We thank Dr. Ricardo Dolmetsch (Stanford University School of Medicine; present affiliation: Novartis Institutes for Biomedical Research) for helping to develop this analysis and Dr. Matthew Wood, Takuma Aihara, and Dongsook Kim for their critical reading of the manuscript.

All animal procedures were performed with the approval of the University of Tsukuba Animal Care and Use Committee (IACUC).
1. Dissection
<ol>
	<li>To prepare embryonic day 13-14 ICR or C57BL/6 fetuses, euthanize pregnant mice by cervical dislocation. Remove the uterus and put it into a petri dish with Hank&#39;s Balanced Salt Solution (HBSS). Rinse it well to remove the blood.</li>
	<li>Using forceps, transfer the uterus to a new petri dish with 70% ethanol to sterilize.</li>
	<li>Transfer the uterus to a new petri dish with HBSS. Remove the fetuses from the uterus using sterilized dissecting forceps and surgical scissors.</li>
	<li>Decapitate the fetuses using surgical scissors. Place them into a new petri dish with HBSS.</li>
	<li>Remove the skin and skull and take out the brains using sterilized dissecting forceps. Put them into a petri dish with HBSS. 
	NOTE: From this step on, it is best to conduct the procedure on a clean bench.</li>
	<li>Remove the meninges using sterilized dissecting forceps under a stereo microscope. Cut off the basal ganglia, hippocampus, and cerebellum using forceps. Using curved-tip forceps, transfer the cortices to a new petri dish with HBSS.</li>
	<li>Collect all cortices using a disposable dropper or a 25 mL pipette. Put them into a 15 mL tube and then remove the HBSS by pipetting.</li>
	<li>Add 3 mL of cerebral cortical enzyme solution to the tube containing the cortices. Put this tube into a water bath and incubate at 37 &#176;C for 5 min. 
	NOTE: Prepare a cerebral cortical culture enzyme consisting of Phosphate-Buffered Saline (PBS) with 0.25% trypsin and 1 mM EDTA. Sterilize with a 0.22 &#181;m filter.</li>
	<li>Add a further 3 mL of cerebral cortical enzyme solution into the tube containing the cortices and incubate at 37 &#176;C in a water bath for an additional 5 min.</li>
	<li>Remove the cerebral cortical enzyme solution using a 5 mL pipette and add 5 mL of Dulbecco&#39;s Modified Eagle Medium (DMEM) containing 10% Fetal Bovine Serum (FBS).</li>
	<li>Dissociate the cortices by gently pipetting up and down using a 5 mL pipette with a sterile P1000 tip. Pipette again using a 5 mL pipette with a P200 tip.</li>
	<li>Put a 40 &#181;m nylon cell strainer on a 50 mL tube and filter the suspension to remove the debris.</li>
	<li>Centrifuge at 420 x g for 5 min at RT and then remove the DMEM medium.</li>
	<li>Add 5 mL of cerebral cortical culture medium and gently resuspend the cell pellet. 
	NOTE: Prepare a cerebral cortical culture medium consisting of neuron basal medium supplemented with 1x B-27 supplements, 2 mM L-glutamine, and 100 U/mL penicillin streptomycin.</li>
	<li>Count the cells using a hemocytometer and adjust the cell concentration (as desired) using the cerebral cortical culture medium.</li>
	<li>Plate 3.0 &#215; 106 cortical neurons in 2.0 mL per coated 35 mm glass-bottomed dishes. 
	NOTE: Before plating, coat the 35mmm glass-bottomed dishes with 0.1 mg/mL Poly-D-Lysine hydrobromide (PDL) or 0.01% Poly-L-Ornithine (PLO). Incubate for 3 h &#8211; O/N at 37 &#176;C and wash with PBS at least 3x.</li>
</ol>
2. Imaging Vesicle Motility
<ol>
	<li>Prepare the plasmids that will label each type of vesicle. For instance, use EGFP-Rab5 for early endosomes, EGFP-Rab7 for late endosomes, LAMP-EGFP for lysosomes, and EGFP-LC3 for autophagosomes8,13. 
	NOTE: These plasmids are available on request from tsuruta.fuminori.fn@u.tsukuba.ac.jp. Typically,3 -5 days in vitro&#160;(DIV) neurons can be transfected with these plasmids using transfection reagents.</li>
	<li>Mix 4.0 &#181;g of plasmids with 200 &#181;L of serum-free medium by pipetting. In a separate tube, dilute 8 &#181;L of transfection reagent (see Table of Materials) in 200 &#181;L of serum-free medium. Incubate for 5 min at RT.</li>
	<li>Mix the plasmid solution and transfection regent solution in step 2.2 by pipetting. Incubate for 20 min at RT.</li>
