Superhelix dimensions of a 1868 base pair plasmid determined by scanning force microscopy in air and in aqueous solution
Superhelix dimensions of a 1868 base pair plasmid determined by scanning force microscopy in air and in aqueous solutionKarsten Rippe*, Norbert Mücke and Jörg Langowski
Deutsches Krebsforschungszentrum, Abteilung Biophysik der Makromoleküle, Im Neuenheimer Feld 280, D-69120 Heidelberg, Germany
Received January 22, 1997;Revised and Accepted March 7, 1997
ABSTRACT
We have used scanning force microscopy (SFM) to study the conformation of a 1868 base pair plasmid (p1868) in its open circular form and at a superhelical density of [sigma] = -0.034. The samples were deposited on a mica surface in the presence of MgCl2. DNA images were obtained both in air and in aqueous solutions, and the dimensions of the DNA superhelix were analysed. Evaluation of the whole plasmid yielded average superhelix dimensions of 27 +- 9 nm (outer superhelix diameter D), 107 +- 51 nm (superhelix pitch P), and 54 +- 8o (superhelix pitch angle [alpha]). We also analysed compact superhelical regions within the plasmid separately, and determined values of D = 9.2 +- 3.3 nm, P = 42 +- 13 nm and [alpha] = 63 +- 20o for samples scanned in air or rehydrated in water. These results indicate relatively large conformation changes between superhelical and more open regions of the plasmid. In addition to the analysis of the DNA superhelix dimensions, we have followed the deposition process of open circular p1868 to mica in real time. These experiments show that it is possible to image DNA samples by SFM without prior drying, and that the surface bound DNA molecules retain some ability to change their position on the surface.
INTRODUCTION
Negative supercoiling is an important feature of the DNA conformation of almost all prokaryotes and eukaryotes. Changes in the degree of supercoiling can have severe effects on the viability of a given organism because DNA supercoiling and various DNA associated processes are tightly interrelated [reviewed in (1 )]. For example, DNA supercoiling has been shown to be connected with DNA transcription in several aspects: (i) negative supercoiling can facilitate the melting of the DNA at the transcription start site and thereby stimulate transcriptional activity [e.g. (2 ,3 )], (ii) the transcription process itself leads to the generation of some negative supercoiling behind the polymerase and positive supercoiling in front of the polymerase [reviewed in (4 )] and (iii) DNA supercoiling changes the DNA tertiary structure and can thereby facilitate interactions over large distances between regulatory proteins bound at enhancer sites and the promoter as discussed in ref. (5 ). DNA supercoiling is of similar relevance to other cases like replication or recombination [reviewed in (1 )]. Accordingly, the characterization of the superhelical DNA conformation is important for the understanding of many DNA dependent biological processes.
Structural studies of superhelical DNA are constrained by the flexibility of the molecule. DNAs longer than ~50 nm (the persistence length) behave as flexible filaments, and their structure can only be characterized by average statistical properties. Furthermore, the conformation of the molecule becomes very sensitive to the environment, and processes such as drying and staining used in classical electron microscopy are prone to interfere with the structure determination. One way to circumvent this problem is to apply biophysical methods that characterize the structural properties of an ensemble of DNA molecules in solution, such as light scattering, ultracentrifugation or spectroscopic techniques. These methods are the least invasive way of studying the structure of large DNA molecules but they lack the possibility to observe structural details of single molecules.
Scanning force microscopy (SFM) (also termed atomic force microscopy or AFM) is an emerging technique that can be used for imaging DNA and other biomolecules without further fixing or staining, the only constraint being that the molecule is bound to a surface. SFM and its application for the study of biological macromolecules are described in several recent reviews (6 -9 ) to which the reader is referred for additional information.
Preparation of the DNA sample for SFM imaging can be done by using Mg2+ to bind the DNA to the negatively charged surface of freshly cleaved mica. This process is relatively gentle and slow, and it has been shown that the DNA re-equilibrates on the substrate under the conditions of deposition used here (10 ). No further treatment is required to enhance the contrast and scanning is possible both in air and in liquid. Thus, the native hydration state of the DNA can be preserved and artefacts due to drying of the sample can be avoided. A disadvantage of SFM is that upon binding to the surface, the three-dimensional conformation of the DNA is constrained to two dimensions, and information on the orientation of the molecule in the third dimension is lost. In this respect cryo-electron microscopy (cryo-EM) is advantageous as 3D images of DNA can be obtained from 3D reconstruction of stereo pair micrographs (11 ,12 ). Cryo-EM, however, also constrains the molecule into a 100 nm thin layer of ice, which is less than the extension of typical plasmid DNAs. In addition, it has been argued that the conformation of the DNA can change during the quick freezing process in cryo-EM (13 ). In summary, no single method can be relied on to give all necessary structural information about a flexible biopolymer, and SFM offers the possibility to gain complementary data on the conformation of single DNA molecules in near physiological conditions that are difficult to obtain by other methods.
