We thank Dan McWhorter for narrating the protocol in the video. This work is supported by NIH Grant
5R01NS062182, a grant from American Federation for Aging Research, and institutional funds awarded to HTG.

I.	Procedures
The following techniques should be performed under sterile conditions, in a laminar flow hood, using sterilized tools.
Preparation of glass bottom dishes for organotypic slices
<ol>
<li>Dishes must be prepared in sterile environment and using sterilized tools.</li>
<li>A 150&mu;L drop of slice medium (see recipes) is placed in the center of the glass-bottom part of the dish with care to avoid air bubbles in the medium.</li>
<li>With a disposable syringe equipped with a 23 Gauge needle, multiple spots of glue (Rubber Cement - Elmer's, cat# E904) are placed on the square fringes adjacent to the circular coverslip that occupies the center of the culture dish, while leaving one side unglued for fluid exchange (Figure 1). The listed type of glue is not toxic to the slices or the cells as applied in this protocol. Care must be given to avoid placing any glue on the glass cover slip as this will impinge upon and obstruct imaging of the slices later. A nucleopore membrane (diameter 25mm, pore size 8.0&mu;m - Whatman, cat# 110614) is placed on top of the glass coverslip, using the glue spots to secure it in place. This should be done using a micro forceps, while ensuring that air bubbles are not trapped between the glass coverslip and the membrane.</li>
<li>Add 1 mL slice medium on top of membrane. Dishes are placed in an incubator for 30 minutes and then on ice, until ready to use.</li>
</ol>
Extraction of early postnatal brains
The best results are obtained when slices are prepared from young postnatal mice (P1-P10).
<ol>
<li>Pups are terminally anesthetized (overdosed) by isofluorane or other approved methods. The head can be sprayed with 70% ethanol to increase sterility, followed by rapid decapitation using sharp scissors.</li>
<li>The head is stabilized by clamping the mandible with micro forceps. The skin is excised longitudinally from the neck to the snout. The skull is cut longitudinally and anteriorly starting at the cisterna magna, by making 1 medial and 2 lateral cuts (one on each side - Figure 2A). Care should be taken to minimize contact with underlying cortical tissue as the cranial flaps are removed away from the brain.</li>
<li>To improve stability of the tissue during vibratome sectioning, the lateral-most aspects of the brain are removed by making two sagittal cuts. The caudal aspect of the brain is also removed by making a cut at the rostral base of the cerebellum (Figure 2B).</li>
<li>The two hemispheres are separated by making a smooth cut along the midline fissure, and the two hemispheres are carefully scooped out of the skull and placed, medial surface down, in an embedding mold (Figs. 2C-D).</li>
</ol>
Sectioning of the host brain
<ol>
<li>The two hemispheres in the embedding mold are immediately covered with melted 3% low melting point DNA-grade agarose gel (Fisher, cat# BP1360-100) dissolved in tissue preparation buffer that is maintained at 37&deg;C (see recipes). After 2 minutes of stabilization on a flat horizontal surface to ensure even hardening of the agarose, the molds are positioned on ice to complete the setting.</li>
<li>Once the gel containing the hemispheres is set, it is removed from the mold and trimmed, leaving 2-3mm of gel around the brain tissue.</li>
<li>The gel-embedded tissue is then mounted on the specimen disc of the vibratome, with the medial surface up, and secured with cyanoacrylate adhesive (Krazy glue or equivalent). Care must be taken to apply only minimal amounts of glue, as too much will have toxic effects on slices and migrating cells. Too much glue on the sides of the block will also impede the cutting, causing potential damage in the tissue.</li>
<li>The disc is installed in the vibratome specimen tray filled with ice-cold tissue preparation medium.</li>
<li>The tissue is sectioned at 150&mu;m thickness, with the vibratome speed set on a slow-to-medium range (will depend on the vibratome used, and thus must be determined independently for optimal results). In our hands, the vibration frequency is optimal when set at maximum. The first few slices may be discarded.</li>
<li>As soon as the RMS-containing slices are released from the blade (the RMS is visible by the naked eye as a grey U-shaped structure extending from the SEZ to the OB), they are carefully teased out of the gel mold, and scooped using a small flathead spatula. The slices are placed on the nucleopore membrane of the glass-bottom dishes containing slice medium (Figure 3A-B). A typical early postnatal mouse brain will yield approximately 2-3 RMS sections per hemisphere when cut at 150 &mu;m. It is critical that handling the slices is kept to a minimum as they are very fragile. The dishes with the slices are then transferred to an incubator.</li>
</ol>
Donor brain sectioning and RMS transplantation
<ol>
<li>Donor brains (brains expressing fluorescent reporters in migrating neuroblasts) are sectioned at 250&mu;m thickness and slices are collected in ice-cold tissue preparation buffer.</li>