	<li>Add serum-free medium and 400 &#181;L of the plasmid solution from step 2.3 to each coated glass-bottom dish. Incubate the neurons at 37 &#176;C in a 5% CO2 incubator for 30 min. Replace the medium with 2 mL of cerebral cortical culture medium. Incubate the neurons at 37 &#176;C in a 5% CO2 incubator for 1-2 days.</li>
	<li>After 1 - 2 d of transfection, select the transfected cells for image acquisition. 
	NOTE: Premature neurons less than 7 DIV have a tendency to exhibit dynamic vesicle motility. Select neurons that moderately express the fluorescence probe because clear images cannot be obtained when a highly expressing neuron is selected. Also, select a typical neuronal shape, such as pyramidal neurons, which have long axons and intricate dendrites, for ease of identification of the axon and dendrites (see Figure 1A).</li>
	<li>Image acquisition using a time-lapse imaging system: 
	For imaging, use a fluorescence microscope equipped with a Charge-Coupled Device (CCD) camera with 40X Plan Apo 0.95NA or 100X Plan Apo VC NA1.4 objective lenses. Control the temperature at 37 &#176;C using an incubation system. Acquire neuron images at one frame/5 s over a 100 s period controlled by the image acquisition software. Alternatively, acquire images at shorter intervals and for longer time periods, such as at one frame per 2 s over a 300-s period. 
	NOTE: A variety of applications are available for taking sequential images, many of which would be suitable for this method. Therefore, no particular application is recommended; rather, each researcher should use whatever application is available.</li>
</ol>
3. Image Analysis
<ol>
	<li>Open all sequential images in ImageJ software with Manual Tracking14. 
	NOTE: To date, Manual Tracking has been widely used in tracking experiments15,16,17,18,19. Manual Tracking was developed by Dr. Fabrice Cordeli&#232;res. Detailed written instructions are available online20. For the analysis, the quality of the imaging data is improved with a sufficient number of images. It is recommended to use more than 20 images.</li>
	<li>To combine the sequential images, choose &#60;Image&#62; &#8594; &#60;Stacks&#62; &#8594; &#60;Images to Stack&#62;. Click &#60;OK&#62;.</li>
	<li>Choose &#60;Plugins&#62; &#8594; &#60;Manual Tracking&#62;; a tracking window will pop up.</li>
	<li>Click on the checkbox of &#60;Show Parameters?&#62; and define the tracking parameters in the parameters section. 
	NOTE: Several parameters can be set, including time interval, x/y calibration, z calibration, search square size for centering, dot size, line width, and font size. In this simplified assay, both time interval and x/y calibration were defined. Under other experimental circumstances, it may be necessary to define other parameters as well. The parameters used here are as follows: the &#60;Time interval&#62; is 5 s, and the &#60;x/y calibration&#62; is either 0.26642 &#181;m for a 40X Plan Apo 0.95NA objective lens or 0.10657 &#181;m for a 100X Plan Apo VC NA1.4 objective lens.</li>
	<li>After the parameters are defined, click on &#60;add Track&#62; to start a new track.</li>
	<li>After determining the vesicles of interest (e.g., the vesicles indicated by blue and red arrowheads in Figure 1A (right panel)), click on the center of the signals in the sequential images to record the xy coordinates. The results of the recorded xy coordinates, distance, and velocity show up in a new window. 
	NOTE: In the case of lysosomes, large bright fluorescent signals (e.g., Figure 1A (right panel), orange arrowhead) are often observed in neurons. As these signals occasionally exhibit either accumulated vesicles or pathological varicosities, these signals should be avoided for quantifying motility.</li>
	<li>Repeat step 3.6 to collect the xy coordinates of vesicles from all sequential images; each image automatically advances to the successive image after the reference point is selected.</li>
	<li>Export these data to a new table (e.g., a spreadsheet). Use an appropriate software to create a suitable graph (see Table of Materials). 
	NOTE: To analyze the data, select the &#60;Distance&#62; column, sum up all values of this column, and show a motility distance as a bar graph, such as in Figure 1B. Additionally, repeat this step, calculate the average of the total motility distance, and show as a bar graph, as in Figure 1C. The software to create a bar graph and waveform data are shown in the Table of Materials.</li>
</ol>

