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Journal of Biomolecular Structure and Dynamics Publication details, including instructions for authors and subscription information: http://www.tandfonline.com/loi/tbsd20
Atomic Force Microscopy Imaging of Double Stranded DNA and RNA Yuri L. Lyubchenko
a b c
d
, Alexander A. Gall , Lyuda
c
c
S. Shlyakhtenko , Rodney E. Harrington , Bertram b
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L. Jacobs , Patrick I. Oden & Stuart M. Lindsay a
a
Departments of Physics
b
Departments of Microbiology , Arizona State University , Tempe , AZ , 85287 , USA c
Department of Biochemistry , University of Nevada Reno , Reno , NV , 89557 , USA d
MicroProbe Corp , 1725 St. SE#104, Bothell , WA , 98021 , USA Published online: 21 May 2012.
To cite this article: Yuri L. Lyubchenko , Alexander A. Gall , Lyuda S. Shlyakhtenko , Rodney E. Harrington , Bertram L. Jacobs , Patrick I. Oden & Stuart M. Lindsay (1992) Atomic Force Microscopy Imaging of Double Stranded DNA and RNA, Journal of Biomolecular Structure and Dynamics, 10:3, 589-606, DOI: 10.1080/07391102.1992.10508670 To link to this article: http://dx.doi.org/10.1080/07391102.1992.10508670
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Journal of Biomolecular Structure & Dynamics, ISSN 0739-1102 Volume 10. Issue Number 3 (1992), ®Adenine Press (1992).
Atomic Force Microscopy Imaging of Double Stranded DNA and RNA Yuri L. Lyubchenko*1•2•3 , Alexander A. Ga114 , Lyuda S. Sh~akhtenko 3 Rodney E. Harrington 3, Bertram L. Jacobs , Patrick I. Oden1 and Stuart M. Lindsay1 Downloaded by [Purdue University] at 13:15 13 April 2015
1
Departments of Physics and 2Microbiology Arizona State University Tempe, AZ 85287, USA 3
Department of Biochemistry University of Nevada Reno Reno NV 89557, USA ~icroProbe Corp 1725 St. SE# 104 Bothell, WA 98021, USA
Abstract A procedure for imaging long DNA and double stranded RNA (dsRNA) molecules using Atomic Force Microscopy (AFM) is described. Stable binding of double stranded DNA molecules to the flat mica surface is achieved by chemical modification of freshly cleaved mica under mild conditions with 3-aminopropyltriethoxy silane. We have obtained striking images of intact lambda DNA, Hind III restriction fragments oflambda DNA and dsRNA from reovirus. These images are stable under repeated scanning and measured contour lengths are accurate to within a few percent. This procedure leads to strong DNA attachment, allowing imaging underwater. The widths of the DNA images lie in the range of20 to 80nm for data obtained in air with commercially available probes. The work demonstrates that AFM is now a routine tool for simple measurements such as a length distribution. Improvement of substrate and sample preparation methods are needed to achieve yet higher resolution.
Introduction Scanning Probe Microscopy, which mostly includes Scanning Tunneling Microscopy (STM) and Atomic Force Microscopy (AFM), is a relatively new tool which *Corresponding author at the Department of Physics, Arizona State University, Tempe, AZ 852871504, USA Lyubchenko, Y.L. and Shlyakhtenko, L.S. are on leave from the Institute ofMolecular Genetics, Russian Academy of Sciences, Moscow, Russia.
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has enormous potential importance to structural biology. The STM and AFM offer unique advantages in their potential for very high resolution of DNA. RNA and their complexes with proteins and small ligands in the absence of stains, shadows and labels. Furthermore, the instruments can be operated in air or liquids. The latter is particularly important for resolving fully hydrated structures. They are theoretically capable of resolving structural details at the level of atomic dimensions, provided the specimen is dynamically stable. The prototype STM instrument was conceived by Binnig, Rohrer and coworkers (1) an invention for which Binnig and Rohrer were awarded the 1986 Nobel Prize in Physics (2). The first AFM was a direct descendent of this early instrument (3), but capable of imaging nonconducting as well as conducting surfaces. Although these instruments are, in principle, capable of imaging biological molecules and ultra structures to nearly atomic resolution and, hence, have great potential in such applications as micro-structural studies of DNA. RNA and other biological macromolecules, this potential has, so far, not been realized. For example, artifactual images may have been a problem in certain STM studies of macromolecules adsorbed on graphite surface (4,5), and the underlying theory of image contrast is not well understood. This appears to be a more serious problem with the STM, although the problem is not trivial with the AFM. However, an immediate practical limitation of the application of AFM and STM to structural studies of biological macromolecules is sample preparation. The macromolecules must be tethered to the substrate surface in order to avoid resolution-limiting motion occasioned by the sweeping tip during scanning (6). A number of procedures for sample preparation have been described, but, to our knowledge, no routine and reliable methods have been reported until recently. Most STM images of DNA and RNA published recentlywere obtained bydryingoutofDNAsolutions of a fewmg/ml onto scanning substrates. DNA aggregates are often stable, but do not allow one to image individual molecule (see reviews 5,7 and references therein). Lindsay et al. (5) have developed the use of electrochemical methods for holding DNA in place on a gold electrode as it is imaged with an STM. The DNA is held onto to a positively charged electrode and imaged under potential control (8). This procedure yields high resolution (better than can be obtained with the AFM at present) and the images are not subject to the artifacts that have plagued graphite surfaces (4). It has even proved possible to obtain images of single bases in based stacked aggregates of purines (9). However, the adsorption process is highly length selective, favoring small molecules. Furthermore, it offers no possibility of imaging large insulating clusters (such as DNA-protein complexes). Another approach to tethering DNA is chemical modification of the substrate. The first attempt was made by Cricenti et al.. (10). We have recently suggested a method for covalently tethering DNA to a graphite substrate (11). However, the graphite surface is not a suitable medium for STM studies ofbiopolymers (in our view) because of the problem of artifacts (4,5). The AFM permits insulating materials to be used as substrates, and freshly cleaved mica produces a much better surface than most graphite samples. We therefore set out to devise a procedure for chemical modification
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ofthe mica surface for studies of DNA (and other biopolymers) (12,13,14). The procedure we have developed is simple and robust and even works well underwater(l4) Here, we will describe some of its strengths and weaknesses. Over the same period of time, other workers have devised alternative strategies for reliable tethering of DNA for AFM studies (15-20). It is of interest to compare the published results to date. The first AFM study of DNA exploited the adhesion of DNA to wet glass (21 ). Subsequent work used mica surfaces treated with AIC13 (22). Resolution was limited by the tendency of the AFM tip to push DNA around (6) so better "tethering" of the molecule to the substrate is required. An ionic treatment of mica for binding DNA has been described by Bustamante et al.. (15) and Vesenka eta/.. (16). This permits reliable imaging in an AFM with a resolution approaching tens of nm. Hansma et a/. ( 17) have used this approach to obtain a significant improvement in resolution by imaging under a covering layer of propanol. Yang eta/. (18) used a conventional cytochrome-c modification of DNA to hold the molecules onto a carbon film that had been deposited on mica. Thundat eta/. (19) and Zenhausem eta/. (20) have reported that reliable results were obtained simply by applying the DNA to mica from an ammonium acetate (19) or MgC12 (20) buffer. Bustamante et al. (15) report that specially manufactured super sharp tips had to be used for reliable imaging. These tips have also been used by Hansma et al. (17). The other reports (12-14,19,20) were based on direct use of commercial probes with no special modification. The super-sharp tips do give better resolution in most cases, and resolution is also improved by imaging under propanol (17) or water (14). However, comparison of the published images suggest that other factors are important also. In this paper, we describe our procedure for imaging long DNA molecules chemically bound to mica. We give examples of a number of DNA and RNA images, images obtained in water, images of complex self-assembeled aggregates of DNA, and also some illustrations of effects caused by changes in the tip during scanning. We end with a discussion of our results (and those of the other groups) in order to clarifY the strengths and weaknesses of this new technique in its present state of development Materials and Methods Mica Modification Procedure Modification in liquid
We used freshly cleaved strips of ruby mica. Pieces of mica were incubated in freshly prepared solutions of 3-aminopropyltriethoxy silane (APTES, Aldrich, USA) in toluene or dimethylformamide for several hours with mild shaking followed by rinsing with pure solvent and drying. Dried strips were immersed into methyl iodide (Aldrich, USA) for one hour and dried again. Modification In Vapors
Freshly cleaved strips of mica were left in the APTES atmosphere created in a 21
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·~t
Mel
1--o-y1(CH2 ) 3 NH 2 OEt
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A Figure 1: (A) The steps in modification ofmica substrate-s with 3-aminopropyltriethoxy silane (APTES). Me is the methyl group and Et is the ethyl group. The silicon oxide group on the left is part of the mica surface. (B) Fraction of total DNA bound against exposure time. (II) data for unmodified mica, (.6.) mica treated with APTES alone and(*) mica methylated after APTES treatment. The standard deviation of data is the symbol size unless otherwise indicated.
glass desiccator containing a small pool of APTES under ambient conditions for 2 hours. The methylation procedure was the same as above.