<li>Slices are immediately placed under a dissecting microscope with epifluorescence capability, and the RMS is gently microdissected, using micro forceps. One forceps is used to stabilize the slice while the other is used to make small cuts around the RMS until it is released from the slice. The excised RMS is then cut into small explants (approximately 200-500 &mu;m in diameter) prior to transplantation.</li>
<li>Glass-bottom dishes containing host slices positioned on the nucleopore filters are removed from the incubator and placed under the dissecting microscope. Using visible light, the RMS is clearly identified and a small incision is made in the initial segment of the RMS.</li>
<li>Using a pipettor equipped with a 20 &mu;L tip, a single donor RMS explant is transferred to the incised site in the host RMS. The explant is gently pushed into the incision to establish contact between the 2 tissues. To ensure this contact is stable, the explants are pushed slightly in between the slice and the nucleopore membrane.</li>
<li>Once all host slices with the RMS are transplanted, the dishes are returned to the incubator for at least 1 hour to allow the sections to settle on the membrane. Neuroblasts should begin to migrate from the explant into the host RMS after approximately 1-2 hours.</li>
</ol>
Time-lapse imaging of neuronal migration
<ol>
<li>The glass-bottom dishes are transferred from the incubator to the incubated chamber on the microscope (Figure 3C). Fluorescent neuroblasts can be imaged at intervals ranging from 0 to 10 minutes depending on the type of analysis desired. For example we image cytoskeletal dynamics during the migratory cycle of individual neuroblasts by setting our intervals at less than or equal to 2.5 minutes. However, population dynamics such as orientation and speed of migration are better captured at intervals ranging from 5 to 10 minutes. The choice of objectives is highly variable depending on the brand of microscope. For a Nikon C1 confocal microscope, we find that the 20x dry lens (Nikon Pan Fluor, N.A. 0.75, W.D. 0.35mm) is most suitable for our analyses. For best results on this confocal system, the pinhole is opened to a medium size (60&mu;m diameter). For most appropriate migratory behavior, imaging is confined to cells deep within the thickness of the RMS, at least 20 &mu;m away from either of the cut surfaces of the slice. Laser power should be optimized so that it is at a minimum, but that the details of individual neuroblasts remain visible.</li>
<li>Once the imaging is completed slices can be fixed with ice cold, and freshly prepared 4% paraformaldehyde, immunostained, and mounted on slides for further imaging. Since we do not coat our membranes with any adherent substrates such as laminin, the slices usually float away from the membrane once they are immersed in the staining buffers. Sections are imaged while maintaining their 150 &mu;m thickness. It is also possible to cryopreserve and freeze the slices and use a cryostat to obtain thinner sections of the original slices. However, this will result in increase the incidence of artifacts in staining as well as alter the integrity of the tissue.</li>
</ol>
II.	 Materials/equipment
Preparation of glass bottom dishes for organotypic slices
<ul>
<li>Small syringes (1 mL)</li>
<li>23-gauge needles</li>
<li>Nucleopore Track-Etch Membrane - diameter 25mm, pore size 8.0&mu;m - Whatman, cat# 110614</li>
<li>Glass Bottom Culture Dishes - 35mm petri dish, 14mm Microwell, No. 1.5 coverglass - MatTech, cat# P35G-1.5-14-C</li>
<li>Rubber Cement - Elmer's, cat# E904</li>
<li>Basal Medium Eagle - Gibco, cat# 21010</li>
<li>1M Hepes (pH7.4)</li>
<li>1M D-glucose</li>
<li>100mM CaCl2</li>
<li>100mM MgSO4</li>
<li>1M NaHCO3</li>
<li>dH2O</li>
<li>200mM L-glutamine</li>
<li>Penicillin-streptomycin</li>
</ul>
Brain extraction and embedding
<ul>
<li>Anesthetic (isofluorane, etc.)</li>
<li>Microwave oven</li>
<li>Low Melting Agarose - Fisher, cat# BP1360-100</li>
<li>Krazy glue - cat#KG585</li>
<li>Peel-A-Way Disposable Embedding Molds (R-40) - 22mmx40mm rectangular, 20mm deep - Polysciences, cat# 18646C</li>
</ul>
Brain sectioning and RMS transplantation
<ul>
<li>Vibratome - Leica VT1000S and all accessory components for slice preparation</li>
<li>10X Hank's Balanced Salt Solution - Gibco, cat# 14185</li>
<li>Microdissecting forceps #5 - Roboz, cat# RS-4976</li>
<li>Microspatula - Fisher, cat# 21-401-15</li>
<li>Stereomicroscope</li>
</ul>
Time-lapse imaging of organotypic slices
<ul>
<li>Humidified Incubator, 5% CO2</li>
<li>Inverted Microscope equipped with incubator chamber and long distance working objectives (NA of 0.6 or higher)</li>
</ul>
III.	Recipes
Buffer solution for tissue dissection and slice preparation (tissue preparation buffer)
<table border="1">
<tbody>
 <tr>
 <td>Stock solution</td>
 <td>Volume</td>
 <td>Final Concetration</td>
 </tr>
 <tr>
 <td>10X HBSS</td>
 <td>50 mL</td>
 <td>1X</td>
 </tr>
 <tr>
 <td>1M Hepes (pH 7.4)</td>
 <td>1.25 mL</td>
 <td>2.5mM</td>
 </tr>
 <tr>
 <td>1M D-Glucose</td>
 <td>15 mL</td>
 <td>30mM</td>
 </tr>
 <tr>
 <td>1M CaCl2</td>
 <td>0.5 mL</td>
 <td>1mM</td>
 </tr>
 <tr>
 <td>1M MgSO4</td>
 <td>0.5 mL</td>
 <td>1mM</td>
 </tr>
 <tr>
 <td>1M NaHCO3</td>
 <td>2 mL</td>
 <td>4mM</td>
 </tr>
 <tr>
 <td>dH2O</td>
 <td>430.75 mL</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>
Filter sterilize with a 0.2 &mu;m filter and store at 4&deg;C.

