We would like to thank Erica Lorenzetti for technical assistance on the injections and Irene Costantini for making figure 1. The research leading to these results has received funding from LASERLABEUROPE (Grant 284464, European Commission&#8217;s Seventh Framework Programme). This research project has also been supported by the Italian Ministry for Education, University and Research in the framework of the Flagship Project NANOMAX and by Italian Ministry of Health in the framework of the &#8220;Stem Cells Call for Proposals.&#8221; This work is part of the research activities of the European Flagship Human Brain Project and has been carried out in the framework of the International Center of Computational Neurophotonics foundation supported by &#8220;Ente Cassa di Risparmio di Firenze&#8221;.

1. Axonal Labeling
<ol>
	<li>Climbing fibers can be labeled by injecting either organic dyes conjugated to high molecular weight dextrans or plasmid/viruses that induce the expression of fluorescent proteins26-29. In this protocol, the organic dye Alexa Fluor Dextran 488 is injected into the inferior olive to label climbing fibers and visualize them in the cerebellar cortex (Figure 1). All the procedures described here have been approved by the Italian Ministry of Health.</li>
	<li>Prepare the glass capillary tube by pulling it on a micropipette puller. Trim the tip of the glass capillary tube with scissors till the external diameter is about 30 &#181;m.</li>
	<li>Load the capillary with 1-2 &#181;l of the dextran-conjugated dye.</li>
	<li>Before starting, sterilize all the surgical instruments in an autoclave. During the procedure, reduce risks of contamination by using a glass bead sterilizer.</li>
	<li>Anesthetize the mouse (9-12 weeks) by intraperitoneal injection of ketamine (90 mg/kg) and xylazine (9 mg/kg). Ensure the animal is fully sedated using tail and/or toe pinches. Use vet ointment on eyes to prevent dryness while under anesthesia.</li>
	<li>Shave the hairs above the neck with either a razor blade or hair removal cream. Swab the surgical site twice with betadine, alternated with ethanol 70%.</li>
	<li>Place the mouse on a stereotaxic holder. Keep the animal warm with a heating pad.</li>
	<li>Gently push the earbars on the bone next to the ears so that the head is held firmly. Rotate the head to form a 140&#176; angle with the body. The desired orientation can be obtained by gently changing the height of the nose holder rod on the stereotactic holder.</li>
	<li>Apply some drops of lidocaine on the skin between the ears. Use forceps or a razor blade to perform a vertical incision of about 1 cm on the skin below the occiput.</li>
	<li>Gently separate the subcutaneous tissue and muscles by blunt tweezers under the dissection microscope. Push down the horizontal muscle bundle and hold apart the two vertical muscles fascicles in order to expose the dura over the foramen magnum.</li>
	<li>Using extreme caution, dissect the dura with a syringe needle or very sharp forceps to expose the brainstem. A small quantity of cerebro-spinal fluid may spill out when the dura has been cut.</li>
	<li>On the stereotactic micromanipulator, rotate the holder of the capillary 45&#176; from the vertical. Place the capillary at the midline, at the midpoint between the caudal edge of the cerebellar cortex and the first cervical vertebra. Insert the pipette at a depth of 2 mm from the surface.</li>
	<li>Deliver a 0.5-1.5 &#956;l volume of dye over 10-15 min. A microinjection dispensing system allows controlled pressure delivery of the liquid in the capillary (Pulse pressure = 20 psi; pulse duration = 3-4 msec; frequency of pulse delivery = 12 Hz).</li>
	<li>Leave the capillary in place for 15 min, then slowly (in 2-3 min) remove it. Delicately re-align the muscles and suture the skin above. Keep the muscles well hydrated by applying some saline throughout the whole procedure.</li>
	<li>To avoid dehydration, inject subcutaneously 0.5 ml of 0.9% NaCl. Keep the animal in a heated cage until full recovery from anesthesia.</li>
	<li>Subcutaneously administer Carprofen (5 mg/kg) daily for 2-3 days after the surgery.</li>
</ol>
2. Optical Window on the Cerebellar Cortex
NOTE: The dye is transported in 2 weeks from the inferior olivary nucleus up to the cerebellar cortex to the CFs terminals (see Figure 1). The sparsely labeled CFs can be visualized under the permanent cranial window&#160;that can be performed as follows:
<ol>
	<li>Sterilize all the surgical instruments.</li>
	<li>Anesthetize the mouse as described above. Use toe pinches to check the animal is fully sedated.</li>
	<li>Shave the hair above the head, wipe the skin with alternating swipes of 70% alcohol and betadine and apply eye ointment. Place the mouse in a stereotaxic frame and position the ear bars in order to firmly hold the head.</li>
	<li>Subcutaneously administer Dexamethasone (0.2 mg/kg) and Carprofen (5 mg/kg) to prevent swelling of the brain and possible inflammations at the cranial window site.</li>
	<li>Apply a drop of lidocaine on the skin above the skull and cut a flap, from between the ears to above the eyes. Apply lidocaine solution again onto the periosteum before scraping it with a scalpel.</li>
