Name: ros live cell imaging during neuronal development
Uploaded: 2021-02-09T14:30:00.000+00:00
Duration: 9 min 25 s
Description: Watch this Scientific Journal Video about ROS Live Cell Imaging During Neuronal Development at JoVE.com

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;C. Thawed laminin should be stored at 4 &#176;C at all times. Do not keep laminin solutions at room temperature or refreeze, as it decreases the neurite outgrowth promoting activity of the protein.
Dissecting zebrafish retina at the proper time is important to yield enough RGCs that extend axons. Zebrafish RGCs are born at 28 hpf and extend axons out of retina starting 32-36 hpf. Therefore, it is critical to plate RGCs after 28 hpf, preferably between 32-36 hpf, to allow for full differentiation of RGCs. Cultures prepared at earlier time points may not yield enough RGCs or will yield RGCs that may not extend axons within 24 hours after plating. It is highly recommended to change the ZFCM(-) media when imaging for more than 30 min. As this media lacks phenol red, pH changes are not visible and may need to be considered. Exchange media very carefully to avoid dislodging neurons during the process (Step 4.7).
Finally, changing the media to E3 with PTU but without methylene blue is necessary by 24 hpf. PTU prevents pigmentation for enhanced larval transparency required for in vivo fluorescence imaging; however, earlier treatment with PTU can cause developmental defects26. Methylene blue is removed from E3 media to reduce autofluorescence, mainly originating from the yolk (Step 5.1).
This method allows cytoplasmic detection of H2O2. However, subcellular compartmentalization of H2O2 contributes to local signaling, and cytoplasmic H2O2-levels may not provide sufficient information on specific signaling roles of H2O2. As roGFP2-Orp1 is a genetically encoded biosensor, it can be targeted for tissue-specific expression (e.g., neurons, astrocytes, glial cells, etc.) or specific subcellular locations (e.g., plasma membrane, mitochondria, nucleus, etc.). As roGFP2-Orp1 is a GFP-based biosensor, the combination with other fluorescent probes is limited to emission wavelengths outside the green spectrum. Since GFP is widely used as transgenesis marker, utilization of GFP-based biosensors can be limited. To address this challenge, red-fluorescent proteins for H2O2-detection have been developed recently27. Furthermore, roGFP2-Orp1 is transiently expressed in the method presented here. Hence, the mRNA and protein expression will decay over time. We were still able to detect the signal in 5 dpf larvae; however, transgenic fish lines need to be established for stable expression of the biosensor throughout development and into adulthood.
Several dye-based assays are available for detecting intracellular ROS (e.g., H2DCF-DA), or specifically, H2O2 (PF6-AM)28,29. Using a genetically encoded biosensor, such as roGFP2-Orp1, has several advantages over dye-based methods. First, it allows detection of both transient changes in oxidative status because of reversibility as well as at specific subcellular locations. Hence, the dynamic changes are well monitored with high degree of spatial resolution30. Second, it generally exhibits less toxicity to cells or tissues when compared to dye molecules. Finally, the biosensor can be further genetically manipulated to meet the needs of specific experiments31. roGFP2-Orp1 is a ratiometric biosensor, since it has two excitation and one emission peak. Upon interaction with H2O2, 405 nm excitation increases, and 480 nm excitation decreases. Hence, the readout is the ratio of the intensity values acquired by 405 and 480 nm excitation, respectively. Because this probe is ratiometric, application of a second probe becomes unnecessary to correct for potential volume effects. Furthermore, since this probe involves only expression of one polypeptide, it does not suffer from potential problems of differential expression of more than one protein, such as in the case of intermolecular Fluorescence Resonance Energy Transfer (FRET).
The roGFP2-Orp1 is a versatile H2O2 probe to study ROS production and signaling. In live cells, this probe exhibits dynamic range of 4.8 (around 4 in transgenic zebrafish embryos)15,32. However, the redox state of roGFP2-Orp1 is subject to alterations by endogenous thioredoxin and glutaredoxin systems; hence, more sensitive probes were developed by replacing Orp1 with another yeast peroxiredoxin, Tsa233,34. While roGFP2-Orp1 is useful to detect changes in H2O2 in zebrafish embryos, the sensitivity is mostly limited to higher concentrations of exogenously applied H2O224,31, or to potent stimuli. For instance, slit2 treatment increased the roGFP2-Orp1 signal in wild type RGCs, but not in nox2 mutant RGCs35. On the other hand, there was no difference in basal H2O2 levels between wildtype and nox2 mutant RGCs and 2 dpf embryos24,35. Hence, to detect basal levels or small changes in H2O2, higher sensitivity probes will be needed.
HyPer is another widely used genetically encoded and ratiometric H2O2-specific biosensor36. The sensitivities of both probes are similar; however, roGFP2-Orp1 exhibited slower responsiveness to H2O2 changes15. Therefore, precise detection of real-time changes might require HyPer instead of roGFP2-Orp1. Whereas initial HyPer versions are affected by pH changes, roGFP2-Orp1 is not16,36. Hence, it is critical to include a pH control probe (SypHer3s) while using HyPer37. The most recent HyPer probe, HyPer7, overcame many of the drawbacks of existing H2O2 probes: HyPer7 has a higher pH stability, increased brightness, faster kinetics and higher dynamic range38. Furthermore, HyPer7 is validated both in vitro and in vivo, showing the subcellular compartmentalization and monitoring real time H2O2 changes in zebrafish embryos during wound healing38.
The methodology presented here can be expanded to study the role of ROS signaling both in vitro and in vivo. roGFP2-Orp1 expressing neurons can be subjected to different treatments to study their effect on H2O2 signaling. For instance, we showed that RGCs respond to changes in H2O2 levels in real time (Figure 4), suggesting that the biosensor can detect immediate changes in H2O2. However, determining absolute intracellular H2O2 concentrations would require extensive calibration of the system. The detection limit of the biosensor is also unclear. Additionally, the detailed spatial distribution of the biosensor in the cell is not completely known, which could interfere with the interpretation of the results. Furthermore, determining the exact location signal from roGFP2-Orp1 could reveal whether there is site-specific H2O2 production that contributes to localized intracellular signaling.
Creating transgenic fish expressing roGFP2-Orp1 has numerous advantages. The biosensor can be expressed ubiquitously, or under a tissue-specific promoter for studying cell-specific H2O2 signaling by using Tol2 transgenesis in zebrafish39. The role of H2O2 in development can be studied by ubiquitous expression of the biosensor. For instance, transgenic zebrafish embryos can be monitored to assess changes in H2O2 levels and how these changes contribute to different developmental stages. On the other hand, tissue-specific expression of the roGFP2-Orp1 focuses on the effects of H2O2 production in cells or tissues of interest. Tissue or cell-specific promoters can be used to drive expression of roGFP2-Orp1 in specific cells. Furthermore, the Gal4/UAS system can be used for sparse labeling of individual neurons. Hence, this approach allows detecting H2O2 levels at a cellular resolution&#160;in vivo. Finally, stable transgenic lines can be used for high-throughput drug screening that involves ROS signaling and oxidative stress in zebrafish embryos.

