This work was supported by the National Institute for Mental Health, and National Science Foundation. Special thanks to Sive lab members for many useful discussions and constructive criticism, and to Olivier Paugois for expert fish husbandry.

1. Preparing for Microinjection
<ol>
 <li>Prepare microinjection needles by pulling capillary tubes using Sutter instruments needle puller.</li>
 <li>Load microinjection needle with fluorescent dye (FITC-Dextran).</li>
 <li>Mount needle on micromanipulator and microinjection apparatus.</li>
 <li>Carefully break microinjection needle using forceps to roughly 2 &mu;m in width, however, this will vary depending on your microinjector setup. For our microinjection needles, this corresponds to the first region of the needle from the tip that does not bend.</li>
 <li>Measure drop size in oil, adjusting injection time and pressure, so that each injection delivers 1 nl. Example settings for Harvard Apparatus microinjector are: P balance = 1.4 psi, P out = 1.4 psi, P inject = 22.9 psi, P clear = 67.8 psi with an injection time of .4 to .7 sec. The diameter of our needle using these settings is roughly 2 &mu;m. However, settings will be microinjector specific, and vary according to needle diameter.</li>
</ol>
2. Preparing the Embryos
<ol>
 <li>Coat 2 dishes with 1% agarose in water for each condition, poke holes into agarose with a 1-200 &mu;l pipette tip, and remove agarose plugs. Fill dishes with embryo media.</li>
 <li>Using forceps, dechorionate embryos that are 18 hpf or older under a stereomicroscope. Embryos are staged according to Kimmel et al.11 .</li>
 <li>Transfer dechorionated embryos into first agarose coated dish.</li>
 <li>To anesthetize embryos, add tricaine (0.1 mg/ml) to the dish until embryos stop moving (made according to Westerfield12).</li>
</ol>
3. Injecting the Brain Ventricles
<ol>
 <li>Orient embryos so you are looking at their dorsal side by putting the tail of the embryo into the hole. If your micromanipulator is on the right, then move the embryo so that the forebrain is to the left and hindbrain to the right.</li>
 <li>Position needle at widest point of hindbrain ventricle.</li>
 <li>Carefully pierce roof plate of hindbrain ventricle being sure not to go through the depth of the brain into the yolk (Figure 1A).</li>
 <li>Inject 1-2 nl of fluorescent dye into the ventricles making sure the dye fills the whole length of the brain ventricles.</li>
 <li>Transfer embryos to the second agarose coated dish filled with embryo media and re-anesthetize as described in 2.4.</li>
 <li>IMMEDIATELY start imaging, as described in section 4 in order to get a time zero image.</li>
</ol>
4. Imaging
<ol>
 <li>Orient embryos with their tail in the hole as described in 3.1.</li>
 <li>Use a dissecting microscope with both transmitted and fluorescent light to take a brightfield dorsal image. Keep the magnification constant between imaging of different embryos. This allows for direct comparison of analyses performed using Image J (5.2-6).</li>
 <li>WITHOUT MOVING the embryo, microscope or dish, take a corresponding fluorescent image.</li>
 <li>Repeat for each embryo at desired time points.</li>
</ol>
5. Quantification of Dye Movement
<ol>
 <li>Merge brightfield and fluorescent images in Photoshop as previously described by Gutzman and Sive10.</li>
 <li>Measure the distance the dye front moves in Image J software available at <a href="http://rsbweb.nih.gov/ij/" target="_blank">http://rsbweb.nih.gov/ij/</a>.</li>
 <li>Open merged file in Image J and use line tool to draw a line from the forebrain hinge-point to dye front at a 10-20 &deg; angle from neuroepithelium (Figure 1A). This region was chosen because it is the first and most noticeable site of dye leaking out of the wild type neuroepithelium.</li>
 <li>Select measurement tool to calculate length of line.</li>
 <li>Repeat for each time-point.</li>
 <li>Calculate net distance the dye front moved over time by subtracting distance at t=0 from other time points.</li>
 <li>Plot on graph.</li>
</ol>
6. Representative Results
An example of results obtained in a neuroepithelial permeability assay using wild type embryos is shown in Figure 1B-D. To accurately differentiate permeability, it is useful to test dyes with different molecular weightsto identify a size that is only slightly leaky in wild type or control embryos (Figure 2). This allows for identification of genetic mutants or environmental conditions that either increase or decrease permeability (Figure 1D, green and red lines respectively). For the 24 hpf zebrafish neuroepithelium, 70 kDa FITC Dextran leaks slowly over 2 hr, whereas 2,000 kDa does not and 10 kDa almost immediately leaks out. Therefore 70 kDa is the ideal molecular weight to identify conditions that both increase and decrease neuroepithelial permeability.
If the needle misses the ventricular lumen, fluorescence will appear outside the brain at t=0 (for an example see Gutzman and Sive, 200910). These embryos should be discarded since the injected dye was not initially contained within the brain and no clear conclusion regarding movement of the dye and permeability of the neuropeithelium can be made.
Finally, if embryos have small ventricles or un-inflated brain ventricles, pre-injection of ventricles with a saline solution can be done prior to injection of the fluorescent dye. This inflates the ventricles making subsequent visualization of the ventricles easier when injecting with the fluorescent dye. Proper controls must be performed to determine whether injection of saline disrupts normal neural tube development.
<img alt="Figure 1" src="/files/ftp_upload/4242/4242fig1.jpg" /> 
 Figure 1. Time course of different molecular weight dyes. (A) Experimental Diagram. First, fluorescent dye is injected into the ventricles. X = position of needle for injection. Next dorsal images are captured over time. Finally, the distance moved by the dye front from the forebrain hinge-point is measured (represented by a red line). (B-C) Merged brightfield and fluorescent dorsal images at 22 hpf (t = 0 min, B) and 24 hpf (t =120 min, C). White line indicates distance of the dye front from forebrain ventricle. (D) Hypothetical sample permeability data. Blue = wild type or controls, red = sample with decreased permeability relative to control, and green = sample with increased permeability relative to control.
<img alt="Figure 2" src="/files/ftp_upload/4242/4242fig2.jpg" /> 
 Figure 2. Measurement of neuroepithelial permeability to different molecular weight dyes. (A-E) Dorsal merged brightfield and fluorescent images of 22 hpf wild type embryos at t=0 min after injection with FITC-dextran of the following molecular weights: 10 kDa (A), 40 kDa (B), 70 kDa (C), 500 kDa (D) and 2,000 kDa (E). (A'-E') Same embryo as in (A-E) at t=120 min at 24 hpf. Anterior to left. F= forebrain, M= midbrain, H= hindbrain. Asterisk = ear.