Culture medium for organotypic slices, tissue transplantation and imaging (slice medium)
<table border="1">
<tbody>
 <tr>
 <td>Stock solution</td>
 <td>Volume</td>
 <td>Final Concentration</td>
 </tr>
 <tr>
 <td>Basal Medium Eagle</td>
 <td>35 mL</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tissue preparation buffer</td>
 <td>12.9 mL</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>1M D-Glucose</td>
 <td>1.35 mL</td>
 <td>20mM</td>
 </tr>
 <tr>
 <td>200mM L-glutamine</td>
 <td>0.25 mL</td>
 <td>1mM</td>
 </tr>
 <tr>
 <td>Penicillin-streptomycin</td>
 <td>0.5 mL</td>
 <td>100units/ mL penicillin and 0.1mg/ mL streptomycin</td>
 </tr>
 </tbody>
</table>

Filter sterilize with a 0.2 &mu;m filter and store at 4&deg;C.

Preparation of Low-melting point agarose gel
Low-melting-point agarose is diluted in tissue preparation buffer at 0.3g/ mL in a 50 mL conical tube (see recipes). The tube is microwaved in increments of 5-10 seconds at high-power. Number of increments is dependent on total volume; for 10 mL, three increments (10-8-5 seconds, each) should suffice. The tube cap is carefully unscrewed between heating increments to release air pressure and avoid explosion of the tube. Caution must be taken as the tube content will be very hot. Once the agarose is completely dissolved, the tube is maintained in a 37&deg;C water bath for at least 5 minutes to allow the temperature to stabilize prior to use. Prolonged exposure to room temperature will harden the gel. Although this must be avoided as much as possible, hardened gel can be reheated and re-melted for immediate use within 24 hours of initial preparation.
Immunohistochemistry on organotypic slices
Following imaging on the confocal microscope, slices may be fixed overnight at 4&deg;C with 4% formaldehyde in PBS. Sections are then blocked overnight at 4&deg;C, in 10% goat serum with 1% Triton X (Sigma, Cat. # S26-36-23) in PBS followed by overnight incubation with primary antibodies at 4&deg;C. Fluorescently-tagged goat secondary antibodies are used for visualization (all diluted 1:1000, 1 hour incubation at room temperature). Labeled slices are thoroughly washed 5-6 times with ice cold PBS prior to mounting on glass slides and coverslipping.
IV. Representative Results
Our organotypic slice culture protocol has been thoroughly tested and optimized over that last few years for consistency in migration pattern and orientation. Analysis of cells emigrating from explants obtained from mice in which the expression of the red fluorescent protein, Td-tomato, is induced under the Nestin promoter (Nestin-Td tomato), reveals highly oriented and rapid migration of tdTomato+ neuroblasts into the host RMS (Figure 4A). High-magnification time-lapse analysis illustrates excellent resolution of the entire length of a migrating neuroblast during a 20-minute imaging session (Figure 4B).
Slices with donor Td-tomato+ cells were fixed and immunostained for different cellular components within the RMS. GFAP+ astrocytes and CD31+ blood vessels were revealed using fluorescent immunohistochemistry (Figure 5A). High-magnification analysis of slices stained for the cytoskeletal proteins actin and tubulin reveal non-uniform expression of these components by a cell in the midst of migration (Figure 5B).
Antibodies used in these examples: rabbit anti-RFP (Abcam, 1:250), rabbit anti-GFAP (Dako, 1:1000), rat anti-CD31(BD Pharmigen, 1:100), mouse anti-actin (Santa Cruz, 1:500), rabbit anti-tubulin (Sigma, 1:1000), goat anti-mouse Cy3 (Chemicon, 1:1000), goat anti-rabbit AlexaFluor 647 (Invitrogen, 1:1000), goat anti-rat AlexaFluor 488 (Invitrogen, 1:1000), goat anti-rabbit AlexaFluor 488 (Invitrogen, 1:1000).
