eoe sub articles

EoE Sub Articles

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
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 thrice with sterile, distilled H2O and three times with ethanol 99%. Then dry the coverslips at 37 &#176;C or under a 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>Using sterile forceps, carefully remove the AXIS Axon Isolation Devices from the package 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 may be damaged by the pressure applied.</li>
	<li>Sterilize the AXIS Axon Isolation Devices by immersing them in ethanol 70%. Then dry the 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 a well within a 6-well plate.</li>
	<li>Place the AXIS Axon Isolation Device onto the coverslip and press gently but firmly with forceps with bent ends to allow full adhesion between the isolation device and the glass coverslip.</li>
	<li>In each culture chamber, pipette 150 &#956;l of poly-D-lysine (0.1 mg/ml in sterile, distilled H2O). Place the microfluidic devices under a vacuum for 5 min, 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 at 37 &#176;C.</li>
	<li>Wash chambers three times with sterile, distilled H2O.</li>
	<li>Fill chambers with 150 &#956;l laminin working solution (Sigma-Aldrich, 5 &#956;g/ml, in phosphate-buffered saline (PBS) or serum-free medium), and incubate O/N at 37 &#176;C.</li>
	<li>Prepare 50 ml of medium for trigeminal ganglia cultures with the following composition: 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 with the following composition: 40 ml Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM)-F12, 10 ml fetal bovine serum (FBS, final concentration: 20%), 100 U/ml penicillin/streptomycin, 2 mM L-glutamine, 150 &#956;g/ml ascorbic acid.</li>
</ol>
2. Mouse Embryo Generation and Dissection
<ol>
	<li>Determine embryonic age based on the 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. Position the mouse&#39;s neck between the first and second fingers on a grid, and decisively pull the tail.</li>
	<li>Dissect the skin around the lower abdomen and open the abdomen using scissors. Locate the uterus: during the late stages of pregnancy, the uterus abundantly fills the abdominal cavity.</li>
	<li>Dissect out the uterus and place it in a tube filled with PBS on ice. When on ice, the tissue can be left for several hours. Discard the corpse of the mother according to institutional guidelines.</li>
	<li>Dissect 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 the lower jaw precisely, 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 trigeminal ganglia (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. Then remove 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 (Figure 2B). Use forceps to separate the TG from the trigeminal nerves. Eliminate the remnants of the trigeminal projections using the dissection needles as knives. 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 knives, 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 2C. Isolate the tooth germs using dissection needles and remove excess 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 &#956;l of the respective media.</li>
	<li>With forceps, gently transfer the dissected TG and tooth germs into the holes created by punching (Figure 2D). Make sure that the tooth germs do not float and that they sink until they contact the coverslips.</li>
	<li>Culture the samples in an 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>Co-cultures can be easily imaged by time-lapse microscopy during the culture period and can be maintained for over 10 days.</li>
	<li>After the culture period, wash the chambers by pipetting 150 &#956;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 &#956;l of paraformaldehyde 4% (in PBS) in one well per chamber; incubate at RT for 15 min. Wash the chambers twice with PBS as described in 3.6.</li>
	<li>Proceed with further analysis.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>AXIS Axon Isolation Devices</td>
			<td>Millipore</td>
			<td>AX15010-TC</td>
			<td>&#160;Microchannels of different lengths are available</td>
		</tr>
		<tr>
			<td>Laminin</td>
			<td>Sigma Aldrich&#160;</td>
			<td>L2020</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurobasal&#160;</td>
			<td>Gibco&#160;</td>
			<td>21103-049</td>
			<td></td>
		</tr>
		<tr>
			<td>B27</td>
			<td>Gibco&#160;</td>
			<td>17504</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant Mouse beta-NGF&#160;</td>
			<td>R&#38;D Systems&#160;</td>
			<td>1156-NG-100&#160;</td>
			<td>Human and Rat beta-NGF (R&#38;D Systems) are equivalent</td>
		</tr>
		<tr>
			<td>DMEM-F12&#160;</td>
			<td>Gibco&#160;</td>
			<td>11320-033</td>
			<td></td>
		</tr>
	</tbody>
</table>

analyzing tooth germ and trigeminal ganglia interactions using a microfluidic co-culture system

