Name: visualization of thalamocortical axon branching and synapse formation in organotypic cocultures
Uploaded: 2018-03-28T12:30:00
Description: Watch this Scientific Journal Video about Visualization of Thalamocortical Axon Branching and Synapse Formation in Organotypic Cocultures at JoVE.com

This work was supported by MEXT KAKENHI on Innovative Areas Mesoscopic Neurocircuitry 23115102 and Dynamic Regulation of Brain Function by Scrap and Build System 16H06460 to N.Y.&#160;We also thank Gabriel Hand for critical reading.

All experiments were performed according to the guidelines established by the animal welfare committees of Osaka University and the Japan Neuroscience Society.
1. Organotypic cocultures of the thalamus and cerebral cortex
Note: For the detailed procedure, refer to the original publications7,8,13. All procedures should be performed under sterile conditions. Sprague-Dawley (SD)...

<ol>
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The current protocol is also a powerful tool to study developmental aspects of growing axons other than of the TC projection11. For instance, a combination of cortical slice culture and the electroporation technique allows visualizing individual axonal morphology of cortical neurons and long term observation9,18.
By using the current protocol, the roles of interesting genes in axon branching and synapse formatio...

The thalamocortical (TC) projection in the mammalian brain is a suitable system to investigate axon guidance and targeting mechanisms. During development, sensory TC axons grow in the cortical plate, and form branches and synapses preferentially in layer IV of the primary sensory areas in the cerebral cortex1,2. Even after establishment of fundamental connections, axonal arbors and synaptic terminals are remodeled depending on environmental changes3,4. However, how TC axon morphology is dynamically altered is poorly understood. One of the main reasons is the lack of an adequate technique to observe structural changes at a single cell level. Although recent developments in microscopy, such as two-photon microscopy, have allowed direct observation of living cortical neurons in vivo, there are still technical limitations for capturing the overall TC trajectories5,6. Therefore, in vitro methods for live imaging of TC axons would provide powerful tools for structural analyses of axon branching and synapse formation.
Our group for the first time established a static slice culture method with permeable membrane7. Using this method, a rat cortical slice was cocultured with a sensory thalamic block, and lamina-specific TC connections were recapitulated in this organotypic cocultures7,8. Sparse labeling with a fluorescent protein further allowed us to observe TC axon growth and branch formation9,10,11. Recently, we have developed a novel method for simultaneous imaging of branching and synapse formation of individual TC axons in the organotypic cocultures12. To visualize TC axons and presynaptic sites simultaneously, DsRed and EGFP-tagged synaptophysin (SYP-EGFP) were co-transfected into a small number of thalamic neurons by electroporation of the organotypic coculture. The current method facilitates morphological analysis of TC axons and allows for long-term observation, which can be used to show the causal relationship between axon branching and synapse formation.

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			<td class="xl20" height="40" style="width:113pt;height:30.0pt" width="151">35 mm petri dishes</td>
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			<td class="xl20" height="40" style="width:113pt;height:30.0pt" width="151">Millicell-CM insert</td>
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			<td class="xl20" height="40" style="width:113pt;height:30.0pt" width="151">100 mm petri dishes</td>
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			<td class="xl20" height="59" style="width:113pt;height:44.25pt" width="151">Disposable sterile plastic pipettes</td>
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			<td class="xl20" height="54" style="width:113pt;height:40.5pt" width="151">Glass capillary: OD 1.2 mm</td>
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			<td class="xl20" height="40" style="width:113pt;height:30.0pt" width="151">1 mL syringe</td>
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			<td class="xl20" height="40" style="width:113pt;height:30.0pt" width="151">Confocal microscope</td>
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			<td class="xl20" height="40" style="width:113pt;height:30.0pt" width="151">Sprague-Dawley (SD) rat</td>
			<td class="xl20" style="width:89pt" width="119">Japan SLC and Nihon-Dobutsu</td>
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			<td class="xl20" height="40" style="width:113pt;height:30.0pt" width="151">Microsurgery scissors</td>
			<td class="xl20" style="width:89pt" width="119">Natsume&#160;</td>
			<td class="xl20" style="width:84pt" width="112">MB-54-1</td>
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visualization of thalamocortical axon branching and synapse formation in organotypic cocultures

