Publications by authors named "Jan Ellenberg"

142 Publications

Super-Resolution Spatial Proximity Detection with Proximity-PAINT.

Angew Chem Int Ed Engl 2021 Jan 9;60(2):716-720. Epub 2020 Nov 9.

Faculty of Physics and Center for Nanoscience, LMU Munich, Geschwister-Scholl-Platz 1, 80539, Munich, Germany.

Visualizing the functional interactions of biomolecules such as proteins and nucleic acids is key to understanding cellular life on the molecular scale. Spatial proximity is often used as a proxy for the direct interaction of biomolecules. However, current techniques to visualize spatial proximity are either limited by spatial resolution, dynamic range, or lack of single-molecule sensitivity. Here, we introduce Proximity-PAINT (pPAINT), a variation of the super-resolution microscopy technique DNA-PAINT. pPAINT uses a split-docking-site configuration to detect spatial proximity with high sensitivity, low false-positive rates, and tunable detection distances. We benchmark and optimize pPAINT using designer DNA nanostructures and demonstrate its cellular applicability by visualizing the spatial proximity of alpha- and beta-tubulin in microtubules using super-resolution detection.
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http://dx.doi.org/10.1002/anie.202009031DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC7839522PMC
January 2021

LifeTime and improving European healthcare through cell-based interceptive medicine.

Nature 2020 11 7;587(7834):377-386. Epub 2020 Sep 7.

Department of Human Genetics, KU Leuven, Leuven, Belgium.

Here we describe the LifeTime Initiative, which aims to track, understand and target human cells during the onset and progression of complex diseases, and to analyse their response to therapy at single-cell resolution. This mission will be implemented through the development, integration and application of single-cell multi-omics and imaging, artificial intelligence and patient-derived experimental disease models during the progression from health to disease. The analysis of large molecular and clinical datasets will identify molecular mechanisms, create predictive computational models of disease progression, and reveal new drug targets and therapies. The timely detection and interception of disease embedded in an ethical and patient-centred vision will be achieved through interactions across academia, hospitals, patient associations, health data management systems and industry. The application of this strategy to key medical challenges in cancer, neurological and neuropsychiatric disorders, and infectious, chronic inflammatory and cardiovascular diseases at the single-cell level will usher in cell-based interceptive medicine in Europe over the next decade.
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http://dx.doi.org/10.1038/s41586-020-2715-9DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC7656507PMC
November 2020

Chemogenetic Control of Nanobodies.

Nat Methods 2020 03 17;17(3):279-282. Epub 2020 Feb 17.

Department of Chemical Biology, Max Planck Institute for Medical Research, Heidelberg, Germany.

We introduce an engineered nanobody whose affinity to green fluorescent protein (GFP) can be switched on and off with small molecules. By controlling the cellular localization of GFP fusion proteins, the engineered nanobody allows interrogation of their roles in basic biological processes, an approach that should be applicable to numerous previously described GFP fusions. We also outline how the binding affinities of other nanobodies can be controlled by small molecules.
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http://dx.doi.org/10.1038/s41592-020-0746-7DOI Listing
March 2020

MINFLUX nanoscopy delivers 3D multicolor nanometer resolution in cells.

Nat Methods 2020 02 13;17(2):217-224. Epub 2020 Jan 13.

Department of NanoBiophotonics, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany.

The ultimate goal of biological super-resolution fluorescence microscopy is to provide three-dimensional resolution at the size scale of a fluorescent marker. Here we show that by localizing individual switchable fluorophores with a probing donut-shaped excitation beam, MINFLUX nanoscopy can provide resolutions in the range of 1 to 3 nm for structures in fixed and living cells. This progress has been facilitated by approaching each fluorophore iteratively with the probing-donut minimum, making the resolution essentially uniform and isotropic over scalable fields of view. MINFLUX imaging of nuclear pore complexes of a mammalian cell shows that this true nanometer-scale resolution is obtained in three dimensions and in two color channels. Relying on fewer detected photons than standard camera-based localization, MINFLUX nanoscopy is poised to open a new chapter in the imaging of protein complexes and distributions in fixed and living cells.
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http://dx.doi.org/10.1038/s41592-019-0688-0DOI Listing
February 2020

Photoactivation of silicon rhodamines via a light-induced protonation.

