This work was supported by NIH grants R37 AI34867 and R01 GM064625 to N.W.A. We thank Dr. R. Tweten from the University of Oklahoma for the SLO expression plasmid.

1. Transcriptional Silencing of ASM
<ol>
 <li>Seed 1.5 x 105 HeLa cells in 2 ml of DMEM growth media (DMEM high glucose with 10% fetal bovine serum (FBS), 2 mM L-glutamine, 1% Penn-Strep) on 35 mm glass bottom dishes (MatTek) and incubate overnight at 37 &deg;C in a 5% CO2 incubator. </li>
 <li>On the following day, aspirate off the growth media and replace it with 2 ml DMEM reduced serum medium (DMEM with 4% FBS), at least 1 hr before adding the siRNA transfection mixture. </li>
 <li>Prepare 4 tubes for the siRNA oligo transfection mixture. In tubes A and C, add 250 &mu;l Optimem reduced serum and 4 &mu;l Lipofectamine RNAiMax, per 35 mm dish to be transfected. In tube B, add 250 &mu;l of Optimem reduced serum and 8 &mu;l (160 pmoles) of control medium GC content oligo (Control siRNA), per 35 mm dish to be transfected. In tube D, add 250 &mu;l of Optimem reduced serum and 8 &mu;l (160 pmoles) of SMPD1 oligo (siASM), per 35 mm dish to be transfected. Incubate the tubes at room temperature for 5 min. </li>
 <li>Combine tubes A and B to form the transfection complex for Control siRNA, and tubes C and D for the ASM siRNA . Incubate the reactions at room temperature for 20 min. </li>
 <li>Add the Control siRNA and ASM siRNA transfection reaction mixtures slowly, dropwise, into the respective 35-mm glass bottom dishes and incubate at 37 &deg;C/5% CO2. At 24 hr post-transfection, gently aspirate off media and replace with fresh DMEM growth media. For efficient ASM knockdown, the dishes should be further incubated at 37 &deg;C/5% CO2 for about 31 hr (total 55 hr) before live imaging. </li>
</ol>
2. Preparation of Reagents for Live Microscopy
<ol>
 <li>Prepare a solution of recombinant pore-forming toxin Streptolysin O (SLO) at a concentration of 100 ng/&mu;l in cold 1x PBS without Ca2+ and Mg2+. The SLO used in these assays is a constitutively active cysteine mutant that does not require reducing agents for activity, kindly provided by Dr. Rod Tweten, University of Oklahoma. The toxin was expressed in E. coli BL21 DE3 and purified under native conditions, as described previously16. </li>
 <li>Prepare the FM1-43 stock solution: Resuspend FM1-43 lyophilized powder in DMSO to create a 25 mM solution. </li>
 <li>Prepare Solution A: Dilute the FM1-43 stock solution in cold DMEM with Ca2+ to a final concentration of 4 &mu;M (1.5 ml are needed per MatTek dish) and store on ice until needed.</li>
 <li>Prepare Solution B: Dilute the FM1-43 stock solution in cold DMEM without Ca2+ and 10 mM EGTA (Ca2+-free DMEM) to a final concentration of 4 &mu;M (1.5 ml needed per MatTek dish) and store on ice until needed.</li>
 <li>Pre-warm (37 &deg;C) 1 ml aliquots of Solution A and Solution B in Eppendorf tubes. </li>
 <li>Keep on ice solutions of DMEM with Ca2+, and Ca2+-free DMEM. </li>
</ol>
3. Live Cell Imaging of FM1-43 Influx into SLO Treated and Untreated Cells
<ol>
 <li>Fill a medium size shallow metal container with crushed ice, invert it into a large glass container with ice, and add additional ice leaving only the bottom surface of the metal container exposed. Place a wet paper towel on the exposed metal bottom to allow for even thermal transfer. </li>
