Anomalous Laterally Stressed Kinetically Trapped DNA Surface Conformations
Corresponding Author: Dmitry V. Klinov
Nano-Micro Letters,
Vol. 13 (2021), Article Number: 130
Abstract
Up to now, the DNA molecule adsorbed on a surface was believed to always preserve its native structure. This belief implies a negligible contribution of lateral surface forces during and after DNA adsorption although their impact has never been elucidated. High-resolution atomic force microscopy was used to observe that stiff DNA molecules kinetically trapped on monomolecular films comprising one-dimensional periodically charged lamellar templates as a single layer or as a sublayer are oversaturated by sharp discontinuous kinks and can also be locally melted and supercoiled. We argue that kink/anti-kink pairs are induced by an overcritical lateral bending stress (> 30 pNnm) inevitable for the highly anisotropic 1D-1D electrostatic interaction of DNA and underlying rows of positive surface charges. In addition, the unexpected kink-inducing mechanical instability in the shape of the template-directed DNA confined between the positively charged lamellar sides is observed indicating the strong impact of helicity. The previously reported anomalously low values of the persistence length of the surface-adsorbed DNA are explained by the impact of the surface-induced low-scale bending. The sites of the local melting and supercoiling are convincingly introduced as other lateral stress-induced structural DNA anomalies by establishing a link with DNA high-force mechanics. The results open up the study in the completely unexplored area of the principally anomalous kinetically trapped DNA surface conformations in which the DNA local mechanical response to the surface-induced spatially modulated lateral electrostatic stress is essentially nonlinear. The underlying rich and complex in-plane nonlinear physics acts at the nanoscale beyond the scope of applicability of the worm-like chain approximation.
Highlights:
1 DNA kinking is inevitable for the highly anisotropic 1D–1D electrostatic interaction with the one-dimensionally periodically charged surface.
2 The double helical structure of the DNA kinetically trapped on positively charged monomolecular films comprising the lamellar templates is strongly laterally stressed and extremely perturbed at the nanometer scale.
3 The DNA kinetic trapping is not a smooth 3D—> 2D conformational flattening but is a complex nonlinear in-plane mechanical response (bending, tensile and unzipping) driven by the physics beyond the scope of the applicability of the linear worm-like chain approximation.
Keywords
Download Citation
Endnote/Zotero/Mendeley (RIS)BibTeX
- J.P. Peters, L.J. Maher, DNA curvature and flexibility in vitro and in vivo. Q. Rev. Biophys. 43, 23–63 (2010). https://doi.org/10.1017/S0033583510000077
- A. Vologodskii, M.D. Frank-Kamenetskii, Strong bending of the DNA double helix. Nucleic Acids Res. 41, 6785–6792 (2013). https://doi.org/10.1093/nar/gkt396
- A.P. Fields, E.A. Meyer, A.E. Cohen, Euler buckling and nonlinear kinking of double-stranded DNA. Nucleic Acids Res. 41, 9881–9890 (2013). https://doi.org/10.1093/nar/gkt739
- J. Wang, H. Qu, G. Zocchi, Critical bending torque of DNA is a materials parameter independent of local base sequence. Phys. Rev. E 88, 032712 (2013). https://doi.org/10.1103/PhysRevE.88.032712
- C. Bustamante, Z. Bryant, S.B. Smith, Ten years of tension: single-molecule DNA mechanics. Nature 421, 423–427 (2003). https://doi.org/10.1038/nature01405
- P. Gross, N. Laurens, L.B. Oddershede, U. Bockelmann, E.J.G. Peterman et al., Quantifying how DNA stretches, melts and changes twist under tension. Nat. Phys. 7, 731–736 (2011). https://doi.org/10.1038/nphys2002
- J.F. Marko, S. Neukirch, Global force-torque phase diagram for the DNA double helix: Structural transitions, triple points, and collapsed plectonemes. Phys. Rev. E. 88, 062722 (2013). https://doi.org/10.1103/PhysRevE.88.062722
- J.F. Marko, DNA Mechanics. in Nuclear Architecture and Dynamics, eds. C. Lavelle, J.-M.Victor (Elsevier, 2018), pp. 3–40.