<img src="/files/ftp_upload/2486/2486fig1.jpg" alt="Figure 1" /> 
Figure 1. Preparation of glass-bottom dishes for organotypic slices. Multiple spots of glue are placed around the circular glass-bottom component of the dish (red), leaving one side open for exchange of medium from underneath the filter. A 150&mu;L drop of slice medium is placed in the center of the glass coverslip. A nucleopore membrane (blue) is then applied, shiny side down, while ensuring that no air bubbles are trapped between the glass coverslip and the membrane. One milliliter of slice medium (grey) is spread on top of membrane, and dishes are incubated at 37&deg; C prior to use.
<img src="/files/ftp_upload/2486/2486fig2.jpg" alt="Figure 2" /> 
Figure 2. Brain extraction and preparation for sectioning. (A) The skull is exposed by incising the scalp from the neck to the snout (dotted line along the midline). The skull is then cut longitudinally and anteriorly starting at the cisterna magna, by making 1 medial and 2 lateral cuts (one on each side; 2A). (B) The lateral-most aspects of the cortex and the caudal aspect of the CNS are resected to improve stability of the tissue during vibratome sectioning. (C-D) The two hemispheres are then separated and placed medial face down in an embedding mold prior to application of 3% agarose gel dissolved in tissue preparation buffer.
<img src="/files/ftp_upload/2486/2486fig3.jpg" alt="Figure 3" /> 
Figure 3. Brain sectioning and cross-transplantation. (A) The host tissue is sectioned at 150&mu;m thickness, and the RMS-containing sections are carefully positioned flat on the nucleopore membrane of cold glass-bottom dishes. (B) Donor brains (brains expressing fluorescent reporters in the RMS) are sectioned at 250&mu;m thickness, and the slices are collected in ice-cold tissue preparation buffer. The donor RMS is microdissected and cut into small explants. Using a pipettor equipped with a 20 &mu;L tip, individual RMS explants are transferred to an incised site in the host RMS. (C) After 1-2 hours of incubation, dishes are transferred to an incubated stage on a confocal microscope and migration is captured using time-lapse imaging. The photomicrograph is a representative low-magnification image of a typical slice (grey) set up 1 hour after transplantation (red explant from tdTomato+ RMS of a donor mouse; red dotted lines outline the RMS in the host slice).
<img src="/files/ftp_upload/2486/2486fig4.jpg" alt="Figure 4" /> 
Figure 4. Migration of neuroblasts from explants into the host RMS. (A) Nestin-tdTomato+ neuroblasts (red) migrate from the explants into the host RMS (dotted green line) 1 hour after transplantation. tdTomato+ cells invading the RMS of the host organotypic slices move in a highly oriented and rapid manner away from the SEZ and toward the OB. (B) The migratory cycle can be observed in high-power time-lapse images of a neuroblast over approximately a 20 minute period. Scale bar = 10 &mu;m.
<img src="/files/ftp_upload/2486/2486fig5.jpg" alt="Figure 5" /> 
Figure 5. Immunohistochemical assessment of organotypic slices. Explanted neuroblasts (red) were fixed in the midst of migration 12 hours post transplantation. (A) Fluorescent immunohistochemical staining of the slice reveals a dense pool of GFAP+ astrocytes (blue) and scattered CD31+ blood vessels (green) within the RMS of the host slice. (B) The cytoskeleton of an isolated tdTomato+ migrating cell (red) in a host RMS is revealed by co-immunostaining using antibodies against actin (blue) and tubulin (green).

