The authors wish to thank the Academia Sinica Inflammation Core Facility, IBMS for research support. The core facility is funded by the Academia Sinica Core Facility and Innovative Instrument Project (AS-CFII-111-213). The authors thank the Common Equipment Core Facility of the Institute of Biomedical Sciences (IBMS), Academia Sinica (AS) for assisting the image acquisition.

1. AlPcS2a stock preparation
<ol>
	<li>Dissolve 10 mg of AlPcS2a in 400 &#181;L of 0.1 M NaOH. To improve solubility, heat the solution at 50 &#176;C and vortex.</li>
	<li>Mix the solution with 4 mL of phosphate-buffered saline (PBS). Then, filter the solution with a 0.22 &#181;m filter to remove the insoluble precipitates.</li>
	<li>Measure the concentration of the solution through a UV-Vis spectrophotometer. The extinction coefficient of AlPcS2a at 672 nm is 4 x 104 cm-1 M-1. Dilute the AlPcS2a solution with PBS to make a 1 mM solution. Make 1 mL aliquots and store at -20 &#176;C.</li>
</ol>
2. Transfection
NOTE: Gal3-GFP is applied as an indicator for live-cell imaging of lysosomal rupture.
<ol>
	<li>Maintain HeLa cell culture in DMEM supplemented with 10% Fetal Bovine Serum (FBS). After washing the cells with PBS, trypsinize the cells, then resuspend the cells in DMEM supplemented with 10% FBS. 
	NOTE: The method is applicable for most of the attached cell lines, including HeLa and A549 cell lines. In principle, despite not testing suspension cells, CALI assays could be performed with them.</li>
	<li>Dilute 300 ng of Gal3-GFP plasmid in 25 &#181;L of reduced serum medium, and then add 0.6 &#181;L of P3000 reagent. Gently mix. Dilute 0.45 &#181;L of Lipofectamine 3000 reagent in 25 &#181;L of reduced serum medium. Gently mix.</li>
	<li>Incubate for 5 min at room temperature. Mix diluted DNA and diluted Lipofectamine 3000, and then incubate for 10 min at room temperature.</li>
	<li>Mix the DNA-Lipofectamine mixture, 3 x 105 suspended HeLa cells, and 200 &#181;L of DMEM+10% FBS, and then seed the cells on the central glass region of a 35 mm glass button dish.</li>
</ol>
3. AlPcS2a staining
<ol>
	<li>To allow cell attachment, incubate the cells at 37 &#176;C in the incubator supplied with 5% CO2&#160;for at least 4 h.</li>
	<li>Dilute AlPcS2a in DMEM supplemented with 10% FBS to make a 1 &#181;M AlPcS2a solution. Pre-warm the AlPcS2a-containing medium in a 37 &#176;C water bath. Replace the culture medium with the AlPcS2a-containing medium.</li>
	<li>Incubate the cells at 37 &#176;C in the incubator supplied with 5% CO2 overnight (16-18 h).</li>
	<li>The following day, wash the cells twice with PBS to remove extracellular AlPcS2a. Replace the medium with fresh medium, and then incubate at 37 &#176;C for 4 h to allow residual dye along the endocytic pathway to accumulate into lysosomes. 
	&#8203;NOTE: To stain endosomes, incubate cells in 1 &#181;M AlPcS2a in DMEM containing 10% FBS for 15 min at 37 &#176;C. Next, wash cells twice with PBS and incubate them in pre-warmed medium before imaging .</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63962/63962fig01.jpg" /> 
Figure 1. A schematic figure representing the selective IV damage with AlPcS2a. The figure shows the schematics of selective IV damage. <a href="https://www.jove.com/files/ftp_upload/63962/63962fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63962/63962fig02.jpg" /> 
Figure 2. Lysosomal staining with AlPcS2a. Lysosomes labeled overnight with 1 &#181;M AlPcS2a in HeLa cells are positively stained with 50 nM green fluorescent dye. Scale bar: 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/63962/63962fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Sample imaging and light illumination
NOTE: IV damage within a subcellular region of a single cell or all the AlPcS2a-labeled cells in a culture dish can be performed. Bulk illumination of the whole culture dish allows quantitative study of this damage, including biochemical studies.
<ol>
	<li>Manipulate single cells by damaging lysosomes within a single cell, as described below.
	<ol>
		<li>Place the culture dish on the stage of a confocal microscope. Use the 488 nm and 561 nm lasers for exciting GFP and AlPcS2a, respectively.</li>
		<li>Circle a 5 x 5 &#181;m2 region of interest (ROI) on the AlPcS2a-labeled vesicles which are selected to be damaged.</li>
		<li>Pulse illuminate the ROI with a 633 nm laser of 0.21 mW for 70 repeats. Monitor the formation of Gal3 puncta as an indicator of lysosomal membrane permeabilization.</li>
	</ol>
	</li>
	<li>Damage all labeled cells in a culture dish as described below.
	<ol>
		<li>Place the culture dish on a platform. Illuminate the dish with 660 nm of collimated LED light (near-infrared light is ideal base on the excitation spectrum of AlPcS2a).</li>
		<li>Place the culture dish on the microscope and monitor the indicators of membrane permeabilization as mentioned in step 4.1.3.</li>
	</ol>
	</li>
	<li>Assess injury of IVs using the following indicators.
	<ol>
		<li>The contents of endosome or lysosome are most highly glycosylated, which are exposed to cytosol upon IV rupture. Rapidly label these using cytosolic glycan-binding galectins, including galectin-1, galectin-3, galectin-8, and galectin-916,20.</li>
		<li>The size of the wound can be determined by the release of membrane-impermeable dye as described in21.</li>
		<li>The difference between endosomes and lysosomes are their luminal pH value. Lysosomal rupture perturbs the luminal pH, which can be measured by using pH-sensitive dye, such as FITC-dextran3.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63962/63962fig03.jpg" /> 
Figure 3. AlPcS2a-mediated CALI induces local recruitment of TagRFP-galectin-3 (Gal3) to the lysosomes within the illumination region. Lysosomes in TagRFP-galectin-3-expressed HeLa cells were stained overnight with 1 &#181;M AlPcS2a, followed by focusing illumination with near-infrared light (633 nm) within the yellow square. AlPcS2a signal within the yellow square is photobleached, accompanied by the formation of TagRFP-galectin-3 puncta. Scale bar: 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/63962/63962fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors declare no conflict of interest.