DNA Samples Preparation Modified mica strips were immersed into DNA or RNA in Tris-HCl buffer solution
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Fraction of Total DNA Bound (%)
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Figure 1: continued
(pH 7), containing lOmM Tris-HCl, 10-20 mM NaCl, 5 mM EDTA and incubated at room temperature for a specified time which varied usually between 1 and 2 hours. The concentration of DNA was varied in different experiments between 0.1 and O.Ql flg/ml for different samples. The lowest concentration, 0.01 flg/ml, was used in experiments with lambda DNA The concentration ofHind m fragments oflambda DNA was around 0.1 flg/ml. Both DNA samples were purchased from New England BioLabs, USA and used without additional purification. After the adsorption stage had been completed, the samples were rinsed with deionized water (NanoPure water system, Barnstead, 2555 Kerper Bvd. Dubuque, lA), blotted at the edge and vacuum dried.
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dsRNA Samples
Reovirus genome dsRNA was isolated from purified virions by a standard technique (23). Briefly, the virus was disrupted by extraction with phenol and nucleic acids were concentrated by precipitation with ethanol. Genomic dsRNA was purified by gel filtration (24).
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Radiolabeled Assay
Mica substrates were incubated for different times in a solution of 32P-labeled Hpall restriction fragments ofPBR322 plasmid DNA (New England Biolabs) at a concentration of0.2 ng/ml in 10 mM Tris-HCl buffer (pH7.5). Ratios of bound to total DNA were calculated from the radioactivity of the rinsed mica plates and of the DNA solution before and after the binding procedure. Apparatus
Imaging was carried out on a N anoScope II STM/AFM from Digital Instruments (6780 Cortona Drive, Santa Barbara, CA) using commercial AFM cantilevers, also from Digital Instruments (spring constant 0.58 and 0.12N/m). The radius of curvature of the end of these tips has been estimated directly from images of whisker crystals (25) and it lies between 20 and 50 nm. The liquid cell used for AFM in this work was an electrochemistry cell fabricated by Digital Instruments, Inc. Results and Discussion Basis of the Mica Modification Procedure
The treatment of the mica substrate is shown schematically in Figure IA The amino groups of APTES are bound covalently to the freshly cleaved mica surface, giving it similar properties to an anion exchanger used in affinity chromatography. Its anionic affinity is enhanced by the substitution of methyl groups for the amino protons using methyl iodide (Step 2). On contact with water (Step 3), the ethoxy groups of the covalently bound APTES moieties are hydrolyzed to OH. We have obtained good AFM images using both the methylated surface (step 2) and the aminopropyl surface (step 2 omitted). Binding of DNA to Modified Mica
We checked the modification procedure with 32P-labeled Hpall restriction fragments of pBR322 ~lasmid DNA Mica substrates, treated with APTES, were incubated in a solution of 2P- DNA for a given period, washed thoroughly and the radioactivity of both mica samples and the DNA solutions was determined. These data yield the fraction of total DNA molecules initially in solution that became bound by the surface. Results comparing non-treated mica, mica modified using APTES only (step I on Figure lA) and mica modified by APTES followed by methylation (step 2) are shown in Figure 1B. DNA binding to the unmodified mica surface is insignificant
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Figure 2: AFM images of mica substrates modified with different concentrations of APTES. Immediately after cleaving, mica plates were treated as described in Figure 1. The surfaces were immersed in toluene solutions offresh APTES at different concentrations, incubated overnight, rinsed with pure toluene and dried. (A) 0.1 M APTES; (B) 0.1 mM APTES; (C) O.l!!M APTES. Bar size is 1 11m.
(radioactivity of the samples is very close to the background). Mter 4 hours of incubation, 0.25% of total DNA molecules were bound to the surface modified with APTES alone. This fraction was increased by an order of magnitude when the surface was methylated. We have investigated the nature of the forces holding DNA on the surface by incubation of DNNmica samples in highly concentrated salt solutions. Mica samples incubated for4 hours in 0.2M NaCl solution resulted in desorption of 50% of DNA molecules. In 2M NaCl, only 20% of DNA remained on the surface. These results suggest that (relatively nonspecific) electrostatic interaction is a predominant force holding DNA molecules onto the aminopropyl mica surface. Suiface Structure ofAP-Mica
AFM images of a mica surface modified with APTES at different concentrations (AP mica) are shown in Figure 2. At high APTES concentration (0.1 M) the surface is extensively and somewhat irregularly modified (Figure 2A) but the irregularities are reduced with decreasing APTES concentration (Figure 2B) so that at 0.1 f.lM the surface is uniform to within the resolution of the AFM images (Figure 2C). The dark bands that cross the image are instrumental artifacts which occur at large scan amplitudes. They do not rotate as the sample is rotated relative to the scan head and a series of small range scans taken over the same area verifies that the surface is flat. These dark bands vary with alignment of the AFM optics, and we believe that they are caused by interference between the reflected beam and laser light scattered from objects on the substrate. Imaging of DNA as shown below utilized mica surfaces treated with fresh preparations of APTES at concentrations that did not exceed 0.1 flM. Imaging of Lambda DNA Restriction Fragments
Here we illustrate the procedure with long linear fragments of DNA Treated substrates were incubated with a solution of Hind III restriction fragments oflambda DNA, rinsed with water, dried and scanned with the AFM in air to yield the images shown in Figure 3. Typical results for this sample are shown in Figure 3A The background is uneven, owing to the interference fringes described above. The most
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Figure 3: (A) AFM image of Hind III restriction fragments of A. phage DNA; raw data showing background fluctuations. Bars are 2 11m markers. (B) shows the same area after high pass filtering to remove background variations. (C) shows the same area as in (B) after 30 min. of continuous scanning at contact forces between 40 and 90 nN. (D) images of well separated fragments including the 23.1 kb restriction fragment (marked with an arrow).
noticeable features are patches that appear to be due to aggregated DNA However, fine filaments can be seen crossing the substrate between the aggregates. These individual strands generally can be followed easily and measured strand lengths correspond well with known restriction fragments. The height of the strands is -2 nm (this can vary considerably), but their width is always much greater. We find that the width ofimages depends upon the tip used in the AFM, as might be expected (6). For a spherical tip (radius R) scanning a cylindrical molecule, it is straightforward to show (13) that the width of the resulting image is 4-j(Rr). For a parabolic tip, the result(15) is [4(R+r).J(r(R-r)]IR. By taking traces across images such as that shown in Figure 3A, we find that the full-width of the DNA imaged in air varies from 20 to 50nm (and sometimes, even broader). This is consistent with the range oftip radii
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Figure 4: AFM images of whole lambda DNA (B), (C) and (D) are high pass filtered. (A) is raw data. The convoluted molecule in (A) starts at the point labeled "1" and finishes at the point labeled "2". A more extended molecule is shown in (B) and (C) (the path of the molecule is pointed to by arrows). Two adjacent images were taken and the overlap is indicated by the common point labeled "X". (C) is a zoomin on the end of the molecule (shown in a box labeled "D" on (C).
determined experimentally. Bustamante eta/. (15) and Hansmaetal. (17) have also demonstrated that the radius of the AFM tip influences the resolution of isolated molecules. However, our discussion will show that other factors contribute to this apparent broadening effect. The instrumental scanning artifacts observed in Figure 2C correspond to several nm of topography and limit the contrast with which individual strands can be displayed. Figure 3B shows the same scan as in Figure 3A after it has been high-pass filtered to remove background variations. Individual strands can be followed easily. Figure 3C is an image of the same area after continuous scanning for 30 minutes at
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Figure 5: AFM images of dsRNA from reovirus. (A) 3.5 f.im X 3.5 J.lm scanning area and (B) 1.75 J.lm X 1.75 J.1ffi scanning area.
