This work was supported by the Department of Biotechnology (BT/PR32489/BRB/10/1786/2019 to SRGS); Science and Engineering Research Board (CRG/2019/000281 to SRGS); DBT-NBACD (BT/HRD-NBA-NWB/38/2019-20 to SRGS) and IISc-DBT partnership program (to SRGS). Infrastructure in the department was supported by DST-FIST, DBT, and UGC. AMB was supported by DBT-JRF (DBT/2015/IISc/NJ-02).

The protocol involves the seeding of melanocytes followed by transfection of the plasmids. Further steps include fixation, immunostaining, imaging, and analysis of the cells to measure the length and number of REs. The detailed description of the protocol is given below.
1. Seeding of mouse melanocytes on pre-treated coverslips
<ol>
	<li>Coat the glass coverslips in a Petri dish (i.e., 4 - 5 in a 35 mm dish) with basement membrane matrix medium (1:20 in complete RPMI medium: RPMI + 10% heat inactivated FBS + 1x Glutamine + 1x Antibiotic mix) and dry it in the tissue culture hood for 15 min. Wash the coverslips once with 1x PBS before use.</li>
	<li>Maintain the wild type mouse melanocytes (melan-Ink4a-Arf-1 from C57BL/6J, a/a, Ink4a-Arf-/- mice, described in20 and available at The Welcome Trust Functional Genomics Cell Bank) in a Petri dish (i.e., 35 or 60 mm dish) supplemented with complete RPMI medium.
	<ol>
		<li>Wash the cells twice with 1x PBS and add 0.5 or 1 mL of Trypsin-EDTA (0.25%) solution for trypsinizing cells. Incubate the cells at 37 &#176;C for 2 - 5 min for the detachment from the Petri dish.</li>
		<li>Add 1 - 2 mL of complete RPMI medium, suspend and then transfer the cells to a centrifuge tube.</li>
	</ol>
	</li>
	<li>Centrifuge the cell suspension at 4 &#176;C, 376 x g for 5 min and then resuspend the pellet in 1x PBS.</li>
	<li>Repeat the centrifugation step and resuspend the cells in 1 mL of complete RPMI medium.</li>
	<li>Seed the cells on the basement membrane medium-coated coverslips at 50-60% confluency (approximately 6 x 105 cells on a 35 mm cell culture dish containing 4 - 5 coverslips). Always add 200 nM of PMA (add 5 &#181;L of working stock 40 &#181;M phorbol 12-myristate 13-acetate) to the plated cell suspension in complete RPMI medium.</li>
	<li>After seeding, incubate the plate at 37 &#176;C for 12- 24 h.</li>
</ol>
2. Transfection of cells with the STX13 plasmids
<ol>
	<li>Use the following reagents: pEGFP-C1-STX13WT and pEGFP-C1-STX13&#916;129 (described in12). mCherry-Rab11, was a kind gift from Gra&#231;a Raposo, Institut Curie, Paris (described in reference19). Anti-TYRP1 antibody from ATCC (TA99).</li>
	<li>After 12 - 24 h of seeding, transfect the cells with plasmids using a lipid-based transfection reagent. For a 35 mm dish, take 5 &#181;L of the transfection reagent in 250 &#181;L of OPTI-MEM medium in a microcentrifuge tube and take approximately 200 ng of each plasmid in 250 &#181;L of OPTI-MEM.</li>
	<li>Incubate the tubes containing DNA and the transfection reagent for 5 min. Mix without repeated pipetting (total volume will be about 500 &#181;L). Incubate for 30 min at room temperature. Perform hand tapping of the tube every 10 min for 30 min at RT.</li>
	<li>During the incubation, wash the cells twice with 1x PBS, once with OPTI-MEM and then add 1 mL of OPTI-MEM to the cells.</li>
	<li>Post 30 min of incubation, add the transfection reagent-DNA mix to the cells in a dropwise manner by covering the dish.</li>
	<li>Incubate the cells at 37 &#176;C for 6 h. Aspirate the OPTI-MEM medium with transfection reagent and add complete RPMI medium supplemented with 200 nM of PMA.</li>
	<li>Incubate the cells at 37 &#176;C for 48 h.</li>
</ol>
3. Fixation of the cells
NOTE: The following procedure is performed outside the tissue culture hood.
<ol>
	<li>After 48 h of transfection, wash the cells twice with 1x PBS and then fix the cells with 3% formaldehyde (freshly prepared in 1x PBS) for 30 min.</li>
	<li>After fixation, wash the cells twice with 1x PBS and store the coverslips in 1x PBS until further use. Alternatively, cells can be mounted on glass slides (see below) or stored at 4 &#176;C.</li>
</ol>
4. Immunostaining of the cells
<ol>
	<li>Prepare a humid chamber: place paraffin film cut piece on moist filter paper in a Petri dish, covered with aluminum foil.</li>