- F.H.C. Crick, A. Klug, Kinky helix. Nature 255, 530–533 (1975). https://doi.org/10.1038/255530a0
- Q. Du, C. Smith, N. Shiffeldrim, M. Vologodskaia, A. Vologodskii, Cyclization of short DNA fragments and bending fluctuations of the double helix. Proc. Natl. Acad. Sci. USA 102, 5397–5402 (2005). https://doi.org/10.1073/pnas.0500983102
- T.A. Lionberger, D. Demurtas, G. Witz, J. Dorier, T. Lillian et al., Cooperative kinking at distant sites in mechanically stressed DNA. Nucleic Acids Res. 39, 9820–9832 (2011). https://doi.org/10.1093/nar/gkr666
- F. Lankaš, R. Lavery, J.H. Maddocks, Kinking occurs during molecular dynamics simulations of small DNA minicircles. Structure 14, 1527–1534 (2006). https://doi.org/10.1016/j.str.2006.08.004
- J. Curuksu, M. Zacharias, R. Lavery, K. Zakrzewska, Local and global effects of strong DNA bending induced during molecular dynamics simulations. Nucleic Acids Res. 37, 3766–3773 (2009). https://doi.org/10.1093/nar/gkp234
- J.P.K. Doye, T.E. Ouldridge, A.A. Louis, F. Romano, P. Sulc et al., Coarse-graining DNA for simulations of DNA nanotechnology. Phys. Chem. Chem. Phys. 15, 20395–20414 (2013). https://doi.org/10.1039/C3CP53545B
- P. Cong, L. Dai, H. Chen, J.R.C. van der Maarel, P.S. Doyle et al., Revisiting the Anomalous Bending Elasticity of Sharply Bent DNA. Biophys. J. 109, 2338–2351 (2015). https://doi.org/10.1016/j.bpj.2015.10.016
- R.M. Harrison, F. Romano, T.E. Ouldridge, A.A. Louis, J.P.K. Doye, Identifying physical causes of apparent enhanced cyclization of short DNA molecules with a coarse-grained model. J. Chem. Theory Comput. 15, 4660–4672 (2019). https://doi.org/10.1021/acs.jctc.9b00112
- C. Rivetti, M. Guthold, C. Bustamante, Scanning force microscopy of DNA deposited onto mica: equilibration versus kinetic trapping studied by statistical polymer chain analysis. J. Mol. Biol. 264, 919–932 (1996). https://doi.org/10.1006/jmbi.1996.0687
- M. Sushko, A. Shluger, C. Rivetti, Simple model for DNA adsorption onto a mica surface in 1:1 and 2:1 electrolyte solutions. Langmuir 22, 7678–7688 (2006). https://doi.org/10.1021/la060356+
- P.A. Wiggins, T. van der Heijden, F. Moreno-Herrero, A. Spakowitz, R. Phillips et al., High flexibility of DNA on short length scales probed by atomic force microscopy. Nat. Nanotechnol. 1, 137–141 (2006). https://doi.org/10.1038/nnano.2006.63
- A.K. Mazur, M. Maaloum, Atomic force microscopy study of DNA flexibility on short length scales: smooth bending versus kinking. Nucleic Acids Res. 42, 14006–14012 (2014). https://doi.org/10.1093/nar/gku1192
- T. Brouns, H. De Keersmaecker, S.F. Konrad, N. Kodera, T. Ando et al., Free energy landscape and dynamics of supercoiled DNA by high-speed atomic force microscopy. ACS Nano 12, 11907–11916 (2018). https://doi.org/10.1021/acsnano.8b06994
- A. Podestà, M. Indrieri, D. Brogioli, G.S. Manning, P. Milani et al., Positively Charged Surfaces Increase the Flexibility of DNA. Biophys. J. 89, 2558–2563 (2005). https://doi.org/10.1529/biophysj.105.064667
- J.-H. Jeon, J. Adamcik, G. Dietler, R. Metzler, Supercoiling induces denaturation bubbles in circular DNA. Phys. Rev. Lett. 105, 208101 (2010). https://doi.org/10.1103/PhysRevLett.105.208101
- Y.L. Lyubchenko, L.S. Shlyakhtenko, T. Ando, Imaging of nucleic acids with atomic force microscopy. Methods 54, 274–283 (2011). https://doi.org/10.1016/j.ymeth.2011.02.001
- B. Akpinar, P.J. Haynes, N.A.W. Bell, K. Brunner, A.L.B. Pyne et al., PEGylated surfaces for the study of DNA–protein interactions by atomic force microscopy. Nanoscale 11, 20072–20080 (2019). https://doi.org/10.1039/C9NR07104K
- N. Severin, J. Barner, A.A. Kalachev, J.P. Rabe, Manipulation and overstretching of genes on solid substrates. Nano Lett. 4, 577–579 (2004). https://doi.org/10.1021/nl035147d
- J. Adamcik, D.V. Klinov, G. Witz, S.K. Sekatskii, G. Dietler, Observation of single-stranded DNA on mica and highly oriented pyrolytic graphite by atomic force microscopy. FEBS Lett. 580, 5671–5675 (2006). https://doi.org/10.1016/j.febslet.2006.09.017
- D. Klinov, B. Dwir, E. Kapon, N. Borovok, T. Molotsky et al., High-resolution atomic force microscopy of duplex and triplex DNA molecules. Nanotechnology 18, 225102 (2007). https://doi.org/10.1088/0957-4484/18/22/225102
- E.V. Dubrovin, M. Schächtele, T.E. Schäffer, Nanotemplate-directed DNA segmental thermal motion. RSC Adv. 6, 79584–79592 (2016). https://doi.org/10.1039/C6RA14383K
- Spontaneous large-angle bends in short DNA contour segments which are many times more prevalent than predicted by the canonical WLC model have been reported for the freely equilibrated DNA on mica using AFM imaging in air [19], however the normal bending angle distribution with no noticeable deflection from WLC model has been found in in-situ AFM measurements carried out in solution [20].
