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Octopus (Optics Clustered to OutPut Unique Solutions) celebrated in June 2020 its 10th birthday. Based at Harwell, near Oxford, Octopus is an open access, peer reviewed, national imaging facility that offers successful U.K. applicants supported access to single molecule imaging, confocal microscopy, several flavours of superresolution imaging, light sheet microscopy, optical trapping and cryoscanning electron microscopy. Managed by a multidisciplinary team, Octopus has so far assisted >100 groups of U.K. and international researchers. Cross-fertilisation across fields proved to be a strong propeller of success underpinned by combining access to top-end instrumentation with a strong programme of imaging hardware and software developments. How Octopus was born, and highlights of the multidisciplinary output produced during its 10-year journey are reviewed below, with the aim of celebrating a myriad of collaborations with the U.K. scientific community, and reflecting on their scientific and societal impact.

On the plasma membrane, Ras is organized into laterally segregated proteo-lipid complexes called nanoclusters. The extent of Ras nanoclustering correlates with its signaling output, positioning nanocluster as dynamic signaling gain modulators. Recent evidence suggests that stacked dimers of Ras and Raf are elemental units at least of one type of Ras nanocluster. However, it is still incompletely understood, in which physiological contexts nanoclustering is regulated and which constituents are parts of nanocluster. Nonetheless, disruption of nanoclustering faithfully diminishes Ras activity in cells, suggesting Ras nanocluster as potential drug targets.While there are several methods available to study Ras nanocluster , fluorescence or Forster resonance energy transfer (FRET ) between fluorescently labeled, nanoclustered Ras proteins is a relatively simple readout. FRET measurements using fluorescence lifetime imaging microscopy (FLIM ) have proven to be robust and sensitive to determine Ras nanoclustering changes. Loss of FRET that emerges due to nanoclustering reports on all processes upstream of Ras nanoclustering, i.e., also on proper trafficking or lipid modification of Ras. Here we report our standard FLIM-FRET protocol to measure nanoclustering-dependent FRET of Ras in mammalian cells. Importantly, nanoclustering-dependent FRET is one of the few methods that can detect differences between the Ras isoforms.

Protein-protein interactions (PPIs) control various key processes in cells. Fluorescence lifetime imaging microscopy (FLIM) combined with Forster resonance energy transfer (FRET) provide accurate information about PPIs in live cells. FLIM-FRET relies on measuring the fluorescence lifetime decay of a FRET donor at each pixel of the FLIM image, providing quantitative and accurate information about PPIs and their spatial cellular organizations. We propose here a detailed protocol for FLIM-FRET measurements that we applied to monitor PPIs in live Pseudomonas aeruginosa in the particular case of two interacting proteins expressed with highly different copy numbers to demonstrate the quality and robustness of the technique at revealing critical features of PPIs. This protocol describes in detail all the necessary steps for PPI characterization - starting from bacterial mutant constructions up to the final analysis using recently developed tools providing advanced visualization possibilities for a straightforward interpretation of complex FLIM-FRET data.

The nuclear factor-erythroid 2 p45-related factor 2 (Nrf2)-mediated stress response is a major cellular defense mechanism against endogenous and exogenous oxidants, electrophiles, and pro-inflammatory agents. A number of Nrf2 inducers are being developed to therapeutically stimulate this pathway. Inducers are typically sensed by Kelch-like ECH-associated protein 1 (Keap1), a negative regulator and a binding partner of Nrf2. Modifications of Keap1 by oxidants or electrophiles, or its targeting by compounds that disrupt its interaction with Nrf2, alter the conformation of the Keap1-Nrf2 protein complex, which initiates the accumulation of Nrf2 required for mounting a stress response. To detect conformational changes in the Keap1-Nrf2 complex in live cells, we have developed a procedure based on Fluorescence Lifetime Imaging-Förster Resonance Energy Transfer (FLIM-FRET). The procedure includes a FLIM time course in cells expressing fluorescently-tagged Nrf2 and Keap1, followed by an extended analysis pipeline that quantifies changes in fluorescence lifetime of labeled Nrf2. The analysis visualizes and removes intensity-dependent bias in fluorescence lifetime measured with the Time-Correlated Single Photon Counting (TCSPC) approach, thereby improving the accuracy of quantification. The throughput is increased by the whole-experiment analysis within the newly developed FLIM dataset tool (FLIMDAST) and by the time-lapse FLIM described here. This pipeline is also suitable for applications beyond the Nrf2 field that assess small changes in fluorescence lifetime of objects with variable fluorescence intensities measured using TCSPC-based FLIM. © 2020 The Authors. Basic Protocol 1: Lipofectamine 2000 transfection Alternate Protocol 1: Calcium phosphate transfection Basic Protocol 2: Time course with individual FLIM Alternate Protocol 2: Time course with time-lapse FLIM Support Protocol: Measuring Instrument Response Function (IRF) Basic Protocol 3: Data analysis in SPCImage Basic Protocol 4: Data processing in ImageJ/FIJI Basic Protocol 5: Experiment analysis in FLIMDAST.

Significance Chromatin dynamics play a central role in the DNA damage response (DDR). A longstanding experimental obstacle was the lack of technology capable of visualizing chromatin dynamics at double-strand break (DSB) sites. Here, we describe biophysical methods that quantify spatiotemporal chromatin compaction dynamics in living cells. Using these tools, we investigate dynamic remodeling of chromatin architecture at DSB loci and find that chromatin “opening” and “compacting” events facilitate repair factor access while demarcating the lesion from the surrounding nuclear environment. Furthermore, we identify regulatory roles for key DDR enzymes in this process. Finally, we demonstrate method utility through physical, pharmacological, and genetic manipulation of the chromatin environment, which collectively demonstrates the potential for its use in future studies of chromatin biology., To investigate how chromatin architecture is spatiotemporally organized at a double-strand break (DSB) repair locus, we established a biophysical method to quantify chromatin compaction at the nucleosome level during the DNA damage response (DDR). The method is based on phasor image-correlation spectroscopy of histone fluorescence lifetime imaging microscopy (FLIM)-Förster resonance energy transfer (FRET) microscopy data acquired in live cells coexpressing H2B-eGFP and H2B-mCherry. This multiplexed approach generates spatiotemporal maps of nuclear-wide chromatin compaction that, when coupled with laser microirradiation-induced DSBs, quantify the size, stability, and spacing between compact chromatin foci throughout the DDR. Using this technology, we identify that ataxia–telangiectasia mutated (ATM) and RNF8 regulate rapid chromatin decompaction at DSBs and formation of compact chromatin foci surrounding the repair locus. This chromatin architecture serves to demarcate the repair locus from the surrounding nuclear environment and modulate 53BP1 mobility.

To overcome limitations of indirect immunofluorescence, a new method is presented to employ the ostensible disadvantage of cross-labeling secondary antibodies by separation of the fluorescence signals via spectral FLIM-FRET. The undesirable cross-labeling among secondary antibodies leads to the generation of new characteristic FRET emission spectra including a change in the donor lifetime. We used a spectrally resolved FLIM detection system with pulse interleaved excitation. The combined spectral FLIM-FRET and pattern-matching analysis forms an excellent tool for use in indirect immunofluorescence that overcomes the undesirable effect of secondary antibody cross-labeling by assigning separate color channels to cross-labeled fluorescent antibodies.

Förster resonance energy transfer (FRET)-based sensors have been powerful tools in cell biologists’ toolkit for decades. Informed by fundamental understanding of fluorescent proteins, protein-protein interactions, and the structural biology of reporter components, researchers have been able to employ creative design approaches to build sensors that are uniquely capable of probing a wide range of phenomena in living cells including visualization of localized calcium signaling, sub-cellular activity gradients, and tension generation to name but a few. While FRET sensors have significantly impacted many fields, one must also be cognizant of the limitations to conventional, intensity-based FRET measurements stemming from variation in probe concentration, sensitivity to photo-bleaching, and bleed-through between the FRET fluorophores. Fluorescence lifetime imaging microscopy (FLIM) largely overcomes the limitations of intensity based FRET measurements. In general terms, FLIM measures the time, which for the reporters described in this chapter is nanoseconds (ns), between photon absorption and emission by a fluorophore. When FLIM is applied to FRET sensors (FLIM-FRET), measurement of the donor fluorophore lifetime provides valuable information such as FRET efficiency and the percentage of reporters engaged in FRET. This chapter introduces fundamental principles of FLIM-FRET towards informing the practical application of the technique and, using two established FRET reporters as proofs of concept, outlines how to use a commercially available FLIM system.