	<li>Prepare 25 &#181;L of primary antibody solution (0.2% saponin in 1x PBS, 0.1% BSA in 1x PBS and 0.02% Sodium azide in 1x PBS). Add antibody at a dilution of 1:200. Add this solution as a drop on a paraffin film in the humid chamber.</li>
	<li>Carefully lift the coverslip with forceps, invert it on this drop of primary antibody staining solution and then cover the lid of the humid chamber. Incubate at room temperature for 30 min.</li>
	<li>Similarly, prepare the secondary antibody solution at a dilution of 1:500 and place it on paraffin film next to the coverslip in the humid chamber. For staining the nucleus, add DAPI (1:20,000 to 1:30,000) to the solution.</li>
	<li>Using forceps, carefully pick up the coverslip from the primary antibody solution and dip it thrice in 1x PBS (in a glass beaker).</li>
	<li>Tap the coverslip on tissue paper to remove the excess PBS on the coverslip. Place it on secondary antibody staining solution in the humid chamber and do not expose it to light due to the presence of fluorescently tagged antibodies in the solution.</li>
	<li>Incubate the coverslip again for 30 min at room temperature. Please always keep a note the side of the coverslip that has the cells throughout these steps.</li>
	<li>After the incubation, carefully pick up the coverslip from the secondary antibody solution and then dip it thrice in 1x PBS. Further, tap the coverslip on tissue paper to remove the excess PBS on the coverslip.</li>
	<li>Place 12 &#181;L of Fluoromount-G mounting reagent onto a glass slide and carefully place the stained coverslip (facing towards the glass) on the mounting reagent. Invert the glass slide on tissue paper and then gently press.</li>
</ol>
5. Fluorescence microscopy of the cells
<ol>
	<li>Image the stained cells under Bright-field (BF) and fluorescence (IF) filters using an inverted fluorescence microscope equipped with a CCD camera using 60x (oil) apochromatic objective or any other microscope with a similar configuration.</li>
</ol>
6. Quantification of overlap between the RE localized proteins and melanosomes:
NOTE: The following steps are followed for the quantification of Mander&#39;s overlap coefficient between the proteins using Fiji software (freely downloadable from the link: https://imagej.net/software/fiji/). Use the TIFF image with multiple channels.
<ol>
	<li>Open the raw image. Go to the Image option, select Color | Split channels, and use the two channels for analysis.</li>
	<li>Open the JACoP plugin in the Plugin option.</li>
	<li>Set the threshold for both the channels such that all the bright spots are selected and the background is eliminated.</li>
	<li>Go to Analysis option, select M1 and M2 coefficients for getting Mander&#39;s overlap coefficient.</li>
	<li>Press the Analyze option in the JACoP Plugin and see the result that displays Mander&#39;s overlap coefficient.</li>
</ol>
7. Quantification of recycling endosomes&#39; tubular number and length:
NOTE: The following steps are followed for the quantification of the number and length of the tubules using Fiji software.
<ol>
	<li>Open the raw image Go to Image option, select Color | Split channels and use the desired channel for analysis.</li>
	<li>Go to Image again, select Type | Convert to 8-bit image.</li>
	<li>Then go to Plugins, select Analyze | Tubeness. Set sigma value at 0.1075 . Press Ok.</li>
	<li>Go to Image again, select Type | Convert to 8 bit image.</li>
	<li>Go to Image | Adjust | Threshold (Use the same threshold values for all the images. Images should have approximately equal intensity).</li>
	<li>Go to Process | Binary | Convert to mask.</li>
	<li>Go to Process, select Binary and then select Skeletonize.</li>
	<li>Go to Analyze, select Skeleton and choose Analyze Skeleton. In result and output | select (a) Calculate the largest shortest path | (b) show detailed information | (c) display labeled skeleton. Press Ok. 
	NOTE: The result will open in the tabulated format. The column of Average branch length shows the length of all the different tubules in the cell that is selected (set scale for getting values in micrometres).</li>
	<li>For obtaining the number of tubules in the cell, go to Analyze, and select Analyze particle option. Press Ok. 
	NOTE: In the obtained results, the count column shows the number of tubules in that particular cell.</li>
	<li>Save the data and analyze.</li>
</ol>