<ol>
	<li>Harrington, M. J., Hong, E., Brewster, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+analysis+of+neurulation:+first+impressions+do+not+count.">Comparative analysis of neurulation: first impressions do not count.</a> Mol. Reprod. Dev. 76, 954-965 (2009).</li><li>Lowery, L. A., Sive, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strategies+of+vertebrate+neurulation+and+a+re-evaluation+of+teleost+neural+tube+formation.">Strategies of vertebrate neurulation and a re-evaluation of teleost neural tube formation.</a> Mech. Dev. 121, 1189-1197 (2004).</li><li>Salehi, Z., Mashayekhi, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+cerebrospinal+fluid+on+neural+cell+survival+in+the+developing+chick+cerebral+cortex:+an+in+vivo+study.">The role of cerebrospinal fluid on neural cell survival in the developing chick cerebral cortex: an in vivo study.</a> Eur. J. Neurol. 13, 760-764 (2006).</li><li>Martin, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+embryonic+brain+development+in+rats+requires+the+trophic+influence+of+cerebrospinal+fluid.">Early embryonic brain development in rats requires the trophic influence of cerebrospinal fluid.</a> Int. J. Dev. Neurosci. 27, 733-740 (2009).</li><li>Lehtinen, M. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+cerebrospinal+fluid+provides+a+proliferative+niche+for+neural+progenitor+cells.">The cerebrospinal fluid provides a proliferative niche for neural progenitor cells.</a> Neuron. 69, 893-905 (2011).</li><li>Gato, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Embryonic+cerebrospinal+fluid+regulates+neuroepithelial+survival,+proliferation,+and+neurogenesis+in+chick+embryos.">Embryonic cerebrospinal fluid regulates neuroepithelial survival, proliferation, and neurogenesis in chick embryos.</a> Anat. Rec. A. Discov. Mol. Cell Evol. Biol. 284, 475-484 (2005).</li><li>Lowery, L. A., Sive, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Totally+tubular:+the+mystery+behind+function+and+origin+of+the+brain+ventricular+system.">Totally tubular: the mystery behind function and origin of the brain ventricular system.</a> Bioessays. 31, 446-458 (2009).</li><li>Lowery, L. A., Sive, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Initial+formation+of+zebrafish+brain+ventricles+occurs+independently+of+circulation+and+requires+the+nagie+oko+and+snakehead/atp1a1a.1+gene+products.">Initial formation of zebrafish brain ventricles occurs independently of circulation and requires the nagie oko and snakehead/atp1a1a.1 gene products.</a> Development. 132, 2057-2067 (2005).</li><li>Zhang, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishment+of+a+neuroepithelial+barrier+by+Claudin5a+is+essential+for+zebrafish+brain+ventricular+lumen+expansion.">Establishment of a neuroepithelial barrier by Claudin5a is essential for zebrafish brain ventricular lumen expansion.</a> Proc. Natl. Acad. Sci. U.S.A. 107, 1425-1430 (2010).</li><li>Gutzman, J. H., Sive, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+brain+ventricle+injection.">Zebrafish brain ventricle injection.</a> J. Vis. Exp. , (2009).</li><li>Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., Schilling, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+embryonic+development+of+the+zebrafish.">Stages of embryonic development of the zebrafish.</a> Dev. Dyn. 203, 253-310 (1995).</li><li>Westerfield, M., Sprague, J., Doerry, E., Douglas, S., Grp, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Zebrafish+Information+Network+(ZFIN):+a+resource+for+genetic,+genomic+and+developmental+research.">The Zebrafish Information Network (ZFIN): a resource for genetic, genomic and developmental research.</a> Nucleic Acids Research. 29, 87-90 (2001).</li></ol>

