Name: measuring the induced membrane voltage with di-8-anepps
Uploaded: 2009-11-19T20:00:00
Duration: 5 min 52 s
Description: Watch this Scientific Journal Video about Measuring the Induced Membrane Voltage with Di-8-ANEPPS at JoVE.com

This work was supported by the Slovenian Research Agency with project Z2-9229 and program P2-0249. This video represents the supplementary material for the "Electroporation-based Technologies and Treatments" scientific workshop and postgraduate course, organized biannually by the Faculty of Electrical Engineering at the University of Ljubljana, Slovenia.

Part I: Preliminary steps
<ol>
<li>In this experiment Chinese hamster ovary cell line (CHO-K1) is used. Cells are plated in Lab-Tek II chambers (2 wells, 4 cm2 each) (Nalge Nunc, Germany) at ~0.7x105 cells/ml in HAM-F12 culture medium supplemented with 8% fetal calf serum, 0.15 mg/ml L-glutamine, 16 mg/ml gentamicin (all from Sigma-Aldrich, Steinheim, Germany), and 200 units/ml Crystacillin (Pliva, Zagreb, Croatia), and incubated in 5% CO2 at 37&deg;C. Alternatively, cells can also be plated on #1 glass cover slips (0.13 to 0.16 mm thick), coated with a cellular adhesive such as polylysine.</li>
<li>Incubate the cells in their culture medium. Incubation lasting 2 to 4 hours yields cells that are still roughly spherical, but firmly attached to the surface with a small part of their membrane. Alternatively, after 16 to 20 hours of incubation, cells are fully attached to the surface and have more complex shapes, but most of them are still not dividing.</li>
<li>Prepare a 10 mM stock solution of di-8-ANEPPS (Invitrogen, Eugene, Oregon, USA) by adding 843 &mu;l of DMSO (Sigma-Aldrich, Steinheim, Germany) to 5 mg of the dye in the original Invitrogen vial. The stock solution can be stored in a refrigerator at 4&deg;C for several months. Before starting the experiments, warm up the the solution until the crystals of DMSO dissolve.</li>
<li>Some cell lines may require the use of pluronic to ease the dye incorporation into the cell membrane. Pluronic can be purchased in 20% stock solution in DMSO (F-127, Invitrogen, Eugene, Oregon, USA), or a stock solution of the same concentration can be prepared by dissolving pluronic in DMSO. Stock solution of pluronic can be stored at room temperature.</li>
</ol>
Part II: Loading the cells with di-8-ANEPPS
<ol>
<li>Mix 3 &mu;l of 10 mM di-8-ANEPPS and 2.5 &mu;l of 20% pluronic in 1 ml of the Spinner modification (calcium-depleted version) of the Minimum Essential Medium SMEM (medium M8167 or M4767, Sigma-Aldrich, Steinheim, Germany) in a 1.5 ml Eppendorf tube. This yields a "loading solution" containing approximately 30 &mu;M di-8-ANEPPS and 0.05% of pluronic. For other cell types, their native culture media can be used instead of SMEM.</li>
<li>Replace the culture medium in the Lab-Tek chamber with the loading solution. Transfer the chamber to the refrigerator for 10 min at 4&deg;C. At this temperature, internalization of the dye through the plasma membrane is largely inhibited.</li>
<li>After staining, gently wash the excess dye two to three times with pure SMEM. </li>
<li>Leave 1.5 ml of SMEM in the chamber.</li>
</ol>
Part III: Experiment and image acquisition
<ol>
<li>The cells are observed using a fluorescence microscope (in our case Zeiss AxioVert 200, Zeiss, Germany) equipped with an oil immersion objective (x63, NA 1.4), a monochromator (Polychrome IV, Visitron, Germany) and a cooled CCD camera (VisiCam 1280, Visitron, Germany). The images are acquired with MetaFluor 7.1.1 and processed in MetaMorph 7.1.1 (both Molecular Devices, Downingtown, PA, USA), but other similar acquisition software can also be used. Set the excitation wavelength in the acquisition software to 490 nm and choose a suitable emission filter for ANEPPS, for example, a band-pass filter centered at 605 nm (605/55m, dichroic 565 DCXR, Chroma, Rockingham, VT, USA). If a monochromator is not available, an excitation filter centered at 490 nm can be used instead.</li>
<li>Place the chamber with cells onto the microscope stage, position the electrodes at the bottom of the chamber and connect them to the pulse generator. The medium should cover the electrodes.</li>
<li>To maintain cell viability and reduce the heating, the pulse should be of sufficiently low amplitude and short duration. In this experiment a square pulse with 35 V amplitude and 50 ms duration (low enough to avoid electroporation) is generated using a DC voltage supply and a custom-made microprocessor-controlled switcher device. Alternatively, a commercial pulse generator, such as 33210A (Agilent, Santa Clara, CA, USA) connected to an amplifier or a commercial stimulator, such as S88 (Grass, West Warwick, RI, USA), can be used. The pulse is delivered to two parallel Pt/Ir wire electrodes with 0.8 mm diameter and 4 mm distance between them, creating an electric field of approximately 88 V/cm between the electrodes and inducing &Delta;&Phi;. The exact field distribution between the electrodes used in this experiment is described elsewhere [1].</li>
<li>Find the cells of interest. Apply a single electric pulse or a sequence of pulses. For each pulse, acquire two fluorescence images: one immediately before the pulse (the control image), and one during the pulse (the pulse image). Due to the low response of di-8-ANEPPS, the changes in the fluorescence are difficult to discern by the naked eye and become apparent only after processing. The pulse delivery must be synchronized with the image acquisition, which is a feature of the acquisition software. To avoid photobleaching and possible heating, the illumination can be limited to the duration of the pulse.</li>
</ol>
Part IV: Image processing and analysis
<ol>
<li>Open the images in MetaMorph software. For each pulse, subtract the background in both the control and the pulse image. </li>
<li>Choose a cell and set the region of interest so that it corresponds to the membrane. Measure the fluorescence intensities along this region in the control and pulse image and transfer the values to a spreadsheet. </li>
<li>For each pulse, subtract the control data from the pulse data, and divide the result by control data to obtain the relative fluorescence changes. If a sequence of pulses is applied, the values of relative fluorescence changes determined for each pulse can be averaged to get a more reliable measurement.</li>
<li>Transform the relative fluorescence changes into &Delta;&Phi; using a calibration curve. A rough estimation of this curve can be obtained from the literature, but for higher accuracy, it has to be measured for each particular setup. For the cells and the setup used in this experiment a 6% decrease in fluorescence corresponds to a 100 mV increase in &Delta;&Phi; [2].</li>
<li>Finally, plot the voltage as a function of the relative arc length in a graphing software package such as Excel (Microsoft Corp., Redmond, WA, USA), Sigma Plot (Systat Software Inc., San Jose, CA, USA), or Origin (OriginLab Corp., Northampton, MA, USA). The curve can also be smoothed using a suitable filter, such as the moving average filter.</li>
</ol>