contact forces between 40 and 90 nN. Figures 3B and 3C are virtually identical, indicating the stability of DNA binding to this substrate, even at this fairly large contact force. Figure 3D shows an area in which many well-separated fragments can be found. In cases where the molecular contours can be followed without ambiguity, their lengths correspond to the known Hind III restriction fragments oflambda DNA. In particular, note that the whole ofthe 23.1 kb fragment is seen clearly in this one scan (arrow, Figure 3D). Its contour length is 7.5 ± 0.3 !Jm which is close to the 8 !Jmexpected if the fragment maintains a B-form base stacking. (A-form stacking would result in a length of 6.6 !Jm.) Imaging of the Whole Lambda DNA
The complete Aphage genome of 48,502 bp (16.4 !Jm length, assuming B-form DNA) is shown in Figure 4A. The molecule follows a very convoluted path with many loops (starting at "1" and finishing at "2", Figure 4A). In order to obtain a contour length, we used the public domain software NIH Image (1.40) to track it pixel by pixel. The measured size was 16.4± l!Jm, in good agreement with the expected length. A quite different type of DNA topography was obtained on a different sample (from a different preparation) and is shown in Figure 4B. The molecule is much less convoluted (its path is marked by the vertical arrows). It starts at the left of the image and continues off the right hand side. It is followed by moving the scan to the right and reimaging as shown in Figure 4C. A feature on the surface has been marked with an "X" on both Figures 4B and 4C, so that the two images can be indexed. The molecule continues on to the right, where it ends in a fairly convoluted set of loops. The end (enclosed in the box labeled "D" on Figure 4C) is shown scanned at higher magnification in Figure 4D. Using all three of these images together with the contour-measurement tools in NIH Image, we estimate the contour
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Figure 6: AFM images oflambda DNA in air (A) and under water (B). Samples were scanned using the liquid cell with contact forces ~ 60nN for scanning in air and 10-12 nN for scanning under water.
length of the whole molecule as 15± 1.0 IJ.ID, also in satisfactory agreement with that expected for intact A. phage DNA. (The widths of these images lay between 20 and 40nm). The fact that images of these large DNA molecules are both stable and reproducible and lead to such high accuracy in contour length measurements makes AFM an attractive method for physical mapping applications. AFM Studies of dsRNA
We have investigated dsRNA sample extracted from reovirus (26). This virus is represented by a multisegment genome (27). Our goal in imaging these molecules was to demonstrate the application of this method for routine length measurements. These are more difficult to make by conventional means for RNA because of the lack of restriction fragments with which to calibrate gels. Images of dsRNA molecules are shown on Figures 5A and 5B. The substrate preparation procedure was identical to that used for DNA. Figure 5A shows the data for a scanning area of 3.51J.m X 3.51J.m. This image clearly demonstrates polydispersity of the RNA sample (corresponding to the various fragments of the genome). Zooming in on parts of such pictures permits more accurate length measurements as shown in Figure 5B (1.7 11m X l.71J.m scan). For example, the length of molecule 1 on Figure 5B is 0.85 11m and molecule 2 is 0.4l!J.ID which corresponds to 3.03 kb and 1.45 kb respectively (using 2.8A as a rise-to-residue distance for dsRNA (28)). Statistics were obtained for a large number of molecules and complete data will be published elsewhere (26). The results showed that RNA length varies between 0.3 and 1.5 IJ.ID, which is in a good agreement with the data obtained using other techniques (27). Thus, the procedure we have described makes AFM a routine technique for length measurements of DNA and RNA. DNA Imaging Under Water
Using APTES treated mica substrates, we have been able to image lambda DNA
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Figure 7: AFM images oflambda DNA (A) 10 llffi X 10 j.lm scanning area. (B) 4.5j.lm X 4.5j.lm scanning area. (C,D) 1.5 j.lm X 1.5 j.lm area. (A), (B) and (C) are high pass filtered, (D) is raw data.
underwater (14). Other work described to date has been carried out in air, or under a poor solvent like propanol, because the DNA was not stable under water (17). (Actually, the first AFM images obtained were taken on glass under water (21) but they were of poor quality and difficult to reproduce). We have published images of the same molecule taken both under water and in air elsewhere (14). Images obtained in air varied in full width from 20 to 80 nm. Images obtained in water varied from I 0 to 25 nm in full width. This result is similar to the narrowing observed by Hansma et al. when imaging under propanol (they also used custom-made super-sharp tips, the combination producing a resolution record that approached 3nm on occasion (17)). These observations imply that the resolution cannot be controlled by the tip radius alone (the same tip was used for imaging in air and under water). At very high resolution, the deformation of the tip and substrate may be important (14,29). However, given the elastic properties of silicon nitride (the tip material) and mica
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Figure 8: Images of DNA toroids (scale indicated by bars). All images are raw data.
(the substrate material), deformations on the order of tens of nm are simply not possible at contact forces of -nN (14,29-31). Other factor(s) must play a role in change of resolution between experiments in air and experiments in water. DNA and RNA Morphology on AP-Mica Swface
In addition, we have noted interesting effects which can alter the fidelity of AFM imaging in capturing solution DNA conformations using modified mica substrates as described here. They appear to result from uneven surface characteristics of the mica substrates, presumably due to non-uniform mica modification with APTES. Examples are shown in Figures 6A(scanned in air) and 6B (scanned underwater). Common features of the AFM images under water for this sample are blurred regions which indicate localized DNA mobility during scanning. However, the overall binding of the DNA is strong as the images are stable during repeated scanning, i.e. they retain their characteristic features and do not change their position. The molecules also do not disappear if allowed to remain under water for long periods of time. We believe that the images of Figures 6A and 6B are of DNA molecules that are bound strongly to the surface at certain places and relatively weakly at others. The weakly bound regions interact dynamically with the scanning tip producing the regions oflow resolution observed. This explanation is in accord with the experiments using radio labelled DNA to determine overall binding, which did not reveal the difference between this substrate and any other ones. Another phenomenon which can be observed on an under-modified surface is extensive DNA agregation. The aggregates observed on the AP-mica surface are of interest in their own right. Obvious examples of extensive condensation are shown in Figures 7A and zoomed images of this sample are shown in Figures 7B,7C and 7D. The substrates used for this sample preparation are found to bind the DNA -50-fold less efficiently as determined using radiolabeling methods. In Figure 7A, we show a surface which is heavily deposited with condensed DNA "blobs". The results of zooming-in and scanning over the upper part of this area are shown in Figure 7B. Figures 7B,C and D are zoomed in at random to show the typical morphology of this particular sample. Figure 7D has not been high pass filtered in order to show the appearance of the raw data on the AFM screen. These pictures
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Figure 9: Demonstrating changes owing to AFM tip contamination. Both images show the same fragments of A. DNA but B is broadened considerably after a particle is picked up by the scanning tip.
show blobs of200-400 nm in diameter, many of which are interconnected by DNA strands. Figures 7B,7C,7D clearly show that the blobs consist of highly localized, tightly coiled DNA strands which can originate at various points in a linear DNA molecule. The images are essentially indistinguishable after 30 minutes of continuous scanning (not shown), which suggests that the condensed regions are bound strongly to the modified mica surface. The scans shown in Figures 7A-7D are obtained in air. When these or similar scans are made under water, only condensed regions are present and no interconnections are observed (not shown) suggesting that the number of attachment points for isolated strands is often inadequate to stabilize them under water in these more weakly treated substrates. Under conditions of extreme aggregation on the scanning surface, some condensed molecules assume a toroidal conformation. Images of such toroids are shown in Figures 8A-8C. The center hole ofthis toroid in Figure 8D is -100 nm, which is close to the persistence length ofDNA in solution (28,32-41 ). Analogous toroid-like structures are formed by dsRNA as well (26). Thus, the binding DNA to AP-mica may be accompanied by self condensation of DNA Condensation of DNA is of fundamental importance, because the DNA in viral capsids, bacterial nucleoids, and chromosomes of higher organisms occupies 104 10 times less volume than it does when free in solution. A considerable amount of experimental and theoretical effort during past decades has been directed at understanding the physical mechanisms underlying condensation transitions in DNA (literature is abundant, see e.g. 33-41). Scheiman and co-workers (34,35) were the first to demonstrate that multivalent cations induces the condensation of DNA in vitro. Later electron microscopy (37 -41) studies permitted visualization of DNA aggregates
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as beaded fibers, sheroids, toroids, and rods. Our studies showthat the interaction of double helical nucleic acids with a surface carrying positively charged amino groups can be accompanied by folding. Because DNA and RNA folding morphologies in nuclear and viral environments are not yet well understood, and the role of surface structure on this process has not been studied before, the study of condensation phenomenon on surfaces analogous to AP-mica might shed a new light on the problem of DNA and RNA folding inside the cell.
