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 β-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 β-galactosidase. The activity of β-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.
18 Related JoVE Articles!
Imaging Plasma Membrane Deformations With pTIRFM
Institutions: Wayne State University.
To gain novel insights into the dynamics of exocytosis, our group focuses on the changes in lipid bilayer shape that must be precisely regulated during the fusion of vesicle and plasma membranes. These rapid and localized changes are achieved by dynamic interactions between lipids and specialized proteins that control membrane curvature. The absence of such interactions would not only have devastating consequences for vesicle fusion, but a host of other cellular functions that involve control of membrane shape. In recent years, the identity of a number of proteins with membrane-shaping properties has been determined. What remains missing is a roadmap of when, where, and how they act as fusion and content release progress.
Our understanding of the molecular events that enable membrane remodeling has historically been limited by a lack of analytical methods that are sensitive to membrane curvature or have the temporal resolution to track rapid changes. PTIRFM satisfies both of these criteria. We discuss how pTIRFM is implemented to visualize and interpret rapid, submicron changes in the orientation of chromaffin cell membranes during dense core vesicle (DCV) fusion. The chromaffin cells we use are isolated from bovine adrenal glands. The membrane is stained with a lipophilic carbocyanine dye,1,1'-dioctadecyl-3,3,3',3'-tetramethylindodicarbocyanine, 4-chlorobenzenesulfonate, or diD. DiD intercalates in the membrane plane with a "fixed" orientation and is therefore sensitive to the polarization of the evanescent field. The diD-stained cell membrane is sequentially excited with orthogonal polarizations of a 561 nm laser (p-pol, s-pol). A 488 nm laser is used to visualize vesicle constituents and time the moment of fusion. Exocytosis is triggered by locally perfusing cells with a depolarizing KCl solution. Analysis is performed offline using custom-written software to understand how diD emission intensity changes relate to fusion pore dilation.
Biochemistry, Issue 86, Chromaffin Cells, Lipid Bilayers, Microscopy, Fluorescence, Polarization, Exocytosis, membrane, TIRF, pTIRF, chromaffin, polarization, vesicle
Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins
Institutions: National Institutes of Health, University of North Carolina at Chapel Hill , Rutgers University .
Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo
imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo
cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.
Cellular Biology, Issue 79, Microscopy, Confocal Microscopy, Fluorescence, Multiphoton, Exocytosis, Cell Biology, animal biology, animal models, Intravital Microscopy, Salivary glands, Exocytosis, In Vivo Imaging
Assessing the Secretory Capacity of Pancreatic Acinar Cells
Institutions: Weizmann Institute of Science.
Pancreatic acinar cells produce and secrete digestive enzymes. These cells are organized as a cluster which forms and shares a joint lumen. This work demonstrates how the secretory capacity of these cells can be assessed by culture of isolated acini. The setup is advantageous since isolated acini, which retain many characteristics of the intact exocrine pancreas can be manipulated and monitored more readily than in the whole animal. Proper isolation of pancreatic acini is a key requirement so that the ex vivo
culture will represent the in vivo
nature of the acini. The protocol demonstrates how to isolate intact acini from the mouse pancreas. Subsequently, two complementary methods for evaluating pancreatic secretion are presented. The amylase secretion assay serves as a global measure, while direct imaging of pancreatic secretion allows the characterization of secretion at a sub-cellular resolution. Collectively, the techniques presented here enable a broad spectrum of experiments to study exocrine secretion.
Cellular Biology, Issue 90, Cell biology, Exocrine secretion, Pancreatic acinar cells, Primary culture, Exocytosis, Actin, Lifeact
Optimized Negative Staining: a High-throughput Protocol for Examining Small and Asymmetric Protein Structure by Electron Microscopy
Institutions: The Molecular Foundry.
