The work was funded by the University of Zurich. The authors would like to thank Estrela Neto and Dr. Meriem Lamghari for helping in establishing the co-culture conditions.

All mice were maintained and handled according to the Swiss Animal Welfare Law and in compliance with the regulations of the Cantonal Veterinary office, Zurich.
1. Preparation of Dissection Material, Culture Media, Microfluidic Devices
<ol>
	<li>Autoclave micro-dissection forceps and scissors (121 &#176;C, sterilization time: 20 min) and store them in a sterile container.</li>
	<li>Sterilize glass coverslips (24 mm x 24 mm) by incubating them in 1 M HCl for 24 hr on an orbital shaker at 37 &#176;C. Wash them three times with sterile, distilled H2O and three times with ethanol 99%. Dry then the coverslips at 37 &#176;C or under sterile flow hood. Finally, autoclave or expose the coverslips to UV light (30 min) to complete the sterilization. Coverslips can then be stored in ethanol 70%.</li>
	<li>Remove carefully the AXIS Axon Isolation Devices from the package using sterile forceps and place them in a sterile Petri dish.</li>
	<li>Using a sterile biopsy punch (&#248;: 1mm) create one hole per sample to be cultured (Figure 1) in correspondence of the culture chambers. 
	NOTE: Do not punch too close to the microgrooves, as they might be damaged by the pressure applied.</li>
	<li>Sterilize the AXIS Axon Isolation Devices by immerging them in ethanol 70%. Dry then AXIS Axon Isolation Devices and coverslips completely under a sterile flow hood. Wait a minimum of 3 hr before proceeding. 
	NOTE: incomplete drying will result in defective assembly of the microfluidic devices.</li>
	<li>Place each coverslip into a 35 mm Petri dish or into a well within a 6-wells plate.</li>
	<li>Place the AXIS Axon Isolation Device onto the coverslip and press gently but firmly with a forceps with bent ends in order to allow full adhesion between the isolation device and the glass coverslip.</li>
	<li>In each culture chamber, pipette 150 &#181;l of poly-D-lysine (0.1 mg/ml in sterile, distilled H2O). Place the microfluidic devices under vacuum for 5 min, in order to remove all the air from the culture chambers.</li>
	<li>If air can still be seen within the chambers, re-pipette the poly-D-lysine solution into the chambers.</li>
	<li>Incubate the devices with poly-D-lysine O/N&#160;at 37 &#176;C.</li>
	<li>Wash chambers three time with sterile, distilled H2O.</li>
	<li>Fill chambers with 150 &#181;l laminin working solution (Sigma-Aldrich, 5 &#181;g/ml, in PBS or serum-free medium), and incubate O/N at 37 &#176;C.</li>
	<li>Prepare 50 ml of medium for trigeminal ganglia cultures 37 , composed as follows: 48 ml Neurobasal medium, 1 ml B-27, 100 U/ml penicillin/streptomycin, 2 mM L-glutamine, 5 ng/ml nerve growth factor (NGF), 0.25 pM cytosine arabinoside.</li>
	<li>Prepare 50 ml of medium for tooth germ cultures 37, composed as follows: 40 ml DMEM-F12, 10 ml foetal bovine serum (FBS, final concentration: 20%), 100 U/ml penicillin/streptomycin, 2 mM L-glutamine, 150 &#181;g/ml ascorbic acid.</li>
</ol>
2. Mouse Embryo Generation and Dissection
<ol>
	<li>Determine embryonic age according to vaginal plug (vaginal plug: embryonic day of development 0.5, E0.5) and confirm it via morphological criteria. For this protocol, we generally use E14.5-E17.5 mouse embryos.</li>
	<li>Clean the dissection area and the stereoscope with ethanol 70%.</li>
	<li>Sacrifice the pregnant mother via cervical dislocation. Block the neck of the mouse with the first and second finger onto a grid, and pull with decision the tail.</li>
