This work was supported by a Hypatia fellowship from the Radboud University Medical Center, a Career Development Award from the Human Frontier Science Program, the Gravitation Programme 2013 from the Netherlands Organization for Scientific Research (NWO; ICI-024.002.009), a VIDI grant from NWO (ALW VIDI 864.14.001), and a Starting Grant from the European Research Council (ERC) under the European Union&#39;s Seventh Framework Programme (Grant Agreement Number 336479).

1. Preparation of the Microscope Samples
<ol>
	<li>Expression of Qa-SNARE syntaxin 4 fused to mCitrine (syntaxin 4-mCitrine; donor fluorophore) and the R-SNARE VAMP3 fused to mCherry (VAMP3-mCherry) 
	NOTE: Other SNAREs with fluorescent proteins fused to their C-terminal transmembrane helices can also be used. Instead of mCitrine-mCherry, other donor-acceptor pairs of spectrally separated fluorophores can also be used (e.g., CFP-YFP).
	<ol>
		<li>Grow HeLa cells and divide 900,000 HeLa cells over three 35 mm-diameter glass bottom dishes suitable for microscopy. 
		NOTE: In principle, this protocol can be adapted for other eukaryotic cell types and microscopy dishes.
		<ol>
			<li>Maintain HeLa cell cultures in T75 flasks with 10 mL of high glucose Dulbecco&#39;s Modified Eagle&#39;s medium (DMEM), supplemented with 10% fetal calf serum (FCS) and 1% antibiotic-antimycotic (containing amphotericin B, penicillin, and streptomycin) at 37 &#176;C and 5% CO2 in a cell culture incubator.</li>
			<li>Replenish the medium twice a week and split the cells 1:10 once a week or when the cells reach 85-90% confluency to new T75 flasks (500,000 cells/flask, on average).</li>
			<li>Remove the DMEM and wash the HeLa monolayers twice with 8 mL of sterile phosphate-buffered saline (PBS) for 3 min.</li>
			<li>Remove the PBS and add 2 mL of PBS with 2 mM ethylenediaminetetraacetic acid (EDTA).</li>
			<li>Incubate the cells for 5 min at 37 &#176;C and 5% CO2 in cell culture incubator and subsequently lightly agitate the flask to separate HeLa cells from the bottom of the flask.</li>
			<li>Flush the cells out with 10 mL of medium, transfer the suspension to a 15 mL tube and remove a 10 &#181;L aliquot for counting.</li>
			<li>Dilute the aliquot 1:1 with 0.4% trypan blue stain and count the cells with a B&#252;rker hemocytometer9. 
			NOTE: Count two 4 x 4 squares and multiple the cell number by 10,000 for the number of cells/mL.</li>
			<li>After cell counting, take 900,000 cells from the 15 mL tube, divide them over the 3 glass bottom dishes (see step 1.1.1), and prepare for transfection.</li>
		</ol>
		</li>
		<li>Transfect the cells by electroporation10 (see Table of Materials) with syntaxin 4-mCitrine construct (donor only, Sample #1), syntaxin 4-mCitrine together with VAMP3-mCherry (Sample #2), and syntaxin 3 fused to both mCitrine and mCherry (Sample #3). 
		NOTE: Other cell transfection methods can also be used11. The tandem construct in sample #3 is a positive control for estimating the shortest lifetime expectable (i.e., maximal expectable FRET).</li>
	</ol>
	</li>
	<li>Culture the cells overnight in high glucose DMEM supplied with 10% FCS and 1% antibiotic-antimycotic at 37 &#176;C with 5% CO2 in a cell culture incubator.</li>
	<li>Aspirate DMEM and wash the cells once with live cell imaging medium (140 mM NaCl, 2.5 mM KCl, 1.8 mM CaCl2, 1 mM MgCl2, 20 mM HEPES, pH = 7.4, mOsm = 300; see Table of Materials) for 2 min and subsequently place the cells in fresh imaging medium.</li>
</ol>
2. Recording of the FLIM Data
<ol>
	<li>Place Sample #2 under a time-domain FLIM confocal microscope with a pulsed excitation source for the donor fluorophore (i.e., mCitrine) and time-resolved data acquisition. Keep the cells at 37 &#176;C with a heated stage. 
	NOTE: The confocal microscope used in this protocol is equipped with a 63X 1.20 NA water immersion objective, a pulsed white light laser (80 MHz pulsing, &#60; 100 ps pulse duration), a photon multiplier tube (PMT), and a Time-Correlated Single Photon Counting (TCSPC) system (see Table of Materials for specific setup used). Other FLIM microscopes can also be used (e.g., single-photon avalanche diodes (SPAD) instead of PMTs, single-wavelength pulsed lasers instead of a white-light pulsed lasers).</li>
	<li>Select a cell with visible expression of the mCitrine and mCherry-labeled SNARE proteins and record a confocal image of 256 &#215; 256 pixels with simultaneous excitation of both fluorophores prior to each FLIM measurement. A pixel step of about 2-fold smaller than the diffraction limited spatial resolution of the microscope (~ 200 nm) should be used. For mCitrine, excite at 516 nm and collect emission from 521-565 nm; for mCherry, excite at 610 nm and collect emission from 613-668 nm. 
	NOTE: Do not position the imaging plane too close to the bottom of the dish (within ~ 2 &#181;m distance), as this will result in a reflection peak in the fluorescence lifetime histogram. Simultaneous excitation of the donor and acceptor fluorophores is preferred, as this allows to overcome the potential problem of sample movement. However, sequential excitation can be used as well.</li>
	<li>Record a FLIM image by clicking Run FLIM with the FLIM image being recorded with the same dimensions and spatial resolution as the confocal image recorded at step 2.2. Record the image at least 50,000 photons for whole-cell FLIM analysis, or at least 400 photons/pixel. Only excite the mCitrine donor fluorophore and not the mCherry acceptor fluorophore, and thus excite at 516 nm and collect emission from 521-565 nm.</li>
	<li>Repeat steps 2.1-2.3 for multiple cells and for the other Sample #1 (donor only) and Sample #3 (the mCitrine-mCherry tandem construct).</li>
	<li>Record an instrument response function (IRF). Tune the monochromator of the emission detector to the excitation wavelength (e.g., record emission from 510-550 nm) and then record a FLIM image with the back scattering from a clean glass cover slip by clicking the button Run FLIM. Use the same excitation wavelength (516 nm) and laser power as used for recording the cell data at steps 2.1-2.4. 
	NOTE: If multiple peaks are visible in the IRF or if the emission monochromators cannot be tuned, the IRF can also be measured with a solution of fluorescent dye with ultrafast lifetime, for instance by fluorescence quenching of an organic dye in a saturated aqueous solution of potassium iodide. Moreover, it is not strictly required to record an IRF, because the decay slope of the fluorescence histograms can also be fitted without deconvolution of the IRF. However, an IRF enables to correct for the timing characteristics of the photon detector used and thereby makes the fits of the lifetime histograms more accurate.</li>
</ol>
3. Conversion of the Photon Recordings to FLIM Images
<ol>
	<li>Download and install the PT32ICS conversion software5. 
	NOTE: The lifetime data can also be analyzed by other software, including software from several microscope vendors.</li>
	<li>Configure the PT32ICS software.
	<ol>
		<li>Set the Size to 256 &#215; 256 pixels image size via Settings | Size.</li>
		<li>Set the Channel to 2 via Settings | Size. 
		NOTE: The channel of the mCitrine emission is exclusively dependent on the configuration of the microscope. A wrong channel will result in a FLIM image with no photons (i.e., black image) and as a result an empty &#39;LifetimeTable.txt&#39;-file. The correct channel can be identified by trial-and-error.</li>
		<li>Set Output to ImageJ (xyz) (or TRI2 (xyt)) for generating FLIM images (step 3.4).</li>
	</ol>
	</li>
	<li>Press Convert and load one or more photon traces (the .pt3 files). To select multiple photon traces, keep the Ctrl key pressed. 
	NOTE: The newer 64 bit .ptu format can also be converted.</li>
	<li>Ensure that the PT32ICS software generates the following files in the same folder as where the pt3 files are saved: one photon stack per .pt3 photon trace in image cytometry standard format (.ics), one bitmap file per .pt3 photon trace (.bmp), a text file per conversion (_Report.txt) containing information on the photon statistics (i.e., the total number of photons in each .pt3 photon trace, the time resolution of detector), and a second text file per conversion (_LifetimeTable.txt) containing the total fluorescence lifetime histograms in tab-delimited format for each .pt3 photon trace. 
	NOTE: The .ics file can be used for single pixel fitting in order to generate FLIM images, for instance with TRI2 software12,13 or the SLIM curve fitting library in FIJI ImageJ14,15. This file can also be used for Phasor analysis16, for instance with the Time Gated Phasor plugin for FIJI from Spechron. The bitmap file contains no lifetime information. The second text file is used for the whole-cell FLIM analysis in step 4.</li>
</ol>
4. Fitting of Fluorescence Lifetime Histograms for Whole-cell FLIM Analysis
NOTE: This step (deconvoluted fitting of the IRF) requires software capable of deconvoluted fitting. Fitting the slopes of the fluorescence histograms without deconvolution of the IRF can be done with other software as well.
<ol>
	<li>Open the data analysis software program capable of fitting with deconvolution (Table of Materials) and import the text file containing the histogram for each photon trace (_LifetimeTable.txt) via File | Import | Single ASCII.</li>
	<li>Reorganize the table such that the IRF is in the second column of the table (using Copy and Paste). Place the IRF in the second column B next to the time values in the first column A.</li>
	<li>Determine the quality of the recorded photon traces by selecting all columns by pressing Ctrl + A and selecting Plot | Multi-panel | 9 panel. Do not analyze photon traces with a high reflection peak (Figure 4F).</li>
	<li>Select all columns containing the quality-controlled lifetime histograms that will be fitted as well as the IRF in column B using Ctrl key and load the non-linear fitting via Analysis | Fitting | Nonlinear Curve Fit | Open Dialog.</li>
	<li>Load the deconvoluted fit function (available in the Supplementary File 1) by adding this function to the analysis software (via Category | User Defined | Function | Add). Select the fit function file &#39;FLIM_convoluted_IRF.fdf&#39;. This function fits the fluorescence lifetime histograms with a mono-exponential decay function deconvoluted with the IRF (i.e., the second column B in the table) (Equation 1): 
	<img alt="Equation 1" src="/files/ftp_upload/56745/56745eq1.jpg" />&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(Equation 1) 
	with t the time, &#964; the apparent fluorescence lifetime, A the amplitude, and y0 the offset. 
	NOTE: Another option is to fit the lifetime histograms with a bi-exponential fit function (Equation 2): 
	<img alt="Equation 2" src="/files/ftp_upload/56745/56745eq2.jpg" />&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(Equation 2) 
	By fixing the lifetime of the slow component (&#964;1) to the donor only condition (step 1.1.2: Sample #1) and that of the fast component (&#964;2) to the tandem construct (Sample #3), this allows estimation of the fraction of SNARE proteins in complex (F) from the amplitudes of the slow (A1) and fast components (A2; see Discussion; Equation 3): 
	<img alt="Equation 3" src="/files/ftp_upload/56745/56745eq3.jpg" />&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(Equation 3) 
	The fit function file for deconvoluted bi-exponential fitting &#39;FLIM_convoluted_IRF_biexp.fdf&#39; is available in the Supplementary File 2.</li>
	<li>Select Fitted Curves | X Data Type as Same as Input Data. 
	NOTE: This will result in the appropriate x-axis scaling of the fitted curves.</li>
	<li>Fit the curves by pressing Fit. 
	NOTE: This will convert the fit and generate a &#39;report sheet&#39; in the array with a table containing the fluorescence lifetimes, offsets, and amplitudes. It will also generate a data sheet with the fitted curves and the residuals of the fit.</li>
</ol>

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The authors have nothing to disclose.

This protocol demonstrates the use of FRET-FLIM for visualization of SNARE interactions between syntaxin 4 and VAMP3 in live HeLa cells. Syntaxin 4 is a Qa-SNARE protein predominantly locating at the plasma membrane where it mediates exocytosis1,2,20,21. VAMP3 is an R-SNARE which is mainly described to locate at recycling endosomal compartments and mediates trafficking to other endosomes as well as to the plasma membrane1,2,20. However, the FRET-FLIM assay can be readily adapted for studying other SNARE proteins. The only condition is that these SNAREs contain a C-terminal transmembrane helix, which is the case for most SNARE proteins by far1,2. In addition, the protocol described here can be adapted for visualization of SNARE complexes in any eukaryotic cell type, including plants and yeast. In this protocol, we used the shortening of the fluorescence lifetime of the donor fluorophore as a measure of FRET. As a complementary approach, the lifetime of the acceptor fluorophore could be examined, because the sensitized emission causes a distinct rise phase which provides unambiguous proof that resonance energy transfer occurs.
Currently, the FRET-FLIM technique may be not able to visualize SNARE complexes in lysosomal compartments. For the syntaxin 3-mCitrine-mCherry tandem construct, the mCherry fluorescence can often be found more accumulated in a juxtanuclear area, which likely corresponds to lysosomal compartments, whereas the mCitrine signal is more abundant in the cellular periphery5. A similar juxtanuclear accumulation of mCherry compared to mCitrine was observed, when the same SNARE proteins fused to these fluorescent proteins were co-expressed5. Lysosomes are characterized by an extremely low pH (&#60; 4) and a high activity of proteolytic enzymes. The juxtanuclear accumulation of mCherry is likely caused by a higher resistance of the mCherry fluorophore to lysosomal degradation compared to the mCitrine fluorophore. It is not due to pH-quenching of mCitrine, as juxtanuclear accumulation of mCherry also occurs upon fixation of the cells5. Thus, the FRET-FLIM technique underestimates the amount of FRET in the juxtanuclear (lysosomal) regions and this would require other fluorescent reporter proteins that survive the harsh conditions within the lumens of lysosomes.
FRET-FLIM in principle allows to obtain a (semi-)quantitative estimate of the fraction of SNAREs in complex5. As we explained in this protocol, this requires the fitting of the fluorescence lifetime histograms with double-exponential decay functions (Equation 2), where the amplitude of the fast component is proportional to the fraction of SNAREs in complex (Equation 3). However, such fitting with a two-component model is technically challenging. Fitting with multiple free fit parameters (two fluorescence lifetimes and two amplitudes) requires a very large number of photons, especially since the parameters will influence each other and small errors in the lifetime will affect the amplitudes and vice versa. To overcome these fitting problems, the fluorescence lifetimes of the slow component can be fixed to the lifetime of the donor only condition (i.e., no FRET; only mCitrine present) and that of the fast component to the lifetime of the tandem construct (maximal expectable FRET). However, this should also be interpreted with care, because the fluorescence lifetimes may not be the same as in these control states, and could deviate because of multiple reasons (self-quenching, dipole orientation, variations in the microenvironment). Multiple SNARE complexes within close proximity (&#60; 10 nm) can result in distance-dependent FRET, the same principle that allows FRET to be used as a &#34;molecular ruler&#34;, but in this case, obscures the quantification of SNARE complexes. Moreover, a quantitative estimate is not always meaningful, because the labeled SNAREs compete with endogenous (unlabeled) SNAREs. As a consequence, the expression level of mCherry-labeled SNARE is a main determinant for the percentage FRET5. Because of all these caveats, it is recommended to fit the fluorescent lifetime histograms with a mono-exponential decay function (Equation 1). This has the advantage that it does not require any a priori knowledge of the lifetimes and the resulting apparent average fluorescent lifetime provides a solid measure for SNARE complexing5.
Nevertheless, it is expected that quantitative FRET-FLIM imaging by two-component fitting models will have potent future applications. SNARE-encoding genes within the chromosome can be fused with fluorescent reporter proteins, for example by CRISPR/CAS9. This results in the fluorescent labeling of endogenous SNARE proteins, with endogenous protein levels and no background of unlabeled SNAREs, and thereby allows for a meaningful quantitative estimation of the fraction of SNARE complexes by FRET-FLIM. While the expression levels of endogenous SNAREs might be quite low and give relatively low fluorescent signals, it is expected that a sufficient number of photons can be obtained, especially for the whole cell FLIM (which requires only a few 1,000s of photons). Moreover, these FRET-FLIM measurements can be performed with more sensitive avalanche photodiode detectors which will also result in higher fluorescence signals and better photon statistics.