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.bdigitaluss.remotexs.co/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.bdigitaluss.remotexs.co/files/ftp_upload/62165/62165fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><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&#039;s L-15 Medium with phenol red</td><td>Gibco/Fisher Scientific</td><td>11-415-064</td><td></td></tr><tr><td>Leibovitz&#039;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&amp;amp;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 &amp;micro;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 be detected with this system (Figure 4)24.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62165/62165fig04.jpg" /> 
Figure 4:&#160;H2O2 imaging in cultured zebrafish RGCs with roGFP2-Orp1. (A) Representative 405/480 nm ratio images of cultured RGC neurons from 34 hpf old zebrafish embryos. Neuron on the left was treated with serum-free control medium and neuron on the right was treated with 100 &#181;M H2O2 for 30 min. Scale bar = 10 &#181;m. (B) Bar graphs showing the average H2O2 levels in the wild type zebrafish RGC growth cones before and after treatment with either control or 100 &#181;M H2O2. The data were normalized to control treatment. Data are shown mean &#177; SEM. Number of growth cones are shown in parentheses for each group. Two-tailed Wilcoxon matched-pairs signed rank test; **p = 0.0009. The figure has been modified from our previous publication24. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/62165/62165fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
H2O2 levels can also be determined in whole zebrafish embryos (Figure 5). As the mRNA is injected during the one-cell stage, all tissues express the roGFP2-Orp1 biosensor. In Figure 5A, we show the head region of zebrafish larvae at two different time points, focusing on the H2O2 levels in the retina, as an example. We have previously shown that addition of H2O2 increases the ratio values in real time, while addition of the reducing agent DTT reverses this effect in vivo24. Here, we measured the basal levels of H2O2 in the same zebrafish embryos at 2 dpf and 5 dpf. At 2 dpf, the ratio values were significantly lower than at 5 dpf (p&#60;0.0001; Figure 5B). Furthermore, each animal showed a different level of increase in their retinal H2O2 content (Figure 5C).
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62165/62165fig05.jpg" /> 
Figure 5: In vivo&#160;H2O2 imaging in zebrafish larvae with roGFP2-Orp1. (A)&#160;DIC and 405/480 nm ratio images of 2 and 5 dpf zebrafish larvae. ROI is defined as retina (yellow lines) to measure ratio values. The same larvae were analyzed at 2 and 5 dpf. Scale bars = 50 &#181;m. (B) Measurement of average H2O2 levels in retinas of 2 and 5 dpf zebrafish. At 5 dpf, H2O2-levels are increased significantly in the retina compared to 2 dpf. (C) Line graph of measurements of individual fish. Each animal exhibits an increase in the H2O2-levels in their retina from 2 to 5 dpf. Data are shown as mean &#177; SEM. Number of larvae analyzed are shown in parentheses for each group. Two-tailed Wilcoxon matched-pairs signed rank test, ****p&#60;0.0001. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/62165/62165fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

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

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 ...