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Tips Changes During Scanning Our discussion of resolution has, thus far, been limited to changes seen as a function of the tip geometry, medium and sample. However, it is important to realize that significant changes can occur during the course of an experiment, either because the tip becomes contaminated, or because its geometry is altered during scanning. This is illustrated in Figure 9 where we show two images obtained over the same patch of A.-Hind III fragments scanned in air. The images were taken a few minutes apart, with no deliberate changes in scanning conditions and no obvious tip-crash. A dramatic loss of resolution is evident. Note how the background assumes a characteristic motif, presumably owing to structure on the end of the tip. Clearly, it would be incorrect to interpret the periodic' structure' along the path of the molecule in Figure 9B as a feature of the DNA
Conclusions We have shown that APTES treated mica surfaces make a simple and reliable substrate for AFM studies of nucleic acids. They have the novel property of permitting AFM imaging underwater. The technique is reliable and easy enough that we expect the AFM will be used in certain routine assays. For example, length distributions are very easy to obtain. The data set for dsRNA measurements described in ref. 26 was obtained in about one hour. Limited resolution causes little loss of accuracy and is actually an advantage in visualizing very long polymers. With the development of suitable markers, we expect that the AFM will play a role in genomic mapping. The processes that determine resolution are not yet fully understood. Certainly, we find a trend which is in agreement with that reported by Hansma et al.: Both sharper tips and imaging under a liquid generally yield better resolution ( 17). Nonetheless, resolution cannot be determined simply by the macroscopic tip profile. It seems unlikely that control of the tip morphology at the micron level will have profound influence on nm surface features. Indeed, Bustamante et al. ( 15) argue that the relevant factor is the influence of the overall tip geometry on long range forces. These scale with the radius of the tip (42). Since these long range (generally attractive) forces must be balanced by the repulsive contact force even when the measured net contact force is zero, this can result in stress at the tip which is adequate to deform it substantially (43). The actual contactforce is the sum ofthe ofthe measured contactforce and the force needed to balance the long-range attractive interactions. Therefore, it is advantageous to minimize these interactions by using a macroscopically' sharp' tip.
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Table I Summarizing some of the AFM images of nucleic acids reported to date. Resolutions are either full widths measured from images or as reported by authors. Conventional tips refers to the standard pyramidal tips supplied by Digital Instruments Inc. or Park Scientific Inc. Notes: a) Humidity reported as affecting resolution. b) Sample electrically grounded. SUBSTRATE APTES-Mica APTES-Mica APTES-Mica Glow-discharged Mica Glow discharged Mica Carbon-Mica
MgC12-Mica Na-Acetate-Mica
SAMPLE A.-DNA ds-RNA A.-DNA plasmids plasmids plasmidscytochrome coated plasmids plasmids
REF
MEDIUM Air Air Water Humidity Controlled Air Propanol
TIP Conventional Conventional Conventional 'Super-tip'
RESOLUTION 20-40nm I0-20nm 10-25nm ~12nm•
13 26 14 15
'Super-tip'
3-8nm
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Air
Conventional
4-6nmh
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~18nm
20 19
Conventional Air Humidity Conventional Controlled Air
~15nm•
Likewise, imaging under a liquid eliminates capillary forces owing to liquid films on the surface of the sample (17). However, we have shown how the same molecule imaged with the same tip yields images whose width can change from nearly 80 to 20nm when the sample is covered with water (14). It is difficult to understand how deformation of the tip and substrate could be large enough to change the contact area by this much (provided that AP - mica in air is as rigid as non-modified one). Indeed, we have recently studied very fine structure on the gold surface, concluding that the contact area almost certainly involved more than a single atom, but was not much more than lnm (43). The intrinsic floppiness of the molecule will play a role, but since DNA is only 2nm in width, it is difficult to understand how this can make much contribution to the width of the observed images. Another factor is the strength of the binding ofthe molecule to the substrate. Here, the tip geometry plays a role, but only indirectly, in as much as it influences contact force. We have tried super-sharp tips (a gift from V. Elings of Digital Instruments) finding that they yield improved resolution, but only by a factor of 2 or 3, much less than the nominallOfold improvement in tip radius. We summarize our results (and those of others) in Table I where we list the various approaches that have been used to tether nucleic acids, the tips used, the medium and the reported resolution. We report a the typical range of image widths seen in a given set of our experiments in order to emphasize the fact that resolution can vary considerably even for a given preparation (perhaps a consequence of uncontrolled contamination). It can be seen that imaging under water does result in a significant improvement The work with 'supertips' (15-17) demonstrates that they give superior resolution. Indeed, Hansma eta/. (17) have, on occasion, obtained images whose width approaches the intrinsic width of the molecule). The results ofYang eta/. ( 18) stand out as rather remarkable. They used conventional tips and imaged in air. Furthermore, they
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imaged a cytochrome-c coated preparation of the sort commonly used in electron microscopy. Their images show a 6nm resolution with remarkable repeatability! Since this is close to the intrinsic width of the DNA-cytochrome complex, it may be that their images show almost no tip broadening. They report that this resolution is obtained with almost all the cantilevers used. Interestingly, they also report that the mica had to be electrically grounded and that the replacement of formamide with ammonium acetate caused resolution to be lost. The former factor might have increased tip-substrate interactions, while the latter might have improved binding to the substrate. Reliable imaging of nucleic acids by AFM is now possible on an almost routine basis. However, as the results presented here demonstrate, there is still room for improvement and a demonstratable need for further understanding of the factors that limit resolution. Acknowledgements
We are grateful to Dr. S. Kazakov (Yale University) for helpful discussions about mica chemistry; Y. Li (Arizona State University) for help in AFM experiments; U. Knipping (Arizona State University) for the help in image processing. Virgil Elings (of Digital Instruments Inc) who kindly gave us some 'super-tips' to evaluate. Financial support: research grants from NIH (REH), NSF (DIR8920053) and ONR (N000149Jl455) (SML), Hatch support from the NAES (REH), grant CA48654 from the US Public Health Service (BJL). References and Footnotes
1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.
Binnig, G, Rohrer, H., Gerber, Ch. and Weibel, E., Phys. Rev. Lett. 49, 57-61 (1982). Binnig, G. and Rohrer, H., Rev. Modem Phys. 59,615-626 (1987). Binnig, G., Quate, C. F. and Gerber, Ch., Phys. Rev. Lett. 56, 930-933 (1986). C1emer, C.R. and Beebe, T.P., Jr., Science 251, 640-642 (1991). Lindsay, S.M., In Scanning Tunneling Microscopy: Theory, Techniques and Applications, (J. Bonnell, ed) VCH Press (in press). Keller, D., Applied Physics, in press (1992). Engel, A, Ann. Rev. Biophys. Biophys. Chern. 20,79-108 (1991). Lindsay, S.M., Tao, NJ., DeRose, J.A, Oden, P.l., Lyubchenko, Y.L., Sh1yakhtenko, L.S and Harrington, R.E., Biophys. J. 61, 1570-1584 (1992). Tao, N.J., DeRose, J.A and Lindsay, S.M. Submitted toJ. Chern. Phys. (1992). Cricenti, A Se1ci, S, Felici, A C., Generosi, R., Gori, E., Djaczenko, W. and Chiarotti, G., Science 245, 1226-1227 (1989). Lyubchenko, Y.L., Lindsay, S.M., DeRose, J.A and Thundat, T., J. Vac. Sci. Techno/. B9, 12881290 (1991). Lyubchenko, Y.L., Lindsay, S.M., Gall, AA, Sh1yakhtenko, L.S., R.E. Harrington, Biophys. J. 61, A149 (1991). Lindsay, S.M., Lyubchenko, Y.L., Gall, AA, Sh1yakhtenko, L.S., R.E. Harrington, SPIE Proc. Intemat. Symp. Laser Spectr. 1639, 84-90 (1992). Lyubchenko, Y.L., Shlyakhtenko, L.S., Harrington, R.E., Oden, P.l and Lindsay, S.M., Proc. Nat/. Acad. Sci. USA, submitted. Bustamante, C. Vesenka, J., Tang, C.L., Rees, W., Gutfo1d, M. and Keller, R., Biochemistry 31, 22-26 (1992). Vesenka, J. Tang, C.L., Gutford, M., Keller, D. and Bustamante, C., Ultramicroscopy 42-44 12431249 (1992).
