We thank the PRIMACEN platform (Normandie Rouen University, France) for imaging equipment and Mr. Arnaud Arabo, Mrs Julie Maucotel and Mrs Martine Dubois, for animal housing and care. We thank Mrs. Celeste Nicola for lending her voice to the videotaping of the protocol. This work was supported by Seinari Normandy maturation program, Fondation AVC under the aegis of the FRM, Normandie Rouen University and Inserm. The Normandy Region and the European Union (3R project). Europe gets involved in Normandy with European Regional Development Fund (ERDF).

All procedures were performed under the supervision of H. Castel in accordance with the French Ethical Committee and guidelines of European Parliament Directive 2010/63/EU and the Council for the Protection of Animals Used for Scientific Purposes. This project was approved by the local CENOMEXA and the national ethic committees on animal research and testing. Male C57Bl/6J Rj mice (Janvier), aged 8&#8211;12 weeks, were housed under controlled standard environmental conditions: 22 &#176;C &#177; 1 &#176;C, 12 hours/12 hours light/dark cycle, and water and food available ad libitum.
1. Setup of SAH surgery and preparation for injection
<ol>
	<li>Before the beginning of surgery, pull an adequate number of glass capillaries by using a micropipette puller. The injection pipette should exhibit an inner diameter of 0.86 mm and outer diameter of 1.5 mm.</li>
	<li>Prepare the artificial cerebrospinal fluid (aCSF) for the sham condition.
	<ol>
		<li>Prepare a solution with 119 mM NaCl, 2.5 mM KCl, 1 mM NaH2PO4, 1.3 mM MgCl2, 10 mM glucose, 26.2 mM NaHCO3 in H2O, pH 7.4.</li>
		<li>Gas aCSF with 95% O2 and 5% CO2 for 15 min, and then add 2.5 mM CaCl2.</li>
		<li>Sterilize the oxygenated aCSF with a 0.22 &#181;m filter apparatus. The aCSF solution can be stable for 3-4 weeks at 4 &#176;C. If contamination (solution becoming cloudy) or a deposit formation has appeared, discard and make fresh aCSF.</li>
	</ol>
	</li>
	<li>Collection of blood from a homologous mouse donor
	<ol>
		<li>Isolate the carotid artery along the trachea and collect the maximum amount of blood by puncture of the carotid artery.</li>
		<li>In practice, place the mouse into an anesthesia chamber and load the chamber with 5% isoflurane until the animal loses consciousness.</li>
		<li>Check the lack of reflexes by clamping one of two hind limbs to allow the setting of the surgical experimental procedure.</li>
		<li>Coat a 1 mL syringe with a heparin solution by using a 26 G needle (heparin sodium). This will prevent blood coagulation during the next steps.</li>
		<li>Install the animal positioned in dorsal decubitus with legs apart, and the nose in an anesthesia mask (anesthesia maintenance with 2 to 2.5% isoflurane).</li>
		<li>Isolate the carotid artery along the trachea by dissecting the omohyoid muscle longitudinally. Once artery isolated, insert the needle towards the heart with the help of the microdissecting hook and forceps and collect the maximum of blood via puncture of the carotid artery (60 &#181;L is needed per SAH mouse).</li>
		<li>Sacrifice the anesthetized donor mouse immediately after blood collection by using cervical dislocation.</li>
	</ol>
	</li>
</ol>
2. Animal (8-10-week-old C57BL/6J male mice) preparation
<ol>
	<li>Weigh each mouse precisely using an electronic balance. In the current study, mice would have body weight within the range of 20 to 25 grams just before surgery.</li>
	<li>As previously explained (see steps 1.3.2 and 1.3.3), induce anesthesia of mice to be operated.</li>
	<li>Shave the neck and the space between the ears with a suitable electric clipper.</li>
	<li>Install the animal positioned in ventral decubitus with legs apart and the nose in an anesthesia mask (anesthesia maintenance with 2 and 2.5% isoflurane) on a stereotactic frame.</li>
	<li>Check that the mouse is sleeping and that his head is properly blocked.</li>
	<li>Subcutaneously inject 100 &#181;L of buprenorphine (0.1 mg/kg) with a 26 G needle in the lower back, to avoid pain after awakening.</li>
	<li>Prevent dry eyes by using protective liquid gel and maintain an intrarectal temperature of 37 &#176;C by using an auto-regulated electric blanket.</li>