Structural determination of proteins is rather challenging for proteins with molecular masses between 40 - 200 kDa. Considering that more than half of natural proteins have a molecular mass between 40 - 200 kDa1,2
, a robust and high-throughput method with a nanometer resolution capability is needed. Negative staining (NS) electron microscopy (EM) is an easy, rapid, and qualitative approach which has frequently been used in research laboratories to examine protein structure and protein-protein interactions. Unfortunately, conventional NS protocols often generate structural artifacts on proteins, especially with lipoproteins that usually form presenting rouleaux artifacts. By using images of lipoproteins from cryo-electron microscopy (cryo-EM) as a standard, the key parameters in NS specimen preparation conditions were recently screened and reported as the optimized NS protocol (OpNS), a modified conventional NS protocol 3
. Artifacts like rouleaux can be greatly limited by OpNS, additionally providing high contrast along with reasonably high‐resolution (near 1 nm) images of small and asymmetric proteins. These high-resolution and high contrast images are even favorable for an individual protein (a single object, no average) 3D reconstruction, such as a 160 kDa antibody, through the method of electron tomography4,5
. Moreover, OpNS can be a high‐throughput tool to examine hundreds of samples of small proteins. For example, the previously published mechanism of 53 kDa cholesteryl ester transfer protein (CETP) involved the screening and imaging of hundreds of samples 6
. Considering cryo-EM rarely successfully images proteins less than 200 kDa has yet to publish any study involving screening over one hundred sample conditions, it is fair to call OpNS a high-throughput method for studying small proteins. Hopefully the OpNS protocol presented here can be a useful tool to push the boundaries of EM and accelerate EM studies into small protein structure, dynamics and mechanisms.
Environmental Sciences, Issue 90, small and asymmetric protein structure, electron microscopy, optimized negative staining
Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures
Institutions: Max Plank Institute of Molecular Plant Physiology, University of Hohenheim.
Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling processes. Extracting sterol-enriched membrane microdomains from plant cells for proteomic analysis is a difficult task mainly due to multiple preparation steps and sources for contaminations from other cellular compartments. The plasma membrane constitutes only about 5-20% of all the membranes in a plant cell, and therefore isolation of highly purified plasma membrane fraction is challenging. A frequently used method involves aqueous two-phase partitioning in polyethylene glycol and dextran, which yields plasma membrane vesicles with a purity of 95% 1
. Sterol-rich membrane microdomains within the plasma membrane are insoluble upon treatment with cold nonionic detergents at alkaline pH. This detergent-resistant membrane fraction can be separated from the bulk plasma membrane by ultracentrifugation in a sucrose gradient 2
. Subsequently, proteins can be extracted from the low density band of the sucrose gradient by methanol/chloroform precipitation. Extracted protein will then be trypsin digested, desalted and finally analyzed by LC-MS/MS. Our extraction protocol for sterol-rich microdomains is optimized for the preparation of clean detergent-resistant membrane fractions from Arabidopsis thaliana
We use full metabolic labeling of Arabidopsis thaliana
suspension cell cultures with K15
as the only nitrogen source for quantitative comparative proteomic studies following biological treatment of interest 3
. By mixing equal ratios of labeled and unlabeled cell cultures for joint protein extraction the influence of preparation steps on final quantitative result is kept at a minimum. Also loss of material during extraction will affect both control and treatment samples in the same way, and therefore the ratio of light and heave peptide will remain constant. In the proposed method either labeled or unlabeled cell culture undergoes a biological treatment, while the other serves as control 4
Empty Value, Issue 79, Cellular Structures, Plants, Genetically Modified, Arabidopsis, Membrane Lipids, Intracellular Signaling Peptides and Proteins, Membrane Proteins, Isotope Labeling, Proteomics, plants, Arabidopsis thaliana, metabolic labeling, stable isotope labeling, suspension cell cultures, plasma membrane fractionation, two phase system, detergent resistant membranes (DRM), mass spectrometry, membrane microdomains, quantitative proteomics
Direct Imaging of ER Calcium with Targeted-Esterase Induced Dye Loading (TED)
Institutions: University of Wuerzburg, Max Planck Institute of Neurobiology, Martinsried, Ludwig-Maximilians University of Munich.