Source: Pagella, P., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/53114">Analysis of Developing Tooth Germ Innervation Using Microfluidic Co-culture Devices.</a> J. Vis. Exp. (2015).
This video demonstrates the utilization of a microfluidic device to study the interactions between tooth germ pulp and trigeminal ganglia. It showcases the placement of tissues inside the microfluidic device and provides a detailed visualization of trigeminal neurite growth toward the tooth germ through the device&#39;s microgrooves. Additionally, it highlights the role of tooth-derived molecular factors in promoting or inhibiting neurite extension into the tooth germ pulp, which is crucial for ensuring efficient postnatal tooth innervation.

<img alt="Figure 1" src="/files/ftp_upload/22432/22432_Figure1.jpg" /> 
Figure 1. Views of mounted microfluidic co-culture device (A) from above; (B) partially lateral. The culture chambers (cc) are highlighted in red (eosin) and blue (toluidine blue); microgrooves (mg) can be seen as a white line between the culture chambers. Ganglia and co-cultured organs/tissues are placed into the appropriate culture chamber via the punched holes (ph) in the device. Media are added and removed via the wells (mw).
<img alt="Figure 2" src="/files/ftp_upload/22432/22432_Figure2.jpg" /> 
Figure 2. (A) Lateral view of a mouse embryo head (embryonic day of development E14.5). The black line indicates where the lower jaw should be cut in order to preserve the integrity of both trigeminal ganglia and tooth germs. (B) Mouse embryo head (E14.5) after removal of lower jaw, skull, and telencephalon. Black arrows indicate trigeminal ganglia (TG). (C) The lower jaw of mouse embryo (E14.5) after removal of the tongue. Black arrows indicate the localization of the different tooth germs (Inc: incisor; FM: first molars). (D) Schematic representation of the microfluidic co-culture device.

Analyzing Tooth Germ and Trigeminal Ganglia Interactions Using a Microfluidic Co-culture System

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Preparation of Dissection ...

Take a microfluidic device coated with a synthetic polymer and an adhesion protein.
The device consists of wells and interconnected culture chambers separated by microgrooves.&#160;&#160;
The chambers contain punched holes for tissue placement.
Obtain the trigeminal ganglia, a collection of trigeminal neuron cell bodies, and tooth germs, prenatal structures that develop into adult teeth, from a mouse embryo.
Place the tissues in the holes and add the respective culture media.
Nutrients and vitamins in the tooth germ media facilitate tooth germ survival during culturing.
The coated device surface and growth factors in the trigeminal ganglia media facilitate neurite and axonal outgrowth.
The axons reach the tooth germ chamber through the microgrooves.&#160;&#160;&#160;&#160;&#160;&#160;&#160;
The tooth-derived signals lead to neuronal repulsion, allowing the axons to surround the tooth germ without innervating it.
Wash the culture chambers with buffer and fix the tissues with a fixative.
The co-culture is now ready for further analysis.

analyzing-tooth-germ-trigeminal-ganglia-interactions-using

Nanyang Technological University

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neuronal Culture Techniques

Co-culture and Interaction Studies

36 Views. Source: Pagella, P., et al. Analysis of Developing Tooth Germ Innervation Using Microfluidic Co-culture Devices. J. Vis. Exp. (2015). This video demonstrates the utilization of a microfluidic device to study the interactions between tooth germ pulp and trigeminal ganglia. It showcases the placement of tissues inside the microfluidic device and provides a detailed visualization of trigeminal neurite growth toward the tooth germ through the device's microgrooves. Additionally, it highlights t...