AlPcS2a binds to the plasma membrane, then is internalized by endocytosis and eventually accumulates in lysosomes. AlPcS2a can thus be localized in the subcellular compartments by adjusting the incubation duration. A limitation of this methodology is that only a sub-population of IVs could be labeled by AlPcS2a through endocytosis because there are many other membrane sources of IVs, such as ER and Golgi apparatus. In addition, the selective labeling of AlPcS2a into early or late endosomes would be challenging, however, fluorescent markers can be used to differentiate early-formed endosomes from late-formed ones during subcellular imaging. CALI-induced IV damage can be induced with a variety of dyes3,24.
The intensity of damage can be controlled by the power and illumination duration of light. Higher intensity of damage may result in cell shrinking or necrosis. Here, several indicators of IV damage have been shown, which could be used to determine the experimental conditions. Additionally, these indicators may also serve as markers for damaged-IVs and provide information about their subcellular distribution over time. 
Clinically, AlPcS2a is designed to improve the delivery of therapeutic agents in a site and time-specific manner through photochemical internalization (PCI)25. However, unavoidable side effects, such as cell apoptosis or differentiation, may also accompany the PCI treatment26. Since such effects may be the consequence of induction of lysosomal damage, the provided assays present a potential tool for preclinical use.
In this study, the AlPcS2a-mediated CALI protocols describe the selective control of the damage size (local and part of endosomes or lysosomes). Scientists can monitor their subcellular behavior, while the rest of the IVs remain intact and could be treated as a control. This methodology has been applied in several studies. For example, damaged lysosomes are labeled sequentially with ubiquitin and microtubule-associated protein light chain 3 (LC3), and undergo autophagic clearance3. Furthermore, galectin-3 and galectin-8 labeling onto damaged endosomes are controlled by cell surface glycan27. Since AlPcS2a has been used in vivo for PCI, AlPcS2a-mediated CALI could also be used in vivo. This protocol provides a robust tool to study cell biology using AlPcS2a-mediated CALI. These studies support that the AlPcS2a-mediated CALI is a robust tool for cell biology studies.