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Experiments on animals were performed in accordance with the guidelines and regulations set forth by North Carolina State University and its College of Veterinary Medicine.
No conflicts of interest declared.

Neuronal migration in the RMS is an essential component of postnatal neurogenesis in the olfactory bulbs 1. Migration through the RMS occurs in a plane tangential to the surface of the brain. Tangentially migrating neuroblasts are distinct from radially migrating cells based on the location of their progenitor source, as well as the divergent fate of their final neuronal products 1, 2, 3. The relatively pure population of tangentially migrating cells in the postnatal RMS makes this anatomically de...

an organotypic slice assay for high-resolution time-lapse imaging of neuronal migration in the postnatal brain

Neurogenesis in the postnatal brain depends on maintenance of three biological events: proliferation of progenitor cells, migration of neuroblasts, as well as differentiation and integration of new neurons into existing neural circuits. For postnatal neurogenesis in the olfactory bulbs, these events are segregated within three anatomically distinct domains: proliferation largely occurs in the subependymal zone (SEZ) of the lateral ventricles, migrating neuroblasts traverse through the rostral migratory stream (RMS), and new neurons differentiate and integrate within the olfactory bulbs (OB). The three domains serve as ideal platforms to study the cellular, molecular, and physiological mechanisms that regulate each of the biological events distinctly. This paper describes an organotypic slice assay optimized for postnatal brain tissue, in which the extracellular conditions closely mimic the in vivo environment for migrating neuroblasts. We show that our assay provides for uniform, oriented, and speedy movement of neuroblasts within the RMS. This assay will be highly suitable for the study of cell autonomous and non-autonomous regulation of neuronal migration by utilizing cross-transplantation approaches from mice on different genetic backgrounds.