	<li>Treat the posterior neck shaved area with an antiseptic solution (povidone-iodine or chlorhexidine by using a sterile cotton road).</li>
	<li>Pre-sterilize all instruments touching the prepared skin/subcutaneous tissue (heating to 200 &#176;C for 2 hours) and handle aseptically.</li>
</ol>
3. SAH induction
<ol>
	<li>On the first day (D-1)
	<ol>
		<li>Cut a 1 cm incision with thin scissors in the posterior neck, followed by the separation of muscles along the midline to access the cisterna magna.</li>
		<li>Cut the tip of the empty glass pipette with thin scissors. Then, adapt to a syringe connected to a flexible silicone connector.</li>
		<li>Transfer 60 &#181;L of blood or aCSF (for SAH or sham condition, respectively) in a 0.5 mL tube using a precision micropipette.</li>
		<li>Suck into the glass pipette the 60 &#181;L of blood for the SAH condition or 60 &#181;L of aCSF for the sham condition.</li>
		<li>For injection, install the pipette on the stereotactic frame using a ring or blue-tack and slowly bring the pipette tip to the membrane at the interface with the cisterna magna.</li>
		<li>Slowly insert the pipette tip through the atlanto-occipital membrane into the cisterna magna, using a micro-manipulator of the stereotactic frame.</li>
		<li>Connect the pipette previously filled with blood or aCSF to the syringe ready for pressure induction.</li>
		<li>Inject by pressing the plunger at a low rate around 10 &#181;L/min, to avoid acute intracranial pressure.</li>
		<li>During injection, closely monitor the respiratory rate and the rectal temperature.</li>
		<li>At the end of the injection, carefully take off the pipette via the micro-manipulator and visually ensure that there is no leak during withdrawal.</li>
		<li>Achieve hemostasis using an absorbable hemostat and run two sutures with braided non-absorbable suturing thread.</li>
		<li>Immediately after surgery, isolate and position the mouse in decline decubitus and cover it with a survival blanket in an open box for the duration of recovery.</li>
	</ol>
	</li>
	<li>Second day of induction (D0)
	<ol>
		<li>After 24 hours, induce anesthesia (see steps 1.3.2 and 1.3.3). Subcutaneously inject 100 &#181;L of buprenorphine again (0.1 mg/kg) and prevent dry eyes by using protective liquid gel (see steps 2.7 and 2.8).</li>
		<li>Install the animal on the stereotactic frame as the day before.</li>
		<li>Carefully remove the sutures with microscissors.</li>
		<li>Prepare the atlanto-occipital membrane as before and apply antiseptic preparation on the shaved area of the neck with a sterile cotton rod.</li>
		<li>Inject 30 &#181;L of blood or aCSF at a low rate (see steps 3.1.2 to 3.1.8). Monitor the respiratory rate and the rectal temperature.</li>
		<li>At the end of the injection, carefully take off the pipette and control the absence of blood leak during withdrawal.</li>
		<li>Achieve hemostasis and run two sutures with braided absorbable suturing thread.</li>
	</ol>
	</li>
</ol>
4. Postoperative follow-up and end of the experiment
<ol>
	<li>Immediately after surgery, isolate and position the mouse in decline decubitus with a survival blanket on its back in an open box during recovery.</li>
	<li>Weigh and carefully observe daily the behavior of each mouse until sacrifice (e.g., D7 post-surgery).</li>
	<li>Among humane endpoints, a significant weight loss (&#62;15% of the weight) is classically noticed. A &#34;hunched back&#34; posture, slow movements, prostration, abnormal vocalizations of hurt and/or significant aggressive behavior are also important signs of animal suffering. If any of these signs or a combination of signs appears, the monitoring of the animal is reinforced within hours of their appearance. If the animal&#39;s welfare worsens or does not improve within 48 hours, it will be considered that a level of intolerable suffering is reached, and euthanasia is carried out.</li>
	<li>At the time of choice, sacrifice anesthetized mice by decapitation, and harvest brains for further analyses.</li>
	<li>Perform euthanasia (decapitation) after isoflurane anesthesia (5%).</li>
</ol>

The authors have nothing to disclose.