	<li>Dissect the skin around the lower abdomen, and open the abdomen using scissors. Locate the uterus: during such late stages of pregnancy, the uterus abundantly fill the abdominal cavity.</li>
	<li>Dissect out the uterus and place in a tube filled with PBS on ice. When on ice, the tissue can be left for several hr. Discard the corpse of the mother according to institutional guidelines</li>
	<li>Dissect out the embryos from the uterus and free them from their extraembryonic tissues. Place the embryos in PBS on ice.</li>
	<li>Decapitate the embryos using scissors, and separate the lower jaw from the rest of the head using micro-dissection scissors (Figure 2A). Remove precisely the lower jaw, without damaging the trigeminal ganglia; as the latter are localized in close proximity to the lower jaw, their accidental damage is possible. Preserve the lower jaw and the rest of the head in cold PBS, on ice.</li>
	<li>To dissect TG, take the head and place it onto a dissection glass Petri dish, previously filled with cold PBS. Using the forceps, remove the skin and the skull. Remove then the telencephalon and the cerebellum by placing forceps below the telencephalon and lift; the telencephalon and the cerebellum will flip together, leaving the bottom of the skull exposed.</li>
	<li>Localize the trigeminal ganglia (shown in Figure 2B). Use the forceps to separate the TG from the trigeminal nerves. Eliminate the remnants of the trigeminal projections using the dissection needles as knifes. Place the dissected TG in a Petri dish filled with cold PBS and keep them on ice.</li>
	<li>To dissect embryonic teeth, place the lower jaw, previously separated from the skull, onto the dissection glass Petri dish, filled with cold PBS. Using dissection needles as knifes, remove the tongue and the skin surrounding the jaw. Separate the left and the right hemi-jaws by cutting along the midline of the jaw. The tooth germs are easily visible, as shown in Figure 1C. Isolate the tooth germs using dissection needles and remove the excess of non-dental tissues. Place the dissected tooth germs in a Petri dish filled with cold PBS and keep them on ice.</li>
</ol>
3. Microfluidic Co-cultures
<ol>
	<li>After dissection, remove laminin from the microfluidic devices. Fill the chambers with 200 &#181;l of the respective media.</li>
	<li>With forceps, transfer gently the dissected TG and tooth germs into the holes created by punching (Figure 1D). Make sure that the tooth germs do not float and that they sink until they contact the coverslips.</li>
	<li>Culture the samples in incubator at 37 &#176;C, 5% CO2.</li>
	<li>Change the culture medium every 48 hr. Do not empty the chambers completely, and do not pipette directly into the culture chambers. Complete emptying of chambers would result in the formation of air bubbles within the chambers; direct pipetting into the chamber would result in axonal damage. To avoid these issues, remove the medium pointing the pipette towards the external side of the wells; similarly, pipette the fresh medium on the side of the wells located opposite to the chambers.</li>
	<li>During the culture period, co-cultures can be easily imaged by time-lapse microscopy. Co-cultures can be maintained for over 10 days.</li>
	<li>After the culture period, wash the chambers by pipetting 150 &#181;l of PBS into one well per chamber, and letting PBS flow through the chambers three times.</li>
	<li>Remove the PBS and fix the samples by pipetting 150 &#181;l of paraformaldehyde 4% (in PBS) in one well per chamber; incubate at RT for 15 min.</li>
	<li>Wash the chambers twice with PBS as described in 3.6.</li>
	<li>Proceed with further analysis.</li>
</ol>

