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>

<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. The regulation of vesicle size is likely involved in the frequency of fusion-fission steps. It may be possible that specific vesicles that do not move dynamically easily accept other vesicles easily and terminate their movements. Thus, selecting vesicles is an important process for this analysis.
Several free plug-in software packages are available for tracking objects21. For instance, the TrackMate package can track objects22 and can be modified by numerical software, providing a range of mathematical information. On the other hand, it is slightly complicated to use for the relatively simple image analysis in this protocol. Mtrack2 is another plug-in and is based on the MultiTracker plug-in23. Mtrack2 is also useful because it can identify targets and then determine which targets in following frames are the closest. This approach is useful for quantifying enormous volumes of data; however, manual tracking may be more reliable for smaller volumes of image data. Therefore, we recommend Manual Tracking for quantifying smaller sets of data and acknowledge that it may be better to use other software to monitor larger volumes.
Several precautions should be taken in this approach. First, the sampling rate should be high in order to obtain strict data. This is because some vesicles exhibit rapid movement and abrupt changes, such as fusion-fission, and appearance-disappearance events are occasionally observed, indicating that frequent imaging is crucial to avoid miss the sign of any changes. Another point is that the use of high-end microscopes, such as scanning-disk confocal microscopes and total internal reflection fluorescence microscopes, may be key to obtaining high-resolution, interesting data. It is recommended to take images at shorter intervals using a high-end microscope, although here, images were taken every 5 s due to equipment limitations.
Substantial studies have suggested a biological link between membrane trafficking and neuronal disorders2. For example, it has been demonstrated that lysosomal defects are involved in severe neuronal disorders, such as Niemann-Pick disease and Gaucher disease24. Intriguingly, several studies have reported that lysosomal movement and localization are involved in the onset of lysosomal storage disorder6. It is possible that vesicle motility can be used as a biomarker for these disorders. One interesting approach may arise from the availability of human induced Pluripotent Stem Cells (iPSCs). Investigating vesicle motility using iPSCs-derived neurons may reveal a biomarker for lysosomal storage disorders. This approach provides some clues to the processes that underlie the risk of disease.
Since this assay is simple, it is easy to obtain motility data. Also, it is applicable to the analysis of other cellular events, including migration and morphological changes. It is important to note that this approach has several limitations due to its simplicity. As such, it is necessary to employ other approaches, including electrophysiology, biochemistry, and electron microscope analysis, to understand the precise physiological relevance of neuronal trafficking. Nevertheless, this approach is a valuable method that is suitable for researchers who want to analyze trafficking in a straightforward and time-effective manner.

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Neurobasal medium</td>
			<td>Thermo Fisher</td>
			<td>21103-049</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s modified eagle medium&#160;</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&#39;s balanced salt solution</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</td>
			<td>Sigma</td>
			<td>P7280</td>
			<td></td>
		</tr>
		<tr>
			<td>poly-L-ornithine&#160;</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 &#34;Max&#34;</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-lysosome vesicles are highly mobile, even in resting neurons8. Indeed, these results demonstrate both resting and moving vesicles in the same dendrite (Figure 1A and B). Normally, resting vesicles exhibit vibratory movement, like Brownian motion, in neuronal dendrites. In contrast, moving vesicles exhibit irregular motility. We speculate that each vesicle has a different compositional feature that underlies the determination of function, even in lysosome vesicles8. Thus, the classification of vesicles using this approach may provide useful information. Using this system, it is easy to quantify the average motility of these vesicles (Figure 1C) and to analyze the vesicle properties using this system in a time-effective manner.
<img alt="Figure 1" src="/files/ftp_upload/55488/55488fig1.jpg" /> 
Figure 1: Lysosome Motility in Neuronal Dendrites. (A) Mouse cortical neurons (4 DIV) were transfected with LAMP-EGFP plasmid for 2 days. This image shows the motility of LAMP-EGFP-containing vesicles in the dendrites. The inset shows the time-lapse image of the LAMP-EGFP-positive vesicles. The red arrowhead indicates a moving vesicle, and the blue arrowhead indicates a resting vesicle. The orange arrowheads indicate larger fluorescent signals, as referred to above. Scale bar = 20 &#181;m. (B) The time-dependent x-axis position of the LAMP-EGFP-positive vesicles from Figure A. The vesicle in the red circle labeled (b) is moving, and the vesicle in the blue circle labeled (a) is resting. Scale bar: 5 &#181;m (left). The bar graph indicates the total distance of LAMP-EGFP movement (right). (C) The bar graph indicates the average motility distance of LAMP-EGFP-containing vesicles. (n = 5, mean &#177;standard error of the mean [SEM], **p &#60;0.01 by Student&#39;s t-test). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55488/55488fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

