Application of Fluorescence Lifetime Imaging Microscopy of DNA Binding Dyes to Assess Radiation-Induced Chromatin Compaction Changes

Abdollahi, Elham; Taucher-Scholz, G.; Jakob, Burkhard

doi:10.3390/ijms19082399

Cited by 15 publications

(16 citation statements)

References 63 publications

(114 reference statements)

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“…Local spikes in the count rate at detection rates below 10% were reported to affect the measured lifetime and require mathematical corrections ( 33 ). To test whether such a correction should be applied to our measurements, we used the dataset where we detected a correlation between count rate and FRET efficiency (ALC1 + TRIM33; Supplementary Figure S3G ).…”

Section: Resultsmentioning

confidence: 99%

“…The sample size ( N ) for each experiment is listed in Supplementary Table S1 along with median values, 25th and 75th percentiles, and mean and confidence intervals. Detection rate-dependent correction of lifetimes (only for data in Supplementary Figure S3G and I ) was performed based on τ corr ≈ τ /(1 − P /4), where τ is the measured lifetime and P is the average number of photons in a given laser period ( 33 ). In our case, the laser repetition rate was 40 MHz such that a laser period amounts to 25 ns.…”

Section: Methodsmentioning

confidence: 99%

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Direct measurement of protein–protein interactions by FLIM-FRET at UV laser-induced DNA damage sites in living cells

Kaufmann

Herbert

Hackl

et al. 2020

Nucleic Acids Research

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Protein–protein interactions are essential to ensure timely and precise recruitment of chromatin remodellers and repair factors to DNA damage sites. Conventional analyses of protein–protein interactions at a population level may mask the complexity of interaction dynamics, highlighting the need for a method that enables quantification of DNA damage-dependent interactions at a single-cell level. To this end, we integrated a pulsed UV laser on a confocal fluorescence lifetime imaging (FLIM) microscope to induce localized DNA damage. To quantify protein–protein interactions in live cells, we measured Förster resonance energy transfer (FRET) between mEGFP- and mCherry-tagged proteins, based on the fluorescence lifetime reduction of the mEGFP donor protein. The UV-FLIM-FRET system offers a unique combination of real-time and single-cell quantification of DNA damage-dependent interactions, and can distinguish between direct protein–protein interactions, as opposed to those mediated by chromatin proximity. Using the UV-FLIM-FRET system, we show the dynamic changes in the interaction between poly(ADP-ribose) polymerase 1, amplified in liver cancer 1, X-ray repair cross-complementing protein 1 and tripartite motif containing 33 after DNA damage. This new set-up complements the toolset for studying DNA damage response by providing single-cell quantitative and dynamic information about protein–protein interactions at DNA damage sites.

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Section: Resultsmentioning

confidence: 99%

Section: Methodsmentioning

confidence: 99%

Direct measurement of protein–protein interactions by FLIM-FRET at UV laser-induced DNA damage sites in living cells

Kaufmann

Herbert

Hackl

et al. 2020

Nucleic Acids Research

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“…Roughly a fraction of half of the dose can be considered to be delivered to the track centre and penumbra, respectively. Cell irradiation with X-Ray was performed as described before 57,58 . Briefly, a 35 kV X-ray tube (GE Inspection Technology, Ahrensburg, Germany) which was operated at 80 mA delivering a dose rate of 0.85 Gy/s at the cell layer was used.…”

Section: Methodsmentioning

confidence: 99%

Differential Repair Protein Recruitment at Sites of Clustered and Isolated DNA Double-Strand Breaks Produced by High-Energy Heavy Ions

