The authors would like to acknowledge the Universit&#224; degli Studi di Milano for support to Eliana Di Cairano (Post-doctoral fellowship) and Stefania Moretti (Ph.D. fellowship). This work was supported by the University Research Program PUR to C.P.
We would like to thank Prof. Jeremy M. Henley, School of Biochemistry, University of Bristol, United Kingdom, for the pHluorin construct and Dr. Dotti Francesco for assistance in data analysis, and Silvia Marsicano for technical assistance.

1. Cell Culture and Transfection
<ol>
	<li>SH-SY5Y cell culture 
	NOTE: The experiments have been performed using the human neuroblastoma SH- SY5Y (ATCC# CRL-2266)15. SH-SY5Y cells grow as a mixture of floating clusters and adherent cells. Follow the instructions reported in the protocol (cell density, splitting ratio, etc.) to have cells that grow firmly attached to glass cover, which is crucial for TIRFM.
	<ol>
		<li>Before starting, under the laminar flow biosafety cabinet, make the opportune volume of sterile phosphate buffer saline solution (PBS) and culture medium.
		<ol>
			<li>Make 50 ml of PBS with concentrations of 150 mM NaCl, 24 mM phosphate buffer, pH 7.4. Filter the solution.</li>
			<li>Make 50 ml of cell medium from Dulbecco&#8217;s modified Eagle medium (DMEM) with high glucose, 10% heat-inactivated fetal bovine serum (FBS), penicillin (100 U/ml), streptomycin (100 &#181;g/ml), L-glutamine (2 mM), and sodium pyruvate (1 mM). Filter the solution.</li>
		</ol>
		</li>
		<li>Remove complete growth medium and wash the cells with 3 ml of PBS.</li>
		<li>Incubate cells with 2 ml of 0.05% trypsin-ethylenediaminetetraacetic acid (EDTA) (for 6 cm Petri dish) for 5 min at 37 &#176;C, 5% CO2 and detach cells using pipette.</li>
		<li>Inactivate trypsin by adding 2 ml of DMEM, and collect cells by centrifugation at 300 x g for 5 min.</li>
		<li>Remove the supernatant, add 1 ml of DMEM to the pellet and pipette the solution up and down sufficiently to disperse cells into a single cell suspension.</li>
		<li>Split them 1:4 in a new 6 cm diameter Petri dish containing 3 ml of complete medium. Maintain cells in culture in 6 cm diameter Petri dishes, at 37 &#176;C in a 5% CO2 incubator. Sub-culture once a week or when they have covered 80 - 90% of the surface area.</li>
	</ol>
	</li>
	<li>SH-SY5Y cell plating for imaging
	<ol>
		<li>For TIRFM experiments, plate cells onto glass covers. Employ glass covers with 0.17 &#177; 0.005 mm thickness and a 1.5255 &#177; 0.00015 refractive index. Before starting, prepare the glass coverslips as follow:
		<ol>
			<li>Clean glass covers with 90% ethanol, O/N.</li>
			<li>Rinse them thoroughly in distilled water (three changes of distilled water). Dry glass covers in a drying oven.</li>
			<li>Place covers in glass Petri dishes and sterilize in a preheated oven at 200 &#176;C for 3 hr.</li>
		</ol>
		</li>
		<li>The day before transfection, place each coverslip in a 3.5 cm Petri dish, add 1 ml of culture medium and incubate at 37 &#176;C in a 5% CO2 incubator.</li>
		<li>Trypsinize cells as described in steps 1.1.3 - 1.1.5, suspend the cell pellet in 1 ml of complete medium and count. Calculate the correct volume of cell suspension to add to each Petri dish to yield 3 x 105 cells/well. This density is required for optimal cell growth and efficient transfection. Incubate at 37 &#176;C in a 5% CO2 incubator O/N.</li>
	</ol>
	</li>
	<li>SH-SY5Ytransfection by polyethylenimine (PEI) 
	NOTE: To visualize synaptic vesicles dynamics, pCB6 vector containing synapto-pHluorin has been used. The synapto-pHluorin has been generated by in frame fusion of a pH-sensitive variant of the green fluorescent protein (GFP)16 and the vesicular membrane protein synaptobrevin 2. The construct has been extensively employed to investigate synaptic vesicle properties within neurons9.
	<ol>
		<li>Before starting transfection, make 10 ml of the following solutions. Keep the solutions as maximal as 1 month.
		<ol>
			<li>Make a 150 mM NaCl solution. Adjust to pH 5.5 with 0.01 N HCl.</li>
			<li>Make a PEI solution at 10% polyethylenimine (PEI; 25 kDa linear) in 150 mM NaCl solution. The pH of solution rises to 8.8. Adjust pH to 7.8 with 0.01 N HCl.</li>
		</ol>
		</li>
		<li>24 hr after plating, remove the medium and refresh with 1.5 ml of complete medium. Keep the cells at 37 &#176;C, in a 5% CO2 incubator.</li>
		<li>Under the laminar flow biosafety cabinet, in a 1.5 ml microfuge tube, add 3 &#181;g of plasmid DNA to 25 &#956;l of 150 mM NaCl solution and 100 &#956;l of PEI solution per 3.5 cm Petri dish.</li>
		<li>Vortex for 10 sec, then incubate the DNA/PEI mixture for 30 min at RT.</li>
		<li>Carefully add the DNA/PEI mixture to the Petri dish containing coverslips with cells and gently shake to equally distribute the reagent in the Petri dish.</li>
		<li>After 4 hr change the medium and incubate the cells O/N at 37 &#176;C in a 5% CO2 incubator. Perform imaging experiments 24 - 48 hr after transfection.</li>
	</ol>
	</li>
</ol>
2. Cell Imaging by Total Internal Reflection Fluorescence Microscopy (TIRFM)
<ol>
	<li>Imaging set-up
	<ol>
		<li>Perform TIRF imaging with the set-up described in Figure 1. It comprises a motorized inverted microscope (Figure 1, inset A), the laser source (Figure 1, inset B) and the TIRF-slider (Figure 1, inset C). Reach TIRFM illumination through a high numerical aperture (NA 1.45 Alpha Plan-Fluar) 100X oil, immersion objective.</li>