No conflicts of interest declared.

We demonstrate the ability to quantify permeability of the living embryonic zebrafish brain as determined for an injected dye of a given molecular weight. Our observation that the embryonic zebrafish neuroepithelium is differentially permeable to dyes of differing molecular weights suggests that the dye is moving via paracellular permeability. However, we cannot rule out the possibility of a transcellular contribution to the observed permeability. This technique could be applied to any other tubular structure as long as both the inside and outside of the tube can be seen and the lumen can be injected. However, identification of the ideal molecular weight for other organs will need to be determined as this may differ between tissues and developmental stages .
This assay will enable further investigation into the role of epithelial permeability during lumen inflation and the regulation of lumen size. In addition, this technique will help characterize alterations in epithelial permeability associated with disorders such as hydrocephalus and polycystic kidney disease.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Dextran, Fluorescein, Anionic, Lysine Fixable</td>
 <td>Invitrogen</td>
 <td>D7136, D7137, D1822, D1820, D1845</td>
 </tr>
 <tr>
 <td>Tricaine powder</td>
 <td>Sigma</td>
 <td>A5040</td>
 </tr>
 <tr>
 <td>Capillary Tubes</td>
 <td>FHC Inc.</td>
 <td>30-30-1</td>
 </tr>
 </tbody>
</table>

an assay for permeability of the zebrafish embryonic neuroepithelium

The brain ventricular system is conserved among vertebrates and is composed of a series of interconnected cavities called brain ventricles, which form during the earliest stages of brain development and are maintained throughout the animal's life. The brain ventricular system is found in vertebrates, and the ventricles develop after neural tube formation, when the central lumen fills with cerebrospinal fluid (CSF) 1,2. CSF is a protein rich fluid that is essential for normal brain development and function3-6.
In zebrafish, brain ventricle inflation begins at approximately 18 hr post fertilization (hpf), after the neural tube is closed. Multiple processes are associated with brain ventricle formation, including formation of a neuroepithelium, tight junction formation that regulates permeability and CSF production. We showed that the Na,K-ATPase is required for brain ventricle inflation, impacting all these processes 7,8, while claudin 5a is necessary for tight junction formation 9. Additionally, we showed that "relaxation" of the embryonic neuroepithelium, via inhibition of myosin, is associated with brain ventricle inflation.
To investigate the regulation of permeability during zebrafish brain ventricle inflation, we developed a ventricular dye retention assay. This method uses brain ventricle injection in a living zebrafish embryo, a technique previously developed in our lab10, to fluorescently label the cerebrospinal fluid. Embryos are then imaged over time as the fluorescent dye moves through the brain ventricles and neuroepithelium. The distance the dye front moves away from the basal (non-luminal) side of the neuroepithelium over time is quantified and is a measure of neuroepithelial permeability (Figure 1). We observe that dyes 70 kDa and smaller will move through the neuroepithelium and can be detected outside the embryonic zebrafish brain at 24 hpf (Figure 2).
This dye retention assay can be used to analyze neuroepithelial permeability in a variety of different genetic backgrounds, at different times during development, and after environmental perturbations. It may also be useful in examining pathological accumulation of CSF. Overall, this technique allows investigators to analyze the role and regulation of permeability during development and disease.