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The authors have nothing to disclose.

This protocol introduces the procedure for quantifying vesicle motility. In primary neurons, endosomes and lysosomes tend to show high motility in younger neurons (4-6 DIV). Given that neurons must deliver some components to the leading edges to elongate neural processes, membrane trafficking should occur dynamically during this stage. Thus, it is important to use younger neurons to observe dynamic motility in neuronal dendrites. In addition, larger vesicles do not tend to exhibit high motility, even in younger neurons. ...

Precise control of endosome-lysosome trafficking is indispensable for regulating neuronal function. Notably, the dynamic movements of these vesicles are a key factor underlying the regulation of neuronal morphology, development, and survival. Defects in this system cause severe neuronal disorders1,2. The molecular mechanisms that link vesicle trafficking to neuronal diseases are considered complicated, and several groups have sought to examine this relevance. For instance, it has been reported that perturbed late endosome motility is significantly associated with Niemann-Pick C disease3, an inherited neurodegenerative disorder caused by lysosome defects. Another example is a mutation in a lysosomal Ca2+ channel, trpml1, which impairs lysosomal motility, resulting in lysosomal storage diseases4,5,6. Our group has reported that dysregulation of PtdIns(3,5)P2 turnover suppresses endosome and lysosome motility in neurons, leading to an increase in vulnerability to the stress response7,8. The metabolic regulation of PtdIns(3,5)P2, which mostly localizes on late endosomes and lysosomes, plays an important role in a wide variety of cellular functions, including vesicle trafficking and fusion-fission processes9,10. Since impaired PtdIns(3,5)P2 turnover causes severe neurodegeneration11,12, the aberrant regulation of endosome-lysosome motility could be a key factor for understanding the pathogenesis of neurodegeneration. An investigation of the molecular mechanisms that underlie vesicle motility may thus provide promising clues that can deepen our understanding of several neuronal disorders.
In this paper, we introduce a valuable method to quantify vesicle motility in neurons using a free software package called Manual Tracking. The aim was to develop a fast quantification method to analyze vesicle motility. This quantification is directed through a standard approach of clicking on a reference point in each frame of a time-lapse movie. The use of the Manual Tracking software makes this approach quite simple and of broad utility, unlike other applications. Furthermore, this approach is also applicable to other cells, such as glial cells. Although this method is primitive, it can be applied to various analyses, including of cellular motility and morphological change. For instance, after defining a reference point across a sequence of images, information on the positions of the reference points and the time at each position can be extracted from sequential images using data analysis and image-processing software. Taken together, this method is simple but powerful and contributes to the development of improved efficiency in studies based on membrane trafficking, such as those examining endosome-lysosome function.