Membrane trafficking is a central feature of eukaryotic cells, where membrane vesicles bud off from a donor organelle and then move to and fuse with a target organelle1,2. Except for mitochondria, all these membrane fusion steps are catalyzed by members of the SNARE protein family1,2. The SNARE protein family consists of about 36 members in mammalian cells and about 20 members in yeast2. SNARE proteins contain one or two ~52 residues-long, natively unstructured regions, called SNARE-motifs. SNARE proteins are often tethered to membranes by a C-terminal transmembrane helix1,2. SNAREs can be categorized based on the central residues they contribute to the SNARE complex into arginine (R) and glutamine (Q) SNAREs1,2. Membrane fusion is driven by the interaction of 3 or 4 cognate SNAREs that together contribute 4 SNARE-motifs and are distributed over both the donor and acceptor membrane1,2. A SNARE complex consists of one R-SNARE motif and three Q-SNARE motifs (termed Qa, Qb, and Qc). Complex formation starts at the N-termini of the SNARE-motifs, forming a so-called trans-SNARE-complex, and proceeds towards the C-termini, forming a tight &#945;-helical coiled-coil bundle called a cis-SNARE-complex. The complex formation of even a single SNARE complex drags the donor and acceptor membranes together and overcomes the energy barrier for membrane fusion3.
Assigning distinct SNARE complexes to specific transport routes within cells is often technically challenging. Although SNAREs clearly contribute to the specificity of membrane trafficking, they are promiscuous, functionally redundant, and their functions overlap1,2. Because of this, perturbation experiments targeting SNAREs, such as by gene knockout, RNA interference, the introduction of blocking antibodies, or with soluble SNARE fragments acting as dominant negatives, frequently do not result in clear phenotypes as other SNAREs compensate2,4. Moreover, it is difficult to differentiate specific membrane fusion steps from upstream trafficking events, because SNAREs can be involved in multiple transport routes2. Localization studies of SNAREs by microscopy approaches, using immunolabeling or genetic fusion to fluorescent reporter proteins, suffer from the problems that: (i) SNAREs locate to multiple organelles as they often mediate multiple trafficking steps, and (ii) their localizations do not automatically mean they are functionally engaged in SNARE complex formation. Finally, SNARE complexes can be identified using immunoprecipitation experiments using one of the SNAREs as bait and the other SNAREs as targets, but this does not allow assignment of these complexes to specific organelles or trafficking routes. Thus, currently, there is no alternative technique to visualize SNARE complexes with organellar resolution. Immunofluorescence is not able to prove SNARE interactions but can show only the presence or absence of co-localization, while immunoprecipitation only can show SNARE interactions in the whole cell population but not assign the organelles where these interactions occur.
To overcome these limitation, a novel method allowing for the quantitative visualization of SNARE complexes within live cells with organellar resolution was recently developed by Verboogen et al.5 This method is based on the expression of pairs of SNAREs with spectrally shifted fluorescent proteins fused C-terminally to their transmembrane helices. After completion of membrane fusion and formation of a cis-SNARE complex, these fluorophores at the C-termini of the transmembrane helices are immediately juxtaposed to each other. The fluorophores are then well within the F&#246;rster distance (typically &#60;5 nm), resulting in FRET from the green-shifted donor fluorophore to the red-shifted acceptor fluorophore5,6. FRET results in quenching of the donor fluorophore and an increased emission of the acceptor fluorophore that can be measured from the ratios of the donor and acceptor emission (ratiometric FRET). However, ratiometric FRET between two different molecules is challenging, because of fluorescence cross-talk and different levels of donor and acceptor SNAREs at different organelles and among cells7,8. FRET can also be measured from the fluorescence lifetime, which is the time between the excitation and the emission of a photon. If the donor fluorophore can release its energy by FRET, this competing process result in an apparent shortening of the fluorescence lifetime. This can be measured by FLIM7,8. Lifetime FRET is much more robust than ratiometric FRET for measuring interactions between two different molecules, as the fluorescence lifetime is an intrinsic property of a fluorophore and is insensitive to its concentration. Moreover, FRET is by approximation quantitative, because the efficiency of FRET is inversely proportional to the sixth power of the distance between the donor and acceptor fluorophores (essentially a step function). Therefore, by fitting the fluorescence lifetime histograms recorded by FLIM with a double-component decay model, FRET-FLIM allows for the (semi-)quantitative estimation of the fraction of SNARE molecules engaged in SNARE complex formation5.
Recently, this FRET-FLIM method was used by Verboogen et al. to visualize SNARE complex formation in primary dendritic cells of the immune system5. It was shown that upon encountering a pathogenic stimulus, dendritic cells reroute their membrane trafficking accompanied by an increased complexing of the R-SNARE vesicle-associated membrane protein (VAMP) 3 with the Qa-SNARE syntaxin 4 specifically at the plasma membrane. This increased SNARE complex formation is likely required to meet the increased secretory capacity for the secretion of inflammatory cytokines such as interleukin-65. This protocol describes the experimental steps needed for the acquisition of FRET-FLIM data for the visualization and (semi-)quantitative measurement of SNARE complexes. It is explained how to fit the whole-cell fluorescence lifetime histograms with mono- and bi-exponential decay functions, resulting in the apparent fluorescence lifetime as a quantitative estimate to SNARE interactions. In this protocol, the widely used HeLa cell line is used as an example, but the method can be readily extended to study SNARE complexes in other eukaryotic cells.