- V.V. Prokhorov, D.V. Klinov, A.A. Chinarev, A.B. Tuzikov, I.V. Gorokhova et al., High-resolution atomic force microscopy study of hexaglycylamide epitaxial structures on graphite. Langmuir 27, 5879–5890 (2011). https://doi.org/10.1021/la103051w
- R. García, R. Perez, Dynamic atomic force microscopy methods. Surf. Sci. Rep. 47, 197–301 (2002). https://doi.org/10.1016/S0167-5729(02)00077-8
- V.V. Prokhorov, S.A. Saunin, Probe-surface interaction mapping in amplitude modulation atomic force microscopy by integrating amplitude-distance and amplitude-frequency curves. Appl. Phys. Lett. 91, 023122 (2007). https://doi.org/10.1063/1.2756271
- A. Mikhaylov, S.K. Sekatskii, G. Dietler, D.N.A. Trace, A comprehensive software for polymer image processing. J. Adv. Microsc. Res. 8, 241–245 (2013). https://doi.org/10.1166/jamr.2013.1164
- M. Frank-Kamenetskii, How the double helix breathes. Nature 328, 17–18 (1987). https://doi.org/10.1038/328017a0
- J.P. Rabe, S. Buchholz, Commensurability and mobility in two-dimensional molecular patterns on graphite. Science 253, 424–427 (1991). https://doi.org/10.1126/science.253.5018.424
- V.V. Prokhorov, K. Nitta, The AFM observation of linear chain and crystalline conformations of ultrahigh molecular weight polyethylene molecules on mica and graphite. J. Polym. Sci. Part B Polym. Phys. 48, 766–777 (2010). https://doi.org/10.1002/polb.21944
- W. Hoyer, D. Cherny, V. Subramaniam, T.M. Jovin, Rapid self-assembly of α-synuclein observed by in situ atomic force microscopy. J. Mol. Biol. 340, 127–139 (2004). https://doi.org/10.1016/j.jmb.2004.04.051
- C. Whitehouse, J. Fang, A. Aggeli, M. Bell, R. Brydson et al., Adsorption and self-assembly of peptides on mica substrates. Angew. Chem. Int. Ed. 44, 1965–1968 (2005). https://doi.org/10.1002/anie.200462160
- D. Bagrov, Y. Gazizova, V. Podgorsky, I. Udovichenko, A. Danilkovich et al., Morphology and aggregation of RADA-16-I peptide studied by AFM, NMR and molecular dynamics simulations. Biopolymers 106, 72–81 (2016). https://doi.org/10.1002/bip.22755
- C. Charbonneau, J.M. Kleijn, M.A. Cohen Stuart, Subtle charge balance controls surface-nucleated self-assembly of designed biopolymers. ACS Nano 8, 2328–2335 (2014). https://doi.org/10.1021/nn405799t
- S.L. Brenner, V.A. Parsegian, A physical method for deriving the electrostatic interaction between rod-like polyions at all mutual angles. Biophys. J. 14, 327–334 (1974). https://doi.org/10.1016/S0006-3495(74)85919-9
- M.D. Frank-Kamenetskii, V.V. Anshelevich, A.V. Lukashin, Polyelectrolyte model of DNA. Sov. Phys. Usp. 30, 317–330 (1987). https://doi.org/10.1070/PU1987v030n04ABEH002833
- A.G. Cherstvy, Electrostatic interactions in biological DNA-related systems. Phys. Chem. Chem. Phys. 13, 9942–9968 (2011). https://doi.org/10.1039/C0CP02796K
- J. Lipfert, S. Doniach, R. Das, D. Herschlag, Understanding nucleic acid–ion interactions. Ann. Rev. Biochem. 83, 813–841 (2014). https://doi.org/10.1146/annurev-biochem-060409-092720
- D. Stigter, An electrostatic model for the dielectric effects, the adsorption of multivalent ions, and the bending of B-DNA. Biopolymers 46, 503–516 (1998)
- I. Rouzina, V.A. Bloomfield, DNA bending by small, mobile multivalent cations. Biophys. J. 74, 3152–3164 (1998). https://doi.org/10.1016/S0006-3495(98)78021-X
- A.I. Dragan, C.M. Read, E.N. Makeyeva, E.I. Milgotina, M.E.A. Churchill et al., DNA binding and bending by HMG boxes: energetic determinants of specificity. J. Mol. Biol. 343, 371–393 (2004). https://doi.org/10.1016/j.jmb.2004.08.035
- C. Rivetti, C. Walker, C. Bustamante, Polymer chain statistics and conformational analysis of DNA molecules with bends or sections of different flexibility. J. Mol. Biol. 280, 41–59 (1998). https://doi.org/10.1006/jmbi.1998.1830