		<li>For TIRFM illumination, employ a multi-line (458/488/514 nm) 100 mW argon-ion laser. Using a mono mode fiber, introduce the linearly polarized laser light into the beam path, via the TIRF slider. Insert the TIRF slider into the luminous field diaphragm plane of the reflected-light beam path.
		<ol>
			<li>For wide field illumination, connect the microscope to a conventional mercury short-arc lamp HBO white light. A polarization-maintaining double prism in the slider ensures the simultaneous combination of TIRF illumination and white light.</li>
		</ol>
		</li>
		<li>Filter the laser light with an excitation filter (band width 488/10 nm) mounted on a filter wheel, introduced into the laser path. Employ a high speed, software-controlled, shutter to allow fast control of laser illumination. For pHluorin analysis, mount a band pass 525/50 nm emission filter. Capture digital images (512 x 512 pixels) on a cooled Fast CCD camera with the Image ProPlus software.</li>
	</ol>
	</li>
	<li>Achieving TIRF illumination (Figure 2)
	<ol>
		<li>Turn on the lasers, the computer, the camera, the filter wheel, and the shutter controllers; then, wait 20 min before starting the experiment as the lasers need to warm up and stabilize.</li>
		<li>Before imaging, make the opportune volume of the following solutions.
		<ol>
			<li>Make 50 ml of Krebs (KRH) solution at 125 mM NaCl, 5 mM KCl, 1.2 mM MgSO4, 1.2 mM KH2PO4, 25 mM 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES) (buffered to pH 7.4), 2 mM CaCl2, and 6 mM glucose.</li>
			<li>Make 10 ml of KCl-KRH solution (pH 7.4) at 80 mM NaCl, 50 mM KCl, 1.2 mM MgSO4, 1.2 mM KH2PO4, 25 mM HEPES (buffered to pH 7.4), 2 mM CaCl2, and 6 mM glucose.</li>
		</ol>
		</li>
		<li>Remove the glass cover with transfected cells and insert it in the appropriate imaging chamber. Assemble the chamber and add 500 &#181;l of KRH solution in the center of the glass.</li>
		<li>Add oil over the objective. Place the imaging chamber on the stage of the microscope and position the objective under the glass coverslip. Position the safe cover over the sample.</li>
		<li>In epifluorescence mode, focus on the coverslip (upper surface) and choose transfected cells placed in the chamber center. Select cells whose fluorescent signal can be clearly recorded using an exposure time below 80 msec.</li>
		<li>Under software control, switch to TIRF illumination in live mode.</li>
		<li>To set the TIRF configuration, check the position of the beam that emerges out of the objective, on the sample cover (Figure 2B). When the beam is positioned in the center of the objective lens (Figure 2A, left), a spot is visible in the center of the TIRF sample cover (Figure 2B, left) and the cell is imaged in epifluorescence mode (several focus planes, high background fluorescence; Figure 2C, left).</li>
		<li>To reach the critical angle, move the focused spot in the Y direction (forward or backward;&#160;Figure 2B, center) using the angle adjustment screw on the TIRF slider (Figure 1C). When the beam converges on the sample plane at an angle larger than the critical angle (Figure 2A, right), the spot disappears and a straight, thin, focused line is evident in the middle of the sample cover (Figure 2B, right).</li>
		<li>To fine-tune the TIRF angle use the cell sample (Figure 2C). Watch the fluorescence image on the video, at this stage, an epifluorescence-like image is still visible. Gently, move the screw until TIRF condition is achieved: only one optical plane of the cell is in focus (i.e., the plasma membrane in contact with the cover-slip), this results in a flat image with high contrast (Figure 2C, right).</li>
	</ol>
	</li>
	<li>Sample imaging
	<ol>
		<li>Set the single-channel time-lapse experiment. To minimize photobleaching, capture the image using low exposure time and high gain. Appropriate exposure times are between 40 - 80 msec. Acquire images at 1 - 2 Hz sampling frequency. Vesicle kinetics may be better appreciated sampling at higher frequency (10 Hz). The regular time of observation is usually 2 min.</li>
		<li>Add 500 &#181;l of KRH solution and record cells in TIRFM mode. This is the resting condition. Save the time sequential images.</li>
		<li>Focus on the same cell and record under the same conditions of resting (laser power, time exposure, frame number). After five frames, add 500 &#181;l of KCl-KRH solution and keep KCl in the chamber. This is the stimulated condition; save the time sequential images.</li>
	</ol>
	</li>
</ol>
3. Image Analysis and Data Processing
NOTE: To analyze images, macros have been developed in the lab, based on existing functions of the image analysis software; similar macros are available online (URL provided in Table of Materials and Equipment).
<ol>
	<li>Fluorescence intensity quantification
	<ol>
		<li>Use a &#8220;Sequence fluorescence intensity&#8221; macro for fluorescence intensity quantification in a region of interest (ROI) of the image, over the course of the movie.</li>
		<li>Open the time-sequential images. Go to the macro menu and select &#39;Sequence fluorescence intensity&#39;. In the &#39;Analysis&#39; window appears &#8220;select the ROI&#8221;.</li>
