The research reported in this publication was supported by the National Institute of General Medical Sciences of the National Institutes of Health under award number R21GM127965.

NOTE: For yeast cloning and endogenous tagging please refer to our recent publication10.
1. Preparation of yeast cell samples for imaging
<ol>
	<li>Prepare a liquid overnight culture of a w303 yeast strain. Using a sterile wooden stick, spot a small amount of yeast cells from an agar plate containing yeast extract&#8211;peptone&#8211;dextrose into a culture tube with ~2 mL of synthetic complete dextrose (SCD) medium. Incubate the tube overnight in a shaking incubator at 270 rpm and 30 &#176;C.</li>
	<li>Perform a 1:50 morning dilution of the cells in SCD. Continue to culture the diluted cells for 4 h at 30 &#176;C in a 270 rpm shaking incubator, allowing the cells to grow in exponential phase and to reach an optical density (OD) of ~0.6. 
	NOTE: The procedure can vary here depending on which metabolic state is being studied. BODIPY conjugates do not require cells in the exponential growth phase. However, be cognizant of autofluorescence from dead cells during the stationary phase, as it can cause a background signal too strong to analyze single BODIPY-DII emitters.</li>
	<li>For studying fasting cells, grow the yeast culture for 2 days without exchange of media.</li>
	<li>At ~30 min prior to plating the cells, incubate a chambered coverglass with 80 &#181;L of 0.8 mg/mL sterile Concanavalin A (ConA) in deionized H2O at room temperature. After 30 min, wash the coverglass three times with deionized H2O.</li>
	<li>At ~30 min before imaging, pipette the cells on the chambered coverglass, with the correct volume of fresh SCD to achieve an optical density of ~0.12 (typically 60 &#181;L yeast culture at OD ~0.6 in 240 &#181;L SCD). Let the cells settle and adhere to the ConA surface for 30 min.</li>
	<li>Add the desired BODIPY conjugate directly to the chambered coverglass at a final concentration of ~100 nM. 
	NOTE: A BODIPY concentration optimization experiment may be required depending on BODIPYs local density in a particular cellular compartment.</li>
</ol>
2. Preparation of mammalian cells for SMLM imaging
<ol>
	<li>Maintain the mammalian U2OS cells in non-fluorescent DMEM with 10% fetal bovine serum, 4 mM glutamine, 1 mM sodium pyruvate and 1% penicillin-streptomycin antibiotics in a T25 flask. 
	NOTE: Cells can also be maintained in DMEM with 10% fetal bovine serum and 1% penicillin-streptomycin antibiotics, however, the medium needs to be exchanged before imaging with a non-autofluorescent solution.</li>
	<li>Split the cells at 70-80% confluency 1:5 in a single well of an 8-well plate. Culture the cells in the 8-well plate for 12 to 24 h before imaging.</li>
	<li>Add BODIPY-C12, LysoTracker Green or any other BODIPY conjugate at a final concentration of 100&#8201;nM (stock solutions in dimethyl sulfoxide [DMSO]) 10&#8201;min prior to imaging. This time can vary based on the desired experiment. 
	NOTE: Imaging can be performed at ambient temperature (23 &#176;C) using live cell imaging solution. However, imaging at 37 &#176;C and 5% CO2 with non-fluorescent DMEM mixed with 10% fetal bovine serum, 4 mM glutamine, 1 mM sodium pyruvate and 1% penicillin-streptomycin antibiotics is preferred to keep cells closer to physiological conditions and to make biological conclusions.</li>
</ol>
3. Equipment preparation
<ol>
	<li>Mount the appropriate filter sets in the emission path based on the emission color of BODIPY being used. 
	NOTE: A quad-band dichroic mirror (zt/405/488/561/640rdc) first separates the excitation from the emission light. The green emission (525 nm) and red emission (595 nm) are then split by a dichroic long-pass beam splitter (T562lpxr) followed by band-pass filters ET525/50 in the green channel and ET 595/50 in the red channel. The two channels are then projected to different areas of the same camera chip. Similarly, the red emission (595 nm) and the far-red emission are first split by a dichroic long-pass beam splitter (FF652-Di01) followed by band-pass filters ET 610/75 in the red and FF731/137 in the far-red channel.</li>
	<li>Turn on the microscope, microscope stage, lasers (488 nm, 561 nm) and camera. Here an inverted microscope with a perfect focus system and an EMCCD camera cooled to -68 &#176;C are used.</li>
	<li>Add a drop of immersion oil on the microscope objective.</li>
	<li>Open the Hal4000 software (see the Table of Materials) that controls the LED light for bright field imaging, laser powers, laser shutters and camera settings for imaging. Set the EMCCD gain to 30 and the camera temperature to -68 &#176;C. Prepare the camera and corresponding software to record movies at 20 Hz. 
	NOTE: This technique applies to any wide-field microscope capable of photo-activated localization microscopy (PALM) and stochastic optical reconstruction microscopy (STORM) imaging. Corresponding software may vary.</li>
	<li>Turn on the microscope stage heater and set it to a temperature of 37 &#176;C and to a CO2 level of 5%. Adjust the objective correction collar accordingly.</li>
	<li>Mount the sample on the microscope stage and focus until the focusing system engages. Move the stage using the stage controller until healthy cells appear in the field of view. 
	NOTE: For imaging with yeast cells at room temperature, there is no need to turn on the heater or&#160;CO2 control.</li>