Despite the intensity of the research in the field of SAH and the development of therapeutic strategies such as endovascular and pharmacological treatment options increasing over the past twenty years, mortality remains high within the first week of hospital admission and reaches about 50% during the following 6 months24,25. This current preclinical model by daily double injection of homologous arterial blood into the cisterna magna has been reco...

Subarachnoid hemorrhage (SAH) accounts for up to 5% of all stroke cases and constitutes a relatively common pathology with an incidence of 7.2 to 9 patients per 100,000 per year, with a mortality rate of 20%-60% depending on the study1,2,3. In the acute phase, the mortality is attributable to the severity of bleeding, rebleeding, cerebral vasospasm (CVS) and/or medical complications4. In survivors, early brain injury (EBI) is associated with parenchymal extension of hemorrhage and abrupt increase in intracranial pressure, which may result in primary cerebral ischemia5 and immediate death in about 10%-15% of cases6. After the initial &#34;acute&#34; stage of SAH, the prognosis depends on the occurrence of &#34;secondary&#34; or delayed cerebral ischemia (DCI), detected in nearly 40% of patients by cerebral computed tomography, and in up to 80% of patients after magnetic resonance imaging (MRI)7,8. In addition to the CVS occurring between 4 to 21 days after aneurysm rupture in a majority of SAH patients, DCI9 may result from multifactorial diffuse brain lesions secondary to microthrombosis formation, reduced cerebral perfusion, neuroinflammation, and cortical spreading depression (CSD)10,11,12,13. This affects 30% of SAH survivors and impacts cognitive functions including visual memory, verbal memory, reaction time, and executive, visuospatial and language functions14 impairing daily life15. Current standard therapies to prevent CVS and/or the poor cognitive outcomes in SAH patients are based on the blockage of Ca2+ signaling and vasoconstriction by using Ca2+ channel inhibitors as Nimodipine. However, more recent clinical trials targeting vasoconstriction revealed dissociation between patient&#8217;s neurological outcome and prevention of CVS16, suggesting more complex pathophysiological mechanisms involved in SAH-long-term consequences. Therefore, there is a medical need for greater understanding of the number of pathological events accompanying SAH and the development of valid and standardized animal models to test original therapeutic interventions.
The rupture of an intracranial aneurysm mostly responsible for SAH in humans is likely difficult to mimic in preclinical animal models. Currently, the aneurysm rupture and SAH situation can tentatively be tested by the perforation of the middle cerebral artery (endovascular puncture model) responsible for CVS and sensitivomotor dysfunctions in mice17,18. Due to the lack of any possible control over the onset of bleeding and the diffusion of blood in this model, other methods have been developed in rodents to generate SAH models without endovascular rupture. More precisely, they consist of the direct administration of arterial blood into the subarachnoid space through a single or a double injection in the magna cisterna19 or a single injection into the prechiasmatic cistern20. The main advantage of these mouse models without endovascular rupture is the possibility to reproducibly master the surgical procedure and the quality and quantity of the injected blood sample. Another advantage of this model over the model by endovascular perforation in particular is the preservation of the general well-being of the animal. As a matter of fact, this surgery is less invasive and technically less challenging than that required to generate a carotid wall rupture. In this last model, the animal has to be intubated and mechanically ventilated, while a monofilament is inserted in the external carotid artery, and advanced into the internal carotid artery. This likely leads to transient ischemia due to vessel obstruction by the wire path. Consequently, the co-morbidity (moribund state, important pain and death) associated with surgery is less important in double injection model compared with endovascular perforation model. In addition to being a more consistent SAH, the double direct injection method complies with the animal welfare in research and testing (reduced time under anesthesia, pain from tissue disruption in surgery and distress) and leads to a minimum total number of animals used for the protocol study and personnel training.
Moreover, this allows implementation of the same protocol to transgenic mice, leading to an optimized pathological understanding of the SAH and the possibility of comparative testing of potential therapeutic compounds. Here, we present a standardized mouse model of subarachnoid hemorrhage (SAH) by a double daily consecutive injection of autologous arterial blood into the cisterna magna in 6-8 weeks-old male C57Bl/6J mice. The main advantage of this model is the control of the bleeding volume compared with the endovascular perforation model, and the reinforcing of the bleeding event without a drastic increase of intracranial pressure21. Recently, the double direct injection of blood into the cisterna magna has been well described on the experimental and physiopathological issues in mice. Indeed, we recently demonstrated CVS of large cerebral arteries (basilar (BA), middle (MCA) and anterior (ACA) cerebral arteries), cerebrovascular fibrin deposition and cell apoptosis from day 3 (D3) to 10 (D10), circulation defects of paravascular cerebrospinal fluid accompanied by altered sensitivomotor and cognitive functions in mice, 10 days post-SAH in this model22. Thus, it makes this model mastered, validated and characterized for short-term and long-lasting events post-SAH. It should be ideally suitable for prospective identification of new targets and for studies on potent and efficient therapeutic strategies against SAH-associated complications.

