This study was made possible by the invaluable contributions of Stephanie Tam who provided support in assisting with the surgeries. We extend our sincere gratitude for her help. The National Institutes of Health (AG066162) for support of this research.

AppNL-Fmice (3 months old) that express human amyloid plaque at a physiological level were used for this study though this system can be used with any rodent model. All animal procedures were approved by the Animal Care Committee of CAMH (Protocol #843) and were in accordance with the ethical standards of the Canadian Council on Animal Care guidelines. Mice were bred in-house and kept on a 12-h light-dark cycle with ad libitum access to chow and water.
1. Procedure for Cisterna magna (CM) injection
<ol>
	<li>Surgical procedure
	<ol>
		<li>Place the mouse (APPNL-F, both sexes, 3 months) in an anesthesia induction chamber for 1 min. 
		NOTE: Flush the chamber with 5% isoflurane mixed with 1% oxygen for at least 1 min.</li>
		<li>After the mouse is anesthetized, remove the animal from the chamber and place it on a fresh surgical drape and maintain anesthesia (isoflurane, 2.5%-3%) by placing a nasal cone over the nose. 
		NOTE: Check depth of anesthesia by pinching the toe to ensure no reflexes.</li>
		<li>Apply ophthalmic ointment to prevent dryness. Then, subcutaneously inject 0.1 mL of bupivacaine (local anesthetic; 1-2 mg/kg 0.125%, with a dilution ratio of 1:2) to the incision site and Metacam (analgesic; 5 mg/kg, with a dilution ratio of 1:10). Keep the animal hydrated by injecting 0.5 mL of saline subcutaneously before doing the procedure to compensate for any potential blood loss during surgery.</li>
		<li>For a clear incision site, shave the neck at the base of the occipital bone, wipe down the surface with sterile PBS and transfer the subject to the stereotaxic frame in the prone position. 
		NOTE: Ensure that anesthesia is maintained through the stereotaxic nose cone.</li>
		<li>Place the nose bar from the stereotaxic frame on top of the mouse to provide stability and ensure the upper teeth are fastened.</li>
		<li>Position the animal head at a 120&#176; angle from the body to elevate and distend the nape to expose the surgical area of interest.</li>
		<li>Clean the surgical site with Betadine scrub 3x, 70% ethanol 3x, and betadine solution 1x with sterile 2 inch x 2 inch gauze. 
		NOTE: Gloves and instruments should be sterile before the procedure. A sterile drape should also be placed over the incision site.</li>
		<li>Make a small incision (1 cm) using a scalpel. 
		NOTE: This should reveal the midline of the nape of the neck.</li>
		<li>Carefully run the scalpel along the midline to cut through the muscles. Then, using forceps gently separate the muscles by pulling them apart to the left and right. 
		NOTE: This should reveal the Cisterna magna (inverted triangle) which is located beneath the base of the skull (Figure 1).</li>
	</ol>
	</li>
	<li>Preparation of elastase 
	NOTE: The package of elastase (powder form) that was ordered contained 250 Units (U). The amount of the elastase to be injected in each animal is 2.5 &#181;L. The amount of elastase that is supposed to be in the 2.5 &#181;L should be 15 milliUnits mU.
	<ol>
		<li>First calculate the concentration of the solution to be injected. 
		Concentration = mass/ volume 
		=&#160;15 mU/2.5 &#181;L 
		=&#160;6 mU/&#181;L 
		Therefore, the concentration that is required is 6 mU/&#181;L.</li>
		<li>The quantity of the elastase is 250 U and the study works with milliUnits, so convert the units to milliUnits which gives 250000 mU.</li>
		<li>After having a mass of 250000 mU and concentration of 6 mU/uL, figure out the volume (PBS) that would require diluting the elastase and maintaining a concentration of 6 mU/uL.</li>
		<li>Using the same formula above, rearrange to solve for volume: 
		Volume = mass/Conc. 
		= 250000&#160;mU/ 6 mU/&#181;L 
		&#8203;=&#160;41666.67 &#181;L</li>
		<li>Since it is hard to pipette 41666.67 &#181;L, simply convert it to milliliters by dividing it by 1000 to get 41.66667 mL to obtain a 1x solution.
