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Abstract Background Fluorescence loss in photobleaching (FLIP) is a widely used imaging technique, which provides information about protein dynamics in various cellular regions. In FLIP, a small cellular region is repeatedly illuminated by an intense laser pulse, while images are taken with reduced laser power with a time lag between the bleaches. Despite its popularity, tools are lacking for quantitative analysis of FLIP experiments. Typically, the user defines regions of interest (ROIs) for further analysis which is subjective and does not allow for comparing different cells and experimental settings. Results We present two complementary methods to detect and quantify protein transport and aggregation in living cells from FLIP image series. In the first approach, a stretched exponential (StrExp) function is fitted to fluorescence loss (FL) inside and outside the bleached region. We show by reaction–diffusion simulations, that the StrExp function can describe both, binding/barrier–limited and diffusion-limited FL kinetics. By pixel-wise regression of that function to FL kinetics of enhanced green fluorescent protein (eGFP), we determined in a user-unbiased manner from which cellular regions eGFP can be replenished in the bleached area. Spatial variation in the parameters calculated from the StrExp function allow for detecting diffusion barriers for eGFP in the nucleus and cytoplasm of living cells. Polyglutamine (polyQ) disease proteins like mutant huntingtin (mtHtt) can form large aggregates called inclusion bodies (IB’s). The second method combines single particle tracking with multi-compartment modelling of FL kinetics in moving IB’s to determine exchange rates of eGFP-tagged mtHtt protein (eGFP-mtHtt) between aggregates and the cytoplasm. This method is self-calibrating since it relates the FL inside and outside the bleached regions. It makes it therefore possible to compare release kinetics of eGFP-mtHtt between different cells and experiments. Conclusions We present two complementary methods for quantitative analysis of FLIP experiments in living cells. They provide spatial maps of exchange dynamics and absolute binding parameters of fluorescent molecules to moving intracellular entities, respectively. Our methods should be of great value for quantitative studies of intracellular transport.

Background: Fluorescence loss in photobleaching (FLIP) is a widely used imaging technique, which provides information about protein dynamics in various cellular regions. In FLIP, a small cellular region is repeatedly illuminated by an intense laser pulse, while images are taken with reduced laser power with a time lag between the bleaches. Despite its popularity, tools are lacking for quantitative analysis of FLIP experiments. Typically, the user defines regions of interest (ROIs) for further analysis which is subjective and does not allow for comparing different cells and experimental settings. Results: We present two complementary methods to detect and quantify protein transport and aggregation in living cells from FLIP image series. In the first approach, a stretched exponential (StrExp) function is fitted to fluorescence loss (FL) inside and outside the bleached region. We show by reaction–diffusion simulations, that the StrExp function can describe both, binding/barrier–limited and diffusion-limited FL kinetics. By pixel-wise regression of that function to FL kinetics of enhanced green fluorescent protein (eGFP), we determined in a user-unbiased manner from which cellular regions eGFP can be replenished in the bleached area. Spatial variation in the parameters calculated from the StrExp function allow for detecting diffusion barriers for eGFP in the nucleus and cytoplasm of living cells. Polyglutamine (polyQ) disease proteins like mutant huntingtin (mtHtt) can form large aggregates called inclusion bodies (IB’s). The second method combines single particle tracking with multi-compartment modelling of FL kinetics in moving IB’s to determine exchange rates of eGFP-tagged mtHtt protein (eGFP-mtHtt) between aggregates and the cytoplasm. This method is self-calibrating since it relates the FL inside and outside the bleached regions. It makes it therefore possible to compare release kinetics of eGFP-mtHtt between different cells and experiments. Conclusions: We present two complementary methods for quantitative analysis of FLIP experiments in living cells. They provide spatial maps of exchange dynamics and absolute binding parameters of fluorescent molecules to moving intracellular entities, respectively. Our methods should be of great value for quantitative studies of intracellular transport. [ABSTRACT FROM AUTHOR]