Endosomes, a type of intracellular vesicle (IV), are formed by endocytosis and then mature into lysosomes. Various intracellular signal pathways are involved in the formation of IVs; additionally, different intrinsic and extrinsic stimuli can damage IVs (e.g., pathogens can escape from the bounded membrane during infection and enter into cytoplasm1). This is usually accompanied by the rupture of endocytotic vesicles2. Therefore, the techniques for targeting and damaging IVs can be used in related studies3.
Photodynamic therapy (PDT) is a light-dependent therapy to combat diseases by killing tumors or pathogens4. In PDT, targeted cells are labeled with non-toxic chromophores, called photosensitizers, that can be locally activated by light illumination5,6. Photosensitizers absorb energy from light and transform into an excited singlet state, leading to the long-lived excited triplet state. Photosensitizers of the triplet state can undergo electron or energy transfer and form reactive oxygen species (ROS) in the presence of oxygen, and can spatially destroy labeled cells within the illumination region7. The consequence varies depending on the power of light8. By controlling the concentration of photosensitizers and the intensity of light illumination, targeted biomolecules can be selectively inactivated without cell lysis, termed as chromophore-assisted light inactivation (CALI)9. With the significant development of photosensitizers that can selectively label various subcellular targets, CALI has become a valuable tool to control light-mediated inactivation of biomolecules for small biomolecules such as nucleotides and proteins, as well as organelles such as mitochondria and endo-lysosomes3,10,11,12,13.
Compared to CALI, chemical or physical methods are also used to impair membranes, such as bacterial toxin14,15 and Leu-Leu-OMe16 treatment for lysosomal damage. However, these methods display bulk impairment of IVs within cells. Robust photosensitizers (i.e., Al(III) phthalocyanine chloride disulfonic acid (AlPcS2a)) are used in CALI; AlPcS2a, targeting the lysosomes through endocytosis, is used to rupture endosomes or lysosomes in a controlled region17. AlPcS2a is a cell membrane-impermeable phthalocyanine-based chromophore that binds to lipid on plasma membrane and is internalized through endocytosis and eventually accumulates within the lysosome through the endocytic pathway18. It absorbs light within a near-infrared spectral region and generates singlet oxygen, a major ROS generated by excited AlPcS2a18. Singlet oxygen decaying rapidly limits its diffusion and reaction distance within a tiny region in cells (approximately 10-20 nm)19. By adjusting the duration of AlPcS2a incubation and light illumination, spatiotemporal control of the damage of IVs within a subcellular area is allowed. CALI therefore becomes a powerful tool for examining the consequences of IV damage, and the formation and regulation of IVs.
In this study, a specific protocol of CALI using AlPcS2a as a photosensitizer is addressed. This protocol can be applied to various types of IVs, including endosomes and lysosomes, and used to examine the follow-up responses after membrane rupture. HeLa cells expressing fluorophore-conjugated galectin-316,20 revealed after lysosome rupture are used to demonstrate this protocol.

