eoe sub articles

EoE Sub Articles

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Animal preparation
<ol>
	<li>Induce anesthesia by injecting ketamine (70 mg/kg) and xylazine (16 mg/kg) intraperitoneally. Maintain normal body temperature, contributing to quick induction of deep anesthesia. Test for adequate sedation with a pain stimulus, such as a toe pinch, and verify the absence of a reaction.</li>
	<li>Carefully shave the neck hair of the mouse with a razor, clean it with 70% ethanol followed by betadine/chlorhexidine, and apply 1% lidocaine on the skin surface for local pain control.</li>
	<li>Place the mouse in a supine position. Use tape to fix the limbs and tail, gently stretching the skin of the neck to the opposite side of the surgery. Simultaneously, elevate the neck slightly.</li>
	<li>Use ophthalmic ointment (e.g., 5% dexpanthenol) to prevent dehydration of the eyes during the operation.</li>
</ol>
2. SAH induction
<ol>
	<li>Open the neck skin with a sterile scalpel, from the chin to the upper edge of the breastbone (1.5 cm), and bluntly separate salivary glands from their surrounding connective tissue.</li>
	<li>Separate the muscle group along one side [in this case, the right side] of the trachea, exposing the common carotid artery (CCA) sheath covered with nourishing blood vessels and venules. The CCA and the nerves are located in close proximity to each other.</li>
	<li>Dissociate the CCA and leave a free 8-0 silk suture around the CCA without ligating it in advance. Pay attention to the protection of the vagal nerve, as it is easily damaged (Figure 1A).</li>
	<li>A triple bifurcation of the CCA, the internal carotid artery (ICA), and the external carotid artery (ECA) is visible along the lower posterior third of the diastasis. Dissect the distal end of the ECA and ligate the vessel twice as far distaally as possible.</li>
	<li>Disconnect the ECA at the midpoint of the twice ligated segment, creating a vessel stump.</li>
	<li>Prearrange one ligation for the filament around the ECA stump, do not close it until successful filament insertion.</li>
	<li>Use a suture or micro clip to occlude the ICA and CCA temporarily (Figure 1B).</li>
	<li>Make a small incision (approximately half of the ECA diameter) in the ECA using microvascular scissors. Insert a 5-0 (alternatively 4-0) prolene filament into the ECA and advance it into the CCA.</li>
	<li>Close the ligature on the ECA slightly while loosening the micro clip on the ICA and CCA (Figure 1C).</li>
	<li>Gently pull back on the filament and adjust the ECA stump in the cranial direction, invaginating the filament through the bifurcation into the ICA (Figure 1D).</li>
	<li>Point the filament tip medially at an angle of ~30&#176; to the tracheal midline and ~30&#176; to the horizontal plane. Push the filament forward inside the ICA. After reaching the anterior cerebral artery (ACA)- middle cerebral artery (MCA) bifurcation, resistance is encountered (~9 mm).</li>
	<li>Advance the filament 3 mm further, perforating the right ACA. Promptly withdraw the filament to the ECA stump, allowing blood flow into the subarachnoid space.</li>
	<li>Keep the filament in this position for about 10 s (Figure 1E). The presence of muscle tremors, ipsilateral miosis, gasping for breath, altered heart rhythm, and urinary incontinence can be supporting evidence of successful surgery.</li>
	<li>Temporarily close the CCA to avoid excess blood loss. Pull out the filament instantly and ligate the ECA with the prearranged suture. Reopen the CCA and allow reperfusion and further effusion of blood into the subarachnoid space (Figure 1F).</li>
	<li>After checking for bleeding leakage, disinfect the skin surrounding the wound to prevent postoperative skin infections, and suture the wound with a non-absorbable 4-0 polyester fiber suture.</li>
	<li>Place the mouse in a thermal box until consciousness is regained. Wait until the animal is fully awake and ensure it has regained sufficient consciousness to maintain sternal recumbency. Do not return animals to the company of other mice until fully recovered.</li>
	<li>Administer 200-300 mg/kg body weight&#160;paracetamol for postoperative pain relief.</li>
	<li>Check on the mice daily after surgery.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Eye cream</td>
			<td>Bayer</td>
			<td>815529836</td>
			<td>Bepanthen</td>
		</tr>
		<tr>
			<td>Ligation suture (5-0)</td>
			<td>SMI</td>
			<td></td>
			<td>Silk black USP</td>
		</tr>
		<tr>
			<td>Light source for microscope</td>
			<td>Zeiss</td>
			<td>CL 6000 LED</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>CP-pharma</td>
			<td>797-037</td>
			<td>100 mg/mL</td>
		</tr>
		<tr>
			<td>Paracetamol (40 mg/mL)</td>
			<td>bene Arzneimittel</td>
			<td>4993736</td>
			<td></td>
		</tr>
		<tr>
			<td>Prolene filament (5-0)</td>
			<td>Erhicon</td>
			<td>EH7255</td>
			<td></td>
		</tr>
		<tr>
			<td>Razor</td>
			<td>Wella</td>
			<td>HS61</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical instrument (Fine Scissors)</td>
			<td>FST</td>
			<td>14060-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical instrument (forceps#1)</td>
			<td>AESCULAP</td>
			<td>FM001R</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical instrument (forceps#2)</td>
			<td>AESCULAP</td>
			<td>FD2855R</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical instrument (forceps#3)</td>
			<td>Hammacher</td>
			<td>HCS 082-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical instrument (Needle holder)</td>
			<td>FST</td>
			<td>91201-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical instrument (Vannas Spring Scissors)</td>
			<td>FST</td>
			<td>15000-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical microscope</td>
			<td>Zeiss</td>
			<td>Stemi 2000 C</td>
			<td></td>
		</tr>
		<tr>
			<td>Wounding suture(4-0)</td>
			<td>Erhicon</td>
			<td>CB84D</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylavet</td>
			<td>CP-pharma</td>
			<td>797-062</td>
			<td>20 mg/mL</td>
		</tr>
	</tbody>
</table>

