This work was supported by the National Institutes of Health (Grant R01NS117701), National Science Foundation (Grant 1146944-IOS), the Indiana Traumatic Spinal Cord and Brain Injury Research Fund (Grant 20000289), the Purdue Research Foundation (Grant 209911), and the Office of the Executive Vice President for Research and Partnerships at Purdue University (Grant 210362). We thank Dr. Cory J. Weaver and Haley Roeder for establishing zebrafish RGC culture protocol. We thank Haley Roeder additionally for providing the data of Figure 4. We thank Leah Biasi and Kenny Nguyen for the help with RGC culture. We thank Gentry Lee for editing the text. We thank Dr. Tobias Dick for providing roGFP2-Orp1 and Dr. Qing Deng for pCS2+ vector containing roGFP2-Orp1. Figure 2&#160;is created with Biorender.com.

<ol>
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Authors declare that they have no conflict of interest.

There are several critical steps that need&#160;attention throughout this protocol. We believe considering these points will improve the experimental flow. For primary RGC culture, the sterility of the ZFCM(-) is very important, since this culture media does not contain antibiotics and contamination can occur before or during imaging. To avoid it, we advise to prepare and use ZFCM(-) only inside a biosafety cabinet and make fresh ZFCM(-) media regularly (Step 1.5). In addition, laminin stocks should be kept at -80 &#176;...

Reactive oxygen species (ROS) signaling regulates development and functioning of the nervous system1. An important cellular ROS source are NADPH oxidases (NOX), which are transmembrane proteins generating superoxide and hydrogen peroxide (H2O2)2. NOX enzymes are found throughout the central nervous system (CNS), and NOX-derived ROS contribute to neuronal development3,4,5,6. Maintenance and differentiation of neural stem cells, establishing neuronal polarity, neurite outgrowth, and synaptic plasticity have been shown to require adequate levels of ROS7,8,9,10,11. On the other hand, uncontrolled production of ROS by NOXes contribute to neurodegenerative disorders including Alzheimer&#39;s Disease, multiple sclerosis, and traumatic brain injury12,13,14. Hence, production of physiologically relevant ROS is critical to maintaining healthy conditions.
Development of genetically encoded biosensors facilitated the detection of cellular ROS greatly. One important advantage of genetically encoded biosensors is the increased temporal and spatial resolution of the ROS signal, as these sensors can be specifically targeted to distinct locations. Redox-sensitive GFP (roGFP) is one type of such ROS biosensors. The roGFP2-Orp1 variant specifically detects H2O2 through its Orp1 domain, which is a glutathione peroxiredoxin family protein from yeast15,16. The oxidation of the Orp1 protein is transferred to roGFP2 to alter its conformation (Figure 1A). The probe exhibits two excitation peaks near 405 nm and 480 nm, and a single emission peak at 515 nm. Upon oxidation, the fluorescence intensity around excitation peaks changes: while 405 nm excitation increases, 480 nm excitation decreases. Thus, roGFP2-Orp1 is a ratiometric biosensor, and H2O2-levels are detected by the ratio of fluorescence intensities excited at two different wavelengths (Figure 1B). Overall, roGFP2-Orp1 is a versatile tool for ROS imaging that can be utilized efficiently in vivo.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62165/62165fig01.jpg" /> 
Figure 1: Schematic representation and excitation spectra of roGFP2-Orp1. (A) Oxidant transfer occurs between Orp1 and roGFP2 in response to H2O2, leading to conformational changes in roGFP2. (B) The excitation spectra of the roGFP2-Orp1 exhibits two excitation peaks at 405 nm and 480 nm and single emission peak at 515 nm. Upon oxidation by H2O2, the 405 nm excitation increases while 480 nm excitation decreases. This results in a ratiometric readout for H2O2 presence. The figure has been modified from Bilan and Belousov (2017)16 and Morgan et al. (2011)25. <a href="https://www.jove.com/files/ftp_upload/62165/62165fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The Danio rerio (zebrafish) model system has several advantages for applying genetically encoded biosensors. The optical transparency of the embryos and larvae enables non-invasive in vivo&#160;imaging. New imaging tools are being developed to achieve higher resolution and deeper penetration17. Furthermore, there are established tools for genetic manipulation (ectopic mRNA expression, Tol2 transgenesis, etc.) and genome editing (TALENs, CRISPR/Cas9, etc.), which promotes the generation of transgenic animals18. As the zebrafish embryos develop outside of the mother, this system further allows easier access and manipulation of the embryos. For instance, one-cell stage injections and drug treatments can easily be done.
Here, we used zebrafish to transiently express the H2O2-specific biosensor roGFP2-Orp1 by injecting in vitro-transcribed mRNA. These embryos can be used for both in vitro imaging of cultured neurons and in vivo imaging (Figure 2). We describe a protocol for dissecting and plating retinal ganglion cells (RGCs) from zebrafish embryos followed by assessing H2O2-levels in cultured neurons. Then, we present a method for in vivo imaging of roGFP2-Orp1-expressing embryos and larvae using confocal microscopy. This approach not only allows to determine physiological H2O2-levels but also potential changes occurring in different developmental stages or conditions. Overall, this system provides a reliable method for detecting H2O2 in live cells and animals to study the role of H2O2 in development, health, and disease.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62165/62165fig03.jpg" /> 
Figure 2. Outline of the experimental approach. Briefly, after embryo collection, roGFP2-Orp1 mRNA is injected into the yolk of one-cell stage zebrafish embryos. Developing embryos can be used for both (A) in vitro and (B) in vivo imaging. (A) GFP-positive embryos are used to dissect retinas for RGC collection at 34 hpf. Dissociated RGCs are plated on PDL/laminin-coated coverslips in ZFCM (+) media. Growth cone imaging can be conducted as RGCs extend their axons after 6-24 h of plating. Cells can be subjected to different treatments to measure the potential changes in H2O2-levels. Here, we measured H2O2-levels in the growth cones of RGCs (red). (B) GFP positive embryos are used for in vivo imaging. At the desired age, embryos can be anesthetized and mounted on 35 mm glass bottom dishes for confocal imaging. Here, embryos are mounted ventrally for retinal imaging. Schematic shows retinal development in zebrafish. RGCs form ganglion cell layer (GCL), which is the innermost layer in retina. RGC axons develop into optic nerve to cross midline, forming optic chiasm. Then, RGC axons grow dorsally to make synapses in the optic tectum in the midbrain. <a href="https://www.jove.com/files/ftp_upload/62165/62165fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>35-mm culture dish</td>
			<td>Sarstedt</td>
			<td>83-3900</td>
			<td></td>
		</tr>
		<tr>
			<td>35-mm glass bottom dish</td>
			<td>MatTek</td>
			<td>P35G-1.5-10-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>Fisher Scientific</td>
			<td>BP160-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate Glass Capillary Tubes</td>
			<td>Sutter/Fisher Scientific</td>
			<td>NC9029378</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Chloride Dihydrate</td>
			<td>Fisher Scientific</td>
			<td>C79-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover glass</td>
			<td>Corning</td>
			<td>2850-22</td>
			<td></td>
		</tr>
		<tr>
			<td>Disposable Petri Dishes (100 x 15 mm)</td>
			<td>VWR</td>
			<td>25384-094</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>ThermoFisher Scientific</td>
			<td>26140087</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma Aldich</td>
			<td>G7528</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma Aldich</td>
			<td>H4034</td>
			<td></td>
		</tr>
		<tr>
			<td>Injection Mold</td>
			<td>Adaptive Science Tools</td>
			<td>TU-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted Microscope</td>
			<td>Nikon</td>
			<td>TE2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Laminin</td>
			<td>ThermoFisher Scientific</td>
			<td>23017-015</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser Scanning Confocal Microscopy</td>
			<td>Zeiss</td>
			<td>710</td>
			<td></td>
		</tr>
		<tr>
			<td>Leibovitz&#39;s L-15 Medium with phenol red</td>
			<td>Gibco/Fisher Scientific</td>
			<td>11-415-064</td>
			<td></td>
		</tr>
		<tr>
			<td>Leibovitz&#39;s L-15 Medium without phenol red</td>
			<td>Gibco/Fisher Scientific</td>
			<td>21-083-027</td>
			<td></td>
		</tr>
		<tr>
			<td>Low melting agarose</td>
			<td>Promega</td>
			<td>V2111</td>
			<td></td>
		</tr>
		<tr>
			<td>mMessage mMachine SP6 Transcription Kit</td>
			<td>Invitrogen</td>
			<td>AM1340</td>
			<td></td>
		</tr>
		<tr>
			<td>NotI</td>
			<td>New England Biolabs</td>
			<td>R0189S</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Hyclone/Fisher Scientific</td>
			<td>SH3025601</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin/streptomycin</td>
			<td>ThermoFisher Scientific</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr>
			<td>Phenol Red</td>
			<td>Sigma Aldich</td>
			<td>P0290</td>
			<td></td>
		</tr>
		<tr>
			<td>Phenylthiourea (PTU)</td>
			<td>Sigma Aldich</td>
			<td>P7629</td>
			<td></td>
		</tr>
		<tr>
			<td>Pneumatic PicoPump</td>
			<td>World Precision Instruments</td>
			<td>PV820</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-D-Lysine (PDL)</td>
			<td>Sigma Aldich</td>
			<td>P7280</td>
			<td></td>
		</tr>
		<tr>
			<td>QiaQUICK PCR Purification Kit</td>
			<td>QIAGEN</td>
			<td>28104</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant mouse slit2</td>
			<td>R&#38;D Systems</td>
			<td>5444-SL-050</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Pyruvate</td>
			<td>Sigma Aldich</td>
			<td>P5280</td>
			<td></td>
		</tr>
		<tr>
			<td>Steritop 0.22 &#181;m filter</td>
			<td>Millipore</td>
			<td>S2GPT05RE</td>
			<td></td>
		</tr>
		<tr>
			<td>TE Buffer</td>
			<td>Ambion</td>
			<td>AM9860</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine Methanesulfonate</td>
			<td>Sigma Aldich</td>
			<td>E10521</td>
			<td></td>
		</tr>
		<tr>
			<td>Vertical Pipette Puller</td>
			<td>David Kopf Instruments</td>
			<td>700C</td>
			<td></td>
		</tr>
	</tbody>
</table>

