. Microelectromech. Syst. 2002, 11, 765-774. (114) Drechsler, U.; Burer, N.; Despont, M.; Durig, U.; B, G.; Robin, F.; Vettiger, P. J. Microelectron. Eng. 2003, 67, 397-404. (115) Lee, C. S.; Nam, H.-J.; Kim, Y.-S.; Jin, W.-H.; Cho, S.-M.; Bu, J.-u. Appl. Phys. Lett. 2003, 83, 4839-4841. (116) Bo, X.-Z.; Rokhinson, L. P.; Yin, H.; Tsui, D. C.; Sturm, J. C. Appl. Phys. Lett. 2002, 81, 3263-3265. (117) Legrand, B.; Deresmes, D.; Stievenard, D. J. Vac. Sci. Technol., B 2002, 20, 862-870. (118) Luscher, S.; Fuhrer, A.; Held, R.; Heinzel, T.; Ensslin, K.; Bichler, M.; Wegscheider, W. Microelectron. J. 2002, 33, 319-321. (119) Lyuksyutov, S. F.; Vaia, R. A.; Paramonov, P. B.; Juhl, S.; Waterhouse, L.; Ralich, R. M.; Sigalov, G.; Sancaktar, E. Nat. Mater. 2003, 2, 468-472. (120) Cavallini, M.; Biscarini, F.; Leon, S.; Zerbetto, F.; Bottari, G.; Leigh, D. A. Science 2003, 299, 531. (121) Crook, R.; Graham, A. C.; Smith, C. G.; Farrer, I.; Beere, H. E.; Ritchie, D. A. Nature 2003, 424, 751-754. (122) Chen, E.-Q.; Jing, A. J.; Weng, X.; Huang, P.; Lee, S.-W.; Cheng, S. Z. D.; Hsiao, B. S.; Yeh, F. Polymer 2003, 44, 6051-6058. (123) Jiang, Y.; Jin, X.-G.; Han, C. C.; Li, L.; Wang, Y.; Chan, C.-M. Langmuir 2003, 19, 8010-8018. (124) Mogi, R.; Inaba, M.; Iriyama, Y.; Abe, T.; Ogumi, Z. J. Electrochem. Soc. 2002, 149, A385-A390. (125) Yang, X.; Perry, S. S. Surf. Sci. 2002, 506, L261-L267. (126) Namazu, T.; Isono, Y.; Tanaka, T. J. Microelectromech. Syst. 2002, 11, 125-135. (127) Yan, L.; Iwasaki, H. Jpn. J. Appl. Phys., Part 1 2002, 41, 75567559. (128) Lo, Y.-S.; Simons, J.; Beebe, T. P., Jr. J. Phys. Chem. B 2002, 106, 9847-9852. (129) Trawick, M. L.; Angelescu, D. E.; Chaikin, P. M.; Valenti, M. J.; Register, R. A. Rev. Sci. Instrum. 2003, 74, 1390-1392. (130) Lin, W.-M.; Koehler, R.; Suchaneck, G.; Gerlach, G. Jpn. J. Appl. Phys., Part 1 2002, 41, 7239-7241. (131) Nakabeppu, O.; Suzuki, T. J. Therm. Anal. Calorim. 2002, 69, 727-737. (132) Florescu, D. I.; Mourokh, L. G.; Pollak, F. H.; Look, D. C.; Cantwell, G.; Li, X. J. Appl. Phys. 2002, 91, 890-892. (133) Gu, Y. Q.; Ruan, X. L.; Han, L.; Zhu, D. Z.; Sun, X. Y. Int. J. Thermophys. 2002, 23, 1115-1124. (134) Shi, L.; Majumdar, A. J. Heat Transfer 2002, 124, 329-337. (135) Thurber, K. R.; Harrell, L. E.; Smith, D. D. J. Appl. Phys. 2003, 93, 4297-4299. (136) Lefevre, S.; Volz, S.; Saulnier, J.-B.; Fuentes, C.; Trannoy, N. Rev. Sci. Instrum. 