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17. Hansma, H.G., Vesenka, J., Siegerist, C., Kelderman, G., Morett, H., Sinsheimer, R.L., Eilings, V., Bustamante, C. and Hansma, P.K, Science 256, 1180-1184 (1992). 18. Yang, J., Takeyasu, K. and Shao, Z., FEES Letters 301, 173-176 (1992). 19. Thundat, T., Warmack, R.J., Allison, D.P., Lourenco, AJ., Ferrell, T.L. and Bottomly, L.A.,.!. Vac. Sci. Techno/., in press (1992). 20. Zenhausern, F., Adrian, M., Heggeler-Bordier, B.T., Emch, R., Jobin, M., Taborelli, M. and Descouts, P.,J. Struct. Bioi. 108, 69-73 (1992). 21. Lindsay, S.M., Nagahara, L.A, Thundat, T., Knipping, U., Rill, R.L., Drake, B., Prater,C.B., Weisenborn, AL., Gould, SAC. and Hansma, P.K, J. Biomol. Struct. Dyn. 7, 279-287 ( 1989). 22. Weisenborn, AL., Gaub, H.E., Hansma, H.G., Sinsheimer, R.L., Kelderman, G.L. and Hansma, P.K, Scanning Microsc. 4, 511-516 (1990). 23. Ramig, R.F., Cross, R.K. and Fields, B.N.,J. Viral. 22,726-733 (1977). 24. Langland, J.O., Pettiford, S.M. and Jacobs, B.L. (Submitted). 25. Oden, P.l., Nagahara, L.A, Graham, J.J., Jin Pan, Tao, N.J., Li, Y., Thundat, T., DeRose, J.A and Lindsay, S.M., Ultramicroscopy 42-44, 580-586 (1992). 26. Lyubchenko, Y.L., Jacobs, B.L. and Lindsay, S.M., Nucl. Acids Res 20, 3983-3986 (1992). 27. In: Fundamental Virology, Raven Press,New York (1986). 28. Livshits, M.A, Amosova, O.A and Lyubchenko, Y.L., J. Biomol. Sruct. Dyn. 7, 1237-1249 (1990). 29. Burnham, N.A. and Colton, R.T. In Scanning Tunneling Microscopy and Spectroscopy: Theory, Techniques and Applications, Editor Bonne!, D. , VCH Press, New York, in press ( 1992). 30. Person, B.N.J, Chern. Phys. Lett. 141,366-368 (1992). 31. Tao, N.J., Lindsay, S.M. and Lees, S., Biophys. J. (in press). 32. Hagerman, P .J., Annu. Rev. Biophys. Biophys. Chern. 17, 265-286 (1988). 33. Reich, Z., Ghirlando, R., Minsky, A, Biochemistry 30, 7828 (1991 ). 34. Gosule, L.C. and Schellman, J.A, Nature 259, 333-335 (1976). 35. Chattoraj, D.K, Gosule, L.C. and Schellman, J.A, J. Mol. Bioi. 121, 327-337 (1979). 36. Evdokimov, Y.M., Pyatigorskaya, T.L, Polyvtsev, 0., Akimenko, N.M., Kadykov, VA and Varshavsky, Ya.M., Nucl. Acids Res. 3, 2353 ( 1976). 37. Wilson, R.W. and Bloomfield, VA, Biochemistry 18,2192-2196 (1979). 38. Lerman, L.S., Wilkerson, L.S., Venable, J.H., Jr., and Robinson, B.H., J. Mol. Bioi. 108, 271-293 (1976). 39. Widom, J. and Baldwin, R.L., J. Mol. Bioi. 144, 431-453 ( 1980). 40. Arscott, P.l., Li, A-H., and Bloomfield, V.A, Biopolymers 30, 619-630 (1990). 41. Plum, G.E., Arscott, P.l. and Bloomfield, V.A, Biopolymers 30, 631-643 (1990). 42. Israelachvilli, J.N., Intermolecular and Surface Forces, Academic Press, New York, (1985). 43. Oden, P.l., Tao, N.J. and Lindsay, S.M., submitted. Date Received: May 3, 1992
Communicated by the Editor E. Trifonov
Journal of Biomolecular Structure and Dynamics Publication details, including instructions for authors and subscription information: http://www.tandfonline.com/loi/tbsd20
Atomic Force Microscopy Imaging of Double Stranded DNA and RNA Yuri L. Lyubchenko
a b c
d
, Alexander A. Gall , Lyuda
c
c
S. Shlyakhtenko , Rodney E. Harrington , Bertram b
a
L. Jacobs , Patrick I. Oden & Stuart M. Lindsay a
a
Departments of Physics
b
Departments of Microbiology , Arizona State University , Tempe , AZ , 85287 , USA c
Department of Biochemistry , University of Nevada Reno , Reno , NV , 89557 , USA d
MicroProbe Corp , 1725 St. SE#104, Bothell , WA , 98021 , USA Published online: 21 May 2012.
To cite this article: Yuri L. Lyubchenko , Alexander A. Gall , Lyuda S. Shlyakhtenko , Rodney E. Harrington , Bertram L. Jacobs , Patrick I. Oden & Stuart M. Lindsay (1992) Atomic Force Microscopy Imaging of Double Stranded DNA and RNA, Journal of Biomolecular Structure and Dynamics, 10:3, 589-606, DOI: 10.1080/07391102.1992.10508670 To link to this article: http://dx.doi.org/10.1080/07391102.1992.10508670
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Journal of Biomolecular Structure & Dynamics, ISSN 0739-1102 Volume 10. Issue Number 3 (1992), ®Adenine Press (1992).
Atomic Force Microscopy Imaging of Double Stranded DNA and RNA Yuri L. Lyubchenko*1•2•3 , Alexander A. Ga114 , Lyuda S. Sh~akhtenko 3 Rodney E. Harrington 3, Bertram L. Jacobs , Patrick I. Oden1 and Stuart M. Lindsay1 Downloaded by [Purdue University] at 13:15 13 April 2015
1
Departments of Physics and 2Microbiology Arizona State University Tempe, AZ 85287, USA 3
Department of Biochemistry University of Nevada Reno Reno NV 89557, USA ~icroProbe Corp 1725 St. SE# 104 Bothell, WA 98021, USA
Abstract A procedure for imaging long DNA and double stranded RNA (dsRNA) molecules using Atomic Force Microscopy (AFM) is described. Stable binding of double stranded DNA molecules to the flat mica surface is achieved by chemical modification of freshly cleaved mica under mild conditions with 3-aminopropyltriethoxy silane. We have obtained striking images of intact lambda DNA, Hind III restriction fragments oflambda DNA and dsRNA from reovirus. These images are stable under repeated scanning and measured contour lengths are accurate to within a few percent. This procedure leads to strong DNA attachment, allowing imaging underwater. The widths of the DNA images lie in the range of20 to 80nm for data obtained in air with commercially available probes. The work demonstrates that AFM is now a routine tool for simple measurements such as a length distribution. Improvement of substrate and sample preparation methods are needed to achieve yet higher resolution.
Introduction Scanning Probe Microscopy, which mostly includes Scanning Tunneling Microscopy (STM) and Atomic Force Microscopy (AFM), is a relatively new tool which *Corresponding author at the Department of Physics, Arizona State University, Tempe, AZ 852871504, USA Lyubchenko, Y.L. and Shlyakhtenko, L.S. are on leave from the Institute ofMolecular Genetics, Russian Academy of Sciences, Moscow, Russia.