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

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

Analysis of Dendritic Spine Morphology in Cultured CNS Neurons

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These structures are rich in actin and have been shown to be highly dynamic. In response to classical Hebbian plasticity 
as well as neuromodulatory signals, dendritic spines can change shape and number, which is thought to be critical for the refinement of neural 
circuits and the processing and storage of information within the brain. Within dendritic spines, a complex network of proteins link extracellular 
signals with the actin cyctoskeleton allowing for control of dendritic spine morphology and number. Neuropathological studies have demonstrated that 
a number of disease states, ranging from schizophrenia to autism spectrum disorders, display abnormal dendritic spine morphology or numbers. 
Moreover, recent genetic studies have identified mutations in numerous genes that encode synaptic proteins, leading to suggestions that these 
proteins may contribute to aberrant spine plasticity that, in part, underlie the pathophysiology of these disorders. In order to study the potential 
role of these proteins in controlling dendritic spine morphologies/number, the use of cultured cortical neurons offers several advantages. Firstly, 
this system allows for high-resolution imaging of dendritic spines in fixed cells as well as time-lapse imaging of live cells. Secondly, this in 
vitro system allows for easy manipulation of protein function by expression of mutant proteins, knockdown by shRNA constructs, or pharmacological 
treatments. These techniques allow researchers to begin to dissect the role of disease-associated proteins and to predict how mutations of these 
proteins may function in vivo.

Numerous recent studies have identified mutations in synaptic proteins associated with brain pathologies. Primary cultured cortical neurons offer great flexibility in examining the effects of these disease-associated proteins on dendritic spine morphology and motility.

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These ...

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 ...

Inducing Dendritic Growth in Cultured Sympathetic Neurons

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs 4-6. Altered patterns of dendritic growth and plasticity are associated with impaired neurobehavioral function in experimental models 7, and are thought to contribute to clinical symptoms observed in both neurodevelopmental disorders 8-10 and neurodegenerative diseases 11-13. Such observations underscore the functional importance of precisely regulating dendritic morphology, and suggest that identifying mechanisms that control dendritic growth will not only advance understanding of how neuronal connectivity is regulated during normal development, but may also provide insight on novel therapeutic strategies for diverse neurological diseases.
Mechanistic studies of dendritic growth would be greatly facilitated by the availability of a model system that allows neurons to be experimentally switched from a state in which they do not extend dendrites to one in which they elaborate a dendritic arbor comparable to that of their in vivo counterparts. Primary cultures of sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rodents provide such a model. When cultured in defined medium in the absence of serum and ganglionic glial cells, sympathetic neurons extend a single process which is axonal, and this unipolar state persists for weeks to months in culture 14,15. However, the addition of either bone morphogenetic protein-7 (BMP-7) 16,17 or Matrigel 18 to the culture medium triggers these neurons to extend multiple processes that meet the morphologic, biochemical and functional criteria for dendrites. Sympathetic neurons dissociated from the SCG of perinatal rodents and grown under defined conditions are a homogenous population of neurons 19 that respond uniformly to the dendrite-promoting activity of Matrigel, BMP-7 and other BMPs of the decapentaplegic (dpp) and 60A subfamilies 17,18,20,21. Importantly, Matrigel- and BMP-induced dendrite formation occurs in the absence of changes in cell survival or axonal growth 17,18.
Here, we describe how to set up dissociated cultures of sympathetic neurons derived from the SCG of perinatal rats so that they are responsive to the selective dendrite-promoting activity of Matrigel or BMPs.

We describe a protocol for using bone morphogenetic protein-7 (BMP-7) or Matrigel to selectively induce dendritic growth in primary sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rats.

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs ...

Large-scale Recording of Neurons by Movable Silicon Probes in Behaving Rodents

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of neurons and circuits requires methods with the spatial selectivity and temporal resolution appropriate for mechanistic analysis of neural ensembles in the behaving animal, i.e. recording of representatively large samples of isolated single neurons. Ensemble monitoring of neuronal activity has progressed remarkably in the past decade in both small and large-brained animals, including human subjects1-11. Multiple-site recording with silicon-based devices are particularly effective because of their scalability, small volume and geometric design.
Here, we describe methods for recording multiple single neurons and local field potential in behaving rodents, using commercially available micro-machined silicon probes with custom-made accessory components. There are two basic options for interfacing silicon probes to preamplifiers: printed circuit boards and flexible cables. Probe supplying companies (<a href="http://www.neuronexustech.com/" >http://www.neuronexustech.com/</a>; <a href="http://www.sbmicrosystems.com/">http://www.sbmicrosystems.com/</a>; <a href="http://www.acreo.se/">http://www.acreo.se/</a>) usually provide the bonding service and deliver probes bonded to printed circuit boards or flexible cables. Here, we describe the implantation of a 4-shank, 32-site probe attached to flexible polyimide cable, and mounted on a movable microdrive. Each step of the probe preparation, microdrive construction and surgery is illustrated so that the end user can easily replicate the process.