		<li>Choose one of the selection tools in the menu to create the ROI. Place 3 ROIs in regions of the cell membrane without spots (background ROI). Employ this &#8220;background ROI&#8221; to evaluate the photobleaching and to set the threshold for fusion event analysis (Figure 3A).</li>
	</ol>
	</li>
	<li>Photobleaching correction and threshold determination (Figure 3B)
	<ol>
		<li>To evaluate the photobleaching, open the fluorescence intensity rows &#8220;background ROIs&#8221;, (Figure 3Ba). Normalize the fluorescence intensity values in each frame to the initial intensity value (F0) (F/F0) (Figure 3Bb). Average the values.</li>
		<li>Highlight the average data and create a line plot using the chart menu options.</li>
		<li>From the data analysis menu, select &#8220;trendline&#8221; to open the plot analysis dialog. Select the type of regression. Set &#8220;exponential&#8221; regression. Then select &#8220;display equation on chart&#8221;. In the graph window, the exponential equation appears and the parameter values are automatically assigned, (Figure 3Bc).</li>
		<li>Apply the exponential correction to the intensity values in each frame as follow: 
		Fn (corrected) = Fn / exp(-n * a) 
		Fn = experimental fluorescence intensity measured at frame n; n = number of frames; a = bleaching factor (constant expressing&#160;the rate of intensity loss due to photobleaching;&#160;Figure 3Bd).</li>
		<li>To set the threshold, open a normalized and corrected &#8220;background ROI&#8221;, calculate the average fluorescence signal and its standard deviation (SD). The average value plus 3 SD represents the threshold (Figure 3Be). Use this threshold for data analysis.</li>
	</ol>
	</li>
	<li>Selection of fusion events using a semiautomatic procedure
	<ol>
		<li>Open the time-sequential images with image analysis software. Apply a Gaussian filter to the active image sequence.</li>
		<li>Analyze images using the tool &#8220;count objects&#8221; or a macro which allows the selection of an object whose pixels have average fluorescence intensity within a defined range. Set the intensity range manually, using the threshold function (go to the bar menu, set measure &#8594; threshold to highlight the area of interest). An adequate threshold is 30% over the local fluorescent background signal.</li>
		<li>Apply a macro &#8220;Filters objects&#8221; to select only objects meeting the following criteria:
		<ol>
			<li>Apply ranges option (min and max inclusive) for aspect. Aspect reports the ratio between the major axis and the minor axis of the ellipse equivalent to the object. Aspect is always &#8805;1. Adequate values are min = 1, max = 3.</li>
			<li>Apply ranges for diameter. Diameter reports the average length of the diameters measured at two degree intervals joining two outline points and passing through the centroid of the object. Set the range in pixels (or in &#181;m, if using a calibrated system).</li>
			<li>Define the optimal range in preliminary experiments: select manually the spots of interest and then measure their diameter using the plot profile function.</li>
		</ol>
		</li>
		<li>Select &#8220;display objects&#8221;: selected objects will appear superimposed to the TIRFM image (Figure 4B).</li>
		<li>Include in the analysis only those spots that show a short (one to three frames) transient increase in fluorescence intensity, immediately followed by a marked loss of signal (transient spots). Employ the circular selection to create a ROI approximately one-spot diameter radially around the selected vesicle/spots (experimental ROIs). Perform this step manually.</li>
		<li>With the ROIs selected, calculate the average fluorescence intensity of each ROI over the course of the movie.</li>
	</ol>
	</li>
	<li>Data analysis (Figure 3C-D)
	<ol>
		<li>Export the time-course of the fluorescence changes measured in each &#8220;experimental ROI&#8221; to a spreadsheet; (Figure 3Da). Normalize the intensity value in each frame to the initial fluorescence intensity (F/F0), (Figure 3Db).</li>
		<li>Apply the exponential correction to the intensity values in each frame as reported in step 3.2.4 (Figure 3Dc).</li>
		<li>To calculate the total number of fusion events (peak number), the time each fusion occurs (peak width) and the amplitude of fluorescent peak (peak height and AUC) apply logical functions using spreadsheet or math packages. An example of fusion event analysis using logical formulas is reported in Figure 3Dd and 3De.</li>
		<li>Assume the increase of fluorescence intensity exceeding the threshold (average background fluorescence intensity &#177; 3SD) as vesicle fusion to the plasma membrane and the resulting peak as a fusion event.</li>
		<li>Calculate the peak width as difference between the last and the first x value of each peak. Multiply this value for 1 / (sampling frequency). Consider this value as the time of vesicle fusion and adhesion at the plasma membrane before vesicle re-acidification and recycling (Figure 3Dd).</li>
		<li>Calculate the whole-cell AUC as a sum of values over threshold. Consider this value as net fluorescent change during the recording time due to the spontaneous (resting) or evoked (stimulated) synaptic activity.</li>
		<li>Calculate the peak height as the difference between the maximal y value of each peak and the threshold. Consider this value as indicative of the fusion type (single vs. simultaneous/sequential fusion or transient vs. full fusion).</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