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The authors declare that they have no competing financial interests.

Previous in vitro studies of tooth innervation were based on conventional co-cultures of trigeminal ganglia and dental tissues or cells 26,28,29. These studies were conducted to investigate mainly the attractive effects of these cells or tissues on sensory axons 38. Although bringing significant advances in the field, several technical issues were raised. Tooth germs start to degenerate after few days of culture 37. Based on these observations, growing neurons and teeth in the sa...

Innervation plays a key role in the development, homeostasis and regeneration of organs and tissues 1,2. Furthermore, innervation is involved in the regulation of stem cell proliferation, mobilization and differentiation 3&#8211;5. Indeed, recent studies realised in tissues of the orofacial complex have shown that parasympathetic nerves are necessary for epithelial progenitor cells function during the development and regeneration of the salivary glands 6,7. Similarly, it has been demonstrated that innervation is necessary for the development and maintenance of taste buds 8&#8211;11. Thus, it is important to analyse the yet neglected roles of innervation in the development of other important orofacial organs and tissues such as teeth.
In spite of the rich innervation of adult teeth, and in contrast to all other organs and tissues of the body, developing teeth start to be innervated at the earliest postnatal stages. Teeth develop as a result of sequential and reciprocal interactions between the oral ectoderm and cranial neural crest-derived mesenchyme. These interactions give rise to epithelial-derived ameloblasts and mesenchyme-derived odontoblasts that are responsible for the formation of enamel and dentin, respectively 12. Sensory nerves from the trigeminal ganglia and sympathetic nerves from the superior cervical ganglia innervate the adult teeth 13&#8211;15. During embryogenesis, nerve fibres emanating from the trigeminal ganglia project towards the developing tooth germs and progressively surround them but they do not penetrate into the dental papilla mesenchyme 13. Nerve fibres enter the dental pulp mesenchyme at more advanced developmental stages that correlate with odontoblast differentiation and dentin matrix deposition 16. Dental pulp innervation is completed soon after tooth eruption in the oral cavity 13. Previous studies have revealed that various semaphorins and neurotrophins are involved in the regulation of innervation during odontogenesis 16&#8211;19. Earlier studies have clearly demonstrated that innervation is a prerequisite for tooth formation in fishes 20. More recent studies have shown that homeostasis of dental mesenchyme stem cells in mouse incisors is regulated by sensory nerves via secretion of sonic hedgehog (shh) 21. Nevertheless, the role of innervation in tooth initiation, development and regeneration is still highly controversial in mammals 22&#8211;24.
A plethora of in vivo studies have provided important information about the patterns of innervation of dental tissues during development and repair processes of various animal models 13,25,26. However, most of these approaches are not optimal to highlight the molecular basis of the interactions between nerve fibres and target organs and tissues. Co-cultures constitute a valuable method to investigate and manipulate the interactions between nerve fibres and teeth in a controlled and isolated environment 26&#8211;29. At the same time, co-culturing is subject to various technical adjustments. For example, nerves and specific dental tissues (e.g., dental pulp, dental follicle, dental epithelium) often require different culture media in order to guarantee tissue survival for long periods of time 30&#8211;32.
In the last decades, conventional co-cultures using the same culture medium have been performed for very short periods (e.g., two days) to investigate the attractive or repulsive effects of developing oral and dental tissues on sensory nerve fibres 27&#8211;29. However, extension of the culture period is required to investigate the effects of innervation on tooth morphogenesis and cytodifferentiation, and to study the dynamics of nerve fibres branching within target organs. Therefore, non-contiguous co-cultures would be more suitable to perform studies on neuronal-dental tissues interactions.
Microfluidics systems allow co-cultures of neurons and different cell types in their appropriate culture media. In these devices, dental tissues and neurons are separated in different compartments, while allowing the growth of axons from the neural cell bodies through microchannels towards the compartment containing their target tissue 33. Microfluidic devices have been already used to study the interactions between neurons and microglia 34,35, as well as cell to cell interactions in cancer and neovascularization 35. Moreover, these systems have been used to study the interactions between dorsal root ganglia and osteoblasts 36.
We have recently demonstrated that trigeminal ganglia (TG) and teeth are able to survive for long periods of time when co-cultured in microfluidic devices 37. Moreover, we have demonstrated that teeth from different developmental stages maintain in these in vitro conditions the same repulsive or attractive effects on trigeminal innervation that they show in vivo 37. This protocol provides information about a simple, powerful and flexible way to co-culture ganglia/nerves and target tissues and to study the roles of specific molecules on such interactions in a controlled and isolated environment.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>AXIS Axon Isolation Devices</td>
			<td>Millipore</td>
			<td>AX15010-TC</td>
			<td>Microchannels of different lenght are available</td>
		</tr>
		<tr>
			<td>Laminin</td>
			<td>Sigma Aldrich</td>
			<td>L2020</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurobasal</td>
			<td>Gibco</td>
			<td>21103-049</td>
			<td></td>
		</tr>
		<tr>
			<td>B27</td>
			<td>Gibco</td>
			<td>17504</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant Mouse beta-NGF</td>
			<td>R&#38;D Systems</td>
			<td>1156-NG-100</td>
			<td>Human and Rat beta-NGF (R&#38;D Systems) are equivalent</td>
		</tr>
		<tr>
			<td>DMEM-F12</td>
			<td>Gibco</td>
			<td>11320-033</td>
			<td></td>
		</tr>
	</tbody>
</table>

analysis of developing tooth germ innervation using microfluidic co-culture devices

Innervation plays a key role in the development, homeostasis and regeneration of organs and tissues. However, the mechanisms underlying these phenomena are not well understood yet. In particular, the role of innervation in tooth development and regeneration is neglected.
Several in vivo studies have provided important information about the patterns of innervation of dental tissues during development and repair processes of various animal models. However, most of these approaches are not optimal to highlight the molecular basis of the interactions between nerve fibres and target organs and tissues.
Co-cultures constitute a valuable method to investigate and manipulate the interactions between nerve fibres and teeth in a controlled and isolated environment. In the last decades, conventional co-cultures using the same culture medium have been performed for very short periods (e.g., two days) to investigate the attractive or repulsive effects of developing oral and dental tissues on sensory nerve fibres. However, extension of the culture period is required to investigate the effects of innervation on tooth morphogenesis and cytodifferentiation.
Microfluidics systems allow co-cultures of neurons and different cell types in their appropriate culture media. We have recently demonstrated that trigeminal ganglia (TG) and teeth are able to survive for a long period of time when co-cultured in microfluidic devices, and that they maintain in these conditions the same innervation pattern that they show in vivo.
On this basis, we describe how to isolate and co-culture developing trigeminal ganglia and tooth germs in a microfluidic co-culture system.This protocol describes a simple and flexible way to co-culture ganglia/nerves and target tissues and to study the roles of specific molecules on such interactions in a controlled and isolated environment.

These results show that isolated trigeminal ganglia can grow in one compartment of the microfluidic device and, in addition, that the development of the isolated tooth germs is sustained for a long period of time in the other compartment of the microfluidic device. Different culture media are used in the two compartments, and the microgrooves between the two compartments allow extension of axon from the trigeminal ganglion towards the developing tooth germs. Figure 3 represents a visualization of neurofi...

Watch this Scientific Journal Video about Analysis of Developing Tooth Germ Innervation Using Microfluidic Co-culture Devices at JoVE.com

Analysis of Developing Tooth Germ Innervation Using Microfluidic Co-culture Devices

Co-cultures represent a valuable method to study the interactions between nerves and target tissues and organs. Microfluidic systems allow co-culturing ganglia and whole developing organs or tissues in different culture media, thus representing a valuable tool for the in vitro study of the crosstalk between neurons and their targets.

Innervation plays a key role in the development, homeostasis and regeneration of organs and tissues. However, the mechanisms underlying these phenomena ...

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JoVE Journal

Neuroscience

8.2K Views.  University of Zurich. Co-cultures represent a valuable method to study the interactions between nerves and target tissues and organs. Microfluidic systems allow co-culturing ganglia and whole developing organs or tissues in different culture media, thus representing a valuable tool for the in vitro study of the crosstalk between neurons and their targets.

Video: Analysis of Developing Tooth Germ Innervation Using Microfluidic Co-culture Devices

Bilaminar Co-culture of Primary Rat Cortical Neurons and Glia

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or co-culture model. This system is suitable for a variety of experimental needs requiring either a glass or plastic growth substrate and can also be used for culture of other types of neurons.
Rat cortical neurons obtained from the late embryonic stage (E17) are plated on glass coverslips or tissue culture dishes facing a feeder layer of glia grown on dishes or plastic coverslips (known as Thermanox), respectively. The choice between the two configurations depends on the specific experimental technique used, which may require, or not, that neurons are grown on glass (e.g. calcium imaging versus Western blot). The glial feeder layer, an astroglia-enriched secondary culture of mixed glia, is separately prepared from the cortices of newborn rat pups (P2-4) prior to the neuronal dissection.
A major advantage of this culture system as compared to a culture of neurons only is the support of neuronal growth, survival, and differentiation provided by trophic factors secreted from the glial feeder layer, which more accurately resembles the brain environment in vivo. Furthermore, the co-culture can be used to study neuronal-glial interactions1.
At the same time, glia contamination in the neuronal layer is prevented by different means (low density culture, addition of mitotic inhibitors, lack of serum and use of optimized culture medium) leading to a virtually pure neuronal layer, comparable to other established methods1-3. Neurons can be easily separated from the glial layer at any time during culture and used for different experimental applications ranging from electrophysiology4, cellular and molecular biology5-8, biochemistry5, imaging and microscopy4,6,7,9,10. The primary neurons extend axons and dendrites to form functional synapses11, a process which is not observed in neuronal cell lines, although some cell lines do extend processes.
A detailed protocol of culturing rat hippocampal neurons using this co-culture system has been described previously4,12,13. Here we detail a modified protocol suited for cortical neurons. As approximately 20x106 cells are recovered from each rat embryo, this method is particularly useful for experiments requiring large numbers of neurons (but not concerned about a highly homogenous neuronal population). The preparation of neurons and glia needs to be planned in a time-specific manner. We will provide the step-by-step protocol for culturing rat cortical neurons as well as culturing glial cells to support the neurons.

Here we provide a protocol for culturing rat cortical neurons in the presence of a glial feeder layer. The cultured neurons establish polarity and create synapses, and can be separated from the glia for use in various applications, such as electrophysiology, calcium imaging, cell survival assays, immunocytochemistry, and RNA/DNA/protein isolation.

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or ...

A Functional Motor Unit in the Culture Dish: Co-culture of Spinal Cord Explants and Muscle Cells

Human primary muscle cells cultured aneurally in monolayer rarely contract spontaneously because, in the absence of a nerve component, cell differentiation is limited and motor neuron stimulation is missing1. These limitations hamper the in vitro study of many neuromuscular diseases in cultured muscle cells. Importantly, the experimental constraints of monolayered, cultured muscle cells can be overcome by functional innervation of myofibers with spinal cord explants in co-cultures.
Here, we show the different steps required to achieve an efficient, proper innervation of human primary muscle cells, leading to complete differentiation and fiber contraction according to the method developed by Askanas2. To do so, muscle cells are co-cultured with spinal cord explants of rat embryos at ED 13.5, with the dorsal root ganglia still attached to the spinal cord slices. After a few days, the muscle fibers start to contract and eventually become cross-striated through innervation by functional neurites projecting from the spinal cord explants that connecting to the muscle cells. This structure can be maintained for many months, simply by regular exchange of the culture medium.
The applications of this invaluable tool are numerous, as it represents a functional model for multidisciplinary analyses of human muscle development and innervation. In fact, a complete de novo neuromuscular junction installation occurs in a culture dish, allowing an easy measurement of many parameters at each step, in a fundamental and physiological context.
Just to cite a few examples, genomic and/or proteomic studies can be performed directly on the co-cultures. Furthermore, pre- and post-synaptic effects can be specifically and separately assessed at the neuromuscular junction, because both components come from different species, rat and human, respectively. The nerve-muscle co-culture can also be performed with human muscle cells isolated from patients suffering from muscle or neuromuscular diseases3, and thus can be used as a screening tool for candidate drugs. Finally, no special equipment but a regular BSL2 facility is needed to reproduce a functional motor unit in a culture dish. This method thus is valuable for both the muscle as well as the neuromuscular research communities for physiological and mechanistic studies of neuromuscular function, in a normal and disease context.

Cultured muscle cells are an inadequate model to recapitulate innervated muscle in vivo. A functional motor unit can be reproduced in vitro by innervation of differentiated human primary muscle cells using rat embryo spinal cord explants. This article describes how co-cultures of spinal cord explants and muscle cells are established.

Human primary muscle cells cultured  aneurally in monolayer rarely contract spontaneously because, in the absence of  a nerve component, cell ...

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

Imaging Analysis of Neuron to Glia Interaction in Microfluidic Culture Platform (MCP)-based Neuronal Axon and Glia Co-culture System

Proper neuron to glia interaction is critical to physiological function of the central nervous system (CNS). This bidirectional communication is sophisticatedly mediated by specific signaling pathways between neuron and glia1,2 . Identification and characterization of these signaling pathways is essential to the understanding of how neuron to glia interaction shapes CNS physiology. Previously, neuron and glia mixed cultures have been widely utilized for testing and characterizing signaling pathways between neuron and glia. What we have learned from these preparations and other in vivo tools, however, has suggested that mutual signaling between neuron and glia often occurred in specific compartments within neurons (i.e., axon, dendrite, or soma)3. This makes it important to develop a new culture system that allows separation of neuronal compartments and specifically examines the interaction between glia and neuronal axons/dendrites. In addition, the conventional mixed culture system is not capable of differentiating the soluble factors and direct membrane contact signals between neuron and glia. Furthermore, the large quantity of neurons and glial cells in the conventional co-culture system lacks the resolution necessary to observe the interaction between a single axon and a glial cell.
	In this study, we describe a novel axon and glia co-culture system with the use of a microfluidic culture platform (MCP). In this co-culture system, neurons and glial cells are cultured in two separate chambers that are connected through multiple central channels. In this microfluidic culture platform, only neuronal processes (especially axons) can enter the glial side through the central channels. In combination with powerful fluorescent protein labeling, this system allows direct examination of signaling pathways between axonal/dendritic and glial interactions, such as axon-mediated transcriptional regulation in glia, glia-mediated receptor trafficking in neuronal terminals, and glia-mediated axon growth. The narrow diameter of the chamber also significantly prohibits the flow of the neuron-enriched medium into the glial chamber, facilitating probing of the direct membrane-protein interaction between axons/dendrites and glial surfaces.

Imaging Analysis of Neuron to Glia Interaction in Microfluidic Culture Platform MCP-based Neuronal Axon and Glia Co-culture System

This study describes the procedures of setting up a novel neuronal axon and (astro)glia co-culture platform. In this co-culture system, manipulation of direct interaction between a single axon (and single glial cell) becomes feasible, allowing mechanistic analysis of the mutual neuron to glial signaling.

Proper  neuron to glia interaction is critical to physiological function of the central  nervous system (CNS). This bidirectional communication is ...

Functional and Morphological Assessment of Diaphragm Innervation by Phrenic Motor Neurons

This protocol specifically focuses on tools for assessing phrenic motor neuron (PhMN) innervation of the diaphragm at both the electrophysiological and morphological levels. Compound muscle action potential (CMAP) recording following phrenic nerve stimulation can be used to quantitatively assess functional diaphragm innervation by PhMNs of the cervical spinal cord in vivo in anesthetized rats and mice. Because CMAPs represent simultaneous recording of all myofibers of the whole hemi-diaphragm, it is useful to also examine the phenotypes of individual motor axons and myofibers at the diaphragm NMJ in order to track disease- and therapy-relevant morphological changes such as partial and complete denervation, regenerative sprouting and reinnervation. This can be accomplished via whole-mount immunohistochemistry (IHC) of the diaphragm, followed by detailed morphological assessment of individual NMJs throughout the muscle. Combining CMAPs and NMJ analysis provides a powerful approach for quantitatively studying diaphragmatic innervation in rodent models of CNS and PNS disease.

Compound muscle action potential recording quantitatively assesses functional diaphragm innervation by phrenic motor neurons. Whole-mount diaphragm immunohistochemistry assesses morphological innervation at individual neuromuscular junctions. The goal of this protocol is to demonstrate how these two powerful methodologies can be used in various rodent models of spinal cord disease.

This protocol specifically focuses on tools for assessing phrenic motor neuron (PhMN) innervation of the diaphragm at both the electrophysiological and ...

High-resolution Volume Imaging of Neurons by the Use of Fluorescence eXclusion Method and Dedicated Microfluidic Devices

Volume is an important parameter regarding physiological and pathological characteristics of neurons at different time scales. Neurons are quite unique cells regarding their extended ramified morphologies and consequently raise several methodological challenges for volume measurement. In the particular case of in vitro neuronal growth, the chosen methodology should include sub-micrometric axial resolution combined with full-field observation on time scales from minutes to hours or days. Unlike other methods like cell shape reconstruction using confocal imaging, electrically-based measurements or Atomic Force Microscopy, the recently developed Fluorescence eXclusion method (FXm) has the potential to fulfill these challenges. However, although being simple in its principle, implementation of a high-resolution FXm for neurons requires multiple adjustments and a dedicated methodology. We present here a method based on the combination of fluorescence exclusion, low-roughness multi-compartments microfluidic devices, and finally micropatterning to achieve in vitro measurements of local neuronal volume. The high resolution provided by the device allowed us to measure the local volume of neuronal processes (neurites) and the volume of some specific structures involved in neuronal growth, such as growth cones (GCs).

Volume is an important parameter regarding physiological and pathological characteristics of cells. We describe a fluorescent exclusion method allowing full-field measurement of in vitro neuronal volume with sub-micrometric axial resolution required for the analysis of neurites and dynamic structures implied in neuronal growth.

Volume is an important parameter regarding physiological and pathological characteristics of neurons at different time scales. Neurons are quite unique ...

Data Acquisition and Analysis In Brainstem Evoked Response Audiometry In Mice

Brainstem evoked response audiometry (BERA) is of central relevance in the clinical neurophysiology. As other evoked potential (EP) techniques, such as visually evoked potentials (VEPs) or somatosensory evoked potentials (SEPs), the auditory evoked potentials (AEPs) are triggered by the repetitive presentation of identical stimuli, the electroencephalographic (EEG) response of which is subsequently averaged resulting in distinct positive (p) and negative (n) deflections. In humans, both the amplitude and the latency of individual peaks can be used to characterize alterations in synchronization and conduction velocity in the underlying neuronal circuitries. Importantly, AEPs are also applied in basic and preclinical science to identify and characterize the auditory function in pharmacological and genetic animal models. Even more, animal models in combination with pharmacological testing are utilized to investigate for potential benefits in the treatment of sensorineural hearing loss (e.g., age- or noise-induced hearing deficits). Here we provide a detailed and integrative description of how to record auditory brainstem-evoked responses (ABRs) in mice using click and tone-burst application. A specific focus of this protocol is on pre-experimental animal housing, anesthesia, ABR recording, ABR filtering processes, automated wavelet-based amplitude growth function analysis, and latency detection.

Brainstem evoked response audiometry is an important tool in clinical neurophysiology. Nowadays, brainstem evoked response audiometry is also applied in the basic science and preclinical studies involving both pharmacological and genetic animal models. Here we provide a detailed description of how auditory brainstem responses can be successfully recorded and analyzed in mice.

Brainstem evoked response audiometry (BERA) is of central relevance in the clinical neurophysiology. As other evoked potential (EP) techniques, such as ...

Migration, Chemo-Attraction, and Co-Culture Assays for Human Stem Cell-Derived Endothelial Cells and GABAergic Neurons

Role of brain vasculature in nervous system development and etiology of brain disorders is increasingly gaining attention. Our recent studies have identified a special population of vascular cells, the periventricular endothelial cells, that play a critical role in the migration and distribution of forebrain GABAergic interneurons during embryonic development. This, coupled with their cell-autonomous functions, alludes to novel roles of periventricular endothelial cells in the pathology of neuropsychiatric disorders like schizophrenia, epilepsy, and autism. Here, we have described three different in vitro assays that collectively evaluate the functions of periventricular endothelial cells and their interaction with GABAergic interneurons. Use of these assays, particularly in a human context, will allow us to identify the link between periventricular endothelial cells and brain disorders. These assays are simple, low cost, and reproducible, and can be easily adapted to any adherent cell type.

We present three simple in vitro assays-the long-distance migration assay, the co-culture migration assay, and chemo-attraction assay-that collectively evaluate the functions of human stem cell derived periventricular endothelial cells and their interaction with GABAergic interneurons.

Role of brain vasculature in nervous system development and etiology of brain disorders is increasingly gaining attention. Our recent studies have ...

Generation of Oligodendrocytes and Oligodendrocyte-Conditioned Medium for Co-Culture Experiments

In the central nervous system, oligodendrocytes are well-known for their role in axon myelination, that accelerates the propagation of action potentials through saltatory conduction. Moreover, an increasing number of reports suggest that oligodendrocytes interact with neurons beyond myelination, notably through the secretion of soluble factors. Here, we present a detailed protocol allowing purification of oligodendroglial lineage cells from glial cell cultures also containing astrocytes and microglial cells. The method relies on overnight shaking at 37 &#176;C, which allows selective detachment of the overlying oligodendroglial cells and microglial cells, and the elimination of microglia by differential adhesion. We then describe the culture of oligodendrocytes and production of oligodendrocyte-conditioned medium (OCM). We also provide the kinetics of OCM treatment or oligodendrocytes addition to purified hippocampal neurons in co-culture experiments, studying oligodendrocyte-neuron interactions.

Herein, we display an efficient method for the purification of oligodendrocytes and production of oligodendrocyte-conditioned medium that can be used for co-culture experiments.

In the central nervous system, oligodendrocytes are well-known for their role in axon myelination, that accelerates the propagation of action potentials ...

Axonal Transport of Organelles in Motor Neuron Cultures using Microfluidic Chambers System

Motor neurons (MNs) are highly polarized cells with very long axons. Axonal transport is a crucial mechanism for MN health, contributing to neuronal growth, development, and survival. We describe a detailed method for the use of microfluidic chambers (MFCs) for tracking axonal transport of fluorescently labeled organelles in MN axons. This method is rapid, relatively inexpensive, and allows for the monitoring of intracellular cues in space and time. We describe a step by step protocol for: 1) Fabrication of polydimethylsiloxane (PDMS) MFCs; 2) Plating of ventral spinal cord explants and MN dissociated culture in MFCs; 3) Labeling of mitochondria and acidic compartments followed by live confocal imagining; 4) Manual and semiautomated axonal transport analysis. Lastly, we demonstrate a difference in the transport of mitochondria and acidic compartments of HB9::GFP ventral spinal cord explant axons as a proof of the system validity. Altogether, this protocol provides an efficient tool for studying the axonal transport of various axonal components, as well as a simplified manual for MFC usage to help discover spatial experimental possibilities.

Axonal transport is a crucial mechanism for motor neuron health. In this protocol we provide a detailed method for tracking the axonal transport of acidic compartments and mitochondria in motor neuron axons using microfluidic chambers.

Motor neurons (MNs) are highly polarized cells with very long axons. Axonal transport is a crucial mechanism for MN health, contributing to neuronal ...