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 &#39;syntaxin 4-mCitrine&#39;</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 &#39;VAMP3-mCherry&#39;</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 &#39;syntaxin 3-mCitrine-mCherry&#39;</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&#160;</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&#39;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 &#39;Software&#39;-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 &#39;Software&#39;-section of www.membranetrafficking.com</td>
		</tr>
		<tr>
			<td>B&#252;rker Haemocytometer</td>
			<td>VWR</td>
			<td>630-1541</td>
			<td></td>
		</tr>
		<tr>
			<td>HeLa cells</td>
			<td>ATCC</td>
			<td>ATCC&#160;CCL-2</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>B Braun Melsungen&#160; 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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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 ...

visualizing-intracellular-snare-trafficking-fluorescence-lifetime

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Cell Membrane Structure and Functions

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9.4K 조회수.  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.

비디오: Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

Visualisation and Quantification of Intracellular Interactions of Neisseria meningitidis and Human &alpha;-actinin by Confocal Imaging

The Opc protein of Neisseria meningitidis (Nm, meningococcus) is a surface-expressed integral outer membrane protein, which can act as an adhesin and an effective invasin for human epithelial and endothelial cells. We have identified endothelial surface-located integrins as major receptors for Opc, a process which requires Opc to first bind to integrin ligands such as vitronectin and via these to the cell-expressed receptors1. This process leads to bacterial invasion of endothelial cells2. More recently, we observed an interaction of Opc with a 100kDa protein found in whole cell lysates of human cells3. We initially observed this interaction when host cell proteins separated by electrophoresis and blotted on to nitrocellulose were overlaid with Opc-expressing Nm. The interaction was direct and did not involve intermediate molecules. By mass spectrometry, we established the identity of the protein as &alpha;-actinin. As no surface expressed &alpha;-actinin was found on any of the eight cell lines examined, and as Opc interactions with endothelial cells in the presence of serum lead to bacterial entry into the target cells, we examined the possibility of the two proteins interacting intracellularly. For this, cultured human brain microvascular endothelial cells (HBMECs) were infected with Opc-expressing Nm for extended periods and the locations of internalised bacteria and &alpha;-actinin were examined by confocal microscopy. We observed time-dependent increase in colocalisation of Nm with the cytoskeletal protein, which was considerable after an eight hour period of bacterial internalisation. In addition, the use of quantitative imaging software enabled us to obtain a relative measure of the colocalisation of Nm with &alpha;-actinin and other cytoskeletal proteins. Here we present a protocol for visualisation and quantification of the colocalisation of the bacterium with intracellular proteins after bacterial entry into human endothelial cells, although the procedure is also applicable to human epithelial cells.

Visualisation and Quantification of Intracellular Interactions of Neisseria meningitidis and Human &#945;-actinin by Confocal Imaging

Neisseria meningitidis (Nm), a gram negative human-specific respiratory pathogen, can bind to human &alpha;-actinin. Here we present a protocol for visualisation of colocalisation of the bacterium with intracellular &alpha;-actinin after bacterial entry into human brain microvascular endothelial cells (HBMECs).

의 세포 상호 작용 Visualisation 및 부량 Neisseria meningitidis</em공촛점 이미징에 의해 α - actinin>과 인간

The Opc protein of Neisseria meningitidis (Nm, meningococcus) is a surface-expressed integral outer membrane protein, which can act as an adhesin and an ...

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면역학 및 감염병학

Microwave Assisted Rapid Diagnosis of Plant Virus Diseases by Transmission Electron Microscopy

Investigations of ultrastructural changes induced by viruses are often necessary to clearly identify viral diseases in plants. With conventional sample preparation for transmission electron microscopy (TEM) such investigations can take several days1,2 and are therefore not suited for a rapid diagnosis of plant virus diseases. Microwave fixation can be used to drastically reduce sample preparation time for TEM investigations with similar ultrastructural results as observed after conventionally sample preparation3-5. Many different custom made microwave devices are currently available which can be used for the successful fixation and embedding of biological samples for TEM investigations5-8. In this study we demonstrate on Tobacco Mosaic Virus (TMV) infected Nicotiana tabacum plants that it is possible to diagnose ultrastructural alterations in leaves in about half a day by using microwave assisted sample preparation for TEM. We have chosen to perform this study with a commercially available microwave device as it performs sample preparation almost fully automatically5 in contrast to the other available devices where many steps still have to be performed manually6-8 and are therefore more time and labor consuming. As sample preparation is performed fully automatically negative staining of viral particles in the sap of the remaining TMV-infected leaves and the following examination of ultrastructure and size can be performed during fixation and embedding.

This study describes a method that allows the rapid and clear diagnosis of plant virus diseases in about half a day by using a combination of microwave assisted plant sample preparation for transmission electron microscopy and negative staining methods.

전자 레인지 전송 전자 현미경으로 식물 바이러스 질병의 신속한 진단을 도와

Investigations of ultrastructural changes induced by viruses are often necessary to clearly identify viral diseases in plants. With conventional sample ...

Single Cell Measurements of Vacuolar Rupture Caused by Intracellular Pathogens

Shigella flexneri are pathogenic bacteria that invade host cells entering into an endocytic vacuole. Subsequently, the rupture of this membrane-enclosed compartment allows bacteria to move within the cytosol, proliferate and further invade neighboring cells. Mycobacterium tuberculosis is phagocytosed by immune cells, and has recently been shown to rupture phagosomal membrane in macrophages. We developed a robust assay for tracking phagosomal membrane disruption after host cell entry of Shigella flexneri or Mycobacterium tuberculosis. The approach makes use of CCF4, a FRET reporter sensitive to &beta;-lactamase that equilibrates in the cytosol of host cells. Upon invasion of host cells by bacterial pathogens, the probe remains intact as long as the bacteria reside in membrane-enclosed compartments. After disruption of the vacuole, &beta;-lactamase activity on the surface of the intracellular pathogen cleaves CCF4 instantly leading to a loss of FRET signal and switching its emission spectrum. This robust ratiometric assay yields accurate information about the timing of vacuolar rupture induced by the invading bacteria, and it can be coupled to automated microscopy and image processing by specialized algorithms for the detection of the emission signals of the FRET donor and acceptor. Further, it allows investigating the dynamics of vacuolar disruption elicited by intracellular bacteria in real time in single cells. Finally, it is perfectly suited for high-throughput analysis with a spatio-temporal resolution exceeding previous methods. Here, we provide the experimental details of exemplary protocols for the CCF4 vacuolar rupture assay on HeLa cells and THP-1 macrophages for time-lapse experiments or end points experiments using Shigella flexneri as well as multiple mycobacterial strains such as Mycobacterium marinum, Mycobacterium bovis, and Mycobacterium tuberculosis.

We describe a method for tracking the endomembrane rupture elicited by the intracellular bacteria Shigella flexneri and Mycobacterium tuberculosis upon host cell invasion. Our assay makes use of CCF4, a host cytoplasmic FRET probe in live or fixed cells. This reporter is degraded by an enzyme activity present on the bacterial surface.

액포 파열의 단일 세포 측정은 세포 내 병원균에 의해 발생

Shigella flexneri are pathogenic bacteria that invade host cells entering into an endocytic vacuole. Subsequently, the rupture of this membrane-enclosed ...

Discovery of New Intracellular Pathogens by Amoebal Coculture and Amoebal Enrichment Approaches

Intracellular pathogens such as legionella, mycobacteria and Chlamydia-like organisms are difficult to isolate because they often grow poorly or not at all on selective media that are usually used to cultivate bacteria. For this reason, many of these pathogens were discovered only recently or following important outbreaks. These pathogens are often associated with amoebae, which serve as host-cell and allow the survival and growth of the bacteria. We intend here to provide a demonstration of two techniques that allow isolation and characterization of intracellular pathogens present in clinical or environmental samples: the amoebal coculture and the amoebal enrichment. Amoebal coculture allows recovery of intracellular bacteria by inoculating the investigated sample onto an amoebal lawn that can be infected and lysed by the intracellular bacteria present in the sample. Amoebal enrichment allows recovery of amoebae present in a clinical or environmental sample. This can lead to discovery of new amoebal species but also of new intracellular bacteria growing specifically in these amoebae. Together, these two techniques help to discover new intracellular bacteria able to grow in amoebae. Because of their ability to infect amoebae and resist phagocytosis, these intracellular bacteria might also escape phagocytosis by macrophages and thus, be pathogenic for higher eukaryotes.

