Visualizing Atoms, Molecules and Surfaces by Scanning

In the Laboratory

Visualizing Atoms, Molecules, and Surfaces by Scanning Probe Microscopy

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Kimberly Aumann, Karen J. C. Muyskens, and Kumar Sinniah* Department of Chemistry and Biochemistry, Calvin College, Grand Rapids, MI 49546; *[email protected]

The past two decades have seen a proliferation of research articles based on the techniques of scanning probe microscopy (SPM). This technique has provided visually stunning three-dimensional images of materials that have been beneficial to a wide range of scientific disciplines. Given its ability to visualize atoms, molecules, and surface features such as steps, pits, and surface reactions, this technique has appeal to an undergraduate setting. In fact, this Journal has published several articles that describe specific experiments using the SPM (1–4). A multidisciplinary SPM course was also introduced for undergraduate students at Arizona State University, and the essence of the course was published in “NSF Highlights” (5). In this paper, we wish to provide several examples of SPM experiments that we have carried out during a three-week undergraduate introductory course in scanning probe microscopy during our January interim term and as independent undergraduate research projects. Our goal is to give the reader a flavor for the broad range of experiments that can be done at the undergraduate level using SPM. Six students ranging from first-year to senior-level were enrolled in the interim course. The interim course was divided roughly in half: an introductory component and a project component. In the introductory component, students had both a theoretical and a laboratory introduction to the techniques of scanning tunneling microscopy (STM) and atomic force microscopy (AFM). Several Internet resources were made available as class texts, and a number of review articles were also provided to the students (6–10). The laboratory aspect included one-on-one training of each student on the scanning probe microscope. Students were taught the different modes of imaging in AFM, such as contact mode and intermittent contact mode (tapping mode). The students were also given increased independence in the lab so they could learn how to choose a cantilever based on the type of experiment and how to discriminate between a “good” image and an image containing tip artifacts. In the project component of the course, students were grouped in pairs and allowed to choose an independent project based on their interest from a list of instructor-chosen experiments. The students were required to update the rest of the class on their progress each day as well as present formal oral and written final reports on their projects. The scanning probe microscope used in our experiments was a Nanoscope IIIa multimode instrument (Digital Instruments) with capabilities of running the instrument in the AFM, STM, or Magnetic Force Microscopy mode. Three different scanners (0.4-µm, 12-µm, and 125-µm) were made available to the students. Vibration isolation consisted of either an air table (AFM experiments) or a concrete block suspended by bungee cords attached to a tripod (STM experiments). All AFM experiments were performed in air, and STM experiments were conducted both in air and in a drop of liquid. If AFM experiments are conducted in a liquid

medium with undergraduate students, we highly recommend using the O-ring provided with the fluid cell. The risk of damaging an expensive scanner is considerably higher when not using the O-ring, and setting up of experiments using liquids without an O-ring should be left in the hands of a more experienced student or researcher. Given the smaller volumes used in STM experiments, the scanners are usually not in danger of being short-circuited.1 In addition to the cost of the instrument itself, the major cost associated with running a scanning probe microscopy course is the high cost of SPM and STM probes. One way to control the cost is to utilize used AFM probes for training purposes: with the aid of a tip cleaner (Bioforce Laboratories Inc.) or a UV lamp, many old probes can be reused without noticeable tip artifacts. For example, our students used old tapping-mode tips during the training phase when they were learning to mount the tips into the tip holder.2 For the STM work, we have used both commercial tips (Pt– Ir, Digital Instruments) and our own mechanically-cut tips from Pt–Rh wire (87:13 percent Pt:Rh, Omega) using a wire cutter to cut the wire. The experiments described below can either be adapted to an upper-level laboratory sequence or run separately as part of a microscopy course. In choosing these experiments, our goal was to provide students with a broad array of experiments, since the student participants in the interim course were majors from several disciplines (engineering, chemistry, biochemistry, and biology). Details of the experimental procedures are provided in the supplemental section available from JCE Online.W Investigating DNA Molecules on Mica and AP–Mica Visualizing DNA in stretched, supercoiled, or relaxed forms provides undergraduate students in biology or biochemistry a sense of topology and function of this important molecule. Much of what is known about DNA comes from a typical biology or biochemistry text at the undergraduate level displaying scanning electron microscopy (SEM) images of DNA. The AFM has been successful in imaging both circular and linear forms of DNA on a mica surface with less sample preparation than required for the SEM (11–14). However, following appropriate sample preparation methods is a must when using the AFM to visualize DNA on a mica surface. Since both mica and DNA are negatively charged, the presence of divalent cations provides a necessary step for immobilizing DNA onto a mica surface. A recent method developed to immobilize DNA is to treat the mica surface with 3-aminopropyltriethoxy silane (AP), which provides a positive charge to the mica surface, thus enabling DNA to adhere to the surface more readily (15–16). In this project, students prepare an AP–mica surface, develop a protocol for imaging DNA on the mica surface, and compare the coverage

