The authors would like to thank the Biophysics group and Jacob Piehler at the University of Osnabr&#252;ck for continuous support, Wladislaw Kohl for technical assistance and preparation of material, and the CellNanOs board for providing microscopes for use. The project was funded by the SFB 944.

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The authors have nothing to disclose.

Here, a technique for dual color single molecule localization of mobile membrane proteins was presented. Following the protocol, membrane proteins are fused to self-labeling proteins that react with the rhodamine dyes TMR and SiR conjugated to their respective substrates. Rhodamine dyes are bright and photostable and thus allow for repetitive imaging1. For successful performance, several conditions and critical topics have to be kept in mind.
First, it is important to choose appropriate filters and splitters to separate the signals from TMR and SiR clearly. To reduce background from outside the cell, a precleaning and coating of the coverslip with PLL-PEG-RGD was helpful. The final concentration of the dye-substrate must be tested by considering the affinity of the self-labeling enzyme for its substrate and the accessibility of the enzyme inside the cell. Nano- to picomolar concentrations were sufficient for mitochondrial proteins17&#160;and stress granules13. For other organelles, the required concentrations need to be tested. If the staining is too strong at the beginning of the recording and no single molecules are distinguishable, it is best to wait until bleaching has reduced the amount of fluorescent dyes so that single particles (SP) can be discerned. We have found that concentrations of dyes higher than 30 nM for BG-substrates and 1 nM for HTL substrates during the staining procedure, or longer staining times in order to increase the number of labeled molecules, are not feasible. For each experiment, the intensity of the excitation laser has to be adapted, since the subcellular location of the dye influences its fluorescence behavior (quantum yield, bleaching, blinking)5 due to different environmental conditions such as pH and redox state3. It further must be mentioned that the background is higher in dual-color experiments when compared to single color imaging. This influences the precision of localization. Therefore, it is critical to optimize the laser power to achieve good signal to noise (S/N) ratios but low photobleaching rates. Typical laser power densities in HILO microscopy are in the range of 25 &#177; 8 kW/cm2. For fine-tuning of excitation power, different sets of neutral density filters can be used if laser diodes are not directly modulated and not modulated via an acousto-optical modulator. It is recommended to use special 2 mm thick mirrors for TIRF to reduce beam distortions. Especially under TIR and HILO conditions, very efficient blocking of excitation light is needed. For single molecule imaging and localization with subpixel accuracy, a proper spatial sampling frequency (Nyquist-Shannon sampling theorem) is required. This should be twice as high as the maximum spatial frequency defined by the resolution limit of the imaging system37. Using an oil immersion objective with a high numerical aperture (NA &#62;1.4), the resolution, d = &#955;/(2 &#215; NA), is typically at 200-250 nm for orange to far-red dyes. Thus, considering the physical pixel size of the detector, the magnification of the imaging optics should result in an image pixel size of approximately 100 nm. For an EMCCD camera with a pixel size of 16 x 16 &#181;m2 in combination with a 150X objective, the pixel size is 106.7 nm and thus adequate.
Generally, it is suggested to use stable transfected cells because this has the advantage that a steady amount of tagged proteins is expressed in most of the cells24. However, for dual-color localization and tracking experiments, this would require a double transfection and double selection with different antibiotics. Therefore, it is often more feasible to transiently transfect an already stable cell line with a second plasmid encoding the additional tagged protein of interest.
For mitochondria, movement and fragmentation is a critical issue. Fragmented or moving organelles should not be recorded or analyzed. Movement can be checked by overlaying rendered images of localized molecules from the beginning and the end of the experiment, with two different (false) colors. A homogeneous distribution of the signals from the organelles in the overlay indicates that organelles have not moved. Generally, room temperature reduces the movement of cellular compounds and structures.