 <li>Take one Control siRNA-treated MatTek dish (Sample 1) and place it on the cold wet paper towel. Gently aspirate off all media and wash 3x with cold Ca2+-free DMEM. After the last wash, aspirate off all media in the dish. </li>
 <li>To image cell-associated FM1-43 in cells with no SLO wounding (Sample 1) in the presence of Ca2+, add 180 &mu;l of cold Solution B to the center glass coverslip of the 35-mm glass bottom dish and incubate for 5 min on ice (Table 1). </li>
 <li>Place MatTek dish on the stage of a confocal microscope heated to 37 &deg;C and equipped with an environmental chamber (which maintains temperature, humidity and CO2 levels). Find the field of cells that will be imaged looking through a 40X NA 1.3 objective (Nikon). If available, turn on PerfectFocus or a similar device to correct for thermal drift. </li>
 <li>Add 1 ml of prewarmed Solution A gently to the side of the 35-mm dish and replace cover of environmental chamber (Table 1). </li>
 <li>Acquire images for 4 min at 1 frame every 3 sec using appropriate imaging software (Volocity in the UltraView Vox Perkin Elmer spinning disk confocal microscopy system). Excitation of FM1-43 is accomplished with a 488 nm laser line using a dichroic filter with a 488 nm band pass, and emission discrimination is achieved through use of a 527(W55) emission filter. Laser power levels and camera sensitivity are set to achieve image exposures of 100 msec. </li>
 <li>Repeat steps 3.2 to 3.6 to detect FM1-43 in Control siRNA-treated cells with no SLO wounding (Sample 2) in the absence of Ca2+, except that at step 3.5 Solution B should be added (Table 1). </li>
 <li>To measure FM1-43 influx into Control siRNA-treated cells wounded with SLO either in the presence (Sample 3) or absence of Ca2+ (Sample 4), add 180 &mu;l of cold Solution B containing SLO to the center glass coverslip of the 35-mm glass bottom dish and incubate for 5 min on ice as in step 3.3. Repeat steps 3.2 through 3.6; adjust adding either Solution A or Solution B at step 3.5 (Table 1). </li>
 <li>To determine the role of ASM in plasma membrane repair, repeat steps 3.2-3.6 using ASM siRNA-treated cells for all conditions (Samples 5-8) (Table 1).</li>
 <li>To determine the role of extracellular recombinant sphingomyelinase (SM) on the kinetics of plasma membrane repair (reflected by the inhibition in FM1-43 influx) (Samples 9-10), SM is added (0-50 &mu;U) to either Solution A or Solution B in step 3.5 and then added to cells during live cell imaging in a total volume of 1 ml (Table 1). </li>
</ol>
4. Quantification of FM1-43 Influx Into Cells
<ol>
 <li>After movies have been acquired, measure the intracellular fluorescence intensity of FM1-43 using image analysis software (Volocity Suite, PerkinElmer). Draw a region of interest in the cytosolic region of each cell in the field and apply the software tools to determine the mean FM1-43 fluorescence intensity throughout all frames of the video. </li>
 <li>To adjust for variations in the initial FM1-43 staining, the data for each video is expressed as fold increase in fluorescence intensity over time, and plotted for each different condition. The fold increase in fluorescence of each individual timepoint per cell is calculated as FT/F0, where FT is the mean fluorescence at the specific timepoint, and F0 is the mean fluorescence at t = 0 (Figures 1-2). </li>
</ol>