ros live cell imaging during neuronal development

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the different ROS, hydrogen peroxide (H2O2) is best characterized with respect to roles in cellular signaling. H2O2 has been implicated during the development in several species. For example, a transient increase in H2O2 has been detected in zebrafish embryos during the first days following fertilization. Furthermore, depleting an important cellular H2O2 source, NADPH oxidase (NOX), impairs nervous system development such as the differentiation, axonal growth, and guidance of retinal ganglion cells (RGCs) both in&#160;vivo and in vitro. Here, we describe a method for imaging intracellular H2O2 levels in cultured zebrafish neurons and whole larvae during development using the genetically encoded H2O2-specific biosensor, roGFP2-Orp1. This probe can be transiently or stably expressed in zebrafish larvae. Furthermore, the ratiometric readout diminishes the probability of detecting artifacts due to differential gene expression or volume effects. First, we demonstrate how to isolate and culture RGCs derived from zebrafish embryos that transiently express roGFP2-Orp1. Then, we use whole larvae to monitor H2O2 levels at the tissue level. The sensor has been validated by the addition of H2O2. Additionally, this methodology could be used to measure H2O2 levels in specific cell types and tissues by generating transgenic animals with tissue-specific biosensor expression. As zebrafish facilitate genetic and developmental manipulations, the approach demonstrated here could serve as a pipeline to test the role of H2O2 during neuronal and general embryonic development in vertebrates.

Cultured zebrafish RGCs extend axons within 1d. A representative 405/480 ratio image of the H2O2-biosensor is shown in Figure 4A. The cell body, axon, and growth cones are clearly visible in individual neurons. These neurons can be subjected to different treatments over time to monitor H2O2 changes. We previously found that adding 100 &#181;M H2O2 to culture media increases the ratio values, showing that real-time changes can ...

Watch this Scientific Journal Video about ROS Live Cell Imaging During Neuronal Development at JoVE.com

ROS Live Cell Imaging During Neuronal Development

This protocol describes the use of a genetically encoded hydrogen peroxide (H2O2)-biosensor in cultured zebrafish neurons and larvae for assessing the physiological signaling roles of H2O2 during nervous system development. It can be applied to different cell types and modified with experimental treatments to study reactive oxygen species (ROS) in general development.