This paper presents a protocol to image and analyze vesicles dynamics in secreting cells, using fluorescent cDNA-encoded vectors and TIRFM. Key elements of successful imaging by TIRFM are the selection of the cellular model and cell transfection with genetically-encoded optical indicators of vesicle release and recycling.
TIRFM is ideally suited for cells growing adherent to a glass cover and sufficiently flat to allow stable visualization of membranes and fusion events. Vesicles should ideall...

Chemical synaptic transmission between neurons is a major mechanism of communication in the nervous system. It relies on the release of neurotransmitters through a dynamic cycle of vesicle fusion and retrieval at the presynaptic site. Many of the proteins involved in vesicle dynamics have been identified; however, their specific contribution to the phenomenon remains to be clarified1.
Our understanding is partly limited by the fact that the most widely used assays for exo/endocytosis are not always the most appropriate. Several studies related to vesicle fusion and dynamics rely on electrophysiological techniques. This technique provides an optimal temporal resolution and is excellent for investigating the initial fusion of vesicles to the plasma membrane but is unable to detect many of the underlying molecular events that support presynaptic function. Electron microscopy, on the other side, provides the finest morphological description of each singular step, but the dynamic aspect of the event cannot be captured, because samples must be fixed in order to be analyzed.
The advent of new optical recording techniques2,3, in combination with advances in fluorescent molecular probes development4-6, enables the visualization of exocytic processes in live cells, thus providing new levels of information about the synaptic structure and function.
Initial studies exploited activity-dependent styryl dyes (FM1&#8211;43 and related organic dyes)7,8. State-of-the-art imaging techniques employ pH-sensitive variants of the Green Fluorescent Protein (GFP) (pHluorin) tethered to luminal vesicles proteins9. These probes are normally switched off when present in the vesicles because of the low luminal pH. After fusion with the plasma membrane, the vesicle interior is exposed to the neutral extracellular space, the pH abruptly increases, relieves the proton-dependent quenching of pHluorin and the fluorescent signal rapidly appears. As the change in pHluorin is faster than the fusion event, by monitoring fluorescence increases, vesicle fusion with the membrane can be measured and analyzed. Because surface pHluorin-tagged molecules are endocytosed, the fluorescence signal subsequently returns to basal level, therefore the same construct may be used also to monitor vesicle recycling9.
While the vesicle-tagged pH-sensor ensures the visualization only of those vesicles that really fuse with the plasma membrane, imaging at high spatial and temporal resolution is required to describe in details the steps involved in the exo/endocytic&#160;processes. The optical technique that provides the necessary spatio-temporal resolution is total internal reflection fluorescence microscopy (TIRFM), an application of fluorescence microscopy10.
Total internal reflection occurs at the interface between the glass cover-slip and the sample. When the light path reaches the glass cover-slip with an incident angle larger than the critical angle, the excitation light is not transmitted into the sample but is completely reflected back. Under these conditions, an evanescent light wave forms at the interface and propagates in the medium with less optical density (the sample). As the intensity of the evanescent field decays exponentially with distance from the interface (with a penetration depth of about 100 nm) only the fluorophores in closest proximity to the cover-slip can be excited while those further away from the boundary are not. In cells transfected with GFP-constructs, this depth corresponds to proteins expressed on the plasma membrane or in vesicular structures approaching it. As fluorophores in the cell interior cannot be excited, the background fluorescence is minimized, and an image with a very high signal/background ratio is formed11.
Several characteristics make TIRFM the technique of choice for monitoring vesicles dynamics. The perfect contrast and the high signal-to-noise-ratio allow the detection of very low signals deriving from single vesicles. Chip-based image acquisition in each frame provides the temporal resolution necessary to detect highly dynamic processes. Finally, the minimal exposure of cells to light at any other plane in the sample strongly reduces phototoxicity and enables long lasting time-lapse recording12.
Data analysis remains the most challenging and crucial aspect of this technique. The simplest way to monitor vesicle fusion is to measure the accumulation of reporter fluorescent proteins at the cell surface, over time13. As fusion increases, net fluorescence signal increases as well. However, this method may underestimate the process, particularly in large cells and in resting conditions, because endocytosis and photobleaching processes offset the increase in fluorescence intensity due to vesicle exocytosis. An alternative method is to follow each single fusion event14. This latter method is very sensitive and can reveal important details about the fusion mechanisms. However, it requires the manual selection of single events, because completely automated procedures to follow vesicles and to register the fluctuation of their fluorescent signals are not always available. Observation of vesicle dynamics requires sampling cells at high frequency. This generates a large amount of data that can hardly be analyzed manually.
The proposal of this paper is to optimize the TIRFM imaging technique for monitoring the basal and stimulated neurotransmitter release in the SH-5YSY neuroblastoma cell line, and to describe, step-by-step, a procedure developed in the laboratory to analyze data, both at whole-cell and single-vesicle levels.

<table><tbody><tr><td colspan="4">Equipment</td></tr><tr><td>Axio Observer Z1</td><td><a href="http://www.zeiss.com/microscopy/en_de/products/light-microscopes/axio-observer-for-biology.html#introduction" target="_blank"><img src="/img/new_window.png" /> Zeiss</a></td><td>491912-9850-000</td><td>inverted microscope</td></tr><tr><td>Multiline Argon Laser Lasos 77</td><td><a href="http://www.lasos.com/products/argon-ion-laser" target="_blank"><img src="/img/new_window.png" /> Lasos</a></td><td>00000-1312-752</td><td>multi-line (458/488/514 nm), 100&amp;nbsp;mW argon-ion laser</td></tr><tr><td>Laser TIRF slider</td><td><a href="http://www.zeiss.com/microscopy/en_de/products/imaging-systems/single-molecular-imaging-laser-tirf-3.html" target="_blank"><img src="/img/new_window.png" /> Zeiss</a></td><td>423681-9901-000</td><td></td></tr><tr><td>100X Objective</td><td><a href="https://www.micro-shop.zeiss.com/?l=en&amp;amp;p=us&amp;amp;f=o&amp;amp;a=v&amp;amp;m=a&amp;amp;id=421190-9900-000&amp;amp;ss=1" target="_blank"><img src="/img/new_window.png" /> Zeiss</a></td><td>421190-9900-000</td><td>Oil, NA 1.45 Alpha-Plan</td></tr><tr><td>CCD Camera RetigaSRV Fast 1394</td><td><a href="http://www.qimaging.com/products/datasheets/Retiga-SRV.pdf" target="_blank"><img src="/img/new_window.png" /> QImaging</a></td><td></td><td></td></tr><tr><td>LAMBDA 10-3 optical filter changer with SmartShutter</td><td><a href="http://www.sutter.com/IMAGING/lambda103.html" target="_blank"><img src="/img/new_window.png" /> Sutter Instrument Company</a></td><td></td><td></td></tr><tr><td colspan="4">Software</td></tr><tr><td>Image ProPlus 6.3 Software</td><td><a href="http://www.mediacy.com/index.aspx?page=IPP" target="_blank"><img src="/img/new_window.png" /> Media Cybernetics</a></td><td></td><td>spot selection, ROI selection, fluorescence intensity determination</td></tr><tr><td>Excel</td><td><a href="http://office.microsoft.com/it-it/excel/" target="_blank"><img src="/img/new_window.png" /> Microsoft</a></td><td></td><td>photobleaching correction, whole-cell and single-vesicle analyses</td></tr><tr><td>GraphPad Prism 4.00</td><td><a href="http://www.graphpad.com/scientific-software/prism/" target="_blank"><img src="/img/new_window.png" /> GraphPad Software, Inc.</a></td><td></td><td>statistical analysis</td></tr></tbody></table>

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.

The TIRF imaging and data analysis procedures described are designed to study vesicles dynamics in cellular systems. This technique can be used to determine the effects of signaling molecules and drugs on fusion events and neurotransmitter vesicle dynamics17. Using GFP-tagged plasma membrane proteins, the TIRFM analysis has been employed to characterize the constitutive trafficking of GFP-tagged glutamate transporters in glial and epithelial cells18,19.
To validate the im...

Watch this Scientific Journal Video about TIRFM and pH-sensitive GFP-probes to Evaluate Neurotransmitter Vesicle Dynamics in SH-SY5Y Neuroblastoma Cells: Cell Imaging and Data Analysis at JoVE.com

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

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 ...

tirfm-ph-sensitive-gfp-probes-to-evaluate-neurotransmitter-vesicle

Research

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Neuroscience

10.8K Views.  Università degli Studi di Milano. 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.

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

Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an experimental approach with excellent spatial and temporal resolutions. Moreover, such an approach must also have the ability to distinguish receptors localized on the PM from those in intracellular compartments. Most importantly, detecting receptors in a single vesicle requires outstanding detection sensitivity, since each vesicle carries only a small number of receptors. Standard approaches for examining receptor trafficking include surface biotinylation followed by biochemical detection, which lacks both the necessary spatial and temporal resolutions; and fluorescence microscopy examination of immunolabeled surface receptors, which requires chemical fixation of cells and therefore lacks sufficient temporal resolution1-6 . To overcome these limitations, we and others have developed and employed a new strategy that enables visualization of the dynamic insertion of receptors into the PM with excellent spatial and temporal resolutions 7-17 . The approach includes tagging of a pH-sensitive GFP, the superecliptic pHluorin 18, to the N-terminal extracellular domain of the receptors. Superecliptic pHluorin has the unique property of being fluorescent at neutral pH and non-fluorescent at acidic pH (pH &lt; 6.0). Therefore, the tagged receptors are non-fluorescent when within the acidic lumen of intracellular trafficking vesicles or endosomal compartments, and they become readily visualized only when exposed to the extracellular neutral pH environment, on the outer surface of the PM. Our strategy consequently allows us to distinguish PM surface receptors from those within intracellular trafficking vesicles. To attain sufficient spatial and temporal resolutions, as well as the sensitivity required to study dynamic trafficking of receptors, we employed total internal reflection fluorescent microscopy (TIRFM), which enabled us to achieve the optimal spatial resolution of optical imaging (~170 nm), the temporal resolution of video-rate microscopy (30 frames/sec), and the sensitivity to detect fluorescence of a single GFP molecule. By imaging pHluorin-tagged receptors under TIRFM, we were able to directly visualize individual receptor insertion events into the PM in cultured neurons. This imaging approach can potentially be applied to any membrane protein with an extracellular domain that could be labeled with superecliptic pHluorin, and will allow dissection of the key detailed mechanisms governing insertion of different membrane proteins (receptors, ion channels, transporters, etc.) to the PM.

By tagging the extracellular domains of membrane receptors with superecliptic pHluorin, and by imaging these fusion receptors in cultured mouse neurons, we can directly visualize individual vesicular insertion events of the receptors to the plasma membrane. This technique will be instrumental in elucidating the molecular mechanisms governing receptor insertion to the plasma membrane.

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an ...

Use of pHluorin to Assess the Dynamics of Axon Guidance Receptors in Cell Culture and in the Chick Embryo

During development, axon guidance receptors play a crucial role in regulating axons sensitivity to both attractive and repulsive cues. Indeed, activation of the guidance receptors is the first step of the signaling mechanisms allowing axon tips, the growth cones, to respond to the ligands. As such, the modulation of their availability at the cell surface is one of the mechanisms that participate in setting the growth cone sensitivity. We describe here a method to precisely visualize the spatio-temporal cell surface dynamics of an axon guidance receptor both in vitro and in vivo in the developing chick spinal cord. We took advantage of the pH-dependent fluorescence property of a green fluorescent protein (GFP) variant to specifically detect the fraction of the axon guidance receptor that is addressed to the plasma membrane. We first describe the in vitro validation of such pH-dependent constructs and we further detail their use in vivo, in the chick spinal chord, to assess the spatio-temporal dynamics of the axon guidance receptor of interest.

We describe here the use of a pH-sensitive green fluorescent protein variant, pHluorin, to study the spatio-temporal dynamics of axon guidance receptors trafficking at the cell surface.&#160;The pHluorin-tagged receptor is expressed both in cell culture and in vivo, using electroporation of the chick embryo.

During development, axon guidance receptors play a crucial role in regulating axons sensitivity to both attractive and repulsive cues. Indeed, activation ...

Nucleofection of Rodent Neuroblasts to Study Neuroblast Migration In vitro

The subventricular zone (SVZ) located in the lateral wall of the lateral ventricles plays a fundamental role in adult neurogenesis. In this restricted area of the brain, neural stem cells proliferate and constantly generate neuroblasts that migrate tangentially in chains along the rostral migratory stream (RMS) to reach the olfactory bulb (OB). Once in the OB, neuroblasts switch to radial migration and then differentiate into mature neurons able to incorporate into the preexisting neuronal network. Proper neuroblast migration is a fundamental step in neurogenesis, ensuring the correct functional maturation of newborn neurons. Given the ability of SVZ-derived neuroblasts to target injured areas in the brain, investigating the intracellular mechanisms underlying their motility will not only enhance the understanding of neurogenesis but may also promote the development of neuroregenerative strategies.
This manuscript describes a detailed protocol for the transfection of primary rodent RMS postnatal neuroblasts and the analysis of their motility using a 3D in vitro migration assay recapitulating their mode of migration observed in vivo. Both rat and mouse neuroblasts can be quickly and efficiently transfected via nucleofection with either plasmid DNA, small hairpin (sh)RNA or short interfering (si)RNA oligos targeting genes of interest. To analyze migration, nucleofected cells are reaggregated in &#39;hanging drops&#39; and subsequently embedded in a three-dimensional matrix. Nucleofection per se does not significantly impair the migration of neuroblasts. Pharmacological treatment of nucleofected and reaggregated neuroblasts can also be performed to study the role of signaling pathways involved in neuroblast migration.

Nucleofection of Rodent Neuroblasts to Study Neuroblast Migration In vitro

Neuroblast migration is a crucial step in postnatal neurogenesis. The protocol described here can be used to investigate the role of candidate regulators of neuroblast migration by employing DNA/small hairpin RNA (shRNA) nucleofection and a 3D migration assay with neuroblasts isolated from the rodent postnatal rostral migratory stream.

The subventricular zone (SVZ) located in the lateral wall of the lateral ventricles plays a fundamental role in adult neurogenesis. In this restricted ...

Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze membrane fusion within eukaryotic cells. The SNARE protein family consists of about 36 different members. Specific intracellular transport routes are catalyzed by specific sets of 3 or 4 SNARE proteins that thereby contribute to the specificity and fidelity of membrane trafficking. However, studying the precise function of SNARE proteins is technically challenging, because SNAREs are highly abundant and functionally redundant, with most SNAREs having multiple and overlapping functions. In this protocol, a new method for the visualization of SNARE complex formation in live cells is described. This method is based on expressing SNARE proteins C-terminally fused to fluorescent proteins and measuring their interaction by F&#246;rster resonance energy transfer (FRET) employing fluorescence lifetime imaging microscopy (FLIM). By fitting the fluorescence lifetime histograms with a multicomponent decay model, FRET-FLIM allows (semi-)quantitative estimation of the fraction of the SNARE complex formation at different vesicles. This protocol has been successfully applied to visualize SNARE complex formation at the plasma membrane and at endosomal compartments in mammalian cell lines and primary immune cells, and can be readily extended to study SNARE functions at other organelles in animal, plant, and fungal cells.

This protocol describes a new method allowing for the quantitative visualization of complex formation of SNARE proteins, based on F&#246;rster resonance energy transfer, and fluorescence lifetime imaging microscopy.

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze ...

Immunology and Infection

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

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

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

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

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

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

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

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

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

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

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

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

Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe dementia (FTLD) is dysfunction of communication between neurons and motor neurons and muscle. The underlying process of synaptic transmission involves membrane depolarization-dependent synaptic vesicle fusion and the release of neurotransmitters into the synapse. This process occurs through localized calcium influx into the presynaptic terminals where synaptic vesicles reside. Here, the protocol describes fluorescence-based live-imaging methodologies that reliably report depolarization-mediated synaptic vesicle exocytosis and presynaptic terminal calcium influx dynamics in cultured neurons.
Using a styryl dye that is incorporated into synaptic vesicle membranes, the synaptic vesicle release is elucidated. On the other hand, to study calcium entry, Gcamp6m is used, a genetically encoded fluorescent reporter. We employ high potassium chloride-mediated depolarization to mimic neuronal activity. To quantify synaptic vesicle exocytosis unambiguously, we measure the loss of normalized styryl dye fluorescence as a function of time. Under similar stimulation conditions, in the case of calcium influx, Gcamp6m fluorescence increases. Normalization and quantification of this fluorescence change are performed in a similar manner to the styryl dye protocol. These methods can be multiplexed with transfection-based overexpression of fluorescently tagged mutant proteins. These protocols have been extensively used to study synaptic dysfunction in models of FUS-ALS and C9ORF72-ALS, utilizing primary rodent cortical and motor neurons. These protocols easily allow for rapid screening of compounds that may improve neuronal communication. As such, these methods are valuable not only for the study of ALS but for all areas of neurodegenerative and developmental neuroscience research.

Two related methods are described to visualize subcellular events required for synaptic transmission. These protocols enable the real-time monitoring of the dynamics of presynaptic calcium influx and synaptic vesicle membrane fusion using live-cell imaging of in vitro cultured neurons.

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe ...

Utilizing pHluorin-tagged Receptors to Monitor Subcellular Localization and Trafficking

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular organelles and those localized on the plasma membrane. Traditional fluorescence-based measurements lack the capability to distinguish membrane proteins residing in different organelles. Cutting edge methodologies transcend traditional methods by coupling pH-sensitive fluorophores with total internal reflection fluorescence microscopy (TIRFM). TIRF illumination excites the sample up to approximately 150 nm from the glass-sample interface, thus decreasing background, increasing the signal to noise ratio, and enhancing resolution. The excitation volume in TIRFM encompasses the plasma membrane and nearby organelles such as the peripheral ER. Superecliptic pHluorin (SEP) is a pH sensitive version of GFP. Genetically encoding SEP into the extracellular domain of a membrane protein of interest positions the fluorophore on the luminal side of the ER and in the extracellular region of the cell. SEP is fluorescent when the pH is greater than 6, but remains in an off state at lower pH values. Therefore, receptors tagged with SEP fluoresce when residing in the endoplasmic reticulum (ER) or upon insertion in the plasma membrane (PM) but not when confined to a trafficking vesicle or other organelles such as the Golgi. The extracellular pH can be adjusted to dictate the fluorescence of receptors on the plasma membrane. The difference in fluorescence between TIRF images at neutral and acidic extracellular pH for the same cell corresponds to a relative number of receptors on the plasma membrane. This allows a simultaneous measurement of intracellular and plasma membrane resident receptors. Single vesicle insertion events can also be measured when the extracellular pH is neutral, corresponding to a low pH trafficking vesicle fusing with the plasma membrane and transitioning into a fluorescent state. This versatile technique can be exploited to study localization, expression, and trafficking of membrane proteins.

Labeling the extracellular domain of a membrane protein with a pH sensitive fluorophore, superecliptic pHluorin (SEP), allows subcellular localization, expression, and trafficking to be determined. Imaging SEP-labeled proteins with total internal reflection fluorescence microscopy (TIRFM) enables the quantification of protein levels in the peripheral ER and plasma membrane.

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular ...

Biology

Live Imaging of Synaptic Vesicle Recycling in the Neuromuscular Junction of Dissected Larval Zebrafish

Neuronal communication is mediated by synaptic transmission, which depends primarily on the release of neurotransmitters stored in synaptic vesicles (SVs) in response to an action potential (AP). Since SVs are recycled locally at the presynaptic terminal, coordination of SV exocytosis and endocytosis is important for sustained synaptic transmission. A pH-sensitive green fluorescent protein, called pHluorin, provides a powerful tool to monitor SV exo/endocytosis by targeting it to the SV lumen. However, tracking AP-driven SV recycling with the pHluorin-based probes is still largely limited to in vitro culture preparations because the introduction of genetically encoded probes and subsequent optical imaging is technically challenging in general for in vivo animal models or tissue preparations. Zebrafish is a model system offering valuable features, including ease of genetic manipulation, optical clarity, and rapid external development. We recently generated a transgenic zebrafish that highly expresses a pHluorin-labeled probe at motor neuron terminals and developed a protocol to monitor AP-driven SV exo/endocytosis at the neuromuscular junction (NMJ), a well-established synapse model that forms in vivo. In this article, we show how to prepare larval zebrafish NMJ preparation suitable for pHluorin imaging. We also show that the preparation allows time-lapse imaging under conventional upright epifluorescence microscope, providing a cost-effective platform for analyzing NMJ function.

The zebrafish larval neuromuscular junction is an attractive model for studying synaptic physiology. It is amenable to many experimental techniques, including electrophysiology and optical imaging. Here, we describe a protocol for imaging synaptic transmission using a pHluorin-based probe under an upright epifluorescence microscope.

Neuronal communication is mediated by synaptic transmission, which depends primarily on the release of neurotransmitters stored in synaptic vesicles (SVs) ...

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

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