Nat Commun 2019 10 8;10(1):4580. Epub 2019 Oct 8.

Department of Chemical Biology, Max Planck Institute for Medical Research, Jahnstrasse 29, 69120, Heidelberg, Germany.

Photoactivatable fluorophores are important for single-particle tracking and super-resolution microscopy. Here we present a photoactivatable fluorophore that forms a bright silicon rhodamine derivative through a light-dependent protonation. In contrast to other photoactivatable fluorophores, no caging groups are required, nor are there any undesired side-products released. Using this photoactivatable fluorophore, we create probes for HaloTag and actin for live-cell single-molecule localization microscopy and single-particle tracking experiments. The unusual mechanism of photoactivation and the fluorophore's outstanding spectroscopic properties make it a powerful tool for live-cell super-resolution microscopy.
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http://dx.doi.org/10.1038/s41467-019-12480-3DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6783549PMC
October 2019

Nuclear pores as versatile reference standards for quantitative superresolution microscopy.

Nat Methods 2019 10 27;16(10):1045-1053. Epub 2019 Sep 27.

EMBL, Cell Biology and Biophysics, Heidelberg, Germany.

Quantitative fluorescence and superresolution microscopy are often limited by insufficient data quality or artifacts. In this context, it is essential to have biologically relevant control samples to benchmark and optimize the quality of microscopes, labels and imaging conditions. Here, we exploit the stereotypic arrangement of proteins in the nuclear pore complex as in situ reference structures to characterize the performance of a variety of microscopy modalities. We created four genome edited cell lines in which we endogenously labeled the nucleoporin Nup96 with mEGFP, SNAP-tag, HaloTag or the photoconvertible fluorescent protein mMaple. We demonstrate their use (1) as three-dimensional resolution standards for calibration and quality control, (2) to quantify absolute labeling efficiencies and (3) as precise reference standards for molecular counting. These cell lines will enable the broader community to assess the quality of their microscopes and labels, and to perform quantitative, absolute measurements.
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http://dx.doi.org/10.1038/s41592-019-0574-9DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6768092PMC
October 2019

Direct Visualization of Single Nuclear Pore Complex Proteins Using Genetically-Encoded Probes for DNA-PAINT.

Angew Chem Int Ed Engl 2019 09 21;58(37):13004-13008. Epub 2019 Aug 21.

Faculty of Physics and Center for Nanoscience, LMU Munich, Geschwister-Scholl-Platz 1, 80539, Munich, Germany.

The nuclear pore complex (NPC) is one of the largest and most complex protein assemblies in the cell and, among other functions, serves as the gatekeeper of nucleocytoplasmic transport. Unraveling its molecular architecture and functioning has been an active research topic for decades with recent cryogenic electron microscopy and super-resolution studies advancing our understanding of the architecture of the NPC complex. However, the specific and direct visualization of single copies of NPC proteins is thus far elusive. Herein, we combine genetically-encoded self-labeling enzymes such as SNAP-tag and HaloTag with DNA-PAINT microscopy. We resolve single copies of nucleoporins in the human Y-complex in three dimensions with a precision of circa 3 nm, enabling studies of multicomponent complexes on the level of single proteins in cells using optical fluorescence microscopy.
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http://dx.doi.org/10.1002/anie.201905685DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6771475PMC
September 2019

Determining cellular CTCF and cohesin abundances to constrain 3D genome models.

Elife 2019 06 17;8. Epub 2019 Jun 17.

Department of Molecular and Cell Biology, Li Ka Shing Center for Biomedical and Health Sciences, CIRM Center of Excellence, University of California, Berkeley, Berkeley, United States.