	<li>Use the dental drill to thin a semicircular region of skull above the cerebellum. The window is usually placed central and towards the back suture, so that the skull depth is almost constant throughout the perimeter of the semicircle. Positioning the window too close to the lambda suture and repetitive drilling over this very thick region may result in overheating of the brain and formation of edemas.</li>
	<li>Apply hemostatic sponge on the skull in case of bleeding.</li>
	<li>Cut a coverglass in two halves with the help of a razor blade and tweezers. Check that the coverglass is slightly bigger than the isle of bone and has a similar shape before removing the flap of bone with the tweezers; if not, reshape the coverglass or choose another one. Do not touch the dura mater with the tweezers while drawing off the bone. Carefully lay down the coverglass upon the dura. NOTE: The adherence of the whole surface of the glass on the dura is foretelling of long lasting cranial windows.</li>
	<li>Remove air bubbles under the glass by repeatedly rinsing with saline solution and drying with cotton sticks. This will also help removing some residual blood that might be spilled out from small capillaries after positioning the glass. Seal the window with a mixture of acrylic glue and dental cement30.</li>
	<li>Keep the animal on a heating pad in a recovery chamber until fully recovered from anesthesia. Do not return mice in the home cage until fully recovered; do not leave them unattended until they regained sufficient consciousness to maintain sternal recumbency.</li>
	<li>Subcutaneously administer Carprofen (5 mg/kg) daily for 2-3 days after the surgery.</li>
</ol>
3. In Vivo Multi-photon Laser Axotomy
<ol>
	<li>Wait 2-7 days from the surgery before performing the first imaging session. The custom built two-photon microscope used here is provided with a Ti:Sapphire laser source (120 fs width pulses, 90 MHz repetition rate) which is first scanned by a pair of galvanometric mirrors and then focused onto the specimen by a water immersion 20X objective (NA 0.95, WD 2 mm). A closed-loop piezoelectric stage performs axial displacements up to 400 &#956;m of the objective. Photomultiplier tubes collect the fluorescence signal.</li>
	<li>Anesthetize the animal, then position the head in the stereotactic holder so that the cranial window lays parallel to the objective focal plane.</li>
	<li>Explore with TPF imaging the cortex under the cranial window. Choose the axon to be damaged amongst the brightest ones.</li>
	<li>Acquire a Z-stack (usually the field of view is 100 x&#160;100 &#181;m2, at 512 x&#160;512 pixels, 2 &#181;m z-axis step) to obtain a 3D-reconstructions of the targeted axon. Within the imaged volume, choose the lesion site on an axonal branch that lies mostly parallel to the focal plane. This orientation allows to rapidly score morphological changes, which are predictive of a successful dissection.</li>
	<li>Irradiate with a high energy dose the selected point on a distal portion of a CF. This is done through the Labview software: increase the laser power 5-10x&#160;more than the power used for imaging (&#8776; 300 mW, measured after the objective lens), set the position of the galvanometric mirrors to do a line scan (~4-10 &#181;m) on the lesion site and open the laser shutter for 400 - 600 msec. Orientate the line scan perpendicular to the CF plane to completely transect the axon. Irradiating brighter spots (e.g., varicosities) usually helps dissecting the axon. The wavelength for laser axotomy is the same used for imaging (920 nm).</li>
	<li>Usually the fluorescence of the axon in the irradiated area transiently decreases due to photobleaching. Acquire a stack of the same region few minutes after the irradiation to ensure the ablation has been well performed. If the region started swelling and the axon formed bead-like structures, the neuron has been successfully dissected. In this case, in a few hours (from few tens of min to 24 hr) the distal portion of the axon will degenerate and disappear completely.</li>
	<li>Unfortunately the energy dose necessary to produce a complete axotomy is not always predictable. If the only outcome of the irradiation is photobleaching, the fluorescence should be rapidly recovered due to dye diffusion, and the irradiation should be repeated again to effectively dissect the axon. Also, in case signs of swelling do not appear within 5 - 20 min, or the swelling is followed by recovery (analogously to what has been described for laser dendrotomy, see Allegra Mascaro et al.23), try again with new scans increasing the dwell time and/or the laser power.</li>
	<li>Monitor the eventual remodeling of the axon in the days that follow the laser axotomy with two-photon in vivo imaging (Figures 2 and 3). The vascular pattern is used as reference pattern to retrieve the dissected axon in the following days.</li>
	<li>Subcutaneously administer Carprofen (5 mg/kg) every imaging session for the whole duration of the experiment.</li>
</ol>