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		<col />
		<col />
		<col />
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	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">absorbable hemostat</td>
			<td style="width:201px;">Ethicon</td>
			<td style="width:161px;">Surgicel</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">absorbable suturing thread</td>
			<td style="width:201px;">Ethicon</td>
			<td style="width:161px;">Vicryl 5.0</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">auto-regulated electric blanket</td>
			<td style="width:201px;">Harvard Apparatus</td>
			<td style="width:161px;">50-7087-F</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">bluetack for capillary fixation</td>
			<td style="width:201px;">UHU</td>
			<td style="width:161px;">Patafix</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">electronic balance</td>
			<td style="width:201px;">Denver Instrument</td>
			<td style="width:161px;">MXX-2001</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:247px;">glass capillaries</td>
			<td style="width:201px;">Harvard Apparatus</td>
			<td style="width:161px;">GC150F-15</td>
			<td style="width:167px;">inner diameter 0.86 mm 
			outer diameter 1.5 mm</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">isoflurane vaporizer</td>
			<td style="width:201px;">Phymep</td>
			<td style="width:161px;">V100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">micropipette puller</td>
			<td style="width:201px;">Sutter Instrument Company</td>
			<td style="width:161px;">P-97</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">needle 26 G</td>
			<td style="width:201px;">BD microbalance</td>
			<td style="width:161px;">300300</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">non absorbable suturing thread</td>
			<td style="width:201px;">Peters surgical</td>
			<td style="width:161px;">Filapeau 4.0</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">stereotaxic frame</td>
			<td style="width:201px;">David Kopf instruments</td>
			<td style="width:161px;">Model 902</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:247px;">surgical equipment</td>
			<td style="width:201px;">Kent scientific</td>
			<td style="width:161px;">clamp, microscissors, thin scissors</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">syringe 20 mL TERUMO</td>
			<td style="width:201px;">Thermofisher</td>
			<td>11866071</td>
			<td></td>
		</tr>
	</tbody>
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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.

Experimental timeline, procedure, follow-up and mortality 
Figure 1A and Figure 1B summarize the SAH model protocol by double intracisternal injection of blood. Briefly, on the first day of SAH induction (D-1), 60 &#181;L of blood withdrawn from a homologous mouse or 60 &#181;L of artificial cerebrospinal fluid (aCSF) were injected into the cisterna magna in SAH or sham conditions, respectively. The next day (D0), 30 &#181;L of blood withdr...

Watch this Scientific Journal Video about Double Direct Injection of Blood into the Cisterna Magna as a Model of Subarachnoid Hemorrhage at JoVE.com

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

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.

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

double-direct-injection-blood-into-cisterna-magna-as-model

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

10.2K Views.  University of Rouen Normandy, INSERM U1239, DC2N, Institute for Research and Innovation in Biomedicine (IRIB), Rouen, France. 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.