MATERIALS AND METHODS
DNA plasmid p1868 is a derivative of pUC18 and was purified from E.coli HB 101 as described previously (35 ,36 ). Details on the construction of the plasmid and the purification procedure are given elsewhere (Hammermann,M. et al., manuscript submitted). Gel electrophoretic analysis showed that the superhelical fraction used here consisted of ~80% of topoisomers -6 corresponding to a superhelical density of [sigma] = -0.034 in 40 mM Tris-acetate, pH 8.0 and 1 mM EDTA. In addition ~10% of the material was present as topoisomer -7 and another 10% was in the open circular form. The SFM images were obtained with a Nanoscope III (Digital Instruments, Santa Barbara, CA) operating in the `tapping mode' following one of three different protocols:
Protocol I
DNA samples were prepared by deposition of 10 or 15 [mu]l of a 2-5 nM DNA solution in 10 mM HEPES-KOH, pH 8.0, 10 mM MgCl2 and30 mM NaCl onto a piece of freshly cleaved mica (Plano GmbH, Wetzlar, Germany). The mica disc was washed immediately by dropping distilled water onto the surface and then drying the sample in a nitrogen stream. Images were recorded in air at ambient humidity using etched Si-probes (type Nanosensors) purchased from L.O.T. Oriel (Darmstadt, Germany) with a force constant 17-64 N/m, a thickness of 3.5-5.0 [mu]m, a resonance frequency between 250 and 400 kHz and a tip curvature radius of ~10 nm (specifications given by the manufacturer).
Protocol II
The DNA samples were prepared as described in protocol I and the mica disc was mounted into the SFM liquid cell. Then the samples were rehydrated by injecting H2O supplemented with 0.01% NP-40 into the liquid cell. Although not required, the presence of 0.01% NP-40 was found to facilitate the imaging process; stable images were usually obtained after 5-10 min. A buffer without 0.01% NP-40 (5 mM HEPES-KOH, pH 8.0, 2 mM MgCl2) was also successfully used to rehydrate and image the samples (data not shown). Tapping in liquid was done with silicon cantilevers type ultralever from Park Scientific Instruments (Sunnyvale, CA) with a thickness of ~0.8 [mu]m, using the C tip which has a typical force constant of 1 N/m, a tip curvature radius of 10 nm and a resonance frequency in air of ~140 kHz (specifications given by the manufacturer). For scanning in liquid a vibration frequency of ~20 kHz was usually used.
Protocol III
According to protocol I, a sample with a 6.8 kb open circular plasmid at a low DNA concentration was prepared so that only a few molecules per 4 * 4 [mu]m scan were present. The sample was rehydrated in 5 mM HEPES-KOH, pH 8.0 buffer supplemented with 2 mM MgCl2, and imaging was done as described in protocol II. After stable images of the 6.8 kb plasmid had been obtained, a solution of the much smaller open circular p1868 plasmid at a concentration of 4 nM was injected into the SFM liquid cell. After a few minutes of re-equilibration, images were recorded every ~3 min to follow the deposition process of the plasmid and to detect any rearrangements of DNA molecules on the surface. This protocol avoided drying of the DNA. To determine the DNA double helix and the superhelix dimensions, 20 superhelical plasmids of images scanned in air (protocol I) and 20 molecules rehydrated in water (protocol II) were evaluated. Height, width and length measurements were made with the installed Nanoscope Software and with the program NIH image version 1.57. The dimensions of the superhelix were determined according to two strategies: the average dimensions of the superhelix were calculated according to the formalism described in ref. (37 ). In addition, direct SFM measurements were used to evaluate regions of the plasmid that showed a continuous plectonemic tract.
RESULTS
SFM images of the p1868 plasmid are displayed in Figures 1 , 2 and 3 . For the superhelical plasmids ([sigma] = -0.034) in Figure 1 C and D a 2 * 2 [mu]m overview is presented, and in Figure 2 C and D magnified images of single molecules are depicted. The open circular plasmids are shown in Figure 1 A and B (overview), Figure 2 A and B (magnification of single molecules), and in Figure 3 (time course demonstrating the binding to the surface). The images were made by using the three different protocols that are described under Materials and Methods.