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has enormous potential importance to structural biology. The STM and AFM offer unique advantages in their potential for very high resolution of DNA. RNA and their complexes with proteins and small ligands in the absence of stains, shadows and labels. Furthermore, the instruments can be operated in air or liquids. The latter is particularly important for resolving fully hydrated structures. They are theoretically capable of resolving structural details at the level of atomic dimensions, provided the specimen is dynamically stable. The prototype STM instrument was conceived by Binnig, Rohrer and coworkers (1) an invention for which Binnig and Rohrer were awarded the 1986 Nobel Prize in Physics (2). The first AFM was a direct descendent of this early instrument (3), but capable of imaging nonconducting as well as conducting surfaces. Although these instruments are, in principle, capable of imaging biological molecules and ultra structures to nearly atomic resolution and, hence, have great potential in such applications as micro-structural studies of DNA. RNA and other biological macromolecules, this potential has, so far, not been realized. For example, artifactual images may have been a problem in certain STM studies of macromolecules adsorbed on graphite surface (4,5), and the underlying theory of image contrast is not well understood. This appears to be a more serious problem with the STM, although the problem is not trivial with the AFM. However, an immediate practical limitation of the application of AFM and STM to structural studies of biological macromolecules is sample preparation. The macromolecules must be tethered to the substrate surface in order to avoid resolution-limiting motion occasioned by the sweeping tip during scanning (6). A number of procedures for sample preparation have been described, but, to our knowledge, no routine and reliable methods have been reported until recently. Most STM images of DNA and RNA published recentlywere obtained bydryingoutofDNAsolutions of a fewmg/ml onto scanning substrates. DNA aggregates are often stable, but do not allow one to image individual molecule (see reviews 5,7 and references therein). Lindsay et al. (5) have developed the use of electrochemical methods for holding DNA in place on a gold electrode as it is imaged with an STM. The DNA is held onto to a positively charged electrode and imaged under potential control (8). This procedure yields high resolution (better than can be obtained with the AFM at present) and the images are not subject to the artifacts that have plagued graphite surfaces (4). It has even proved possible to obtain images of single bases in based stacked aggregates of purines (9). However, the adsorption process is highly length selective, favoring small molecules. Furthermore, it offers no possibility of imaging large insulating clusters (such as DNA-protein complexes). Another approach to tethering DNA is chemical modification of the substrate. The first attempt was made by Cricenti et al.. (10). We have recently suggested a method for covalently tethering DNA to a graphite substrate (11). However, the graphite surface is not a suitable medium for STM studies ofbiopolymers (in our view) because of the problem of artifacts (4,5). The AFM permits insulating materials to be used as substrates, and freshly cleaved mica produces a much better surface than most graphite samples. We therefore set out to devise a procedure for chemical modification
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ofthe mica surface for studies of DNA (and other biopolymers) (12,13,14). The procedure we have developed is simple and robust and even works well underwater(l4) Here, we will describe some of its strengths and weaknesses. Over the same period of time, other workers have devised alternative strategies for reliable tethering of DNA for AFM studies (15-20). It is of interest to compare the published results to date. The first AFM study of DNA exploited the adhesion of DNA to wet glass (21 ). Subsequent work used mica surfaces treated with AIC13 (22). Resolution was limited by the tendency of the AFM tip to push DNA around (6) so better "tethering" of the molecule to the substrate is required. An ionic treatment of mica for binding DNA has been described by Bustamante et al.. (15) and Vesenka eta/.. (16). This permits reliable imaging in an AFM with a resolution approaching tens of nm. Hansma et a/. ( 17) have used this approach to obtain a significant improvement in resolution by imaging under a covering layer of propanol. Yang eta/. (18) used a conventional cytochrome-c modification of DNA to hold the molecules onto a carbon film that had been deposited on mica. Thundat eta/. (19) and Zenhausem eta/. (20) have reported that reliable results were obtained simply by applying the DNA to mica from an ammonium acetate (19) or MgC12 (20) buffer. Bustamante et al. (15) report that specially manufactured super sharp tips had to be used for reliable imaging. These tips have also been used by Hansma et al. (17). The other reports (12-14,19,20) were based on direct use of commercial probes with no special modification. The super-sharp tips do give better resolution in most cases, and resolution is also improved by imaging under propanol (17) or water (14). However, comparison of the published images suggest that other factors are important also. In this paper, we describe our procedure for imaging long DNA molecules chemically bound to mica. We give examples of a number of DNA and RNA images, images obtained in water, images of complex self-assembeled aggregates of DNA, and also some illustrations of effects caused by changes in the tip during scanning. We end with a discussion of our results (and those of the other groups) in order to clarifY the strengths and weaknesses of this new technique in its present state of development Materials and Methods Mica Modification Procedure Modification in liquid
We used freshly cleaved strips of ruby mica. Pieces of mica were incubated in freshly prepared solutions of 3-aminopropyltriethoxy silane (APTES, Aldrich, USA) in toluene or dimethylformamide for several hours with mild shaking followed by rinsing with pure solvent and drying. Dried strips were immersed into methyl iodide (Aldrich, USA) for one hour and dried again. Modification In Vapors
Freshly cleaved strips of mica were left in the APTES atmosphere created in a 21
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1--o-y1(CH2 ) 3 NH 2 OEt
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A Figure 1: (A) The steps in modification ofmica substrate-s with 3-aminopropyltriethoxy silane (APTES). Me is the methyl group and Et is the ethyl group. The silicon oxide group on the left is part of the mica surface. (B) Fraction of total DNA bound against exposure time. (II) data for unmodified mica, (.6.) mica treated with APTES alone and(*) mica methylated after APTES treatment. The standard deviation of data is the symbol size unless otherwise indicated.
glass desiccator containing a small pool of APTES under ambient conditions for 2 hours. The methylation procedure was the same as above.
DNA Samples Preparation Modified mica strips were immersed into DNA or RNA in Tris-HCl buffer solution
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(pH 7), containing lOmM Tris-HCl, 10-20 mM NaCl, 5 mM EDTA and incubated at room temperature for a specified time which varied usually between 1 and 2 hours. The concentration of DNA was varied in different experiments between 0.1 and O.Ql flg/ml for different samples. The lowest concentration, 0.01 flg/ml, was used in experiments with lambda DNA The concentration ofHind m fragments oflambda DNA was around 0.1 flg/ml. Both DNA samples were purchased from New England BioLabs, USA and used without additional purification. After the adsorption stage had been completed, the samples were rinsed with deionized water (NanoPure water system, Barnstead, 2555 Kerper Bvd. Dubuque, lA), blotted at the edge and vacuum dried.
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dsRNA Samples
Reovirus genome dsRNA was isolated from purified virions by a standard technique (23). Briefly, the virus was disrupted by extraction with phenol and nucleic acids were concentrated by precipitation with ethanol. Genomic dsRNA was purified by gel filtration (24).
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Radiolabeled Assay
Mica substrates were incubated for different times in a solution of 32P-labeled Hpall restriction fragments ofPBR322 plasmid DNA (New England Biolabs) at a concentration of0.2 ng/ml in 10 mM Tris-HCl buffer (pH7.5). Ratios of bound to total DNA were calculated from the radioactivity of the rinsed mica plates and of the DNA solution before and after the binding procedure. Apparatus
Imaging was carried out on a N anoScope II STM/AFM from Digital Instruments (6780 Cortona Drive, Santa Barbara, CA) using commercial AFM cantilevers, also from Digital Instruments (spring constant 0.58 and 0.12N/m). The radius of curvature of the end of these tips has been estimated directly from images of whisker crystals (25) and it lies between 20 and 50 nm. The liquid cell used for AFM in this work was an electrochemistry cell fabricated by Digital Instruments, Inc. Results and Discussion Basis of the Mica Modification Procedure
The treatment of the mica substrate is shown schematically in Figure IA The amino groups of APTES are bound covalently to the freshly cleaved mica surface, giving it similar properties to an anion exchanger used in affinity chromatography. Its anionic affinity is enhanced by the substitution of methyl groups for the amino protons using methyl iodide (Step 2). On contact with water (Step 3), the ethoxy groups of the covalently bound APTES moieties are hydrolyzed to OH. We have obtained good AFM images using both the methylated surface (step 2) and the aminopropyl surface (step 2 omitted). Binding of DNA to Modified Mica
We checked the modification procedure with 32P-labeled Hpall restriction fragments of pBR322 ~lasmid DNA Mica substrates, treated with APTES, were incubated in a solution of 2P- DNA for a given period, washed thoroughly and the radioactivity of both mica samples and the DNA solutions was determined. These data yield the fraction of total DNA molecules initially in solution that became bound by the surface. Results comparing non-treated mica, mica modified using APTES only (step I on Figure lA) and mica modified by APTES followed by methylation (step 2) are shown in Figure 1B. DNA binding to the unmodified mica surface is insignificant
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Figure 2: AFM images of mica substrates modified with different concentrations of APTES. Immediately after cleaving, mica plates were treated as described in Figure 1. The surfaces were immersed in toluene solutions offresh APTES at different concentrations, incubated overnight, rinsed with pure toluene and dried. (A) 0.1 M APTES; (B) 0.1 mM APTES; (C) O.l!!M APTES. Bar size is 1 11m.
(radioactivity of the samples is very close to the background). Mter 4 hours of incubation, 0.25% of total DNA molecules were bound to the surface modified with APTES alone. This fraction was increased by an order of magnitude when the surface was methylated. We have investigated the nature of the forces holding DNA on the surface by incubation of DNNmica samples in highly concentrated salt solutions. Mica samples incubated for4 hours in 0.2M NaCl solution resulted in desorption of 50% of DNA molecules. In 2M NaCl, only 20% of DNA remained on the surface. These results suggest that (relatively nonspecific) electrostatic interaction is a predominant force holding DNA molecules onto the aminopropyl mica surface. Suiface Structure ofAP-Mica
AFM images of a mica surface modified with APTES at different concentrations (AP mica) are shown in Figure 2. At high APTES concentration (0.1 M) the surface is extensively and somewhat irregularly modified (Figure 2A) but the irregularities are reduced with decreasing APTES concentration (Figure 2B) so that at 0.1 f.lM the surface is uniform to within the resolution of the AFM images (Figure 2C). The dark bands that cross the image are instrumental artifacts which occur at large scan amplitudes. They do not rotate as the sample is rotated relative to the scan head and a series of small range scans taken over the same area verifies that the surface is flat. These dark bands vary with alignment of the AFM optics, and we believe that they are caused by interference between the reflected beam and laser light scattered from objects on the substrate. Imaging of DNA as shown below utilized mica surfaces treated with fresh preparations of APTES at concentrations that did not exceed 0.1 flM. Imaging of Lambda DNA Restriction Fragments
Here we illustrate the procedure with long linear fragments of DNA Treated substrates were incubated with a solution of Hind III restriction fragments oflambda DNA, rinsed with water, dried and scanned with the AFM in air to yield the images shown in Figure 3. Typical results for this sample are shown in Figure 3A The background is uneven, owing to the interference fringes described above. The most
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Figure 3: (A) AFM image of Hind III restriction fragments of A. phage DNA; raw data showing background fluctuations. Bars are 2 11m markers. (B) shows the same area after high pass filtering to remove background variations. (C) shows the same area as in (B) after 30 min. of continuous scanning at contact forces between 40 and 90 nN. (D) images of well separated fragments including the 23.1 kb restriction fragment (marked with an arrow).
