Bioenergetic Alterations of Metabolic Redox Coenzymes as NADH,FAD and FMN by Means of Fluorescence Lifetime Imaging Techniques

Metabolic FLIM (fluorescence lifetime imaging) is used to image bioenergetic status in cells and tissue. Whereas an attribution of the fluorescence lifetime of coenzymes as an indicator for cell metabolism is mainly accepted, it is debated whether this is valid for the redox state of cells. In this regard, an innovative algorithm using the lifetime characteristics of nicotinamide adenine dinucleotide (phosphate) (NAD(P)H) and flavin adenine dinucleotide (FAD) to calculate the fluorescence lifetime induced redox ratio (FLIRR) has been reported so far. We extended the FLIRR approach and present new results, which includes FLIM data of the various enzymes, such as NAD(P)H, FAD, as well as flavin mononucleotide (FMN). Our algorithm uses a two-exponential fitting procedure for the NAD(P)H autofluorescence and a three-exponential fit of the flavin signal. By extending the FLIRR approach, we introduced FLIRR1 as protein-bound NAD(P)H related to protein-bound FAD, FLIRR2 as protein-bound NAD(P)H related to free (unbound) FAD and FLIRR3 as protein-bound NAD(P)H related to protein-bound FMN. We compared the significance of extended FLIRR to the metabolic index, defined as the ratio of protein-bound NAD(P)H to free NAD(P)H. The statistically significant difference for tumor and normal cells was found to be highest for FLIRR1.     

A novel fluorescence lifetime imaging system that optimizes photon efficiency

Fluorescence lifetime imaging (FLIM) is a powerful microscopy technique for providing contrast of biological and other systems by differences in molecular species or their environments. However, the cost of equipment and the complexity of data analysis have limited the application of FLIM. We present a mathematical model and physical implementation for a low cost digital frequency domain FLIM (DFD-FLIM) system, which can provide lifetime resolution with quality comparable to time-correlated single photon counting methods. Our implementation provides data natively in the form of phasors. On the basis of the mathematical model, we present an error analysis that shows the precise parameters for maximizing the quality of lifetime acquisition, as well as data to support this conclusion. The hardware and software of the proposed DFD-FLIM method simplifies the process of data acquisition for FLIM, presents a new interface for data display and interpretation, and optimizes the accuracy of lifetime determination.     

Correlated fluorescence lifetime and spectral measurements in living cells

Studies of proteins' interaction in cells by FRET can take benefit from two important photo-physical properties describing fluorescent proteins: fluorescence emission spectrum and fluorescence lifetime. These properties provide specific and complementary information about the tagged proteins and their environment. However, none of them taken individually can completely quantify the involved fluorophore characteristics due to their multiparametric dependency with molecular environment, experimental conditions, and interpretation complexity. A solution to get a better understanding of the biological process implied at the cellular level is to combine the spectral and temporal fluorescence data acquired simultaneously at every cell region under investigation. We present the SLiM-SPRC160, an original temporal/spectral acquisition system for simultaneous lifetime measurements in 16 spectral channels directly attached to the descanned port of a confocal microscope with two-photon excitation. It features improved light throughput, enabling low-level excitation and minimum invasivity in living cells studies. To guarantee a fairly good level of accuracy and reproducibility in the measurements of fluorescence lifetime and spectra on living cells, we propose a rigorous protocol for running experiments with this new equipment that preserves cell viability. The usefulness of SLiM approach for the precise determination of overlapping fluorophores is illustrated with the study of known solutions of rhodamine. Then, we describe reliable FRET experiments in imaging mode realized in living cells using this protocol. We also demonstrate the benefit of localized fluorescence spectrum-lifetime acquisitions for the dynamic study of fluorescent proteins. proteins.     

Sensitivity of CFP/YFP and GFP/mCherry pairs to donor photobleaching on FRET determination by fluorescence lifetime imaging microscopy in living cells

Fluorescent protein-based FRET is a powerful method for visualizing protein-protein interactions and biochemical reactions in living cells. It can be difficult, however, to avoid photobleaching when observing fluorescent cells under the microscope, especially those expressing CFP. We compared the sensitivity of two protein-based FRET pairs to light-induced fluorescence changes in the donor, on FRET determination by fluorescence lifetime imaging microscopy (FLIM). Thanks to the very low excitation light levels of the time- and space-correlated single photon counting (TSCSPC) method, FLIM acquisitions were achieved without donor photobleaching. Here, we show that photobleaching of CFP by a mercury lamp under the microscope induced a decrease in the mean fluorescence lifetime, which interfered with FRET determination between CFP and YFP. Importantly, the range of light-induced variation of the mean fluorescence lifetime of CFP was not proportional to the decrease in the steady state fluorescence intensity and varied from cell to cell. The choice of the CFP/YFP pair therefore requires that the cells be observed and analyzed at very low light levels during the whole FRET experiment. In contrast, the GFP/mCherry pair provided an accurate FRET measurement by FLIM, even if some GFP photobleaching took place. We thus demonstrate that CFP can be an unreliable donor for FRET determination in living cells, due to its photosensitivity properties. We demonstrate that the GFP/mCherry pair is better suited for FRET measurement by FLIM in living cells than the CFP/YFP pair.     