- J. Baschnagel, H. Meyer, J. Wittmer, I. Kulic, H. Mohrbach et al., Semiflexible chains at surfaces: worm-like chains and beyond. Polymers 8, 286 (2016). https://doi.org/10.3390/polym8080286
- A. Milchev, K. Binder, Linear dimensions of adsorbed semiflexible polymers: what can be learned about their persistence length? Phys. Rev. Lett. 123, 128003 (2019). https://doi.org/10.1103/PhysRevLett.123.128003
- D.J. Bonthuis, S. Gekle, R.R. Netz, Dielectric profile of interfacial water and its effect on double-layer capacitance. Phys. Rev. Lett. 107, 166102 (2011). https://doi.org/10.1103/PhysRevLett.107.166102
- L. Fumagalli, A. Esfandiar, R. Fabregas, S. Hu, P. Ares et al., Anomalously low dielectric constant of confined water. Science 360, 1339–1342 (2018). https://doi.org/10.1126/science.aat4191
- W. Reisner, J.N. Pedersen, R.H. Austin, DNA confinement in nanochannels: physics and biological applications. Reports Prog. Phys. 75, 106601 (2012). https://doi.org/10.1088/0034-4885/75/10/106601
- S. Dasgupta, D.P. Allison, C.E. Snyder, S. Mitra, Base-unpaired regions in supercoiled replicative form DNA of coliphage M13. J. Biol. Chem. 252, 5916–5923 (1977)
- D.I. Cherny, T.M. Jovin, Electron and scanning force microscopy studies of alterations in supercoiled DNA tertiary structure. J. Mol. Biol. 313, 295–307 (2001). https://doi.org/10.1006/jmbi.2001.5031
- A.A. Kornyshev, S. Leikin, Electrostatic interaction between long, rigid helical macromolecules at all interaxial angles. Phys. Rev. E 62, 2576–2596 (2000). https://doi.org/10.1103/PhysRevE.62.2576
- E. Skoruppa, S.K. Nomidis, J.F. Marko, E. Carlon, Bend-induced twist waves and the structure of nucleosomal DNA. Phys. Rev. Lett. 121(8), 088101 (2018). https://doi.org/10.1103/PhysRevLett.121.088101
- B. Feng, R.P. Sosa, A.K.F. Mårtensson, K. Jiang, A. Tong et al. Hydrophobic catalysis and a potential biological role of DNA unstacking induced by environment effects. Proc. Natl. Acad. Sci. 116(35), 17169–17174 (2019). https://doi.org/10.1073/pnas.1909122116
- C. Danilowicz, Y. Kafri, R.S. Conroy, V.W. Coljee, J. Weeks et al., Measurement of the phase diagram of DNA unzipping in the temperature-force plane. Phys. Rev. Lett. 93, 078101 (2004). https://doi.org/10.1103/PhysRevLett.93.078101
- J. Camunas-Soler, M. Ribezzi-Crivellari, F. Ritort, Elastic properties of nucleic acids by single-molecule force spectroscopy. Annu. Rev. Biophys. 45, 65–84 (2016). https://doi.org/10.1146/annurev-biophys-062215-011158
- M. Manghi, N. Destainville, Physics of base-pairing dynamics in DNA. Phys. Rep. 631, 1–41 (2016). https://doi.org/10.1016/j.physrep.2016.04.001
- N. Bosaeus, A. Reymer, T. Beke-Somfai, T. Brown, M. Takahashi et al., A stretched conformation of DNA with a biological role? Q. Rev. Biophys. 50, e11 (2017). https://doi.org/10.1017/S0033583517000099
- R.N. Irobalieva, J.M. Fogg, D.J. Catanese Jr., T. Sutthibutpong, M. Chen et al., Structural diversity of supercoiled DNA. Nat. Commun. 6, 8440 (2015). https://doi.org/10.1006/10.1038/ncomms9440
- A.V. Vologodskii, A.V. Lukashin, V.V. Anshelevich, M.D. Frank-Kamenetskii, Fluctuations in superhelical DNA. Nucleic Acids Res. 6, 967–982 (1979). https://doi.org/10.1093/nar/6.3.967
- T.B. Liverpool, S.A. Harris, C.A. Laughton, Supercoiling and denaturation of DNA loops. Phys. Rev. Lett. 100, 238103 (2008). https://doi.org/10.1103/PhysRevLett.100.238103
- F. Sicard, N. Destainville, P. Rousseau, C. Tardin, M. Manghi, Dynamical control of denaturation bubble nucleation in supercoiled DNA minicircles. Phys. Rev. E 101, 012403 (2020). https://doi.org/10.1103/PhysRevE.101.012403