<table><tbody><tr><td>&lt;strong&gt;Reagent&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Al(III) Phthalocyanine Chloride Disulfonic acid (AlPcS&lt;sub&gt;2a&lt;/sub&gt;)</td><td>Frontier Scientific</td><td>P40632</td><td></td></tr><tr><td>Culture dish</td><td>ibidi</td><td>812128-200</td><td></td></tr><tr><td>Culture Medium</td><td></td><td></td><td>DMEM supplemented with 10% FBS and 100 U/mL penicillin G and 100 mg/mL Streptomycin</td></tr><tr><td>DMEM</td><td>Gibco</td><td>11965092</td><td></td></tr><tr><td>FBS</td><td>Thermo Fisher Scientific</td><td>A4736301</td><td></td></tr><tr><td>Gal3-GFP plasmid</td><td>addgene</td><td></td><td></td></tr><tr><td>Lipofectamine 3000 kit</td><td>Thermo Fisher Scientific</td><td>L3000008</td><td></td></tr><tr><td>LysoTracker Green DND-26</td><td>Thermo Fisher Scientific</td><td>L7526</td><td>green fluorescent dye</td></tr><tr><td>Multiwall plate</td><td>perkinelmer</td><td>PK-6005550</td><td></td></tr><tr><td>NaOH</td><td>Thermo Fisher Scientific</td><td>Q15895</td><td></td></tr><tr><td>OptiMEM</td><td>Thermo Fisher Scientific</td><td>31985070</td><td></td></tr><tr><td>Penicillin-streptomycin</td><td>Gibco</td><td>15140163</td><td></td></tr><tr><td>Phosphate-Buffered Saline (PBS)</td><td>Gibco</td><td>21600-069</td><td>137 mM NaCl, 2.7 mM KCl, 10mM Na2HPO4, 1.8 mM KH2PO4</td></tr><tr><td>&lt;strong&gt;Cell line&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>HeLa Cell Line</td><td>ATCC</td><td>CCL-2</td><td>The methods are applicable for most of the attached cell lines. Conditions must be determined individually.</td></tr><tr><td>&lt;strong&gt;Equipments&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>0.22 &amp;micro;m Filter</td><td>Merck</td><td>SLGV013SL</td><td></td></tr><tr><td>Collimated LED Light&amp;nbsp; (660nm)</td><td>Thorlabs</td><td>M660L3-C1 and DC2100</td><td>Near-infared light is ideal base on the excitation spectrum of AlPcS2a.</td></tr><tr><td>Confocal microscopy</td><td>Carl Zeiss</td><td>LSM 780</td><td>An incubation system is required for long-term imaging.</td></tr><tr><td>NanoDrop 2000/2000c Spectrophotometers</td><td>Thermo Fisher Scientific</td><td></td><td></td></tr><tr><td>Red LED light</td><td>Tholabs</td><td>M660L4-C1</td><td></td></tr></tbody></table>

a photodynamic approach to study function of intracellular vesicle rupture

Intracellular vesicles (IVs) are formed through endocytosis of vesicles into cytoplasm. IV formation is involved in activating various signal pathways through permeabilization of IV membranes and the formation of endosomes and lysosomes. A method named chromophore-assisted laser inactivation (CALI) is applied to study the formation of IVs and the materials in controlling IV regulation. CALI is an imaging-based photodynamic methodology to study the signaling pathway induced by membrane permeabilization. The method allows spatiotemporal manipulation of the selected organelle to be permeabilized in a cell. The CALI method has been applied to observe and monitor specific molecules through the permeabilization of endosomes and lysosomes. The membrane rupture of IVs is known to selectively recruit glycan-binding proteins, such as galectin-3. Here, the protocol describes the induction of IV rupture by AlPcS2a and the use of galectin-3 as a marker to label impaired lysosomes, which is useful in studying the downstream effects of IV membrane rupture and their downstream effects under various situations.

A schematic figure representing the AlPcS2a-induced damage of IV, including endosome and lysosome, has been shown (Figure 1).
Commercially available markers can be used to determine the AlPcS2a staining conditions. For example, AlPcS2a puncta and green fluorescent dye22 colocalization (Figure 2).
Fluorophore-labeled galectin-3 can be applied as an indicator for monitoring IV damage (Figure 3). Moreover, the location of Gal3 puncta could also be tracked for assaying the downstream signaling pathway, including lysosome repair and lysophagy3,23. The rapid accumulation of Gal3 from cytosol to damaged IV would significantly increase the intensity of Gal3, which would make the intensity of the Gal3 puncta saturated if the contrast of the images is adjusted to show cytosolic Gal3. In fact, the images also showed a slight decrease in cytosolic Gal3.
In summary, these results indicate that AlPcS2a-based CALI is able to control local lysosomal rupture within the ROI, leaving the rest of the lysosomes intact.

Watch this Scientific Journal Video about A Photodynamic Approach to Study Function of Intracellular Vesicle Rupture at JoVE.com

A Photodynamic Approach to Study Function of Intracellular Vesicle Rupture

AlPcS2a-mediated chromophore-assisted laser inactivation (CALI) is a powerful tool for studying spatiotemporal damage of intracellular vesicles (IVs) in live cells.

Intracellular vesicles (IVs) are formed through endocytosis of vesicles into cytoplasm. IV formation is involved in activating various signal pathways ...