Video: Analyzing Tooth Germ and Trigeminal Ganglia Interactions Using a Microfluidic Co-culture System - Concept

Rewiring Neuronal Circuits: A New Method for Fast Neurite Extension and Functional Neuronal Connection

Brain and spinal cord injury may lead to permanent disability and death because it is still not possible to regenerate neurons over long distances and accurately reconnect them with an appropriate target. Here a procedure is described to rapidly initiate, elongate, and precisely connect new functional neuronal circuits over long distances. The extension rates achieved reach over 1.2 mm/h, 30-60 times faster than the in vivo rates of the fastest growing axons from the peripheral nervous system (0.02 to 0.04 mm/h)28 and 10 times faster than previously reported for the same neuronal type at an earlier stage of development4. First, isolated populations of rat hippocampal neurons are grown for 2-3 weeks in microfluidic devices to precisely position the cells, enabling easy micromanipulation and experimental reproducibility. Next, beads coated with poly-D-lysine (PDL) are placed on neurites to form adhesive contacts and pipette micromanipulation is used to move the resulting bead-neurite complex. As the bead is moved, it pulls out a new neurite that can be extended over hundreds of micrometers and functionally connected to a target cell in less than 1 h. This process enables experimental reproducibility and ease of manipulation while bypassing slower chemical strategies to induce neurite growth. Preliminary measurements presented here demonstrate a neuronal growth rate far exceeding physiological ones. Combining these innovations allows for the precise establishment of neuronal networks in culture with an unprecedented degree of control. It is a novel method that opens the door to a plethora of information and insights into signal transmission and communication within the neuronal network as well as being a playground in which to explore the limits of neuronal growth. The potential applications and experiments are widespread with direct implications for therapies that aim to reconnect neuronal circuits after trauma or in neurodegenerative diseases.

This procedure describes how to rapidly initiate, extend and connect neurites organized in microfluidic chambers using poly-D-lysine-coated beads fixed to micropipettes that guide neurite elongation.

Brain and spinal cord injury may lead to permanent disability and death because it is still not possible to regenerate neurons over long distances and ...

JoVE Journal

Developing HiPSC Derived Serum Free Embryoid Bodies for the Interrogation of 3-D Stem Cell Cultures Using Physiologically Relevant Assays

Although a number of in vitro disease models have been developed using hiPSCs, one limitation is that these two-dimensional (2-D) systems may not represent the underlying cytoarchitectural and functional complexity of the affected individuals carrying suspected disease variants. Conventional 2-D models remain incomplete representations of in vivo-like structures and do not adequately capture the complexity of the brain. Thus, there is an emerging need for more 3-D hiPSC-based models that can better recapitulate the cellular interactions and functions seen in an in vivo system.
Here we report a protocol to develop a 3-D system from undifferentiated hiPSCs based on the serum free embryoid body (SFEB). This 3-D model mirrors aspects of a developing ventralized neocortex and allows for studies into functions integral to living neural cells and intact tissue such as migration, connectivity, communication, and maturation. Specifically, we demonstrate that the SFEBs using our protocol can be interrogated using physiologically relevant and high-content cell based assays such as calcium imaging, and multi-electrode array (MEA) recordings without cryosectioning. In the case of MEA recordings, we demonstrate that SFEBs increase both spike activity and network-level bursting activity during long-term culturing. This SFEB protocol provides a robust and scalable system for the study of developing network formation in a 3-D model that captures aspects of early cortical development.

Here we report a protocol to develop a three-dimensional (3-D) system from human induced-pluripotent stem cells (hiPSCs) called the serum free embryoid body (SFEB). This 3-D model can be used like an organotypic slice culture to model human cortical development and for the physiological interrogation of developing neural circuits.

Although a number of in vitro disease models have been developed using hiPSCs, one limitation is that these two-dimensional (2-D) systems may not ...

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 ...

Analyzing Tooth Germ and Trigeminal Ganglia Interactions Using a Microfluidic Co-culture System

Transcript

Analyzing Tooth Germ and Trigeminal Ganglia Interactions Using a Microfluidic Co-culture System

Rewiring Neuronal Circuits: A New Method for Fast Neurite Extension and Functional Neuronal Connection

Developing HiPSC Derived Serum Free Embryoid Bodies for the Interrogation of 3-D Stem Cell Cultures Using Physiologically Relevant Assays

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