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The authors declare that they have no conflict of interest.

Recycling endosomes are a cohort of endocytic organelles, and they mediate the recycling of cargo to the cell surface in all cell types21,22,23,24,25. In specialized cell types such as melanocytes, these organelles partly divert their trafficking route towards the melanosomes for their biogenesis3,16<s...

Biosynthesis of melanin pigments occurs in melanosomes, a melanocyte-specific lysosome-related organelle (LRO) that co-exists with conventional lysosomes. The endocytic system plays a key role in the biogenesis of melanosomes, required for skin color and photoprotection against ionizing radiation1,2,3. During this process, the melanin synthesizing enzymes are sorted on early/sorting endosomes and then transported to premature melanosomes through tubular or vesicular endosomes called recycling endosomes (REs)4,5,6,7,8,9,10. The targeting and fusion of these organelles regulate the maturation of fully functional pigmented melanosomes7,11,12,13,14. Defects in the formation of these organelles or cargo sorting to these organelles cause oculocutaneous albinism and other clinical phenotypes, observed in Hermansky-Pudlak syndrome15,16.
Here we describe a simple microscopy-based technique to study and analyze the REs. In this method, we have taken advantage of a transmembrane protein, Qa-SNARE Syntaxin (STX)13 that resides on recycling endosomes17 and cycles between sorting endosomes and melanosomes in melanocytes12,18. Further, deletion of N-terminal unstructured regulatory domain (namely SynN or STX13&#916;129) allows the SNARE to get stuck in melanosomes, which measures the forward trafficking pathway towards the melanosome12. We have used a known recycling endosomal marker Rab GTPase (Rab)11 in our studies14,19. Fluorescence imaging of the proteins GFP-STX13WT, GFP-STX13&#916;129, mCherry-Rab11, and TYRP1 in wild type melanocytes followed by quantification of their relative localization will provide the nature and dynamics of REs in addition to their targeting to melanosomes. Thus, this is a simple technique that can be used to visualize and measure the dynamics of REs in melanocytes.

<table><tbody><tr><td>anti-TYRP1 antibody (TA99)</td><td>ATCC</td><td>HB-8704</td><td></td></tr><tr><td>Fluoromount-G</td><td>Southern Biotech</td><td>0100-01</td><td></td></tr><tr><td>Lipofectamine 2000</td><td>ThermoFisher Scientific</td><td>11668-500</td><td></td></tr><tr><td>Matrigel matrix</td><td>BD Biosciences</td><td>356231</td><td></td></tr><tr><td>OPTI-MEM</td><td>ThermoFisher Scientific</td><td>022600-050</td><td></td></tr><tr><td>Phorbol 12-myristate 13-acetate</td><td>Sigma-Aldrich</td><td>P8139</td><td></td></tr><tr><td>RPMI Medium 1640</td><td>ThermoFisher Scientific</td><td>31800-022</td><td></td></tr></tbody></table>

the microscopy-based assay to study and analyze the recycling endosomes using snare trafficking