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Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

1.1K Views.  Academia Sinica. AlPcS2a-mediated chromophore-assisted laser inactivation (CALI) is a powerful tool for studying spatiotemporal damage of intracellular vesicles (IVs) in live cells.

Video: A Photodynamic Approach to Study Function of Intracellular Vesicle Rupture

Long-term Live-cell Imaging to Assess Cell Fate in Response to Paclitaxel

Live-cell imaging is a powerful technique that can be used to directly visualize biological phenomena in single cells over extended periods of time. Over the past decade, new and innovative technologies have greatly enhanced the practicality of live-cell imaging. Cells can now be kept in focus and continuously imaged over several days while maintained under 37 &#176;C and 5% CO2 cell culture conditions. Moreover, multiple fields of view representing different experimental conditions can be acquired simultaneously, thus providing high-throughput experimental data. Live-cell imaging provides a significant advantage over fixed-cell imaging by allowing for the direct visualization and temporal quantitation of dynamic cellular events. Live-cell imaging can also identify variation in the behavior of single cells that would otherwise have been missed using population-based assays. Here, we describe live-cell imaging protocols to assess cell fate decisions following treatment with the anti-mitotic drug paclitaxel. We demonstrate methods to visualize whether mitotically arrested cells die directly from mitosis or slip back into interphase. We also describe how the fluorescent ubiquitination-based cell cycle indicator (FUCCI) system can be used to assess the fraction of interphase cells born from mitotic slippage that are capable of re-entering the cell cycle. Finally, we describe a live-cell imaging method to identify nuclear envelope rupture events.

Live-cell imaging provides a wealth of information on single cells or whole populations that is unattainable by fixed cell imaging alone. Here, live-cell imaging protocols to assess cell fate decisions following treatment with the anti-mitotic drug paclitaxel are described.

Live-cell imaging is a powerful technique that can be used to directly visualize biological phenomena in single cells over extended periods of time. Over ...

Cancer Research

Single Cell Measurements of Vacuolar Rupture Caused by Intracellular Pathogens

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

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

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

Membrane Remodeling of Giant Vesicles in Response to Localized Calcium Ion Gradients

In a wide variety of fundamental cell processes, such as membrane trafficking and apoptosis, cell membrane shape transitions occur concurrently with local variations in calcium ion concentration. The main molecular components involved in these processes have been identified; however, the specific interplay between calcium ion gradients and the lipids within the cell membrane is far less known, mainly due to the complex nature of biological cells and the difficultly of observation schemes. To bridge this gap, a synthetic approach is successfully implemented to reveal the localized effect of calcium ions on cell membrane mimics. Establishing a mimic to resemble the conditions within a cell is a severalfold problem. First, an adequate biomimetic model with appropriate dimensions and membrane composition is required to capture the physical properties of cells. Second, a micromanipulation setup is needed to deliver a small amount of calcium ions to a particular membrane location. Finally, an observation scheme is required to detect and record the response of the lipid membrane to the external stimulation. This article offers a detailed biomimetic approach for studying the calcium ion-membrane interaction, where a lipid vesicle system, consisting of a giant unilamellar vesicle (GUV) connected to a multilamellar vesicle (MLV), is exposed to a localized calcium gradient formed using a microinjection system. The dynamics of the ionic influence on the membrane were observed using fluorescence microscopy and recorded at video frame rates. As a result of the membrane stimulation, highly curved membrane tubular protrusions (MTPs) formed inside the GUV, oriented away from the membrane. The described approach induces the remodeling of the lipid membrane and MTP production in an entirely contactless and controlled manner. This approach introduces a means to address the details of calcium ion-membrane interactions, providing new avenues to study the mechanisms of cell membrane reshaping.

We present a technique for contactless micromanipulation of vesicles, using localized calcium ion gradients. Microinjection of a calcium ion solution, in the vicinity of a giant lipid vesicle, is utilized to remodel the lipid membrane, resulting in the production of membrane tubular protrusions.

In a wide variety of fundamental cell processes, such as membrane trafficking and apoptosis, cell membrane shape transitions occur concurrently with local ...