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K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cholesterol-dependent+cytolysins,+a+family+of+versatile+pore-forming+toxins.">Cholesterol-dependent cytolysins, a family of versatile pore-forming toxins.</a> Infect. Immun. 73, 6199-6209 (2005).</li><li>Zha, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sphingomyelinase+treatment+induces+ATP-independent+endocytosis.">Sphingomyelinase treatment induces ATP-independent endocytosis.</a> J. Cell Biol. 140, 39-47 (1998).</li><li>Buckley, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crossing+three+membranes.+Channel+formation+by+aerolysin.">Crossing three membranes. Channel formation by aerolysin.</a> FEBS Lett. 307, 30-33 (1992).</li><li>Parker, M. 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The authors declare they have no competing financial interests.

Earlier studies on mechanical injury and plasma membrane repair relied on diverse mechanisms of injuring cells, ranging from dropping glass beads, micropipetting, shearing, scratching or scraping. The readout of all these assays was qualitative rather than quantitative, and did not give precise information on the kinetics of the repair mechanism. The ability of cells to reseal their plasma membrane after these forms of injury was measured by the presence or absence of tracers added to the extracellular fluid in the cyt...

It has been known for several decades that plasma membrane repair after mechanical injury is a Ca2+-dependent process1,2. Later studies showed that Ca2+ influx through the wound triggers a vigorous process of exocytosis of intracellular vesicles at the site of injury, which is required for resealing3,4. The rate of loss of a fluorescent dye loaded into the cytoplasm of cells was used to assess the speed of repair, and concluded that resealing was completed within &lt;30 sec after injury5. Two models were initially proposed to explain the requirement for exocytosis in plasma membrane repair: 1) the "patch" model, which suggested that Ca2+ influx through the lesion triggers initially homotypic fusion of intracellular vesicles, forming a large "patch" that would then be applied to the plasma membrane to reseal the wound6 and 2) the tension reduction model, which proposed that membrane added by Ca2+-dependent exocytosis in the vicinity of the wound would reduce plasma membrane tension, facilitating resealing of the bilayer7. The role of Ca2+-triggered exocytosis in plasma membrane repair was further reinforced by studies showing that lysosomes contain a Ca2+ sensor molecule, synaptotagmin VII, which facilitates their exocytosis and plasma membrane repair in injured cells8,9,10,11.
However, additional evidence has indicated that exocytosis alone was not sufficient to promote plasma membrane repair. In addition to mechanical tears on the plasma membrane, a frequent form of cell injury is permeabilization by pore-forming toxins produced by bacteria12,13 or immune cells14,15. Unlike mechanical tears, pore-forming proteins insert themselves on the plasma membrane forming a stable, protein-lined pore that cannot be resealed simply by applying a membrane "patch" or by reducing membrane tension. Intriguingly, studies revealed that mammalian cells have an efficient mechanism to repair their plasma membrane after permeabilization with pore-forming proteins, and this process also requires the presence of extracellular Ca2+ 12. This finding raised the question of whether transmembrane pore removal from the cell surface was also a rapid process, as observed with mechanical wounds5. Surprisingly, our recent studies revealed that the resealing of cells permeabilized with pore-forming toxins has very similar properties to the repair of mechanical wounds: the process requires extracellular Ca2+, and is completed within ~30 sec. Investigating this process in more detail, we recently learned that in addition to Ca2+-regulated exocytosis of lysosomes, plasma membrane repair involves a rapid form of endocytosis, which is triggered by release of the lysosomal enzyme acid sphingomyelinase (ASM) and is essential not only for the removal of transmembrane pores, but also for the repair of mechanical wounds16.
To determine the kinetics of cell resealing after permeabilization with pore-forming proteins, in our laboratory we adapted a live imaging methodology that had been used previously to assess resealing of laser-injured isolated muscle fibers17. This assay relies on properties of the lipophilic dye FM1-43, which stably intercalates into the outer leaflet of lipid bilayers increasing in fluorescence intensity. When the plasma membrane bilayer is disrupted extracellular dye gains access to intracellular membranes, providing a sensitive assay to detect plasma membrane injury and repair18,16,19,20. To adapt this assay for the assessment of cell resealing after permeabilization with pore-forming proteins, we pre-incubated cells at 4 &deg;C with the bacterial toxin streptolysin O (SLO), which binds to membrane cholesterol21. Synchronous cell permeabilization can then be easily achieved by moving the cells from ice to warm medium in a heated microscope stage, which activates the oligomerization and change in conformation that leads to transmembrane pore formation. An advantage of this approach, over the previously published assays using laser wounding, is that a larger number of cells can be analyzed simultaneously in a microscopic field, providing better sampling of the cell population. Given the mechanistic similarities between the cell resealing process seen after mechanical injury and permeabilization with pore-forming toxins, the assay we describe here provides a very versatile and powerful method for dissecting factors involved in the fundamental process of plasma membrane repair. As an example, we show that it is possible to use this assay to identify Ca2+ dependent and independent steps of the repair process.