Visualization of calcium dynamics is important to understand the role of calcium in cell physiology. To examine calcium dynamics, synthetic fluorescent Ca2+
indictors have become popular. Here we demonstrate TED (= targeted-esterase induced dye loading), a method to improve the release of Ca2+
indicator dyes in the ER lumen of different cell types. To date, TED was used in cell lines, glial cells, and neurons in vitro
. TED bases on efficient, recombinant targeting of a high carboxylesterase activity to the ER lumen using vector-constructs that express Carboxylesterases (CES). The latest TED vectors contain a core element of CES2 fused to a red fluorescent protein, thus enabling simultaneous two-color imaging. The dynamics of free calcium in the ER are imaged in one color, while the corresponding ER structure appears in red. At the beginning of the procedure, cells are transduced with a lentivirus. Subsequently, the infected cells are seeded on coverslips to finally enable live cell imaging. Then, living cells are incubated with the acetoxymethyl ester (AM-ester) form of low-affinity Ca2+
indicators, for instance Fluo5N-AM, Mag-Fluo4-AM, or Mag-Fura2-AM. The esterase activity in the ER cleaves off hydrophobic side chains from the AM form of the Ca2+
indicator and a hydrophilic fluorescent dye/Ca2+
complex is formed and trapped in the ER lumen. After dye loading, the cells are analyzed at an inverted confocal laser scanning microscope. Cells are continuously perfused with Ringer-like solutions and the ER calcium dynamics are directly visualized by time-lapse imaging. Calcium release from the ER is identified by a decrease in fluorescence intensity in regions of interest, whereas the refilling of the ER calcium store produces an increase in fluorescence intensity. Finally, the change in fluorescent intensity over time is determined by calculation of ΔF/F0
Cellular Biology, Issue 75, Neurobiology, Neuroscience, Molecular Biology, Biochemistry, Biomedical Engineering, Bioengineering, Virology, Medicine, Anatomy, Physiology, Surgery, Endoplasmic Reticulum, ER, Calcium Signaling, calcium store, calcium imaging, calcium indicator, metabotropic signaling, Ca2+, neurons, cells, mouse, animal model, cell culture, targeted esterase induced dye loading, imaging
Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons
Institutions: The Jackson Laboratory.
A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an experimental approach with excellent spatial and temporal resolutions. Moreover, such an approach must also have the ability to distinguish receptors localized on the PM from those in intracellular compartments. Most importantly, detecting receptors in a single vesicle requires outstanding detection sensitivity, since each vesicle carries only a small number of receptors. Standard approaches for examining receptor trafficking include surface biotinylation followed by biochemical detection, which lacks both the necessary spatial and temporal resolutions; and fluorescence microscopy examination of immunolabeled surface receptors, which requires chemical fixation of cells and therefore lacks sufficient temporal resolution1-6
. To overcome these limitations, we and others have developed and employed a new strategy that enables visualization of the dynamic insertion of receptors into the PM with excellent spatial and temporal resolutions 7-17
. The approach includes tagging of a pH-sensitive GFP, the superecliptic pHluorin 18
, to the N-terminal extracellular domain of the receptors. Superecliptic pHluorin has the unique property of being fluorescent at neutral pH and non-fluorescent at acidic pH (pH < 6.0). Therefore, the tagged receptors are non-fluorescent when within the acidic lumen of intracellular trafficking vesicles or endosomal compartments, and they become readily visualized only when exposed to the extracellular neutral pH environment, on the outer surface of the PM. Our strategy consequently allows us to distinguish PM surface receptors from those within intracellular trafficking vesicles. To attain sufficient spatial and temporal resolutions, as well as the sensitivity required to study dynamic trafficking of receptors, we employed total internal reflection fluorescent microscopy (TIRFM), which enabled us to achieve the optimal spatial resolution of optical imaging (~170 nm), the temporal resolution of video-rate microscopy (30 frames/sec), and the sensitivity to detect fluorescence of a single GFP molecule. By imaging pHluorin-tagged receptors under TIRFM, we were able to directly visualize individual receptor insertion events into the PM in cultured neurons. This imaging approach can potentially be applied to any membrane protein with an extracellular domain that could be labeled with superecliptic pHluorin, and will allow dissection of the key detailed mechanisms governing insertion of different membrane proteins (receptors, ion channels, transporters, etc.) to the PM.
Neuroscience, Issue 69, Cellular Biology, Bioengineering, Medicine, primary cultured mouse neuron, superecliptic pHluorin, receptor, plasma membrane insertion, total internal reflection fluorescence microscopy, neurons, mice, pHlourin-tagged, plasma membrane
Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes
Institutions: European Neuroscience Institute Göttingen.
The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1
; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+
). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii
). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v
), process that can be followed by monitoring the fluorescence intensity decrease (destaining).
Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3
. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.
JoVE Neuroscience, Issue 36, Photoconversion, FM1-43, Electron Microscope, Fluorescence, Drosophila, NMJ
A Microplate Assay to Assess Chemical Effects on RBL-2H3 Mast Cell Degranulation: Effects of Triclosan without Use of an Organic Solvent
Institutions: University of Maine, Orono, University of Maine, Orono.
Mast cells play important roles in allergic disease and immune defense against parasites. Once activated (e.g.
by an allergen), they degranulate, a process that results in the exocytosis of allergic mediators. Modulation of mast cell degranulation by drugs and toxicants may have positive or adverse effects on human health. Mast cell function has been dissected in detail with the use of rat basophilic leukemia mast cells (RBL-2H3), a widely accepted model of human mucosal mast cells3-5
. Mast cell granule component and the allergic mediator β-hexosaminidase, which is released linearly in tandem with histamine from mast cells6
, can easily and reliably be measured through reaction with a fluorogenic substrate, yielding measurable fluorescence intensity in a microplate assay that is amenable to high-throughput studies1
. Originally published by Naal et al.1
, we have adapted this degranulation assay for the screening of drugs and toxicants and demonstrate its use here.
Triclosan is a broad-spectrum antibacterial agent that is present in many consumer products and has been found to be a therapeutic aid in human allergic skin disease7-11
, although the mechanism for this effect is unknown. Here we demonstrate an assay for the effect of triclosan on mast cell degranulation. We recently showed that triclosan strongly affects mast cell function2
. In an effort to avoid use of an organic solvent, triclosan is dissolved directly into aqueous buffer with heat and stirring, and resultant concentration is confirmed using UV-Vis spectrophotometry (using ε280
= 4,200 L/M/cm)12
. This protocol has the potential to be used with a variety of chemicals to determine their effects on mast cell degranulation, and more broadly, their allergic potential.
Immunology, Issue 81, mast cell, basophil, degranulation, RBL-2H3, triclosan, irgasan, antibacterial, β-hexosaminidase, allergy, Asthma, toxicants, ionophore, antigen, fluorescence, microplate, UV-Vis
Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy
Institutions: University of Maine.
Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.
Basic Protocol, Issue 82, Microscopy, Super-resolution imaging, Multicolor, single molecule, FPALM, Localization microscopy, fluorescent proteins
Examination of Synaptic Vesicle Recycling Using FM Dyes During Evoked, Spontaneous, and Miniature Synaptic Activities
Institutions: University of Iowa Carver College of Medicine, University of Bath.
Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using styryl FM dyes, which reveal membrane turnover. Conventional protocols for the use of FM dyes were designed for analyzing neurons following stimulated (evoked) synaptic activity. Recently, protocols have become available for analyzing the FM signals that accompany weaker synaptic activities, such as spontaneous or miniature synaptic events. Analysis of these small changes in FM signals requires that the imaging system is sufficiently sensitive to detect small changes in intensity, yet that artifactual changes of large amplitude are suppressed. Here we describe a protocol that can be applied to evoked, spontaneous, and miniature synaptic activities, and use cultured hippocampal neurons as an example. This protocol also incorporates a means of assessing the rate of photobleaching of FM dyes, as this is a significant source of artifacts when imaging small changes in intensity.
Neuroscience, Issue 85, Presynaptic Terminals, Synaptic Vesicles, Microscopy, Biological Assay, Nervous System, Endocytosis, exocytosis, fluorescence imaging, FM dye, neuron, photobleaching
Reconstitution of a Kv Channel into Lipid Membranes for Structural and Functional Studies
Institutions: University of Texas Southwestern Medical Center at Dallas.
To study the lipid-protein interaction in a reductionistic fashion, it is necessary to incorporate the membrane proteins into membranes of well-defined lipid composition. We are studying the lipid-dependent gating effects in a prototype voltage-gated potassium (Kv) channel, and have worked out detailed procedures to reconstitute the channels into different membrane systems. Our reconstitution procedures take consideration of both detergent-induced fusion of vesicles and the fusion of protein/detergent micelles with the lipid/detergent mixed micelles as well as the importance of reaching an equilibrium distribution of lipids among the protein/detergent/lipid and the detergent/lipid mixed micelles. Our data suggested that the insertion of the channels in the lipid vesicles is relatively random in orientations, and the reconstitution efficiency is so high that no detectable protein aggregates were seen in fractionation experiments. We have utilized the reconstituted channels to determine the conformational states of the channels in different lipids, record electrical activities of a small number of channels incorporated in planar lipid bilayers, screen for conformation-specific ligands from a phage-displayed peptide library, and support the growth of 2D crystals of the channels in membranes. The reconstitution procedures described here may be adapted for studying other membrane proteins in lipid bilayers, especially for the investigation of the lipid effects on the eukaryotic voltage-gated ion channels.