- C. Leung, A. Bestembayeva, R. Thorogate, J. Stinson, A. Pyne et al., Atomic force microscopy with nanoscale cantilevers resolves different structural conformations of the DNA double helix. Nano Lett. 12, 3846–3850 (2012). https://doi.org/10.1021/nl301857p
- S. Ido, K. Kimura, N. Oyabu, K. Kobayashi, M. Tsukada et al., Beyond the helix pitch: direct visualization of native DNA in aqueous solution. ACS Nano 7, 1817–1822 (2013). https://doi.org/10.1017/S0033583517000099
- P. Ares, M.E. Fuentes-Perez, E. Herrero-Galan, J.M. Valpuesta, A. Gil et al., High resolution atomic force microscopy of double-stranded RNA. Nanoscale 8, 11818–11826 (2016). https://doi.org/10.1039/C5NR07445B
- Y.F. Dufrêne, T. Ando, R. Garcia, D. Alsteens, D. Martinez-Martin et al., Imaging modes of atomic force microscopy for application in molecular and cell biology. Nat. Nanotechnol. 12, 295–307 (2017). https://doi.org/10.1038/nnano.2017.45
- K. Kuchuk, L. Katrivas, A. Kotlyar, U. Sivan, Sequence-dependent deviations of constrained dna from canonical B-form. Nano Lett. 19, 6600–6603 (2019). https://doi.org/10.1021/acs.nanolett.9b02863
References
J.P. Peters, L.J. Maher, DNA curvature and flexibility in vitro and in vivo. Q. Rev. Biophys. 43, 23–63 (2010). https://doi.org/10.1017/S0033583510000077
A. Vologodskii, M.D. Frank-Kamenetskii, Strong bending of the DNA double helix. Nucleic Acids Res. 41, 6785–6792 (2013). https://doi.org/10.1093/nar/gkt396
A.P. Fields, E.A. Meyer, A.E. Cohen, Euler buckling and nonlinear kinking of double-stranded DNA. Nucleic Acids Res. 41, 9881–9890 (2013). https://doi.org/10.1093/nar/gkt739
J. Wang, H. Qu, G. Zocchi, Critical bending torque of DNA is a materials parameter independent of local base sequence. Phys. Rev. E 88, 032712 (2013). https://doi.org/10.1103/PhysRevE.88.032712
C. Bustamante, Z. Bryant, S.B. Smith, Ten years of tension: single-molecule DNA mechanics. Nature 421, 423–427 (2003). https://doi.org/10.1038/nature01405
P. Gross, N. Laurens, L.B. Oddershede, U. Bockelmann, E.J.G. Peterman et al., Quantifying how DNA stretches, melts and changes twist under tension. Nat. Phys. 7, 731–736 (2011). https://doi.org/10.1038/nphys2002
J.F. Marko, S. Neukirch, Global force-torque phase diagram for the DNA double helix: Structural transitions, triple points, and collapsed plectonemes. Phys. Rev. E. 88, 062722 (2013). https://doi.org/10.1103/PhysRevE.88.062722
J.F. Marko, DNA Mechanics. in Nuclear Architecture and Dynamics, eds. C. Lavelle, J.-M.Victor (Elsevier, 2018), pp. 3–40.
F.H.C. Crick, A. Klug, Kinky helix. Nature 255, 530–533 (1975). https://doi.org/10.1038/255530a0
Q. Du, C. Smith, N. Shiffeldrim, M. Vologodskaia, A. Vologodskii, Cyclization of short DNA fragments and bending fluctuations of the double helix. Proc. Natl. Acad. Sci. USA 102, 5397–5402 (2005). https://doi.org/10.1073/pnas.0500983102
T.A. Lionberger, D. Demurtas, G. Witz, J. Dorier, T. Lillian et al., Cooperative kinking at distant sites in mechanically stressed DNA. Nucleic Acids Res. 39, 9820–9832 (2011). https://doi.org/10.1093/nar/gkr666
F. Lankaš, R. Lavery, J.H. Maddocks, Kinking occurs during molecular dynamics simulations of small DNA minicircles. Structure 14, 1527–1534 (2006). https://doi.org/10.1016/j.str.2006.08.004
J. Curuksu, M. Zacharias, R. Lavery, K. Zakrzewska, Local and global effects of strong DNA bending induced during molecular dynamics simulations. Nucleic Acids Res. 37, 3766–3773 (2009). https://doi.org/10.1093/nar/gkp234
J.P.K. Doye, T.E. Ouldridge, A.A. Louis, F. Romano, P. Sulc et al., Coarse-graining DNA for simulations of DNA nanotechnology. Phys. Chem. Chem. Phys. 15, 20395–20414 (2013). https://doi.org/10.1039/C3CP53545B