Achieving a quantitative and predictive understanding of 3D genome architecture remains a major challenge, as it requires quantitative measurements of the key proteins involved. Here, we report the quantification of CTCF and cohesin, two causal regulators of topologically associating domains (TADs) in mammalian cells. Extending our previous imaging studies (Hansen et al., 2017), we estimate bounds on the density of putatively DNA loop-extruding cohesin complexes and CTCF binding site occupancy. Furthermore, co-immunoprecipitation studies of an endogenously tagged subunit (Rad21) suggest the presence of cohesin dimers and/or oligomers. Finally, based on our cell lines with accurately measured protein abundances, we report a method to conveniently determine the number of molecules of any Halo-tagged protein in the cell. We anticipate that our results and the established tool for measuring cellular protein abundances will advance a more quantitative understanding of 3D genome organization, and facilitate protein quantification, key to comprehend diverse biological processes.
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http://dx.doi.org/10.7554/eLife.40164DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6579579PMC
June 2019

Absolute quantification of cohesin, CTCF and their regulators in human cells.

Elife 2019 06 17;8. Epub 2019 Jun 17.

Research Institute of Molecular Pathology (IMP), Vienna Biocenter (VBC), Vienna, Austria.

The organisation of mammalian genomes into loops and topologically associating domains (TADs) contributes to chromatin structure, gene expression and recombination. TADs and many loops are formed by cohesin and positioned by CTCF. In proliferating cells, cohesin also mediates sister chromatid cohesion, which is essential for chromosome segregation. Current models of chromatin folding and cohesion are based on assumptions of how many cohesin and CTCF molecules organise the genome. Here we have measured absolute copy numbers and dynamics of cohesin, CTCF, NIPBL, WAPL and sororin by mass spectrometry, fluorescence-correlation spectroscopy and fluorescence recovery after photobleaching in HeLa cells. In G1-phase, there are ~250,000 nuclear cohesin complexes, of which ~ 160,000 are chromatin-bound. Comparison with chromatin immunoprecipitation-sequencing data implies that some genomic cohesin and CTCF enrichment sites are unoccupied in single cells at any one time. We discuss the implications of these findings for how cohesin can contribute to genome organisation and cohesion.
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http://dx.doi.org/10.7554/eLife.46269DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6606026PMC
June 2019

Mysteries in embryonic development: How can errors arise so frequently at the beginning of mammalian life?

PLoS Biol 2019 03 6;17(3):e3000173. Epub 2019 Mar 6.

Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany.

Chromosome segregation errors occur frequently during female meiosis but also in the first mitoses of mammalian preimplantation development. Such errors can lead to aneuploidy, spontaneous abortions, and birth defects. Some of the mechanisms underlying these errors in meiosis have been deciphered but which mechanisms could cause chromosome missegregation in the first embryonic cleavage divisions is mostly a "mystery". In this article, we describe the starting conditions and challenges of these preimplantation divisions, which might impair faithful chromosome segregation. We also highlight the pending research to provide detailed insight into the mechanisms and regulation of preimplantation mitoses.
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http://dx.doi.org/10.1371/journal.pbio.3000173DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6422315PMC
March 2019

Gain of CTCF-Anchored Chromatin Loops Marks the Exit from Naive Pluripotency.

Cell Syst 2018 11 7;7(5):482-495.e10. Epub 2018 Nov 7.

European Molecular Biology Laboratory (EMBL), Genome Biology Unit, Meyerhofstrasse 1, Heidelberg 69117, Germany. Electronic address:

The genome of pluripotent stem cells adopts a unique three-dimensional architecture featuring weakly condensed heterochromatin and large nucleosome-free regions. Yet, it is unknown whether structural loops and contact domains display characteristics that distinguish embryonic stem cells (ESCs) from differentiated cell types. We used genome-wide chromosome conformation capture and super-resolution imaging to determine nuclear organization in mouse ESC and neural stem cell (NSC) derivatives. We found that loss of pluripotency is accompanied by widespread gain of structural loops. This general architectural change correlates with enhanced binding of CTCF and cohesins and more pronounced insulation of contacts across chromatin boundaries in lineage-committed cells. Reprogramming NSCs to pluripotency restores the unique features of ESC domain topology. Domains defined by the anchors of loops established upon differentiation are enriched for developmental genes. Chromatin loop formation is a pervasive structural alteration to the genome that accompanies exit from pluripotency and delineates the spatial segregation of developmentally regulated genes.
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http://dx.doi.org/10.1016/j.cels.2018.09.003DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6327227PMC
November 2018

A call for public archives for biological image data.