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 ...

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 ...

Real-time Bioluminescence Imaging of Notch Signaling Dynamics during Murine Neurogenesis

Notch signaling regulates the maintenance of neural stem/progenitor cells by cell-cell interactions. The components of Notch signaling exhibit dynamic expression. Notch signaling effector Hes1 and the Notch ligand Delta-like1 (Dll1) are expressed in an oscillatory manner in neural stem/progenitor cells. Because the period of the oscillatory expression of these genes is very short (2 h), it is difficult to monitor their cyclic expression. To examine such rapid changes in the gene expression or protein dynamics, fast response reporters are required. Because of its fast maturation kinetics and high sensitivity, the bioluminescence reporter luciferase is suitable to monitor rapid gene expression changes in living cells. We used a destabilized luciferase reporter for monitoring the promoter activity and a luciferase-fused reporter for visualization of protein dynamics at single cell resolution. These bioluminescence reporters show rapid turnover and generate very weak signals; therefore, we have developed a highly sensitive bioluminescence imaging system to detect such faint signals. These methods enable us to monitor various gene expression dynamics in living cells and tissues, which are important information to help understand the actual cellular states.

Neural stem/progenitor cells exhibit various expression dynamics of Notch signaling components that lead to different outcomes of cellular events. Such dynamic expression can be revealed by real-time monitoring, not by static analysis, using a highly sensitive bioluminescence imaging system that enables visualization of rapid changes in gene expressions.

Notch signaling regulates the maintenance of neural stem/progenitor cells by cell-cell interactions. The components of Notch signaling exhibit dynamic ...

Developmental Biology

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 ...

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 ...

Live Imaging of Primary Cerebral Cortex Cells Using a 2D Culture System

During cerebral cortex development, progenitor cells undergo several rounds of symmetric and asymmetric cell divisions to generate new progenitors or postmitotic neurons. Later, some progenitors switch to a gliogenic fate, adding to the astrocyte and oligodendrocyte populations. Using time-lapse video-microscopy of primary cerebral cortex cell cultures, it is possible to study the cellular and molecular mechanisms controlling the mode of cell division and cell cycle parameters of progenitor cells. Similarly, the fate of postmitotic cells can be examined using cell-specific fluorescent reporter proteins or post-imaging immunocytochemistry. More importantly, all these features can be analyzed at the single-cell level, allowing the identification of progenitors committed to the generation of specific cell types. Manipulation of gene expression can also be performed using viral-mediated transfection, allowing the study of cell-autonomous and non-cell-autonomous phenomena. Finally, the use of fusion fluorescent proteins allows the study of symmetric and asymmetric distribution of selected proteins during division and the correlation with daughter cells fate. Here, we describe the time-lapse video-microscopy method to image primary cerebral cortex murine cells for up to several days and analyze the mode of cell division, cell cycle length and fate of newly generated cells. We also describe a simple method to transfect progenitor cells, which can be applied to manipulate genes of interest or simply label cells with reporter proteins.

Live imaging is a powerful tool to study cellular behaviors in real time. Here, we describe a protocol for time-lapse video-microscopy of primary cerebral cortex cells that allows a detailed examination of the phases enacted during the lineage progression from primary neural stem cells to differentiated neurons and glia.

During cerebral cortex development, progenitor cells undergo several rounds of symmetric and asymmetric cell divisions to generate new progenitors or ...

Live Imaging Followed by Single Cell Tracking to Monitor Cell Biology and the Lineage Progression of Multiple Neural Populations

Understanding the mechanisms that control critical biological events of neural cell populations, such as proliferation, differentiation, or cell fate decisions, will be crucial to design therapeutic strategies for many diseases affecting the nervous system. Current methods to track cell populations rely on their final outcomes in still images and they generally fail to provide sufficient temporal resolution to identify behavioral features in single cells. Moreover, variations in cell death, behavioral heterogeneity within a cell population, dilution, spreading, or the low efficiency of the markers used to analyze cells are all important handicaps that will lead to incomplete or incorrect read-outs of the results. Conversely, performing live imaging and single cell tracking under appropriate conditions represents a powerful tool to monitor each of these events. Here, a time-lapse video-microscopy protocol, followed by post-processing, is described to track neural populations with single cell resolution, employing specific software. The methods described enable researchers to address essential questions regarding the cell biology and lineage progression of distinct neural populations.