P. Cong, L. Dai, H. Chen, J.R.C. van der Maarel, P.S. Doyle et al., Revisiting the Anomalous Bending Elasticity of Sharply Bent DNA. Biophys. J. 109, 2338–2351 (2015). https://doi.org/10.1016/j.bpj.2015.10.016
R.M. Harrison, F. Romano, T.E. Ouldridge, A.A. Louis, J.P.K. Doye, Identifying physical causes of apparent enhanced cyclization of short DNA molecules with a coarse-grained model. J. Chem. Theory Comput. 15, 4660–4672 (2019). https://doi.org/10.1021/acs.jctc.9b00112
C. Rivetti, M. Guthold, C. Bustamante, Scanning force microscopy of DNA deposited onto mica: equilibration versus kinetic trapping studied by statistical polymer chain analysis. J. Mol. Biol. 264, 919–932 (1996). https://doi.org/10.1006/jmbi.1996.0687
M. Sushko, A. Shluger, C. Rivetti, Simple model for DNA adsorption onto a mica surface in 1:1 and 2:1 electrolyte solutions. Langmuir 22, 7678–7688 (2006). https://doi.org/10.1021/la060356+
P.A. Wiggins, T. van der Heijden, F. Moreno-Herrero, A. Spakowitz, R. Phillips et al., High flexibility of DNA on short length scales probed by atomic force microscopy. Nat. Nanotechnol. 1, 137–141 (2006). https://doi.org/10.1038/nnano.2006.63
A.K. Mazur, M. Maaloum, Atomic force microscopy study of DNA flexibility on short length scales: smooth bending versus kinking. Nucleic Acids Res. 42, 14006–14012 (2014). https://doi.org/10.1093/nar/gku1192
T. Brouns, H. De Keersmaecker, S.F. Konrad, N. Kodera, T. Ando et al., Free energy landscape and dynamics of supercoiled DNA by high-speed atomic force microscopy. ACS Nano 12, 11907–11916 (2018). https://doi.org/10.1021/acsnano.8b06994
A. Podestà, M. Indrieri, D. Brogioli, G.S. Manning, P. Milani et al., Positively Charged Surfaces Increase the Flexibility of DNA. Biophys. J. 89, 2558–2563 (2005). https://doi.org/10.1529/biophysj.105.064667
J.-H. Jeon, J. Adamcik, G. Dietler, R. Metzler, Supercoiling induces denaturation bubbles in circular DNA. Phys. Rev. Lett. 105, 208101 (2010). https://doi.org/10.1103/PhysRevLett.105.208101
Y.L. Lyubchenko, L.S. Shlyakhtenko, T. Ando, Imaging of nucleic acids with atomic force microscopy. Methods 54, 274–283 (2011). https://doi.org/10.1016/j.ymeth.2011.02.001
B. Akpinar, P.J. Haynes, N.A.W. Bell, K. Brunner, A.L.B. Pyne et al., PEGylated surfaces for the study of DNA–protein interactions by atomic force microscopy. Nanoscale 11, 20072–20080 (2019). https://doi.org/10.1039/C9NR07104K
N. Severin, J. Barner, A.A. Kalachev, J.P. Rabe, Manipulation and overstretching of genes on solid substrates. Nano Lett. 4, 577–579 (2004). https://doi.org/10.1021/nl035147d
J. Adamcik, D.V. Klinov, G. Witz, S.K. Sekatskii, G. Dietler, Observation of single-stranded DNA on mica and highly oriented pyrolytic graphite by atomic force microscopy. FEBS Lett. 580, 5671–5675 (2006). https://doi.org/10.1016/j.febslet.2006.09.017
D. Klinov, B. Dwir, E. Kapon, N. Borovok, T. Molotsky et al., High-resolution atomic force microscopy of duplex and triplex DNA molecules. Nanotechnology 18, 225102 (2007). https://doi.org/10.1088/0957-4484/18/22/225102
E.V. Dubrovin, M. Schächtele, T.E. Schäffer, Nanotemplate-directed DNA segmental thermal motion. RSC Adv. 6, 79584–79592 (2016). https://doi.org/10.1039/C6RA14383K
Spontaneous large-angle bends in short DNA contour segments which are many times more prevalent than predicted by the canonical WLC model have been reported for the freely equilibrated DNA on mica using AFM imaging in air [19], however the normal bending angle distribution with no noticeable deflection from WLC model has been found in in-situ AFM measurements carried out in solution [20].
