Letter pubs.acs.org/NanoLett
Atomic Force Microscopy with Nanoscale Cantilevers Resolves Different Structural Conformations of the DNA Double Helix Carl Leung,*,† Aizhan Bestembayeva,†,‡ Richard Thorogate,† Jake Stinson,†,‡ Alice Pyne,†,§ Christian Marcovich,†,∥ Jinling Yang,⊥ Ute Drechsler,# Michel Despont,# Tilo Jankowski,∇ Martin Tschöpe,∇ and Bart W. Hoogenboom*,†,‡ †
London Centre for Nanotechnology, University College London, 17−19 Gordon Street, London WC1H 0AH, United Kingdom Department of Physics and Astronomy, University College London, Gower Street, London WC1E 6BT, United Kingdom § National Physical Laboratory, Hampton Road, Teddington TW11 0LW, United Kingdom ∥ ́ Ecole Polytechnique, 91128 Palaiseau Cedex, France ⊥ Institute of Semiconductors, Chinese Academy of Sciences, Beijing 100083, P. R. China # IBM Research Division, Zurich Research Laboratory, Säumerstrasse 4, 8803 Rüschlikon, Switzerland ∇ JPK Instruments AG, Bouchéstrasse 12, 12435 Berlin, Germany ‡
S Supporting Information *
ABSTRACT: Structural variability and flexibility are crucial factors for biomolecular function. Here we have reduced the invasiness and enhanced the spatial resolution of atomic force microscopy (AFM) to visualize, for the first time, different structural conformations of the two polynucleotide strands in the DNA double helix, for single molecules under nearphysiological conditions. This is achieved by identifying and tracking the anomalous resonance behavior of nanoscale AFM cantilevers in the immediate vicinity of the sample.
KEYWORDS: Atomic force microscopy, molecular resolution, DNA, nanoscale cantilevers, frequency and phase modulation
A
molecules in solution. But even then, most biomolecules and complexes appear as mobile three-dimensional (3D) structures with large (nanometer-scale) protrusions that can be easily dislodged by the scanning tip. This problem underpinned the development of amplitude-modulation AFM (“tapping mode”)9where the cantilever probe is vertically oscillated above the sample to minimize lateral drag forcesand the development of other techniques that aim to improve the sensitivity of the probe in close proximity to the sample.5,10−14 In this respect, the miniaturization of AFM cantilevers8,15−17 (Figure 1) has yielded unambiguous advantages, as it reduces the noise in detecting the tip−sample forces and facilitates faster scanning due to the increased resonance frequencies. Another promising development has been frequency-modulation AFM, which tracks the tip−sample interaction via minute changes in the cantilever resonance frequency (instead of its amplitude, such as tapping mode). In combination with highly
tomic force microscopy (AFM) is a unique tool for the nanometer-scale visualization of single biomolecules in their native environment, that is, in aqueous solution. To achieve its remarkable, up to atomic resolution,1−5 AFM scans the sample surface with a sharp tip that is mounted on a flexible cantilever. By monitoring the cantilever deflection and controlling the compressive force exerted by the scanning tip, a topographical image of the sample surface can be obtained in aqueous solution. Atomic resolution on flat surfaces has thus been obtained,1−5 as well as subnanometer resolution on membrane proteins in densely packed, two-dimensional (2D) arrays.2,6,7 AFM is therefore a potentially ideal platform for realtime and subnanometer-resolution imaging of the various and complex structures adopted by single and fully functional biological molecules,8 thus providing a direct view on their intricate function. However, this potential is far from being fully exploited, since only few biological molecules arrange to form flat surfaces that are appropriate for high-resolution AFM. Such nanoscale experiments require a minimum fixation (typically by physisorption) of the sample to ausually flatsubstrate, thus reducing the blurring effect of thermal motion of the © 2012 American Chemical Society
Received: May 17, 2012 Revised: June 21, 2012 Published: June 26, 2012 3846
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sensitive optical deflection detectors, it now routinely succeeds in imaging solid−liquid interfaces at atomic resolution1−3 and has been applied to a range of biological samples.3,18,19 It is conceptually simple and has the additional advantage of a straightforward conversion of measured frequency shifts to tip− sample forces.20 Nevertheless, in spite of these developments, which have by now been implemented as standard options on several commercial systems, there has been no substantial improvement in spatial resolution on benchmark biomolecular samples since the mid-1990s, for example, on the membrane protein bacteriorhodopsin6 and on DNA.9,21 In particular, it remains a grand challenge to resolve biologically relevant variations in molecular substructure and subnanometer-scale conformational changes on fragile and corrugated samples with nonhomogeneous topography. Since the spatial resolution on such samples is largely determined by the minimum load force at which sufficient contrast can be obtained, we here set out to minimize the invasiness of the AFM probe for measurements in aqueous solution. To achieve this aim, we have reduced the thickness of the cantilevers to the nanometer scale and their lateral dimensions to the order of 1 μm (Figure 1 and Supporting Information (SI), Table S1)16 to reduce the noise when detecting the tip− sample forces and characterized and tracked their resonance behavior near the sample surface. As will be shown below, the
Figure 1. Miniaturized cantilever driven by photothermal actuation. (a) Scanning electron microscopy image of a 11 μm long and 250 nm tick cantilever16 (inset), compared to a conventional NCH cantilever (Nanosensors, Neuchatel, Switzerland) on the same scale. (b) Miniaturized cantilever with a superposed sketch (not to scale) representing part of the optics23 that is used for both the interferometric detection and the photothermal actuation11,24 of the cantilever in this work. (c) Resonance curve of a small cantilever obtained by standard piezo-acoustic actuation (blue, dotted) and by photothermal (red) actuation. With photothermal actuation, the resonance curve can be directly overlaid with the thermal fluctuations (green) of the cantilever.
Figure 2. AFM by tracking the resonance frequency of miniaturized (MHz-frequency) cantilevers. (a) Thermal noise of a small cantilever (spring constant k = 13 N/m, resonance frequency f 0 = 1.52 MHz, tip height 1.4 μm) compared with a conventional cantilever (k = 16 N/m, f 0 = 0.14 MHz, tip height 16 μm) as displayed in Figure 1, at 4.4 μm and a 130 nm tip−sample distance. Continuous lines indicate harmonic-oscillator fits, and arrows indicate the resonance frequencies. (b) Resonance frequency of the miniaturized cantilever as a function of tip−sample distance, as predicted by hydrodynamic theory (Green−Sader),27 as measured by tracking the phase of the photothermally actuated cantilever (oscillation amplitude a = 0.75 nm) and as determined from its thermal noise at various tip−sample distances. (c) Resonance frequency on closest tip−sample approach and the corresponding peak force between the tip and the sample (see SI). Inset: Atomic resolution on mica (color scale 0.12 nm). (d) Trimers of bacteriorhodopsin molecules in a 2D lattice, including a ∼10 nm scale defect (color scale 0.5 nm).
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in the MHz frequency range around the cantilever resonance, whichdue to their macroscopic originwere prone to drift. On approaching the miniaturized cantilever probe to the sample, its resonance frequency does not remain constant, butbecause of hydrodynamic interactions between the cantilever beam and the sample surface27it decreases rapidly over a range of several micrometers before the tip and sample make physical contact. Over this range, the (normalized) frequency shift (Figure 2a) for our miniaturized cantilevers is an order of magnitude larger than for conventionally sized cantilevers (widths ≳30 μm, SI, Figure S1). Upon contact, there is a much smaller upturn of the resonance frequency due to the short-range (≲1 nm) repulsive tip−sample interactions on which the AFM image contrast relies (Figure 2b,c). For the miniaturized cantilevers, the mesoscale hydrodynamics and nanometer-scale tip−sample interactions thus have competing effects on the resonance frequency over the typical ≲1 μm vertical scan range of a high-resolution AFM experiment. Similarly, the cantilever hydrodynamics near the surface leads to increasingly rapid variations of the quality factor of the cantilever (SI, Figure S1).26 Since, over distances of ≲1 μm, the variation in these hydrodynamic interactions scales inversely to the cantilever width, these effects will be even more dramatic for cantilevers of which the lateral dimensions are reduced to the ∼100 nm scale, which is an ongoing development to further increase their resonance frequencies and enhance the signal-tonoise ratio.28 These effects can partly be mitigated by increasing the ratio between tip height and cantilever width.26,27 In practice, however, this ratio is limited to well below unity, for reasons of tip stability and complications in the manufacturing process. When using miniaturized cantilevers in an amplitudemodulation (“tapping-mode”) AFM experiment, the competition between cantilever−surface and tip−surface interactions translates into increased ambiguity on the appropriate imaging parameters. In an experiment where the tip−sample distance control is based on frequency tracking, this competition can have disastrous consequences for the tip (and possibly for the sample), for the following reason. To approach the tip to the sample, the target value (“set-point”) for the frequency shift should be set slightly higher than its value at several hundred nanometers (or more) above the surface. Because of the drop in resonance frequency closer to the surface, such a set-point implies a huge frequency shift with respect to the cantilever resonance in the last few nanometers before contact. This results in a destructively large tip−sample force. We circumvent this problem by continuously adjusting a reference frequency during the tip−sample approach, such that the frequency shift in contact is measured with respect to the resonance frequency only a few nanometers above the surface. Thus tracking the tip−sample interaction via