Axon branching and synapse formation are crucial processes for establishing precise neuronal circuits. During development, sensory thalamocortical (TC) axons form branches and synapses in specific layers of the cerebral cortex. Despite the obvious spatial correlation between axon branching and synapse formation, the causal relationship between them is poorly understood. To address this issue, we recently developed a method for simultaneous imaging of branching and synapse formation of individual TC axons in organotypic cocultures.
This protocol describes a method which consists of a combination of an organotypic coculture and electroporation. Organotypic cocultures of the thalamus and cerebral cortex facilitate gene manipulation and observation of axonal processes, preserving characteristic structures such as laminar configuration. Two distinct plasmids encoding DsRed and EGFP-tagged synaptophysin (SYP-EGFP) were co-transfected into a small number of thalamic neurons by an electroporation technique. This method allowed us to visualize individual axonal morphologies of TC neurons and their presynaptic sites simultaneously. The method also enabled long-term observation which revealed the causal relationship between axon branching and synapse formation.

The experiment described here aims to reveal the relationship between TC axon branching and synapse formation. To simultaneously visualize axonal trajectories and locations of presynaptic sites, single or a few thalamic cells in organotypic cocultures were transfected with two plasmids encoding SYP-EGFP and DsRed using electroporation. During the second week in culture, individually distinguishable TC axons were clearly labeled by DsRed (Figure 3). Only the a...

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Visualization of Thalamocortical Axon Branching and Synapse Formation in Organotypic Cocultures

This protocol describes a method for simultaneous imaging of thalamocortical axon branching and synapse formation in organotypic cocultures of the thalamus and cerebral cortex. Individual thalamocortical axons and their presynaptic terminals are visualized by a single cell electroporation technique with DsRed and GFP-tagged synaptophysin.

Axon branching and synapse formation are crucial processes for establishing precise neuronal circuits. During development, sensory thalamocortical (TC) ...

The overall goal of this method is to study dynamic morphological changes of axon branching and synapse formation in the thalamocortical projection. This method can help answer key questions in the field of neurobiology, such as new neural circuit formation. The main advantage of this technique is simultaneous observation of axon growth and the synapse formation in individual living cells.This method can provide insight into the relationship between axon branching and synapse formation in the thalamocortical projections. It can also be applied to other neural circuits, such as intrinsic connection in the cerebral cortex. To begin this procedure, sterilize all the surgical instruments with 70%ethanol for 10 minutes or longer.Next, dissect the whole brain quickly from a postnatal day-two rat under ice-cold anesthesia. Then, place the brain into a 100-millimeter petri dish containing 40 milliliters of cold Hank's Balanced Salt Solution. Under a binocular microscope, remove the pia matter carefully with fine forceps, and cut the primary visual and somatosensory cortex into 300 to 400-micron thick slices using microsurgical scissors.After that, transfer the cortical slices onto the membrane filter of the culture insert using a disposable plastic pipette. Adjust the positions of the slices near the center of the culture insert with the disposable plastic pipette, and remove excess medium from the insert. Maintain the slices in the serum-containing culture medium at 37 degrees Celsius, and culture them alone for one day before placing a thalamic block from an E15 embryo next to it.Adjust the level of the medium slightly below the surface of the slice, so that the slices can receive sufficient supply of both the gas and the medium. This is crucial for the viability of the slices. In this step, make the glass capillaries with an electrode puller.Then break the tips of the glass capillaries for plasmid injection. For the electrode pipettes, grind and polish the tips of the glass capillaries with sandpaper. And then fire polish them to achieve an inner diameter of 50 to 200 microns.Next, attach the electrode pipette and the ejection pipette to the manipulators. Transfer five microliters of plasmid solution to a petri dish. Then draw the solution into the electrode pipette with capillary action.Insert the 0.2-millimeter diameter electrode into the electrode pipette. After connecting the ejection pipette to a one-milliliter syringe with a plastic tube, transfer five microliters of plasmid solution to a petri dish again. Then slowly draw the solution into the ejection pipette using the syringe.Following that, transfer the coculture on the electroporation stage and insert the one-millimeter diameter electrode into the culture medium. Subsequently, place the ejection pipette on the cultured thalamic block. Push the syringe manually after the tip touches the surface of the thalamic block.Place the electrode pipette on the thalamic block immediately after the retraction of the ejection pipette. Now, deliver the electrical pulses. Monitor the amplitude and the number of trains on the oscilloscope.Afterward, retract the electrode pipette and return the dish to the incubator. To observe the sample, place the culture dish on the stage of an upright confocal microscope equipped with two filter sets for EGFP and DsRed. Then take images of 10 to 25 optical sections, at one to seven-micron step sizes, with a 20x long working distance objective lens.Select individually-distinguishable labeled axons, which express both DsRed and Syp-EGFP. Shown here is an organotypic coculture of thalamic and cortical slices after 12 DIV. Here is a magnified image of the boxed region.And here is a DsRed-labeled thalamic cell extending an axon into the cortical slice, and forming branches in upper layers. These images show the DsRed-labeled thalamocortical axon and Syp-EGFP puncta. Discrete accumulation of Syp-EGFP was observed along a single thalamocortical axon.DsRed plus Syp-EGFP plasmids were introduced into cultured thalamic cells at one DIV using electroporation. This axon was imaged daily over four days. Axon branching and synapse formation were dynamic, with addition and elimination.Following these procedures, electroporation with interesting genes together with Syp-EGFP and DsRed can be performed in order to investigate the role of the gene in axon branching and synapse formations. This technique paves the way for researchers in the field of the neurobiology to explore the mechanisms of dynamic changes of neural circuits.