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Measurements of the induced membrane (transmembrane) voltage, &Delta;&Phi;, can be important in various experimental settings, such as studies of voltage-gated membrane channels, action potential propagation, cardiac cell stimulation, or cell membrane electroporation [3, 4, 5, 6, 7]. With simple cell shapes, &Delta;&Phi; can be calculated analytically. For example, for a spherical cell, &Delta;&Phi; is given by the Schwan s equation, which states that the voltage is proportional to the field strength and cell size and follows the cosine function along the membrane [8, 9]. For more complicated cell shapes, &Delta;&Phi; can deviate considerably from the cosine and must be determined either numerically, using a computer [2, 10, 11], or experimentally, using a potentiometric dye [12, 13, 14, 15].
One of the potentiometric dyes widely used for this purpose is di-8-ANEPPS (di-8-butyl-amino-naphthyl-ethylene-pyridinium-propyl-sulfonate), a fast dye with excitation and emission spectra dependent on the membrane voltage, which allows noninvasive observations of the variations of the &Delta;&Phi; on the cell membrane and to measure its value. In this video, we show an experimental approach for determination of &Delta;&Phi; by using di-8-ANEPPS.
The dye was developed by Professor Leslie Loew and colleagues [13, 14] at the University of Connecticut and belongs to the class of fast-response dyes. di-8-ANEPPS is nonfluorescent in water and becomes strongly fluorescent when it incorporates into the lipid bilayer of the cell membrane. A change in &Delta;&Phi; results in a change of the intramolecular charge distribution and corresponding changes in the spectral profile and intensity of the dye's fluorescence. The fluorescence intensity of di-8-ANEPPS varies proportionally to the change of &Delta;&Phi;; the response of the dye is linear for voltages ranging from &#8722;280 mV to +250 mV [4, 16]. Relatively small changes in fluorescence of the dye, uneven membrane staining, and dye internalization make di-8-ANEPPS less suitable for absolute measurements of membrane voltage, e.g. its resting component, although such efforts were also reported [17]. It is, however, suitable for measuring larger changes in membrane voltage, such as the onset of induced membrane voltage in nonexcitable cells exposed to external electric fields [12, 13], or action potentials in excitable cells [4, 5]. Although not applied here, di-8-ANEPPS also allows determination of &Delta;&Phi; by ratiometric measurements of fluorescence excitation [18] or emission [19], which increases the sensitivity of the response and reduces the abovementioned effects. As di-8-ANEPPS stains the membrane, it can also be used plainly as a membrane marker [2].
One of the drawbacks of the dye is that it is prone to photobleaching, so that prolonged exposures to the strong light should be avoided. Calibration of the dye is performed with either (i) potassium ionophore valinomycin and a set of different potassium concentrations in external medium [2,18], or (ii) patch-clamp in voltage clamp mode [17].
Finally, with the measurements of &Delta;&Phi; on spherical cells and cells of more complex shapes, the video demonstrates the influence of the cell shape on the amplitude and spatial distribution of &Delta;&Phi;. Thus for spherical cells &Delta;&Phi; is close to a cosine, in agreement with Schwan's equation, while for more complicated cell shapes the spatial distribution of &Delta;&Phi; is more intricate [20]...

