© Copyright 2000 by the American Chemical Society
VOLUME 104, NUMBER 26, JULY 6, 2000
LETTERS Local Solvation Shell Measurement in Water Using a Carbon Nanotube Probe Suzanne P. Jarvis,* Takayuki Uchihashi,† Takao Ishida,‡ and Hiroshi Tokumoto Joint Research Center for Atom Technology, Higashi 1-1-4, Tsukuba, Ibaraki 305-0046, Japan
Yoshikazu Nakayama Department of Physics and Electronics, Osaka Prefecture UniVersity, 1-1 Gakuen-Cho Sakai, Osaka 599-8531, Japan ReceiVed: April 27, 2000
Combining a carbon nanotube probe and a highly sensitive dynamic measurement scheme has enabled us to use atomic force microscopy to measure oscillatory forces in water on approaching a surface that has been laterally characterized on a nanometer scale. This opens up the possibility of investigating water layers under a variety of experimental conditions and as a function of precise lateral position on any surface including biological membranes and macromolecules. Among the many and varied roles of water layers are effects on biomolecular adhesion, colloid dispersion, and tribology, which can now be investigated with nanometer lateral resolution and with a wider range of materials than that previously provided by surface forces appartus.
1. Introduction Oscillatory forces between two approaching surfaces in a solvent have long been the subject of study due to their possible influence on any surface-surface interactions mediated through a liquid or in the presence of a fluid film. Of particular interest is water, due to its omnipresence in all but the most stringently controlled environments and its role as the primary medium for biological interactions.1 The observed breakdown in continuum theories at short range (usually of the order of 4-8 molecular diameters) can readily be explained as being due to confinement when the liquid is squeezed between two macroscopic, molecularly smooth surfaces, forcing it into discrete numbers of layers known as * Author to whom correspondence should be addressed. Email: spjarvis@ jrcat.or.jp. † Present address: Department of Electronics, Himeji Institute of Technology, Himeji 671-2201, Japan. ‡ Present address: Mechanical Engineering Laboratory and PRESTOJST, Tsukuba, Ibaraki 305-8564, Japan.
solvation shells. However, this describes the very unique experimental situation found in the surface forces apparatus (SFA) and further simulations were necessary in order to rationalize the subsequent measurement of solvation shells by atomic force microscopy (AFM) where one of the surfaces is approaching molecular dimensions.2,3 In the latter case the oscillatory force curves can be explained in terms of the AFM tip sampling the density profiles of the solvent in the region under the tip. Theoretical simulations have also addressed the issue of how such structuring of the fluid may influence the resolution and imaging mechanisms of AFM in ambient and liquid immersion.4 The instrument most commonly used for the investigation of solvation shells is the SFA,5 which measures forces between two smooth crossed cylinders. The cylinders usually have radii of the order of 1 cm, and the separation between them is measured directly using white light interferometry. This is the only apparatus thus far to demonstrate the continuous measurement of solvation forces in water as a function of surface-
10.1021/jp001616d CCC: $19.00 © 2000 American Chemical Society Published on Web 06/13/2000
6092 J. Phys. Chem. B, Vol. 104, No. 26, 2000 surface separation.6 The instrument is reliant on having a very low surface roughness over a very large interaction area. Thus, most measurements have been limited to mica, a hydrophilic and chemically unreactive surface with no lateral characterization of the two surfaces possible or relevant. With magnetically activated AFM it has been possible to resolve molecular layers of large molecules such as octamethylcyclotetrasiloxane (OMCTS) and n-dodecanol.7 With this method, magnetic material is deposited directly behind an AFM tip on the backside of the cantilever so that the tip position can be controlled by the addition of a magnetic field. The lever can be vibrated in an oscillating magnetic field in order to make dynamic measurements.8 One expected consequence of the success of this technique was a rapid exploitation of the experimental advantages over SFA such as various surface materials that can be studied and simultaneous lateral characterization. However, at present the literature is restricted to measurements as a function of separation between a silicon tip and a mica or graphite surface. Further, using magnetically activated AFM it has not yet been possible to reproduce the solvation shell measurements in water9 measured by SFA. We suspect that this is, in part, due to the long averaging times necessary in order to obtain a sufficiently sensitive signal-tonoise ratio using a lock-in amplifier and also because of the low aspect ratio tips commonly used. One report of water layer and/or hydrated ion measurements using static AFM is that of Cleveland et al.10 They show that with sufficiently long measurement times and sufficient stability it is possible even with static measurements to pinpoint different energy minima close to ionic crystals in water by using the thermal noise of the cantilever. Unfortunately, due to the long averaging times needed for this technique it is not readily applicable to location sensitive investigations. 