induction of a subarachnoid hemorrhage in a mouse model through endovascular perforation

Source: Liu, S., et al. <a href="https://app.jove.com/v/63150">Endovascular Perforation Model for Subarachnoid Hemorrhage Combined with Magnetic Resonance Imaging (MRI).</a>&#160;J. Vis. Exp. (2021).
The video demonstrates the induction of subarachnoid hemorrhage via endovascular perforation in a mouse model. An incision is made in the neck region of the anesthetized mouse to access the common carotid artery and its bifurcation. A filament is inserted through the arteries to perforate an intracranial artery branch, inducing subarachnoid hemorrhage.

<img alt="Figure 1" src="/files/ftp_upload/23102/23102_Figure1.jpg" /> 
Figure 1:&#160;Step-by-step images of surgical technique.&#160;(A) Depiction of the exposed right carotid artery anatomy: the CCA and its bifurcation into ICA and ECA are identified, as well as the small branches of the ECA (OA and STA). (B) The ECA is mobilized from the surrounding tissue and ligated with two sutures before cutting it. A third ligation needs to ...

Induction of a Subarachnoid Hemorrhage in a Mouse Model through Endovascular Perforation

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Animal ...

Take an anesthetized mouse with a shaved neck.
Disinfect the neck and make a midline incision.
Expose the common carotid artery, or CCA, which bifurcates into the external carotid artery, or ECA, and internal carotid artery, or ICA.
Ligate the ECA and temporarily occlude both the CCA and ICA to restrict blood flow.
Incise the ECA, insert a filament, and advance it into the CCA.
Reopen the CCA and ICA to restore blood flow. Redirect the filament through the ICA toward the brain to perforate the anterior cerebral artery, a branch of the ICA.
The perforation causes bleeding, or hemorrhage, into the subarachnoid space, a fluid-filled area surrounding the brain. Confirm the hemorrhage by observing clinical symptoms.
Close the CCA to stop blood flow, remove the filament, and ligate the ECA incision.
Reopen the CCA to continue subarachnoid bleeding.
Suture the skin and allow the mouse to recover for analysis.

induction-subarachnoid-hemorrhage-mouse-model-through-endovascular

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neuropathology

Cerebrovascular Diseases

39 vues. Source : Liu, S., et al. Modèle de perforation endovasculaire pour l’hémorragie sous-arachnoïdienne combiné à l’imagerie par résonance magnétique (IRM).J. Vis. Exp. (2021). La vidéo démontre l’induction de l’hémorragie sous-arachnoïdienne par perforation endovasculaire dans un modèle murin. Une incision est pratiquée dans la région du cou de la souris anesthésiée pour accéder à l’artère carotide commune et à sa bifurcation. Un filament est inséré dans les artère...

Vidéo: Induction d’une hémorragie sous-arachnoïdienne dans un modèle murin par perforation endovasculaire - Concept

A Volumetric Method for Quantification of Cerebral Vasospasm in a Murine Model of Subarachnoid Hemorrhage

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor determining patient outcome and is therefore frequently taken as a study endpoint. However, in small animal studies on SAH, quantification of cerebral vasospasm is a major challenge. Here, an ex vivo method is presented that allows quantification of volumes of entire vessel segments, which can be used as an objective measure to quantify cerebral vasospasm. In a first step, endovascular casting of the cerebral vasculature is performed using a radiopaque casting agent. Then, cross-sectional imaging data are acquired by micro computed tomography. The final step involves 3-dimensional reconstruction of the virtual vascular tree, followed by an algorithm to calculate center lines and volumes of the selected vessel segments. The method resulted in a highly accurate virtual reconstruction of the cerebrovascular tree shown by a diameter-based comparison of anatomical samples with their virtual reconstructions. Compared with vessel diameters alone, the vessel volumes highlight the differences between vasospastic and non-vasospastic vessels shown in a series of SAH and sham-operated mice.