Nat Methods 2018 11;15(11):849-854

European Molecular Biology Laboratory, European Bioinformatics Institute (EMBL-EBI), Wellcome Genome Campus, Cambridge, UK.

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http://dx.doi.org/10.1038/s41592-018-0195-8DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6884425PMC
November 2018

Multivariate Control of Transcript to Protein Variability in Single Mammalian Cells.

Cell Syst 2018 10 17;7(4):398-411.e6. Epub 2018 Oct 17.

Department of Molecular Life Sciences, University of Zurich, Zürich, Switzerland. Electronic address:

A long-standing question in quantitative biology is the relationship between mRNA and protein levels of the same gene. Here, we measured mRNA and protein abundance, the phenotypic state, and the population context in thousands of single human cells for 23 genes by combining a unique collection of cell lines with fluorescently tagged endogenous genomic loci and quantitative immunofluorescence with branched DNA single-molecule fluorescence in situ hybridization and computer vision. mRNA and protein abundance displayed a mean single-cell correlation of 0.732 at steady state. Single-cell outliers of linear correlations are in a specific phenotypic state or population context. This is particularly relevant for interpreting mRNA-protein relationships during acute gene induction and turnover, revealing a specific adaptation of gene expression at multiple steps in single cells. Together, we show that single-cell protein abundance can be predicted by multivariate information that integrates mRNA level with the phenotypic state and microenvironment of a particular cell.
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http://dx.doi.org/10.1016/j.cels.2018.09.001DOI Listing
October 2018

Experimental and computational framework for a dynamic protein atlas of human cell division.

Nature 2018 09 10;561(7723):411-415. Epub 2018 Sep 10.

European Molecular Biology Laboratory (EMBL), Heidelberg, Germany.

Essential biological functions, such as mitosis, require tight coordination of hundreds of proteins in space and time. Localization, the timing of interactions and changes in cellular structure are all crucial to ensure the correct assembly, function and regulation of protein complexes. Imaging of live cells can reveal protein distributions and dynamics but experimental and theoretical challenges have prevented the collection of quantitative data, which are necessary for the formulation of a model of mitosis that comprehensively integrates information and enables the analysis of the dynamic interactions between the molecular parts of the mitotic machinery within changing cellular boundaries. Here we generate a canonical model of the morphological changes during the mitotic progression of human cells on the basis of four-dimensional image data. We use this model to integrate dynamic three-dimensional concentration data of many fluorescently knocked-in mitotic proteins, imaged by fluorescence correlation spectroscopy-calibrated microscopy. The approach taken here to generate a dynamic protein atlas of human cell division is generic; it can be applied to systematically map and mine dynamic protein localization networks that drive cell division in different cell types, and can be conceptually transferred to other cellular functions.
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http://dx.doi.org/10.1038/s41586-018-0518-zDOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6556381PMC
September 2018

Modified aptamers enable quantitative sub-10-nm cellular DNA-PAINT imaging.

Nat Methods 2018 09 20;15(9):685-688. Epub 2018 Aug 20.

Department of Physics and Center for Nanoscience, Ludwig Maximilian University, Munich, Germany.

Although current implementations of super-resolution microscopy are technically approaching true molecular-scale resolution, this has not translated to imaging of biological specimens, because of the large size of conventional affinity reagents. Here we introduce slow off-rate modified aptamers (SOMAmers) as small and specific labeling reagents for use with DNA points accumulation in nanoscale topography (DNA-PAINT). To demonstrate the achievable resolution, specificity, and multiplexing capability of SOMAmers, we labeled and imaged both transmembrane and intracellular targets in fixed and live cells.
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http://dx.doi.org/10.1038/s41592-018-0105-0DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6345375PMC
September 2018

Dual-spindle formation in zygotes keeps parental genomes apart in early mammalian embryos.