2003, 74, 2418-2423. (137) Jena, B. P., Hoerber, J. K. H., Eds. Atomic Force Microscopy in Cell Biology; Methods in cell biology 68; Academic Press: San Diego, 2002. (138) Fotiadis, D.; Scheuring, S.; Muller, S. A.; Engel, A.; Muller, D. J. Micron 2002, 33, 385-397. (139) Hoerber, J. K. H.; Miles, M. J. Science 2003, 302, 1002-1005. (140) Zlatanova, J.; Leuba, S. H. J. Mol. Biol. 2003, 331, 1-19. (141) Hansma, H.; Kasuya, K.; Oroudjev, E. Curr. Opin. Struct. Biol., in press. (142) Mueller, D. J.; Janovjak, H.; Lehto, T.; Kuerschner, L.; Anderson, K. Prog. Biophys. Mol. Biol. 2002, 79, 1-43. (143) Werten, P. J. L.; Remigy, H. W.; de Groot, B. L.; Fotiadis, D.; Philippsen, A.; Stahlberg, H.; Grubmuller, H.; Engel, A. FEBS Lett. 2002, 529, 65-72. (144) Fotiadis, D.; Liang, Y.; Filipek, S.; Saperstein, D. A.; Engel, A.; Palczewski, K. Nature 2003, 421, 127-128. (145) DePace, A. H.; Weissman, J. S. Nat. Struct. Biol. 2002, 9, 389396. (146) Kundu, B.; Maiti, N. R.; Jones, E. M.; Surewicz, K. A.; Vanik, D. L.; Surewicz, W. K. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 12069-12074. (147) Scheibel, T.; Parthasarathy, R.; Sawicki, G.; Lin, X.-M.; Jaeger, H.; Lindquist Susan, L. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 4527-4532. (148) Yan, H.; LaBean, T. H.; Feng, L.; Reif, J. H. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 8103-8108. (149) Yan, H.; Park, S. H.; Finkelstein, G.; Reif, J. H.; LaBean, T. H. Science 2003, 301, 1882-1884. (150) Yan, H.; Zhang, X.; Shen, Z.; Seeman, N. C. Nature 2002, 415, 62-65. (151) Keren, K.; Krueger, M.; Gilad, R.; Ben-Yoseph, G.; Sivan, U.; Braun, E. Science 2002, 297, 72-75. (152) Keren, K.; Berman, R. S.; Buchstab, E.; Sivan, U.; Braun, E. Science 2003, 302, 1380-1382. 3442

Analytical Chemistry, Vol. 76, No. 12, June 15, 2004

(153) Gomez-Navarro, C.; Moreno-Herrero, F.; de Pablo, P. J.; Colchero, J.; Gomez-Herrero, J.; Baro, A. M. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 8484-8487. (154) Sampaolese, B.; Bergia, A.; Scipioni, A.; Zuccheri, G.; Savino, M.; Samori, B.; De Santis, P. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 13566-13570. (155) Sitko, J. C.; Mateescu, E. M.; Hansma, H. G. Biophys J 2003, 84, 419-431. (156) Kato, M.; McAllister, C. J.; Hokabe, S.; Shimizu, N.; Lyubchenko, Y. L. Eur. J. Biochem. 