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		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>di-8-ANEPPS</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>D-3167</td>
<td>potentiometric fluorescent dye</td>
</tr>
<tr>
<td>pluronic</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>P3000MP</td>
<td>potassium ionophore</td>
</tr>
<tr>
<td>DMSO</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>D2650</td>
<td>&nbsp;</td>
<tr>
<td>SMEM</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>M8167 or M4767</td>
<td>Spinner modification of the Minimum Essential Medium</td>
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<td>Ham-F12</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>N4888</td>
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<td>fetal calf serum</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
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<td>crystacillin</td>
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<td>Pliva</td>
<td>625110</td>
<td>antibiotic</td>
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<td>gentamicin</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>G1397</td>
<td>antibiotic</td>
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<td>Lab-Tek II</td>
<td>&nbsp;</td>
<td>Nalge Nunc</td>
<td>155379</td>
<td>chamber</td>
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<td>DC voltage supply</td>
<td>&nbsp;</td>
<td>Elektro-Automatik</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>microprocessor-controlled switcher</td>
<td>&nbsp;</td>
<td>Custom made</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>electrodes</td>
<td>&nbsp;</td>
<td>Custom made</td>
<td>&nbsp;</td>
<td>Pt/Ir</td>
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measuring the induced membrane voltage with di-8-anepps

Placement of a cell into an external electric field causes a local charge redistribution inside and outside of the cell in the vicinity of the cell membrane, resulting in a voltage across the membrane. This voltage, termed the induced membrane voltage (also induced transmembrane voltage, or induced transmembrane potential difference) and denoted by &Delta;&Phi;, exists only as long as the external field is present. If the resting voltage is present on the membrane, the induced voltage superimposes (adds) onto it. By using one of the potentiometric fluorescent dyes, such as di-8-ANEPPS, it is possible to observe the variations of &Delta;&Phi; on the cell membrane and to measure its value noninvasively. di-8-ANEPPS becomes strongly fluorescent when bound to the lipid bilayer of the cell membrane, with the change of the fluorescence intensity proportional to the change of &Delta;&Phi;. This video shows the protocol for measuring &Delta;&Phi; using di-8-ANEPPS and also demonstrates the influence of cell shape on the amplitude and spatial distribution of &Delta;&Phi;.