	<li>Turn on the appropriate lasers for the excitation of monomers as well as dimers. For BODIPY green or LysoTracker green, we use a 561 nm laser to excite DII states for SMLM and a 488 nm laser to excite monomers for conventional fluorescence. 
	NOTE: For BODIPY red, use a 640 nm laser to excite DII states for SMLM and a 561 nm laser to excite the monomers for conventional fluorescence. For BODIPY red, adjust the 561 nm and 640 nm laser powers to visualize bulk fluorescence in the red channel and single molecule bursts in the far-red channel. The typical power for 561 nm is ~0.06 W/cm2 and ~5 kW/cm2 for 640 nm. For BODIPY green, expect to also see conventional fluorescence in the green emission channel under 561 nm excitation. For BODIPY red, expect to see conventional fluorescence in the red channel under 640 nm excitation. This signal arises from anti-Stokes emission, which becomes useful for monomer/dimer co-localization images with continuous laser excitation.</li>
</ol>
4. Data acquisition
<ol>
	<li>Load laser shutter sequences for the excitation of monomers as well as dimers. 
	NOTE: We typically use nine single molecule excitation frames at 561 nm&#160;followed by one conventional excitation frame at 488 nm. This offers a brighter conventional fluorescence signal and avoids leaking of another 488 nm excitable fluorophore such as green fluorescent protein (GFP) into the red single-molecule detection channel in multi-color imaging applications. Alternatively, turn on the 561 nm laser continuously and rely on the anti-Stokes emission for conventional images in the shorter wavelength channel.</li>
	<li>Tune the laser powers such that single molecule fluorescence bursts are detected in the red-shifted emission channel under 561 nm excitation, and conventional fluorescence appears in the green emission channel with 488 nm excitation. Typical laser powers of the 561 nm laser will be around 0.8-1 kW/cm2 for SMLM, and 0.035-0.07 W/cm2 for the 488 nm laser in the conventional fluorescence imaging mode.</li>
	<li>Choose a destination folder for movies and record 5,000-20,000 acquisition frames to collect enough localizations for reconstructing super-resolution images.</li>
	<li>Move to different fields of view and repeat the steps above to collect data from more cells.</li>
</ol>
5. Data analysis and single-molecule tracking
<ol>
	<li>Load the movie into a SMLM analysis software. 
	NOTE: Any software18 can be used. We use INSIGHT (see the Table of Materials) and cross-validate the results using the ThunderSTORM19 plugin for imageJ (Fiji).</li>
	<li>Visually screen the movie and adjust contrast settings such that single-molecule fluorescence&#160;blinking is visible. If needed restrict the region or the frame range for SMLM data analysis if parts of the sample are continuously fluorescing.</li>
	<li>Set single molecule identification parameters for fitting with 2D Gaussian PSFs (ROI: 7 x 7 pixels with pixel size 160 nm, width 260-650 nm, height &#62; 50 photons). Visually screen through some example frames to check the identification parameters and to reliably detect the distinct single molecule fluorescence&#160;bursts (see Figure 1C). 
	NOTE: Certain identification parameters such as height and width can be slightly adjusted to optimize the recognition of visually perceived single molecule fluorescence signals.</li>
	<li>Perform SMLM image analysis with the optimized identification parameters and then render each single molecule as a 2D Gaussian whose width is weighted by the inverse square root of the detected number of photons.</li>
	<li>Assess the quality of the data. Use restricted frame ranges to observe single molecule distributions at more specific instances in time. This can account for organelle movement during data acquisition.</li>
	<li>To further analyze the spatial distribution and dynamics of the molecule distributions, export the obtained molecule list containing the coordinates, frames of appearance, photons, widths and heights of the&#160;localizations. Import the molecule list in custom written analysis procedures.</li>
	<li>For obtaining spatial information of the single molecule distribution, calculate the radial distribution function &#961;(r), which represents the density of localizations as a function of the radial distance20. To obtain &#961;(r), calculate unique pair-wise distances of all localizations, construct the histogram with bins centered at ri with height H(ri) and with a width dr and divide by 2&#960;ri*dr; (&#961;(ri) = H(ri)/&#960;(ri+dr)2-ri2). The radial distribution function can then be used to quantify and compare the degree of clustering as well as the characteristic size of clusters.</li>
	<li>To obtain dynamic information about the diffusion of molecules, link localizations that appear for example within 3 pixels (0.48 &#181;m) in consecutive data acquisition frames to create single molecule traces. 
	NOTE: The linking distance will depend on the diffusion of molecules and the density of localizations. The maximum linking distance can be estimated by analyzing the density of localizations in each frame21,22. The average density was determined to be 0.043 localizations per &#181;m2; thus a 0.48 &#181;m radius was within a low enough density to ensure that different molecules were not linked together.</li>
	<li>Average the displacements for different lag times &#916;t from multiple traces lasting at least three lag times to create a mean squared displacement (MSD) vs. &#916;t plot. Fit the MSD vs. &#916;t curve with the equation MSD = 4D&#916;t + &#963;2 to calculate the average diffusion coefficient D3.</li>
</ol>

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The authors declare no competing interests.

In this protocol, we demonstrated how conventional BODIPY conjugates can be used to obtain SMLM images with an order of magnitude improvement in spatial resolution. This method is based on exploiting previously reported, red-shifted DII states of conventional BODIPY dyes, which transiently form through bi-molecular encounters. These states can be specifically excited and detected with red-shifted wavelengths and are sparse and short-lived enough for SMLM. By tuning the concentration of BODIPY monomers along wi...

Single-molecule localization microscopy (SMLM) techniques such as stochastic optical reconstruction microscopy (STORM) and photo-activated localization microscopy (PALM) have emerged as methods for generating super-resolution images with information beyond Abbe&#8217;s optical diffraction limit1,2 and for tracking the dynamics of single biomolecules3,4. One of the requirements for probes compatible with SMLM is the ability to control the number of active fluorophores at any time to avoid spatial overlap of their point spread functions (PSF). In each of the thousands of data acquisition frames, the location of each fluorescent fluorophores is then determined with ~20 nm precision by fitting its corresponding point-spread function. Traditionally, the on-off blinking of fluorophores has been controlled through stochastic photoswitching1,2,5 or chemically induced intrinsic blinking6. Other approaches include the induced activation of fluorogens upon transient binding to a fluorogen-activating protein7,8 and the programmable binding-unbinding of labeled DNA oligomers in total internal reflection fluorescence (TIRF) or light sheet excitation9. Recently, we reported a novel and versatile labeling strategy for SMLM10 in which previously reported red-shifted dimeric (DII) states of conventional boron di-pyromethane (BODIPY) conjugates11,12,13 are transiently forming and become specifically excited and detected with red-shifted wavelengths.
BODIPYs are widely used dyes with hundreds of variants that specifically label sub-cellular compartments and biomolecules14,15,16. Because of their ease of use and applicability in living cells, BODIPY variants are commercially available for conventional fluorescence microscopy. Here, we describe a detailed and optimized protocol on how the hundreds of commercially available BODIPY conjugates can be used for live-cell SMLM. By tuning the concentration of BODIPY monomers and by optimizing the excitation laser powers, imaging and data analysis parameters, high-quality super-resolution images and single molecule tracking data is obtained in living cells. When used at low concentration (25-100 nM), BODIPY conjugates can be simultaneously used for SMLM in the red-shifted channel and for correlative conventional fluorescence&#160;microscopy in the conventional emission channel. The obtained single molecule data can be analyzed to quantify the spatial organization of immobile structures and to extract the diffusive states of molecules in living cells17. The availability of BODIPY probes in both green and red forms allows for multi-color imaging when used in the right combination with other compatible fluorophores.
In this report, we provide an optimized protocol for acquiring and analyzing live-cell SMLM data using BODIPY-C12, BODIPY (493/503), BODIPY-C12 red and lysotracker-green in multiple colors. We resolve fatty acids and neutral lipids in living yeast and mammalian cells with ~30 nm resolution. We further demonstrate that yeast cells&#160;regulate the spatial distribution of externally added fatty acids depending on their metabolic state. We find that added BODIPY-fatty acids (FA) localize to the endoplasmic reticulum (ER) and lipid droplets (LDs) under fed conditions whereas BODIPY-FAs form non-LD clusters in the plasma membrane upon fasting. We further extend the application of this technique to image lysosomes and LDs in living mammalian cells. Our optimized protocol for SMLM using conventional BODIPY conjugates can be a useful resource to study biological processes at the nanoscale with the myriad available BODIPY conjugates.

<table><tbody><tr><td>BODIPY C12</td><td>ThermoFisher</td><td>D3822</td><td>Green fatty acid analog</td></tr><tr><td>BODIPY C12 Red</td><td>ThermoFisher</td><td>D3835</td><td>Red fatty acid analog</td></tr><tr><td>BODIPY(493/503)</td><td>ThermoFisher</td><td>D3922</td><td>Neutral lipid marker</td></tr><tr><td>Concanavalin A</td><td>Sigma-Aldrich</td><td>C2010</td><td>Cell immobilization on glass surface</td></tr><tr><td>Drop-out Mix Complete w/o nitrogen base</td><td>US Biological</td><td>D9515</td><td>Amino acids for SCD</td></tr><tr><td>Dextrose</td><td>Sigma-Aldrich</td><td>G7021</td><td>Carbon source for SCD</td></tr><tr><td>Eight Well</td><td>Cellvis</td><td>C8-1.58-N</td><td>Chambered Coverglasses</td></tr><tr><td>Eight Well, Lb-Tek II</td><td>Sigma-Aldrich</td><td></td><td>Chambered Coverglasses</td></tr><tr><td>ET525/50</td><td>Chroma</td><td></td><td>Bandpass filter</td></tr><tr><td>ET595/50</td><td>Chroma</td><td></td><td>Bandpass filter</td></tr><tr><td>ET610/75</td><td>Chroma</td><td></td><td>Bandpass filter</td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>Gibco</td><td>26140079</td><td>Serum</td></tr><tr><td>FF652</td><td>Semrock</td><td></td><td>Beam splitter</td></tr><tr><td>FF731/137</td><td>Semrock</td><td></td><td>Bandpass filter</td></tr><tr><td>FluoroBrite DMEM</td><td>ThermoFisher</td><td>A1896701</td><td>Cell culture medium</td></tr><tr><td>Hal4000</td><td>Zhuang Lab, Harvard University</td><td></td><td>Data acquisition software</td></tr><tr><td>Ixon89Ultra DU-897U</td><td>Andor</td><td></td><td>EMCCD camera for photon detection</td></tr><tr><td>Laser 405, 488, 561, 640 nm</td><td>CW-OBIS</td><td></td><td>Lasers for excitation</td></tr><tr><td>Insight3</td><td>Zhuang Lab, Harvard University</td><td></td><td>Single molecule localization software</td></tr><tr><td>L-Glutamine</td><td>Gibco</td><td>25030-081</td><td>Amino acid required for cell culture</td></tr><tr><td>live-cell imaging solution</td><td>ThermoFisher</td><td>A14291DJ</td><td>Imaging buffer</td></tr><tr><td>Lysotracker Green</td><td>ThermoFisher</td><td>L7526</td><td>Bodipy based lysosome marker</td></tr><tr><td>Mammalian ATCC U2OS cells (Manassas, VA)</td><td>Dr. Jochen Mueller (University of Minnesota)</td><td></td><td></td></tr><tr><td>Nikon-CFI Apo 100 1.49 N.A</td><td>Nikon</td><td></td><td>Oil immersion objective</td></tr><tr><td>Penicillin streptomycin</td><td>Gibco</td><td>15140-122</td><td>Antibiotics</td></tr><tr><td>Sodium Pyruvate</td><td>Gibco</td><td>11360-070</td><td>Supplement for cell culture</td></tr><tr><td>T562lpxr</td><td>Chroma</td><td></td><td>Beam splitter</td></tr><tr><td>Trypsin-EDTA</td><td>Gibco</td><td>15400-054</td><td>Dissociation of adherent cell</td></tr><tr><td>W303 MATa strain</td><td>Horizon-Dharmacon</td><td>YSC1058</td><td>Parental yeast strain</td></tr><tr><td>Yeast Nitrogen Base</td><td>Sigma-Aldrich</td><td>Y1250</td><td>Nitrogen base without amino-acids</td></tr><tr><td>zt405/488/561/640rdc</td><td>Chroma</td><td></td><td>Quadband dichroic mirror</td></tr></tbody></table>