Recycling endosomes (REs) are tubular-vesicular organelles generated from early/sorting endosomes in all cell types. These organelles play a key role in the biogenesis of melanosomes, a lysosome-related organelle produced by melanocytes. REs deliver the melanocyte-specific cargo to premature melanosomes during their formation. Blockage in the generation of REs, observed in several mutants of Hermansky-Pudlak syndrome, results in hypopigmentation of skin, hair, and eye. Therefore, studying the dynamics (refer to number and length) of REs is useful to understand the function of these organelles in normal and disease conditions. In this study, we aim to measure the RE dynamics using a resident SNARE STX13.

Quantification of STX13&#916;129 mutant localization to the melanosomes 
Immunofluorescence microscopy of STX13 in mouse wild type melanocytes showed GFP-STX13WT localized as vesicular and tubular structures and GFP-STX13&#916;129 localized as ring-like structures in addition to the cell surface (Figure 1A). Further, intracellular ring-like GFP-STX13&#916;129 showed colocalization with the mel...

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The Microscopy-Based Assay to Study and Analyze the Recycling Endosomes using SNARE Trafficking

Recycling endosomes are part of the endosomal tubular network. Here we present a method to quantify the dynamics of recycling endosomes using GFP-STX13 as an organelle marker.

Recycling endosomes (REs) are tubular-vesicular organelles generated from early/sorting endosomes in all cell types. These organelles play a key role in ...

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3.5K Views.  Indian Institute of Science, Bangalore KA 560012, India. Recycling endosomes are part of the endosomal tubular network. Here we present a method to quantify the dynamics of recycling endosomes using GFP-STX13 as an organelle marker.

Video: The Microscopy-Based Assay to Study and Analyze the Recycling Endosomes using SNARE Trafficking

Analysis of Endocytic Uptake and Retrograde Transport to the Trans-Golgi Network Using Functionalized Nanobodies in Cultured Cells

Transport of proteins and membranes from the cell surface to the Golgi and beyond is essential for homeostasis, organelle identity and physiology. To study retrograde protein traffic, we have recently developed a versatile nanobody-based toolkit to analyze transport from the cell surface to the Golgi complex, either by fixed and live cell imaging, by electron microscopy, or biochemically. We engineered functionalized anti-green fluorescent protein (GFP) nanobodies &#8212; small, monomeric, high-affinity protein binders &#8212; that can be applied to cell lines expressing membrane proteins of interest with an extracellular GFP moiety. Derivatized nanobodies bound to the GFP reporters are specifically internalized and transported piggyback along the reporters&#39; sorting routes. Nanobodies were functionalized with fluorophores to follow retrograde transport by fluorescence microscopy and live imaging, with ascorbate peroxidase 2 (APEX2) to investigate the ultrastructural localization of reporter-nanobody complexes by electron microscopy, and with tyrosine sulfation (TS) motifs to assess kinetics of trans-Golgi network (TGN) arrival. In this methodological article, we outline the general procedure to bacterially express and purify functionalized nanobodies. We illustrate the powerful use of our tool using the mCherry- and TS-modified nanobodies to analyze endocytic uptake and TGN arrival of cargo proteins.

Retrograde transport of proteins from the cell surface to the Golgi is essential to maintain membrane homeostasis. Here, we describe a method to biochemically analyze cell surface-to-Golgi transport of recombinant proteins using functionalized nanobodies in HeLa cells.

Transport of proteins and membranes from the cell surface to the Golgi and beyond is essential for homeostasis, organelle identity and physiology. To ...