<table border="1" fo:keep-together.within-page="always">
		<tbody>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Reagent</td>
		 </tr>
		 <tr>
			<td>HeLa 229 cell line</td>
			<td>ATCC</td>
			<td>CCL2-1</td>
			<td></td>
		 </tr>
		 <tr>
			<td>DMEM High Glucose </td>
			<td>Invitrogen</td>
			<td>11965</td>
			<td></td>
		 </tr>
		 <tr>
			<td>DMEM High Glucose No Calcium</td>
			<td>Invitrogen</td>
			<td>20168</td>
			<td></td>
		 </tr>
		 <tr>
			<td>FM1-43</td>
			<td>Invitrogen</td>
			<td>T3163</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Optimem Reduced Serum</td>
			<td>Invitrogen</td>
			<td>31985</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Lipofectamine RNAiMax</td>
			<td>Invitrogen</td>
			<td>13778</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Control medium GC content RNAi oligo</td>
			<td>Invitrogen</td>
			<td>12935300</td>
			<td></td>
		 </tr>
		 <tr>
			<td>SMPD1 RNAi oligo</td>
			<td>Invitrogen</td>
			<td>HSS143988</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Fetal Bovine Serum Heat-inactivated</td>
			<td>Gemini-Bioproducts</td>
			<td>100-106</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Sphingomyelinase from Bacillus cereus</td>
			<td>Sigma</td>
			<td>S7651</td>
			<td></td>
		 </tr>
		 <tr>
			<td>35 mm-glass bottom dishes</td>
			<td>MatTek</td>
			<td>P35G-0-14</td>
			<td></td>
		 </tr>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Material</td>
		 </tr>
		 <tr>
			<td>Inverted microscope</td>
			<td>Nikon</td>
			<td>Eclipse Ti</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Camera</td>
			<td>Hamamatsu Photonics</td>
			<td>C9100-50</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Spinning disk confocal microscope</td>
			<td>PerkinElmer</td>
			<td>UltraViewVoX</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Software analysis software</td>
			<td>PerkinElmer</td>
			<td>Volocity Suite</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Environmental chamber</td>
			<td>Pathology Devices</td>
			<td>LiveCell System Chamber</td>
		 </tr>
		</tbody>
 </table>

live imaging assay for assessing the roles of ca2+ and sphingomyelinase in the repair of pore-forming toxin wounds

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+.

Low concentrations of bacterial sphingomyelinase (SM) rescue plasma membrane repair in cells depleted in lysosomal acid sphingomyelinase.
Using the FM1-43 imaging assay, we previously showed that cells deficient for the lysosomal enzyme ASM and exposed to SLO wounding in the presence of Ca2+ have a plasma membrane defect is rescued by extracellular addition of the recombinant human enzyme19. Since the human lysosomal ASM has a low pH optimum, we investigated whether resealin...

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Live Imaging Assay for Assessing the Roles of Ca2+ and Sphingomyelinase in the Repair of Pore-forming Toxin Wounds

Live imaging of cells exposed to the lipophilic dye FM1-43 allows precise determination of the kinetics by which pore-forming toxins are removed from the plasma membrane. This is a sensitive assay that can be used to assess requirements for Ca2+, sphingomyelinase and other factors on plasma membrane repair.

Live Imaging Assay for Assessing the Roles of Ca2+ and Sphingomyelinase in the Repair of Pore-forming Toxin Wounds

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 ...

live-imaging-assay-for-assessing-roles-ca2-sphingomyelinase-repair

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11.6K Views.  University of Maryland. Live imaging of cells exposed to the lipophilic dye FM1-43 allows precise determination of the kinetics by which pore-forming toxins are removed from the plasma membrane. This is a sensitive assay that can be used to assess requirements for Ca2+, sphingomyelinase and other factors on plasma membrane repair.

Video: Live Imaging Assay for Assessing the Roles of Ca2+ and Sphingomyelinase in the Repair of Pore-forming Toxin Wounds

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Techniques for Imaging Ca2+ Signaling in Human Sperm

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in live cells. Here we describe the method used in our laboratories for loading suspensions of human sperm with Ca2+-reporting dyes and measuring the fluorescence signal during physiological stimulation. Motile cells are isolated by direct swim-up and incubated under capacitating conditions for 0-24 h, depending upon the experiment. The cell-permeant AM (acetoxy methyl ester) ester form of the Ca2+-reporting dye is then added to a cell aliquot and a period of 1 h is allowed for loading of the dye into the cytoplasm. We use visible wavelength dyes to minimize photo-damage to the cells, but this means that ratiometric recording is not possible. Advantages and disadvantages of this approach are discussed. During the loading period cells are introduced into an imaging chamber and allowed to adhere to a poly-D-lysine coated coverslip. At the end of the loading period excess dye and loose cells are removed by connection of the chamber to the perfusion apparatus. The chamber is perfused continuously, stimuli and modified salines are then added to the perfusion header. Experiments are recorded by time-lapse acquisition of fluorescence images and analyzed in detail offline, by manually drawing regions of interest. Data are normalized to pre-stimulus levels such that, for each cell (or part of a cell), a graph showing the Ca2+ response as % change in fluorescence is obtained.