the resonance frequency of miniaturized cantilevers (see also the SI), we obtain images on flat surfaces such as mica and 2D crystals of bacteriorhodopsin (Figure 2c,d) that are in good agreement with the highest-resolution data available in the literature.1,2,4,5,7 To demonstrate the advantage of our method for imaging biomolecules of larger 3D structural complexity, we have adsorbed DNA plasmids on mica and resolved their molecular substructure. DNA can be probed in real time by AFM for its structural variability arising from thermal fluctuations, its interactions with individual proteins and ligands, and due to twisting, bending, and stretching.9,21,29,30 However, submolecular details of DNA have yet to be reproducibly resolved in
Figure 3. Minimally invasive imaging of plasmid DNA. (a) Amplitudemodulation (“Ampl.”, set-point ≳95% of free amplitude) and frequency-tracking (“Freq.”, phase set-point 2°) AFM images of the same molecule with the same miniaturized cantilever, displayed on the same color scale (2.8 nm). The amplitude of the cantilever oscillation was 1.0 ± 0.1 nm while imaging, and other parameters for both images were adjusted for optimum contrast and stability. The measured heights are compared via average cross sections of the DNA along the roughly straight segment marked in the AFM images. The molecule appears significantly wider and higher in the frequency-tracking data (orange) than in amplitude-modulation ("tapping mode") data (blue), with the height approaching the expected 2.0 nm diameter of B-DNA. (b) With our method, the measurement is sufficiently accurate to extract realistic heights for single-stranded (ss) DNA and doublestranded B-DNA. In addition, there are higher segments that may be identified as intramolecular triplex DNA (H-DNA, see cartoon inset), all in the same (75 × 75 nm2) image. The cross sections were taken along the lines marked by arrows in the AFM image (color scale: 3.4 nm). See the SI, Figure S2, for a larger-scale image.
resonance behavior of such miniaturized cantilevers is dramatically different from that of conventionally sized cantilevers, butwhen accurately trackedfacilitates a minimally invasive method of probing molecules with complex 3D structures. We demonstrate this potential using plasmid DNA as a benchmark sample, adsorbed in a 2D projection of its native 3D configuration,22 and show that we can reproducibly resolve the two oligonucleotide strands of the double helix in different structural conformations. Technically speaking, we use constant-amplitude phasemodulation AFM (see SI).25 This is equivalent to frequencymodulation AFM1,2,26 as long as the frequency shifts are small compared to the width of the resonance, since in this range the phase is a linear function of the frequency. Cantilever characterization and AFM were performed on a home-built microscope with a Fabry−Perot interferometer to detect the cantilever deflection.23 For all measurements in this study, the sample, the cantilever, and the front lens surface of the interferometer were completely immersed in buffer solution. For characterization of their resonance behavior near the surface, the small cantilevers were actuated photothermally,11,24 avoiding the many spurious mechanical resonances (Figure 1c) 3848
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Figure 4. High-resolution AFM on plasmid DNA. (a) 3486 bp plasmid DNA, which on magnification shows a right-handed double helix, superposed to substantial height differences on and along the DNA. (b) Profiles along right-handed (RH) and left-handed (LH) DNA, acquired along the lines marked by the red and blue arrows in the insets of A and D. (c) Elongated configuration of the 3486 bp plasmid DNA displayed at the same scale as part a. (d) On subsequent magnifications of the dashed rectangle in c, an elongated left-handed double helix is resolved. Color scale: 1.5 nm (a); 1.1 nm (a, inset); 1.5 nm (c); 1.1 nm (d); 0.7 nm (d, inset). Green arrows indicate the two strands of the double helix, separated by the minor groove (depth ≲0.1 nm). The major groove (depth ∼0.2 nm) separates the subsequent turns of the double helix.