Watch this Scientific Journal Video about An Assay for Permeability of the Zebrafish Embryonic Neuroepithelium at JoVE.com

An Assay for Permeability of the Zebrafish Embryonic Neuroepithelium

We describe a live whole animal quantitative measurement for permeability of the embryonic zebrafish brain. The technique analyzes the ability to retain cerebrospinal fluid and molecules of different molecular weights within the neural tube lumen and quantifies their movement out of the ventricles. This method is useful for determining differences in epithelial permeability and maturation during development and disease.

The brain ventricular system is conserved among vertebrates and is composed of a series of interconnected cavities called brain ventricles, which form ...

an-assay-for-permeability-of-the-zebrafish-embryonic-neuroepithelium

Nanyang Technological University

Research

JoVE Journal

Neuroscience

9.4K Views.  Massachusetts Institute of Technology. We describe a live whole animal quantitative measurement for permeability of the embryonic zebrafish brain. The technique analyzes the ability to retain cerebrospinal fluid and molecules of different molecular weights within the neural tube lumen and quantifies their movement out of the ventricles. This method is useful for determining differences in epithelial permeability and maturation during development and disease.

Video: An Assay for Permeability of the Zebrafish Embryonic Neuroepithelium

Establishing Embryonic Mouse Neural Stem Cell Culture Using the Neurosphere Assay

In mammalians, stem cells acts as a source of undifferentiated cells to maintain cell genesis and renewal in different tissues and organs during the life span of the animal. They can potentially replace cells that are lost in the aging process or in the process of injury and disease. The existence of neural stem cells (NSCs) was first described by Reynolds and Weiss (1992) in the adult mammalian central nervous system (CNS) using a novel serum&#8208;free culture system, the neurosphere assay (NSA). Using this assay, it is also feasible to isolate and expand NSCs from different regions of the embryonic CNS. These in vitro expanded NSCs are multipotent and can give rise to the three major cell types of the CNS. While the NSA seems relatively simple to perform, attention to the procedures demonstrated here is required in order to achieve reliable and consistent results. This video practically demonstrates NSA to generate and expand NSCs from embryonic day 14-mouse brain tissue and provides technical details so one can achieve reproducible neurosphere cultures. The procedure includes harvesting E14 mouse embryos, brain microdissection to harvest the ganglionic eminences, dissociation of the harvested tissue in NSC medium to gain a single cell suspension, and finally plating cells in NSA culture. After 5-7 days in culture, the resulting primary neurospheres are passaged to further expand the number of the NSCs for future experiments.

This video protocol demonstrates the application of the neurosphere assay for the isolation and expansion of neural stem cells from the ganglionic eminences of embryonic day 14-mouse brain.

In mammalians, stem cells acts as a source of undifferentiated cells to maintain cell genesis and renewal in different tissues and organs during the life ...

The Neuroblast Assay: An Assay for the Generation and Enrichment of Neuronal Progenitor Cells from Differentiating Neural Stem Cell Progeny Using Flow Cytometry

Neural stem cells (NSCs) can be isolated and expanded in large-scale, using the neurosphere assay and differentiated into the three major cell types of the central nervous system (CNS); namely, astrocytes, oligodendrocytes and neurons. These characteristics make neural stem and progenitor cells an invaluable renewable source of cells for in vitro studies such as drug screening, neurotoxicology and electrophysiology and also for cell replacement therapy in many neurological diseases. In practice, however, heterogeneity of NSC progeny, low production of neurons and oligodendrocytes, and predominance of astrocytes following differentiation limit their clinical applications. Here, we describe a novel methodology for the generation and subsequent purification of immature neurons from murine NSC progeny using fluorescence activated cell sorting (FACS) technology. Using this methodology, a highly enriched neuronal progenitor cell population can be achieved without any noticeable astrocyte and bona fide NSC contamination. The procedure includes differentiation of NSC progeny isolated and expanded from E14 mouse ganglionic eminences using the neurosphere assay, followed by isolation and enrichment of immature neuronal cells based on their physical (size and internal complexity) and fluorescent properties using flow cytometry technology. Overall, it takes 5-7 days to generate neurospheres and 6-8 days to differentiate NSC progeny and isolate highly purified immature neuronal cells.