2002, 269, 3632-3636. (157) Potaman, V. N.; Bissler, J. J.; Hashem, V. I.; Oussatcheva, E. A.; Lu, L.; Shlyakhtenko, L. S.; Lyubchenko, Y. L.; Matsuura, T.; Ashizawa, T.; Leffak, M.; Benham, C. J.; Sinden, R. R. J. Mol. Biol. 2003, 326, 1095-1111. (158) Liu, W.; Montana, V.; Chapman Edwin, R.; Mohideen, U.; Parpura, V. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 1362113625. (159) Pinnaduwage, L. A.; Gehl, A.; Hedden, D. L.; Muralidharan, G.; Thundat, T.; Lareau, R. T.; Sulchek, T.; Manning, L.; Rogers, B.; Jones, M.; Adams, J. D. Nature 2003, 425, 474. (160) Arntz, Y.; Seelig, J. D.; Lang, H. P.; Zhang, J.; Hunziker, P.; Ramseyer, J. P.; Meyer, E.; Hegner, M.; Gerber, C. Nanotechnology 2003, 14, 86-90. (161) Dutta, P.; Tipple, C. A.; Lavrik, N. V.; Datskos, P. G.; Hofstetter, H.; Hofstetter, O.; Sepaniak, M. J. Anal. Chem. 2003, 75, 23422348. (162) Lang, H. P.; Hegner, M.; Meyer, E.; Gerber, C. Nanotechnology 2002, 13, R29-R36. (163) Cherian, S.; Thundat, T. Appl. Phys. Lett. 2002, 80, 2219-2221. (164) Dufour, I.; Fadel, L. Sens. Actuators, B 2003, B91, 353-361. (165) Vidic, A.; Then, D.; Ziegler, C. Ultramicroscopy 2003, 97, 407416. (166) Naik, T.; Longmire, E. K.; Mantell, S. C. Sens. Actuators, A 2003, A102, 240-254. (167) Headrick, J. J.; Sepaniak, M. J.; Lavrik, N. V.; Datskos, P. G. Ultramicroscopy 2003, 97, 417-424. (168) Zhang, Y.; Ji, H.-F.; Brown, G. M.; Thundat, T. Anal. Chem. 2003, 75, 4773-4777. (169) Zhou, J.; Li, P.; Zhang, S.; Long, Y.; Zhou, F.; Huang, Y.; Yang, P.; Bao, M. Sens. Actuators, B 2003, B94, 337-342. (170) Domanski, K.; Grabiec, P.; Marczewski, J.; Gotszalk, T.; Ivanov, T.; Abedinov, N.; Rangelow, I. W. J. Vac. Sci. Technol., B 2003, 21, 48-52. (171) Marie, R.; Thaysen, J.; Christensen, C. B. V.; Boisen, A. J. Microelectron. Eng. 2003, 67-68, 893-898. (172) Subramanian, A.; Oden, P. I.; Kennel, S. J.; Jacobson, K. B.; Warmack, R. J.; Thundat, T.; Doktycz, M. J. Appl. Phys. Lett. 2002, 81, 385-387. (173) Lange, D.; Hagleitner, C.; Hierlemann, A.; Brand, O.; Baltes, H. Anal. Chem. 2002, 74, 3084-3095. (174) Davis, Z. J.; Abadal, G.; Helbo, B.; Hansen, O.; Campabadal, F.; Perez-Murano, F.; Esteve, J.; Figueras, E.; Verd, J.; Barniol, N.; Boisen, A. Sens. Actuators, A 2003, A105, 311-319. (175) Calleja, M.; Rasmussen, P. A.; Johansson, A.; Boisen, A. Proc. SPIE Int. Soc. Opt. Eng. 2003, 5116, 314-321. (176) Adams, J. D.; Parrott, G.; Bauer, C.; Sant, T.; Manning, L.; Jones, M.; Rogers, B.; McCorkle, D.; Ferrell, T. L. Appl. Phys. Lett. 2003, 83, 3428-3430. (177) Humphris, A. D. L.; Antognozzi, M.; McMaster, T. J.; Miles, M. J. Langmuir 2002, 18, 1729-1733. (178) Lim, R.; Li, S. F. Y.; O’Shea, S. J. Langmuir 2002, 18, 61166124. (179) Kappl, M.; Butt, H.