<table><tbody><tr><td>Plasmid DNA &#039;syntaxin 4-mCitrine&#039;</td><td>Addgene</td><td>ID 92422</td><td>Other SNAREs with fluorescent proteins fused to their C-terminal transmembrane helices can also be used. Instead of mCitrine-mCherry, other donor-acceptor pairs of spectrally separated fluorophores can also be used (e.g., CFP-YFP).</td></tr><tr><td>Plasmid DNA &#039;VAMP3-mCherry&#039;</td><td>Addgene</td><td>ID 92423</td><td>Other SNAREs with fluorescent proteins fused to their C-terminal transmembrane helices can also be used. Instead of mCitrine-mCherry, other donor-acceptor pairs of spectrally separated fluorophores can also be used (e.g., CFP-YFP).</td></tr><tr><td>Plasmid DNA &#039;syntaxin 3-mCitrine-mCherry&#039;</td><td>Addgene</td><td>ID 92426</td><td>Positive control for maximum achievable FRET.</td></tr><tr><td>Hela cells</td><td></td><td></td><td></td></tr><tr><td>35 mm glass bottom dishes&amp;nbsp;</td><td>Willco Wells</td><td>HBST-3522</td><td>Other live cell imaging chambers will work as a substitute</td></tr><tr><td>Dulbecco&#039;s Modified Eagle Medium</td><td>Gibco, Life Technologies</td><td>31966-021</td><td></td></tr><tr><td>Fetal calf serum</td><td>Greiner Bio-one</td><td>758093</td><td></td></tr><tr><td>Antibiotic-antimycotic solution</td><td>Gibco, Life Technologies</td><td>15240-062</td><td>Pen/Strep will work as a substitute</td></tr><tr><td>Live cell imaging medium</td><td>Thermo Fisher Scientific</td><td>A14291DJ</td><td>Any other live cell imaging solution will work, as long as fluorescence from the medium is prevented</td></tr><tr><td>Leica SP8 confocal microscope with a 63x 1.20 NA water immersion objective</td><td>Leica</td><td>SP8</td><td>Other confocal microscopes capable of time-domain FLIM can also be used</td></tr><tr><td>Pulsed white light laser</td><td>Leica</td><td>SP8</td><td>Other pulsed laser sources can also be used</td></tr><tr><td>Time-Correlated Single Photon Counting (TCSPC) system</td><td>PicoQuant</td><td>PicoHarp 300</td><td></td></tr><tr><td>PT32ICS conversion software</td><td></td><td></td><td>Available at the &#039;Software&#039;-section of www.membranetrafficking.com</td></tr><tr><td>data analysis software programme capable of deconvolution</td><td>Originlabs</td><td>OriginPro 2016</td><td></td></tr><tr><td>Fiji ImageJ</td><td></td><td></td><td></td></tr><tr><td>Custom-made Fiji ImageJ macro for convolution of FLIM image with Intensity</td><td>Fiji ImageJ</td><td></td><td>Available at the &#039;Software&#039;-section of www.membranetrafficking.com</td></tr><tr><td>B&amp;uuml;rker Haemocytometer</td><td>VWR</td><td>630-1541</td><td></td></tr><tr><td>HeLa cells</td><td>ATCC</td><td>ATCC&amp;nbsp;CCL-2</td><td></td></tr><tr><td>PBS</td><td>B Braun Melsungen&amp;nbsp; AG</td><td>362 3140</td><td></td></tr><tr><td>EDTA 2 mM</td><td>Merck</td><td>108417</td><td>CAS: 60-00-4</td></tr><tr><td>15 mL tubes</td><td>Greiner Bio-one</td><td>188271</td><td></td></tr><tr><td>Trypan blue</td><td>Sigma Aldrich</td><td>93595</td><td>CAS: 72-57-1</td></tr><tr><td>NEON cell electroporation device</td><td>Thermo Fisher Scientific</td><td>MPK5000S</td><td></td></tr></tbody></table>

visualizing intracellular snare trafficking by fluorescence lifetime imaging microscopy

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze membrane fusion within eukaryotic cells. The SNARE protein family consists of about 36 different members. Specific intracellular transport routes are catalyzed by specific sets of 3 or 4 SNARE proteins that thereby contribute to the specificity and fidelity of membrane trafficking. However, studying the precise function of SNARE proteins is technically challenging, because SNAREs are highly abundant and functionally redundant, with most SNAREs having multiple and overlapping functions. In this protocol, a new method for the visualization of SNARE complex formation in live cells is described. This method is based on expressing SNARE proteins C-terminally fused to fluorescent proteins and measuring their interaction by F&#246;rster resonance energy transfer (FRET) employing fluorescence lifetime imaging microscopy (FLIM). By fitting the fluorescence lifetime histograms with a multicomponent decay model, FRET-FLIM allows (semi-)quantitative estimation of the fraction of the SNARE complex formation at different vesicles. This protocol has been successfully applied to visualize SNARE complex formation at the plasma membrane and at endosomal compartments in mammalian cell lines and primary immune cells, and can be readily extended to study SNARE functions at other organelles in animal, plant, and fungal cells.

The rationale of the assay for measuring SNARE interactions by FRET-FLIM is shown in Figure 1. The C-termini of the transmembrane helices of cognate SNARE proteins are fused to a pair of spectrally shifted fluorescent proteins (e.g., mCitrine and mCherry). The formation of a cis-SNARE complex upon membrane fusion results in these fluorescent proteins becoming immediately juxtaposed to each other and FRET. Figure 2 shows representative confocal images of HeLa cells expressing fluorescently-labeled SNARE proteins. Shown are a cell expressing syntaxin 4-mCitrine (donor fluorophore) with VAMP3-mCherry (acceptor fluorophore), as well as control conditions of cells expressing only the syntaxin 4-mCitrine construct (donor only; no FRET) and syntaxin 3 fused to both mCitrine and mCherry in tandem (maximal expectable FRET). Figure 3 shows the accompanying fluorescence lifetime and FLIM images generated by fitting the lifetime histograms of each pixel with mono-exponential decay functions (Figure 3A-B) and bi-exponential decay functions (Figure 3C-D). Figure 4A shows the IRF of our setup measured by back scattering of a microscope cover glass. In Figure 4B-E, representative fluorescence lifetime histograms of the whole cell FLIM analysis together with accompanying fit curves for mono-exponential decay functions are shown. Figure 4F shows a fluorescent lifetime histogram of an experiment imaged too close to the surface of the microscope cover glass, resulting in a prominent reflection peak. Figure 4G shows a lifetime histogram with representative fit with a bi-exponential decay function.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56745/56745fig1.jpg" /> 
Figure 1: Scheme of the rationale for visualizing SNARE complexes by FRET. (A) Structural model of the neuronal SNAREs (protein database 3HD717) vesicle-associated membrane protein (VAMP) 2 (blue; R), syntaxin 1 (red; Qa-SNARE), and SNAP25 (green; contains both a Qb- and Qc-SNARE motif). The C-terminus of the transmembrane helix of syntaxin-1 is conjugated to mCitrine (donor fluorophore; protein database 3DQ118). The C-terminus of the transmembrane helix of VAMP2 is conjugated to mCherry (acceptor fluorophore; protein database 2H5Q19). (B) Scheme of SNARE mediated membrane fusion resulting in FRET. After membrane fusion by formation of a cis-SNARE complex, the SNAREs are immediately juxtaposed to each other resulting in FRET. <a href="https://cloudflare.jove.com/files/ftp_upload/56745/56745fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56745/56745fig2.jpg" /> 
Figure 2: Expression of SNARE proteins fused to fluorescent proteins. First column: donor fluorophore, excited at 516 nm. Second column: acceptor fluorophore, excited at 610 nm. (A) Donor only condition. Representative confocal image of a HeLa cell expressing syntaxin 4-mCitrine (donor fluorophore; green in merge). The acceptor channel (second column) shows the fluorescence cross-talk. (B) Negative control (no FRET). Representative confocal image of a HeLa cell expressing both syntaxin 4-mCitrine with VAMP3-mCherry (acceptor fluorophore; magenta in merge). (C) Positive control (maximal expectable FRET). Representative confocal image of a HeLa cell expressing the syntaxin 3-mCitrine-mCherry tandem construct. Note that the fluorescence signal of the mCitrine and mCherry signals does not completely overlap, likely because of a lower resistance of mCitrine to lysosomal degradation compared to mCherry (see Discussion section). BF: bright field. Scale bars, 20 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56745/56745fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56745/56745fig3.jpg" /> 
Figure 3: FLIM images of SNARE complex formation. (A) Representative fluorescence lifetime images of the cells shown in Figure 2. The images were generated by first converting the photon traces recorded by a TCSPC system (.pt3) to the image cytometry standard (.ics) using the PT32ICS software. Single pixel fitted fluorescence lifetime images were then generated using the TRI2 software12,13 with thresholding from 15-100% intensity, 7 pixel circular binning and monoexponential fitting algorithm (Marquardt). The color indicates the average apparent fluorescent lifetime. (B) FLIM images where the fluorescence lifetime images (shown in panel A) were convoluted with the fluorescence intensities of the mCitrine donor fluorophore (shown in Figure 2). The convolution was performed with FIJI ImageJ using a custom-made macro (see Table of Materials). (C) Fluorescence lifetime and FLIM images of the HeLa cell expressing both syntaxin 4-mCitrine and VAMP3-mCherry from panels A-B, but now with fitting with bi-exponential decay curves (with lifetimes fixed to the control conditions; see step 4.4 in protocol). The pixel colors indicate the estimated fractions F of syntaxin 4 in complex with VAMP3 (Equation 3). (D) Same as panel B, but for the bi-exponential fit. The convolution was performed with FIJI ImageJ using a custom-made macro (see Table of Materials). <a href="https://cloudflare.jove.com/files/ftp_upload/56745/56745fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56745/56745fig4.jpg" /> 
Figure 4: Whole cell FLIM analysis. (A) The instrument response function (IRF) of the setup. The IRF was measured using the back scattering on the glass-water interface (using a clean glass microscopy dish containing water). (B-E) Whole cell lifetime histograms for the cells shown in Figure 2 and Figure 3. All photons present in the images were pooled. The curves were deconvoluted with the IRF and fitted with mono-exponential decay functions (Equation 1). For these cells, fluorescence lifetimes were obtained of 2.82 ns (cells expressing only syntaxin 4-mCitrine; donor only; panel B), 2.09 ns (cells co-expressing syntaxin 4-mCitrine with VAMP3-mCherry; panel C), and 2.08 ns (cells expressing the syntaxin 3-mCitrine-mCherry tandem construct; maximal expectable FRET control; panel D). The inlays show the same graphs, but now with logarithmic scaling of the y-axis. Panel E shows an overlay of the decay curves of panels B-D. (F) Example of a fluorescent lifetime histogram recorded too close to the surface of the microscope cover slip. This results in a large reflection peak (depicted by the yellow shaded area). (G) Same as panel C, but now fitting with a bi-exponential decay curve with lifetimes fixed to the control conditions (see step 4.4 in protocol). The amplitudes of the fast (A1) and slow (A2) components were 14.42 and 0.01, respectively, resulting in an estimated fraction of SNAREs in complex F of 0.99 (Equation 3). <a href="https://cloudflare.jove.com/files/ftp_upload/56745/56745fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary File 1. Function file FLIM_convoluted_IRF <a href="http://cloudflare.jove.com/files/ftp_upload/56745/FLIM_convoluted_IRF.FDF">Please click here to download this file.</a>
Supplementary File 2. Function file FLIM_convoluted_IRF_biexp <a href="http://cloudflare.jove.com/files/ftp_upload/56745/FLIM_convoluted_IRF_biexp.FDF">Please click here to download this file.</a>

Watch this Scientific Journal Video about Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy at JoVE.com

Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

This protocol describes a new method allowing for the quantitative visualization of complex formation of SNARE proteins, based on F&#246;rster resonance energy transfer, and fluorescence lifetime imaging microscopy.

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze ...

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Cell Membrane Structure and Functions

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9.4K Views.  Radboud University Medical Center. This protocol describes a new method allowing for the quantitative visualization of complex formation of SNARE proteins, based on Förster resonance energy transfer, and fluorescence lifetime imaging microscopy.

Video: Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

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

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular level. To date the shear complexity of biological systems has caused precise single-molecule experimentation to be far too demanding, instead focusing on studies of single systems using relatively crude bulk ensemble-average measurements. However, many important processes occur in the living cell at the level of just one or a few molecules; ensemble measurements generally mask the stochastic and heterogeneous nature of these events. Here, using advanced optical microscopy and analytical image analysis tools we demonstrate how to monitor proteins within a single living bacterial cell to a precision of single molecules and how we can observe dynamics within molecular complexes in functioning biological machines. The techniques are directly relevant physiologically. They are minimally-perturbative and non-invasive to the biological sample under study and are fully attuned for investigations in living material, features not readily available to other single-molecule approaches of biophysics. In addition, the biological specimens studied all produce fluorescently-tagged protein at levels which are almost identical to the unmodified cell strains ("genomic encoding"), as opposed to the more common but less ideal approach for generating significantly more protein than would occur naturally ('plasmid expression'). Thus, the actual biological samples which will be investigated are significantly closer to the natural organisms, and therefore the observations more relevant to real physiological processes.

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Here we demonstrate the protocols for performing single-molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified.