V.V. Prokhorov, D.V. Klinov, A.A. Chinarev, A.B. Tuzikov, I.V. Gorokhova et al., High-resolution atomic force microscopy study of hexaglycylamide epitaxial structures on graphite. Langmuir 27, 5879–5890 (2011). https://doi.org/10.1021/la103051w
R. García, R. Perez, Dynamic atomic force microscopy methods. Surf. Sci. Rep. 47, 197–301 (2002). https://doi.org/10.1016/S0167-5729(02)00077-8
V.V. Prokhorov, S.A. Saunin, Probe-surface interaction mapping in amplitude modulation atomic force microscopy by integrating amplitude-distance and amplitude-frequency curves. Appl. Phys. Lett. 91, 023122 (2007). https://doi.org/10.1063/1.2756271
A. Mikhaylov, S.K. Sekatskii, G. Dietler, D.N.A. Trace, A comprehensive software for polymer image processing. J. Adv. Microsc. Res. 8, 241–245 (2013). https://doi.org/10.1166/jamr.2013.1164
M. Frank-Kamenetskii, How the double helix breathes. Nature 328, 17–18 (1987). https://doi.org/10.1038/328017a0
J.P. Rabe, S. Buchholz, Commensurability and mobility in two-dimensional molecular patterns on graphite. Science 253, 424–427 (1991). https://doi.org/10.1126/science.253.5018.424
V.V. Prokhorov, K. Nitta, The AFM observation of linear chain and crystalline conformations of ultrahigh molecular weight polyethylene molecules on mica and graphite. J. Polym. Sci. Part B Polym. Phys. 48, 766–777 (2010). https://doi.org/10.1002/polb.21944
W. Hoyer, D. Cherny, V. Subramaniam, T.M. Jovin, Rapid self-assembly of α-synuclein observed by in situ atomic force microscopy. J. Mol. Biol. 340, 127–139 (2004). https://doi.org/10.1016/j.jmb.2004.04.051
C. Whitehouse, J. Fang, A. Aggeli, M. Bell, R. Brydson et al., Adsorption and self-assembly of peptides on mica substrates. Angew. Chem. Int. Ed. 44, 1965–1968 (2005). https://doi.org/10.1002/anie.200462160
D. Bagrov, Y. Gazizova, V. Podgorsky, I. Udovichenko, A. Danilkovich et al., Morphology and aggregation of RADA-16-I peptide studied by AFM, NMR and molecular dynamics simulations. Biopolymers 106, 72–81 (2016). https://doi.org/10.1002/bip.22755
C. Charbonneau, J.M. Kleijn, M.A. Cohen Stuart, Subtle charge balance controls surface-nucleated self-assembly of designed biopolymers. ACS Nano 8, 2328–2335 (2014). https://doi.org/10.1021/nn405799t
S.L. Brenner, V.A. Parsegian, A physical method for deriving the electrostatic interaction between rod-like polyions at all mutual angles. Biophys. J. 14, 327–334 (1974). https://doi.org/10.1016/S0006-3495(74)85919-9
M.D. Frank-Kamenetskii, V.V. Anshelevich, A.V. Lukashin, Polyelectrolyte model of DNA. Sov. Phys. Usp. 30, 317–330 (1987). https://doi.org/10.1070/PU1987v030n04ABEH002833
A.G. Cherstvy, Electrostatic interactions in biological DNA-related systems. Phys. Chem. Chem. Phys. 13, 9942–9968 (2011). https://doi.org/10.1039/C0CP02796K
J. Lipfert, S. Doniach, R. Das, D. Herschlag, Understanding nucleic acid–ion interactions. Ann. Rev. Biochem. 83, 813–841 (2014). https://doi.org/10.1146/annurev-biochem-060409-092720
D. Stigter, An electrostatic model for the dielectric effects, the adsorption of multivalent ions, and the bending of B-DNA. Biopolymers 46, 503–516 (1998)
I. Rouzina, V.A. Bloomfield, DNA bending by small, mobile multivalent cations. Biophys. J. 74, 3152–3164 (1998). https://doi.org/10.1016/S0006-3495(98)78021-X
A.I. Dragan, C.M. Read, E.N. Makeyeva, E.I. Milgotina, M.E.A. Churchill et al., DNA binding and bending by HMG boxes: energetic determinants of specificity. J. Mol. Biol. 343, 371–393 (2004). https://doi.org/10.1016/j.jmb.2004.08.035
C. Rivetti, C. Walker, C. Bustamante, Polymer chain statistics and conformational analysis of DNA molecules with bends or sections of different flexibility. J. Mol. Biol. 280, 41–59 (1998). https://doi.org/10.1006/jmbi.1998.1830
J. Baschnagel, H. Meyer, J. Wittmer, I. Kulic, H. Mohrbach et al., Semiflexible chains at surfaces: worm-like chains and beyond. Polymers 8, 286 (2016). https://doi.org/10.3390/polym8080286
A. Milchev, K. Binder, Linear dimensions of adsorbed semiflexible polymers: what can be learned about their persistence length? Phys. Rev. Lett. 123, 128003 (2019). https://doi.org/10.1103/PhysRevLett.123.128003
D.J. Bonthuis, S. Gekle, R.R. Netz, Dielectric profile of interfacial water and its effect on double-layer capacitance. Phys. Rev. Lett. 107, 166102 (2011). https://doi.org/10.1103/PhysRevLett.107.166102
L. Fumagalli, A. Esfandiar, R. Fabregas, S. Hu, P. Ares et al., Anomalously low dielectric constant of confined water. Science 360, 1339–1342 (2018). https://doi.org/10.1126/science.aat4191
W. Reisner, J.N. Pedersen, R.H. Austin, DNA confinement in nanochannels: physics and biological applications. Reports Prog. Phys. 75, 106601 (2012). https://doi.org/10.1088/0034-4885/75/10/106601
S. Dasgupta, D.P. Allison, C.E. Snyder, S. Mitra, Base-unpaired regions in supercoiled replicative form DNA of coliphage M13. J. Biol. Chem. 252, 5916–5923 (1977)
D.I. Cherny, T.M. Jovin, Electron and scanning force microscopy studies of alterations in supercoiled DNA tertiary structure. J. Mol. Biol. 313, 295–307 (2001). https://doi.org/10.1006/jmbi.2001.5031
A.A. Kornyshev, S. Leikin, Electrostatic interaction between long, rigid helical macromolecules at all interaxial angles. Phys. Rev. E 62, 2576–2596 (2000). https://doi.org/10.1103/PhysRevE.62.2576
E. Skoruppa, S.K. Nomidis, J.F. Marko, E. Carlon, Bend-induced twist waves and the structure of nucleosomal DNA. Phys. Rev. Lett. 121(8), 088101 (2018). https://doi.org/10.1103/PhysRevLett.121.088101
B. Feng, R.P. Sosa, A.K.F. Mårtensson, K. Jiang, A. Tong et al. Hydrophobic catalysis and a potential biological role of DNA unstacking induced by environment effects. Proc. Natl. Acad. Sci. 116(35), 17169–17174 (2019). https://doi.org/10.1073/pnas.1909122116
C. Danilowicz, Y. Kafri, R.S. Conroy, V.W. Coljee, J. Weeks et al., Measurement of the phase diagram of DNA unzipping in the temperature-force plane. Phys. Rev. Lett. 93, 078101 (2004). https://doi.org/10.1103/PhysRevLett.93.078101
J. Camunas-Soler, M. Ribezzi-Crivellari, F. Ritort, Elastic properties of nucleic acids by single-molecule force spectroscopy. Annu. Rev. Biophys. 45, 65–84 (2016). https://doi.org/10.1146/annurev-biophys-062215-011158
M. Manghi, N. Destainville, Physics of base-pairing dynamics in DNA. Phys. Rep. 631, 1–41 (2016). https://doi.org/10.1016/j.physrep.2016.04.001
N. Bosaeus, A. Reymer, T. Beke-Somfai, T. Brown, M. Takahashi et al., A stretched conformation of DNA with a biological role? Q. Rev. Biophys. 50, e11 (2017). https://doi.org/10.1017/S0033583517000099
R.N. Irobalieva, J.M. Fogg, D.J. Catanese Jr., T. Sutthibutpong, M. Chen et al., Structural diversity of supercoiled DNA. Nat. Commun. 6, 8440 (2015). https://doi.org/10.1006/10.1038/ncomms9440
A.V. Vologodskii, A.V. Lukashin, V.V. Anshelevich, M.D. Frank-Kamenetskii, Fluctuations in superhelical DNA. Nucleic Acids Res. 6, 967–982 (1979). https://doi.org/10.1093/nar/6.3.967
T.B. Liverpool, S.A. Harris, C.A. Laughton, Supercoiling and denaturation of DNA loops. Phys. Rev. Lett. 100, 238103 (2008). https://doi.org/10.1103/PhysRevLett.100.238103
F. Sicard, N. Destainville, P. Rousseau, C. Tardin, M. Manghi, Dynamical control of denaturation bubble nucleation in supercoiled DNA minicircles. Phys. Rev. E 101, 012403 (2020). https://doi.org/10.1103/PhysRevE.101.012403
C. Leung, A. Bestembayeva, R. Thorogate, J. Stinson, A. Pyne et al., Atomic force microscopy with nanoscale cantilevers resolves different structural conformations of the DNA double helix. Nano Lett. 12, 3846–3850 (2012). https://doi.org/10.1021/nl301857p
S. Ido, K. Kimura, N. Oyabu, K. Kobayashi, M. Tsukada et al., Beyond the helix pitch: direct visualization of native DNA in aqueous solution. ACS Nano 7, 1817–1822 (2013). https://doi.org/10.1017/S0033583517000099
P. Ares, M.E. Fuentes-Perez, E. Herrero-Galan, J.M. Valpuesta, A. Gil et al., High resolution atomic force microscopy of double-stranded RNA. Nanoscale 8, 11818–11826 (2016). https://doi.org/10.1039/C5NR07445B
Y.F. Dufrêne, T. Ando, R. Garcia, D. Alsteens, D. Martinez-Martin et al., Imaging modes of atomic force microscopy for application in molecular and cell biology. Nat. Nanotechnol. 12, 295–307 (2017). https://doi.org/10.1038/nnano.2017.45
K. Kuchuk, L. Katrivas, A. Kotlyar, U. Sivan, Sequence-dependent deviations of constrained dna from canonical B-form. Nano Lett. 19, 6600–6603 (2019). https://doi.org/10.1021/acs.nanolett.9b02863