Förster Resonance Energy Transfer (FRET) has become an immensely powerful tool to profile intra- and inter-molecular interactions. Through fusion of genetically encoded fluorescent proteins (FPs) researchers have been able to detect protein oligomerization, receptor activation, and protein translocation among other biophysical phenomena. Recently, two bright monomeric red fluorescent proteins, mRuby3 and mScarlet-I, have been developed. These proteins offer much improved physical properties compared to previous generations of monomeric red FPs that should help facilitate more general adoption of Green/Red FRET. Here we assess the ability of these two proteins, along with mCherry, to act as a FRET acceptor for the bright, monomeric, green-yellow FP mNeonGreen using intensiometric FRET and 2-photon Fluorescent Lifetime Imaging Microscopy (FLIM) FRET techniques. We first determined that mNeonGreen was a stable donor for 2-photon FLIM experiments under a variety of imaging conditions. We then tested the red FP’s ability to act as FRET acceptors using mNeonGreen-Red FP tandem construct. With these constructs we found that mScarlet-I and mCherry are able to efficiently FRET with mNeonGreen in spectroscopic and FLIM FRET. In contrast, mNeonGreen and mRuby3 FRET with a much lower efficiency than predicted in these same assays. We explore possible explanations for this poor performance and determine mRuby3’s protein maturation properties are a major contributor. Overall, we find that mNeonGreen is an excellent FRET donor, and both mCherry and mScarlet-I, but not mRuby3, act as practical FRET acceptors, with the brighter mScarlet-I out performing mCherry in intensiometric studies, but mCherry out performing mScarlet-I in instances where consistent efficiency in a population is critical.

This protocol was designed to quantitatively measure small-molecule displacement of proteins in live mammalian cells using fluorescence lifetime imaging microscopy-Forster resonance energy transfer (FLIM-FRET). Tumour cell survival is often dependent on anti-apoptotic proteins, which bind to and inhibit pro-apoptotic proteins, thus preventing apoptosis. Small-molecule inhibitors that selectively target these proteins (termed BH3-mimetics) are therefore a promising avenue for the treatment of several cancers. Previous techniques used to study the efficacy of these drugs often use truncated versions of both pro- and anti-apoptotic proteins, as they are membrane bound and hydrophobic in nature. As a result, the true efficacy of these drugs to displace full-length pro-apoptotic proteins in their native environment within a cell is poorly understood. This protocol describes FLIM-FRET methods to directly measure the displacement (or lack of displacement) of full-length Bcl-2 family proteins in live mammalian cells.

The Bcl-2 proteins control cell death via interchanging interactions within the Bcl-2 family. Fluorescence lifetime imaging microscopy (FLIM) is used to detect Forster resonance energy transfer (FRET) between two fluorescent-fusion proteins in live cells. FLIM-FRET has been used to detect specific interactions and their disruption, for Bcl-2 family proteins. To date, this has been possible only in low throughput, due to the time required for serial data acquisition. We developed an automated optical system with eight parallel detectors for rapid and efficient data collection. Here we describe how to use this system for FLIM-FRET imaging of Bcl-2 family protein interactions in a 384-well plate format.

Optical imaging assays, especially fluorescence molecular assays, are minimally invasive if not completely noninvasive, and thus an ideal technique to be applied to live specimens. These fluorescence imaging assays are a powerful tool in biomedical sciences as they allow the study of a wide range of molecular and physiological events occurring in biological systems. Furthermore, optical imaging assays bridge the gap between the in vitro cell-based analysis of subcellular processes and in vivo study of disease mechanisms in small animal models. In particular, the application of Förster resonance energy transfer (FRET) and fluorescence lifetime imaging (FLIM), well-known techniques widely used in microscopy, to the optical imaging assay toolbox, will have a significant impact in the molecular study of protein-protein interactions during cancer progression. This review article describes the application of FLIM-FRET to the field of optical imaging and addresses their various applications, both current and potential, to anti-cancer drug delivery and cancer research.

The fundamental theory of Forster resonance energy transfer (FRET) was established in the 1940s. Its great power was only realized in the past 20 years after different techniques were developed and applied to biological experiments. This success was made possible by the availability of suitable fluorescent probes, advanced optics, detectors, microscopy instrumentation, and analytical tools. Combined with state-of-the-art microscopy and spectroscopy, FRET imaging allows scientists to study a variety of phenomena that produce changes in molecular proximity, thereby leading to many significant findings in the life sciences. In this review, we outline various FRET imaging techniques and their strengths and limitations; we also provide a biological model to demonstrate how to investigate protein-protein interactions in living cells using both intensity- and fluorescence lifetime-based FRET microscopy methods.

Fluorescence imaging techniques are extremely powerful tools in cell biology, providing valuable insights into the structure and function of biomolecules in their native environments. In particular, the use of Forster resonance energy transfer (FRET) has become increasingly important to obtain information on interactions on the nanoscale, in turn providing insights into molecular behaviour inside living cells. This review describes the basic principles of FRET and fluorescence lifetime imaging microscopy (FLIM) and their application in analyses of protein interactions inside living fungal cells.