Techniques for Imaging Ca2+ Signaling in Human Sperm

Stimulus-evoked [Ca2+]i signals of individual human sperm are assessed. Motile cells are loaded with Ca2+-sensitive fluorescent dye (AM-ester method) and immobilised in a perfusable chamber. Cells are imaged by time-lapse fluorescence microscopy and stimulated via the perfusing medium. Responses of single cells (or regions) are analysed offline using Excel.

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in ...

Electron Spin Resonance Micro-imaging of Live Species for Oxygen Mapping

This protocol describes an electron spin resonance (ESR) micro-imaging method for three-dimensional mapping of oxygen levels in the immediate environment of live cells with micron-scale resolution1. Oxygen is one of the most important molecules in the cycle of life. It serves as the terminal electron acceptor of oxidative phosphorylation in the mitochondria and is used in the production of reactive oxygen species. Measurements of oxygen are important for the study of mitochondrial and metabolic functions, signaling pathways, effects of various stimuli, membrane permeability, and disease differentiation. Oxygen consumption is therefore an informative marker of cellular metabolism, which is broadly applicable to various biological systems from mitochondria to cells to whole organisms. Due to its importance, many methods have been developed for the measurements of oxygen in live systems. Current attempts to provide high-resolution oxygen imaging are based mainly on optical fluorescence and phosphorescence methods that fail to provide satisfactory results as they employ probes with high photo-toxicity and low oxygen sensitivity. ESR, which measures the signal from exogenous paramagnetic probes in the sample, is known to provide very accurate measurements of oxygen concentration. In a typical case, ESR measurements map the probe's lineshape broadening and/or relaxation-time shortening that are linked directly to the local oxygen concentration. (Oxygen is paramagnetic; therefore, when colliding with the exogenous paramagnetic probe, it shortness its relaxation times.) Traditionally, these types of experiments are carried out with low resolution, millimeter-scale ESR for small animals imaging. Here we show how ESR imaging can also be carried out in the micron-scale for the examination of small live samples. ESR micro-imaging is a relatively new methodology that enables the acquisition of spatially-resolved ESR signals with a resolution approaching 1 micron at room temperature2. The main aim of this protocol-paper is to show how this new method, along with newly developed oxygen-sensitive probes, can be applied to the mapping of oxygen levels in small live samples. A spatial resolution of ~30 x 30 x 100 &mu;m is demonstrated, with near-micromolar oxygen concentration sensitivity and sub-femtomole absolute oxygen sensitivity per voxel. The use of ESR micro-imaging for oxygen mapping near cells complements the currently available techniques based on micro-electrodes or fluorescence/phosphorescence. Furthermore, with the proper paramagnetic probe, it will also be readily applicable for intracellular oxygen micro-imaging, a capability which other methods find very difficult to achieve.

This protocol describes a method for micron-scale three-dimensional imaging of oxygen concentration in the immediate environment of live cells by electron spin resonance microscopy.

This protocol describes an electron spin resonance (ESR) micro-imaging method for three-dimensional mapping of oxygen levels in the immediate environment ...

Live Cell Calcium Imaging Combined with siRNA Mediated Gene Silencing Identifies Ca2+ Leak Channels in the ER Membrane and their Regulatory Mechanisms