Amoebal coculture is a cell culture system using adherent amoebae to selectively grow intracellular pathogens able to resist phagocytic cells such as amoebae and macrophages. It thus represents a key tool to discover new infectious agents. Amoebal enrichment allows discovery of new amoebal species and of their specific intracellular bacteria.

Amoebal 공 배양 및 Amoebal 심화 접근에 의한 새로운 세포 내 병원균의 발견

Intracellular pathogens such as legionella, mycobacteria and Chlamydia-like organisms are difficult to isolate because they often grow poorly or not at ...

Analysis of Yersinia enterocolitica Effector Translocation into Host Cells Using Beta-lactamase Effector Fusions

Many gram-negative bacteria including pathogenic Yersinia spp. employ type III secretion systems to translocate effector proteins into eukaryotic target cells. Inside the host cell the effector proteins manipulate cellular functions to the benefit of the bacteria. To better understand the control of type III secretion during host cell interaction, sensitive and accurate assays to measure translocation are required. We here describe the application of an assay based on the fusion of a Yersinia enterocolitica effector protein fragment (Yersinia outer protein; YopE) with TEM-1 beta-lactamase for quantitative analysis of translocation. The assay relies on cleavage of a cell permeant FRET dye (CCF4/AM) by translocated beta-lactamase fusion. After cleavage of the cephalosporin core of CCF4 by the beta-lactamase, FRET from coumarin to fluorescein is disrupted and excitation of the coumarin moiety leads to blue fluorescence emission. Different applications of this method have been described in the literature highlighting its versatility. The method allows for analysis of translocation in vitro and also in in vivo, e.g., in a mouse model. Detection of the fluorescence signals can be performed using plate readers, FACS analysis or fluorescence microscopy. In the setup described here, in vitro translocation of effector fusions into HeLa cells by different Yersinia mutants is monitored by laser scanning microscopy. Recording intracellular conversion of the FRET reporter by the beta-lactamase effector fusion in real-time provides robust quantitative results. We here show exemplary data, demonstrating increased translocation by a Y. enterocolitica YopE mutant compared to the wild type strain.

Analysis of Yersinia enterocolitica Effector Translocation into Host Cells Using Beta-lactamase Effector Fusions

Effector translocation into host cells via a type III secretion system is a common virulence strategy among gram-negative bacteria. A beta-lactamase effector fusion based assay for quantitative analysis of translocation was applied. In Yersinia infected cells, conversion of a FRET reporter by the beta-lactamase is monitored using laser scanning microscopy.

분석<em&gt; 예 르시 니아의 enterocolitica</em숙주 세포 내로&gt; 이펙터 전좌 베타 - 락타 마제 이펙터의 용해 사용

Many gram-negative bacteria including pathogenic Yersinia spp. employ type III secretion systems to translocate effector proteins into eukaryotic target ...

Measuring Phagosome pH by Ratiometric Fluorescence Microscopy

Phagocytosis is a fundamental process through which innate immune cells engulf bacteria, apoptotic cells or other foreign particles in order to kill or neutralize the ingested material, or to present it as antigens and initiate adaptive immune responses. The pH of phagosomes is a critical parameter regulating fission or fusion with endomembranes and activation of proteolytic enzymes, events that allow the phagocytic vacuole to mature into a degradative organelle. In addition, translocation of H+ is required for the production of high levels of reactive oxygen species (ROS), which are essential for efficient killing and signaling to other host tissues. Many intracellular pathogens subvert phagocytic killing by limiting phagosomal acidification, highlighting the importance of pH in phagosome biology. Here we describe a ratiometric method for measuring phagosomal pH in neutrophils using fluorescein isothiocyanate (FITC)-labeled zymosan as phagocytic targets, and live-cell imaging. The assay is based on the fluorescence properties of FITC, which is quenched by acidic pH when excited at 490 nm but not when excited at 440 nm, allowing quantification of a pH-dependent ratio, rather than absolute fluorescence, of a single dye. A detailed protocol for performing in situ dye calibration and conversion of ratio to real pH values is also provided. Single-dye ratiometric methods are generally considered superior to single wavelength or dual-dye pseudo-ratiometric protocols, as they are less sensitive to perturbations such as bleaching, focus changes, laser variations, and uneven labeling, which distort the measured signal. This method can be easily modified to measure pH in other phagocytic cell types, and zymosan can be replaced by any other amine-containing particle, from inert beads to living microorganisms. Finally, this method can be adapted to make use of other fluorescent probes sensitive to different pH ranges or other phagosomal activities, making it a generalized protocol for the functional imaging of phagosomes.