Average lifetime between the usually bi-exponential double -label specimen and a mono -exponential single donor sample serves as a basis for the calculation of the average energy transfer efficiency (E ). T his semi-quantitative approach however does not fully explore cellular functions, such as endosomal pH differences, specific morphological features, ex amin ing sub-po pulations and the like. W e applied a different, quantitative Regi on -of-Interest (ROI)-based method in 2 live -cell assays by TCSPC FLIM-FRET microscopy: a 5 amino-acid linked FRET standard and mouse pituitary cells expressing a GLPHUL]HG& (%3. -bZip transcription factor in the nucleus , both tagged with Cerulean (C) and Venus (V). ROIs with different selection thresholds were generated and compared. Average lifetimes are similar , but ratios between them and other subtle differences are revealed by compreh ensive distribution information. Following published references, we also explored 3 different methods to calculate FLIM-FRET energy transfer efficiencies for the Cerulean-Venus constructs, producing differences and supporting the long-held notion that is called 'apparent' E . efficiencyFRET's greatest contribution es to be exploringcontinu changes taking place at the cellular level and quantifying differences in relative terms between control and variables .

Fluorescence lifetime imaging microscopy-Forster resonant energy transfer (FLIM-FRET) is a high-resolution technique for the detection of protein interactions in live cells. As the cost of this technology becomes more competitive and methods are devised to extract more information from the FLIM images, this technique will be increasingly useful for studying protein interactions in live cells. Here we demonstrate the use of the ISS-Alba FLIM/FCS confocal microscope, which was custom-built for supervised automation of FLIM data acquisition. We provide a detailed protocol for collecting and analyzing good FLIM-FRET data. As an example, we use FLIM-FRET to detect the interaction between BclXL and Bad at the mitochondrial outer membrane in live MCF7 breast cancer cells.

The number of publications using FLIM/FRET (Fluorescence lifetime imaging and Forster resonance energy transfer) technique is increasing over the past 10 years (Vogel et al. Sci STKE 18:re2, 2006) [1].

Fluorescence & Other Luminescence II 3496-Pos Board B651 Subcellular Imaging of Protein-Protein Interactions in Live Epithelial Cells using Single Particle Tracking-FLIM-FRET Luca Lanzano 1 , Hector Giral 2 , Moshe Levi 2 , Enrico Gratton 1 . University California, Irvine, CA, USA, 2 University Colorado, Denver, CO, USA. Phosphate (Pi) homeostasis is crucial for many cellular processes. Pi absorption in the kidney proximal tubule and in the intestine is mainly mediated by the type II family of the sodium-dependent phosphate co-transporters (NaPi) which are located in the apical membrane microvilli. Interaction with scaffolding PDZ domain containing proteins is thought to play a central role in the regulation of the abundance and activity of the NaPi transporters, in response to dietary and hormonal stimuli. We investigated protein-protein interactions between NaPi and PDZ proteins in cellular models of the proximal tubule and the intestine using the FLIM detection of FRET [1]. Also, we have recently introduced the nSPIRO method which generalizes the principles of Single Particle Tracking to the imaging of extended subcellular structures like the apical membrane microvilli [2]. using this method we can image the microvilli with high resolution even if they are moving. Here we combine the tracking of single microvilli with the phasor analysis of FLIM to quantify FRET interaction at the level of the single microvilli. We explore how the measurement of FRET on single microvilli compares to the measurement performed on the entire apical membrane, and discuss the advan- tages and limitations of this technique. Work supported in part by NIH RO1 DK066029-01A2 (ML, EG), 8P41GM103540 (EG and LL) and P50GM076516 (EG and LL). [1] Giral, Lanzano et al, J Biol Chem (2011) 286, 15032-15042; Giral, Cranston, Lanzano et al, J Biol Chem (2012) epub ahead of print [2] Lanzano L et al, J Biophotonics (2011) 4, 415-42; Lanzano L and Gratton E, Microsc Res Tech (2012) 75, 1253-1264

Extended abstract of a paper presented at Microscopy and Microanalysis 2012 in Phoenix, Arizona, USA, July 29 – August 2, 2012.

We demonstrate the application of Fluorescence Lifetime Imaging (FLIM) to read out Forster resonant energy transfer (FRET) based biosensors for studying the spatio-temporal dynamics of signalling pathways in cells undergoing chemotaxis.