-J. Part. Part. Syst. Charact. 2002, 19, 129143. (180) Zhang, W.; Zhang, X. Prog. Polym. Sci. 2003, 28, 1271-1295. (181) Akhremitchev, B. B.; Bemis, J. E.; Al-Maawali, S.; Sun, Y.; Stebounova, L.; Walker, G. C. Biofouling 2003, 19, 99-104. (182) Chen, F.; Mohideen, U.; Klimchitskaya, G. L.; Mostepanenko, V. M. Phys. Rev. Lett. 2002, 88, 101801. (183) Kudera, M.; Eschbaumer, C.; Gaub, H. E.; Schubert, U. S. Adv. Funct. Mater. 2003, 13, 615-620. (184) Marszalek, P. E.; Li, H.; Oberhauser, A. F.; Fernandez, J. M. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 4278-4283. (185) Zapotoczny, S.; Auletta, T.; de Jong, M. R.; Schoenherr, H.; Huskens, J.; van Veggel, F. C. J. M.; Reinhoudt, D. N.; Vancso, G. J. Langmuir 2002, 18, 6988-6994. (186) Tivanski, A. V.; Bemis, J. E.; Akhremitchev, B. B.; Liu, H.; Walker, G. C. Langmuir 2003, 19, 1929-1934. (187) Dicke, C.; Haehner, G. J. Am. Chem. Soc. 2002, 124, 1261912625. (188) Zepeda, S.; Yeh, Y.; Noy, A. Langmuir 2003, 19, 1457-1461. (189) Mabry, J. C.; Yau, T.; Yap, H.-W.; Green, J.-B. D. Ultramicroscopy 2002, 91, 73-82. (190) Green, J.-B. D. Anal. Chim. Acta 2003, 496, 267-277. (191) Connell, S. D. A.; Allen, S.; Roberts, C. J.; Davies, J.; Davies, M. C.; Tendler, S. J. B.; Williams, P. M. Langmuir 2002, 18, 17191728. (192) Eaton, P.; Smith, J. R.; Graham, P.; Smart, J. D.; Nevell, T. G.; Tsibouklis, J. Langmuir 2002, 18, 3387-3389. (193) Sato, F.; Okui, H.; Akiba, U.; Suga, K.; Fujihira, M. Ultramicroscopy 2003, 97, 303-314. (194) Bottomley, L. A.; Poggi, M. A.; Lillehei, P. T. Proc. Electrochem. Soc. 2003, 2003-15, 297-304.

(195) Akabori, K.-i.; Tanaka, K.; Kajiyama, T.; Takahara, A. Macromolecules 2003, 36, 4937-4943. (196) Bliznyuk, V. N.; Assender, H. E.; Briggs, G. A. D. Macromolecules 2002, 35, 6613-6622. (197) Harmon, M. E.; Kuckling, D.; Frank, C. W. Langmuir 2003, 19, 10660-10665. (198) Hodges, C. S. Adv. Colloid Interface Sci. 2002, 99, 13-75. (199) Bunker, B. C.; Kim, B. I.; Houston, J. E.; Rosario, R.; Garcia, A. A.; Hayes, M.; Gust, D.; Picraux, S. T. Nano Lett. 2003, 3, 17231727. (200) Houston, J. E.; Kim Hyun, I. Acc. Chem. Res. 2002, 35, 547553. (201) Hugel, T.; Holland