JChemEd.chem.wisc.edu • Vol. 80 No. 2 February 2003 • Journal of Chemical Education

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of DNA on AP–mica to the coverage of DNA on a freshly cleaved mica surface. Figure 1 shows representative images of identical amounts of DNA on AP-mica and untreated mica. AP–mica shows dense regions of DNA while the untreated mica shows much less DNA. This experiment demonstrates that DNA does adhere better to AP–mica than to the untreated mica surface. For routine laboratory experiments, however, it is sufficient to prepare and image DNA in the presence of divalent cations on a freshly cleaved mica surface given the success we have had in imaging DNA under these conditions. This experiment can be further expanded to visualize and compare single-stranded DNA (ss-DNA) and double-stranded DNA (ds-DNA). Since the AFM is ideally suited for measuring height changes in the subnanoscale regime, height changes

observed between ss-DNA and ds-DNA should provide students with information about the compliance of such molecules. In performing height-scale measurements, it is important to note that the width of the DNA results in an artificial measurement because of tip convolution, while the height of the DNA can be measured accurately by changing the set-point voltage or amplitude to minimize the loading force applied by the scanning tip to the surface. We have found that the height of the DNA obtained from AFM measurements conducted in air (~ 0.5–1 nm) show values smaller than its actual height (~ 2 nm) most likely due to the DNA– surface interactions or capillary effects that cause higher loading forces. Analysis of Microchip Memory Arrays and Circuitry Scanning probe microscopy is ideally suited to investigate semiconductors for quality control and failure analysis. In many thin-film research laboratories and in industry, AFM

Figure 1. AFM images of φx174 ds-DNA on a mica surface using tapping-mode in air. (a) DNA (10 ng/␮L) on freshly cleaved mica in the presence of Mg2+ ions. Scan size is 1.9-µm × 1.9-µm. (b) DNA (10 ng/␮L) on AP–mica surface. Scan size 1.2-␮m × 1.2-␮m.

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Figure 2. Memory arrays of a Toshiba EPROM memory chip. (a) Contact mode AFM image of the memory array. Scan size is 105-␮m × 105-␮m. (b) Digital camera image at 100-fold magnification. The boxed region shows the area of the AFM scan.

Journal of Chemical Education • Vol. 80 No. 2 February 2003 • JChemEd.chem.wisc.edu

In the Laboratory

is routinely used to measure roughness parameters, which provides an indication of the quality of thin-film surfaces and other semiconductors. In addition, observing defects such as missing atoms, step density, and dislocations provides a measure of the overall quality of the surface, since such defects influence the function of electronic circuits. Since microchips are readily available and inexpensive, the surfaces of integrated circuits (IC) are ideal examples to show students the intricate and complex designs and the layering of integrated circuits. In our interim course, we investigated a microchip memory array and a microchip circuit using the AFM. An AFM image shows a topographic view of the surface of the chip’s top layer. Chips contain many layers below the top surface layer that are not visible in an AFM image, but these layers do, however, affect the top surface layer by pushing up on surface wires. The advantage of using the AFM to image microchips is its ability to identify contaminants and defects that may lead to the failure of a chip.

Since undergraduate students may not be exposed to semiconductor theory or chip design, students are directed to search the database at Intel’s education Web site which contains a number of excellent articles to familiarize them with microchip fabrication and production (17). Figures 2–4 show memory arrays of a Toshiba EPROM memory chip and a Motorola micro-controller. It is rather trivial to image these surfaces in air using either contact-mode or tapping-mode AFM. The images shown below were obtained by students within a short time and were easily reproducible. The Toshiba EPROM memory chip is shown in Figure 2a. The memory arrays are elevated layers above the silicon surface. Owing to the height of the memory arrays, the underlying silicon surface appears dark. Figure 3a also shows an AFM image of memory (RAM), but in a Motorola micro-controller. In this image, the U-shaped tungsten plugs that go down to the transistors are visible. The vertical aluminum wires carrying electricity to and from the chips are also visible.

Figure 3. AFM image of RAM memory in a Motorola micro-controller. (a) Contact mode AFM image in air. Scan size is100-␮m × 80-␮m. (b) RAM memory, 100-fold magnification, using a digital camera. The boxed region shows the area of the AFM scan.