Chemistry

TIRFM and pH-sensitive GFP-probes to Evaluate Neurotransmitter Vesicle Dynamics in SH-SY5Y Neuroblastoma Cells: Cell Imaging and Data Analysis

Synaptic vesicles release neurotransmitters at chemical synapses through a dynamic cycle of fusion and retrieval. Monitoring synaptic activity in real time and dissecting the different steps of exo-endocytosis at the single-vesicle level are crucial for understanding synaptic functions in health and disease.
Genetically-encoded pH-sensitive probes directly targeted to synaptic vesicles and Total Internal Reflection Fluorescence Microscopy (TIRFM) provide the spatio-temporal resolution necessary to follow vesicle dynamics. The evanescent field generated by total internal reflection can only excite fluorophores placed in a thin layer (&#60;150 nm) above the glass cover on which cells adhere, exactly where the processes of exo-endocytosis take place. The resulting high-contrast images are ideally suited for vesicles tracking and quantitative analysis of fusion events.
In this protocol, SH-SY5Y human neuroblastoma cells are proposed as a valuable model for studying neurotransmitter release at the single-vesicle level by TIRFM, because of their flat surface and the presence of dispersed vesicles. The methods for growing SH-SY5Y as adherent cells and for transfecting them with synapto-pHluorin are provided, as well as the technique&#160;to perform TIRFM and imaging. Finally, a strategy aiming to select, count, and analyze fusion events at whole-cell and single-vesicle levels is presented.
To validate the imaging procedure and data analysis approach, the dynamics of pHluorin-tagged vesicles are&#160;analyzed under resting and stimulated (depolarizing potassium concentrations) conditions. Membrane depolarization increases the frequency of fusion events and causes a parallel raise of the net fluorescence signal recorded in whole cell. Single-vesicle analysis reveals modifications of fusion-event behavior (increased peak height and width). These data suggest that potassium depolarization not only induces a massive neurotransmitter release but also modifies the mechanism of vesicle fusion and recycling.
With the appropriate fluorescent probe, this technique can be employed in different cellular systems to dissect the mechanisms of constitutive and stimulated secretion.

This paper provides a method for investigating neurotransmitter vesicle dynamics in neuroblastoma cells, using a synaptobrevin2-pHluorin construct and Total Internal Reflection Fluorescence Microscopy. The strategy developed for image processing and data analysis is also reported.

Synaptic vesicles release neurotransmitters at chemical synapses through a dynamic cycle of fusion and retrieval. Monitoring synaptic activity in real ...

Neuroscience

Cell Membrane Repair Assay Using a Two-photon Laser Microscope

Numerous pathophysiological insults can cause damage to cell membranes and, when coupled with innate defects in cell membrane repair or integrity, can result in disease. Understanding the underlying molecular mechanisms surrounding cell membrane repair is, therefore, an important objective to the development of novel therapeutic strategies for diseases associated with dysfunctional cell membrane dynamics. Many in vitro and in vivo studies aimed at understanding cell membrane resealing in various disease contexts utilize two-photon laser ablation as a standard for determining functional outcomes following experimental treatments. In this assay, cell membranes are subjected to wounding with a two-photon laser, which causes the cell membrane to rupture and fluorescent dye to infiltrate the cell. The intensity of fluorescence within the cell can then be monitored to quantify the cell's ability to reseal itself. There are several alternative methods for assessing cell membrane response to injury, as well as great variation in the two-photon laser wounding approach itself, therefore, a single, unified model of cell wounding would beneficially serve to decrease the variation between these methodologies. In this article, we outline a simple two-photon laser wounding protocol for assessing cell membrane repair in vitro in both healthy and dysferlinopathy patient fibroblast cells transfected with or without a full-length dysferlin plasmid.

Cell membrane wounding via two-photon laser is a widely used method for assessing membrane resealing ability and can be applied to multiple cell types. Here, we describe a protocol for in vitro live-imaging of membrane resealing in dysferlinopathy patient cells following two-photon laser ablation.

Numerous pathophysiological insults can cause damage to cell membranes and, when coupled with innate defects in cell membrane repair or integrity, can ...

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+.

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.

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 ...

Biology

Imaging Cell Membrane Injury and Subcellular Processes Involved in Repair

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.