physiological conditions, with the DNA in its natively coiled 3D structure (or a 2D projection of it)22 and at controlled ionic concentrations. Most images in the literature feature DNA as a long polymer without any noticeable substructure.9,21 Only in exceptional cases could the major groove of the double helix be resolved in liquid, by imaging in isopropanol,9 by creating a surface of artificially dense DNA aggregates,3,31 or by artificially stretching the DNA on a lipid monolayer in a drying process.32 In other words, the helical character of DNA has thus far been barely recognized by AFM and its double-strandedness (as would be apparent by the presence of subsequent major and minor grooves) completely ignored. Upon imaging DNA plasmids with the same miniaturized cantilever probe, we find an immediate and tangible advantage of tracking the resonance frequency instead of amplitude. On imaging DNA in amplitude-modulation (“tapping mode”), the average diameter of the molecule measures only (and typically) ∼1.1 nm, indicating significant compression of the molecule. However, when tracking the tip−sample distance via the resonance frequency, the measured plasmid height of ∼1.8 nm agrees to within the experimental error (±10%) with the 2.0 nm diameter that is known from the crystal structure of B-DNA (Figure 3a). Based on measured heights, we also unambiguously discern single-stranded (ss) and double-stranded (BDNA), as well as a segment with about three times the height of ss-DNA, which may therefore be attributed to triplex HDNA33 (Figure 3b). On a smaller scale, superposed to the ∼2.0 nm protrusion of the DNA above the substrate and to significant height differences at length scales ≳10 nm along the molecule, we discern a tilted double-stranded structure with a periodicity of 3.5 ± 0.4 nm (Figure 4a, inset). Its subnanometer corrugation becomes strikingly clear in line profiles along the DNA (Figure 4b, top) and can immediately be identified as the subsequent major and minor grooves of Watson and Crick’s
double helix (B-DNA, right-handed, helical periodicity 3.6 nm30), demonstrating the superiority of our method by thus resolving the two separate oligonucleotide strands of the DNA. This contrast is obtained in spite of the overall tip convolution that results in the DNA appearing significantly wider than its true diameter (Figure 3a). This suggests that structural or chemical asperities on the tip are responsible for the fine contrast, akin to the mechanism proposed for high-resolution imaging in contact-mode images.7 Intriguingly, a small fraction of the DNA molecules have contour lengths between 1.8−2.2 μm, which is 50−85% longer than the ∼1.2 μm expected and found (Figure 4a) for the BDNA conformation of the 3486 base-pair (bp) plasmid. This suggests that the plasmid is either elongated or dimerized (see SI, Figures S3−S5, for further characterization of the DNA samples by agarose gel electrophoresis and AFM). Similarly elongated (by ∼70%) DNA has been observed in optical tweezers experiments,29,30 where a central question is whether any helical structure persists on overstretching DNA molecules. Here we demonstrate the existence of an elongated helical conformation. The elongated DNA in our experiments (Figure 4c), possibly stabilized by interactions with the Ni2+ ions on the mica substrate, displays a left-handed double helix with a periodicity of 8.0 ± 0.5 nm and distinct major and minor grooves (Figure 4b and d). The latter two observations rule out Z-DNA, which is the best known left-handed DNA conformation (SI, Table S2).34 The periodicity along the molecule is the same for different DNA orientations with respect to the AFM tip, which excludes tip-artifacts as the origin of the observed contrast (Figure 4d, SI, Figure S6). On adsorbed biological molecules that are typical in terms of their corrugation and mobility, we have thus achieved unprecedented spatial resolution by tracking the strongly nonmonotonic resonance behavior of miniaturized AFM3849
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cantilevers near the sample surface. Importantly, the results are sufficiently reproducible to enable identification of the individual oligonucleotide strands in different structural conformations of the DNA double helix. With the advent of several commercial frequency-tracking atomic force microscopes and the increasing availability of miniaturized cantilevers, this method will facilitate the study of a range of fundamental biological processes in the genome, such as changes in its molecular substructure as a function of protein binding and of macromolecular extension and coiling,9,21,29,30 or naturally occurring DNA lesion.35 Moreover, because of the high (MHz) resonance frequencies of these cantilevers, there is a broad scope for time-resolved studies of biomolecular kinetics at subnanometer resolution on soft and uneven samples.
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ASSOCIATED CONTENT
S Supporting Information *
Further experimental details on the atomic force microscope and cantilevers used in this work; procedure to convert changes in phase to frequency shifts and forces; sample preparation, imaging conditions, and image analysis; further characterization of the hydrodynamic interactions between cantilever and sample surface; additional height comparisons for different modes of operation and for left- and right-handed conformations of the double helix; further characterization of the DNA samples by agarose gel electrophoresis and AFM; comparison of the experimental observations to different known structural conformations of the DNA double helix. This material is available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected];
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS We thank P. L. T. M. Frederix, A. Engel, and H. J. Hug for support; Nanoworld AG for free samples of their USC-2 MHz cantilevers; S. Medalion, Y. Rabin, R. Cortini, D. J. Lee, and A. A. Kornyshev for discussions; and S. Howorka, M. Watkins and A. L. Shluger for discussions and proofreading the manuscript. This work was funded by the UK Biotechnology and Biological Sciences Research Council (BB/G011729/1 to B.W.H.).
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REFERENCES
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