We describe methods for large-scale recording of multiple single units and local field potential in behaving rodents with silicon probes. Drive fabrication, probe attachment to the drive and probe implantation processes are illustrated in sufficient details for easy replication.

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of ...

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 ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange (ABE)

Palmitoylation is a post-translational lipid modification involving the attachment of a 16-carbon saturated fatty acid, palmitate, to cysteine residues of substrate proteins through a labile thioester bond [reviewed in1]. Palmitoylation of a substrate protein increases its hydrophobicity, and typically facilitates its trafficking toward cellular membranes. Recent studies have shown palmitoylation to be one of the most common lipid modifications in neurons1, 2, suggesting that palmitate turnover is an important mechanism by which these cells regulate the targeting and trafficking of proteins. The identification and detection of palmitoylated substrates can therefore better our understanding of protein trafficking in neurons.
Detection of protein palmitoylation in the past has been technically hindered due to the lack of a consensus sequence among substrate proteins, and the reliance on metabolic labeling of palmitoyl-proteins with 3H-palmitate, a time-consuming biochemical assay with low sensitivity. Development of the Acyl-Biotin Exchange (ABE) assay enables more rapid and high sensitivity detection of palmitoylated proteins2-4, and is optimal for measuring the dynamic turnover of palmitate on neuronal proteins. The ABE assay is comprised of three biochemical steps (Figure 1): 1) irreversible blockade of unmodified cysteine thiol groups using N-ethylmaliemide (NEM), 2) specific cleavage and unmasking of the palmitoylated cysteine's thiol group by hydroxylamine (HAM), and 3) selective labeling of the palmitoylated cysteine using a thiol-reactive biotinylation reagent, biotin-BMCC. Purification of the thiol-biotinylated proteins following the ABE steps has differed, depending on the overall goal of the experiment.
Here, we describe a method to purify a palmitoylated protein of interest in primary hippocampal neurons by an initial immunoprecipitation (IP) step using an antibody directed against the protein, followed by the ABE assay and western blotting to directly measure palmitoylation levels of that protein, which is termed the IP-ABE assay. Low-density cultures of embryonic rat hippocampal neurons have been widely used to study the localization, function, and trafficking of neuronal proteins, making them ideally suited for studying neuronal protein palmitoylation using the IP-ABE assay. The IP-ABE assay mainly requires standard IP and western blotting reagents, and is only limited by the availability of antibodies against the target substrate. This assay can easily be adapted for the purification and detection of transfected palmitoylated proteins in heterologous cell cultures, primary neuronal cultures derived from various brain tissues of both mouse and rat, and even primary brain tissue itself.

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange ABE

The reversible addition of palmitate to proteins is an important regulator of intracellular protein trafficking. This is of particular interest in neurons where many synaptic proteins are palmitoylated. We utilize a simple biochemical method to detect palmitoylated proteins in cultured neurons, which can be adapted for multiple cell types and tissues.

Palmitoylation is a post-translational lipid  modification involving the attachment of a 16-carbon saturated fatty acid,  palmitate, to cysteine residues ...

Visualization of Endosome Dynamics in Living Nerve Terminals with Four-dimensional Fluorescence Imaging

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis muscle of the garter snake. Raw data comprises time-lapse sequences of 3D z-stacks. Each stack contains 4-20 images acquired with epifluorescence optics at focal planes separated by 400-1,500 nm. Steps in the acquisition of image stacks, such as adjustment of focus, switching of excitation wavelengths, and operation of the digital camera, are automated as much as possible to maximize image rate and minimize tissue damage from light exposure. After acquisition, a set of image stacks is deconvolved to improve spatial resolution, converted to the desired 3D format, and used to create a 4D "movie" that is suitable for variety of computer-based analyses, depending upon the experimental data sought. One application is study of the dynamic behavior of two classes of endosomes found in nerve terminals-macroendosomes (MEs) and acidic endosomes (AEs)-whose sizes (200-800 nm for both types) are at or near the diffraction limit. Access to 3D information at each time point provides several advantages over conventional time-lapse imaging. In particular, size and velocity of movement of structures can be quantified over time without loss of sharp focus. Examples of data from 4D imaging reveal that MEs approach the plasma membrane and disappear, suggesting that they are exocytosed rather than simply moving vertically away from a single plane of focus. Also revealed is putative fusion of MEs and AEs, by visualization of overlap between the two dye-containing structures as viewed in each three orthogonal projections.

Four-dimensional (4D) imaging is utilized to study the behavior and interactions among two types of endosomes in living vertebrate nerve terminals. Movement of these small structures is characterized in three dimensions, permitting confirmation of events such as endosome fusion and exocytosis.

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis ...