This work is supported by the Intramural Research Program of National Institute of Neurological Disorders and Stroke, National Institutes of Health. Project number 1ZIANS003137.

CAUTION: This protocol involves the use of class IV lasers and will require proper training and safety guidelines to be followed. Avoid eye or skin exposure to direct or scattered laser light. 
NOTE: The protocol includes six steps. The workflow is shown in Figure 1A.
1. Labeling Individual Neurons Using the Flip-out Technique
NOTE: The single labeling of LNvs is achieved by expressing mCD8::GFP in single LNvs using flippase -mediated stochastic labeling. The genotype of the fly line is: hs-flp; Pdf-Gal4; UAS-FRT-CD2-stop-FRT-mCD8::GFP11,12,13. The frequency of obtaining a single labeled LNv is around 10%. Fly stocks are maintained in standard medium in circadian- and humidity-controlled 25 &#176;C incubators.
<ol>
	<li>Collect 100-200 eggs within a 2 h window post fertilization on a grape juice plate with yeast supplement. Incubate the embryos at 25 &#176;C for 24 h and collect newly hatched first instar larvae for the next step.</li>
	<li>Heat shock the newly hatched first instar larvae at 37.5 &#176;C for 40 min two times, with a 40 min recovery period in between.</li>
	<li>Culture the larvae at 25 &#176;C with circadian and humidity controls to the desired developmental stage(s).</li>
</ol>
2. Dissecting and Mounting Larval Brain Explants
<ol>
	<li>Dissect larval brains in the physiological external saline solution (120 mM NaCl, 4 mM MgCl2, 3 mM KCl, 10 mM NaHCO3, 10 mM Glucose, 10 mM Sucrose, 5 mM TES, 10 mM HEPES, 2 mM Ca2+, PH 7.2) under a dissection microscope (4.5x magnification power) with two pairs of #5 standard tip dissection forceps (11 cm). Use one pair of forceps to hold the larval body in place and the other to carefully dissect out the brain. Preserve the eye disks, brain lobes and the ventral nerve cord. Remove attached muscles to minimize sample movements during imaging.</li>
	<li>Prepare a glass slide (25 x 75 x 1.0 mm3) and use a syringe to draw a square chamber with vacuum grease.</li>
	<li>Add 20 &#181;L of external saline solution to the square chamber with grease barriers.</li>
	<li>Transfer dissected larval brains into the chamber on the glass slide using forceps. Adjust the position of the brains under the dissection scope to ensure the dorsal side faces up.</li>
	<li>Cover the chamber with a glass cover slip (22 x 22 x 0.15 mm3). The larval brain is now mounted on the slide within a chamber filled with the external saline solution (Figure 1B). 
	NOTE: Pressing gently on the coverslip confines the brains and reduces sample drifting in the subsequent imaging session.</li>
</ol>
3. Time-lapse Live Imaging
NOTE: We perform time-lapse imaging experiments using a confocal microscope equipped with a multiphoton laser. The acquisition parameters need to be adjusted for other imaging setups.
<ol>
	<li>Identify brain explants containing individually labeled neurons using a 40X water immersion objective (NA 1.3) and an epifluorescent light source. For image collection, use a two-photon laser tuned to 920 nm and a non-descanned (NDD) detector.</li>
	<li>Collect images at 512 x 512 pixels per frame and 1 min per Z-stack for 10 min. Adjust the optical and digital zoom to achieve a sufficient x-y-z resolution while ensuring the coverage of the whole dendritic arbor within 1 min (Figure 2, Supplementary Video 1). The settings generate images with a typical x-y-z resolution of 0.11 x 0.11 x 0.25 &#181;m3.</li>
	<li>Collect the time lapse image series within 30 min of the brain dissection. Data with excessive drift or obvious morphological deterioration should be excluded from post- processing and quantification.</li>
</ol>
4. Drift Correction and Deconvolution
NOTE: Depending on the image quality, both steps are optional but strongly recommended.
<ol>
	<li>Open an acquired image file with a drift correction software. Edit microscopic parameters to match those used for the experiment. For the software (see Table of Materials), click Edit | Edit microscopic parameters. Set microscope type as widefield for two-photon images if there is no two-photon option and follow the workflow defined by the software. For example, open the tab Deconvolution | Object Stabilizer, and choose Stabilize time frames.</li>
	<li>Open the drift-corrected image in the deconvolution software and follow its workflow. For the software (see Table of Materials), select the stabilized image and then click Deconvolution | Deconvolution Express. To get a better result, fine-tune the parameters using Deconvolution Wizard.</li>
	<li>Save the deconvolved image in a file type that is supported by the subsequent image annotation software capable of analyzing 4D data and reporting the spatial coordinates of defined spots in the image (see Table of Materials).</li>
</ol>
5. Image Annotation
<ol>
	<li>Open the deconvolved image in the image annotation software. Examine and mark branch tips at all time points in 3D. The image annotation software reports and stores the spatial and temporal coordinates of the marked branch tips. Export the coordinate information as a .csv file for subsequent calculations. 
	NOTE: The following two steps are specific to the annotation software we used (see Table of Materials). The workflow may be different for other software.</li>
	<li>Within the Spots module, click &#34;Skip automatic creation, edit manually&#34; and check the bottom &#34;Auto-connect to selected Spot&#34; checkbox (Figure 3A). Go over the frames in the time series and select a branch for annotation. Hold the Shift key and click the branch terminal tip to add a spot. Click through all time points. 
	NOTE: The image annotation software connects the spots between frames and generates a trajectory automatically. The spatial and temporal information of the branch tips is now associated with marked spots. Repeat these steps until all branch tips are annotated (Figure 3B).</li>
	<li>Within the Spots module, click the Statistics tab, choose Detailed and select Specific Values | Position. The recorded spatial and temporal coordinate information will be on display. Click the Save bottom to export that information as a .csv file (Figure 3C).</li>
</ol>
6. Calculating Dendritic Branch Dynamics
NOTE: Using the coordinate information, displacement of the branch tips in 3D can be easily calculated. In our study, all dendritic branch movements are categorized as Extension or Retraction. The steps below describe how to process the .csv files using custom-written R scripts (Supplementary File) and by adding the directional information through manual editing.
<ol>
	<li>Open the .csv file as a spreadsheet. Select the Track ID column, click the Sort Smallest to Largest option and accept Expand the selection. After sorting, Track ID identifies each unique dendritic branch, and spots from the same branch share the same Track ID. The Time column stores the information of different time frames.</li>
	<li>In the spreadsheet, add a Distance column. Calculate the distance of every two temporally adjacent spots using their coordinates and put the values in the Distance column (Figure 4A).</li>
	<li>Minor movements at the single voxel level. Based on the imaging settings, 0.3 &#181;m are usually artifacts from uncorrected drifting or imperfect image annotation. Filter these movements out by resetting all Distance values smaller than 0.3 &#181;m to 0. Process multiple .csv files manually or use our R script batch column filtering.R (Supplementary File).</li>
	<li>Manually generate a new column named Displacement in the spreadsheet. Copy the values from Distance column to Displacement column. Manually assign the extension and retraction events for each branch tip. If it is an extension, leave the Displacement value unchanged, which is a positive value. If it is a retraction, change the corresponding Displacement value to a negative value (e.g., 0.35 to -0.35) (Figure 4B).</li>
	<li>Generate a new column named Event. In this column, manually sum the Displacement values for individual extension and retraction events (Figure 4B).</li>
	<li>Process the modified spreadsheet and quantify the extension and retraction events based on their displacement values using R script batch column sum.R (Supplementary File). The output parameters include Number of extension events, Number of retraction events, Cumulative length extended, Cumulative length retracted, Net length changed, and Total length traveled.</li>
</ol>

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The authors have nothing to disclose.

Here, we describe a protocol we developed to record and quantify the dynamic behavior of dendritic branches in individually labeled neurons in Drosophila larval brains. Notably, our live imaging protocol contains specific parameters that enable us to capture the whole dendritic arbor of a larval LNv and provide a global view of the dynamic state of its dendrite branches. However, because our quantification methods heavily rely on the annotation of branch terminals, neurons with complex dendritic structures and c...

Dendrites are specialized neuronal compartments that receive and process sensory and synaptic input. The complex and stereotyped structure of dendritic arbors has been under intense investigation since their discovery. A number of model systems, including Xenopus optic tectal neurons, chick retinal ganglion cells, and dendritic arborization (da) neurons in the Drosophila system, have been established to study the development, remodeling and plasticity of neuronal dendrites1,2,3,4. Drosophila ventral lateral neurons (LNvs) are a group of visual projection neurons initially identified for their important functions in circadian regulation of fly behaviors5. Studies also revealed the role of larval LNvs as the direct postsynaptic target of the larval photoreceptors (PRs)6,7. Importantly, culturing developing larvae in different light regimes strongly affects the size of LNvs&#39; dendritic arbors, demonstrating the suitability of LNvs as a new model for studying dendritic plasticity7. Recent work from our group further indicates that both the size of the LNv dendrite and the dynamic behavior of the dendritic branches display experience-dependent plasticity8,9. As part of this work, we developed a new live imaging and quantification protocol to perform analysis on the dendrite dynamics of LNvs from 2nd or 3rd instar larvae.
The transparent nature of the Drosophila larval brain makes it ideal for live imaging. However, the dendritic arbors of LNvs are situated in the densely innervated larval optic neuropil (LON) in the center of the larval brain lobe6. To capture images of fine dendrite branches and filopodia in the intact brain tissue, we utilize two-photon microscopy, which increases the depth of light penetration and reduces the phototoxicity during live imaging experiments10. Using this setup, we successfully performed live imaging experiments on LNvs for over 30 min without observing obvious morphological deterioration of the neuron. In addition, genetic manipulations using the Flip-out technique enabled us to label the individual LNvs with a membrane tagged GFP, which is also critical for monitoring the movements of individual branches11,12,13.
To capture the dynamic behavior of all branches on the LNv dendritic arbor with optimal optic resolution, we performed time-lapse 3D imaging on freshly dissected larval brain explants with a high spatial resolution at 1 min per frame for 10 to 30 min. Developing LNv dendrites are highly dynamic, with a large percentage of the branches displaying observable changes within the 10 min window. This leads to one of the main technical challenges in studying dendrite dynamics, quantifying branch behavior based on the 4D image data sets. Previously established methods have various limitations, including lack of accuracy and excessive time requirement. Therefore, we developed a semi-automatic method that combines image post-processing, manual marking of the branch terminals, and automatic 4D spot tracing using an image annotation software. We calculate the movements of branch terminals at different time points based on the 3D coordinates of the spots. The data are then exported and analyzed to produce quantitative measurements of the branch dynamics. This method accurately assesses the duration and extent of extension and retraction events of existing branches, as well as the formation of new branches, allowing us to monitor dendrite dynamics in a large number of neurons.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chameleon Vision II multiphoton laser</td>
			<td>Coherent</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>high vacuum grease</td>
			<td>Dow Corning</td>
			<td>79751-30</td>
			<td></td>
		</tr>
		<tr>
			<td>LSM 780 two-photon laser scanning confocal microscope</td>
			<td>Carl Zeiss</td>
			<td></td>
			<td>upright configuration</td>
		</tr>
		<tr>
			<td>Microscope Cover Glass</td>
			<td>Fisher Scientific</td>
			<td>12-544-E</td>
			<td></td>
		</tr>
		<tr>
			<td>Superfrost Plus Microscope Slides</td>
			<td>Fisher Scientific</td>
			<td>12-550-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Software</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Excel</td>
			<td>Microsoft</td>
			<td></td>
			<td>for processing .csv files</td>
		</tr>
		<tr>
			<td>Huygens Professional</td>
			<td colspan="2">Scientific Volume Imaging</td>
			<td>for drift correction and deconvolution</td>
		</tr>
		<tr>
			<td>Imaris</td>
			<td>Oxford Instruments</td>
			<td></td>
			<td>for 3D visualization and image annotation</td>
		</tr>
		<tr>
			<td>Reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>TES</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

time-lapse live imaging and quantification of fast dendritic branch dynamics in developing drosophila neurons

Highly motile dendritic filopodia are widely present in neurons at early developmental stages. These exploratory dynamic branches sample the surrounding environment and initiate contacts with potential synaptic partners. Although the connection between dendritic branch dynamics and synaptogenesis is well established, how developmental and activity-dependent processes regulate dendritic branch dynamics is not well understood. This is partly due to the technical difficulties associated with the live imaging and quantitative analyses of these fine structures using an in vivo system. We established a method to study dendrite dynamics using Drosophila larval ventral lateral neurons (LNvs), which can be individually labeled using genetic approaches and are accessible for live imaging. Taking advantage of this system, we developed protocols to capture branch dynamics of the whole dendritic arbor of a single labeled LNv through time-lapse live imaging. We then performed post-processing to improve image quality through drift correction and deconvolution, followed by analyzing branch dynamics at the single-branch level by annotating spatial positions of all branch terminals. Lastly, we developed R scripts (Supplementary File) and specific parameters to quantify branch dynamics using the coordinate information generated by the terminal tracing. Collectively, this protocol allows us to achieve a detailed quantitative description of branch dynamics of the neuronal dendritic arbor with high temporal and spatial resolution. The methods we developed are generally applicable to sparsely labeled neurons in both in vitro and in vivo conditions.

Using the live imaging protocol described above, we capture high resolution image stacks for the subsequent analyses and quantification. Supplementary Video 1 shows the maximum intensity projected (MIP) image series collected from a representative individually labeled LNv. Figure 2B shows the corresponding montage of eight frames of the image series. In both panels, arrowheads mark retraction events and arrows mark extension ...

Watch this Scientific Journal Video about Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons at JoVE.com

Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

Here, we describe the method we employed to image highly motile dendritic filopodia in a live preparation of the Drosophila larval brain, and the protocol we developed to quantify time-lapse 3D imaging datasets for quantitative assessments of dendrite dynamics in developing neurons.

Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

Highly motile dendritic filopodia are widely present in neurons at early developmental stages. These exploratory dynamic branches sample the surrounding ...

time-lapse-live-imaging-quantification-fast-dendritic-branch-dynamics

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6.1K Views.  National Institutes of Health. Here, we describe the method we employed to image highly motile dendritic filopodia in a live preparation of the Drosophila larval brain, and the protocol we developed to quantify time-lapse 3D imaging datasets for quantitative assessments of dendrite dynamics in developing neurons.

Video: Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

Live-imaging of PKC Translocation in Sf9 Cells and in Aplysia Sensory Neurons

Protein kinase Cs (PKCs) are serine threonine kinases that play a central role in regulating a wide variety of cellular processes such as cell growth and learning and memory. There are four known families of PKC isoforms in vertebrates: classical PKCs (&alpha;, &beta;I, &beta;II and &gamma;), novel type I PKCs (&#949; and &eta;), novel type II PKCs (&delta; and &theta;), and atypical PKCs (&zeta; and &iota;). The classical PKCs are activated by Ca2+ and diacylclycerol (DAG), while the novel PKCs are activated by DAG, but are Ca2+-independent. The atypical PKCs are activated by neither Ca2+ nor DAG. In Aplysia californica, our model system to study memory formation, there are three nervous system specific PKC isoforms one from each major class, namely the conventional PKC Apl I, the novel type I PKC Apl II and the atypical PKC Apl III. PKCs are lipid-activated kinases and thus activation of classical and novel PKCs in response to extracellular signals has been frequently correlated with PKC translocation from the cytoplasm to the plasma membrane. Therefore, visualizing PKC translocation in real time in live cells has become an invaluable tool for elucidating the signal transduction pathways that lead to PKC activation. For instance, this technique has allowed for us to establish that different isoforms of PKC translocate under different conditions to mediate distinct types of synaptic plasticity and that serotonin (5HT) activation of PKC Apl II requires production of both DAG and phosphatidic acid (PA) for translocation 1-2. Importantly, the ability to visualize the same neuron repeatedly has allowed us, for example, to measure desensitization of the PKC response in exquisite detail 3. In this video, we demonstrate each step of preparing Sf9 cell cultures, cultures of Aplysia sensory neurons have been described in another video article 4, expressing fluorescently tagged PKCs in Sf9 cells and in Aplysia sensory neurons and live-imaging of PKC translocation in response to different activators using laser-scanning microscopy.

In this video, we demonstrate visualization of PKC translocation in living cells using fluorescently tagged PKCs.

Protein kinase Cs (PKCs) are serine threonine kinases that play a central role in regulating a wide variety of cellular processes such as cell growth and ...

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

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 ...

High-resolution Live Imaging of Cell Behavior in the Developing Neuroepithelium

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central nervous system (CNS). Although much is known about the molecular mechanisms that pattern the spinal cord and elicit neuronal differentiation1, 2, we lack a deep understanding of these early events at the level of cell behavior. It is thus critical to study the behavior of neural progenitors in real time as they undergo neurogenesis.
In the past, real-time imaging of early embryonic tissue has been limited by cell/tissue viability in culture as well as the phototoxic effects of fluorescent imaging. Here we present a novel assay for imaging such tissue for long periods of time, utilizing a novel ex vivo slice culture protocol and wide-field fluorescence microscopy (Fig. 1). This approach achieves long-term time-lapse monitoring of chick embryonic spinal cord progenitor cells with high spatial and temporal resolution.
This assay may be modified to image a range of embryonic tissues3, 4 In addition to the observation of cellular and sub-cellular behaviors, the development of novel and highly sensitive reporters for gene activity (for example, Notch signaling5) makes this assay a powerful tool with which to understand how signaling regulates cell behavior during embryonic development.

Imaging embryonic tissue in real-time is challenging over long periods of time. Here we present an assay for monitoring cellular and sub-cellular changes in chick spinal cord for long periods with high spatial and temporal resolution. This technique can be adapted for other regions of the nervous system and developing embryo.

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central ...

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

Motor Nerve Transection and Time-lapse Imaging of Glial Cell Behaviors in Live Zebrafish

The nervous system is often described as a hard-wired component of the body even though it is a considerably fluid organ system that reacts to external stimuli in a consistent, stereotyped manner, while maintaining incredible flexibility and plasticity. Unlike the central nervous system (CNS), the peripheral nervous system (PNS) is capable of significant repair, but we have only just begun to understand the cellular and molecular mechanisms that govern this phenomenon. Using zebrafish as a model system, we have the unprecedented opportunity to couple regenerative studies with in vivo imaging and genetic manipulation. Peripheral nerves are composed of axons surrounded by layers of glia and connective tissue. Axons are ensheathed by myelinating or non-myelinating Schwann cells, which are in turn wrapped into a fascicle by a cellular sheath called the perineurium. Following an injury, adult peripheral nerves have the remarkable capacity to remove damaged axonal debris and re-innervate targets. To investigate the roles of all peripheral glia in PNS regeneration, we describe here an axon transection assay that uses a commercially available nitrogen-pumped dye laser to axotomize motor nerves in live transgenic zebrafish. We further describe the methods to couple these experiments to time-lapse imaging of injured and control nerves. This experimental paradigm can be used to not only assess the role that glia play in nerve regeneration, but can also be the platform for elucidating the molecular mechanisms that govern nervous system repair.

Although the peripheral nervous system (PNS) is capable of significant repair after injury, little is known about the cellular and molecular mechanisms that govern this phenomenon. Using live, transgenic zebrafish and a reproducible nerve transection assay, we can study dynamic glial cell behaviors during nerve degeneration and regeneration.

The  nervous system is often described as a hard-wired component of the body even  though it is a considerably fluid organ system that reacts to external ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...