<table><tbody><tr><td>Neurobasal medium</td><td>Thermo Fisher</td><td>21103-049</td><td></td></tr><tr><td>Dulbecco&#039;s modified eagle medium (DMEM)</td><td>Wako</td><td>044-29765</td><td></td></tr><tr><td>Opti-MEM</td><td>Thermo Fisher</td><td>31985070</td><td>Serum free medium</td></tr><tr><td>Hank&#039;s balanced salt solution (HBSS)</td><td>Thermo Fisher</td><td>14170112</td><td></td></tr><tr><td>Penicilin Streptmycin</td><td>Thermo Fisher</td><td>15140122</td><td></td></tr><tr><td>L-Glutamine</td><td>Thermo Fisher</td><td>25030081</td><td></td></tr><tr><td>B-27 Supplements</td><td>Thermo Fisher</td><td>17504044</td><td></td></tr><tr><td>2.5% Trypsine</td><td>Thermo Fisher</td><td>15090046</td><td></td></tr><tr><td>Poly-D-Lysine hydrobromide (PDL)</td><td>Sigma</td><td>P7280</td><td></td></tr><tr><td>Poly-L-Ornithine (PLO)</td><td>Sigma</td><td>P4957</td><td></td></tr><tr><td>Nylon cell strainer</td><td>Corning</td><td>431750</td><td></td></tr><tr><td>Lipofectamine 2000</td><td>Thermo Fisher</td><td>11668027</td><td>Transfection reagent</td></tr><tr><td>Polyethyleneimine &quot;Max&quot; (PEI)</td><td>polysciences</td><td>24765</td><td>Transfection reagent. As an alternative to Lipofectamine2000, 0.1mg/mL Polyethyleneimine dissolved in sterilized water is available. But it is low efficiency and high toxicity.</td></tr><tr><td>BIOREVO BZ-9000</td><td>Keyence</td><td>NA</td><td></td></tr><tr><td>Incubation system INUG2-K13</td><td>Tokay Hit</td><td>NA</td><td></td></tr><tr><td>GraphPad Prism version 6.0</td><td>GraphPad Software</td><td>NA</td><td></td></tr><tr><td>Excel version 15</td><td>Microsoft</td><td>NA</td><td></td></tr><tr><td>ImageJ verion 1.47</td><td>NA</td><td>NA</td><td></td></tr></tbody></table>

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.

This assay was designed to measure vesicle dynamics in in vitro culture. This assay was utilized to determine the vesicle motility associated with neuronal morphology and survival. Figure 1A and B display representative data showing lysosome motility in neurons. Cortical neurons were transfected with the lysosome marker, LAMP-EGFP, and observed using a standard fluorescence microscope. It has been previously reported that specific endosome-lysoso...

Watch this Scientific Journal Video about Quantification of Endosome and Lysosome Motilities in Cultured Neurons Using Fluorescent Probes at JoVE.com

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

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, ...

quantification-endosome-lysosome-motilities-cultured-neurons-using

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

8.0K Views.  University of Tsukuba. 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.

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

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

Automated Quantification of Synaptic Fluorescence in C. elegans

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall neural circuit function. Synapse strength critically depends on the abundance of neurotransmitter receptors clustered at synaptic sites on the postsynaptic membrane. Receptor levels are established developmentally, and can be altered by receptor trafficking between surface-localized, subsynaptic, and intracellular pools, representing important mechanisms of synaptic plasticity and neuromodulation. Rigorous methods to quantify synaptically-localized neurotransmitter receptor abundance are essential to study synaptic development and plasticity. Fluorescence microscopy is an optimal approach because it preserves spatial information, distinguishing synaptic from non-synaptic pools, and discriminating among receptor populations localized to different types of synapses. The genetic model organism Caenorhabditis elegans is particularly well suited for these studies due to the small size and relative simplicity of its nervous system, its transparency, and the availability of powerful genetic techniques, allowing examination of native synapses in intact animals.
Here we present a method for quantifying fluorescently-labeled synaptic neurotransmitter receptors in C. elegans. Its key feature is the automated identification and analysis of individual synapses in three dimensions in multi-plane confocal microscope output files, tabulating position, volume, fluorescence intensity, and total fluorescence for each synapse. This approach has two principal advantages over manual analysis of z-plane projections of confocal data. First, because every plane of the confocal data set is included, no data are lost through z-plane projection, typically based on pixel intensity averages or maxima. Second, identification of synapses is automated, but can be inspected by the experimenter as the data analysis proceeds, allowing fast and accurate extraction of data from large numbers of synapses. Hundreds to thousands of synapses per sample can easily be obtained, producing large data sets to maximize statistical power. Considerations for preparing C. elegans for analysis, and performing confocal imaging to minimize variability between animals within treatment groups are also discussed. Although developed to analyze C. elegans postsynaptic receptors, this method is generally useful for any type of synaptically-localized protein, or indeed, any fluorescence signal that is localized to discrete clusters, puncta, or organelles.
The procedure is performed in three steps: 1) preparation of samples, 2) confocal imaging, and 3) image analysis. Steps 1 and 2 are specific to C. elegans, while step 3 is generally applicable to any punctate fluorescence signal in confocal micrographs.