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

Biology

Single-Molecule Imaging of Nuclear Transport

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the last decade. The investigation of molecular interactions with high temporal and spatial resolutions deep within cells has remained challenging due to the inherently weak signals arising from individual molecules. Recent works by Yang et al. demonstrated that narrow-field epifluorescence microscopy allows visualization of nucleocytoplasmic transport at the single molecule level. By the single molecule approach, important kinetics, such as nuclear transport time and efficiency, for signal-dependent and independent cargo molecules have been obtained. Here we described a protocol for the methodological approach with an improved spatiotemporal resolution of 0.4 ms and 12 nm. The improved resolution enabled us to capture transient active transport and passive diffusion events through the nuclear pore complexes (NPC) in semi-intact cells. We expect this method to be used in elucidating other binding and trafficking events within cells.

Single molecule microscopy approch provided novel insights into nuclear transport.

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the ...

TIRFM and pH-sensitive GFP-probes to Evaluate Neurotransmitter Vesicle Dynamics in SH-SY5Y Neuroblastoma Cells: Cell Imaging and Data Analysis

Synaptic vesicles release neurotransmitters at chemical synapses through a dynamic cycle of fusion and retrieval. Monitoring synaptic activity in real time and dissecting the different steps of exo-endocytosis at the single-vesicle level are crucial for understanding synaptic functions in health and disease.
Genetically-encoded pH-sensitive probes directly targeted to synaptic vesicles and Total Internal Reflection Fluorescence Microscopy (TIRFM) provide the spatio-temporal resolution necessary to follow vesicle dynamics. The evanescent field generated by total internal reflection can only excite fluorophores placed in a thin layer (&#60;150 nm) above the glass cover on which cells adhere, exactly where the processes of exo-endocytosis take place. The resulting high-contrast images are ideally suited for vesicles tracking and quantitative analysis of fusion events.
In this protocol, SH-SY5Y human neuroblastoma cells are proposed as a valuable model for studying neurotransmitter release at the single-vesicle level by TIRFM, because of their flat surface and the presence of dispersed vesicles. The methods for growing SH-SY5Y as adherent cells and for transfecting them with synapto-pHluorin are provided, as well as the technique&#160;to perform TIRFM and imaging. Finally, a strategy aiming to select, count, and analyze fusion events at whole-cell and single-vesicle levels is presented.
To validate the imaging procedure and data analysis approach, the dynamics of pHluorin-tagged vesicles are&#160;analyzed under resting and stimulated (depolarizing potassium concentrations) conditions. Membrane depolarization increases the frequency of fusion events and causes a parallel raise of the net fluorescence signal recorded in whole cell. Single-vesicle analysis reveals modifications of fusion-event behavior (increased peak height and width). These data suggest that potassium depolarization not only induces a massive neurotransmitter release but also modifies the mechanism of vesicle fusion and recycling.
With the appropriate fluorescent probe, this technique can be employed in different cellular systems to dissect the mechanisms of constitutive and stimulated secretion.

This paper provides a method for investigating neurotransmitter vesicle dynamics in neuroblastoma cells, using a synaptobrevin2-pHluorin construct and Total Internal Reflection Fluorescence Microscopy. The strategy developed for image processing and data analysis is also reported.

Synaptic vesicles release neurotransmitters at chemical synapses through a dynamic cycle of fusion and retrieval. Monitoring synaptic activity in real ...

Neuroscience

SNARE-mediated Fusion of Single Proteoliposomes with Tethered Supported Bilayers in a Microfluidic Flow Cell Monitored by Polarized TIRF Microscopy

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. During neurotransmitter or hormone release via exocytosis, the fusion pore can transiently open and close repeatedly, regulating cargo release kinetics. Pore dynamics also determine the mode of vesicle recycling; irreversible resealing results in transient, &#34;kiss-and-run&#34; fusion, whereas dilation leads to full fusion. To better understand what factors govern pore dynamics, we developed an assay to monitor membrane fusion using polarized total internal reflection fluorescence (TIRF) microscopy with single molecule sensitivity and ~15 msec time resolution in a biochemically well-defined in vitro system. Fusion of fluorescently labeled small unilamellar vesicles containing v-SNARE proteins (v-SUVs) with a planar bilayer bearing t-SNAREs, supported on a soft polymer cushion (t-SBL, t-supported bilayer), is monitored. The assay uses microfluidic flow channels that ensure minimal sample consumption while supplying a constant density of SUVs. Exploiting the rapid signal enhancement upon transfer of lipid labels from the SUV to the SBL during fusion, kinetics of lipid dye transfer is monitored. The sensitivity of TIRF microscopy allows tracking single fluorescent lipid labels, from which lipid diffusivity and SUV size can be deduced for every fusion event. Lipid dye release times can be much longer than expected for unimpeded passage through permanently open pores. Using a model that assumes retardation of lipid release is due to pore flickering, a pore &#34;openness&#34;, the fraction of time the pore remains open during fusion, can be estimated. A soluble marker can be encapsulated in the SUVs for simultaneous monitoring of lipid and soluble cargo release. Such measurements indicate some pores may reseal after losing a fraction of the soluble cargo.

Here, we present a protocol to detect single, SNARE-mediated fusion events between liposomes and supported bilayers in microfluidic channels using polarized TIRFM, with single molecule sensitivity and ~15 msec time resolution. Lipid and soluble cargo release can be detected simultaneously. Liposome size, lipid diffusivity, and fusion pore properties are measured.

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. ...

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 ...

Utilizing pHluorin-tagged Receptors to Monitor Subcellular Localization and Trafficking

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular organelles and those localized on the plasma membrane. Traditional fluorescence-based measurements lack the capability to distinguish membrane proteins residing in different organelles. Cutting edge methodologies transcend traditional methods by coupling pH-sensitive fluorophores with total internal reflection fluorescence microscopy (TIRFM). TIRF illumination excites the sample up to approximately 150 nm from the glass-sample interface, thus decreasing background, increasing the signal to noise ratio, and enhancing resolution. The excitation volume in TIRFM encompasses the plasma membrane and nearby organelles such as the peripheral ER. Superecliptic pHluorin (SEP) is a pH sensitive version of GFP. Genetically encoding SEP into the extracellular domain of a membrane protein of interest positions the fluorophore on the luminal side of the ER and in the extracellular region of the cell. SEP is fluorescent when the pH is greater than 6, but remains in an off state at lower pH values. Therefore, receptors tagged with SEP fluoresce when residing in the endoplasmic reticulum (ER) or upon insertion in the plasma membrane (PM) but not when confined to a trafficking vesicle or other organelles such as the Golgi. The extracellular pH can be adjusted to dictate the fluorescence of receptors on the plasma membrane. The difference in fluorescence between TIRF images at neutral and acidic extracellular pH for the same cell corresponds to a relative number of receptors on the plasma membrane. This allows a simultaneous measurement of intracellular and plasma membrane resident receptors. Single vesicle insertion events can also be measured when the extracellular pH is neutral, corresponding to a low pH trafficking vesicle fusing with the plasma membrane and transitioning into a fluorescent state. This versatile technique can be exploited to study localization, expression, and trafficking of membrane proteins.