Jakob

Dubiak-Szepietowska

Janiel

et al. 2020

Sci Rep

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DNA double-strand break (DSB) repair is crucial to maintain genomic stability. The fidelity of the repair depends on the complexity of the lesion, with clustered DSBs being more difficult to repair than isolated breaks. Using live cell imaging of heavy ion tracks produced at a high-energy particle accelerator we visualised simultaneously the recruitment of different proteins at individual sites of complex and simple DSBs in human cells. NBS1 and 53BP1 were recruited in a few seconds to complex DSBs, but in 40% of the isolated DSBs the recruitment was delayed approximately 5 min. Using base excision repair (BER) inhibitors we demonstrate that some simple DSBs are generated by enzymatic processing of base damage, while BER did not affect the complex DSBs. The results show that DSB processing and repair kinetics are dependent on the complexity of the breaks and can be different even for the same clastogenic agent. Repair of DNA double-strand breaks is a necessary pathway to maintain genomic stability in mammalian cells 1. The DNA damage response (DDR) signalling cascade is rapid and hierarchically coordinated, with many proteins being recruited to the damage sites at different times and depending on the repair pathway. Complex DSBs, i.e. complex lesions involving multiple strand breaks and/or oxidative damages within two helical turns 2,3 , are more difficult to repair than isolated, frank DSBs 4. Complex DSBs can be produced by ionising radiation (IR) 3,5 , and their fraction and complexity and clustering increases for densely ionising, high-linear energy transfer (LET) radiation such as α-particles and heavy ions 6. Endogenous DSBs (simple DSBs produced during replication) have much higher frequency than IR-induced DSBs at low doses, but the latter include complex DSBs 7. These complex lesions are considered ultimately responsible for the late effects of low doses of IR 8 , including environmental exposures and cosmic radiation risk in space travel. The repair of simple and complex DSBs is therefore crucial to understand the difference between endogenous and exogenous genotoxicity and for modelling the risk related to exposure to low doses IR on Earth 9 and in space 10. Delayed repair of clustered DSBs has been observed by immunostaining of markers of single strand breaks (SSBs), DSBs and base damage in mammalian cells 11. Here we measured the early protein recruitment at sites of simple and clustered, complex DSBs by live cell imaging. Immunostaining on fixed samples is unable to identify the early kinetics and cannot follow the evolution of individual foci, therefore providing only average values 12. High-charge and-energy (HZE) ions offer a unique opportunity for these studies as they simultaneously produce both clustered (along the track) and isolated (off-track) DSBs. In fact, the inner part of the ion track (core) includes primary particle ionizations and low-energy electrons, and results mostly in complex, clustered DSBs for HZE ions at high linear energy transfer (LET), whereas low-LET high-energy ionised ...

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“…In addition, a phasor-based approach to FLIM-FRET analysis of chromatin compaction has been described (Lou et al, 2019). DNA-staining dyes and fluorescent protein reporters have also been used for FLIM analysis of events in the nucleus (Abdollahi et al, 2018;Audugé et al, 2019;Estandarte et al, 2016;Kawanabe et al, 2015;Okkelman et al, 2016;Sparks et al, 2018).…”

Section: Flim For Live Subcellular Imaging In 3dmentioning

confidence: 99%

Luminescence lifetime imaging of three-dimensional biological objects

Dmitriev

Intes

Barroso

2021

Journal of Cell Science

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A major focus of current biological studies is to fill the knowledge gaps between cell, tissue and organism scales. To this end, a wide array of contemporary optical analytical tools enable multiparameter quantitative imaging of live and fixed cells, three-dimensional (3D) systems, tissues, organs and organisms in the context of their complex spatiotemporal biological and molecular features. In particular, the modalities of luminescence lifetime imaging, comprising fluorescence lifetime imaging (FLI) and phosphorescence lifetime imaging microscopy (PLIM), in synergy with Förster resonance energy transfer (FRET) assays, provide a wealth of information. On the application side, the luminescence lifetime of endogenous molecules inside cells and tissues, overexpressed fluorescent protein fusion biosensor constructs or probes delivered externally provide molecular insights at multiple scales into protein–protein interaction networks, cellular metabolism, dynamics of molecular oxygen and hypoxia, physiologically important ions, and other physical and physiological parameters. Luminescence lifetime imaging offers a unique window into the physiological and structural environment of cells and tissues, enabling a new level of functional and molecular analysis in addition to providing 3D spatially resolved and longitudinal measurements that can range from microscopic to macroscopic scale. We provide an overview of luminescence lifetime imaging and summarize key biological applications from cells and tissues to organisms.

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Application of Fluorescence Lifetime Imaging Microscopy of DNA Binding Dyes to Assess Radiation-Induced Chromatin Compaction Changes

Cited by 15 publications

References 63 publications

Direct measurement of protein–protein interactions by FLIM-FRET at UV laser-induced DNA damage sites in living cells

Direct measurement of protein–protein interactions by FLIM-FRET at UV laser-induced DNA damage sites in living cells

Differential Repair Protein Recruitment at Sites of Clustered and Isolated DNA Double-Strand Breaks Produced by High-Energy Heavy Ions

Luminescence lifetime imaging of three-dimensional biological objects

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