Science 2018 07;361(6398):189-193

Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Meyerhofstrasse 1, 69117 Heidelberg, Germany.

At the beginning of mammalian life, the genetic material from each parent meets when the fertilized egg divides. It was previously thought that a single microtubule spindle is responsible for spatially combining the two genomes and then segregating them to create the two-cell embryo. We used light-sheet microscopy to show that two bipolar spindles form in the zygote and then independently congress the maternal and paternal genomes. These two spindles aligned their poles before anaphase but kept the parental genomes apart during the first cleavage. This spindle assembly mechanism provides a potential rationale for erroneous divisions into more than two blastomeric nuclei observed in mammalian zygotes and reveals the mechanism behind the observation that parental genomes occupy separate nuclear compartments in the two-cell embryo.
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http://dx.doi.org/10.1126/science.aar7462DOI Listing
July 2018

Quantitative live and super-resolution microscopy of mitotic chromosomes.

Methods Cell Biol 2018 24;145:65-90. Epub 2018 Apr 24.

European Molecular Biology Laboratory (EMBL), Cell Biology and Biophysics Unit, Heidelberg, Germany. Electronic address:

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http://dx.doi.org/10.1016/bs.mcb.2018.03.014DOI Listing
December 2018

Live imaging of cell division in preimplantation mouse embryos using inverted light-sheet microscopy.

Methods Cell Biol 2018 26;145:279-292. Epub 2018 Apr 26.

European Molecular Biology Laboratory (EMBL), Heidelberg, Germany. Electronic address:

Systematic studies of cell divisions at the beginning of mammalian life are of fundamental importance for our understanding of embryonic development and fertility. However, in the past the challenges of in vitro embryo culture and the embryo's pronounced light sensitivity have precluded a detailed investigation of preimplantation cell divisions. This protocol is based on recent technological breakthroughs in inverted light microscopy tailored for mouse embryology. Due to its reduced light dose, and therefore low phototoxicity, as well as higher acquisition speed, light-sheet microscopy allows extended 3D time-lapse imaging of early embryonic development with very high spatial and temporal resolution. This imaging approach enables imaging of key subcellular structures during the critical cell cycles from the zygote up to the blastocyst stage, with a resolution that allows automatic computational tracking and quantitative analysis of the dynamics of mitotic organelles.
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http://dx.doi.org/10.1016/bs.mcb.2018.03.030DOI Listing
December 2018

The replicative helicase MCM recruits cohesin acetyltransferase ESCO2 to mediate centromeric sister chromatid cohesion.

EMBO J 2018 08 21;37(15). Epub 2018 Jun 21.

Research Institute of Molecular Pathology, Vienna, Austria

Chromosome segregation depends on sister chromatid cohesion which is established by cohesin during DNA replication. Cohesive cohesin complexes become acetylated to prevent their precocious release by WAPL before cells have reached mitosis. To obtain insight into how DNA replication, cohesion establishment and cohesin acetylation are coordinated, we analysed the interaction partners of 55 human proteins implicated in these processes by mass spectrometry. This proteomic screen revealed that on chromatin the cohesin acetyltransferase ESCO2 associates with the MCM2-7 subcomplex of the replicative Cdc45-MCM-GINS helicase. The analysis of ESCO2 mutants defective in MCM binding indicates that these interactions are required for proper recruitment of ESCO2 to chromatin, cohesin acetylation during DNA replication, and centromeric cohesion. We propose that MCM binding enables ESCO2 to travel with replisomes to acetylate cohesive cohesin complexes in the vicinity of replication forks so that these complexes can be protected from precocious release by WAPL Our results also indicate that ESCO1 and ESCO2 have distinct functions in maintaining cohesion between chromosome arms and centromeres, respectively.
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http://dx.doi.org/10.15252/embj.201797150DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6068434PMC
August 2018

Quantitative mapping of fluorescently tagged cellular proteins using FCS-calibrated four-dimensional imaging.