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;

Here, we present an optimized sample preparation, data acquisition and analysis procedure for SMLM using BODIPY conjugates based on the protocol above (Figure 1A). To demonstrate an example of the workflow for acquiring and analyzing SMLM data, we employ BODIPY (493/503) in yeast to resolve LDs below the optical diffraction limit (Figure 1B-F). Examples of the different multi-color imaging modes of BODIPY in conjunction with other probes such as...

Watch this Scientific Journal Video about Conventional BODIPY Conjugates for Live-Cell Super-Resolution Microscopy and Single-Molecule Tracking at JoVE.com

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

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 ...

conventional-bodipy-conjugates-for-live-cell-super-resolution

Universidad San Sebastian

Research

JoVE Journal

Biochemistry

8.2K Views.  University of Minnesota, Twin Cities, Physics and Nanotechnology (PAN). 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.

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

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 ...

Visualizing Protein-DNA Interactions in Live Bacterial Cells Using Photoactivated Single-molecule Tracking

Protein-DNA interactions are at the heart of many fundamental cellular processes. For example, DNA replication, transcription, repair, and chromosome organization are governed by DNA-binding proteins that recognize specific DNA structures or sequences. In vitro experiments have helped to generate detailed models for the function of many types of DNA-binding proteins, yet, the exact mechanisms of these processes and their organization in the complex environment of the living cell remain far less understood. We recently introduced a method for quantifying DNA-repair activities in live Escherichia coli cells using Photoactivated Localization Microscopy (PALM) combined with single-molecule tracking. Our general approach identifies individual DNA-binding events by the change in the mobility of a single protein upon association with the chromosome. The fraction of bound molecules provides a direct quantitative measure for the protein activity and abundance of substrates or binding sites at the single-cell level. Here, we describe the concept of the method and demonstrate sample preparation, data acquisition, and data analysis procedures.

Photoactivated localization microscopy (PALM) combined with single-molecule tracking allows direct observation and quantification of protein-DNA interactions in live Escherichia coli cells.

Protein-DNA interactions are at the heart of many fundamental cellular processes. For example, DNA replication, transcription, repair, and chromosome ...

Immunology and Infection

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation (BiFC-PALM)