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the ...

ros-live-cell-imaging-during-neuronal-development

Research

JoVE Journal

Neuroscience

4.1K Views.  Purdue University. This protocol describes the use of a genetically encoded hydrogen peroxide (H2O2)-biosensor in cultured zebrafish neurons and larvae for assessing the physiological signaling roles of H2O2 during nervous system development. It can be applied to different cell types and modified with experimental treatments to study reactive oxygen species (ROS) in general development.

Video: ROS Live Cell Imaging During Neuronal Development

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Live Imaging of Dorsal Root Axons after Rhizotomy

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. Regeneration of dorsal root (DR) axons into spinal cord is prevented at the dorsal root entry zone (DREZ), the interface between the CNS and PNS. Our understanding of the molecular and cellular events that prevent regeneration at DREZ is incomplete, in part because complex changes associated with nerve injury have been deduced from postmortem analyses. Dynamic cellular processes, such as axon regeneration, are best studied with techniques that capture real-time events with multiple observations of each living animal. Our ability to monitor neurons serially in vivo has increased dramatically owing to revolutionary innovations in optics and mouse transgenics. Several lines of thy1-GFP transgenic mice, in which subsets of neurons are genetically labeled in distinct fluorescent colors, permit individual neurons to be imaged in vivo1. These mice have been used extensively for in vivo imaging of muscle2-4 and brain5-7, and have provided novel insights into physiological mechanisms that static analyses could not have resolved. Imaging studies of neurons in living spinal cord have only recently begun. Lichtman and his colleagues first demonstrated their feasibility by tracking injured dorsal column (DC) axons with wide-field microscopy8,9. Multi-photon in vivo imaging of deeply positioned DC axons, microglia and blood vessels has also been accomplished10. Over the last few years, we have pioneered in applying in vivo imaging to monitor regeneration of DR axons using wide-field microscopy and H line of thy1-YFP mice. These studies have led us to a novel hypothesis about why DR axons are prevented from regenerating within the spinal cord11.
In H line of thy1-YFP mice, distinct YFP+ axons are superficially positioned, which allows several axons to be monitored simultaneously. We have learned that DR axons arriving at DREZ are better imaged in lumbar than in cervical spinal cord. In the present report we describe several strategies that we have found useful to assure successful long-term and repeated imaging of regenerating DR axons. These include methods that eliminate repeated intubation and respiratory interruption, minimize surgery-associated stress and scar formation, and acquire stable images at high resolution without phototoxicity.

An in vivo imaging protocol to monitor primary sensory axons following dorsal root crush is described. The procedures utilize wide-field fluorescence microscopy and thy1-YFP transgenic mice, and permit repeated imaging of axon regeneration over 4 cm in the PNS and axon interactions with the interface of the CNS.

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. ...

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 ...

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of existing mouse lines for genetic cell lineage tracing: a tamoxifen-inducible Cre line and a Cre reporter line expressing dsRed upon Cre-mediated recombination. By using a relatively low level of tamoxifen, we were able to induce recombination in a small number of cells, permitting us to follow individual cell divisions. Additionally, we observed the transcriptional response to Sonic Hedgehog (Shh) signaling using an Olig2-eGFP transgenic line 1-3 and we monitored formation of cilia by infecting the cultured slice with virus expressing the cilia marker, Sstr3-GFP 4. In order to image the neuroepithelium, we harvested embryos at E8.5, isolated the neural tube, mounted the neural slice in proper culturing conditions into the imaging chamber and performed time-lapse confocal imaging. Our ex vivo live imaging method enables us to trace single cell divisions to assess the relative timing of primary cilia formation and Shh response in a physiologically relevant manner. This method can be easily adapted using distinct fluorescent markers and provides the field the tools with which to monitor cell behavior in situ and in real time.

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

Here we develop the tools necessary for ex vivo live imaging to trace single cell divisions in the mouse E8.5 neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of ...

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 ...

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 ...