DISCUSSION
We have studied the conformation of a 1868 bp long plasmid in its superhelical and in its open circular conformation. SFM images were recorded both in air and in aqueous solution and the DNA dimensions were analysed. The values determined here for the apparent width and height of the DNA double helix (Table 1 ) fall within the range obtained in other studies for DNA plasmids bound to a mica surface via Mg2+. For imaging in air apparent widths of 12.3 +- 3.3 nm (16 ), 21.9 +- 3.7 nm (18 ) and 11.2 +- 1.8 nm (39 ) have been determined, and the height was measured to be 1.32 +- 0.34 nm (16 ), 0.79 +- 0.05 nm (18 ), 0.43 +- 0.08 nm (39 ) and 0.54 +- 0.12 nm (40 ). The dimensions of the DNA double helix in aqueous solution were found to be 19 +- 4 nm with a height of 2.5 +- 0.5 nm (41 ).
The apparent width of the DNA double helix is largely dependent on the size of the tip and this parameter therefore is not very meaningful in terms of the real DNA dimensions (Fig. 4 A). The low height for DNA imaged in air, 0.44 +- 0.07 nm instead of 2.4 nm expected for B-DNA, could result from two effects: compression of the molecule by the tip during the scanning process and attractive capillary forces mediated by the thin water layer on the sample surface which result in an apparent reduction of the DNA height. Both factors are likely to be different in water, which would explain the observed increase in the measured DNA height from 0.44 +- 0.07 to 1.14 +- 0.16 nm.
Drying the DNA sample can potentially introduce a conformation change. It is known that at low relative humidity (60-75%) natural DNA sequences will undergo a transition from B-DNA into the A-form. A-DNA is characterized by a different helix geometry and in particular by a shorter axial rise of 0.28 nm/bp instead of 0.34 nm/bp in B-DNA, as determined by X-ray diffraction of DNA fibres (42 ). One would therefore expect a shortening of the DNA contour, detectable in the SFM images, if a transition from B- to A-DNA occurs. Under the conditions studied here we found a helical rise of 0.34 +- 0.01 nm/bp characteristic for B-form DNA with both dried and rehydrated open circular plasmids. This is in agreement with the results from previous studies which reported 0.34 +- 0.01 nm (17 ,20 ,43 ). In addition, it was observed with samples deposited from aqueous buffers to mica that the characteristic B-DNA spacing of 0.34 +- 0.03 nm/bp persisted even in propanol (41 ). These results demonstrate that the binding of the plasmid DNA molecules to the mica surface can prevent the expected transition to the A-form under some conditions.
On the other hand, several studies reported lower helical rise values which would suggest that at least a partial transition to the A-form occurs: a helical rise between 0.28 and 0.33 nm was observed by Bustamante and co-workers for three different plasmids (16 ). Gold-labelled linear DNA molecules measured in air at a relative humidity <10% showed a helical rise of 0.28 nm/bp (40 ), and a value of 0.30 +- 0.01 nm/bp was determined by Thundat et al. (18 ). For a set of eight linear DNA fragments from 350 to 5994 bp Rivetti et al. (10 ) measured a helical rise of 0.312 +- 0.005 nm/bp. However, the authors attributed this discrepancy to the value for B-form DNA to the smoothing procedure applied in their image analysis program and to limitations in the pixel resolution. In a recent SFM study, Hansma et al. (19 ) estimated a value of 0.25 nm/bp for short linear DNA fragments (50, 100 and 200 bp), indicative of an A-DNA conformation. They suggested that the ability of mica-bound DNA strands to undergo a B- to A-transition will depend on the strength of the binding interaction between DNA and mica, which is expected to be higher with large DNA fragments (19 ,41 ). Thus, the varying results of contour length measurements of dried samples in air could reflect differences in the size of the DNA samples and/or the deposition protocol used for binding the DNA to mica.
The average dimensions of the superhelix can be determined from the contour length Lc, the measured length of the superhelix l, the number of nodes n and the number of end loops E as described in ref. (37 ). The values obtained here by SFM (outer superhelix diameter D = 27 +- 9 nm, superhelix radius r = 12.5 +- 4.7 nm, superhelical pitch P = 107 +- 51nm and superhelix pitch angle [alpha] = 54 +- 8o, at [sigma] = -0.034, Table 2 ) can be compared with results of electron microscopy studies of superhelical plasmids (12 ,37 ,44 ). Boles et al. determined values of r = 9.6 nm, P = 99 nm and [alpha] = 59o for a 7 kb plasmid spread in TE buffer at [sigma] = -0.033 (37 ). In another study cryo-electron microscopy was used to analyse pUC18 (2686 bp) at a somewhat higher [sigma] of -0.047, and values of superhelix diameter D = 12 nm, P [approx] 55 nm and [alpha] = 55o were determined in TE buffer (44 ).