In mammalian cells, the endoplasmic reticulum (ER) plays a key role in protein biogenesis as well as in calcium signalling1. The heterotrimeric Sec61 complex in the ER membrane provides an aqueous path for newly-synthesized polypeptides into the lumen of the ER. Recent work from various laboratories suggested that this heterotrimeric complex may also form transient Ca2+ leak channels2-8. The key observation for this notion was that release of nascent polypeptides from the ribosome and Sec61 complex by puromycin leads to transient release of Ca2+ from the ER. Furthermore, it had been observed in vitro that the ER luminal protein BiP is involved in preventing ion permeability at the level of the Sec61 complex9,10. We have established an experimental system that allows us to directly address the role of the Sec61 complex as potential Ca2+ leak channel and to characterize its putative regulatory mechanisms11-13. This system combines siRNA mediated gene silencing and live cell Ca2+ imaging13. Cells are treated with siRNAs that are directed against the coding and untranslated region (UTR), respectively, of the SEC61A1 gene or a negative control siRNA. In complementation analysis, the cells are co-transfected with an IRES-GFP vector that allows the siRNA-resistant expression of the wildtype SEC61A1 gene. Then the cells are loaded with the ratiometric Ca2+-indicator FURA-2 to monitor simultaneously changes in the cytosolic Ca2+ concentration in a number of cells via a fluorescence microscope. The continuous measurement of cytosolic Ca2+ also allows the evaluation of the impact of various agents, such as puromycin, small molecule inhibitors, and thapsigargin on Ca2+ leakage. This experimental system gives us the unique opportunities to i) evaluate the contribution of different ER membrane proteins to passive Ca2+ efflux from the ER in various cell types, ii) characterize the proteins and mechanisms that limit this passive Ca2+ efflux, and iii) study the effects of disease linked mutations in the relevant components.

Live Cell Calcium Imaging Combined with siRNA Mediated Gene Silencing Identifies Ca2+ Leak Channels in the ER Membrane and their Regulatory Mechanisms

The endoplasmic reticulum plays a key role in protein biogenesis and in calcium homeostasis. We have established an experimental system that allows us to address the role of Ca2+ leak channels and to characterize their putative regulatory mechanisms. This system involves siRNA mediated gene silencing and live cell Ca2+ imaging.

In mammalian cells, the endoplasmic reticulum (ER) plays a key role in protein biogenesis as well as in calcium signalling1. The heterotrimeric Sec61 ...

An Explant Assay for Assessing Cellular Behavior of the Cranial Mesenchyme

The central nervous system is derived from the neural plate that undergoes a series of complex morphogenetic movements resulting in formation of the neural tube in a process known as neurulation. During neurulation, morphogenesis of the mesenchyme that underlies the neural plate is believed to drive neural fold elevation. The cranial mesenchyme is comprised of the paraxial mesoderm and neural crest cells. The cells of the cranial mesenchyme form a pourous meshwork composed of stellate shaped cells and intermingling extracellular matrix (ECM) strands that support the neural folds. During neurulation, the cranial mesenchyme undergoes stereotypical rearrangements resulting in its expansion and these movements are believed to provide a driving force for neural fold elevation. However, the pathways and cellular behaviors that drive cranial mesenchyme morphogenesis remain poorly studied. Interactions between the ECM and the cells of the cranial mesenchyme underly these cell behaviors. Here we describe a simple ex vivo explant assay devised to characterize the behaviors of these cells. This assay is amendable to pharmacological manipulations to dissect the signaling pathways involved and live imaging analyses to further characterize the behavior of these cells. We present a representative experiment demonstrating the utility of this assay in characterizing the migratory properties of the cranial mesenchyme on a variety of ECM components.

The cranial mesenchyme undergoes dramatic morphogenic movements that likely provides a driving force for elevation of the neural folds1,2. Here we describe a simple ex vivo explant assay to characterize the cellular behaviors of the cranial mesenchyme during neurulation. This assay has numerous applications including being amenable to pharmacological manipulations and live imaging analyses.

The central nervous system is derived from the neural plate that undergoes a series of complex morphogenetic movements resulting in formation of the ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Autophagy is a cellular response triggered by the lack of nutrients, especially the absence of amino acids. Autophagy is defined by the formation of double membrane structures, called autophagosomes, that sequester cytoplasm, long-lived proteins and protein aggregates, defective organelles, and even viruses or bacteria. Autophagosomes eventually fuse with lysosomes leading to bulk degradation of their content, with the produced nutrients being recycled back to the cytoplasm. Therefore, autophagy is crucial for cell homeostasis, and dysregulation of autophagy can lead to disease, most notably neurodegeneration, ageing and cancer.	
	Autophagosome formation is a very elaborate process, for which cells have allocated a specific group of proteins, called the core autophagy machinery. The core autophagy machinery is functionally complemented by additional proteins involved in diverse cellular processes, e.g. in membrane trafficking, in mitochondrial and lysosomal biology. Coordination of these proteins for the formation and degradation of autophagosomes constitutes the highly dynamic and sophisticated response of autophagy. Live cell imaging allows one to follow the molecular contribution of each autophagy-related protein down to the level of a single autophagosome formation event and in real time, therefore this technique offers a high temporal and spatial resolution.	
	Here we use a cell line stably expressing GFP-DFCP1, to establish a spatial and temporal context for our analysis. DFCP1 marks omegasomes, which are precursor structures leading to autophagosomes formation. A protein of interest (POI) can be marked with either a red or cyan fluorescent tag. Different organelles, like the ER, mitochondria and lysosomes, are all involved in different steps of autophagosome formation, and can be marked using a specific tracker dye. Time-lapse microscopy of autophagy in this experimental set up, allows information to be extracted about the fourth dimension, i.e. time. Hence we can follow the contribution of the POI to autophagy in space and time.

Time-lapse microscopy of fluorescently labeled autophagy markers allows monitoring of the dynamic autophagy response with high temporal resolution. Using specific autophagy and organelle markers in a combination of 3 different colors, we can follow the contribution of a protein to autophagosome formation in a robust spatial and temporal context.

Autophagy is a  cellular response triggered by the lack of nutrients, especially the absence of  amino acids. Autophagy is defined by the formation of ...

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Drosophila eye is widely used as a model for studies of development and neuronal degeneration. With the powerful mitotic recombination technique, elegant genetic screens based on clonal analysis have led to the identification of signaling pathways involved in eye development and photoreceptor (PR) differentiation at larval stages. We describe here the Tomato/GFP-FLP/FRT method, which can be used for rapid clonal analysis in the eye of living adult Drosophila. Fluorescent photoreceptor cells are imaged with the cornea neutralization technique, on retinas with mosaic clones generated by flipase-mediated recombination. This method has several major advantages over classical histological sectioning of the retina: it can be used for high-throughput screening and has proved an effective method for identifying the factors regulating PR survival and function. It can be used for kinetic analyses of PR degeneration in the same living animal over several weeks, to demonstrate the requirement for specific genes for PR survival or function in the adult fly. This method is also useful for addressing cell autonomy issues in developmental mutants, such as those in which the establishment of planar cell polarity is affected.

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Tomato/GFP-FLP/FRT method involves visualizing mosaic photoreceptor cells in living Drosophila. It can be used to follow individual photoreceptor cell fates in the retina for days or weeks. This method is ideal for studies of retinal degeneration and neurodegenerative diseases or photoreceptor cell development.

The Drosophila eye is widely used as a model  for studies of development and neuronal degeneration. With the powerful mitotic  recombination technique, ...

Adipo-Clear: A Tissue Clearing Method for Three-Dimensional Imaging of Adipose Tissue

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte precursors, immune cells, fibroblasts, blood vessels, and nerve projections. Although the molecular control of cell type specification and how these cells interact have been increasingly delineated, a more comprehensive understanding of these adipose-resident cells can be achieved by visualizing their distribution and architecture throughout the whole tissue. Existing immunohistochemistry and immunofluorescence approaches to analyze adipose histology rely on thin paraffin-embedded sections. However, thin sections capture only a small portion of tissue; as a result, the conclusions can be biased by what portion of tissue is analyzed. We have therefore developed an adipose tissue clearing technique, Adipo-Clear, to permit comprehensive three-dimensional visualization of molecular and cellular patterns in whole adipose tissues. Adipo-Clear was adapted from iDISCO/iDISCO+, with specific modifications made to completely remove the lipid stored in the tissue while preserving native tissue morphology. In combination with light-sheet fluorescence microscopy, we demonstrate here the use of the Adipo-Clear method to obtain high-resolution volumetric images&#160;of an entire adipose tissue.

Due to the high lipid content, adipose tissue has been challenging to visualize using traditional histological methods. Adipo-Clear is a tissue clearing technique that allows robust labeling and high-resolution volumetric fluorescent imaging of adipose tissue. Here, we describe the methods for sample preparation, pretreatment, staining, clearing, and mounting for imaging.

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte ...