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Tessmer I. The roles of non-productive complexes of DNA repair proteins with DNA lesions. DNA Repair (Amst) 2023; 129:103542. [PMID: 37453245 DOI: 10.1016/j.dnarep.2023.103542] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/06/2023] [Revised: 06/30/2023] [Accepted: 07/07/2023] [Indexed: 07/18/2023]
Abstract
A multitude of different types of lesions is continuously introduced into the DNA inside our cells, and their rapid and efficient repair is fundamentally important for the maintenance of genomic stability and cellular viability. This is achieved by a number of DNA repair systems that each involve different protein factors and employ versatile strategies to target different types of DNA lesions. Intriguingly, specialized DNA repair proteins have also evolved to form non-functional complexes with their target lesions. These proteins allow the marking of innocuous lesions to render them visible for DNA repair systems and can serve to directly recruit DNA repair cascades. Moreover, they also provide links between different DNA repair mechanisms or even between DNA lesions and transcription regulation. I will focus here in particular on recent findings from single molecule analyses on the alkyltransferase-like protein ATL, which is believed to initiate nucleotide excision repair (NER) of non-native NER target lesions, and the base excision repair (BER) enzyme hOGG1, which recruits the oncogene transcription factor Myc to gene promoters under oxidative stress.
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Affiliation(s)
- Ingrid Tessmer
- Rudolf Virchow Center, University of Würzburg, Josef Schneider Str. 2, 97080 Würzburg, Germany
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2
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Irvin EM, Wang H. Single-molecule imaging of genome maintenance proteins encountering specific DNA sequences and structures. DNA Repair (Amst) 2023; 128:103528. [PMID: 37392578 PMCID: PMC10989508 DOI: 10.1016/j.dnarep.2023.103528] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/08/2023] [Revised: 06/08/2023] [Accepted: 06/21/2023] [Indexed: 07/03/2023]
Abstract
DNA repair pathways are tightly regulated processes that recognize specific hallmarks of DNA damage and coordinate lesion repair through discrete mechanisms, all within the context of a three-dimensional chromatin landscape. Dysregulation or malfunction of any one of the protein constituents in these pathways can contribute to aging and a variety of diseases. While the collective action of these many proteins is what drives DNA repair on the organismal scale, it is the interactions between individual proteins and DNA that facilitate each step of these pathways. In much the same way that ensemble biochemical techniques have characterized the various steps of DNA repair pathways, single-molecule imaging (SMI) approaches zoom in further, characterizing the individual protein-DNA interactions that compose each pathway step. SMI techniques offer the high resolving power needed to characterize the molecular structure and functional dynamics of individual biological interactions on the nanoscale. In this review, we highlight how our lab has used SMI techniques - traditional atomic force microscopy (AFM) imaging in air, high-speed AFM (HS-AFM) in liquids, and the DNA tightrope assay - over the past decade to study protein-nucleic acid interactions involved in DNA repair, mitochondrial DNA replication, and telomere maintenance. We discuss how DNA substrates containing specific DNA sequences or structures that emulate DNA repair intermediates or telomeres were generated and validated. For each highlighted project, we discuss novel findings made possible by the spatial and temporal resolution offered by these SMI techniques and unique DNA substrates.
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Affiliation(s)
| | - Hong Wang
- Toxicology Program, North Carolina State University, Raleigh, NC, USA; Physics Department, North Carolina State University, Raleigh, NC, USA; Center for Human Health and the Environment, North Carolina State University, Raleigh, NC, USA.
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3
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Bangalore DM, Tessmer I. Direct hOGG1-Myc interactions inhibit hOGG1 catalytic activity and recruit Myc to its promoters under oxidative stress. Nucleic Acids Res 2022; 50:10385-10398. [PMID: 36156093 PMCID: PMC9561264 DOI: 10.1093/nar/gkac796] [Citation(s) in RCA: 6] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/08/2022] [Revised: 08/23/2022] [Accepted: 09/02/2022] [Indexed: 11/13/2022] Open
Abstract
The base excision repair (BER) glycosylase hOGG1 (human oxoguanine glycosylase 1) is responsible for repairing oxidative lesions in the genome, in particular oxidised guanine bases (oxoG). In addition, a role of hOGG1 in transcription regulation by recruitment of various transcription factors has been reported. Here, we demonstrate direct interactions between hOGG1 and the medically important oncogene transcription factor Myc that is involved in transcription initiation of a large number of genes including inflammatory genes. Using single molecule atomic force microscopy (AFM), we reveal recruitment of Myc to its E-box promoter recognition sequence by hOGG1 specifically under oxidative stress conditions, and conformational changes in hOGG1-Myc complexes at oxoG lesions that suggest loading of Myc at oxoG lesions by hOGG1. Importantly, our data show suppression of hOGG1 catalytic activity in oxoG repair by Myc. Furthermore, mutational analyses implicate the C28 residue in hOGG1 in oxidation induced protein dimerisation and suggest a role of hOGG1 dimerisation under oxidising conditions in hOGG1-Myc interactions. From our data we develop a mechanistic model for Myc recruitment by hOGG1 under oxidising, inflammatory conditions, which may be responsible for the observed enhanced gene expression of Myc target genes.
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Affiliation(s)
- Disha M Bangalore
- Rudolf Virchow Center, University of Würzburg, Josef Schneider Str. 2, 97080 Würzburg, Germany
| | - Ingrid Tessmer
- Rudolf Virchow Center, University of Würzburg, Josef Schneider Str. 2, 97080 Würzburg, Germany
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Konrad SF, Vanderlinden W, Lipfert J. A High-throughput Pipeline to Determine DNA and Nucleosome Conformations by AFM Imaging. Bio Protoc 2021; 11:e4180. [PMID: 34722827 DOI: 10.21769/bioprotoc.4180] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/28/2021] [Revised: 06/30/2021] [Accepted: 07/28/2021] [Indexed: 11/02/2022] Open
Abstract
Atomic force microscopy (AFM) is a powerful tool to image macromolecular complexes with nanometer resolution and exquisite single-molecule sensitivity. While AFM imaging is well-established to investigate DNA and nucleoprotein complexes, AFM studies are often limited by small datasets and manual image analysis that is slow and prone to user bias. Recently, we have shown that a combination of large scale AFM imaging and automated image analysis of nucleosomes can overcome these previous limitations of AFM nucleoprotein studies. Using our high-throughput imaging and analysis pipeline, we have resolved nucleosome wrapping intermediates with five base pair resolution and revealed how distinct nucleosome variants and environmental conditions affect the unwrapping pathways of nucleosomal DNA. Here, we provide a detailed protocol of our workflow to analyze DNA and nucleosome conformations focusing on practical aspects and experimental parameters. We expect our protocol to drastically enhance AFM analyses of DNA and nucleosomes and to be readily adaptable to a wide variety of other protein and protein-nucleic acid complexes.
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Affiliation(s)
- Sebastian F Konrad
- Department of Physics and Center for Nanoscience, LMU Munich, Amalienstr. 54, 80799 Munich, Germany
| | - Willem Vanderlinden
- Department of Physics and Center for Nanoscience, LMU Munich, Amalienstr. 54, 80799 Munich, Germany
| | - Jan Lipfert
- Department of Physics and Center for Nanoscience, LMU Munich, Amalienstr. 54, 80799 Munich, Germany
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5
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Main KHS, Provan JI, Haynes PJ, Wells G, Hartley JA, Pyne ALB. Atomic force microscopy-A tool for structural and translational DNA research. APL Bioeng 2021; 5:031504. [PMID: 34286171 PMCID: PMC8272649 DOI: 10.1063/5.0054294] [Citation(s) in RCA: 11] [Impact Index Per Article: 3.7] [Reference Citation Analysis] [Abstract] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/16/2021] [Accepted: 06/07/2021] [Indexed: 12/26/2022] Open
Abstract
Atomic force microscopy (AFM) is a powerful imaging technique that allows for structural characterization of single biomolecules with nanoscale resolution. AFM has a unique capability to image biological molecules in their native states under physiological conditions without the need for labeling or averaging. DNA has been extensively imaged with AFM from early single-molecule studies of conformational diversity in plasmids, to recent examinations of intramolecular variation between groove depths within an individual DNA molecule. The ability to image dynamic biological interactions in situ has also allowed for the interaction of various proteins and therapeutic ligands with DNA to be evaluated-providing insights into structural assembly, flexibility, and movement. This review provides an overview of how innovation and optimization in AFM imaging have advanced our understanding of DNA structure, mechanics, and interactions. These include studies of the secondary and tertiary structure of DNA, including how these are affected by its interactions with proteins. The broader role of AFM as a tool in translational cancer research is also explored through its use in imaging DNA with key chemotherapeutic ligands, including those currently employed in clinical practice.