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(12), (2008).</li></ol>

We have nothing to disclose.

This protocol shows how to label neurons of the inferior olive with a fluorescent dye. Subsequently, the method to perform a cranial window on the cerebellar cortex is described. This technique provides optical access to the terminal portion of olivocerebellar neurons, the climbing fibers. Unfortunately, the outcome of both labeling and craniotomy surgery is quite low even in the hands of skilled operators (usually 1 out of 3 mice is labeled, and 1 out of 3 cranial windows remains clear after 1-2 weeks).
<p class="jo...

Axonal transection resulting from mechanical injury, toxic insult or neurodegenerative diseases is usually followed by degeneration of the distal part of the axon that is detached from the cell body10-13. With a few exceptions2,7,14,15, severed axons in the CNS of adult animals are usually unable to activate a regrowth program16.
Little is known about the real-time dynamics of degenerative events at the cellular and subcellular level. The development of new strategies for limiting neuronal damage and promoting neuronal regrowth requires, as a first step, clarifying the mechanism by which singularly injured neuronal cells degenerate and regenerate. This study is most directly addressed by monitoring the dynamics of a single neuron in vivo. While one-photon fluorescence imaging techniques are limited by intense scattering of visible light, two-photon excitation reaches deep cortical layers in live mice with subcellular resolution3,4,17. Taking advantage of transgenic mice in which fluorescent proteins are selectively expressed in subpopulations of neurons18-20, TPF microscopy has been applied to the exploration of synaptic plasticity and axonal elongation during development in vivo21,22.The capability of singularly damaged neurons to regrow after injury can be investigated by coupling in vivo monitoring by two-photon imaging with a model of injury specifically targeted to the axon of interest. Multi-photon absorption of femtosecond pulses has been used to disrupt single dendrites or even single spines5,23. Moreover, this injury paradigm allows cutting single axonal branches without disrupting the contacting dendrite6. Within the context of dissecting the features that allow specific neuronal population to regenerate their axons once damaged, cerebellar climbing fibers (CFs) are a useful model since they retain remarkable plastic properties after injury even in adult animals24,25. Recently, long-term imaging of CFs showed that these axons are capable of regrowing in the days that follow laser axotomy6.
This protocol describes how to label olivocerebellar neurons and their axonal elongation through anterograde tracing. Once the neurons of interest are fluorescently labeled, they can be monitored repeatedly at arbitrary time points for weeks or months under a cranial window. The procedure to dissect single axonal branches by laser axotomy in vivo will then be illustrated.
The techniques presented here provide new insights into the mechanism of axonal remodeling in vivo and may help the development of therapeutic strategies to limit neuronal degeneration and promote axonal regrowth.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Lab standard stereotaxic, rat and mouse</td>
			<td>Stoelting&#160;</td>
			<td>51670</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate glass with filament</td>
			<td>Sutter Instrument Inc</td>
			<td>BF100-50-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Germinator 500 (Glass bead sterilizer)</td>
			<td>Roboz</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microinjection dispense system</td>
			<td>Picospritzer</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Small diameter round cover glass, #1 thickness, 3 mm, 100 pack (CS-3R)</td>
			<td>Warner Instruments&#160;</td>
			<td>64-0720</td>
			<td></td>
		</tr>
		<tr>
			<td>Ti:Sapphire laser, 120 fs width pulses,&#160; 90 MHz repetition rate</td>
			<td>Coherent</td>
			<td>Chameleon&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Spongostan, haemostatic sponge&#160;</td>
			<td>Ferrosan</td>
			<td>MS0005</td>
			<td></td>
		</tr>
		<tr>
			<td>Galvanometric mirrors&#160;</td>
			<td>GSI Lumonics</td>
			<td>VM500+</td>
			<td></td>
		</tr>
		<tr>
			<td>Objective&#160;</td>
			<td>&#160;Olympus</td>
			<td>XLUMPLFLN 20XW</td>
			<td></td>
		</tr>
		<tr>
			<td>Piezoelectric stage&#160;</td>
			<td>Physik Instrumente</td>
			<td>P-721</td>
			<td></td>
		</tr>
		<tr>
			<td>Photomultiplier modules&#160;</td>
			<td>Hamamatsu Photonics</td>
			<td>H7710-13</td>
			<td></td>
		</tr>
		<tr>
			<td>LabVIEW System Design Software</td>
			<td>National Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Voren, 1 mg/ml (dexamethasone-21- isonicotinate)</td>
			<td>Boheringer Ingelheim</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Rymadil (carprofen)&#160;</td>
			<td>Pfizer</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Lidocaine clorohydrate 2%</td>
			<td>ATI</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa488 dextran</td>
			<td>Life Technologies</td>
			<td>D22910</td>
			<td></td>
		</tr>
	</tbody>
</table>

laser nanosurgery of cerebellar axons in vivo

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to understand the mechanism that promotes neuronal regeneration, an in-depth analysis of the neuronal types that can remodel after injury is needed. Several studies showed that damaged climbing fibers are capable of regrowing also in adult animals1,2. The investigation of the time-lapse dynamics of degeneration and regeneration of these axons within their complex environment can be performed by time-lapse two-photon fluorescence (TPF) imaging in vivo3,4. This technique is here combined with laser surgery, which proved to be a highly selective tool to disrupt fluorescent structures in the intact mouse cortex5-9.
This protocol describes how to perform TPF time-lapse imaging and laser nanosurgery of single axonal branches in the cerebellum in vivo. Olivocerebellar neurons are labeled by anterograde tracing with a dextran-conjugated dye and then monitored by TPF imaging through a cranial window. The terminal portion of their axons are then dissected by irradiation with a Ti:Sapphire laser at high power. The degeneration and potential regrowth of the damaged neuron are monitored by TPF in vivo imaging during the days following the injury.

This protocol described how to perform axonal labeling, in vivo imaging and laser axotomy on single neurons. The timeline of the experiment is shown in Figure 1.
An example of CFs labeled with Alexa Fluor 488 Dextran and visualized under the cranial window by in vivo two-photon microscopy is reported in Figure 2. As previously reported6,27, the ascending branches display a high stability throughout the observation period of several...

Watch this Scientific Journal Video about Laser Nanosurgery of Cerebellar Axons In Vivo at JoVE.com

Laser Nanosurgery of Cerebellar Axons In Vivo

Two-photon imaging, coupled to laser nanodissection, are useful tools to study degenerative and regenerative processes in the central nervous system with subcellular resolution. This protocol shows how to label, image, and dissect single climbing fibers in the cerebellar cortex in vivo.

Laser Nanosurgery of Cerebellar Axons In Vivo

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to ...

laser-nanosurgery-of-cerebellar-axons-in-vivo

Research

JoVE Journal

Neuroscience

8.1K Views.  University of Florence.  Двухфотонное изображений, в сочетании с лазерной nanodissection, являются полезными инструментами для изучения дегенеративные и регенеративные процессы в центральной нервной системе с субклеточной резолюции. Этот протокол показывает, как маркировать, изображени...

Video: Laser Nanosurgery of Cerebellar Axons In Vivo

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

In vivo Laser Axotomy in C. elegans

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron is damaged by injury or disease, it may regenerate. Cell-intrinsic and extrinsic factors influence the ability of a neuron to regenerate and restore function. Recently, the nematode C. elegans has emerged as an excellent model organism to identify genes and signaling pathways that influence the regeneration of neurons1-6. The main way to initiate neuronal regeneration in C. elegans is laser-mediated cutting, or axotomy. During axotomy, a fluorescently-labeled neuronal process is severed using high-energy pulses. Initially, neuronal regeneration in C. elegans was examined using an amplified femtosecond laser5. However, subsequent regeneration studies have shown that a conventional pulsed laser can be used to accurately sever neurons in vivo and elicit a similar regenerative response1,3,7.
We present a protocol for performing in vivo laser axotomy in the worm using a MicroPoint pulsed laser, a turnkey system that is readily available and that has been widely used for targeted cell ablation. We describe aligning the laser, mounting the worms, cutting specific neurons, and assessing subsequent regeneration. The system provides the ability to cut large numbers of neurons in multiple worms during one experiment. Thus, laser axotomy as described herein is an efficient system for initiating and analyzing the process of regeneration.

In vivo Laser Axotomy in C. elegans

A protocol to cut neurons in C. elegans with a MicroPoint pulsed laser is presented. We describe setting up the system, immobilizing worms, and severing labeled neurons. Advantages include a relatively low-cost system and the ability to sever neuronal processes or ablate cells in vivo.

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

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 ...

Ex Vivo Imaging of Postnatal Cerebellar Granule Cell Migration Using Confocal Macroscopy

During postnatal development, immature granule cells (excitatory interneurons) exhibit tangential migration in the external granular layer, and then radial migration in the molecular layer and the Purkinje cell layer to reach the internal granular layer of the cerebellar cortex. Default in migratory processes induces either cell death or misplacement of the neurons, leading to deficits in diverse cerebellar functions. Centripetal granule cell migration involves several mechanisms, such as chemotaxis and extracellular matrix degradation, to guide the cells towards their final position, but the factors that regulate cell migration in each cortical layer are only partially known. In our method, acute cerebellar slices are prepared from P10 rats, granule cells are labeled with a fluorescent cytoplasmic marker and tissues are cultured on membrane inserts from 4 to 10 hr before starting real-time monitoring of cell migration by confocal macroscopy at 37 &#176;C in the presence of CO2. During their migration in the different cortical layers of the cerebellum, granule cells can be exposed to neuropeptide agonists or antagonists, protease inhibitors, blockers of intracellular effectors or even toxic substances such as alcohol or methylmercury to investigate their possible role in the regulation of neuronal migration.

Ex Vivo Imaging of Postnatal Cerebellar Granule Cell Migration Using Confocal Macroscopy

During postnatal cerebellum development, immature granule cells originating from the germinal zone exhibit distinct modalities of migration to reach their final destination and to establish neuronal networks. This protocol describes the preparation of cerebellar slices and the confocal macroscopic approach used to investigate the factors that regulate neuronal migration.

During postnatal development, immature granule cells (excitatory interneurons) exhibit tangential migration in the external granular layer, and then ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their post-natal development, abundance, and accessibility, CGNs are an ideal model to study neuronal processes, including neuronal development, neuronal migration, and physiological neuronal activity stimulation. In addition, CGN cultures provide an excellent model for studying different modes of cell death including excitotoxicity and apoptosis. Within a week in culture, CGNs express N-methyl-D-aspartate (NMDA) receptors, a specific ionotropic glutamate receptor with many critical functions in neuronal health and disease. The addition of low concentrations of NMDA in conjunction with membrane depolarization to rodent primary CGN cultures has been used to model physiological neuronal activity stimulation while the addition of high concentrations of NMDA can be employed to model excitotoxic neuronal injury. Here, a method of isolation and culturing of CGNs from 6 day old pups as well as genetic manipulation of CGNs by adenoviruses and lentiviruses are described. We also present optimized protocols on how to stimulate NMDA-induced excitotoxicity, low-potassium-induced apoptosis, oxidative stress and DNA damage following transduction of these neurons.

This protocol describes a simple method for isolating and culturing primary mouse cerebral granule neurons (CGNs) from 6-7 day old pups, efficient transduction of CGNs for loss and gain of function studies, and modelling NMDA-induced neuronal excitotoxicity, low-potassium-induced cell death, DNA-damage, and oxidative stress using the same culture model.

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their ...

In Vivo Multimodal Imaging and Analysis of Mouse Laser-Induced Choroidal Neovascularization Model

Laser-induced choroidal neovascularization (CNV) is a well-established model to mimic the wet form of age-related macular degeneration (AMD). In this protocol, we aim to guide the reader not simply through the technical considerations of generating laser-induced lesions to trigger neovascular processes, but rather focus on the powerful information that can be obtained from multimodal longitudinal in vivo imaging throughout the follow-up period.
The laser-induced mouse CNV model was generated by a diode laser administration. Multimodal in vivo imaging techniques were used to monitor CNV induction, progression and regression. First, spectral domain optical coherence tomography (SD-OCT) was performed immediately after the lasering to verify a break of Bruch&#39;s membrane. Subsequent in vivo imaging using fluorescein angiography (FA) confirmed successful damage of Bruch&#39;s membrane from serial images acquired at the choroidal level. Longitudinal follow-up of CNV proliferation and regression on days 5, 10, and 14 after the lasering was performed using both SD-OCT and FA. Simple and reliable grading of leaky CNV leasions from FA images is&#160;presented. Automated segmentation for measurement of total retinal thickness, combined with manual caliber application for measurement of retinal thickness at CNV sites, allow unbiased evaluation of the presence of edema. Finally, histological verification of CNV is performed using isolectin GS-IB4 staining on choroidal flatmounts. The staining is thresholded, and the isolectin-positive area is calculated with ImageJ.
This protocol is especially useful in therapeutics studies requiring high-throughput-like screening of CNV pathology as it allows fast, multimodal, and reliable classification of CNV pathology and retinal edema. In addition, high resolution SD-OCT enables the recording of other pathological hallmarks, such as the accumulation of subretinal or intraretinal fluid. However, this method does not provide a possibility to automate CNV volume analysis from SD-OCT images, which has to be performed manually.

In Vivo Multimodal Imaging and Analysis of Mouse Laser-Induced Choroidal Neovascularization Model

Here, we present the usefulness of longitudinal in vivo imaging in the follow-up of morphological changes of laser-induced choroidal neovascularization in mice.

Laser-induced choroidal neovascularization (CNV) is a well-established model to mimic the wet form of age-related macular degeneration (AMD). In this ...

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its highly-organized subcellular architecture, we recently developed a new imaging modality, named spectral reflectometry (SpeRe), which enables unprecedented label-free nanoscale imaging of the live myelinated axons in situ. The underlying principle is to obtain nanostructural information by analyzing the reflectance spectrum of the multilayered subcellular structure. In this article, we describe a detailed step-by-step protocol for performing a basic SpeRe imaging of the nervous tissues using a commercial confocal microscopic system, equipped with a white-light laser and a tunable filter. We cover the procedures of sample preparation, acquisition of spectral data, and image processing for obtaining nanostructural information.

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

Here, we present a step-by-step protocol for imaging myelinated axons in a fixed brain slice using a label-free nanoscale imaging technique based on spectral reflectometry.

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its ...

Assessment of Long-term Depression Induction in Adult Cerebellar Slices

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from parallel fibers (PF) to Purkinje cells (PC) is considered the basis for motor learning, and deficiencies of both LTD and motor learning are observed in various gene-manipulated animals. Common motor learning sets, such as adaptation of the optokinetic reflex (OKR), the vestibular-ocular reflex (VOR), and rotarod test were used for evaluation of motor learning ability. However, results obtained from the GluA2-carboxy terminus modified knock-in mice demonstrated normal adaptation of the VOR and the OKR, despite lacking PF-LTD. In that report, induction of LTD was only attempted using one type of stimulation protocol at room temperature. Thus, conditions to induce cerebellar LTD were explored in the same knock-in mutants using various protocols at near physiological temperature. Finally, we found stimulation protocols, by which LTD could be induced in these gene-manipulated mice. In this study, a set of protocols are proposed to evaluate LTD-induction, which will more accurately allow examination of the causal relationship between LTD and motor learning. In conclusion, experimental conditions are crucial when evaluating LTD in gene-manipulated mice.

In some gene-manipulated animals, using a single protocol may fail to induce LTD in cerebellar Purkinje cells, and there may be a discrepancy between LTD and motor learning. Multiple protocols are necessary to assess LTD-induction in gene-manipulated animals. Standard protocols are shown.

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from ...