Watch this Scientific Journal Video about Measuring the Induced Membrane Voltage with Di-8-ANEPPS at JoVE.com

Measuring the Induced Membrane Voltage with Di-8-ANEPPS

External electric field induces a voltage on the membrane of a cell, termed the induced membrane voltage (&Delta;&Phi;). By using the potentiometric dye di-8-ANEPPS, it is possible to measure the &Delta;&Phi; noninvasively. This video shows the protocol for measuring &Delta;&Phi; using di-8-ANEPPS.

Placement of a cell into an external electric field causes a local charge redistribution inside and outside of the cell in the vicinity of the cell ...

measuring-the-induced-membrane-voltage-with-di-8-anepps

Research

JoVE Journal

Biology

Measuring Near Plasma Membrane and Global Intracellular Calcium Dynamics in Astrocytes

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, increasing evidence suggests that astrocytes may also actively participate in brain function through functional interactions with neurons.  However, many fundamental aspects of astrocyte biology remain controversial, unclear and/or experimentally unexplored. One important issue is the dynamics of intracellular calcium transients in astrocytes. This is relevant because calcium is well established as an important second messenger and because it has been proposed that astrocyte calcium elevations can trigger the release of transmitters from astrocytes. However, there has not been any detailed or satisfying description of near plasma membrane calcium signaling in astrocytes. Total internal reflection fluorescence (TIRF) microscopy is a powerful tool to analyze physiologically relevant signaling events within about 100 nm of the plasma membrane of live cells. Here, we use TIRF microscopy and describe how to monitor near plasma membrane and global intracellular calcium dynamics almost simultaneously. The further refinement and systematic application of this approach has the potential to inform about the precise details of astrocyte calcium signaling. A detailed understanding of astrocyte calcium dynamics may provide a basis to understand if, how, when and why astrocytes and neurons undergo calcium-dependent functional interactions.

We describe how to measure near membrane and global intracellular calcium dynamics in cultured astrocytes using total internal reflection and epifluorescence microscopy.

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, ...

Measuring Plasma Membrane Protein Endocytic Rates by Reversible Biotinylation

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins is responsible for a variety of key cellular events, including nutrient delivery, cellular excitability, and chemical signaling.  Many plasma membrane proteins are dynamically regulated by endocytic trafficking, which modulates protein function by altering protein surface expression.  The mechanisms that facilitate protein endocytosis are complex and are not fully understood for many membrane proteins.  In order to fully understand the mechanisms that control the endocytic trafficking of a given protein, it is critical that the protein s endocytic rate be precisely measured.  For many receptors, direct endocytic rate measurements are frequently achieved utilizing labeled receptor ligands.  However, for many classes of membrane proteins, such as transporters, pumps and ion channels, there is no convenient ligand that can be used to measure the endocytic rate.  In the present report, we describe a reversible biotinylation method that we employ to measure the dopamine transporter (DAT) endocytic rate.  This method provides a straightforward approach to measuring internalization rates, and can be easily employed for trafficking studies of most membrane proteins.

Regulated endocytosis governs the cell surface expression levels of the majority of membrane proteins. Here we utilize reducible, membrane impermeant biotinylation reagents to measure the endocytic rate of the dopamine transporter (DAT), a polytopic membrane protein. The method facilitates a straightforward approach to measuring the endocytic rate of most plasma membrane proteins.

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins ...

Measuring Caenorhabditis elegans Life Span in 96 Well Microtiter Plates

Lifespan is a biological process regulated by several genetic pathways. One strategy to investigate the biology of aging is to study animals that harbor mutations in components of age-regulatory pathways. If these mutations perturb the function of the age-regulatory pathway and therefore alter the lifespan of the entire organism, they provide important mechanistic insights1-3.
Another strategy to investigate the regulation of lifespan is to use small molecules to perturb age-regulatory pathways. To date, a number of molecules are known to extend lifespan in various model organisms and are used as tools to study the biology of aging4-16. The number of molecules identified thus far is small compared to the genetic "toolset" that is available to study the biology of aging.
Caenorhabditis elegans is one of the principle models used to study aging because of its excellent genetics and short lifespan of three weeks. More recently, C.elegans has emerged as a model organism for phenotype based drug screens5,7,16-20 because of its small size and its ability to grow in microtiter plates.
Here we present an assay to measure C.elegans lifespan in 96 well microtiter plates. The assay was developed and successfully used to screen large libraries for molecules that extend C.elegans lifespan7. The reliability of the assay was evaluated in multiple tests: first, by measuring the lifespan of wild type animals grown at different temperatures; second, by measuring the lifespan of mutants with altered lifespans; third, by measuring changes in lifespan in response to different concentrations of the antidepressant Mirtazepine. Mirtazepine has previously been shown to extend lifespan in C.elegans7. The results of these tests show that the assay is able to replicate previous findings from other assays and is quantitative. The microtiter format also makes this lifespan assay compatible with automated liquid handling systems and allows integration into automated platforms.