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

Analyzing Dendritic Morphology in Columns and Layers

In many regions of the central nervous systems, such as the fly optic lobes and the vertebrate cortex, synaptic circuits are organized in layers and columns to facilitate brain wiring during development and information processing in developed animals. Postsynaptic neurons elaborate dendrites in type-specific patterns in specific layers to synapse with appropriate presynaptic terminals. The fly medulla neuropil is composed of 10 layers and about 750 columns; each column is innervated by dendrites of over 38 types of medulla neurons, which match with the axonal terminals of some 7 types of afferents in a type-specific fashion. This report details the procedures to image and analyze dendrites of medulla neurons. The workflow includes three sections: (i) the dual-view imaging section combines two confocal image stacks collected at orthogonal orientations into a high-resolution 3D image of dendrites; (ii) the dendrite tracing and registration section traces dendritic arbors in 3D and registers dendritic traces to the reference column array; (iii) the dendritic analysis section analyzes dendritic patterns with respect to columns and layers, including layer-specific termination and planar projection direction of dendritic arbors, and derives estimates of dendritic branching and termination frequencies. The protocols utilize custom plugins built on the open-source MIPAV (Medical Imaging Processing, Analysis, and Visualization) platform and custom toolboxes in the matrix laboratory language. Together, these protocols provide a complete workflow to analyze the dendritic routing of Drosophila medulla neurons in layers and columns, to identify cell types, and to determine defects in mutants.

Here, we show how to analyze dendritic routing of Drosophila medulla neurons in columns and layers. The workflow includes a dual-view imaging technique to improve the image quality and computational tools for tracing, registering dendritic arbors to the reference column array and for analyzing the dendritic structures in 3D space.

In many regions of the central nervous systems, such as the fly optic lobes and the vertebrate cortex, synaptic circuits are organized in layers and ...

In Vivo Calcium Imaging of Lateral-line Hair Cells in Larval Zebrafish

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory stimuli that mechanically deflect apical protrusions called hair bundles. Deflection opens mechanotransduction (MET) channels in hair bundles, leading to an influx of cations, including calcium. This cation influx depolarizes the cell and opens voltage-gated calcium channels located basally at the hair-cell presynapse. In mammals, hair cells are encased in bone, and it is challenging to functionally assess these activities in vivo. In contrast, larval zebrafish are transparent and possess an externally located lateral-line organ that contains hair cells. These hair cells are functionally and structurally similar to mammalian hair cells and can be functionally assessed in vivo. This article outlines a technique that utilizes a genetically encoded calcium indicator (GECI), GCaMP6s, to measure stimulus-evoked calcium signals in zebrafish lateral-line hair cells. GCaMP6s can be used, along with confocal imaging, to measure in vivo calcium signals at the apex and base of lateral-line hair cells. These signals provide a real-time, quantifiable readout of both mechanosensation- and presynapse-dependent calcium activities within these hair cells. These calcium signals also provide important functional information regarding how hair cells detect and transmit sensory stimuli. Overall, this technique generates useful data about relative changes in calcium activity in vivo. It is less well-suited for quantification of the absolute magnitude of calcium changes. This in vivo technique is sensitive to motion artifacts. A reasonable amount of practice and skill are required for proper positioning, immobilization, and stimulation of larvae. Ultimately, when properly executed, the protocol outlined in this article provides a powerful way to collect valuable information about the activity of hair-cells in their natural, fully integrated states within a live animal.

The zebrafish is a model system that has many valuable features including optical clarity, rapid external development, and, of particular importance to the field of hearing and balance, externally located sensory hair cells. This article outlines how transgenic zebrafish can be used to assay both hair-cell mechanosensation and presynaptic function in toto.

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory ...

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are optimized to minimize their impact on behavior. This is first achieved by using a holder that minimizes movement hindrances. The back of the fly&#8217;s head is glued to this holder at an angle that allows optical access to the whole brain while retaining the fly&#8217;s ability to walk, groom, smell, taste and see. The back of the head is dissected to remove tissues in the optical path and muscles responsible for head movement artefacts. The fly brain can subsequently be imaged to record brain activity, for instance using calcium or voltage indicators, during specific behaviors such as walking or grooming, and in response to different stimuli. Once the challenging dissection, which requires considerable practice, has been mastered, this technique allows to record rich data sets relating whole brain activity to behavior and stimulus responses.

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method specifically tailored to image the whole brain of adult Drosophila during behavior and in response to stimuli. The head is positioned to allow optical access to the whole brain, while the fly can move its legs and the antennae, the tip of the proboscis, and the eyes can receive sensory stimuli.

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are ...