Automated Quantification of Synaptic Fluorescence in C. elegans

The abundance of neurotransmitter receptors clustered at synapses strongly influences synaptic strength. This method quantifies fluorescently-labeled neurotransmitter receptors in three dimensions with single-synapse resolution in C. elegans, allowing hundreds of synapses to be rapidly characterized within a single sample without distortions introduced by z-plane projection.

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall ...

Characterizing the Composition of Molecular Motors on Moving Axonal Cargo Using "Cargo Mapping" Analysis

Understanding the mechanisms by which molecular motors coordinate their activities to transport vesicular cargoes within neurons requires the quantitative analysis of motor/cargo associations at the single vesicle level. The goal of this protocol is to use quantitative fluorescence microscopy to correlate (&#8220;map&#8221;) the position and directionality of movement of live cargo to the composition and relative amounts of motors associated with the same cargo. &#8220;Cargo mapping&#8221; consists of live imaging of fluorescently labeled cargoes moving in axons cultured on microfluidic devices, followed by chemical fixation during recording of live movement, and subsequent immunofluorescence (IF) staining of the exact same axonal regions with antibodies against motors. Colocalization between cargoes and their associated motors is assessed by assigning sub-pixel position coordinates to motor and cargo channels, by fitting Gaussian functions to the diffraction-limited point spread functions representing individual fluorescent point sources. Fixed cargo and motor images are subsequently superimposed to plots of cargo movement, to &#8220;map&#8221; them to their tracked trajectories. The strength of this protocol is the combination of live and IF data to record both the transport of vesicular cargoes in live cells and to determine the motors associated to these exact same vesicles. This technique overcomes previous challenges that use biochemical methods to determine the average motor composition of purified heterogeneous bulk vesicle populations, as these methods do not reveal compositions on single moving cargoes. Furthermore, this protocol can be adapted for the analysis of other transport and/or trafficking pathways in other cell types to correlate the movement of individual intracellular structures with their protein composition. Limitations of this protocol are the relatively low throughput due to low transfection efficiencies of cultured primary neurons and a limited field of view available for high-resolution imaging. Future applications could include methods to increase the number of neurons expressing fluorescently labeled cargoes.

Intracellular transport of cargoes, such as vesicles or organelles, is carried out by molecular motor proteins that track on polarized microtubules. This protocol describes the correlation of the directionality of transport of individual cargo particles moving inside neurons, to the relative amount and type of associated motor proteins.

Understanding the mechanisms by which molecular motors coordinate their activities to transport vesicular cargoes within neurons requires the quantitative ...

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 ...

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 ...

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized microtubules. Organelles move along these processes by microtubule motors. Easy maintenance, high sensitivity to RNAi-mediated protein knock-down and efficient procedure for creating stable cell lines make Drosophila S2 cells an ideal model system to study cargo transport by live imaging. The results obtained with S2 cells can be further applied to a more physiologically relevant system: axonal transport in primary neurons cultured from dissociated Drosophila embryos. Cultured neurons grow long neurites filled with bundled microtubules, very similar to S2 processes. Like in S2 cells, organelles in cultured neurons can be visualized by either organelle-specific fluorescent dyes or by using fluorescent organelle markers encoded by DNA injected into early embryos or expressed in transgenic flies. Therefore, organelle transport can be easily recorded in neurons cultured on glass coverslips using living imaging. Here we describe procedures for culturing and visualizing cargo transport in Drosophila S2 cells and primary neurons. We believe that these protocols make both systems accessible for labs studying cargo transport.