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Affiliation(s)
| | - James I. Provan
- Institute of Molecular, Cell, and Systems Biology, University of Glasgow, Glasgow G12 8QQ, United Kingdom
| | | | - Geoffrey Wells
- UCL School of Pharmacy, University College London, London WC1N 1AX, United Kingdom
| | - John A. Hartley
- UCL Cancer Institute, University College London, London WC1E 6DD, United Kingdom
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Emperle M, Bangalore DM, Adam S, Kunert S, Heil HS, Heinze KG, Bashtrykov P, Tessmer I, Jeltsch A. Structural and biochemical insight into the mechanism of dual CpG site binding and methylation by the DNMT3A DNA methyltransferase. Nucleic Acids Res 2021; 49:8294-8308. [PMID: 34289056 PMCID: PMC8373138 DOI: 10.1093/nar/gkab600] [Citation(s) in RCA: 6] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/14/2021] [Revised: 06/23/2021] [Accepted: 06/30/2021] [Indexed: 01/02/2023] Open
Abstract
DNMT3A/3L heterotetramers contain two active centers binding CpG sites at 12 bp distance, however their interaction with DNA not containing this feature is unclear. Using randomized substrates, we observed preferential co-methylation of CpG sites with 6, 9 and 12 bp spacing by DNMT3A and DNMT3A/3L. Co-methylation was favored by AT bases between the 12 bp spaced CpG sites consistent with their increased bending flexibility. SFM analyses of DNMT3A/3L complexes bound to CpG sites with 12 bp spacing revealed either single heterotetramers inducing 40° DNA bending as observed in the X-ray structure, or two heterotetramers bound side-by-side to the DNA yielding 80° bending. SFM data of DNMT3A/3L bound to CpG sites spaced by 6 and 9 bp revealed binding of two heterotetramers and 100° DNA bending. Modeling showed that for 6 bp distance between CpG sites, two DNMT3A/3L heterotetramers could bind side-by-side on the DNA similarly as for 12 bp distance, but with each CpG bound by a different heterotetramer. For 9 bp spacing our model invokes a tetramer swap of the bound DNA. These additional DNA interaction modes explain how DNMT3A and DNMT3A/3L overcome their structural preference for CpG sites with 12 bp spacing during the methylation of natural DNA.
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Affiliation(s)
- Max Emperle
- Institute of Biochemistry and Technical Biochemistry, Department of Biochemistry, University of Stuttgart, Stuttgart, Germany
| | - Disha M Bangalore
- Rudolf Virchow Center for Integrative and Translational Bioimaging, University of Würzburg, Würzburg, Germany
| | - Sabrina Adam
- Institute of Biochemistry and Technical Biochemistry, Department of Biochemistry, University of Stuttgart, Stuttgart, Germany
| | - Stefan Kunert
- Institute of Biochemistry and Technical Biochemistry, Department of Biochemistry, University of Stuttgart, Stuttgart, Germany
| | - Hannah S Heil
- Rudolf Virchow Center for Integrative and Translational Bioimaging, University of Würzburg, Würzburg, Germany
| | - Katrin G Heinze
- Rudolf Virchow Center for Integrative and Translational Bioimaging, University of Würzburg, Würzburg, Germany
| | - Pavel Bashtrykov
- Institute of Biochemistry and Technical Biochemistry, Department of Biochemistry, University of Stuttgart, Stuttgart, Germany
| | - Ingrid Tessmer
- Rudolf Virchow Center for Integrative and Translational Bioimaging, University of Würzburg, Würzburg, Germany
| | - Albert Jeltsch
- Institute of Biochemistry and Technical Biochemistry, Department of Biochemistry, University of Stuttgart, Stuttgart, Germany