Watch this Scientific Journal Video about An Organotypic Slice Assay for High-Resolution Time-Lapse Imaging of Neuronal Migration in the Postnatal Brain at JoVE.com

An Organotypic Slice Assay for High-Resolution Time-Lapse Imaging of Neuronal Migration in the Postnatal Brain

This protocol describes an organotypic slice assay optimized for the postnatal brain and high-resolution time-lapse imaging of neuroblast migration in the rostral migratory stream.

Neurogenesis in the postnatal brain depends on maintenance of three biological events: proliferation of progenitor cells, migration of neuroblasts, as ...

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Research

JoVE Journal

Neuroscience

11.8K Views.  North Carolina State University. Этот протокол описывает органотипической анализа ломтик оптимизирован для послеродового мозга и высоким разрешением, замедленная съемка нейробластов миграции в ростральной миграционных потоков.

Video: An Organotypic Slice Assay for High-Resolution Time-Lapse Imaging of Neuronal Migration in the Postnatal Brain

The Organotypic Hippocampal Slice Culture Model for Examining Neuronal Injury

Organotypic hippocampal slice culture is an in vitro method to examine mechanisms of neuronal injury in which the basic architecture and composition of the hippocampus is relatively preserved 1. The organotypic culture system allows for the examination of neuronal, astrocytic and microglial effects, but as an ex vivo preparation, does not address effects of blood flow, or recruitment of peripheral inflammatory cells. To that end, this culture method is frequently used to examine excitotoxic and hypoxic injury to pyramidal neurons of the hippocampus, but has also been used to examine the inflammatory response. Herein we describe the methods for generating hippocampal slice cultures from postnatal rodent brain, administering toxic stimuli to induce neuronal injury, and assaying and quantifying hippocampal neuronal death.

The organoptypic hippocampal slice culture model is an in vitro model used to examine neuronal injury in a variety of paradigms. In this article, we describe the methods for generating slice cultures and quantifying neuronal injury.

Organotypic hippocampal slice culture is an in vitro method to examine mechanisms of neuronal injury in which the basic architecture and composition of ...

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 ...

Multi-electrode Array Recordings of Neuronal Avalanches in Organotypic Cultures

The cortex is spontaneously active, even in the absence of any particular input or motor output. During development, this activity is important for the migration and differentiation of cortex cell types and the formation of neuronal connections1. In the mature animal, ongoing activity reflects the past and the present state of an animal into which sensory stimuli are seamlessly integrated to compute future actions. Thus, a clear understanding of the organization of ongoing i.e. spontaneous activity is a prerequisite to understand cortex function. 
Numerous recording techniques revealed that ongoing activity in cortex is comprised of many neurons whose individual activities transiently sum to larger events that can be detected in the local field potential (LFP) with extracellular microelectrodes, or in the electroencephalogram (EEG), the magnetoencephalogram (MEG), and the BOLD signal from functional magnetic resonance imaging (fMRI). The LFP is currently the method of choice when studying neuronal population activity with high temporal and spatial resolution at the mesoscopic scale (several thousands of neurons). At the extracellular microelectrode, locally synchronized activities of spatially neighbored neurons result in rapid deflections in the LFP up to several hundreds of microvolts. When using an array of microelectrodes, the organizations of such deflections can be conveniently monitored in space and time. 
Neuronal avalanches describe the scale-invariant spatiotemporal organization of ongoing neuronal activity in the brain2,3. They are specific to the superficial layers of cortex as established in vitro4,5, in vivo in the anesthetized rat 6, and in the awake monkey7. Importantly, both theoretical and empirical studies2,8-10 suggest that neuronal avalanches indicate an exquisitely balanced critical state dynamics of cortex that optimizes information transfer and information processing.
In order to study the mechanisms of neuronal avalanche development, maintenance, and regulation, in vitro preparations are highly beneficial, as they allow for stable recordings of avalanche activity under precisely controlled conditions. The current protocol describes how to study neuronal avalanches in vitro by taking advantage of superficial layer development in organotypic cortex cultures, i.e. slice cultures, grown on planar, integrated microelectrode arrays (MEA; see also 11-14).

A robust way to study neuronal avalanches, i.e. scale-invariant spatio-temporal activity bursts, indicative of critical state dynamics in cortex. Avalanches emerge spontaneously in developing superficial layers of cultured cortex which allows for long-term measurements of the activity with planar integrated multi-electrode arrays (MEA) under precisely controlled conditions.

The cortex is spontaneously active, even in the absence of any particular input or motor output.  During development, this activity is important for the ...

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

High-resolution Functional Magnetic Resonance Imaging Methods for Human Midbrain

Functional MRI (fMRI) is a widely used tool for non-invasively measuring correlates of human brain activity. However, its use has mostly been focused upon measuring activity on the surface of cerebral cortex rather than in subcortical regions such as midbrain and brainstem. Subcortical fMRI must overcome two challenges: spatial resolution and physiological noise. Here we describe an optimized set of techniques developed to perform high-resolution fMRI in human SC, a structure on the dorsal surface of the midbrain; the methods can also be used to image other brainstem and subcortical structures.
High-resolution (1.2 mm voxels) fMRI of the SC requires a non-conventional approach. The desired spatial sampling is obtained using a multi-shot (interleaved) spiral acquisition1. Since, T2* of SC tissue is longer than in cortex, a correspondingly longer echo time (TE ~ 40 msec) is used to maximize functional contrast. To cover the full extent of the SC, 8-10 slices are obtained. For each session a structural anatomy with the same slice prescription as the fMRI is also obtained, which is used to align the functional data to a high-resolution reference volume.
In a separate session, for each subject, we create a high-resolution (0.7 mm sampling) reference volume using a T1-weighted sequence that gives good tissue contrast. In the reference volume, the midbrain region is segmented using the ITK-SNAP software application2. This segmentation is used to create a 3D surface representation of the midbrain that is both smooth and accurate3. The surface vertices and normals are used to create a map of depth from the midbrain surface within the tissue4.
Functional data is transformed into the coordinate system of the segmented reference volume. Depth associations of the voxels enable the averaging of fMRI time series data within specified depth ranges to improve signal quality. Data is rendered on the 3D surface for visualization.
In our lab we use this technique for measuring topographic maps of visual stimulation and covert and overt visual attention within the SC1. As an example, we demonstrate the topographic representation of polar angle to visual stimulation in SC.

This article describes techniques to perform high-resolution functional magnetic resonance imaging with 1.2 mm sampling in human midbrain and subcortical structures using a 3T scanner. Use of these techniques to resolve topographic maps of visual stimulation in the human superior colliculus (SC) is given as an example.

Functional MRI (fMRI) is a widely used tool for non-invasively measuring correlates of human brain activity. However, its use has mostly been focused upon ...

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

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

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

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

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to neuroblasts able to move along the rostral migratory stream (RMS) towards the olfactory bulb (OB). This long-distance migration is required for the subsequent maturation of newborn neurons in the OB, but the molecular mechanisms regulating this process are still unclear. Investigating the signaling pathways controlling neuroblast motility may not only help understand a fundamental step in neurogenesis, but also have therapeutic regenerative potential, given the ability of these neuroblasts to target brain sites affected by injury, stroke, or degeneration.
In this manuscript we describe a detailed protocol for in vivo postnatal electroporation and subsequent time-lapse imaging of neuroblast migration in the mouse RMS. Postnatal electroporation can efficiently transfect SVZ progenitor cells, which in turn generate neuroblasts migrating along the RMS. Using confocal spinning disk time-lapse microscopy on acute brain slice cultures, neuroblast migration can be monitored in an environment closely resembling the in vivo condition. Moreover, neuroblast motility can be tracked and quantitatively analyzed. As an example, we describe how to use in vivo postnatal electroporation of a GFP-expressing plasmid to label and visualize neuroblasts migrating along the RMS. Electroporation of shRNA or CRE recombinase-expressing plasmids in conditional knockout mice employing the LoxP system can also be used to target genes of interest. Pharmacological manipulation of acute brain slice cultures can be performed to investigate the role of different signaling molecules in neuroblast migration. By coupling in vivo electroporation with time-lapse imaging, we hope to understand the molecular mechanisms controlling neuroblast motility and contribute to the development of novel approaches to promote brain repair.

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

Neuroblast migration is a fundamental event in postnatal neurogenesis. We describe a protocol for efficient labeling of neuroblasts by in vivo postnatal electroporation and subsequent visualization of their migration using time-lapse imaging of acute brain slices. We include a description for the quantitative analysis of neuroblast dynamics by video tracking.

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It has helped to describe the different steps of radial migration and characterize the molecular mechanisms controlling this process. To directly and dynamically analyze migrating neurons they have to be traced over time. This protocol describes a workflow that combines in utero electroporation with organotypic slice culture and time-lapse confocal imaging, which allows for a direct examination and dynamic analysis of radially migrating cortical neurons. Furthermore, detailed characterization of migrating neurons, such as migration speed, speed profiles, as well as radial orientation changes, is possible. The method can easily be adapted to perform functional analyses of genes of interest in radially migrating cortical neurons by loss and gain of function as well as rescue experiments. Time-lapse imaging of migrating neurons is a state-of-the-art technique that once established is a potent tool to study the development of the cerebral cortex in mouse models of neuronal migration disorders.

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

This protocol provides instructions for direct observation of radially migrating cortical neurons. In utero electroporation, organotypic slice culture, and time-lapse confocal imaging are combined to directly and dynamically study the effects of overexpression or downregulation of genes of interest in migrating neurons and to analyze their differentiation during development.

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It ...

High-Resolution 3D Imaging of Rabies Virus Infection in Solvent-Cleared Brain Tissue

The visualization of infection processes in tissues and organs by immunolabeling is a key method in modern infection biology. The ability to observe and study the distribution, tropism, and abundance of pathogens inside of organ tissues provides pivotal data on disease development and progression. Using conventional microscopy methods, immunolabeling is mostly restricted to thin sections obtained from paraffin-embedded or frozen samples. However, the limited 2D image plane of these thin sections may lead to the loss of crucial information on the complex structure of an infected organ and the cellular context of the infection. Modern multicolor, immunostaining-compatible tissue clearing techniques now provide a relatively fast and inexpensive way to study high-volume 3D image stacks of virus-infected organ tissue. By exposing the tissue to organic solvents, it becomes optically transparent. This matches the sample&#8217;s refractive indices and eventually leads to a significant reduction of light scattering. Thus, in combination with long free working distance objectives, large tissue sections up to 1 mm in size can be imaged by conventional confocal laser scanning microscopy (CLSM) at high resolution. Here, we describe a protocol to apply deep-tissue imaging after tissue clearing to visualize rabies virus distribution in infected brains in order to study topics like virus pathogenesis, spread, tropism, and neuroinvasion.

Novel, immunostaining-compatible tissue clearing techniques like the ultimate 3D imaging of solvent-cleared organs allow the 3D visualization of rabies virus brain infection and its complex cellular environment. Thick, antibody-labeled brain tissue slices are made optically transparent to increase imaging depth and to enable 3D analysis by confocal laser scanning microscopy.

The visualization of infection processes in tissues and organs by immunolabeling is a key method in modern infection biology. The ability to observe and ...