Nolan, B.; Cattani, A.; Moroder, L.; Seitz, M.; Gaub Hermann, E. Science 2002, 296, 1103-1106. (202) Rabe, U.; Amelio, S.; Kopycinska, M.; Hirsekorn, S.; Kempf, M.; Goken, M.; Arnold, W. Surf. Interface Anal. 2002, 33, 65-70. (203) Benmouna, F.; Dimitrova, T. D.; Johannsmann, D. Langmuir 2003, 19, 10247-10253. (204) Dimitrova, T. D.; Johannsmann, D.; Willenbacher, N.; Pfau, A. Langmuir 2003, 19, 5748-5755. (205) Linke, W. A., Granzier, H., Kellermayer, M. S. Z., Eds. Mechanics of Elastic Biomolecules; Kluwer Academic Publishers: Boston, 2003. (206) Williams, M. C.; Rouzina, I. Curr. Opin. Struct. Biol. 2002, 12, 330-336. (207) Dufrene, Y. F. Curr. Opin. Microbiol. 2003, 6, 317-323. (208) Zhuang, X.; Rief, M. Curr. Opin. Struct. Biol. 2003, 13, 88-97. (209) Allison, D. P.; Hinterdorfer, P.; Han, W. Curr. Opin. Biotechnol. 2002, 13, 47-51. (210) Rief, M.; Grubmuller, H. ChemPhysChem 2002, 3, 255-261. (211) Terentjev, E. M. Nat. Mater. 2002, 1, 149-150. (212) Li, H.; Linke, W. A.; Oberhauser, A. F.; Carrion-Vazquez, M.; Kerkvliet, J. G.; Lu, H.; Marszalek, P. E.; Fernandez, J. M. Nature 2002, 418, 998-1002. (213) Scott, K. A.; Steward, A.; Fowler, S. B.; Clarke, J. J. Mol. Biol. 2002, 315, 819-829. (214) Oberhauser, A. F.; Badilla-Fernandez, C.; Carrion-Vazquez, M.; Fernandez, J. M. J. Mol. Biol. 2002, 319, 433-447. (215) Carrion-Vazquez, M.; Li, H.; Lu, H.; Marszalek Piotr, E.; Oberhauser Andres, F.; Fernandez Julio, M. Nat. Struct. Biol. 2003, 10, 738-743. (216) Fernandez, J. M.; Li, H. B. Science 2004, 303, 1674-1678. (217) Muller, D., J.; Kessler, M.; Oesterhelt, F.; Moller, C.; Oesterhelt, D.; Gaub, H. Biophys. J. 2002, 83, 3578-3588. (218) Janovjak, H.; Kessler, M.; Oesterhelt, D.; Gaub, H.; Mueller, D. J. EMBO J. 2003, 22, 5220-5229. (219) Nevo, R.; Stroh, C.; Kienberger, F.; Kaftan, D.; Brumfeld, V.; Elbaum, M.; Reich, Z.; Hinterdorfer, P. Nat. Struct. Biol. 2003, 10, 553-557. (220) Oroudjev, E.; Soares, J.; Arcdiacono, S.; Thompson, J. B.; Fossey, S. A.; Hansma, H. G. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 6460-6465. (221) Oroudjev, E.; Hayashi, C. Y.; Soares, J.; Arcidiacono, S.; Fossey, S. A.; Hansma, H. G. Mater. Res. Soc. Symp. Proc. 2003, 738, 273-279. (222) Becker, N.; Oroudjev, E.; Mutz, S.; Cleveland, J. P.; Hansma, P. K.; Hayashi, C. Y.; Makarov, D. E.; Hansma, H. G. Nat. Mater. 2003, 2, 278-283. (223) Abu-Lail, N. I.; Camesano, T. A. Langmuir 2002, 18, 40714081.

(224) Abu-Lail, N. I.; Camesano, T. A. Biomacromolecules 2003, 4, 1000-1012. (225) Abu-Lail, N. I.; Camesano, T. A. Environ. Sci. Technol. 2003, 37, 2173-2183. (226) Marshall, B. T.; Long, M.; Piper, J. W.; Yago, T.; McEver, R. P.; Zhu, C. Nature 2003, 423, 190-193. (227) Krautbauer, R.; Pope, L. H.; Schrader, T. E.; Allen, S.; Gaub, H. E. FEBS Lett. 2002, 510, 154-158. (228) Albrecht, C.; Blank, K.; Lalic-Myelthaler, M.; Hirler, S.; Mai, T.; Gilbert, I.; Schiffmann, S.; Bayer, T.; Clausen-Schaumann, H.; Gaub, H. E. Science 2003, 301, 367-370. (229) Bonin, M.; Zhu, R.; Klaue, Y.; Oberstrass, J.; Oesterschulze, E.; Nellen, W. Nucleic Acids Res. 2002, 30, 81-86. (230) Ouyang, M.; Huang, J.-L.; Lieber, C. M. Annu. Rev. Phys. Chem. 2002, 53, 201-220. (231) Kim, D.-H.; Koo, J.-Y.; Kim, J.-J. Phys. Rev. B: Condens. Matter 2003, 68, 113401-113406. (232) Park, J.-Y.; Yaish, Y.; Brink, M.; Rosenblatt, S.; McEuen, P. L. Appl. Phys. Lett. 2002, 80, 4446-4448. (233) Woodside, M. T.; McEuen, P. L. Science 2002, 296, 1098-1101. (234) Bozovic, D.; Bockrath, M.; Hafner, J. H.; Lieber, C. M.; Park, H.; Tinkham, M. Phys. Rev. B: Condens. Matter 2003, 67, 33401-33407. (235) Kim, K. S.; Lim, S. C.; Lee, I. B.; An, K. H.; Bae, D. J.; Choi, S.; Yoo, J.-E.; Lee, Y. H. Rev. Sci. Instrum. 2003, 74, 4021-4025. (236) Thelander, C.; Samuelson, L. Nanotechnology 2002, 13, 108113. (237) Minot, E. D.; Yaish, Y.; Sazonova, V.; Park, J.-Y.; Brink, M.; McEuen, P. L. Phys. Rev. Lett. 2003, 90, 156401-156404. (238) Williams, P. A.; Papadakis, S. J.; Patel, A. M.; Falvo, M. R.; Washburn, S.; Superfine, R. Phys. Rev. Lett. 2002, 89, 2555021-255502-4. (239) Chen, X.; Zhang, S.; Dikin, D. A.; Ding, W.; Ruoff, R. S.; Pan, L.; Nakayama, Y. Nano Lett. 2003, 3, 1299-1304. (240) Lee, J.; Kim, H.; Kahng, S. J.; Kim, G.; Son, Y. W.; Ihm, J.; Kato, H.; Wang, Z. W.; Okazaki, T.; Shinohara, H.; Kuk, Y. Nature 2002, 415, 1005-1008. (241) Cho, Y.; Han, S.; Kim, G.; Lee, H.; Ihm, J. Phys. Rev. Lett. 2003, 90, 106401-106402. (242) Hornbaker, D. J.; Kahng, S. J.; Misra, S.; Smith, B. W.; Johnson, A. T.; Mele, E. J.; Luzzi, D. E.; Yazdani, A. Science 2002, 295, 828-831. (243) Kane, C. L.; Mele, E. J.; Johnson, A. T.; Luzzi, D. E.; Smith, B. W.; Hornbaker, D. J.; Yazdani, A. Phys. Rev. B: Condens. Matter 2002, 66, 235423-1/235421-15. (244) Li, J.; Stevens, R.; Delzeit, L.; Ng, H. T.; Cassell, A.; Han, J.; Meyyappan, M. Appl. Phys. Lett. 2002, 81, 910-912. (245) Lillehei, P. T.; Park, C.; Rouse, J. H.; Siochi, E. J. Nano Lett. 2002, 2, 827-829. (246) Cooper, C. A.; Cohen, S. R.; Barber, A. H.; Wagner, H. D. Appl. Phys. Lett. 2002, 81, 3873-3875. (247) Barber, A. H.; Cohen, S. R.; Wagner, H. D. Appl. Phys. Lett. 2003, 82, 4140-4142. (248) Ding, W.; Eitan, A.; Fisher, F. T.; Chen, X.; Dikin, D. A.; Andrews, R.; Brinson, L. C.; Schadler, L. S.; Ruoff, R. S. Nano Lett. 2003, 3, 1593-1597.

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