Phagosomal pH influences phagosome maturation, oxidant production, phagosomal killing as well as antigen presentation. Here we describe a ratiometric method for measuring time-course and endpoint pH changes in individual phagosomes in living phagocytes using fluorescence microscopy.

비례 형광 현미경으로 Phagosome 산도 측정

Phagocytosis is a fundamental process through which innate immune cells engulf bacteria, apoptotic cells or other foreign particles in order to kill or ...

Non-invasive In Vivo Fluorescence Optical Imaging of Inflammatory MMP Activity Using an Activatable Fluorescent Imaging Agent

This paper describes a non-invasive method for imaging matrix metalloproteinases (MMP)-activity by an activatable fluorescent probe, via in vivo fluorescence optical imaging (OI), in two different mouse models of inflammation: a rheumatoid arthritis (RA) and a contact hypersensitivity reaction (CHR) model. Light with a wavelength in the near infrared (NIR) window (650 - 950 nm) allows a deeper tissue penetration and minimal signal absorption compared to wavelengths below 650 nm. The major advantages using fluorescence OI is that it is cheap, fast and easy to implement in different animal models.
Activatable fluorescent probes are optically silent in their inactivated states, but become highly fluorescent when activated by a protease. Activated MMPs lead to tissue destruction and play an important role for disease progression in delayed-type hypersensitivity reactions (DTHRs) such as RA and CHR. Furthermore, MMPs are the key proteases for cartilage and bone degradation and are induced by macrophages, fibroblasts and chondrocytes in response to pro-inflammatory cytokines. Here we use a probe that is activated by the key MMPs like MMP-2, -3, -9 and -13 and describe an imaging protocol for near infrared fluorescence OI of MMP activity in RA and control mice 6 days after disease induction as well as in mice with acute (1x challenge) and chronic (5x challenge) CHR on the right ear compared to healthy ears.

Non-invasive In Vivo Fluorescence Optical Imaging of Inflammatory MMP Activity Using an Activatable Fluorescent Imaging Agent

This paper explains the application of fluorescent imaging using an activatable optical imaging probe to visualize the in vivo activity of key matrix metalloproteinases in two different experimental models of inflammation.

비 침습적<em&gt; In Vivo</em&gt; 활성화 가능한 형광 이미징 에이전트를 사용한 염증성 MMP 활성의 형광 광학 이미징

This paper describes a non-invasive method for imaging matrix metalloproteinases (MMP)-activity by an activatable fluorescent probe, via in vivo ...

Murine Lymphocyte Labeling by 64Cu-Antibody Receptor Targeting for In Vivo Cell Trafficking by PET/CT

This protocol illustrates the production of 64Cu and the chelator conjugation/radiolabeling of a monoclonal antibody (mAb) followed by murine lymphocyte cell culture and 64Cu-antibody receptor targeting of the cells. In vitro evaluation of the radiolabel and non-invasive in vivo cell tracking in an animal model of an airway delayed-type hypersensitivity reaction (DTHR) by PET/CT are described.
In detail, the conjugation of a mAb with the chelator 1,4,7,10-tetraazacyclododecane-1,4,7,10-tetraacetic acid (DOTA) is shown. Following the production of radioactive 64Cu, radiolabeling of the DOTA-conjugated mAb is described. Next, the expansion of chicken ovalbumin (cOVA)-specific CD4+ interferon (IFN)-&#947;-producing T helper cells (cOVA-TH1) and the subsequent radiolabeling of the cOVA-TH1 cells are depicted. Various in vitro techniques are presented to evaluate the effects of 64Cu-radiolabeling on the cells, such as the determination of cell viability by trypan blue exclusion, the staining for apoptosis with Annexin V for flow cytometry, and the assessment of functionality by IFN-&#947; enzyme-linked immunosorbent assay (ELISA). Furthermore, the determination of the radioactive uptake into the cells and the labeling stability are described in detail. This protocol further describes how to perform cell tracking studies in an animal model for an airway DTHR and, therefore, the induction of cOVA-induced acute airway DHTR in BALB/c mice is included. Finally, a robust PET/CT workflow including image acquisition, reconstruction, and analysis is presented.
The 64Cu-antibody receptor targeting approach with subsequent receptor internalization provides high specificity and stability, reduced cellular toxicity, and low efflux rates compared to common PET-tracers for cell labeling, e.g.64Cu-pyruvaldehyde bis(N4-methylthiosemicarbazone) (64Cu-PTSM). Finally, our approach enables non-invasive in vivo cell tracking by PET/CT with an optimal signal-to-background ratio for 48 h. This experimental approach can be transferred to different animal models and cell types with membrane-bound receptors that are internalized.