Biochemistry

Density Gradient Ultracentrifugation for Investigating Endocytic Recycling in Mammalian Cells

Endosomal trafficking is an essential cellular process that regulates a broad range of biological events. Proteins are internalized from the plasma membrane and then transported to the early endosomes. The internalized proteins could be transited to the lysosome for degradation or recycled back to the plasma membrane. A robust endocytic recycling pathway is required to balance the removal of membrane materials from endocytosis. Various proteins are reported to regulate the pathway, including ADP-ribosylation factor 6 (ARF6). Density gradient ultracentrifugation is a classical method for cell fractionation. After the centrifugation, organelles are sedimented at their isopycnic surface. The fractions are collected and used for other downstream applications. Described here is a protocol to obtain a recycling endosome-containing fraction from transfected mammalian cells using density gradient ultracentrifugation. The isolated fractions were subjected to standard Western blotting for analyzing their protein contents. By employing this method, we identified that the plasma membrane targeting of engulfment and cell motility 1 (ELMO1), a Ras-related C3 botulinum toxin substrate 1 (Rac1) guanine nucleotide exchange factor, is through ARF6-mediated endocytic recycling.

This paper aims to present a protocol for preparing recycling endosomes from mammalian cells using sucrose density gradient ultracentrifugation.

Endosomal trafficking is an essential cellular process that regulates a broad range of biological events. Proteins are internalized from the plasma ...

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.

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 ...

Immunology and Infection

Quantification of Endosome and Lysosome Motilities in Cultured Neurons Using Fluorescent Probes

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, survival, and glial communications. To date, numerous studies have reported that defects in these systems cause various neuronal diseases. Thus, understanding the mechanisms underlying vesicle dynamics may provide influential clues that could aid in the treatment of several neuronal disorders. Here, we describe a method for quantifying vesicle motilities, such as motility distance and rate of movement, using a software plug-in for the ImageJ platform. To obtain images for quantification, we labeled neuronal endosome-lysosome structures with EGFP-tagged vesicle marker proteins and observed the movement of vesicles using a time-lapse microscopy. This method is highly useful and simplify measuring vesicle motility in neurites, such as axons and dendrites, as well as in the soma of both neurons and glial cells. Furthermore, this method can be applied to other cell lines, such as fibroblasts and endothelial cells. This approach could provide a valuable advancement of our understanding of membrane trafficking.

The investigation of membrane trafficking is crucial for understanding neuronal functions. Here, we introduce a method for quantifying vesicle motility in neurons. This is a convenient method that can be adapted to the quantification of membrane trafficking in the nervous system.

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, ...

Neuroscience

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic transmission during repetitive nerve firing. Impaired endocytosis in pathological conditions leads to decreases in synaptic strength and brain functions. Here, we describe methods used to measure synaptic vesicle endocytosis at the mammalian hippocampal synapse in neuronal culture. We monitored synaptic vesicle protein endocytosis by fusing a synaptic vesicular membrane protein, including synaptophysin and VAMP2/synaptobrevin, at the vesicular lumenal side, with pHluorin, a pH-sensitive green fluorescent protein that increases its fluorescence intensity as the pH increases. During exocytosis, vesicular lumen pH increases, whereas during endocytosis vesicular lumen pH is re-acidified. Thus, an increase of pHluorin fluorescence intensity indicates fusion, whereas a decrease indicates endocytosis of the labelled synaptic vesicle protein. In addition to using the pHluorin imaging method to record endocytosis, we monitored vesicular membrane endocytosis by electron microscopy (EM) measurements of Horseradish peroxidase (HRP) uptake by vesicles. Finally, we monitored the formation of nerve terminal membrane pits at various times after high potassium-induced depolarization. The time course of HRP uptake and membrane pit formation indicates the time course of endocytosis.

Synaptic vesicle endocytosis is detected by light microscopy of pHluorin fused with synaptic vesicle protein and by electron microscopy of vesicle uptake.

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic ...