2. Experiment One of the most important aspects of our method introduced here is the utilization of carbon nanotubes as the AFM probe.11 The main purpose being to remove the unwanted hydrodynamic damping effect caused by the bulk of the tip12 and relying on the fact that integral equation theory has shown3 that the range of oscillations is relatively unaffected by the size of a spherical probe. We have used a multiwalled carbon nanotube (MWNT) tip attached to a PtIr-coated silicon lever using amorphous carbon deposition in a specially designed field emission scanning electron microscope system (FE-SEM). The carbon nanotubes, which are prepared by a conventional arc discharge method,13 are aligned on a knife-edge using an alternating current electrophoresis technique.14 Attachment of a tube to the tip is done using two specially designed independent translation stages for the knife-edge and the AFM cantilever.15 With the MWNT and AFM tip in contact, amorphous contaminants within the FE-SEM are deposited, using the focused electron beam, to immobilize the MWNT on the tip. Depending on the resulting length, tubes are sometimes cut by applying a pulsed voltage between the knife-edge and the tube. After attaching the carbon nanotube to the tip, we avoided prolonged FE-SEM observation to minimize carbon contamination. From the SEM pictures taken before and after the experiment (Figure 1A) and from AFM measurements of similar tubes (Figure 1B) we gather that the tube used in the following experiments was 265 nm in length and had a tip radius of approximately 7 nm. The end is assumed to be closed as no pulsing method was implemented to shorten the nanotube in this case. Further, it is reasonable to assume that the tip apex is
Letters
Figure 1. (A) Electron micrograph of the carbon nanotube tip after the experiment. The tip appears slightly larger than the true diameter during the AFM measurements due to carbon deposition during SEM imaging. (B) Close-up AFM image of a multiwalled carbon nanotube typical of the type used as probes.
hydrophobic and of low surface roughness. These factors together with the high aspect ratio and robust construction make it the ideal probe of solvation forces. However, when using such a probe it is important that the measurement technique is sufficiently sensitive to detect the force interaction of molecules over a cross sectional area of less than 300 nm2. This is very small compared with the SFA where the radii of the crossed cylinders are usually of the order of 1 cm. For high sensitivity we have utilized a magnetic activation technique8 operating in both the dynamic off-resonance mode7 and resonance-tracking oscillator mode.16 Although we detect the presence of solvation shells using both methods we found the resonance-tracking oscillator method to give more clear and consistent data due to the superior noise level at reasonable measurement speeds. With this technique the cantilever is used as a resonator in an active feedback circuit. We use a constant amplitude oscillation and thus have ready access to both the frequency shift and dissipation caused by the tip-sample interaction. The latter being qualititatively related to the additional driving voltage to the magnetic coil needed to keep the amplitude constant throughout the experiment. The exact quality factor (Q) value can also be measured in our system by modulating the phase shift between the excitation and response of the cantilever and monitoring the induced shifts of the oscillation frequency. The Q value is defined as the product of the effective mass and the resonant frequency of the subsyem divided by the damping. In the case of AFM in liquid the
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J. Phys. Chem. B, Vol. 104, No. 26, 2000 6093
Figure 2. 12 × 12 nm2 area of the hydrophilic SAM showing faint periodicity and striped regions. The region indicated in the top righthand corner corresponds to the location of the solvation shells measurement shown in Figure 3. Some uncertainty in the exact location is due to thermal drift.
damping factor has three main components; the damping within the free lever, which remains constant during the experiment, long range hydrodynamic damping or squeezing of the liquid between the bulk of the lever and the sample and most importantly for the measurement of solvation shells the damping due to the tip-sample interaction. From the Q value we calculate a mean dispersive tip-water-sample force using the equation
Fdis ) (Qo - Q/QoQ)kl∆y/2
(1)
where kl is the stiffness of the cantilever, ∆y is the driven oscillation amplitude of the cantilever, Q is the quality factor with the tip-sample interaction and Qoi is the quality factor without the tip-sample interaction. The sample consisted of a hydrophilic self-assembled monolayer (SAM) of (COOH(CH2)10-SH on atomically flat Au(111) surface epitaxially grown on mica by vacuum deposition. To prepare the SAM, the Au substrate was immersed in 1 mM solution for more than 24 h. The solvent used was ethanol. After being removed from solution, the Au substrate was rinsed with pure solvent to remove the physisorbed multilayer and immediately placed in a beaker of milli-Q water. After being placed in the AFM, fresh milli-Q water was used in the liquid cell. Cantilevers were assembled with a magnetic particle coated with ultrahigh-vacuum (UHV) compatible epoxy glued behind the tip and a MWNT attached to the end of the tip. The final cantilever assembly had a normal stiffness of 2.2 N/m, resonant frequency of 14.332 kHz, and Q value of 6.8 in water. By using a constant-dissipation, AFM-imaging method,17 the surface was characterized on a nanometer scale within minutes of the solvation force measurements. In this way it was possible to try to relate solvation force measurements directly to specific features on the surface with nanometer-scale lateral resolution. 3. Results and Discussion Figure 2 shows a 12 × 12 nm image of the self-assembled monolayer taken under total immersion in milli-Q water. There seems to be some periodicity in the image of 1-1.2 nm separation. This is larger than the 0.5 nm expected and may be an indication that the molecules are not standing upright under the tip. Unfortunately, for these surfaces we found it difficult to obtain true molecular-resolution images, even by STM in UHV, so we have no alternative means at present to confirm
Figure 3. Graph showing the changes in driving voltage necessary for constant amplitude lever oscillations as a function of sample displacement. The zero on the x-axis is arbitrary. Calibrated Q values and calculated dissipative forces are shown at two values on the curve.
the apparent periodicity. However, stripes seen in this image and in images on a slightly larger scale may well be evidence of a “striped phase” that sometimes occurs on low molecular density SAMs.18 We commonly observed such stripes once the SAM had been immersed in water even when the initial selfassembly had occurred over several days, but not for similar samples observed in air. Figure 3 shows solvation shells measured in the region indicated in Figure 2. The measurement is performed by holding the tip at a particular lateral position and varying the sample height while recording changes in the dissipation (changes in voltage to the coil driver in order to keep the oscillation amplitude of the lever constant). The curves are oscillatory with a periodicity corresponding approximately to the diameter of a water molecule and the oscillatory amplitude (and hence dissipation) increasing for decreasing separation. This may be explained by layering of the water at small tip-sample separations. The dissipation in the dynamic measurement being higher when there are an integer number of layers between the tip and sample at the point of closest approach and lower when this separation does not correspond to an integer number of layers. In other words, higher dissipation occurs on impacting a stiffer arrangement of the water molecules. Although Figure 3 is one of the clearest examples of solvation shells, we observed similar oscillations in 25 of the 50 approach curves taken. Analyzing all the curves yielded a mean solvation shell separation of 2.2 Å. Some lateral dependence was also observed in that the observation of solvation layers was more consistently observed at the higher areas in the images although it is not yet clear why this should be the case. One possibility is that in this region interaction of the water with the hydrophilic end groups dominates over the interaction with the carbon chains, which may be exposed in regions of the SAM where the molecules are lying flat on the surface. By modulating the phase, it is possible to calibrate exact values for the change in Q value of the lever due to the tip interactions. This calibration was performed at two values in Figure 3 and the Q value changes converted to dissipation forces
6094 J. Phys. Chem. B, Vol. 104, No. 26, 2000 using eq 1. Normalizing the calculated value by dividing by the tip radius19,3 places the dissipative forces of the measured solvation shells in the range 1-50 mNm-1. These values lie surprisingly close to the values of the data points measured by Israelachvili et al. in KCl with SFA.6 We had expected our quantitative dispersive force measurements not to correspond directly to static repulsive force measurements: With our dynamic measurements we sample the average viscosity experienced by the tip during its 12 nm p-p oscillation cycle with the averaging weighted toward the interaction at the turning points of the oscillation, that is to say, the point of closest approach. 4. Conclusions One important aspect of these results is that forces appear to scale with the surface dimensions from the mesoscopic, as measured by surface forces apparatus, to the nanoscale, as we observe here. A fact of immediate importance to colloid systems. Also of importance is the observation of solvation shells on a nonrigid surface. Further investigations are underway to see if this extends to biological molecules where thermal fluctuations in the highly compliant structures might be expected to smear out the oscillatory character of the interaction force. Further, it has been suggested that the action of a dehydrating agent or water-structure breaker may be chiefly on the nearby surface rather than on the water.1 The reproducible measurement of water layers by our method should now enable one to study the influence of temperature and structure breakers on a patterned surfaces, a direct method of confirming or disproving the influence of the surface relative to the possible ‘unique’ properties of the water itself.
Letters Acknowledgment. S.P.J. thanks the European Commission for their generous support via a EUS&T fellowship. This work was partially supported by the New Energy and Industrial Technology Development Organization. References and Notes (1) Israelachvili, J. N.; Wennerstro¨m, H. Nature 1996, 379, 219. (2) Luedtke, W. D.; Landman, U. Comput. Mater. Sci. 1992, 1, 1. (3) Gelb, L. D.; Lynden-Bell, R. M. Phys. ReV. B 1994, 49, 2058. (4) Patrick, D. L.; Lynden-Bell, R. M. Surf. Sci. 1997, 380, 224. (5) Tabor, D.; Winterton, R. H. S. Nature 1968, 219, 1120. (6) Israelachvili, J. N.; Pashley, R. M. Nature 1983, 306, 249. (7) O’Shea, S. J.; Welland, M. E.; Pethica, J. B. Chem. Phys. Lett. 1994, 223, 336. (8) Jarvis, S. P.; Oral, A.; Weihs, T. P.; Pethica, J. B. ReV. Sci. Instrum. 1993, 64, 3515. (9) O’Shea, S. J.; Lantz, M. A.; Tokumoto, H. Langmuir 1999, 15, 922. (10) Cleveland, J. P.; Scha¨ffer, T. E.; Hansma, P. K. Phys. ReV. B 1995, 52, R8692. (11) Dai, H.; Hafner, J. H.; Rinzler, A. G.; Colbert, D. T.; Smalley, R. E. Nature 1996, 384, 147. (12) O’Shea, S. J.; Welland, M. E. Langmuir 1998, 14, 4186. (13) Ebbesen, T. W.; Ajayan, P. M. Nature 1992, 358, 220. (14) Yamamoto, K.; Akita, S.; Nakayama, Y. J. Phys. D 1998, 31, 34. (15) Nisijima, H.; Kamo, S.; Akita, S.; Nakayama, Y.; Hohmura, K. I.; Yoshimura, S. H.; Takeyasu, K. Appl. Phys. Lett. 1999, 74, 4061. (16) Du¨rig, U.; Steinauer, H. R.; Blanc, N. J. Appl. Phys. 1997, 82, 3641. (17) Jarvis, S. P.; Tokumoto, H.; Yamada, H.; Kobayashi, K.; Toda, A. Appl. Phys. Lett. 1999, 75, 3883. (18) Naselli, C.; Swalen, J. D.; Rabolt, J. F. J. Chem. Phys. 1989, 90, 3855. (19) Horn, R. G.; Israelachvili, J. N. J. Chem. Phys. 1981, 75, 1400.