Measuring Caenorhabditis elegans Life Span in 96 Well Microtiter Plates

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Lifespan is a biological process regulated by several genetic pathways. One strategy to investigate the biology of aging is to study animals that harbor ...

Crystallization of Membrane Proteins in Lipidic Mesophases

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are implicated in a multitude of malfunctions and diseases. However, a structural and functional understanding of membrane proteins is strongly lagging behind that of their soluble partners, mainly, due to difficulties associated with their solubilization and generation of diffraction quality crystals. Crystallization in lipidic mesophases (also known as in meso or LCP crystallization) is a promising technique which was successfully applied to obtain high resolution structures of microbial rhodopsins, photosynthetic proteins, outer membrane beta barrels and G protein-coupled receptors. In meso crystallization takes advantage of a native-like membrane environment and typically produces crystals with lower solvent content and better ordering as compared to traditional crystallization from detergent solutions. The method is not difficult, but requires an understanding of lipid phase behavior and practice in handling viscous mesophase materials. Here we demonstrate a simple and efficient way of making LCP and reconstituting a membrane protein in the lipid bilayer of LCP using a syringe mixer, followed by dispensing nanoliter portions of LCP into an assay or crystallization plate, conducting pre-crystallization assays and harvesting crystals from the LCP matrix. These protocols provide a basic guide for approaching in meso crystallization trials; however, as with any crystallization experiment, extensive screening and optimization are required, and a successful outcome is not necessarily guaranteed.

The protocols describe the essential steps for obtaining diffraction quality crystals of a membrane protein starting from reconstitution of the protein in a lipidic cubic phase (LCP), finding initial conditions with LCP-FRAP pre-crystallization assays, setting up LCP crystallization trials and harvesting crystals.

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion channels in oocytes. Recordings from the cut-open setup are particularly useful for resolving low magnitude gating currents, rapid ionic current activation, and deactivation. The main benefits over the two-electrode voltage clamp (TEVC) technique include increased clamp speed, improved signal-to-noise ratio, and the ability to modulate the intracellular and extracellular milieu.
Here, we employ the human cardiac sodium channel (hNaV1.5), expressed in Xenopus oocytes, to demonstrate the cut-open setup and protocol as well as modifications that are required to add voltage clamp fluorometry capability.
The properties of fast activating ion channels, such as hNaV1.5, cannot be fully resolved near room temperature using TEVC, in which the entirety of the oocyte membrane is clamped, making voltage control difficult. However, in the cut-open technique, isolation of only a small portion of the cell membrane allows for the rapid clamping required to accurately record fast kinetics while preventing channel run-down associated with patch clamp techniques.
In conjunction with the COVG technique, ion channel kinetics and electrophysiological properties can be further assayed by using voltage clamp fluorometry, where protein motion is tracked via cysteine conjugation of extracellularly applied fluorophores, insertion of genetically encoded fluorescent proteins, or the incorporation of unnatural amino acids into the region of interest1. This additional data yields kinetic information about voltage-dependent conformational rearrangements of the protein via changes in the microenvironment surrounding the fluorescent molecule.

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open Vaseline gap approach is used to obtain low noise recordings of ionic and gating currents from voltage-dependent ion channels expressed in Xenopus oocytes with high resolution of fast channel kinetics. With minor modification, voltage clamp fluorometry can be coupled to the cut-open oocyte protocol.

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion ...

Imaging Plasma Membrane Deformations With pTIRFM

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&#39;-dioctadecyl-3,3,3&#39;,3&#39;-tetramethylindodicarbocyanine, 4-chlorobenzenesulfonate, or&#160;diD. DiD intercalates in the membrane plane with a &#34;fixed&#34; 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&#160;nm laser (p-pol, s-pol). A 488&#160;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.