Nat Protoc 2018 06 24;13(6):1445-1464. Epub 2018 May 24.

EMBL, Heidelberg, Germany.

The ability to tag a protein at its endogenous locus with a fluorescent protein (FP) enables quantitative understanding of protein dynamics at the physiological level. Genome-editing technology has now made this powerful approach routinely applicable to mammalian cells and many other model systems, thereby opening up the possibility to systematically and quantitatively map the cellular proteome in four dimensions. 3D time-lapse confocal microscopy (4D imaging) is an essential tool for investigating spatial and temporal protein dynamics; however, it lacks the required quantitative power to make the kind of absolute and comparable measurements required for systems analysis. In contrast, fluorescence correlation spectroscopy (FCS) provides quantitative proteomic and biophysical parameters such as protein concentration, hydrodynamic radius, and oligomerization but lacks the capability for high-throughput application in 4D spatial and temporal imaging. Here we present an automated experimental and computational workflow that integrates both methods and delivers quantitative 4D imaging data in high throughput. These data are processed to yield a calibration curve relating the fluorescence intensities (FIs) of image voxels to the absolute protein abundance. The calibration curve allows the conversion of the arbitrary FIs to protein amounts for all voxels of 4D imaging stacks. Using our workflow, users can acquire and analyze hundreds of FCS-calibrated image series to map their proteins of interest in four dimensions. Compared with other protocols, the current protocol does not require additional calibration standards and provides an automated acquisition pipeline for FCS and imaging data. The protocol can be completed in 1 d.
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http://dx.doi.org/10.1038/nprot.2018.040DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6609853PMC
June 2018

Generation and validation of homozygous fluorescent knock-in cells using CRISPR-Cas9 genome editing.

Nat Protoc 2018 06 24;13(6):1465-1487. Epub 2018 May 24.

EMBL, Heidelberg, Germany.

Gene tagging with fluorescent proteins is essential for investigations of the dynamic properties of cellular proteins. CRISPR-Cas9 technology is a powerful tool for inserting fluorescent markers into all alleles of the gene of interest (GOI) and allows functionality and physiological expression of the fusion protein. It is essential to evaluate such genome-edited cell lines carefully in order to preclude off-target effects caused by (i) incorrect insertion of the fluorescent protein, (ii) perturbation of the fusion protein by the fluorescent proteins or (iii) nonspecific genomic DNA damage by CRISPR-Cas9. In this protocol, we provide a step-by-step description of our systematic pipeline to generate and validate homozygous fluorescent knock-in cell lines.We have used the paired Cas9D10A nickase approach to efficiently insert tags into specific genomic loci via homology-directed repair (HDR) with minimal off-target effects. It is time-consuming and costly to perform whole-genome sequencing of each cell clone to check for spontaneous genetic variations occurring in mammalian cell lines. Therefore, we have developed an efficient validation pipeline of the generated cell lines consisting of junction PCR, Southern blotting analysis, Sanger sequencing, microscopy, western blotting analysis and live-cell imaging for cell-cycle dynamics. This protocol takes between 6 and 9 weeks. With this protocol, up to 70% of the targeted genes can be tagged homozygously with fluorescent proteins, thus resulting in physiological levels and phenotypically functional expression of the fusion proteins.
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http://dx.doi.org/10.1038/nprot.2018.042DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6556379PMC
June 2018

A quantitative map of human Condensins provides new insights into mitotic chromosome architecture.

J Cell Biol 2018 07 9;217(7):2309-2328. Epub 2018 Apr 9.

Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany

The two Condensin complexes in human cells are essential for mitotic chromosome structure. We used homozygous genome editing to fluorescently tag Condensin I and II subunits and mapped their absolute abundance, spacing, and dynamic localization during mitosis by fluorescence correlation spectroscopy (FSC)-calibrated live-cell imaging and superresolution microscopy. Although ∼35,000 Condensin II complexes are stably bound to chromosomes throughout mitosis, ∼195,000 Condensin I complexes dynamically bind in two steps: prometaphase and early anaphase. The two Condensins rarely colocalize at the chromatid axis, where Condensin II is centrally confined, but Condensin I reaches ∼50% of the chromatid diameter from its center. Based on our comprehensive quantitative data, we propose a three-step hierarchical loop model of mitotic chromosome compaction: Condensin II initially fixes loops of a maximum size of ∼450 kb at the chromatid axis, whose size is then reduced by Condensin I binding to ∼90 kb in prometaphase and ∼70 kb in anaphase, achieving maximum chromosome compaction upon sister chromatid segregation.
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http://dx.doi.org/10.1083/jcb.201801048DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6028534PMC
July 2018

Real-time 3D single-molecule localization using experimental point spread functions.

Nat Methods 2018 05 9;15(5):367-369. Epub 2018 Apr 9.

Cell Biology and Biophysics Unit, European Molecular Biology Laboratory (EMBL), Heidelberg, Germany.

We present a real-time fitter for 3D single-molecule localization microscopy using experimental point spread functions (PSFs) that achieves minimal uncertainty in 3D on any microscope and is compatible with any PSF engineering approach. We used this method to image cellular structures and attained unprecedented image quality for astigmatic PSFs. The fitter compensates for most optical aberrations and makes accurate 3D super-resolution microscopy broadly accessible, even on standard microscopes without dedicated 3D optics.
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http://dx.doi.org/10.1038/nmeth.4661DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC6009849PMC
May 2018

Correlative live and super-resolution imaging reveals the dynamic structure of replication domains.

J Cell Biol 2018 06 23;217(6):1973-1984. Epub 2018 Mar 23.

Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany

Chromosome organization in higher eukaryotes controls gene expression, DNA replication, and DNA repair. Genome mapping has revealed the functional units of chromatin at the submegabase scale as self-interacting regions called topologically associating domains (TADs) and showed they correspond to replication domains (RDs). A quantitative structural and dynamic description of RD behavior in the nucleus is, however, missing because visualization of dynamic subdiffraction-sized RDs remains challenging. Using fluorescence labeling of RDs combined with correlative live and super-resolution microscopy in situ, we determined biophysical parameters to characterize the internal organization, spacing, and mechanical coupling of RDs. We found that RDs are typically 150 nm in size and contain four co-replicating regions spaced 60 nm apart. Spatially neighboring RDs are spaced 300 nm apart and connected by highly flexible linker regions that couple their motion only <550 nm. Our pipeline allows a robust quantitative characterization of chromosome structure in situ and provides important biophysical parameters to understand general principles of chromatin organization.
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http://dx.doi.org/10.1083/jcb.201709074DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC5987722PMC
June 2018

ChromoTrace: Computational reconstruction of 3D chromosome configurations for super-resolution microscopy.

PLoS Comput Biol 2018 03 9;14(3):e1006002. Epub 2018 Mar 9.

European Molecular Biology Laboratory, European Bioinformatics Institute (EMBL-EBI), Wellcome Genome Campus, Hinxton, Cambridge, United Kingdom.

The 3D structure of chromatin plays a key role in genome function, including gene expression, DNA replication, chromosome segregation, and DNA repair. Furthermore the location of genomic loci within the nucleus, especially relative to each other and nuclear structures such as the nuclear envelope and nuclear bodies strongly correlates with aspects of function such as gene expression. Therefore, determining the 3D position of the 6 billion DNA base pairs in each of the 23 chromosomes inside the nucleus of a human cell is a central challenge of biology. Recent advances of super-resolution microscopy in principle enable the mapping of specific molecular features with nanometer precision inside cells. Combined with highly specific, sensitive and multiplexed fluorescence labeling of DNA sequences this opens up the possibility of mapping the 3D path of the genome sequence in situ. Here we develop computational methodologies to reconstruct the sequence configuration of all human chromosomes in the nucleus from a super-resolution image of a set of fluorescent in situ probes hybridized to the genome in a cell. To test our approach, we develop a method for the simulation of DNA in an idealized human nucleus. Our reconstruction method, ChromoTrace, uses suffix trees to assign a known linear ordering of in situ probes on the genome to an unknown set of 3D in-situ probe positions in the nucleus from super-resolved images using the known genomic probe spacing as a set of physical distance constraints between probes. We find that ChromoTrace can assign the 3D positions of the majority of loci with high accuracy and reasonable sensitivity to specific genome sequences. By simulating appropriate spatial resolution, label multiplexing and noise scenarios we assess our algorithms performance. Our study shows that it is feasible to achieve genome-wide reconstruction of the 3D DNA path based on super-resolution microscopy images.
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http://dx.doi.org/10.1371/journal.pcbi.1006002DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC5862484PMC
March 2018

Author Correction: Sister chromatid resolution is an intrinsic part of chromosome organization in prophase.

Nat Cell Biol 2018 04;20(4):503

Cancer Institute of the Japanese Foundation for Cancer Research, Division of Experimental Pathology, 3-8-31 Ariake, Koto-ku, Tokyo, 135-8550, Japan.

In the version of this Letter originally published, the authors omitted a citation of an early study demonstrating topoisomerase-II-dependent sister chromatid resolution. This reference has now been added to the reference list as reference number 28, and the relevant text has been amended as follows to include its citation: 'Resolution must reflect the removal of sister-sister contacts, and we show here that Topo-IIα-mediated release of DNA catenation plays a major role (Fig. 4), in agreement with previous findings, whereas, surprisingly, cohesin dissociation is not strictly required (Fig. 3).' Subsequent references have been renumbered. All online versions of the Letter have been updated to reflect this change.
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http://dx.doi.org/10.1038/s41556-018-0044-0DOI Listing
April 2018

Postmitotic nuclear pore assembly proceeds by radial dilation of small membrane openings.

Nat Struct Mol Biol 2018 01 27;25(1):21-28. Epub 2017 Nov 27.

Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany.

The nuclear envelope has to be reformed after mitosis to create viable daughter cells with closed nuclei. How membrane sealing of DNA and assembly of nuclear pore complexes (NPCs) are achieved and coordinated is poorly understood. Here, we reconstructed nuclear membrane topology and the structures of assembling NPCs in a correlative 3D EM time course of dividing human cells. Our quantitative ultrastructural analysis shows that nuclear membranes form from highly fenestrated ER sheets whose holes progressively shrink. NPC precursors are found in small membrane holes and dilate radially during assembly of the inner ring complex, forming thousands of transport channels within minutes. This mechanism is fundamentally different from that of interphase NPC assembly and explains how mitotic cells can rapidly establish a closed nuclear compartment while making it transport competent.
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http://dx.doi.org/10.1038/s41594-017-0001-9DOI Listing
January 2018

Topologically associating domains and chromatin loops depend on cohesin and are regulated by CTCF, WAPL, and PDS5 proteins.

EMBO J 2017 12 7;36(24):3573-3599. Epub 2017 Dec 7.

Research Institute of Molecular Pathology (IMP), Vienna Biocenter (VBC), Vienna, Austria

Mammalian genomes are spatially organized into compartments, topologically associating domains (TADs), and loops to facilitate gene regulation and other chromosomal functions. How compartments, TADs, and loops are generated is unknown. It has been proposed that cohesin forms TADs and loops by extruding chromatin loops until it encounters CTCF, but direct evidence for this hypothesis is missing. Here, we show that cohesin suppresses compartments but is required for TADs and loops, that CTCF defines their boundaries, and that the cohesin unloading factor WAPL and its PDS5 binding partners control the length of loops. In the absence of WAPL and PDS5 proteins, cohesin forms extended loops, presumably by passing CTCF sites, accumulates in axial chromosomal positions (vermicelli), and condenses chromosomes. Unexpectedly, PDS5 proteins are also required for boundary function. These results show that cohesin has an essential genome-wide function in mediating long-range chromatin interactions and support the hypothesis that cohesin creates these by loop extrusion, until it is delayed by CTCF in a manner dependent on PDS5 proteins, or until it is released from DNA by WAPL.
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http://dx.doi.org/10.15252/embj.201798004DOI Listing
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC5730888PMC
December 2017