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

Autologous Blood Injection to Model Spontaneous Intracerebral Hemorrhage in Mice

Investigation of the pathophysiology of injury after intracerebral
hemorrhage (ICH) requires a reproducible animal model. While ICH
accounts for 10-15% of all strokes, there remains no specific
effective therapy. The autologous blood injection model in mice
involves the stereotaxic injection of arterial blood into the basal
ganglia mimicking a spontaneous hypertensive hemorrhage in man. The
response to hemorrhage can then be studied in vivo and the
neurobehavioral deficits quantified, allowing for description of the
ensuing pathology and the testing of potential therapeutic agents. The
procedure described in this protocol uses the double injection
technique to minimize risk of blood reflux up the needle track, no
anticoagulants in the pumping system, and eliminates all dead space
and expandable tubing in the system.

The autologous blood injection model of intracerebral hemorrhage in mice described in this protocol uses the double injection technique to minimize risk of blood reflux up the needle track, no anticoagulants in the pumping system, and eliminates all dead space and expandable tubing in the system.

Investigation of the pathophysiology of injury after intracerebral
hemorrhage (ICH) requires a reproducible animal model.  While ICH
accounts for 10-15% ...

Cannula Implantation into the Cisterna Magna of Rodents

Cisterna magna cannulation (CMc) is a straightforward procedure that enables direct access to the cerebrospinal fluid (CSF) without operative damage to the skull or the brain parenchyma. In anesthetized rodents, the exposure of the dura mater by blunt dissection of the neck muscles allows the insertion of a cannula into the cisterna magna (CM). The cannula, composed either by a fine beveled needle or borosilicate capillary, is attached via a polyethylene (PE) tube to a syringe. Using a syringe pump, molecules can then be injected at controlled rates directly into the CM, which is continuous with the subarachnoid space. From the subarachnoid space, we can trace CSF fluxes by convective flow into the perivascular space around penetrating arterioles, where solute exchange with the interstitial fluid (ISF) occurs. CMc can be performed for acute injections immediately following the surgery, or for chronic implantation, with later injection in anesthetized or awake, freely moving rodents. Quantitation of tracer distribution in the brain parenchyma can be performed by epifluorescence, 2-photon microscopy, and magnetic resonance imaging (MRI), depending on the physico-chemical properties of the injected molecules. Thus, CMc in conjunction with various imaging techniques offers a powerful tool for assessment of the glymphatic system and CSF dynamics and function. Furthermore, CMc can be utilized as a conduit for fast, brain-wide delivery of signaling molecules and metabolic substrates that could not otherwise cross the blood brain barrier (BBB).

Here we describe a protocol to perform cisterna magna cannulation (CMc), a minimally invasive way to deliver tracers, substrates and signaling molecules into the cerebrospinal fluid (CSF). Combined with different imaging modalities, CMc enables glymphatic system and CSF dynamics assessment, as well as brain-wide delivery of various compounds.

Cisterna magna cannulation (CMc) is a straightforward procedure that enables direct access to the cerebrospinal fluid (CSF) without operative damage to ...

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.

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

Direct Intrathecal Injection of Recombinant Adeno-associated Viruses in Adult Mice

Intrathecal (IT) injection of adeno-associated virus (AAV) has drawn considerable interest in CNS gene therapy by virtue of its safety, noninvasiveness, and excellent transduction efficacy in the CNS.&#160;Previous studies have demonstrated the therapeutic potency of AAV-delivered gene therapy in neurodegenerative disorders by IT administration. However, high rates of unpredictable failure due to the technical limitation of IT administration in small animals have been reported. Here, we established a scoring system to indicate the success extent of lumbar puncture in small animals by adding 1% lidocaine hydrochloride into the injection solution. We further show that the extent of transient weakness following injection can predict the transduction efficiency of AAV. Thus, this IT injection method can be used to optimize therapeutic trials in mouse models of CNS diseases that afflict wide regions of the CNS.

Here we present a direct intrathecal injection technique using 1% lidocaine hydrochloride in a viral solution to ensure efficient adeno-associated virus delivery to small animals and establish a scoring system to predict transduction efficiency in the central nervous system according to the degree of transient weakness induced by lidocaine.

Intrathecal (IT) injection of adeno-associated virus (AAV) has drawn considerable interest in CNS gene therapy by virtue of its safety, noninvasiveness, ...