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JoVE Journal

Neuroscience

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct ...

Inhibitory Synapse Formation in a Co-culture Model Incorporating GABAergic Medium Spiny Neurons and HEK293 Cells Stably Expressing GABAA Receptors

Inhibitory neurons act in the central nervous system to regulate the dynamics and spatio-temporal co-ordination of neuronal networks. GABA (&#947;-aminobutyric acid) is the predominant inhibitory neurotransmitter in the brain. It is released from the presynaptic terminals of inhibitory neurons within highly specialized intercellular junctions known as synapses, where it binds to GABAA receptors (GABAARs) present at the plasma membrane of the synapse-receiving, postsynaptic neurons. Activation of these GABA-gated ion channels leads to influx of chloride resulting in postsynaptic potential changes that decrease the probability that these neurons will generate action potentials. 
During development, diverse types of inhibitory neurons with distinct morphological, electrophysiological and neurochemical characteristics have the ability to recognize their target neurons and form synapses which incorporate specific GABAARs subtypes. This principle of selective innervation of neuronal targets raises the question as to how the appropriate synaptic partners identify each other. 
To elucidate the underlying molecular mechanisms, a novel in vitro co-culture model system was established, in which medium spiny GABAergic neurons, a highly homogenous population of neurons isolated from the embryonic striatum, were cultured with stably transfected HEK293 cell lines that express different GABAAR subtypes. Synapses form rapidly, efficiently and selectively in this system, and are easily accessible for quantification. Our results indicate that various GABAAR subtypes differ in their ability to promote synapse formation, suggesting that this reduced in vitro model system can be used to reproduce, at least in part, the in vivo conditions required for the recognition of the appropriate synaptic partners and formation of specific synapses. Here the protocols for culturing the medium spiny neurons and generating HEK293 cells lines expressing GABAARs are first described, followed by detailed instructions on how to combine these two cell types in co-culture and analyze the formation of synaptic contacts.

Inhibitory Synapse Formation in a Co-culture Model Incorporating GABAergic Medium Spiny Neurons and HEK293 Cells Stably Expressing GABAA Receptors

The molecular mechanisms that co-ordinate the formation of inhibitory GABAergic synapses during ontogeny are largely unknown. To study these processes,we have developed a co-culture model system which incorporates embryonic medium spiny GABAergic neurons cultured together with stably transfected human embryonic kidney 293 (HEK293) cells expressing functional GABAA receptors.

Inhibitory neurons act in the central nervous system to regulate the dynamics and spatio-temporal co-ordination of neuronal networks. GABA ...

Whole-mount Imaging of Mouse Embryo Sensory Axon Projections

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we describe a method to label in situ a subset of dorsal root ganglion (DRG) axon projections to assess their phenotypic characteristics using several genetically manipulated mouse lines. The TrkA-positive neurons are nociceptor neurons, dedicated to the transmission of pain signals. We utilize a TrkAtaulacZ mouse line to label the trajectories of all TrkA-positive peripheral axons in the intact mouse embryo. We further breed the TrkAtaulacZ line onto a Bax null background, which essentially abolishes neuronal apoptosis, in order to assess growth-related questions independently of possible effects of genetic manipulations on neuronal survival. Subsequently, genetically modified mice of interest are bred with the TrkAtaulacZ/Bax null line and are then ready for study using the techniques described herein. This presentation includes detailed information on mouse breeding plans, genotyping at the time of dissection, tissue preparation, staining and clearing to allow for visualization of full-length axonal trajectories in whole-mount preparation.

We present here an optimized protocol to genotype, stain and prepare fetal mice for the imaging of peripheral nociceptor axon projections in the whole animal, as an effective method to assess sensory axon growth phenotypes in developing genetically engineered mice.

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we ...

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

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

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

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

Quadruple Immunostaining of the Olfactory Bulb for Visualization of Olfactory Sensory Axon Molecular Identity Codes

The mouse olfactory system is often used to study mechanisms of neural circuit formation&#12288;because of its simple anatomical structure. An Olfactory Sensory Neuron (OSN) is a bipolar cell with a single dendrite and a single unbranched axon. An OSN expresses only one Olfactory Receptor (OR) gene, OSNs expressing a given type of OR converge their axons to a few sets of invariant glomeruli in the Olfactory Bulb (OB). A remarkable feature of OSN projection is that the expressed ORs play instructive roles in axonal projection. ORs regulate the expression of multiple axon-sorting molecules and generate the combinatorial molecular code of axon-sorting molecules at the OSN axon termini. Thus, to understand the molecular mechanisms of OR-specific axon guidance mechanisms, it is vital to characterize their expression profiles at the OSN axon termini within the same glomerulus. The aim of this article was to introduce methods for collecting as many glomeruli as possible on a single OB section and for performing immunostaining using multiple antibodies. This would allow the comparison and analysis of the expression patterns of axon-sorting molecules without staining variation between OB sections.

Olfactory sensory neurons express a wide variety of axon-sorting molecules to establish proper neural circuitry. This protocol describes an immunohistochemical staining method to visualize combinatorial expressions of axon-sorting molecules at the axon termini of olfactory sensory neurons.

The mouse olfactory system is often used to study mechanisms of neural circuit formation&#12288;because of its simple anatomical structure. An Olfactory ...

Improving the Application of High Molecular Weight Biotinylated Dextran Amine for Thalamocortical Projection Tracing in the Rat

High molecular weight biotinylated dextran amine (BDA) has been used as a highly sensitive neuroanatomical tracer for many decades. Since the quality of its labeling was affected by various factors, here, we provide a refined protocol for the application of high molecular weight BDA for studying optimal neural labeling in the central nervous system. After stereotactic injection of BDA into the ventral posteromedial nucleus (VPM) of the thalamus in the rat through a delicate glass pipette, BDA was stained with fluorescent streptavidin-Alexa (AF) 594 and counterstained with fluorescent Nissl stain AF500/525. On the background of green Nissl staining, the red BDA labeling, including neuronal cell bodies and axonal terminals, was more distinctly demonstrated in the somatosensory cortex. Furthermore, double fluorescent staining for BDA and the calcium-binding protein parvalbumin (PV) was carried out to observe the correlation of BDA labeling and PV-positive interneurons in the cortical target, providing the opportunity to study the local neural circuits and their chemical characteristics. Thus, this refined method is not only suitable for visualizing high quality neural labeling with the high molecular weight BDA through reciprocal neural pathways between the thalamus and cerebral cortex, but also will permit the simultaneous demonstration of other neural markers with fluorescent histochemistry or immunochemistry.

Here, we present a refined protocol to effectively reveal biotinylated dextran amine (BDA) labeling with a fluorescent staining method through a reciprocal neural pathway. It is suitable for analyzing the fine structure of BDA labeling and distinguishing it from other neural elements under a confocal laser scanning microscope.

High molecular weight biotinylated dextran amine (BDA) has been used as a highly sensitive neuroanatomical tracer for many decades. Since the quality of ...

Evaluation of Synapse Density in Hippocampal Rodent Brain Slices

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is tightly regulated during development and neuronal maturation. Importantly, deficits in synapse number can lead to cognitive dysfunction. Therefore, the evaluation of synapse number is an integral part of neurobiology. However, as synapses are small and highly compact in the intact brain, the assessment of absolute number is challenging. This protocol describes a method to easily identify and evaluate synapses in hippocampal rodent slices using immunofluorescence microscopy. It includes a three-step procedure to evaluate synapses in high-quality confocal microscopy images by analyzing the co-localization of pre- and postsynaptic proteins in hippocampal slices. It also explains how the analysis is performed and gives representative examples from both excitatory and inhibitory synapses. This protocol provides a solid foundation for the analysis of synapses and can be applied to any research investigating the structure and function of the brain.

A protocol for accurately identifying and analyzing synapses in hippocampal slices using immunofluorescence is outlined in this article.

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is ...

Visualization and Live Imaging of Oligodendrocyte Organelles in Organotypic Brain Slices Using Adeno-associated Virus and Confocal Microscopy

Neurons rely on the electric insulation and trophic support of myelinating oligodendrocytes. Despite the importance of oligodendrocytes, the advanced tools currently used to study neurons, have only partly been taken on by oligodendrocyte researchers. Cell type-specific staining by viral transduction is a useful approach to study live organelle dynamics. This paper describes a protocol for visualizing oligodendrocyte mitochondria in organotypic brain slices by transduction with adeno-associated virus (AAV) carrying genes for mitochondrial targeted fluorescent proteins under the transcriptional control of the myelin basic protein promoter. It includes the protocol for making organotypic coronal mouse brain slices. A procedure for time-lapse imaging of mitochondria then follows. These methods can be transferred to other organelles and may be particularly useful for studying organelles in the myelin sheath. Finally, we describe a readily available technique for visualization of unstained myelin in living slices by Confocal Reflectance microscopy (CoRe). CoRe requires no extra equipment and can be useful to identify the myelin sheath during live imaging.

Myelinating oligodendrocytes promote rapid action potential propagation and neuronal survival. Described here is a protocol for oligodendrocyte-specific expression of fluorescent proteins in organotypic brain slices with subsequent time-lapse imaging. Further, a simple procedure for visualizing unstained myelin is presented.

Neurons rely on the electric insulation and trophic support of myelinating oligodendrocytes. Despite the importance of oligodendrocytes, the advanced ...

Visualization of Motor Axon Navigation and Quantification of Axon Arborization In Mouse Embryos Using Light Sheet Fluorescence Microscopy

Spinal motor neurons (MNs) extend their axons to communicate with their innervating targets, thereby controlling movement and complex tasks in vertebrates. Thus, it is critical to uncover the molecular mechanisms of how motor axons navigate to, arborize, and innervate their peripheral muscle targets during development and degeneration. Although transgenic Hb9::GFP mouse lines have long served to visualize motor axon trajectories during embryonic development, detailed descriptions of the full spectrum of axon terminal arborization remain incomplete due to the pattern complexity and limitations of current optical microscopy. Here, we describe an improved protocol that combines light sheet fluorescence microscopy (LSFM) and robust image analysis to qualitatively and quantitatively visualize developing motor axons. This system can be easily adopted to cross genetic mutants or MN disease models with Hb9::GFP lines, revealing novel molecular mechanisms that lead to defects in motor axon navigation and arborization.

Here, we describe a protocol for visualizing motor neuron projection and axon arborization in transgenic Hb9::GFP mouse embryos. After immunostaining for motor neurons, we used light sheet fluorescence microscopy to image embryos for subsequent quantitative analysis. This protocol is applicable to other neuron navigation processes in the central nervous system.