noticeable features are patches that appear to be due to aggregated DNA However, fine filaments can be seen crossing the substrate between the aggregates. These individual strands generally can be followed easily and measured strand lengths correspond well with known restriction fragments. The height of the strands is -2 nm (this can vary considerably), but their width is always much greater. We find that the width ofimages depends upon the tip used in the AFM, as might be expected (6). For a spherical tip (radius R) scanning a cylindrical molecule, it is straightforward to show (13) that the width of the resulting image is 4-j(Rr). For a parabolic tip, the result(15) is [4(R+r).J(r(R-r)]IR. By taking traces across images such as that shown in Figure 3A, we find that the full-width of the DNA imaged in air varies from 20 to 50nm (and sometimes, even broader). This is consistent with the range oftip radii
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Figure 4: AFM images of whole lambda DNA (B), (C) and (D) are high pass filtered. (A) is raw data. The convoluted molecule in (A) starts at the point labeled "1" and finishes at the point labeled "2". A more extended molecule is shown in (B) and (C) (the path of the molecule is pointed to by arrows). Two adjacent images were taken and the overlap is indicated by the common point labeled "X". (C) is a zoomin on the end of the molecule (shown in a box labeled "D" on (C).
determined experimentally. Bustamante eta/. (15) and Hansmaetal. (17) have also demonstrated that the radius of the AFM tip influences the resolution of isolated molecules. However, our discussion will show that other factors contribute to this apparent broadening effect. The instrumental scanning artifacts observed in Figure 2C correspond to several nm of topography and limit the contrast with which individual strands can be displayed. Figure 3B shows the same scan as in Figure 3A after it has been high-pass filtered to remove background variations. Individual strands can be followed easily. Figure 3C is an image of the same area after continuous scanning for 30 minutes at
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Figure 5: AFM images of dsRNA from reovirus. (A) 3.5 f.im X 3.5 J.lm scanning area and (B) 1.75 J.lm X 1.75 J.1ffi scanning area.
contact forces between 40 and 90 nN. Figures 3B and 3C are virtually identical, indicating the stability of DNA binding to this substrate, even at this fairly large contact force. Figure 3D shows an area in which many well-separated fragments can be found. In cases where the molecular contours can be followed without ambiguity, their lengths correspond to the known Hind III restriction fragments oflambda DNA. In particular, note that the whole ofthe 23.1 kb fragment is seen clearly in this one scan (arrow, Figure 3D). Its contour length is 7.5 ± 0.3 !Jm which is close to the 8 !Jmexpected if the fragment maintains a B-form base stacking. (A-form stacking would result in a length of 6.6 !Jm.) Imaging of the Whole Lambda DNA
The complete Aphage genome of 48,502 bp (16.4 !Jm length, assuming B-form DNA) is shown in Figure 4A. The molecule follows a very convoluted path with many loops (starting at "1" and finishing at "2", Figure 4A). In order to obtain a contour length, we used the public domain software NIH Image (1.40) to track it pixel by pixel. The measured size was 16.4± l!Jm, in good agreement with the expected length. A quite different type of DNA topography was obtained on a different sample (from a different preparation) and is shown in Figure 4B. The molecule is much less convoluted (its path is marked by the vertical arrows). It starts at the left of the image and continues off the right hand side. It is followed by moving the scan to the right and reimaging as shown in Figure 4C. A feature on the surface has been marked with an "X" on both Figures 4B and 4C, so that the two images can be indexed. The molecule continues on to the right, where it ends in a fairly convoluted set of loops. The end (enclosed in the box labeled "D" on Figure 4C) is shown scanned at higher magnification in Figure 4D. Using all three of these images together with the contour-measurement tools in NIH Image, we estimate the contour
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Figure 6: AFM images oflambda DNA in air (A) and under water (B). Samples were scanned using the liquid cell with contact forces ~ 60nN for scanning in air and 10-12 nN for scanning under water.
length of the whole molecule as 15± 1.0 IJ.ID, also in satisfactory agreement with that expected for intact A. phage DNA. (The widths of these images lay between 20 and 40nm). The fact that images of these large DNA molecules are both stable and reproducible and lead to such high accuracy in contour length measurements makes AFM an attractive method for physical mapping applications. AFM Studies of dsRNA
We have investigated dsRNA sample extracted from reovirus (26). This virus is represented by a multisegment genome (27). Our goal in imaging these molecules was to demonstrate the application of this method for routine length measurements. These are more difficult to make by conventional means for RNA because of the lack of restriction fragments with which to calibrate gels. Images of dsRNA molecules are shown on Figures 5A and 5B. The substrate preparation procedure was identical to that used for DNA. Figure 5A shows the data for a scanning area of 3.51J.m X 3.51J.m. This image clearly demonstrates polydispersity of the RNA sample (corresponding to the various fragments of the genome). Zooming in on parts of such pictures permits more accurate length measurements as shown in Figure 5B (1.7 11m X l.71J.m scan). For example, the length of molecule 1 on Figure 5B is 0.85 11m and molecule 2 is 0.4l!J.ID which corresponds to 3.03 kb and 1.45 kb respectively (using 2.8A as a rise-to-residue distance for dsRNA (28)). Statistics were obtained for a large number of molecules and complete data will be published elsewhere (26). The results showed that RNA length varies between 0.3 and 1.5 IJ.ID, which is in a good agreement with the data obtained using other techniques (27). Thus, the procedure we have described makes AFM a routine technique for length measurements of DNA and RNA. DNA Imaging Under Water
Using APTES treated mica substrates, we have been able to image lambda DNA
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Figure 7: AFM images oflambda DNA (A) 10 llffi X 10 j.lm scanning area. (B) 4.5j.lm X 4.5j.lm scanning area. (C,D) 1.5 j.lm X 1.5 j.lm area. (A), (B) and (C) are high pass filtered, (D) is raw data.
underwater (14). Other work described to date has been carried out in air, or under a poor solvent like propanol, because the DNA was not stable under water (17). (Actually, the first AFM images obtained were taken on glass under water (21) but they were of poor quality and difficult to reproduce). We have published images of the same molecule taken both under water and in air elsewhere (14). Images obtained in air varied in full width from 20 to 80 nm. Images obtained in water varied from I 0 to 25 nm in full width. This result is similar to the narrowing observed by Hansma et al. when imaging under propanol (they also used custom-made super-sharp tips, the combination producing a resolution record that approached 3nm on occasion (17)). These observations imply that the resolution cannot be controlled by the tip radius alone (the same tip was used for imaging in air and under water). At very high resolution, the deformation of the tip and substrate may be important (14,29). However, given the elastic properties of silicon nitride (the tip material) and mica
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Figure 8: Images of DNA toroids (scale indicated by bars). All images are raw data.
(the substrate material), deformations on the order of tens of nm are simply not possible at contact forces of -nN (14,29-31). Other factor(s) must play a role in change of resolution between experiments in air and experiments in water. DNA and RNA Morphology on AP-Mica Swface
In addition, we have noted interesting effects which can alter the fidelity of AFM imaging in capturing solution DNA conformations using modified mica substrates as described here. They appear to result from uneven surface characteristics of the mica substrates, presumably due to non-uniform mica modification with APTES. Examples are shown in Figures 6A(scanned in air) and 6B (scanned underwater). Common features of the AFM images under water for this sample are blurred regions which indicate localized DNA mobility during scanning. However, the overall binding of the DNA is strong as the images are stable during repeated scanning, i.e. they retain their characteristic features and do not change their position. The molecules also do not disappear if allowed to remain under water for long periods of time. We believe that the images of Figures 6A and 6B are of DNA molecules that are bound strongly to the surface at certain places and relatively weakly at others. The weakly bound regions interact dynamically with the scanning tip producing the regions oflow resolution observed. This explanation is in accord with the experiments using radio labelled DNA to determine overall binding, which did not reveal the difference between this substrate and any other ones. Another phenomenon which can be observed on an under-modified surface is extensive DNA agregation. The aggregates observed on the AP-mica surface are of interest in their own right. Obvious examples of extensive condensation are shown in Figures 7A and zoomed images of this sample are shown in Figures 7B,7C and 7D. The substrates used for this sample preparation are found to bind the DNA -50-fold less efficiently as determined using radiolabeling methods. In Figure 7A, we show a surface which is heavily deposited with condensed DNA "blobs". The results of zooming-in and scanning over the upper part of this area are shown in Figure 7B. Figures 7B,C and D are zoomed in at random to show the typical morphology of this particular sample. Figure 7D has not been high pass filtered in order to show the appearance of the raw data on the AFM screen. These pictures
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Figure 9: Demonstrating changes owing to AFM tip contamination. Both images show the same fragments of A. DNA but B is broadened considerably after a particle is picked up by the scanning tip.