Abstract: Hydrogen/deuterium exchange (HDX) monitored by mass spectrometry (HDX-MS) has become a valuable tool in studies of protein dynamics and protein interactions. Isotopic exchange is typically initiated by diluting a protein solution into deuterated buffer at physiological conditions. For MS analysis, the exchange reaction is quenched by acidification and cooling and the labeled protein (or a digest thereof) is analyzed by mass spectrometry. An inevitable deuterium loss occurs during quench conditions (i.e., back-exchange). The primary structure of the peptide influences the back-exchange rate due to steric hindrance by bulky side groups and by inductive and charge effects. Here we show that the back-exchange in histidine repeats (His6) at HDX-MS quench conditions is complete within a few seconds using either acetic acid or formic acid in the quench solution, while aspartic acid repeats (Asp6) retain deuterons for several minutes using formic acid. We employ electron transfer dissociation to obtain residue-specific deuterium levels of the Asp repeat in K2D6IIKIIK using a hybrid linear ion trap-Orbitrap mass spectrometer. Our results show an unexpected uneven distribution of deuterium in the Asp repeat. The implication of the rapid back-exchange of His repeats for HDX-MS protein hydrogen exchange studies is discussed. We also discuss the implications of retained deuterons in the Asp repeat of K2D6IIKIIK when this peptide is used as a probe for the occurrence of hydrogen scrambling. [Copyright &y& Elsevier]

Cellular cholesterol homeostasis relies on precise control of the sterol content of organelle membranes. Obtaining insight into cholesterol trafficking pathways and kinetics by live-cell imaging relies on two conditions. First, one needs to develop suitable analogs that resemble cholesterol as closely as possible with respect to their biophysical and biochemical properties. Second, the cholesterol analogs should have good fluorescence properties. This interferes, however, often with the first requirement, such that the imaging instrumentation must be optimized to collect photons from suboptimal fluorophores, but good cholesterol mimics, such as the intrinsically fluorescent sterols, cholestatrienol (CTL) or dehydroergosterol (DHE). CTL differs from cholesterol only in having two additional double bonds in the ring system, which is why it is slightly fluorescent in the ultraviolet (UV). In the first part of this protocol, we describe how to synthesize and image CTL in living cells relative to caveolin, a structural component of caveolae. In the second part, we explain in detail how to perform time-lapse experiments of commercially available BODIPY-tagged cholesterol (TopFluor-cholesterol ® ; TF-Chol) in comparison to DHE. Finally, using two-photon time-lapse imaging data of TF-Chol, we demonstrate how to use our imaging toolbox SpatTrack for tracking sterol rich vesicles in living cells over time.

Late endosomes and lysosomes (LE/LYSs) play a central role in trafficking of endocytic cargo, secretion of exosomes, and hydrolysis of ingested proteins and lipids. Failure in such processes can lead to lysosomal storage disorders in which a particular metabolite accumulates within LE/LYSs. Analysis of endocytic trafficking relies heavily on quantitative fluorescence microscopy, but evaluation of the huge image data sets is challenging and demands computer-assisted statistical tools. Here, we describe how to use SpatTrack ( www.sdu.dk/bmb/spattrack ), an imaging toolbox, which we developed for quantification of the distribution and dynamics of endo-lysosomal cargo from fluorescence images of living cells. First, we explain how to analyze experimental images of endocytic processes in Niemann Pick C2 disease fibroblasts using SpatTrack. We demonstrate how to quantify the location of the sterol-binding protein NPC2 in LE/LYSs relative to cholesterol -rich lysosomal storage organelles (LSOs) stained with filipin. Second, we show how to simulate realistic vesicle patterns in the cell geometry using Markov Chain Monte Carlo and suitable inter-vesicle and cell-vesicle interaction potentials. Finally, we use such synthetic vesicle patterns as "ground truth" for validation of two-channel analysis tools in SpatTrack, revealing their high reliability. An improved version of SpatTrack for microscopy-based quantification of cargo transport through the endo-lysosomal system accompanies this protocol.