This video protocol demonstrates a novel method for the generation and subsequent purification of neuronal progenitor cells from a renewable source of neural stem cells (NSCs) based on their physical (size and internal granularity) and fluorescent properties using flow cytometry technology.

Neural stem cells (NSCs) can be isolated and expanded in large-scale, using the neurosphere assay and differentiated into the three major cell types of ...

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 ...

Manual Drainage of the Zebrafish Embryonic Brain Ventricles

Cerebrospinal fluid (CSF) is a protein rich fluid contained within the brain ventricles. It is present during early vertebrate embryonic development and persists throughout life. Adult CSF is thought to cushion the brain, remove waste, and carry secreted molecules1,2. In the adult and older embryo, the majority of CSF is made by the choroid plexus, a series of highly vascularized secretory regions located adjacent to the brain ventricles3-5. In zebrafish, the choroid plexus is fully formed at 144 hours post fertilization (hpf)6. Prior to this, in both zebrafish and other vertebrate embryos including mouse, a significant amount of embryonic CSF (eCSF) is present . These data and studies in chick suggest that the neuroepithelium is secretory early in development and may be the major source of eCSF prior to choroid plexus development7.
eCSF contains about three times more protein than adult CSF, suggesting that it may have an important role during development8,9. Studies in chick and mouse demonstrate that secreted factors in the eCSF, fluid pressure, or a combination of these, are important for neurogenesis, gene expression, cell proliferation, and cell survival in the neuroepithelium10-20. Proteomic analyses of human, rat, mouse, and chick eCSF have identified many proteins that may be necessary for CSF function. These include extracellular matrix components, apolipoproteins, osmotic pressure regulating proteins, and proteins involved in cell death and proliferation21-24. However, the complex functions of the eCSF are largely unknown.
We have developed a method for removing eCSF from zebrafish brain ventricles, thus allowing for identification of eCSF components and for analysis of the eCSF requirement during development. Although more eCSF can be collected from other vertebrate systems with larger embryos, eCSF can be collected from the earliest stages of zebrafish development, and under genetic or environmental conditions that lead to abnormal brain ventricle volume or morphology. Removal and collection of eCSF allows for mass spectrometric analysis, investigation of eCSF function, and reintroduction of select factors into the ventricles to assay their function. Thus the accessibility of the early zebrafish embryo allows for detailed analysis of eCSF function during development.

We present a method to collect cerebrospinal fluid (CSF) and to create a system which lacks CSF within the embryonic zebrafish brain ventricular system. This allows for further examination of CSF composition and its requirement during embryonic brain development.

Cerebrospinal fluid  (CSF) is a protein rich fluid contained within the brain ventricles. It is  present during early vertebrate embryonic development and ...

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 ...

Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells

Mauthner cells (M-cells) are large reticulospinal neurons located in the hindbrain of teleost fish. They are key neurons involved in a characteristic behavior known as the C-start or escape response that occurs when the organism perceives a threat. The M-cell has been extensively studied in adult goldfish where it has been shown to receive a wide range of excitatory, inhibitory and neuromodulatory signals1. We have been examining M-cell activity in embryonic zebrafish in order to study aspects of synaptic development in a vertebrate preparation. In the late 1990s Ali and colleagues developed a preparation for patch clamp recording from M-cells in zebrafish embryos, in which the CNS was largely intact2,3,4. The objective at that time was to record synaptic activity from hindbrain neurons, spinal cord neurons and trunk skeletal muscle while maintaining functional synaptic connections within an intact brain-spinal cord preparation. This preparation is still used in our laboratory today. To examine the mechanisms underlying developmental synaptic plasticity, we record excitatory (AMPA and NMDA-mediated)5,6 and inhibitory (GABA and glycine) synaptic currents from developing M-cells. Importantly, this unique preparation allows us to return to the same cell (M-cell) from preparation to preparation to carefully examine synaptic plasticity and neuro-development in an embryonic organism. The benefits provided by this preparation include 1) intact, functional synaptic connections onto the M-cell, 2) relatively inexpensive preparations, 3) a large supply of readily available embryos 4) the ability to return to the same cell type (i.e. M-cell) in every preparation, so that synaptic development at the level of an individual cell can be examined from fish to fish, and 5) imaging of whole preparations due to the transparent nature of the embryos.