Optimizing imaging parameters for the separation of multiple labels in a fluorescence image

A theoretical analysis is presented on how to separate the contributions from individual, simultaneously present fluorophores in a spectrally resolved image. Equations are derived that allow the calculation of the signal‐to‐noise ratio of the estimates for such contributions, given the spectral information on the individual fluorophores, the excitation wavelengths and intensities, and the number and widths of the spectral detection channels. We then ask how such imaging parameters have to be chosen for optimal fluorophore separation. We optimize the signal‐to‐noise ratio or optimize a newly defined ‘figure of merit’, which is a measure of efficiency in the use of emitted photons. The influence of photobleaching on the resolution and on the choice of imaging parameters is discussed, as well as the additional resolution gained by including fluorescence lifetime information. A surprisingly small number of spectral channels are required for an almost optimal resolution, if the borders of these channels are optimally selected. The detailed consideration of photobleaching is found to be essential, whenever there is significant bleaching. Consideration of fluorescence lifetime information (in addition to spectral information) improves results, particularly when lifetimes differ by more than a factor of two.     

Protein‐Specific Imaging of O‐GlcNAcylation in Single Cells

Thousands of intracellular proteins are post‐translationally modified with O‐GlcNAc, and O‐GlcNAcylation impacts the function of modified proteins and mediates diverse biological processes. However, the ubiquity of this important glycosylation makes it highly challenging to probe the O‐GlcNAcylation state of a specific protein at the cellular level. Herein, we report the development of a FLIM–FRET‐based strategy, which exploits the spatial proximity of the O‐GlcNAc moiety and the attaching protein, for protein‐specific imaging of O‐GlcNAcylation in single cells. We demonstrated this strategy by imaging the O‐GlcNAcylation state of tau and β‐catenin inside the cells. Furthermore, the changes in tau O‐GlcNAcylation were monitored when the overall cellular O‐GlcNAc was pharmacologically altered by using the OGT and OGA inhibitors. We envision that the FLIM–FRET strategy will be broadly applicable to probe the O‐GlcNAcylation state of various proteins in the cells.     

Flexible Nanosomes (SECosomes) Enable Efficient siRNA Delivery in Cultured Primary Skin Cells and in the Viable Epidermis of Ex Vivo Human Skin

The extent to which nanoscale‐engineered systems cross intact human skin and can exert pharmacological effects in viable epidermis is controversial. This research seeks to develop a new lipid‐based nanosome that enables the effective delivery of siRNA into human skin. The major finding is that an ultraflexible siRNA‐containing nanosome—prepared using DOTAP, cholesterol, sodium cholate, and 30% ethanol—penetrates into the epidermis of freshly excised intact human skin and is able to enter into the keratinocytes. The nanosomes, called surfactant‐ethanol‐cholesterol‐osomes (SECosomes), show excellent size, surface charge, morphology, deformability, transfection efficiency, stability, and skin penetration capacity after complexation with siRNA. Importantly, these nanosomes have ideal characteristics for siRNA encapsulation, in that the siRNA is stable for at least 4 weeks, they enable highly efficient transfection of in vitro cultured cells, and are shown to transport siRNA delivery through intact human skin where changes in the keratinocyte cell state are demonstrated. It is concluded that increasing flexibility in nanosomes greatly enhances their ability to cross the intact human epidermal membrane and to unload their payload into targeted epidermal cells.     

Retraction of ‘Basu,S., Dahl,K.N. & Rohde,G.K. (2013) Localizing and extracting filament distributions from microscopy images. Journal of Microscopy, 250, 57–67. doi:10.1111/jmi.12018’

The above article, published online on 4 March 2013 in Wiley Online Library ( wileyonlinelibrary.com ), has been retracted by agreement between the authors, the journal Editor in Chief, Tony Wilson, the Royal Microscopical Society and John Wiley & Sons Ltd.  The retraction has been agreed following an investigation by an ad hoc advisory committee of senior faculty members assembled by the College of Engineering, Carnegie Mellon University. The committee concluded that all the parties involved acted honourably, but implicit misunderstandings and unintended miscommunications led to the unfortunate inclusion of several images used in the study without proper approval from the copyright owner.     

A brief history of the octopus imaging facility to celebrate its 10th anniversary

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.