Labeling the extracellular domain of a membrane protein with a pH sensitive fluorophore, superecliptic pHluorin (SEP), allows subcellular localization, expression, and trafficking to be determined. Imaging SEP-labeled proteins with total internal reflection fluorescence microscopy (TIRFM) enables the quantification of protein levels in the peripheral ER and plasma membrane.

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular ...

Probing Structural and Dynamic Properties of Trafficking Subcellular Nanostructures by Spatiotemporal Fluctuation Spectroscopy

Imaging-derived mean square displacement (iMSD) is used to address the structural and dynamic properties of subcellular nanostructures, such as vesicles involved in the endo/exocytotic trafficking of solutes and biomolecules. iMSD relies on standard time-lapse imaging, is compatible with any optical setup, and does not need to dwell on single objects to extract trajectories. From each iMSD trace, a unique triplet of average structural and dynamic parameters (i.e., size, local diffusivity, anomalous coefficient) is calculated and combined to build the "iMSD signature" of the nanostructure under study. 
The potency of this approach is proved here with the exemplary case of macropinosomes. These vesicles evolve in time, changing their average size, number, and dynamic properties passing from early to late stages of intracellular trafficking. As a control, insulin secretory granules (ISGs) are used as a reference for subcellular structures that live in a stationary state in which the average structural and dynamic properties of the whole population of objects are invariant in time. The iMSD analysis highlights these peculiar features quantitatively and paves the way to similar applications at the sub-cellular level, both in the physiological and pathological states.

Imaging-derived mean square displacement (iMSD) analysis is applied to macropinosomes to highlight their intrinsic time-evolving nature in terms of structural and dynamic properties. Macropinosomes are then compared to insulin secretory granules (ISGs) as a reference for subcellular structures with time-invariant average structural/dynamic properties.

Imaging-derived mean square displacement (iMSD) is used to address the structural and dynamic properties of subcellular nanostructures, such as vesicles ...

Live Fluorescence, Inverse Imaging of Cell Ruffling, and Macropinocytosis

Macropinocytosis is a highly conserved but still incompletely understood process that is essential for the uptake and ingestion of fluid, fluid-phase nutrients and other material in cells. The dramatic extension of cell surface ruffles, their closure to form macropinosomes, and the maturation of internalized macropinosomes are key events in this pathway that can be difficult to capture using conventional confocal imaging based on tracking a bolus of fluorescent cargo. Fluorescent dextrans are commonly used experimentally as fluid phase markers for macropinosomes and for other endocytic pathways. A method the lab has adopted to optimize the imaging of dextran uptake involves using live imaging of cells bathed in high concentrations of fluorescent dextran in the medium, with the unlabeled cells appearing in relief (as black). The cell ruffles are highlighted to visualize ruffle closure, and internalized macropinosomes appear as fluorescent vacuoles in the cell interior. This method is optimal for visualizing macropinosome features and allows for easy segmentation and quantification. This paper describes dual-labeling of pathways with different sized dextrans and the co-expression of lipid probes and fluorescent membrane proteins to demark macropinosomes and other endosomes. The detection of internalized dextran at an ultrastructural level using correlative light and electron microscopy (CLEM) is also demonstrated. These cell processes can be imaged using multiple live imaging modalities, including in 3D. Taken together, these approaches optimize macropinosome imaging for many different settings and experimental systems.

This protocol demonstrates dextran imaging in live cells using continuous uptake and inverse images to optimize visualization of ruffling, macropinosome maturation, and analysis of dextran and other cell labelings.

Macropinocytosis is a highly conserved but still incompletely understood process that is essential for the uptake and ingestion of fluid, fluid-phase ...

Enhancing Axonal Transport Analysis in Living C. elegans Using GFP-Tagged RAB-3

Optimizing Visualization of Axonal Transport of Endogenous Cargo by Fluorescence Microscopy in Living Caenorhabditis elegans

Axonal transport is a prerequisite to deliver axonal proteins from their site of synthesis in the neuronal cell body to their destination in the axon. Consequently, loss of axonal transport impairs neuronal growth and function. Studying axonal transport therefore improves our understanding of neuronal cell biology. With recent improvements in CRISPR Cas9 genome editing, endogenous labeling of axonal cargos has become accessible, enabling to move beyond ectopic expression-based visualization of transport. However, endogenous labeling often comes at the cost of low signal intensity and necessitates optimization strategies to obtain robust data. Here, we describe a protocol to optimize the visualization of axonal transport by discussing acquisition parameters and a bleaching approach to improve the signal of endogenous labeled cargo over diffuse cytoplasmic background. We apply our protocol to optimize the visualization of synaptic vesicle precursors (SVPs) labeled by green fluorescent protein (GFP)-tagged RAB-3 to highlight how fine-tuning acquisition parameters can improve the analysis of endogenously labeled axonal cargo in Caenorhabditis elegans (C. elegans).

Author Spotlight: Advanced Techniques for Visualizing Endogenous Axonal Transport Dynamics

The paper describes the optimization of fluorescence microscopy acquisition parameters to visualize the axonal transport of endogenous labeled cargos at single-neuron resolution in a living nematode.

Axonal transport is a prerequisite to deliver axonal proteins from their site of synthesis in the neuronal cell body to their destination in the axon. ...

Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

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