We have developed an intact brain-spinal cord preparation to record and monitor electrical activity via patch clamp recording from the Mauthner neurons and other reticulospinal cells in zebrafish embryos. Thus, we are able to record excitatory and inhibitory synaptic currents, voltage-gated channel activity and action potentials from key neurons in a developing embryo.

Mauthner cells (M-cells) are large  reticulospinal neurons located in the hindbrain of teleost fish. They are key  neurons involved in a characteristic ...

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 ...

An Assay for Measuring the Effects of Ethanol on the Locomotion Speed of Caenorhabditis elegans

Alcohol use disorders are a significant public health concern, for which there are few effective treatment strategies. One difficulty that has delayed the development of more effective treatments is the relative lack of understanding of the molecular underpinnings of the effects of ethanol on behavior. The nematode, Caenorhabditis elegans (C.&#160;elegans), provides a useful model in which to generate and test hypotheses about the molecular effects of ethanol. Here, we describe an assay that has been developed and used to examine the roles of particular genes and environmental factors in behavioral responses to ethanol, in which locomotion is the behavioral output. Ethanol dose-dependently causes an acute depression of crawling on an agar surface. The effects are dynamic; animals exposed to a high concentration demonstrate an initial strong depression of crawling, referred to here as initial sensitivity, and then partially recover locomotion speed despite the continued presence of the drug. This ethanol-induced behavioral plasticity is referred to here as the development of acute functional tolerance. This assay has been used to demonstrate that these two phenotypes are distinct and genetically separable. The straightforward locomotion assay described here is suitable for examining the effects of both genetic and environmental manipulations on these acute behavioral responses to ethanol in C.&#160;elegans.

An Assay for Measuring the Effects of Ethanol on the Locomotion Speed of Caenorhabditis elegans

C. elegans is a useful model for studying the effects of ethanol on behavior. We present a behavioral assay that quantifies the effects of ethanol on the locomotion speed of crawling worms; both initial sensitivity and the development of acute functional tolerance to ethanol can be measured with this assay.

Alcohol use disorders are a significant public health concern, for which there are few effective treatment strategies. One difficulty that has delayed the ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

Assay Development for High Content Quantification of Sod1 Mutant Protein Aggregate Formation in Living Cells

Amyotrophic lateral sclerosis (ALS) is a fatal neurodegenerative disease that can be caused by inherited mutations in the gene encoding copper-zinc superoxide dismutase 1 (SOD1). The structural instability of SOD1 and the detection of SOD1-positive inclusions in familial-ALS patients supports a potential causal role for misfolded and/or aggregated SOD1 in ALS pathology. In this study, we describe the development of a cell-based assay designed to quantify the dynamics of SOD1 aggregation in living cells by high content screening approaches. Using lentiviral vectors, we generated stable cell lines expressing wild-type and mutant A4V SOD1 tagged with yellow fluorescent protein and found that both proteins were expressed in the cytosol without any sign of aggregation. Interestingly, only SOD1 A4V stably expressed in HEK-293, but not in U2OS or SH-SY5Y cell lines, formed aggregates upon proteasome inhibitor treatment. We show that it is possible to quantify aggregation based on dose-response analysis of various proteasome inhibitors, and to track aggregate-formation kinetics by time-lapse microscopy. Our approach introduces the possibility of quantifying the effect of ALS mutations on the role of SOD1 in aggregate formation as well as screening for small molecules that prevent SOD1 A4V aggregation.

We describe a method to quantify the aggregation of misfolded proteins. Our protocol details lentiviral induced stable cell line generation, automated confocal imaging, and image analysis of protein aggregates. As an illustrative application, we studied the effect of small molecules in promoting SOD1 aggregation in a time- and dose-dependent manner.

Amyotrophic lateral sclerosis (ALS) is a fatal neurodegenerative disease that can be caused by inherited mutations in the gene encoding copper-zinc ...