The goal of this protocol is to provide an imaging method to localize and track single membrane proteins inside live cells. We call this method Tracking and Localization Microscopy (TALM)1,2. Like Stochastic Optical Reconstruction Microscopy (STORM)3 and Fluorescence Photoactivation Localization Microscopy ((F)PALM)4,5, TALM is a single molecule-based fluorescence localization technique. However, it is distinct in the way that the mobility of membrane proteins in combination with repetitive imaging of the same labeled molecule at different positions reveals the microcompartment that is accessible for the mobile protein. In other words, the possible localizations of the protein are set by the architecture of the organelle and by the mobility of the protein1. The method is complementary to various other super-resolution techniques6,7,8 because it reveals localization and trajectory maps by imaging mobile proteins. The labeling is based on using genetically engineered fusion proteins that are per se non-fluorescent. These fusion proteins are self-labeling enzymes that react covalently with a substrate conjugated to a dye. This procedure has the advantage that the labeling degree can be controlled by the amount of added substrate. Furthermore, it allows to vary the color of fluorescence, depending on the chosen conjugated dye. Several self-labeling enzyme-tags are available9. Another advantage of using self-labeling enzyme-tags is, that the conjugated dyes usually are more stable and brighter than fluorescent proteins1 and individual proteins therefore can be recorded longer and more precisely until they are bleached. This allows for the recording of trajectories of mobile proteins and the extraction of diffusion coefficients10,11.
Here, we demonstrate the feasibility of TALM with mitochondrial membrane proteins, but it can also be applied for other intra- and extra-cellular membrane proteins, including different cell types12,13. We show that multi-color TALM further allows for the simultaneous distinction of proteins in different subcompartments in complementation to existing super-resolution fluorescence microscopy techniques14,15,16. TALM is compatible with live cell imaging17. The photo-physics of the chosen rhodamines Tetramethylrhodamine (TMR) and Silicon-Rhodamien (SiR), in particular their brightness and stability, allows to record single membrane proteins over multiple frames providing localization (and trajectory) maps. However, TALM is limited for the localization of soluble proteins with high diffusion coefficients since the motion blur is too high and the collected photons per frame are too low for proper localization. Besides, TALM requires less excitation power than for example STORM or Stimulated Emission Depletion (STED) microscopy6,7, reducing phototoxic effects. This is important, since phototoxic stress often affects organellar morphology18&#160;and thus mobility analysis19. In sum, we present multi-color TALM in living cells as a technique that fills a gap between the localization microscopy methods STORM/STED/(F)PALM and techniques that analyze protein mobility such as fluorescence recovery after photobleaching (FRAP)20,21, fluorescence correlation spectroscopy (FCS)22, and fluorescence cross correlation spectroscopy (FCCS)11,23.