A robust protocol to monitor neural populations by time-lapse video-microscopy followed by software-based post-processing is described. This method represents a powerful tool to identify biological events in a selected population during live imaging experiments.

Understanding the mechanisms that control critical biological events of neural cell populations, such as proliferation, differentiation, or cell fate ...

Time-Lapse Imaging of Neuronal Arborization using Sparse Adeno-Associated Virus Labeling of Genetically Targeted Retinal Cell Populations

Discovering mechanisms that pattern dendritic arbors requires methods to visualize, image, and analyze dendrites during development. The mouse retina is a powerful model system for the investigation of cell type-specific mechanisms of neuronal morphogenesis and connectivity. The organization and composition of retinal subtypes are well-defined, and genetic tools are available to access specific types during development. Many retinal cell types also constrain their dendrites and/or axons to narrow layers, which facilitates time-lapse imaging. Mouse retina explant cultures are well suited for live-cell imaging using confocal or multiphoton microscopy, but methods optimized for imaging dendrite dynamics with temporal and structural resolution are lacking. Presented here is a method to sparsely label and image the development of specific retinal populations marked by the Cre-Lox system. Commercially available adeno-associated viruses (AAVs) used here expressed membrane-targeted fluorescent proteins in a Cre-dependent manner. Intraocular delivery of AAVs in neonatal mice produces fluorescent labeling of targeted cell types by 4-5 days post-injection (dpi). The membrane fluorescent signals are detectable by confocal imaging and resolve fine branch structures and dynamics. High-quality videos spanning 2-4 h are acquired from imaging retinal flat-mounts perfused with oxygenated artificial cerebrospinal fluid (aCSF). Also provided is an image postprocessing pipeline for deconvolution and three-dimensional (3D) drift correction. This protocol can be used to capture several cellular behaviors in the intact retina and to identify novel factors controlling neurite morphogenesis. Many developmental strategies learned in the retina will be relevant for understanding the formation of neural circuits elsewhere in the central nervous system.

Here, we present a method for investigating neurite morphogenesis in postnatal mouse retinal explants by time-lapse confocal microscopy. We describe an approach for sparse labeling and acquisition of retinal cell types and their fine processes using recombinant adeno-associated virus vectors that express membrane-targeted fluorescent proteins in a Cre-dependent manner.

Discovering mechanisms that pattern dendritic arbors requires methods to visualize, image, and analyze dendrites during development. The mouse retina is a ...

Visualizing Neuronal and Glial Cell Dynamics in the Developing Mouse Brain

Time-Lapse Imaging of Migrating Neurons and Glial Progenitors in Embryonic Mouse Brain Slices

During the development of the cerebral cortex, neurons and glial cells originate in the ventricular zone lining the ventricle and migrate toward the brain surface. This process is crucial for proper brain function, and its dysregulation can result in neurodevelopmental and psychiatric disorders after birth. In fact, many genes responsible for these diseases have been found to be involved in this process, and therefore, revealing how these mutations affect cellular dynamics is important for understanding the pathogenesis of these diseases. This protocol introduces a technique for time-lapse imaging of migrating neurons and glial progenitors in brain slices obtained from mouse embryos. Cells are labeled with fluorescent proteins using in utero electroporation, which visualizes individual cells migrating from the ventricular zone with a high signal-to-noise ratio. Moreover, this in vivo gene transfer system enables us to easily perform gain-of-function or loss-of-function experiments on the given genes by co-electroporation of their expression or knockdown/knockout vectors. Using this protocol, the migratory behavior and migration speed of individual cells, information that is never obtained from fixed brains, can be analyzed.

Author Spotlight: Exploring Cell Migration and Gene Roles in the Developing Brain

During the development of the cerebral cortex, neurons and glial cells originate in the ventricular zone lining the ventricle and migrate toward the brain surface. Many genes are involved in this process. This protocol introduces the technique for the time-lapse imaging of migrating neurons and glial progenitors.

During the development of the cerebral cortex, neurons and glial cells originate in the ventricular zone lining the ventricle and migrate toward the brain ...

ROS Live Cell Imaging During Neuronal Development

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

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

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Real-time Bioluminescence Imaging of Notch Signaling Dynamics during Murine Neurogenesis

Live Imaging of Drosophila Larval Neuroblasts

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

Live Imaging of Primary Cerebral Cortex Cells Using a 2D Culture System

Live Imaging Followed by Single Cell Tracking to Monitor Cell Biology and the Lineage Progression of Multiple Neural Populations

Time-Lapse Imaging of Neuronal Arborization using Sparse Adeno-Associated Virus Labeling of Genetically Targeted Retinal Cell Populations

Time-Lapse Imaging of Migrating Neurons and Glial Progenitors in Embryonic Mouse Brain Slices