The process of healing injured cells involves trafficking of specific proteins and subcellular compartments to the site of cell membrane injury. This protocol describes assays to monitor these processes.

The ability of injured cells to heal is a fundamental cellular process, but cellular and molecular mechanisms involved in healing injured cells are poorly ...

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 ...

Characterization of MLKL-mediated Plasma Membrane Rupture in Necroptosis

Necroptosis is a programmed cell death pathway triggered by activation of receptor interacting protein kinase 3 (RIPK3), which phosphorylates and activates the mixed lineage kinase-like domain pseudokinase, MLKL, to rupture or permeabilize the plasma membrane. Necroptosis is an inflammatory pathway associated with multiple pathologies including autoimmunity, infectious and cardiovascular diseases, stroke, neurodegeneration, and cancer. Here, we describe protocols that can be used to characterize MLKL as the executioner of plasma membrane rupture in necroptosis. We visualize the process of necroptosis in cells using live-cell imaging with conventional and confocal fluorescence&#160;microscopy, and in fixed cells using electron microscopy, which together revealed the redistribution of MLKL from the cytosol to the plasma membrane prior to induction of large holes in the plasma membrane. We present in vitro nuclear magnetic resonance (NMR) analysis using lipids to identify putative modulators of MLKL-mediated necroptosis. Based on this method, we identified quantitative lipid-binding preferences and phosphatidyl-inositol phosphates (PIPs) as critical binders of MLKL that are required for plasma membrane targeting and permeabilization in necroptosis.

We report methods for characterization of MLKL-mediated plasma membrane rupture in necroptosis including conventional and confocal live-cell microscopy imaging, scanning electron microscopy, and NMR-based lipid binding.

Necroptosis is a programmed cell death pathway triggered by activation of receptor interacting protein kinase 3 (RIPK3), which phosphorylates and ...

Laser Induced Thermoplasmonic Puncture for the Study of Plasma Membrane Repair

A Thermoplasmonic Approach for Investigating Plasma Membrane Repair in Living Cells and Model Membranes

The cell membrane is crucial for cell survival, and ensuring its integrity is essential as the cell experiences injuries throughout its entire life cycle. To prevent damage to the membrane, cells have developed efficient plasma membrane repair mechanisms. These&#160;repair mechanisms can be studied by combining confocal microscopy and nanoscale thermoplasmonics to identify and investigate the role of key proteins, such as annexins, involved in surface repair in living cells and membrane model systems.
The puncturing method employs a laser to induce highly localized heating upon nanoparticle irradiation. The use of near-infrared light minimizes phototoxicity in the biological sample, while the majority of the absorption takes place in the near-infrared resonant plasmonic nanoparticle. This thermoplasmonic method has been exploited for potential photothermal and biophysical research to enhance the understanding of intracellular mechanisms and cellular responses through vesicle and cell fusion studies. The approach has shown to be complementary to existing methods for membrane disruption, such as mechanically, chemically, or optically induced injuries, and provides a high level of control by inflicting extremely localized injuries. The extent of the injury is limited to the vicinity of the spherical nanoparticle, and no detrimental damage occurs along the beam path as opposed to pulsed lasers using different wavelengths. Despite certain limitations, such as the formation of nanobubbles, the thermoplasmonic method offers a unique tool for investigating cellular responses in plasma membrane repair in an almost native environment without compromising cell viability.
When integrated with confocal microscopy, the puncturing method can provide a mechanistic understanding of membrane dynamics in model membrane systems as well as quantitative information on protein responses to membrane damage, including protein recruitment and their biophysical function. Overall, the application of this method to reduced model systems can enhance our understanding of the intricate plasma membrane repair machinery in living cells.

Author Spotlight: Exploring Plasma Membrane Repair Mechanisms with Innovative Thermoplasmonic Puncturing 

The thermoplasmonic puncture method integrates confocal microscopy, optical tweezers, and gold nanoparticles to study protein responses during plasma membrane repair in cells and giant unilamellar vesicles. The technique enables rapid and localized membrane puncture, allowing the identification of key proteins and their functional roles in the intricate plasma membrane repair machinery.

The cell membrane is crucial for cell survival, and ensuring its integrity is essential as the cell experiences injuries throughout its entire life cycle. ...