<table><tbody><tr><td>(2-(4-(2-hydroxyethyl)-1-piperazinyl)-ethanesulfonic acid, 1 M) (HEPES)</td><td>Biochrom</td><td>#1104E</td><td></td></tr><tr><td>DC1: Dichroid beam splitter&amp;nbsp;</td><td>Chroma</td><td>640 dcxr</td><td>NC506031</td></tr><tr><td>DC2: Polychroic Mirror, beamsplitter</td><td>Chroma</td><td>zt405/488/561/640rpc</td><td>discontinued</td></tr><tr><td>Dulbecco&amp;acute;s Phosphate-Buffered Saline (PBS) 1x (w/o Ca &amp;amp; Mg)</td><td>Sigma-Aldrich &amp;amp; Co.</td><td>#RNBF8311</td><td></td></tr><tr><td>Earle&amp;acute;s MEM without phenol red, without L-Glutamine and without NaHCO3 containing 1% FBS, 0.1% HEPES, 0.1% NEAA, 0.1% Alanyl-L-Glutamine and 34.78% sodium hydrogen carbonate (NaHCO3 0.75g/l)</td><td></td><td></td><td>Imaging medium</td></tr><tr><td>Earle&amp;acute;s minimum essential medium (MEM) with phenol red, containing 1% Fetal Bovine Serum Superior (FBS), 0.1% HEPES (2-(4-(2-hydroxyethyl)-1-piperazinyl)-ethanesulfonic acid, 1 M), and 0.1% non-essential amino acids (NEAA)</td><td></td><td></td><td>Growth medium</td></tr><tr><td>EF: Emission filter quadbandpass</td><td>AHF analysentechnik</td><td>F72-866</td><td>Brightline HC 446 nm/523 nm/600 nm/677 nm</td></tr><tr><td>EMCCD camera</td><td>Andor</td><td>Andor iXON 897</td><td>EMCCD camera</td></tr><tr><td>Emission filter QuadView filter cubes, orange</td><td>AHF analysentechnik</td><td>F39-637</td><td>bandpass 582 - 619 nm</td></tr><tr><td>Emission filter QuadView filter cubes, red</td><td>Chroma</td><td></td><td>bandpass 655 - 725 nm (HQ 690/70)</td></tr><tr><td>FBS (Fetal bovine serum) superior</td><td>Biochrom</td><td>S0615</td><td></td></tr><tr><td>Fluorescent beads: TetraSpeck&amp;trade; Microspheres, 0.1 &amp;micro;m, fluorescent blue/green/orange/dark red</td><td>Thermo Fisher Scientific</td><td>T7279</td><td>fluorescent microspheres</td></tr><tr><td>Glutamine</td><td>Biochrom</td><td>#0951C</td><td></td></tr><tr><td>HeLa cells&amp;nbsp;</td><td>DSMZ</td><td>ACC-57</td><td>Cervical carcinoma cells from patient Henrietta Lacks</td></tr><tr><td>Hela cells CI::paGFP, stable</td><td>Muster et al., PLOSOne 2010</td><td></td><td></td></tr><tr><td>Hela cells CV g::Halo7-Tag, stable</td><td>Appelhans et al., NanoLett 2012</td><td></td><td></td></tr><tr><td>Hela cells Tom20::Halo7-Tag, stable</td><td>Appelhans et al., NanoLett 2012</td><td></td><td></td></tr><tr><td>Image splitter</td><td>Photometrics</td><td>Dual-View QV2</td><td>image splitter emission</td></tr><tr><td>Imaging processing software&amp;nbsp;</td><td>ImageJ2 / Fiji</td><td>freeware</td><td></td></tr><tr><td>Immersion Oil - ImmersolTM 518 F (ne = 1.518, ve = 45)</td><td>Carl Zeiss Jena GmbH</td><td>444960-0000-000</td><td></td></tr><tr><td>Inverted epifluorescence microscope</td><td>Olympus IX-71/73/83</td><td></td><td></td></tr><tr><td>Laser 561 nm, 200 mW</td><td>CrystaLaser</td><td>CL-561-200</td><td>561 nm emission</td></tr><tr><td>Laser 642 nm, 140 mW</td><td>Omicron</td><td>Luxx-642-140</td><td>642 nm emission</td></tr><tr><td>MATLAB</td><td>MathWorks</td><td>version R2013a</td><td></td></tr><tr><td>MEM with Earle&#039;s Balanced Salt Solution 2.2 g/L NaHCO3, stable glutamine w/o PR</td><td>Biochrom</td><td>FG-0385</td><td></td></tr><tr><td>MEM with Earle&#039;s Balanced Salt Solution with 2.2 g/L NaHCO3, stable glutamine, Phenolred</td><td>Biochrom</td><td>FG-0325</td><td></td></tr><tr><td>MitoTracker&amp;reg; Deep Red FM</td><td>Thermo Fisher Scientific</td><td>M22426</td><td>dye</td></tr><tr><td>MitoTracker&amp;reg; Green FM</td><td>Thermo Fisher Scientific</td><td>M7514</td><td>dye</td></tr><tr><td>Multi-mode-optical polarization maintaining monomode fiber</td><td>Pointsource/Qioptiq</td><td></td><td>KineFLEX</td></tr><tr><td>NHS-PEG-MAL, Rapp Polymer&amp;nbsp;</td><td>Rapp Polymere GmbH T&amp;uuml;bingen</td><td></td><td>coverslip coating</td></tr><tr><td>non-essential amino acids (NEAA)</td><td>Biochrom</td><td>#0802E</td><td></td></tr><tr><td>PEG 800 (Polyethylene glycol) 10 %</td><td>Carl Roth GmbH</td><td>Art No. 0263.1</td><td>coverslip coating</td></tr><tr><td>Penicillin/Streptomycin</td><td>Biochrom</td><td>#0122E</td><td></td></tr><tr><td>Plasmid for PINK1-Halo7-Tag expression</td><td>Beinlich et al., ACS Chemical Biology 2015</td><td></td><td></td></tr><tr><td>Poly-L-lysine (1.2 mg/ml)</td><td>Sigma-Aldrich &amp;amp; Co.</td><td>Cat. No.P9155</td><td>coverslip coating</td></tr><tr><td>RGD Peptide (Ac-CGRGDS-COOH)</td><td>Coring System Diagnostix GmbH, Gernsheim</td><td></td><td>coverslip coating / Intergrin receptor motif</td></tr><tr><td>Silicon Rhodamine linked to HaloTag&amp;reg;-Ligand (SiRHTL)&amp;nbsp;</td><td>personal gift from Kai Johnson</td><td></td><td>dye</td></tr><tr><td>Software analysis plugin</td><td>self-written C. P. Richter, Biophysik Osnabr&amp;uuml;ck</td><td>SLIMFAST 16g</td><td></td></tr><tr><td>Tetramethylrhodamine / SNAP-Cell&amp;reg; TMR-Star linked to SNAP-Ligand (TMRstar)&amp;nbsp;</td><td>New England Biolab&amp;reg;</td><td>S9105S</td><td>dye</td></tr><tr><td>Tetramethylrhodamine linked to HaloTag&amp;reg;-Ligand (TMRHTL)&amp;nbsp;</td><td>Promega</td><td>G8251</td><td>dye</td></tr><tr><td>TIRF condensor</td><td>Olympus</td><td>Cell^TIRF MITICO System</td><td>TIRF condensor</td></tr><tr><td>TIRF microscope controlling software</td><td>Olympus cellSens 1.12</td><td></td><td></td></tr><tr><td>TIRF objective</td><td>Olympus</td><td>150x oil objective (N.A. 1.45; Olympus UAPO)</td><td></td></tr><tr><td>Trypsin/EDTA 10x</td><td>Biochrom</td><td>#0266</td><td></td></tr><tr><td>Water H2O 99,5 % Rotipuran&amp;reg; Low organic&amp;nbsp;</td><td>Carl Roth GmbH</td><td>Art. No. HN57.1</td><td></td></tr></tbody></table>