Figure 4. (a) and (b) AFM images of the Motorola micro-controller. Image size is 100-␮m × 100-␮m.


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A second region on the Motorola micro-controller was imaged as shown in Figure 4. Both Figures 4a and 4b show the layering beneath the circuit that affects the chip’s topography. A line of tungsten plugs connecting different levels of circuitry can be seen in the lower left corner of Figure 4a. Figure 4b shown in false color exemplifies the different levels underneath the circuit. The light gray areas are the most elevated and the black represents the lowest areas. The students who carried out this project found the study of microchip production to be very interesting, and the images they obtained from the AFM were able to give them precise measurements of surface features.

Bundled actin filaments are shown in Figure 6. The difference between bundled and network filaments is that the bundles appear to be wrapped around each other. Some bundles were found to have widths 5–10 times greater than the width of individual filaments. Individual filaments can be seen branching out from the bundles. Visualizing actin filaments appeals to students in biochemistry, as actin is an essential component of muscle as well as providing control of shape and movement to nonmuscle cells. From a study such as this, students can gain an understanding of the importance of structure and function of this important class of filaments.

Visualizing Filamentous Actin

Self-Assembled Monolayers on Graphite

Actin is an important protein involved in a variety of tasks within the cytoskeleton of different cells in living organisms. When induced by K+, Mg2+, and Na+, actin is polymerized from its globular form into right-handed helical filaments. Filaments thus produced can be immobilized on a mica surface and visualized by AFM (18). We have developed a simple procedure for visualizing actin on a mica surface at room temperature using the AFM; further details are provided in the supplemental section available from JCE Online.W Individual actin filaments, networks, and bundles are easily imaged using this method (Figures 5 and 6). In the networks observed, the actin filaments were several micrometers in length with considerable branching within each filament. The branched filaments have the same height as individual filaments, indicating that the filaments are not coiled around each other. As an exercise, students can be required to measure the branching angles, periodicity of actin filaments (~ 38 nm from our measurements), and heights of individual actin filaments and bundles, and then make comparisons to literature values.

Scanning tunneling microscopy (STM) has been used to obtain images of self-assembled monolayers of organic molecules on surfaces such as graphite and molybdenum disulfide (7, 19–26). A previous article in this Journal described a laboratory investigation of self-assembled monolayers of 11bromoundecanol on graphite (3). In this project students attempt to obtain STM images of three different 30-carbon hydrocarbons (triacontane, triacontanol, and triacontanoic acid) on graphite in order to observe the ways in which the functional group influences the structure and appearance of the self-assembled monolayers. In addition, the students make observations and measurements that show the influence of surface–adsorbate and intermolecular interactions on the structure of the film. The students use a procedure similar to those found in the literature (3, 23, 24) to prepare the samples and take STM images. The goal of the project is to observe various aspects of the structure and dynamics of the self-assembled monolayers. On the larger-scale (100-nm × 100-nm), the students look at the number of domains in the imaged area. If multiple domains exist, as in Figure 7a, the angle between the

Figure 5. Networked actin filaments on mica (25 ␮mol/mL) imaged by tapping-mode AFM. Scan size is 4.2-␮m × 4.2-␮m.

Figure 6. Bundled actin filaments on mica imaged in tapping-mode AFM. Scan size is 3.5-␮m × 3.5-␮m.

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direction of the rows of molecules (lamellae) from one domain to another can be measured. This angle is expected to be 60⬚ or 120⬚ when the hydrocarbon axes are aligned with the graphite lattice, as they are with these long-chained hydrocarbons (19). Students can also look for a Moiré pattern3 in triacontanoic acid as shown in Figure 7. This Moiré pattern has been explained as arising from the incommensurability of the molecular film with the graphite lattice due to the large carboxylic acid group (19). The larger-scale view of the monolayer can also show the dynamic nature of these physisorbed monolayers. Sometimes changes in domain boundaries can be observed over time; sometimes the organic monolayer is no longer observable and only the graphite substrate can be imaged.

Smaller scan sizes such as the images shown in Figures 7b and 8 can be used to determine more of the detailed structure of the monolayer. Imaging the graphite surface underneath the monolayer can provide distance and angle calibration measurements for the molecular film, though approximate angles and distances can be obtained without calibration. Students can measure the width of the lamellae and compare this to the molecular length to show that these molecules form monolayers in which the molecules are aligned on the surface with the molecular axis parallel to the graphite surface (19, 20). Under favorable imaging conditions the angle between the molecular axis and the lamella direction can also be measured. A 90⬚ angle results in the greatest amount of intermolecular van der Waals interactions and is observed for triacontane and triacontanoic acid (Figure 7b), whereas Figure 8 illustrates the 60⬚ angle that is favored for triacontanol due to hydrogen-bonding interactions (21, 25). The students can also try to observe atomically-resolved images. In an atomically-resolved image, the bumps along the molecular axis have been analyzed as corresponding to the hydrogen atoms protruding furthest from the surface (20, 21, 26). Other Experiments In addition to what has been described above, there are several other experiments that we have tried. Oxides such as MgO, calcite, and halite have surfaces that can be easily cleaved and imaged in air or under varying pH conditions. These surfaces show a wide distribution of step heights or wandering steps depending on the cleavage plane. These surfaces can also be used to investigate the kinetics of dissolution in aqueous or buffered solutions. We have also used gold colloids (Sigma) on a mica surface to calibrate heights in the 5-nm and 10-nm region. Such calibration measurements in the nanoscale regime provide students with an appreciation of the reliability of heights measured from features or protrusions especially on compliant surfaces.

Figure 7. STM images of triacontanoic acid on graphite. Scan size is (a) 100-nm × 100-nm; (b) 25-nm × 25-nm. Bar is one molecular length.

Figure 8. STM image of triacontanol on graphite. The bar shows one molecular length. Scan size is 20-nm × 20-nm.


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Hazards

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Students should wear appropriate protective gloves, clothing, and eye protection when handling chemicals and biochemicals, and work in areas with adequate ventilation. The following chemicals are known to be hazardous and MSDS information sheets should be read prior to use for further details and advice on protective measures. 3-Aminopropyltriethoxy silane (AP) is corrosive and is a harmful liquid that targets nerve and liver organs via inhalation, ingestion, or skin absorption. It should be handled using appropriate gloves in a fume hood to avoid skin contact and inhalation of vapors. Furthermore, AP is moisture sensitive and should be stored under nitrogen in a desiccator. 2-Mercaptoethanol must be handled with great care using appropriate gloves in a fume hood: it is highly toxic via inhalation, skin contact, and ingestion; it is readily absorbed through the skin; it is irritating to the eye and skin; and it has a bad odor. Phalloidin must also be handled with great care using appropriate gloves in a fume hood: it is highly toxic via inhalation, skin contact, and ingestion. Ammonium acetate, CaCl2, Na2ATP, KCl, and MgCl2 are all skin and eye irritants and should be handled using gloves. Triacontane, triacontanoic acid, triacontanol, and phenyloctane are all skin and eye irritants whose toxicological properties have not been thoroughly investigated; appropriate chemical safety precautions should be used in handling these chemicals. Ethanol is highly flammable and should be kept away from flames.

Details of the experimental procedure including protocols for imaging of the materials are available in this issue of JCE Online.

Conclusions In this paper we demonstrate several experiments that are feasible with a scanning probe microscope for undergraduate students. All the experiments described in this paper were done by undergraduate students. Without too much effort the experiments described here can be easily adapted to an upper-level undergraduate laboratory course or run with individual students as independent projects. The exposure to SPM techniques at the undergraduate level will certainly enable students to gain insights into areas of science previously not possible. Acknowledgments We wish to thank the students who worked on the projects presented in this paper: Mark Schotanus, Kimberly Aumann, Julia Heetderks, Brad Veldkamp, Nathan Morris, Tae-Sung Kwon, and Carissa Hockema. Eric Arnoys (Calvin College) is thanked for the preparation of filamentous actin, and Elizabeth Howell (Calvin College) is thanked for providing the DNA samples. We also wish to acknowledge the support of a grant from the National Science Foundation (CHE 9871225) to purchase the scanning probe microscope, and support from the donors of the Petroleum Research Fund, administered by the ACS, for partial support of the work presented here.

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Supplemental Material

Notes 1. Scanning probe microscopes with the piezo attached to the probe tip rather than the sample are better suited for experiments in liquids. 2. Owing to the high cost of tapping-mode tips, the instructor mounted the new tapping-mode tips used for the project work. 3. A Moiré pattern forms when two or more patterns are superimposed. See http://www.sandlotscience.com/Moire/Moire_frm.htm (accessed Nov 2002) for an example of a Moiré pattern.