show blobs of200-400 nm in diameter, many of which are interconnected by DNA strands. Figures 7B,7C,7D clearly show that the blobs consist of highly localized, tightly coiled DNA strands which can originate at various points in a linear DNA molecule. The images are essentially indistinguishable after 30 minutes of continuous scanning (not shown), which suggests that the condensed regions are bound strongly to the modified mica surface. The scans shown in Figures 7A-7D are obtained in air. When these or similar scans are made under water, only condensed regions are present and no interconnections are observed (not shown) suggesting that the number of attachment points for isolated strands is often inadequate to stabilize them under water in these more weakly treated substrates. Under conditions of extreme aggregation on the scanning surface, some condensed molecules assume a toroidal conformation. Images of such toroids are shown in Figures 8A-8C. The center hole ofthis toroid in Figure 8D is -100 nm, which is close to the persistence length ofDNA in solution (28,32-41 ). Analogous toroid-like structures are formed by dsRNA as well (26). Thus, the binding DNA to AP-mica may be accompanied by self condensation of DNA Condensation of DNA is of fundamental importance, because the DNA in viral capsids, bacterial nucleoids, and chromosomes of higher organisms occupies 104 10 times less volume than it does when free in solution. A considerable amount of experimental and theoretical effort during past decades has been directed at understanding the physical mechanisms underlying condensation transitions in DNA (literature is abundant, see e.g. 33-41). Scheiman and co-workers (34,35) were the first to demonstrate that multivalent cations induces the condensation of DNA in vitro. Later electron microscopy (37 -41) studies permitted visualization of DNA aggregates
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as beaded fibers, sheroids, toroids, and rods. Our studies showthat the interaction of double helical nucleic acids with a surface carrying positively charged amino groups can be accompanied by folding. Because DNA and RNA folding morphologies in nuclear and viral environments are not yet well understood, and the role of surface structure on this process has not been studied before, the study of condensation phenomenon on surfaces analogous to AP-mica might shed a new light on the problem of DNA and RNA folding inside the cell.
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Tips Changes During Scanning Our discussion of resolution has, thus far, been limited to changes seen as a function of the tip geometry, medium and sample. However, it is important to realize that significant changes can occur during the course of an experiment, either because the tip becomes contaminated, or because its geometry is altered during scanning. This is illustrated in Figure 9 where we show two images obtained over the same patch of A.-Hind III fragments scanned in air. The images were taken a few minutes apart, with no deliberate changes in scanning conditions and no obvious tip-crash. A dramatic loss of resolution is evident. Note how the background assumes a characteristic motif, presumably owing to structure on the end of the tip. Clearly, it would be incorrect to interpret the periodic' structure' along the path of the molecule in Figure 9B as a feature of the DNA
Conclusions We have shown that APTES treated mica surfaces make a simple and reliable substrate for AFM studies of nucleic acids. They have the novel property of permitting AFM imaging underwater. The technique is reliable and easy enough that we expect the AFM will be used in certain routine assays. For example, length distributions are very easy to obtain. The data set for dsRNA measurements described in ref. 26 was obtained in about one hour. Limited resolution causes little loss of accuracy and is actually an advantage in visualizing very long polymers. With the development of suitable markers, we expect that the AFM will play a role in genomic mapping. The processes that determine resolution are not yet fully understood. Certainly, we find a trend which is in agreement with that reported by Hansma et al.: Both sharper tips and imaging under a liquid generally yield better resolution ( 17). Nonetheless, resolution cannot be determined simply by the macroscopic tip profile. It seems unlikely that control of the tip morphology at the micron level will have profound influence on nm surface features. Indeed, Bustamante et al. ( 15) argue that the relevant factor is the influence of the overall tip geometry on long range forces. These scale with the radius of the tip (42). Since these long range (generally attractive) forces must be balanced by the repulsive contact force even when the measured net contact force is zero, this can result in stress at the tip which is adequate to deform it substantially (43). The actual contactforce is the sum ofthe ofthe measured contactforce and the force needed to balance the long-range attractive interactions. Therefore, it is advantageous to minimize these interactions by using a macroscopically' sharp' tip.
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Table I Summarizing some of the AFM images of nucleic acids reported to date. Resolutions are either full widths measured from images or as reported by authors. Conventional tips refers to the standard pyramidal tips supplied by Digital Instruments Inc. or Park Scientific Inc. Notes: a) Humidity reported as affecting resolution. b) Sample electrically grounded. SUBSTRATE APTES-Mica APTES-Mica APTES-Mica Glow-discharged Mica Glow discharged Mica Carbon-Mica
MgC12-Mica Na-Acetate-Mica
SAMPLE A.-DNA ds-RNA A.-DNA plasmids plasmids plasmidscytochrome coated plasmids plasmids
REF
MEDIUM Air Air Water Humidity Controlled Air Propanol
TIP Conventional Conventional Conventional 'Super-tip'
RESOLUTION 20-40nm I0-20nm 10-25nm ~12nm•
13 26 14 15
'Super-tip'
3-8nm
17
Air
Conventional
4-6nmh
18
~18nm
20 19
Conventional Air Humidity Conventional Controlled Air
~15nm•
Likewise, imaging under a liquid eliminates capillary forces owing to liquid films on the surface of the sample (17). However, we have shown how the same molecule imaged with the same tip yields images whose width can change from nearly 80 to 20nm when the sample is covered with water (14). It is difficult to understand how deformation of the tip and substrate could be large enough to change the contact area by this much (provided that AP - mica in air is as rigid as non-modified one). Indeed, we have recently studied very fine structure on the gold surface, concluding that the contact area almost certainly involved more than a single atom, but was not much more than lnm (43). The intrinsic floppiness of the molecule will play a role, but since DNA is only 2nm in width, it is difficult to understand how this can make much contribution to the width of the observed images. Another factor is the strength of the binding ofthe molecule to the substrate. Here, the tip geometry plays a role, but only indirectly, in as much as it influences contact force. We have tried super-sharp tips (a gift from V. Elings of Digital Instruments) finding that they yield improved resolution, but only by a factor of 2 or 3, much less than the nominallOfold improvement in tip radius. We summarize our results (and those of others) in Table I where we list the various approaches that have been used to tether nucleic acids, the tips used, the medium and the reported resolution. We report a the typical range of image widths seen in a given set of our experiments in order to emphasize the fact that resolution can vary considerably even for a given preparation (perhaps a consequence of uncontrolled contamination). It can be seen that imaging under water does result in a significant improvement The work with 'supertips' (15-17) demonstrates that they give superior resolution. Indeed, Hansma eta/. (17) have, on occasion, obtained images whose width approaches the intrinsic width of the molecule). The results ofYang eta/. ( 18) stand out as rather remarkable. They used conventional tips and imaged in air. Furthermore, they
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imaged a cytochrome-c coated preparation of the sort commonly used in electron microscopy. Their images show a 6nm resolution with remarkable repeatability! Since this is close to the intrinsic width of the DNA-cytochrome complex, it may be that their images show almost no tip broadening. They report that this resolution is obtained with almost all the cantilevers used. Interestingly, they also report that the mica had to be electrically grounded and that the replacement of formamide with ammonium acetate caused resolution to be lost. The former factor might have increased tip-substrate interactions, while the latter might have improved binding to the substrate. Reliable imaging of nucleic acids by AFM is now possible on an almost routine basis. However, as the results presented here demonstrate, there is still room for improvement and a demonstratable need for further understanding of the factors that limit resolution. Acknowledgements
We are grateful to Dr. S. Kazakov (Yale University) for helpful discussions about mica chemistry; Y. Li (Arizona State University) for help in AFM experiments; U. Knipping (Arizona State University) for the help in image processing. Virgil Elings (of Digital Instruments Inc) who kindly gave us some 'super-tips' to evaluate. Financial support: research grants from NIH (REH), NSF (DIR8920053) and ONR (N000149Jl455) (SML), Hatch support from the NAES (REH), grant CA48654 from the US Public Health Service (BJL). References and Footnotes
1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.
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Communicated by the Editor E. Trifonov