multi-color localization microscopy of single membrane proteins in organelles of live mammalian cells

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a super-resolution technique that allows for the determination of the microcompartments that are accessible for proteins by generating localization and tracking maps of these proteins. Moreover, by multi-color localization microscopy, the localization and tracking profiles of proteins in different subcompartments are obtained simultaneously. The technique is specific for live cells and is based on the repetitive imaging of single mobile membrane proteins. Proteins of interest are genetically fused with specific, so-called self-labeling tags. These tags are enzymes that react with a substrate in a covalent manner. Conjugated to these substrates are fluorescent dyes. Reaction of the enzyme-tagged proteins with the fluorescence labeled substrates results in labeled proteins. Here, Tetramethylrhodamine (TMR) and Silicon Rhodamine (SiR) are used as fluorescent dyes attached to the substrates of the enzymes. By using substrate concentrations in the pM to nM range, sub-stoichiometric labeling is achieved that results in distinct signals. These signals are localized with ~15&#8211;27 nm precision. The technique allows for multi-color imaging of single molecules, whereby the number of colors is limited by the available membrane-permeable dyes and the repertoire of self-labeling enzymes. We show the feasibility of the technique by determining the localization of the quality control enzyme (Pten)-induced kinase 1 (PINK1) in different mitochondrial compartments during its processing in relation to other membrane proteins. The test for true physical interactions between differently labeled single proteins by single molecule FRET or co-tracking is restricted, though, because the low labeling degrees decrease the probability for having two adjacent proteins labeled at the same time. While the technique is strong for imaging proteins in membrane compartments, in most cases it is not appropriate to determine the localization of highly mobile soluble proteins.