Literature Cited 1. Coury, L. A., Jr.; Johnson, M.; Murphy, T. J. J. Chem. Educ. 1995, 72, 1088. 2. Polar, J. C. J. Chem. Educ. 2000, 77, 1198–1200. 3. Giancarlo, L. C.; Fang, H.; Avila, L.; Fine, L. W.; Flynn, G. W. J. Chem. Educ. 2000, 77, 66–71. 4. Rapp, C. S. J. Chem. Educ. 1997, 74, 1087–1089. 5. Glausinger, W. S.; Ramakrishna, B. L.; Garcia, A. A.; Pizziconi, V. J. Chem. Educ. 1997, 74, 310–311. 6. The following Internet sites provide excellent introductions to the SPM technique: A Practical Guide to Scanning Probe Microscopy. http://www.thermomicro.com/spmguide/contents.htm (accessed Oct 2002); Scanning Probe/Atomic Force Microscopy: Technology Overview and Update. http://www.di.com/ AppNotes/AmLab/AL-SPMmain.html (accessed Oct 2002). 7. Frommer, J. Angew. Chem., Int. Ed. Engl. 1992, 31, 1298– 1328. 8. Hamers, R. J. J. Phys. Chem. 1996, 100, 13106–13123. 9. Rohrer, H. Ultramicroscopy 1992, 42–44, 1–6. 10. Quate, C. F. Surf. Sci. 1994, 299, 980–995. 11. Hansma, H. G.;Vesenka, J.; Siegerist, C.; Kelderman, G.; Morett, H.; Sinsheimer, R. L.; Elings, V.; Bustamante, C.; Hansma, P. K. Science 1992, 256, 1180–1184. 12. Hansma, H. G.; Sinsheimer, R. L.; Li, M.–Q.; Hansma, P. K. Nucleic Acids Res. 1992, 20, 3585. 13. Bezanilla, M.; Drake, B.; Nudler, E.; Kashlev, M.; Hansma, P. K.; Hansma, H. G. Biophys. J. 1994, 67, 2454–2459. 14. Hansma, H. G. in Procedures in Scanning Probe Microscopy; Colton R. J., Engel, A., Frommer, J. E., Gaub, H. E., Gewirth, A. A., Guckenberger, R., Rabe, J., Heckl, W. M., Parkinson, B., Eds.; John Wiley & Sons: New York, 1999; pp 389–393 and references therein. 15. Lyubchenko, Y. L.; Lindsay, S. M. in Procedures in Scanning Probe Microscopy; Colton R. J., Engel, A., Frommer, J. E., Gaub, H. E., Gewirth, A. A., Guckenberger, R., Rabe, J., Heckl, W. M., Parkinson, B., Eds.; John Wiley & Sons: New York, 1999; pp 493–495. 16. Umemura, K.; Ishikawa, M.; Kuroda, R. Anal. Biochem. 2001, 290, 232–237. 17. Intel Innovation in Education. http://www97.intel.com/education/index.asp (accessed Oct 2002). 18. Shao, Z.; Shi, D.; Somlyo, A. V. Biophys. J. 2000, 78, 950– 958.


In the Laboratory 19. Rabe, J. P.; Bucholz, S. Science 1991, 253, 424–427. 20. Cyr, D. M.; Venkataraman, B.; Flynn, G. W. Chem. Mater. 1996, 8, 1600–1615. 21. Claypool, C. L.; Faglioni, F.; Goddard, W. A.; Gray, H. B.; Lewis, N. S.; Marcus, R. A. J. Phys. Chem. B 1997, 101, 5978– 5995. 22. Giancarlo, L. C.; Flynn, G. W. Annu. Rev. Phys. Chem. 1998, 49, 297–336. 23. Yeo, Y. H.; McGonigal, G. C.; Thomson, D. J. in Procedures in Scanning Probe Microscopy; Colton R. J., Engel, A., Frommer, J. E., Gaub, H. E., Gewirth, A. A., Guckenberger, R., Rabe, J., Heckl, W. M., Parkinson, B., Eds.; John Wiley

& Sons: New York, 1999; pp 340–345 and references therein. 24. Venkataraman, B; Flynn, G. W.; Wilbur, J. L.; Folkers, J. P.; Whitesides, G. M. J. Phys. Chem. 1995, 99, 8684–8689; Cyr, D. M.; Venkataraman, B.; Flynn, G. W.; Black, A.; Whitesides, G. M. J. Phys. Chem. 1996, 100, 13747–13759; Giancarlo, L.; Cyr, D. M.; Muyskens, K.; Flynn, G. W. Langmuir 1998, 111, 321–335. 25. Venkataraman, B.; Breen, J. J.; Flynn, G. W. J. Phys. Chem. 1995, 99, 6608–6619. 26. Liang, W.; Whangbo, M.-H.; Wawkuschewski, A.; Cantow, H. J.; Magonov, S. N. Adv. Mater. 1993, 5, 817.


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