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Dubrovin EV, Dadinova LA, Petoukhov MV, Soshinskaya EY, Mozhaev AA, Klinov DV, Schäffer TE, Shtykova EV, Batishchev OV. Spatial organization of Dps and DNA-Dps complexes. J Mol Biol 2021; 433:166930. [PMID: 33713674 DOI: 10.1016/j.jmb.2021.166930] [Citation(s) in RCA: 8] [Impact Index Per Article: 2.7] [Reference Citation Analysis] [Abstract] [Key Words] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/25/2020] [Revised: 02/24/2021] [Accepted: 03/05/2021] [Indexed: 02/04/2023]
Abstract
DNA co-crystallization with Dps family proteins is a fundamental mechanism, which preserves DNA in bacteria from harsh conditions. Though many aspects of this phenomenon are well characterized, the spatial organization of DNA in DNA-Dps co-crystals is not completely understood, and existing models need further clarification. To advance in this problem we have utilized atomic force microscopy (AFM) as the main structural tool, and small-angle X-scattering (SAXS) to characterize Dps as a key component of the DNA-protein complex. SAXS analysis in the presence of EDTA indicates a significantly larger radius of gyration for Dps than would be expected for the core of the dodecamer, consistent with the N-terminal regions extending out into solution and being accessible for interaction with DNA. In AFM experiments, both Dps protein molecules and DNA-Dps complexes adsorbed on mica or highly oriented pyrolytic graphite (HOPG) surfaces form densely packed hexagonal structures with a characteristic size of about 9 nm. To shed light on the peculiarities of DNA interaction with Dps molecules, we have characterized individual DNA-Dps complexes. Contour length evaluation has confirmed the non-specific character of Dps binding with DNA and revealed that DNA does not wrap Dps molecules in DNA-Dps complexes. Angle analysis has demonstrated that in DNA-Dps complexes a Dps molecule contacts with a DNA segment of ~6 nm in length. Consideration of DNA condensation upon complex formation with small Dps quasi-crystals indicates that DNA may be arranged along the rows of ordered protein molecules on a Dps sheet.
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Affiliation(s)
- Evgeniy V Dubrovin
- A.N. Frumkin Institute of Physical Chemistry and Electrochemistry, Russian Academy of Sciences, 31/4 Leninskiy prospekt, Moscow 119071, Russia; Lomonosov Moscow State University, Faculty of Physics, Leninskie Gory 1 bld 2, 119991 Moscow, Russia.
| | - Liubov A Dadinova
- Shubnikov Institute of Crystallography, Federal Scientific Research Centre "Crystallography and Photonics," Russian Academy of Sciences, 119333 Moscow, Russia
| | - Maxim V Petoukhov
- Shubnikov Institute of Crystallography, Federal Scientific Research Centre "Crystallography and Photonics," Russian Academy of Sciences, 119333 Moscow, Russia
| | - Ekaterina Yu Soshinskaya
- Shubnikov Institute of Crystallography, Federal Scientific Research Centre "Crystallography and Photonics," Russian Academy of Sciences, 119333 Moscow, Russia
| | - Andrey A Mozhaev
- Shubnikov Institute of Crystallography, Federal Scientific Research Centre "Crystallography and Photonics," Russian Academy of Sciences, 119333 Moscow, Russia; Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences, 117997 Moscow, Russia
| | - Dmitry V Klinov
- Federal Research and Clinical Center of Physical-Chemical Medicine of Federal Medical Biological Agency, Malaya Pirogovskaya 1a, 119435 Moscow, Russia
| | - Tilman E Schäffer
- University of Tübingen, Institute of Applied Physics, Auf der Morgenstelle 10, 72076 Tübingen, Germany
| | - Eleonora V Shtykova
- Shubnikov Institute of Crystallography, Federal Scientific Research Centre "Crystallography and Photonics," Russian Academy of Sciences, 119333 Moscow, Russia
| | - Oleg V Batishchev
- A.N. Frumkin Institute of Physical Chemistry and Electrochemistry, Russian Academy of Sciences, 31/4 Leninskiy prospekt, Moscow 119071, Russia
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