Multi-color imaging and colocalization analysis can help to determine the sub-organellar localization&#160;of proteins. We demonstrated this earlier with the cytosolic phosphatase and tensin homologue, PINK1, that has different sub-mitochondrial locations due to its processing by mitochondrial proteases17. PINK1 is an important factor guaranteeing mitochondrial functionality34,35. To determine the localization of PINK1 in different mitochondrial compartments in the course of its processing (Figure 4A), multi-color super-resolution microscopy with live cells was performed. Therefore, PINK1 was genetically fused to a self-labeling tag (HaloTag). To determine its localization relative to other proteins in functional and dysfunctional mitochondria, Tom20, as part of the translocase of the outer mitochondrial membrane (TOM)36, was tagged with another self-labeling tag (SNAP-tag). Time series with at least 1,000 frames (96 frames per s) were recorded. The cumulative image of all signals over time from one channel showed that PINK1 was localized in the cytosol and in the mitochondria (Figure 4B, left panel) under normal conditions, while it was retained at the outer membrane of depolarized mitochondria (treatment with 10 &#181;M of the uncoupler carbonyl cyanide m-chlorophenyl hydrazine, CCCP, for 20 min). The corresponding trajectory maps also show this difference in the spatio-temporal organization. As the summed images of localized particles reveal, PINK1 is distributed mainly inside polarized mitochondria (Figure 4C, left upper panel), while Tom20 is localized in the outer mitochondrial membrane (OM) (Figure 4C, left lower panel). This becomes even more clear in the merged image of localized Tom20 and PINK1 particles, where the distribution of the outer membrane protein Tom20 is much broader (Figure 4C, right panel).
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57690/57690fig4.jpg" /> 
Figure 4: Dual color super-resolution microscopy showing the subcompartmental localization of PINK1 in mitochondria. (A) Hypothesized PINK1 shuttling and processing in mitochondria. (B) Localization and trajectory maps of PINK1 under normal conditions and in depolarized mitochondria. (C) Dual-color super-resolution localization microscopy to reveal mitochondrial localization of PINK1 in relation to Tom20. Image series of 1,000 frames recorded with a frame rate of 96 frames per s. Left upper panel: Localization map of PINK1 (labeled via HaloTag with TMRHTL). Left lower panel: localization map of Tom20 (labeled via SNAP-Tag with SiRBG). Right panel: Merge cumulative sum image of PINK1-TMR (red) and Tom20-SiR(blue)localizations. (D) Same data plus localizations of respiratory complex I (CI, green) in the inner mitochondrial membrane. CI was fused to paGFP (CI-paGFP). Right: Mean cross section profile of Tom20 (blue), PINK1 (red), and CI (green) fluorescence in the marked region of the merged image. The mean cross section profiles were obtained by averaging up to 30 parallel oriented cross lines (interval 40 nm). Adapted version reprinted with permission from Beinlich et al.16, copyright (2015) ACS chemical biology. All colors are false colors. Scale bars = 1 &#181;m. <a href="/files/ftp_upload/57690/57690fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
To verify the localization of PINK1 inside mitochondria, a third protein localized in the inner mitochondrial membrane was also imaged. Therefore, the 30 kDa subunit of respiratory complex I (CI) was fused to photoactivable GFP (paGFP) and recorded by FPALM4. The cumulative triple color image and the cross-section distribution show the overlay of CI (green) and PINK1 (red) (Figure 4D), confirming the import of full-length PINK1. PINK1 has an N-terminal mitochondrial targeting sequence and thus the label was put at the C-terminus of the kinase. Co-localization with CI demonstrates that the full length PINK1 had been&#160;imported. The fluorescence distribution along a cross-section shows that a quota of PINK1 particles apparently also colocalized with Tom20 in the outer mitochondrial membrane. Probably, this is PINK1 associated with Tom20, which is the import receptor. As these data demonstrate, several mitochondrial micro-compartments are occupied by subpopulations of PINK1. To address this question of sub-mitochondrial localization by biochemical methods would be much more difficult, since it would require stringent sub-fractionation of the different membranes and a clear separation of soluble components to exclude cross-contamination.
Supplementary Video 1: Footage of single molecule recording, 100 frames, CV-HaloTag/TMRHTL. <a href="/files/ftp_upload/57690/01_SM_raw_data_f-1-100.avi">Please click here to download this file.</a>
Supplementary Video 2: Localized data. <a href="/files/ftp_upload/57690/02_SM_localized_data_all_frames.avi">Please click here to download this file.</a>

Watch this Scientific Journal Video about Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells at JoVE.com

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

Here, we present a protocol for multi-color localization of single membrane proteins in organelles of live cells. To attach fluorophores, self-labeling proteins are used. Proteins, located in different membranes compartments of the same organelle, can be localized with a precision of ~18 nm.

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a ...

multi-color-localization-microscopy-single-membrane-proteins

Nanyang Technological University

Research

JoVE Journal

Biology

8.4K Views.  University of Osnabrück. Hier presenteren we een protocol voor Multi-Color lokalisatie van één membraaneiwitten in organellen van levende cellen. Als u wilt koppelen fluorophores, worden zelf labeling eiwitten gebruikt. Eiwitten, gelegen in verschillende membranen compartimenten van de dezelfde organel, kunnen worden gelokaliseerd met een nauwkeurigheid van ~ 18 nm.

Video: Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

Single-Molecule Tracking Microscopy - A Tool for Determining the Diffusive States of Cytosolic Molecules