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells and cultured neurons are great systems for imaging of motor-driven organelle transport in vivo. Here we describe detailed protocols for culturing both cell types, their imaging and analysis of transport.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized ...

Biology

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 ...

Measuring Axonal Cargo Transport in Mouse Primary Cortical Cultured Neurons

Neuronal cells are highly polarized cells that stereotypically harbor several dendrites and an axon. The length of an axon necessitates efficient bidirectional transport by motor proteins. Various reports have suggested that defects in axonal transport are associated with neurodegenerative diseases. Also, the mechanism of the coordination of multiple motor proteins has been an attractive topic. Since the axon has uni-directional microtubules, it is easier to determine which motor proteins are involved in the movement. Therefore, understanding the mechanisms underlying the transport of axonal cargo is crucial for uncovering the molecular mechanism of neurodegenerative diseases and the regulation of motor proteins. Here, we introduce the entire process of axonal transport analysis, including the culturing of mouse primary cortical neurons, transfection of plasmids encoding cargo proteins, and directional and velocity analyses without the effect of pauses. Furthermore, the open-access software "KYMOMAKER" is introduced, which enables the generation of a kymograph to highlight transport traces according to their direction and allow easier visualization of axonal transport.

The present protocol describes the whole procedure of analysis for axonal transport. In particular, the calculation of transport velocity, not including the pausing, and the visualization method using open-access software "KYMOMAKER" are shown here.

Neuronal cells are highly polarized cells that stereotypically harbor several dendrites and an axon. The length of an axon necessitates efficient ...

Enhancing Axonal Transport Analysis in Living C. elegans Using GFP-Tagged RAB-3

Optimizing Visualization of Axonal Transport of Endogenous Cargo by Fluorescence Microscopy in Living Caenorhabditis elegans

Axonal transport is a prerequisite to deliver axonal proteins from their site of synthesis in the neuronal cell body to their destination in the axon. Consequently, loss of axonal transport impairs neuronal growth and function. Studying axonal transport therefore improves our understanding of neuronal cell biology. With recent improvements in CRISPR Cas9 genome editing, endogenous labeling of axonal cargos has become accessible, enabling to move beyond ectopic expression-based visualization of transport. However, endogenous labeling often comes at the cost of low signal intensity and necessitates optimization strategies to obtain robust data. Here, we describe a protocol to optimize the visualization of axonal transport by discussing acquisition parameters and a bleaching approach to improve the signal of endogenous labeled cargo over diffuse cytoplasmic background. We apply our protocol to optimize the visualization of synaptic vesicle precursors (SVPs) labeled by green fluorescent protein (GFP)-tagged RAB-3 to highlight how fine-tuning acquisition parameters can improve the analysis of endogenously labeled axonal cargo in Caenorhabditis elegans (C. elegans).

Author Spotlight: Advanced Techniques for Visualizing Endogenous Axonal Transport Dynamics

The paper describes the optimization of fluorescence microscopy acquisition parameters to visualize the axonal transport of endogenous labeled cargos at single-neuron resolution in a living nematode.

Axonal transport is a prerequisite to deliver axonal proteins from their site of synthesis in the neuronal cell body to their destination in the axon. ...

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

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Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Automated Quantification of Synaptic Fluorescence in C. elegans

Characterizing the Composition of Molecular Motors on Moving Axonal Cargo Using "Cargo Mapping" Analysis

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

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

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

Measuring Axonal Cargo Transport in Mouse Primary Cortical Cultured Neurons

Optimizing Visualization of Axonal Transport of Endogenous Cargo by Fluorescence Microscopy in Living Caenorhabditis elegans