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and sensitivity in cells because the resolution of conventional light microscopy is diffraction-limited to ~250 nm. By combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM), PPIs can be visualized in cells with single molecule sensitivity and nanometer spatial resolution. BiFC is a commonly used technique for visualizing PPIs with fluorescence contrast, which involves splitting of a fluorescent protein into two non-fluorescent fragments. PALM is a recent superresolution microscopy technique for imaging biological samples at the nanometer and single molecule scales, which uses phototransformable fluorescent probes such as photoactivatable fluorescent proteins (PA-FPs). BiFC-PALM was demonstrated by splitting PAmCherry1, a PA-FP compatible with PALM, for its monomeric nature, good single molecule brightness, high contrast ratio, and utility for stoichiometry measurements. When split between amino acids 159 and 160, PAmCherry1 can be made into a BiFC probe that reconstitutes efficiently at 37 &#176;C with high specificity to PPIs and low non-specific reconstitution. Ras-Raf interaction is used as an example to show how BiFC-PALM helps to probe interactions at the nanometer scale and with single molecule resolution. Their diffusion can also be tracked in live cells using single molecule tracking (smt-) PALM. In this protocol, factors to consider when designing the fusion proteins for BiFC-PALM are discussed, sample preparation, image acquisition, and data analysis steps are explained, and a few exemplary results are showcased. Providing high spatial resolution, specificity, and sensitivity, BiFC-PALM is a useful tool for studying PPIs in intact biological samples.

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation BiFC-PALM

Protein-protein interactions are visualized in cells with nanometer spatial resolution by combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM). Described here is the use of BiFC-PALM for imaging Ras-Raf interactions in U2OS cells for visualizing the nanoscale clustering and diffusion of individual Ras-Raf complexes.

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and ...

Bioengineering

Super-resolution Imaging of the Bacterial Division Machinery

Bacterial cell division requires the coordinated assembly of more than ten essential proteins at midcell1,2. Central to this process is the formation of a ring-like suprastructure (Z-ring) by the FtsZ protein at the division plan3,4. The Z-ring consists of multiple single-stranded FtsZ protofilaments, and understanding the arrangement of the protofilaments inside the Z-ring will provide insight into the mechanism of Z-ring assembly and its function as a force generator5,6. This information has remained elusive due to current limitations in conventional fluorescence microscopy and electron microscopy. Conventional fluorescence microscopy is unable to provide a high-resolution image of the Z-ring due to the diffraction limit of light (~200 nm). Electron cryotomographic imaging has detected scattered FtsZ protofilaments in small C. crescentus cells7, but is difficult to apply to larger cells such as E. coli or B. subtilis. Here we describe the application of a super-resolution fluorescence microscopy method, Photoactivated Localization Microscopy (PALM), to quantitatively characterize the structural organization of the E. coli Z-ring8.
	PALM imaging offers both high spatial resolution (~35 nm) and specific labeling to enable unambiguous identification of target proteins. We labeled FtsZ with the photoactivatable fluorescent protein mEos2, which switches from green fluorescence (excitation = 488 nm) to red fluorescence (excitation = 561 nm) upon activation at 405 nm9. During a PALM experiment, single FtsZ-mEos2 molecules are stochastically activated and the corresponding centroid positions of the single molecules are determined with &lt;20 nm precision. A super-resolution image of the Z-ring is then reconstructed by superimposing the centroid positions of all detected FtsZ-mEos2 molecules.
	Using this method, we found that the Z-ring has a fixed width of ~100 nm and is composed of a loose bundle of FtsZ protofilaments that overlap with each other in three dimensions. These data provide a springboard for further investigations of the cell cycle dependent changes of the Z-ring10 and can be applied to other proteins of interest.

We describe a super-resolution imaging method to probe the structural organization of the bacterial FtsZ-ring, an essential apparatus for cell division. This method is based on quantitative analyses of photoactivated localization microscopy (PALM) images and can be applied to other bacterial cytoskeletal proteins.

Bacterial cell division requires the coordinated  assembly of more than ten essential proteins at midcell1,2. Central  to this process is the formation of ...

Biology

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 ...

Three-dimensional Super Resolution Microscopy of F-actin Filaments by Interferometric PhotoActivated Localization Microscopy (iPALM)

Fluorescence microscopy enables direct visualization of specific biomolecules within cells. However, for conventional fluorescence microscopy, the spatial resolution is restricted by diffraction to ~ 200 nm within the image plane and &#62; 500 nm along the optical axis. As a result, fluorescence microscopy has long been severely limited in the observation of ultrastructural features within cells. The recent development of super resolution microscopy methods has overcome this limitation. In particular, the advent of photoswitchable fluorophores enables localization-based super resolution microscopy, which provides resolving power approaching the molecular-length scale. Here, we describe the application of a three-dimensional super resolution microscopy method based on single-molecule localization microscopy and multiphase interferometry, called interferometric PhotoActivated Localization Microscopy (iPALM). This method provides nearly isotropic resolution on the order of 20 nm in all three dimensions. Protocols for visualizing the filamentous actin cytoskeleton, including specimen preparation and operation of the iPALM instrument, are described here. These protocols are also readily adaptable and instructive for the study of other ultrastructural features in cells.