Single-molecule localization microscopy probes the position and motions of individual molecules in living cells with tens of nanometer spatial and millisecond temporal resolution. These capabilities make single-molecule localization microscopy ideally suited to study molecular level biological functions in physiologically relevant environments. Here, we demonstrate an integrated protocol for both acquisition and processing/analysis of single-molecule tracking data to extract the different diffusive states a protein of interest may exhibit. This information can be used to quantify molecular complex formation in living cells. We provide a detailed description of a camera-based 3D single-molecule localization experiment, as well as the subsequent data processing steps that yield the trajectories of individual molecules. These trajectories are then analyzed using a numerical analysis framework to extract the prevalent diffusive states of the fluorescently labeled molecules and the relative abundance of these states. The analysis framework is based on stochastic simulations of intracellular Brownian diffusion trajectories that are spatially confined by an arbitrary cell geometry. Based on the simulated trajectories, raw single-molecule images are generated and analyzed in the same way as experimental images. In this way, experimental precision and accuracy limitations, which are difficult to calibrate experimentally, are explicitly incorporated into the analysis workflow. The diffusion coefficient and relative population fractions of the prevalent diffusive states are determined by fitting the distributions of experimental values using linear combinations of simulated distributions. We demonstrate the utility of our protocol by resolving the diffusive states of a protein that exhibits different diffusive states upon forming homo- and hetero-oligomeric complexes in the cytosol of a bacterial pathogen.

3D single-molecule localization microscopy is utilized to probe the spatial positions and motion trajectories of fluorescently labeled proteins in living bacterial cells. The experimental and data analysis protocol described herein determines the prevalent diffusive behaviors of cytosolic proteins based on pooled single-molecule trajectories.

Single-molecule localization microscopy probes the position and motions of individual molecules in living cells with tens of nanometer spatial and ...

Biochemistry

From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn&#8217;t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on &#181;m-sized membrane regions in the micro-to-milli-second time range.

Spatial distribution and temporal dynamics of plasma membrane proteins and lipids is a hot topic in biology. Here this issue is addressed by a spatio-temporal image fluctuation analysis that provides conceptually the same physical quantities of single particle tracking, but it uses small molecular labels and standard microscopy setups.

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the ...

Bioengineering

Conventional BODIPY Conjugates for Live-Cell Super-Resolution Microscopy and Single-Molecule Tracking

Single molecule localization microscopy (SMLM) techniques overcome the optical diffraction limit of conventional fluorescence microscopy and can resolve intracellular structures and the dynamics of biomolecules with ~20 nm precision. A prerequisite for SMLM are fluorophores that transition from a dark to a fluorescent state in order to avoid spatio-temporal overlap of their point spread functions in each of the thousands of data acquisition frames. BODIPYs are well-established dyes with numerous conjugates used in conventional microscopy. The transient formation of red-shifted BODIPY ground-state dimers (DII) results in bright single molecule emission enabling single molecule localization microscopy (SMLM). Here we present a simple but versatile protocol for SMLM with conventional BODIPY conjugates in living yeast and mammalian cells. This procedure can be used to acquire super-resolution images and to track single BODIPY-DII states to extract spatio-temporal information of BODIPY conjugates. We apply this procedure to resolve lipid droplets (LDs), fatty acids, and lysosomes in living yeast and mammalian cells at the nanoscopic length scale. Furthermore, we demonstrate the multi-color imaging capability with BODIPY dyes when used in conjunction with other fluorescent&#160;probes. Our representative results show the differential spatial distribution and mobility of BODIPY-fatty acids and neutral lipids in yeast under fed and fasted conditions. This optimized protocol for SMLM can be used with hundreds of commercially available BODIPY conjugates and is a useful resource to study biological processes at the nanoscale far beyond the applications of this work.&#160;

Conventional BODIPY conjugates can be used for live-cell single-molecule localization microscopy (SMLM) through exploitation of their transiently forming, red-shifted ground state dimers. We present an optimized SMLM protocol to track and resolve subcellular neutral lipids and fatty acids in living mammalian and yeast cells at the nanoscopic length scale.

Single molecule localization microscopy (SMLM) techniques overcome the optical diffraction limit of conventional fluorescence microscopy and can resolve ...

Live-Cell Fluorescence Microscopy to Investigate Subcellular Protein Localization and Cell Morphology Changes in Bacteria

Investigations of factors influencing cell division and cell shape in bacteria are commonly performed in conjunction with high-resolution fluorescence microscopy as observations made at a population level may not truly reflect what occurs at a single cell level. Live-cell timelapse microscopy allows investigators to monitor the changes in cell division or cell morphology which provide valuable insights regarding subcellular localization of proteins and timing of gene expression, as it happens, to potentially aid in answering important biological questions. Here, we describe our protocol to monitor phenotypic changes in Bacillus subtilis and Staphylococcus aureus using a high-resolution deconvolution microscope. The objective of this report is to provide a simple and clear protocol that can be adopted by other investigators interested in conducting fluorescence microscopy experiments to study different biological processes in bacteria as well as other organisms.