Direct Injection of a Lentiviral Vector Highlights Multiple Motor Pathways in the Rat Spinal Cord

Introducing proteins of interest into cells in the nervous system is challenging due to innate biological barriers that limit access to most molecules. Injection directly into spinal cord tissue bypasses these barriers, providing access to cell bodies or synapses where molecules can be incorporated. Combining viral vector technology with this method allows for introduction of target genes into nervous tissue for the purpose of gene therapy or tract tracing. Here a virus engineered for highly efficient retrograde transport (HiRet) is introduced at the synapses of propriospinal interneurons (PNs) to encourage specific transport to neurons in the spinal cord and brainstem nuclei. Targeting PNs takes advantage of the numerous connections they receive from motor pathways such as the rubrospinal and reticulospinal tracts, as well as their interconnection with each other throughout spinal cord segments. Representative tracing using the HiRet vector with constitutively active green fluorescent protein (GFP) shows high fidelity details of cell bodies, axons and dendritic arbors in thoracic PNs and in reticulospinal neurons in the pontine reticular formation. HiRet incorporates well into brainstem pathways and PNs but shows age dependent integration into corticospinal tract neurons. In summary, spinal cord injection using viral vectors is a suitable method for introduction of proteins of interest into neurons of targeted tracts.

This protocol demonstrates injection of a retrogradely transportable viral vector into rat spinal cord tissue. The vector is taken up at the synapse and transported to the cell body of target neurons. This model is suitable for retrograde tracing of important spinal pathways or targeting cells for gene therapy applications.

Introducing proteins of interest into cells in the nervous system is challenging due to innate biological barriers that limit access to most molecules. ...

In Vivo Direct Reprogramming of Resident Glial Cells into Interneurons by Intracerebral Injection of Viral Vectors

Converting resident glia in the brain into functional and subtype-specific neurons in vivo provides a step forward towards the development of alternative cell replacement therapies while also creating tools to study cell fate in situ. To date, it has been possible to obtain neurons via in vivo reprogramming, but the precise phenotype of these neurons or how they mature has not been analyzed in detail. In this protocol, we describe a more efficient conversion and cell-specific identification of the in vivo reprogrammed neurons, using an AAV-based viral vector system. We also provide a protocol for functional assessment of the reprogrammed cells&#8217; neuronal maturation. By injecting flip-excision (FLEX) vectors, containing the reprogramming and synapsin-driven reporter genes&#160;to specific cell types in the brain that serve as the target for cell reprogramming. This technique allows for the easy identification of newly reprogrammed neurons. Results show that the obtained reprogrammed neurons functionally mature over time, receive synaptic contacts and show electrophysiological properties of different types of interneurons. Using the transcription factors Ascl1, Lmx1a and Nurr1, the majority of the reprogrammed cells have properties of fast-spiking, parvalbumin-containing interneurons.

This protocol aims at generating directly reprogrammed interneurons in vivo, using an AAV-based viral system in the brain and a FLEX synapsin-driven GFP reporter, which allows for cell identification and further analysis in vivo.

Converting resident glia in the brain into functional and subtype-specific neurons in vivo provides a step forward towards the development of alternative ...

In vitro Modeling for Neurological Diseases using Direct Conversion from Fibroblasts to Neuronal Progenitor Cells and Differentiation into Astrocytes

Research on neurological disorders focuses primarily on the impact of neurons on disease mechanisms. Limited availability of animal models severely impacts the study of cell type specific contributions to disease. Moreover, animal models usually do not reflect variability in mutations and disease courses seen in human patients. Reprogramming methods for generation of induced pluripotent stem cells (iPSCs) have revolutionized patient specific research and created valuable tools for studying disease mechanisms. However, iPSC technology has disadvantages such as time, labor commitment, clonal selectivity and loss of epigenetic markers. Recent modifications of these methods allow more direct generation of cell lineages or specific cell types, bypassing clonal isolation or a pluripotent stem cell state. We have developed a rapid direct conversion method to generate induced Neuronal Progenitor Cells (iNPCs) from skin fibroblasts utilizing retroviral vectors in combination with neuralizing media. The iNPCs can be differentiated into neurons (iNs) oligodendrocytes (iOs) and astrocytes (iAs). iAs production facilitates rapid drug and disease mechanism testing as differentiation from iNPCs only takes 5 days. Moreover, iAs are easy to work with and are generated in pure populations at large numbers. We developed a highly reproducible co-culture assay using mouse GFP+ neurons and patient derived iAs to evaluate potential therapeutic strategies for numerous neurological and neurodegenerative disorders. Importantly, the iA assays are scalable to 384-well format facilitating the evaluation of multiple small molecules in one plate. This approach allows simultaneous therapeutic evaluation of multiple patient cell lines with diverse genetic background. Easy production and storage of iAs and capacity to screen multiple compounds in one assay renders this methodology adaptable for personalized medicine.