Polarization-based Total Internal Reflection Fluorescence Microscopy (pTIRFM) enables real-time detection of cell membrane dynamics. This article describes the implementation of pTIRFM for the study of membrane remodeling during regulated exocytosis. The technique is generalizable to other processes in cell biology that directly or indirectly involve changes in membrane shape.

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 ...

Measuring Glutathione-induced Feeding Response in Hydra

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. The movement of prey organisms causes mechanosensory discharge of the stinging cells called nematocysts from hydra, which are inserted into the prey. The feeding response in hydra, which includes curling of the tentacles to bring the prey towards the mouth, opening of the mouth and consequent engulfing of the prey, is triggered by GSH present in the fluid released from the injured prey. To be able to identify the molecular mechanism of the feeding response in hydra which is unknown to date, it is necessary to establish an assay to measure the feeding response. Here, we describe a simple method for the quantitation of the feeding response in which the distance between the apical end of the tentacle and mouth of hydra is measured and the ratio of such distance before and after the addition of GSH is determined. The ratio, called the relative tentacle spread, was found to give a measure of the feeding response. This assay was validated using a starvation model in which starved hydra show an enhanced feeding response in comparison with daily fed hydra.

Here we describe a simple assay for the quantification of the feeding response in hydra induced by the reduced form of glutathione. This assay relies on measuring the distance between the apical end of the tentacle and mouth of hydra.

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. ...

Microperfusion Technique to Investigate Regulation of Microvessel Permeability in Rat Mesentery

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular mechanisms regulating vascular permeability in cultured endothelial cell monolayers and the functional exchange properties of whole microvascular beds. A method to cannulate and perfuse venular microvessels of rat mesentery and measure the hydraulic conductivity of the microvessel wall is described. The main equipment needed includes an intravital microscope with a large modified stage that supports micromanipulators to position three different microtools: (1) a beveled glass micropipette to cannulate and perfuse the microvessel; (2) a glass micro-occluder to transiently block perfusion and enable measurement of transvascular water flow movement at a measured hydrostatic pressure, and (3) a blunt glass rod to stabilize the mesenteric tissue at the site of cannulation. The modified Landis micro-occlusion technique uses red cells suspended in the artificial perfusate as markers of transvascular fluid movement, and also enables repeated measurements of these flows as experimental conditions are changed and hydrostatic and colloid osmotic pressure difference across the microvessels are carefully controlled. Measurements of hydraulic conductivity first using a control perfusate, then after re-cannulation of the same microvessel with the test perfusates enable paired comparisons of the microvessel response under these well-controlled conditions. Attempts to extend the method to microvessels in the mesentery of mice with genetic modifications expected to modify vascular permeability were severely limited because of the absence of long straight and unbranched microvessels in the mouse mesentery, but the recent availability of the rats with similar genetic modifications using the CRISPR/Cas9 technology is expected to open new areas of investigation where the methods described herein can be applied.

The modified Landis technique enables paired measurement of the hydraulic conductivity of individual microvessels in the mesentery of normal and genetically modified rats under control and test conditions using microperfusion techniques. It provides a convenient method to evaluate mechanisms that regulate microvessel permeability and transvascular exchange under physiological conditions.

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular ...

Ground State Depletion Super-resolution Imaging in Mammalian Cells

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to dramatic leaps in our understanding of how cells function. The development and availability of super-resolution microscopy has considerably extended the limits of optical resolution from ~250-20 nm. Biologists are no longer limited to describing molecular interactions in terms of colocalization within a diffraction limited area, rather it is now possible to visualize the dynamic interactions of individual molecules. Here, we outline a protocol for the visualization and quantification of cellular proteins by ground-state depletion microscopy&#160;for fixed cell imaging. We provide examples from two different membrane proteins, an element of the endoplasmic reticulum translocon, sec61&#946;, and a plasma membrane-localized voltage-gated L-type Ca2+ channel (CaV1.2). Discussed are the specific microscope parameters, fixation methods, photo-switching buffer formulation, and pitfalls and challenges of image processing.

This article outlines a protocol for the detection of one or more plasma and/or intracellular membrane proteins using Ground State Depletion (GSD) super-resolution microscopy in mammalian cells. Here, we discuss the benefits and considerations of using such approaches for the visualization and quantification of cellular proteins.

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to ...