Murine Lymphocyte Labeling by 64Cu-Antibody Receptor Targeting for In Vivo  Cell Trafficking by PET/CT

Following the preparation of a 64Cu-modified monoclonal antibody binding to a transgenic murine T cell receptor, T cells are radiolabeled in vivo, analyzed for viability, functionality, labeling stability and apoptosis, and adoptively transferred into mice with an airway delayed-type hypersensitivity reaction for non-invasive imaging by positron emission tomography/computed tomography (PET/CT).

에 의해 쥐 림프구 라벨링<sup&gt; (64)</sup&gt; CU-항체 수용체는 대한 타겟팅<em&gt; 생체</em&gt; PET / CT에 의한 세포 트래 피킹

This protocol illustrates the production of 64Cu and the chelator conjugation/radiolabeling of a monoclonal antibody (mAb) followed by murine lymphocyte ...

Visualizing Macrophage Extracellular Traps Using Confocal Microscopy

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal microscopy methods to visualize macrophage extracellular traps (METs). These extracellular traps are defined by the presence of extracellular chromatin with co-expression of other components such as granule proteases, citrullinated histones, and peptidyl arginase deiminase (PAD). The expression of METs is generally measured after exposure to a stimulus and compared to un-stimulated samples. Samples are also included for background and isotype control. Cells are analyzed using well-defined image analysis software. Confocal microscopy may be used to define the presence of METs both in vitro and in vivo in lung tissue.

Macrophage extracellular traps are a newly described entity. This article will concentrate on confocal microscopy methods and how they are visualized in vitro and in vivo from lung samples.

Confocal 현미경 검사 법을 사용 하 여 대 식 세포 세포 외 트랩을 시각화

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal ...

Integrate Imaging Flow Cytometry and Transcriptomic Profiling to Evaluate Altered Endocytic CD1d Trafficking

Populational analyses of the morphological and functional alteration of endocytic proteins are challenging due to the demand of image capture at a single cell level and statistical image analysis at a populational level. To overcome this difficulty, we used imaging flow cytometry and transcriptomic profiling (RNA-seq) to determine altered subcellular localization of the cluster of differentiation 1d protein (CD1d) associated with impaired endocytic gene expression in human dendritic cells (DCs), which were exposed to the common lipophilic air pollutant benzo[a]pyrene. The colocalization of CD1d and endocytic marker Lamp1 proteins from thousands of cell images captured with imaging flow cytometry was analyzed using IDEAS and ImageJ-Fiji programs. Numerous cellular images with co-stained CD1d and Lamp1 proteins were visualized after gating on CD1d+Lamp1+ DCs using IDEAS. The enhanced CD1d and Lamp1 colocalization upon BaP exposure was further demonstrated using thresholded scatterplots, tested with Mander&#39;s coefficients for co-localized intensity, and plotted based on the percentage of co-localized areas using ImageJ-Fiji. Our data provide an advantageous instrumental and bioinformatic approach to measure protein colocalization at both single and populational cellular levels, supporting an impaired functional outcome of transcriptomic alteration in pollutant-exposed human DCs.

Imaging flow cytometry provides an ideal approach to detect the morphological and functional alteration of cells at individual and populational levels. Disrupted endocytic function for lipid antigen presentation in pollutant-exposed human dendritic cells was demonstrated with a combined transcriptomic profiling of gene expression and morphological demonstration of protein trafficking.

통합 이미징 Cytometry Transcriptomic 변경된 Endocytic CD1d 매매 평가 하 프로 파일링

Populational analyses of the morphological and functional alteration of endocytic proteins are challenging due to the demand of image capture at a single ...