As judged from EM studies and from Brownian dynamics simulations ~75% of the linking number deficit [Delta]Lk = -6 is expected to be in the writhe component Wr and 25% in the change of twist [Delta]Tw (37 ,44 -46 ). Here we determined a writhe of -3.7 +- 1.6 (air) and -3.0 +- 1.1 (water) corresponding to a change of twist of -2.3 +- 1.6 (air) and -3.0 +- 1.1 (water), respectively (Table 2 ). This would suggest that the writhe contributes only 50-60% to [Delta]Lk. It appears unlikely that the slight differences with respect to the solution conditions used for the determination of [Delta]Lk on the gel (40 mM Tris-acetate, pH 8.0 and 1 mM EDTA) and the SFM sample deposition buffer (10 mM HEPES-KOH, pH 8.0, 10 mM MgCl2 and30 mM NaCl) would lead to the observed value of writhe (47 ). In addition, there is no evidence for a structural transition of the DNA induced by negative supercoiling from the gel electrophoretic analysis. Thus, the observed difference to the expected value of Wr are most likely due to the binding of the DNA to the surface, which might favour some redistribution of writhe and twist under the conditions used here. The somewhat less negative value of Wr determined in water might reflect an increased electrostatic repulsion in the rehydration solution (H2O + 0.01% NP-40) as compared with the deposition conditions (10 mM HEPES-KOH, pH 8.0, 10 mM MgCl2 and30 mM NaCl, ionic strength 70 mM), which could lead to a further decrease of [Delta]Tw. This would be in agreement with the observed dependence of the duplex winding on the ionic environment of the DNA (47 ). That some conformation rearrangement of the mica bound plasmids is possible in liquid can be seen with the DNA samples shown in Figure 3 .
The above values are derived from an analysis that assumes an idealized regular conformation of the superhelix. However, in our studies we usually observe tightly interwound regions that alternate with more open regions of the plasmid, rather than a regular superhelix structure. Instead of averaging over the whole molecule, we therefore measured also the tightly interwound parts of these compact superhelical DNA regions separately, and obtained values of D = 9.2 +- 3.3 nm, r = 3.4 +- 1.7 nm, P = 42 +- 13 nm and [alpha] = 63 +- 20o (Table 3 ). These values are expected to be similar to the superhelix conformation that is formed throughout the whole plasmid molecules upon increasing the superhelical density in the presence of MgCl2. This view is supported by the results from electron microscopy studies. For example, for the above mentioned 7 kb plasmid the superhelix dimensions averaged over the whole plasmid at [sigma] = -0.059 in the presence of 10 mM MgCl2 were found to change to r = 5.8 nm, P = 52 nm and [alpha] = 55o (37 ). By cryo-EM it has been observed that very compact superhelix structures form at [sigma] = -0.06 in the presence of 100 mM NaCl or 10 mM MgCl2 (12 ,44 ). The interpretation of these findings as close contact of DNA double helices in the interwound region, however, was recently challenged by solution measurements (13 ).
The quality of the images obtained by injection of the DNA (Fig. 3 , protocol III) was not as good as with protocols I and II. This is likely to be related to the more difficult sample preparation and to the requirement to record successive images which leaves less possibilities to optimize the instrument settings. Nevertheless, the results show that one can image DNA molecules that have not been dried before, and therefore their native hydration state has been preserved throughout the preparation. This technique has been developed by Hansma and co-workers (31 ,33 ,34 ), albeit by using a somewhat different deposition protocol. In addition, we were able to follow the binding of DNA to the mica surface by SFM. Together with the observation that the DNA plasmid molecules bound to the surface have still retained some ability to change their position and their location on the surface, as has been observed previously (31 ,33 ), this reveals the exciting potential of SFM to follow dynamic processes in real time. This feature is going to be even more important for studies of proteins and their interaction with DNA. For it is known that drying of protein samples often leads to an irreversible loss of protein activity, and recent work in studies of E.coli RNA polymerase (32 ,48 ) demonstrates that the transcription process can be visualized by SFM.
ACKNOWLEDGEMENTS
We thank Katalin Tóth and Nathalie Brun for the p1868 plasmid samples. We are also grateful to Martin Guthold, Achim Schaper and Tom Jovin for their advice, to Helen Hansma for sharing preprints of their SFM work on DNA, to Alexandra Schulz for critical reading of the manuscript and to Ingrid Grummt for her support.
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