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling and cellular processes that control efferocytosis. This quantification can be difficult to perform as apoptotic cells are often efferocytosed piecemeal, thus necessitating methods which can accurately delineate between the efferocytosed portion of an apoptotic target versus residual unengulfed cellular fragments. The approach outlined herein utilizes dual-labeling approaches to accurately quantify the dynamics of efferocytosis and efferocytic capacity of efferocytes such as macrophages. The cytosol of the apoptotic cell is labeled with a cell-tracking dye to enable monitoring of all apoptotic cell-derived materials, while surface biotinylation of the apoptotic cell allows for differentiation between internalized and non-internalized apoptotic cell fractions. The efferocytic capacity of efferocytes is determined by taking fluorescent images of live or fixed cells and quantifying the amount of bound versus internalized targets, as differentiated by streptavidin staining. This approach offers several advantages over methods such as flow cytometry, namely the accurate delineation of non-efferocytosed versus efferocytosed apoptotic cell fractions, the ability to measure efferocytic dynamics by live-cell microscopy, and the capacity to perform studies of cellular signaling in cells expressing fluorescently-labeled transgenes. Combined, the methods outlined in this protocol serve as the basis for a flexible experimental approach that can be used to accurately quantify efferocytic activity and interrogate cellular signaling pathways active during efferocytosis.

Efferocytosis, the phagocytic removal of apoptotic cells, is required to maintain homeostasis and is facilitated by receptors and signaling pathways that allow for the recognition, engulfment, and internalization of apoptotic cells. Herein, we present a fluorescence microscopy protocol for the quantification of efferocytosis and the activity of efferocytic signaling pathways.

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling ...

An Optical Assay for Synaptic Vesicle Recycling in Cultured Neurons Overexpressing Presynaptic Proteins

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane proteins become exposed at the cell surface. One of these proteins is Synaptotagmin-1 (Syt1). An antibody directed against the luminal domain of Syt1, once added to the culture medium, is taken up during the exo-endocytotic cycle. This uptake is proportional to the amount of SV recycling and can be quantified through immunofluorescence. Here, we combine Syt1 antibody uptake with double transfection of cultured hippocampal neurons. This allows us to (1) localize presynaptic sites based on expression of recombinant presynaptic marker Synaptophysin, (2) determine their functionality using Syt1 uptake, and (3) characterize the targeting and effects of a protein of interest, GFP-Rogdi.

We describe an optical assay for synaptic vesicle (SV) recycling in cultured neurons. Combining this protocol with double transfection to express a presynaptic marker and protein of interest allows us to locate presynaptic sites, their synaptic vesicle recycling capacity, and determine the role of the protein of interest.

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane ...

Analysis of SNARE-mediated Membrane Fusion Using an Enzymatic Cell Fusion Assay

The interactions of SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) proteins on vesicles (v-SNAREs) and on target membranes (t-SNAREs) catalyze intracellular vesicle fusion1-4. Reconstitution assays are essential for dissecting the mechanism and regulation of SNARE-mediated membrane fusion5. In a cell fusion assay6,7, SNARE proteins are expressed ectopically at the cell surface. These "flipped" SNARE proteins drive cell-cell fusion, demonstrating that SNAREs are sufficient to fuse cellular membranes. Because the cell fusion assay is based on microscopic analysis, it is less efficient when used to analyze multiple v- and t-SNARE interactions quantitatively.
Here we describe a new assay8 that quantifies SNARE-mediated cell fusion events by activated expression of &beta;-galactosidase. Two components of the Tet-Off gene expression system9 are used as a readout system: the tetracycline-controlled transactivator (tTA) and a reporter plasmid that encodes the LacZ gene under control of the tetracycline-response element (TRE-LacZ). We transfect tTA into COS-7 cells that express flipped v-SNARE proteins at the cell surface (v-cells) and transfect TRE-LacZ into COS-7 cells that express flipped t-SNARE proteins at the cell surface (t-cells). SNARE-dependent fusion of the v- and t-cells results in the binding of tTA to TRE, the transcriptional activation of LacZ and expression of &beta;-galactosidase. The activity of &beta;-galactosidase is quantified using a colorimetric method by absorbance at 420 nm.
The vesicle-associated membrane proteins (VAMPs) are v-SNAREs that reside in various post-Golgi vesicular compartments10-15. By expressing VAMPs 1, 3, 4, 5, 7 and 8 at the same level, we compare their membrane fusion activities using the enzymatic cell fusion assay. Based on spectrometric measurement, this assay offers a quantitative approach for analyzing SNARE-mediated membrane fusion and for high-throughput studies.

We have developed a cell fusion assay that quantifies SNARE-mediated membrane fusion events by activated expression of &beta;-galactosidase.

The interactions of SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor)  proteins on vesicles (v-SNAREs) and on target membranes ...

The Cell-based L-Glutathione Protection Assays to Study Endocytosis and Recycling of Plasma Membrane Proteins

Membrane trafficking involves transport of proteins from the plasma membrane to the cell interior (i.e. endocytosis) followed by trafficking to lysosomes for degradation or to the plasma membrane for recycling. The cell based L-glutathione protection assays can be used to study endocytosis and recycling of protein receptors, channels, transporters, and adhesion molecules localized at the cell surface. The endocytic assay requires labeling of cell surface proteins with a cell membrane impermeable biotin containing a disulfide bond and the N-hydroxysuccinimide (NHS) ester at 4 &#186;C -&#160;a temperature at which membrane trafficking does not occur. Endocytosis of biotinylated plasma membrane proteins is induced by incubation at 37 &#186;C. Next, the temperature is decreased again to 4 &#186;C to stop endocytic trafficking and the disulfide bond in biotin covalently attached to proteins that have remained at the plasma membrane is reduced with L-glutathione. At this point, only proteins that were endocytosed remain protected from L-glutathione and thus remain biotinylated. After cell lysis, biotinylated proteins are isolated with streptavidin agarose, eluted from agarose, and the biotinylated protein of interest is detected by western blotting. During the recycling assay, after biotinylation cells are incubated at 37 &#176;C to load endocytic vesicles with biotinylated proteins and the disulfide bond in biotin covalently attached to proteins remaining at the plasma membrane is reduced with L-glutathione at 4 &#186;C as in the endocytic assay. Next, cells are incubated again at 37 &#176;C to allow biotinylated proteins from endocytic vesicles to recycle to the plasma membrane. Cells are then incubated at 4 &#186;C, and the disulfide bond in biotin attached to proteins that recycled to the plasma membranes is reduced with L-glutathione. The biotinylated proteins protected from L-glutathione are those that did not recycle to the plasma membrane.

Membrane trafficking involves transport of proteins from the plasma membrane to the cell interior (i.e. endocytosis) followed by trafficking to lysosomes for degradation or to the plasma membrane for recycling. Methods described in this article are designed to study endocytosis and recycling of plasma membrane proteins.

Membrane trafficking involves transport of proteins from the plasma membrane to the cell interior (i.e. endocytosis) followed by trafficking to lysosomes ...

An Introduction to Endocytosis and Exocytosis

Cells can take in substances from the extracellular environment by endocytosis and actively release molecules into it by exocytosis. Such processes involve lipid membrane-bound sacs called vesicles. Knowledge of the molecular architecture and mechanisms of both is key to understanding normal cell physiology, as well as the disease states that arise when they become defective. This video will first briefly review a few pivotal discoveries in the history of endo- and exocytosis research. Next, some key questions will be examined, followed by a discussion of the prominent methods used to investigate these problems, including cell labeling, fusion assays, and fluorescence imaging. Finally, it will explore current research being conducted by scientists in the field today.

Endocytosis and Exocytosis; Clathrin Coated Pits and SNAREs

Cells can take in substances from the extracellular environment by endocytosis and actively release molecules into it by exocytosis. Such processes ...