This article provides a step-by-step guide to investigate protein subcellular localization dynamics and to monitor morphological changes using high-resolution fluorescence microscopy in Bacillus subtilis and Staphylococcus aureus.

Investigations of factors influencing cell division and cell shape in bacteria are commonly performed in conjunction with high-resolution fluorescence ...

Immunology and Infection

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Autophagy is a cellular response triggered by the lack of nutrients, especially the absence of amino acids. Autophagy is defined by the formation of double membrane structures, called autophagosomes, that sequester cytoplasm, long-lived proteins and protein aggregates, defective organelles, and even viruses or bacteria. Autophagosomes eventually fuse with lysosomes leading to bulk degradation of their content, with the produced nutrients being recycled back to the cytoplasm. Therefore, autophagy is crucial for cell homeostasis, and dysregulation of autophagy can lead to disease, most notably neurodegeneration, ageing and cancer.	
	Autophagosome formation is a very elaborate process, for which cells have allocated a specific group of proteins, called the core autophagy machinery. The core autophagy machinery is functionally complemented by additional proteins involved in diverse cellular processes, e.g. in membrane trafficking, in mitochondrial and lysosomal biology. Coordination of these proteins for the formation and degradation of autophagosomes constitutes the highly dynamic and sophisticated response of autophagy. Live cell imaging allows one to follow the molecular contribution of each autophagy-related protein down to the level of a single autophagosome formation event and in real time, therefore this technique offers a high temporal and spatial resolution.	
	Here we use a cell line stably expressing GFP-DFCP1, to establish a spatial and temporal context for our analysis. DFCP1 marks omegasomes, which are precursor structures leading to autophagosomes formation. A protein of interest (POI) can be marked with either a red or cyan fluorescent tag. Different organelles, like the ER, mitochondria and lysosomes, are all involved in different steps of autophagosome formation, and can be marked using a specific tracker dye. Time-lapse microscopy of autophagy in this experimental set up, allows information to be extracted about the fourth dimension, i.e. time. Hence we can follow the contribution of the POI to autophagy in space and time.

Time-lapse microscopy of fluorescently labeled autophagy markers allows monitoring of the dynamic autophagy response with high temporal resolution. Using specific autophagy and organelle markers in a combination of 3 different colors, we can follow the contribution of a protein to autophagosome formation in a robust spatial and temporal context.

Autophagy is a  cellular response triggered by the lack of nutrients, especially the absence of  amino acids. Autophagy is defined by the formation of ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and functions despite the intense interorganelle molecular traffic. The techniques described in this paper are powerful means of studying protein and lipid mobility and trafficking in vivo and in their physiological environment. Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) are widely used live-cell imaging techniques for studying intracellular trafficking through the exo-endocytic pathway, the continuity between organelles or subcompartments, the formation of protein complexes, and protein localization in lipid microdomains, all of which can be observed under physiological and pathological conditions. The limitations of these approaches are mainly due to the use of fluorescent fusion proteins, and their potential drawbacks include artifactual over-expression in cells and the possibility of differences in the folding and localization of tagged and native proteins. Finally, as the limit of resolution of optical microscopy (about 200 nm) does not allow investigation of the fine structure of the ER or the specific subcompartments that can originate in cells under stress (i.e. hypoxia, drug administration, the over-expression of transmembrane ER resident proteins) or under pathological conditions, we combine live-cell imaging of cultured transfected cells with ultrastructural analyses based on transmission electron microscopy.

We describe the imaging approaches we use to investigate the distribution and mobility of the transfected fluorescent proteins resident in the endoplasmic reticulum (ER) by means of the confocal imaging of living cells. We also ultrastructurally analyze the effect of their expression on the architecture of this subcellular compartment.

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and ...

Unveiling Mitochondrial Ultrastructure in Neurons Using STED Microscopy

Using Live Cell STED Imaging to Visualize Mitochondrial Inner Membrane Ultrastructure in Neuronal Cell Models

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of reactive oxygen species (ROS). These mitochondria-mediated processes take on specialized roles in neurons, coordinating aerobic metabolism to meet the high energy demands of these cells, modulating Ca2+ signaling, providing lipids for axon growth and regeneration, and tuning ROS production for neuronal development and function. Mitochondrial dysfunction is therefore a central driver in neurodegenerative diseases. Mitochondrial structure and function are inextricably linked. The morphologically complex inner membrane with structural infolds called cristae harbors many molecular systems that perform the signature processes of the mitochondrion. The architectural features of the inner membrane are ultrastructural and therefore, too small to be visualized by traditional diffraction-limited resolved microscopy. Thus, most insights on mitochondrial ultrastructure have come from electron microscopy on fixed samples. However, emerging technologies in super-resolution fluorescence microscopy now provide resolution down to tens of nanometers, allowing visualization of ultrastructural features in live cells. Super-resolution imaging therefore offers an unprecedented ability to directly image fine details of mitochondrial structure, nanoscale protein distributions, and cristae dynamics, providing fundamental new insights that link mitochondria to human health and disease. This protocol presents the use of stimulated emission depletion (STED) super-resolution microscopy to visualize the mitochondrial ultrastructure of live human neuroblastoma cells and primary rat neurons. This procedure is organized into five sections: (1) growth and differentiation of the SH-SY5Y cell line, (2) isolation, plating, and growth of primary rat hippocampal neurons, (3) procedures for staining cells for live STED imaging, (4) procedures for live cell STED experiments using a STED microscope&#160;for reference, and (5) guidance for segmentation and image processing using examples to measure and quantify morphological features of the inner membrane.

Author Spotlight: Decoding Mitochondrial Aging 

This protocol presents a workflow for the propagation, differentiation, and staining of cultured SH-SY5Y cells and primary rat hippocampal neurons for mitochondrial ultrastructure visualization and analysis using stimulated emission depletion (STED) microscopy.

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of ...

Neuroscience

Determination of Plasma Membrane Partitioning for Peripherally-associated Proteins

This method provides a fast approach for the determination of plasma membrane partitioning of any fluorescently-tagged peripherally-associated protein using the profiles of fluorescence intensity across the plasma membrane. Measured fluorescence profiles are fitted by a model for membrane and cytoplasm fluorescence distribution along a line applied perpendicularly to the cell periphery. This model is constructed from the fluorescence intensity values in reference cells expressing a fluorescently-tagged marker for cytoplasm and with FM 4-64-labeled plasma membrane. The method can be applied to various cell types and organisms; however, only plasma membranes of non-neighboring cells can be evaluated. This fast microscopy-based method is suitable for experiments, where subtle and dynamic changes of plasma membrane-associated markers are expected and need to be quantified, e.g., in the analysis of mutant versions of proteins, inhibitor treatments, and signal transduction observations. The method is implemented in a multi-platform R package that is coupled with an ImageJ macro that serves as a user-friendly interface.

Here, we present a protocol to perform a quantitative analysis of the level of plasma-membrane association for fluorescently-tagged peripherally-associated protein. The method is based on the computational decomposition of membrane and cytoplasmic component of signal observed in cells labeled with plasma membrane fluorescent marker.

This method provides a fast approach for the determination of plasma membrane partitioning of any fluorescently-tagged peripherally-associated protein ...

Correlative Microscopy Reveals Stress-Induced Mitochondria-Lysosome Interactions

Dual-color Correlative Light and Electron Microscopy for the Visualization of Interactions between Mitochondria and Lysosomes

Cellular organelles, such as mitochondria and lysosomes, display dynamic structures. Despite the higher resolution of transmission electron microscopy for structural analysis, light microscopy is essential for the visualization of dynamic organelles by target-specific labeling. The following protocol describes a method that combines dual-color correlative light and electron microscopy (CLEM) to observe the interactions between mitochondria and lysosomes. In this study, mitochondria were labeled with mEosEM (Mito-mEosEM) and lysosomes with TMEM192-V5-APEX2. The results obtained from CLEM images enable us to observe the changes in the interactions between mitochondria and lysosomes under external stress conditions. Treatment with bafilomycin (BFA), which inhibits lysosomal function, resulted in an increase in contact between mitochondria and lysosomes, leading to the formation of fragmented mitochondria trapped inside lysosomes. Conversely, treatment with U18666A, which inhibits cholesterol export from lysosomes, caused lysosomes to be surrounded by mitochondria, indicating a distinct form of interaction. This study presents an effective method for observing the interactions between mitochondria and lysosomes in fixed cells. Furthermore, CLEM imaging with dual-color probes offers a powerful tool for future investigations of organelle dynamics and their implications for cell function and pathology.

This protocol describes a mechanism for using correlative light and electron microscopy to visualize the interaction of mitochondria and lysosomes labeled with mEosEM and APEX2, respectively.

Cellular organelles, such as mitochondria and lysosomes, display dynamic structures. Despite the higher resolution of transmission electron microscopy for ...

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

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