The goal of this article is to present a method that allows a 3-dimensional reconstruction of the cerebrovascular tree in mice after micro computed tomography and determination of volumes of entire vessel segments that can be used to quantify cerebral vasospasm in murine models of subarachnoid hemorrhage.

Une méthode volumétrique pour la Quantification du vasospasme cérébral dans un modèle murin d’hémorragie sous-arachnoïdienne

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor ...

Recherche

JoVE Journal

Neurosciences

Double Direct Injection of Blood into the Cisterna Magna as a Model of Subarachnoid Hemorrhage

Among strokes, subarachnoid hemorrhage (SAH) consecutive to the rupture of a cerebral arterial aneurysm represents 5-9% but is responsible for about 30% of the total stroke-related mortality with an important morbidity in terms of neurological outcome. A delayed cerebral vasospasm (CVS) may occur most often in association with a delayed cerebral ischemia. Different animal models of SAH are now being used including endovascular perforation and direct injection of blood into the cisterna magna or even the prechiasmatic cistern, each exhibiting distinct advantages and disadvantages. In this article, a standardized mouse model of SAH by double direct injection of determined volumes of autologous whole blood into the cisterna magna is presented. Briefly, mice were weighed and then anesthetized by isoflurane inhalation. Then, the animal was placed in a reclining position on a heated blanket maintaining a rectal temperature of 37 &#176;C and positioned in a stereotactic frame with a cervical bend of about 30&#176;. Once in place, the tip of an elongated glass micropipette filled with the homologous arterial blood taken from carotid artery of another mouse of the same age and gender (C57Bl/6J) was positioned at a right angle in contact with the atlanto-occipital membrane by means of a micromanipulator. Then 60 &#181;L of blood was injected in the cisterna magna followed by a 30&#176; downward tilt of the animal for 2 minutes. The second infusion of 30 &#181;L of blood into the cisterna magna was performed 24 h after the first one. The individual follow-up of each animal is carried out daily (careful evaluation of weight and well-being). This procedure allows a predictable and highly reproducible distribution of blood, likely accompanied by intracranial pressure elevation that can be mimicked by an equivalent injection of an artificial cerebral spinal fluid (CSF), and represents an acute to mild-model of SAH inducing low mortality.

We described in this protocol a standardized subarachnoid hemorrhage (SAH) mouse model by a double injection of autologous whole-blood into the cisterna magna. The high degree of standardization of the double-injection procedure represents a middle-to-acute model of SAH with relative safety regarding mortality.

Double injection directe de sang dans la Cisterna Magna comme modèle d’hémorragie subarachnoïde

Among strokes, subarachnoid hemorrhage (SAH) consecutive to the rupture of a cerebral arterial aneurysm represents 5-9% but is responsible for about 30% ...

Analysis of Cerebral Vasospasm in a Murine Model of Subarachnoid Hemorrhage with High Frequency Transcranial Duplex Ultrasound

Cerebral vasospasm that occurs in the weeks after subarachnoid hemorrhage, a type of hemorrhagic stroke, contributes to delayed cerebral ischemia. A problem encountered in experimental studies using murine models of SAH is that methods for in vivo monitoring of cerebral vasospasm in mice are lacking. Here, we demonstrate the application of high frequency ultrasound to perform transcranial Duplex sonography examinations on mice. Using the method, the internal carotid arteries (ICA) could be identified. The blood flow velocities in the intracranial ICAs were accelerated significantly after induction of SAH, while blood flow velocities in the extracranial ICAs remained low, indicating cerebral vasospasm. In conclusion, the method demonstrated here allows functional, noninvasive in vivo monitoring of cerebral vasospasm in a murine SAH model.

The aim of this manuscript is to present a sonography-based method that allows in vivo imaging of blood flow in cerebral arteries in mice. We demonstrate its application to determine changes in blood flow velocities associated with vasospasm in murine models of subarachnoid hemorrhage (SAH).

Analyse du vasospasme cérébral dans un modèle murin d’hémorragie sous-arachnoïdienne avec ultrason duplex transcrânien à haute fréquence

Cerebral vasospasm that occurs in the weeks after subarachnoid hemorrhage, a type of hemorrhagic stroke, contributes to delayed cerebral ischemia. A ...

Induction of a Subarachnoid Hemorrhage in a Mouse Model through Endovascular Perforation

Transcription

Induction of a Subarachnoid Hemorrhage in a Mouse Model through Endovascular Perforation