Three-dimensional Super Resolution Microscopy of F-actin Filaments by Interferometric PhotoActivated Localization Microscopy iPALM

We present a protocol for the application of interferometric PhotoActivated Localization Microscopy (iPALM), a 3-dimensional single-molecule localization super resolution microscopy method, to the imaging of the actin cytoskeleton in adherent mammalian cells. This approach allows light-based visualization of nanoscale structural features that would otherwise remain unresolved by conventional diffraction-limited optical microscopy.

Fluorescence microscopy enables direct visualization of specific biomolecules within cells. However, for conventional fluorescence microscopy, the spatial ...

Application of High-speed Super-resolution SPEED Microscopy in Live Primary Cilium

The primary cilium is a microtubule-based protrusion on the surface of many eukaryotic cells and contains a unique complement of proteins that function critically in cell motility and signaling. Since cilia are incapable of synthesizing their own protein, nearly 200 unique ciliary proteins need to be trafficked between the cytosol and primary cilia. However, it is still a technical challenge to map three-dimensional (3D) locations of transport pathways for these proteins in live primary cilia due to the limitations of currently existing techniques. To conquer the challenge, recently we have developed and employed a high-speed virtual 3D super-resolution microscopy, termed single-point edge-excitation sub-diffraction (SPEED) microscopy, to determine the 3D spatial location of transport pathways for both cytosolic and membrane proteins in primary cilia of live cells. In this article, we will demonstrate the detailed setup of SPEED microscopy, the preparation of cells expressing fluorescence-protein-labeled ciliary proteins, the real-time single-molecule tracking of individual proteins in live cilium and the achievement of 3D spatial probability density maps of transport routes for ciliary proteins.

Recently we mapped the three-dimensional (3D) spatial locations of transport routes for various proteins translocating inside primary cilia in live cells. Here this paper details the experimental setup, the process of biological samples and the data analyses for the 3D super-resolution fluorescence imaging approach newly applied in live primary cilia.

The primary cilium is a microtubule-based protrusion on the surface of many eukaryotic cells and contains a unique complement of proteins that function ...

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.

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 ...

Specific Labeling of Mitochondrial Nucleoids for Time-lapse Structured Illumination Microscopy

Mitochondrial nucleoids are compact particles formed by mitochondrial DNA molecules coated with proteins. Mitochondrial DNA encodes tRNAs, rRNAs, and several essential mitochondrial polypeptides. Mitochondrial nucleoids divide and distribute within the dynamic mitochondrial network that undergoes fission/fusion and other morphological changes. High resolution live fluorescence microscopy is a straightforward technique to characterize a nucleoid&#39;s position and motion. For this technique, nucleoids are commonly labeled through fluorescent tags of their protein components, namely transcription factor a (TFAM). However, this strategy needs overexpression of a fluorescent protein-tagged construct, which may cause artifacts (reported for TFAM), and is not feasible in many cases. Organic DNA-binding dyes do not have these disadvantages. However, they always show staining of both nuclear and mitochondrial DNAs, thus lacking specificity to mitochondrial nucleoids. By taking into account the physico-chemical properties of such dyes, we selected a nucleic acid gel stain (SYBR Gold) and achieved preferential labeling of mitochondrial nucleoids in live cells. Properties of the dye, particularly its high brightness upon binding to DNA, permit subsequent quantification of mitochondrial nucleoid motion using time series of super-resolution structured illumination images.

The protocol describes specific labeling of mitochondrial nucleoids with a commercially available DNA gel stain, acquisition of time lapse series of live labeled cells by super-resolution structured illumination microscopy (SR-SIM), and automatic tracking of nucleoid motion.

Mitochondrial nucleoids are compact particles formed by mitochondrial DNA molecules coated with proteins. Mitochondrial DNA encodes tRNAs, rRNAs, and ...