We describe a protocol to reprogram human skin-derived fibroblasts into induced Neuronal Progenitor Cells (iNPCs), and their subsequent differentiation into induced Astrocytes (iAs). This method leads to fast and reproducible generation of iNPCs and iAs in large quantities.

Research on neurological disorders focuses primarily on the impact of neurons on disease mechanisms. Limited availability of animal models severely ...

Massive Pontine Hemorrhage by Dual Injection of Autologous Blood

We provide a protocol to establish a massive pontine hemorrhage model in a rat. Rats weighing about 250 grams were used in this study. One hundred microliters of autologous blood was taken from the tail vein and stereotaxically injected into the pons. The injection process was divided into 2 steps: First, 10 &#181;L of blood was injected into a specific location, anteroposterior position (AP) -9.0 mm; lateral (Lat) 0 mm; vertical (Vert) -9.2 mm, followed by a second injection of the residual blood located at AP -9.0 mm; Lat 0 mm; Vert -9.0 mm with a 20-minute interval. The balance beam test, limb placement test, and the modified Voestch neuroscore were used to evaluate neurological function. Magnetic Resonance Imaging (MRI) was used to assess the volume of hemorrhage in vivo. The symptoms of this model were in line with patients with massive pontine hemorrhage.

We present a protocol to establish a massive pontine hemorrhage model in a rat via dual injection of autologous blood.

We provide a protocol to establish a massive pontine hemorrhage model in a rat. Rats weighing about 250 grams were used in this study. One hundred ...

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).

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

Pre-Chiasmatic, Single Injection of Autologous Blood to Induce Experimental Subarachnoid Hemorrhage in a Rat Model

Despite advances in treatment over the last decades, subarachnoid hemorrhage (SAH) continues to carry a high burden of morbidity and mortality, largely afflicting a fairly young population. Several animal models of SAH have been developed to investigate the pathophysiological mechanisms behind SAH and to test pharmacological interventions. The pre-chiasmatic, single injection model in the rat presented in this article is an experimental model of SAH with a predetermined blood volume. Briefly, the animal is anesthetized, intubated, and kept under mechanical ventilation. Temperature is regulated with a heating pad. A catheter is placed in the tail artery, enabling continuous blood pressure measurement as well as blood sampling. The atlantooccipital membrane is incised and a catheter for pressure recording is placed in the cisterna magna to enable intracerebral pressure measurement. This catheter can also be used for intrathecal therapeutic interventions. The rat is placed in a stereotaxic frame, a burr hole is drilled anteriorly to the bregma, and a catheter is inserted through the burr hole and placed just anterior to the optic chiasm. Autologous blood (0.3 mL) is withdrawn from the tail catheter and manually injected. This results in a rise of intracerebral pressure and a decrease of cerebral blood flow. The animal is kept sedated for 30 min and given subcutaneous saline and analgesics. The animal is extubated and returned to its cage. The pre-chiasmatic model has a high reproducibility rate and limited variation between animals due to the pre-determined blood volume. It mimics SAH in humans making it a relevant model for SAH research.

Subarachnoid hemorrhage continues to carry a high burden of mortality and morbidity in man. To facilitate further research into the condition and its pathophysiology, a pre-chiasmatic, single injection model is presented.

Despite advances in treatment over the last decades, subarachnoid hemorrhage (SAH) continues to carry a high burden of morbidity and mortality, largely ...