Molecular Biology, Issue 77, Biochemistry, Genetics, Cellular Biology, Structural Biology, Biophysics, Membrane Lipids, Phospholipids, Carrier Proteins, Membrane Proteins, Micelles, Molecular Motor Proteins, life sciences, biochemistry, Amino Acids, Peptides, and Proteins, lipid-protein interaction, channel reconstitution, lipid-dependent gating, voltage-gated ion channel, conformation-specific ligands, lipids
Live Imaging Assay for Assessing the Roles of Ca2+ and Sphingomyelinase in the Repair of Pore-forming Toxin Wounds
Institutions: University of Maryland .
Plasma membrane injury is a frequent event, and wounds have to be rapidly repaired to ensure cellular survival. Influx of Ca2+
is a key signaling event that triggers the repair of mechanical wounds on the plasma membrane within ~30 sec. Recent studies revealed that mammalian cells also reseal their plasma membrane after permeabilization with pore forming toxins in a Ca2+
-dependent process that involves exocytosis of the lysosomal enzyme acid sphingomyelinase followed by pore endocytosis. Here, we describe the methodology used to demonstrate that the resealing of cells permeabilized by the toxin streptolysin O is also rapid and dependent on Ca2+
influx. The assay design allows synchronization of the injury event and a precise kinetic measurement of the ability of cells to restore plasma membrane integrity by imaging and quantifying the extent by which the liphophilic dye FM1-43 reaches intracellular membranes. This live assay also allows a sensitive assessment of the ability of exogenously added soluble factors such as sphingomyelinase to inhibit FM1-43 influx, reflecting the ability of cells to repair their plasma membrane. This assay allowed us to show for the first time that sphingomyelinase acts downstream of Ca2+
-dependent exocytosis, since extracellular addition of the enzyme promotes resealing of cells permeabilized in the absence of Ca2+
Cellular Biology, Issue 78, Molecular Biology, Infection, Medicine, Immunology, Biomedical Engineering, Anatomy, Physiology, Biophysics, Genetics, Bacterial Toxins, Microscopy, Video, Endocytosis, Biology, Cell Biology, streptolysin O, plasma membrane repair, ceramide, endocytosis, Ca2+, wounds
Methods for Cell-attached Capacitance Measurements in Mouse Adrenal Chromaffin Cell
Institutions: University of Illinois at Chicago.
Neuronal transmission is an integral part of cellular communication within the brain. Depolarization of the presynaptic membrane leads to vesicle fusion known as exocytosis that mediates synaptic transmission. Subsequent retrieval of synaptic vesicles is necessary to generate new neurotransmitter-filled vesicles in a process identified as endocytosis. During exocytosis, fusing vesicle membranes will result in an increase in surface area and subsequent endocytosis results in a decrease in the surface area. Here, our lab demonstrates a basic introduction to cell-attached capacitance recordings of single endocytic events in the mouse adrenal chromaffin cell. This type of electrical recording is useful for high-resolution recordings of exocytosis and endocytosis at the single vesicle level. While this technique can detect both vesicle exocytosis and endocytosis, the focus of our lab is vesicle endocytosis. Moreover, this technique allows us to analyze the kinetics of single endocytic events. Here the methods for mouse adrenal gland tissue dissection, chromaffin cell culture, basic cell-attached techniques, and subsequent examples of individual traces measuring singular endocytic event are described.
Neuroscience, Issue 92, Cell-attached capacitance measurements, chromaffin cells, single vesicles, endocytosis, exocytosis, clathrin-mediated endocytosis (CME), patch clamp
Imaging Cell Membrane Injury and Subcellular Processes Involved in Repair
Institutions: Children's National Medical Center, George Washington University.
The ability of injured cells to heal is a fundamental cellular process, but cellular and molecular mechanisms involved in healing injured cells are poorly understood. Here assays are described to monitor the ability and kinetics of healing of cultured cells following localized injury. The first protocol describes an end point based approach to simultaneously assess cell membrane repair ability of hundreds of cells. The second protocol describes a real time imaging approach to monitor the kinetics of cell membrane repair in individual cells following localized injury with a pulsed laser. As healing injured cells involves trafficking of specific proteins and subcellular compartments to the site of injury, the third protocol describes the use of above end point based approach to assess one such trafficking event (lysosomal exocytosis) in hundreds of cells injured simultaneously and the last protocol describes the use of pulsed laser injury together with TIRF microscopy to monitor the dynamics of individual subcellular compartments in injured cells at high spatial and temporal resolution. While the protocols here describe the use of these approaches to study the link between cell membrane repair and lysosomal exocytosis in cultured muscle cells, they can be applied as such for any other adherent cultured cell and subcellular compartment of choice.
Biochemistry, Issue 85, cell injury, lysosome exocytosis, repair, calcium, imaging, total internal reflection fluorescence (TIRF) microscopy, laser ablation
Characterizing the Composition of Molecular Motors on Moving Axonal Cargo Using "Cargo Mapping" Analysis
Institutions: The Scripps Research Institute, University of California San Diego, University of California San Diego, University of California San Diego School of Medicine.
Understanding the mechanisms by which molecular motors coordinate their activities to transport vesicular cargoes within neurons requires the quantitative analysis of motor/cargo associations at the single vesicle level. The goal of this protocol is to use quantitative fluorescence microscopy to correlate (“map”) the position and directionality of movement of live cargo to the composition and relative amounts of motors associated with the same cargo. “Cargo mapping” consists of live imaging of fluorescently labeled cargoes moving in axons cultured on microfluidic devices, followed by chemical fixation during recording of live movement, and subsequent immunofluorescence (IF) staining of the exact same axonal regions with antibodies against motors. Colocalization between cargoes and their associated motors is assessed by assigning sub-pixel position coordinates to motor and cargo channels, by fitting Gaussian functions to the diffraction-limited point spread functions representing individual fluorescent point sources. Fixed cargo and motor images are subsequently superimposed to plots of cargo movement, to “map” them to their tracked trajectories. The strength of this protocol is the combination of live and IF data to record both the transport of vesicular cargoes in live cells and to determine the motors associated to these exact same vesicles. This technique overcomes previous challenges that use biochemical methods to determine the average motor composition of purified heterogeneous bulk vesicle populations, as these methods do not reveal compositions on single moving cargoes. Furthermore, this protocol can be adapted for the analysis of other transport and/or trafficking pathways in other cell types to correlate the movement of individual intracellular structures with their protein composition. Limitations of this protocol are the relatively low throughput due to low transfection efficiencies of cultured primary neurons and a limited field of view available for high-resolution imaging. Future applications could include methods to increase the number of neurons expressing fluorescently labeled cargoes.
Neuroscience, Issue 92, kinesin, dynein, single vesicle, axonal transport, microfluidic devices, primary hippocampal neurons, quantitative fluorescence microscopy
Measuring Exocytosis in Neurons Using FM Labeling
The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission. Here we used real-time imaging of vesicles labeled with FM dye to monitor the rate of presynaptic vesicle release. FM4-64 is a red fluorescent amphiphilic styryl dye that embeds into the membranes of synaptic vesicles as endocytosis is stimulated. Lipophilic interactions cause the dye to greatly increase in fluorescence, thus emitting a bright signal when associated with vesicles and a nominal one when in the extracellular fluid. After a wash step is used to help remove external dye within the plasma membrane, the remaining FM is concentrated within the vesicles and is then expelled when exocytosis is induced by another round of electrical stimulation. The rate of vesicles release is measured from the resulting decrease in fluorescence. Since FM dye can be applied external and transiently, it is a useful tool for determining rates of exocytosis in neuronal cultures, especially when comparing the rates between transfected synapses and neighboring control boutons.
Neuroscience, Issue 1, neuron, imaging, exocytosis
Methods for Patch Clamp Capacitance Recordings from the Calyx
Institutions: National Institute of Health.
We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal. Electrical recordings from the presynaptic terminal allow the measurement of action potentials, calcium channel currents, vesicle fusion (exocytosis) and subsequent membrane uptake (endocytosis). The fusion of vesicles containing neurotransmitter causes the vesicle membrane to be added to the cell membrane of the calyx. This increase in the amount of cell membrane is measured as an increase in capacitance. The subsequent reduction in capacitance indicates endocytosis, the process of membrane uptake or removal from the calyx membrane. Endocytosis, is necessary to maintain the structure of the calyx and it is also necessary to form vesicles that will be filled with neurotransmitter for future exocytosis events. Capacitance recordings at the calyx of Held have made it possible to directly and rapidly measure vesicular release and subsequent endocytosis in a mammalian CNS nerve terminal. In addition, the corresponding postsynaptic activity can be simultaneously measured by using paired recordings. Thus a complete picture of the presynaptic and postsynaptic electrical activity at a central nervous system synapse is achievable using this preparation. Here, the methods for slice preparation, morphological features for identification of calyces of Held, basic patch clamping techniques, and examples of capacitance recordings to measure exocytosis and endocytosis are presented.
Neuroscience, Issue 6, membrane fusion, exocytosis, endocytosis