		<ol>
			<li>To make the solution more concentrated,increase it by 10x or 100x.</li>
			<li>Create a 10x solution since it is simpler to work with (see below). 
			&#8203;10x solution ---------- 4.166667 mL (conc.)</li>
		</ol>
		</li>
		<li>Place the 250 U in a 5 mL aliquot tube and add 4.167 mL of sterile PBS into the aliquot tube to make the 10x stock solution. 
		NOTE: Prepare the elastase inside a sterile biosafety cabinet to maintain sterility.</li>
		<li>Pipette 10 &#181;L of stock solution and place in 100 aliquot tubes giving 1000 &#181;L of the 4.167 mL stock solution. Store the tubes at -20 &#176;C.</li>
		<li>Store the remainder of stock solution in aliquot tubes at 100 &#181;L per tube and store in freezer at -20 &#176;C. 
		NOTE: The solutions are stable at -20 &#176;C for up to 6 months with no significant loss of activity. In addition, it is important to note that excessive freezing and thawing of the elastase can reduce stability. Therefore, to prevent that from happening, the stock solution can be divided before freezing such that only one sample needs to be thawed at a time for injection.</li>
		<li>After thawing one of the aliquot tubes that contains 10 &#181;L of a 10x solution, add 90 &#181;L of sterile PBS to get of a 100 &#181;L 1x solution.</li>
		<li>If the aliquot tube that contains 100 &#181;L of a 10x solution was thawed instead, simply add 900 &#181;L of PBS to get 1000 &#181;L, which is a 1x solution. Store either solution at 4 &#176;C for 1 week.</li>
	</ol>
	</li>
	<li>Injecting into the Cisterna magna 
	NOTE: Clean the Hamilton syringe before loading with elastase by loading and aspirating with distilled water 5x, then ethanol 5x, and finally distilled water 5x.
	<ol>
		<li>Load the Hamilton syringe with 2.5 &#181;L of elastase (concentration of 15 mU) and ensure there are no air bubbles.</li>
		<li>Slowly insert the syringe with the bevel facing upward into the center of the Cisterna magna. 
		NOTE: Avoid puncturing any blood vessels that run across the Cisterna magna.</li>
		<li>After the puncture, remove the bevel. This will facilitate the release of a small amount of cerebrospinal fluid (CSF). Use a sterile cotton swab to absorb the excess CSF.</li>
		<li>Reinsert the bevel into the puncture site, slowly inject elastase or vehicle over 1 min into the Cisterna magna and leave the needle in place for 1 min to prevent any leakage of elastase.</li>
		<li>After removing the bevel, close the incision site with a non-absorbable surgical suture (#4-0 with 3/8&#34; needle), and turn off the anesthetic.</li>
		<li>Remove the animal from the stereotaxic frame and place on a warm heat pad set at 37 &#176;C until the subject recovers. 
		NOTE: Another dose of Metacam is recommended after surgery to help with pain during post-surgical recovery.</li>
	</ol>
	</li>
</ol>
2. Silicone rubber compound injection and tissue harvest
<ol>
	<li>Anesthetize the animal with an overdose of avertin (1 mL, 125-250 mg/kg). 
	NOTE: Pinch the toe to ensure reflexes cease.</li>
	<li>Place the animal on a diaper and secure all limbs with surgical tape with the chest cavity facing upward</li>
	<li>Open the chest cavity using forceps and scissors to expose the heart.</li>
	<li>Insert a 23 G blunt needle into the left ventricle of the heart and perfuse with PBS infused with heparin (100 U/mL) for 4 min to clear the blood vessels using a micro-perfusion pump.</li>
	<li>Replace the buffer bottle with one containing 4% paraformaldehyde (PFA)(made up in PBS) and perfuse for another 4 min.</li>
	<li>Prepare the silicone rubber compound (yellow) solution by combining 5 mL of compound with 5 mL of the diluent in a 50 mL tube. Mix thoroughly before use.</li>
	<li>Fill a 10 mL syringe with 10 mL of silicone rubber compo und. Attach the syringe to the hose connected to a 23 G blunt needle, and carefully inject the solution manually into the heart. 
	NOTE: Due to the high viscosity of the silicone rubber compound, it is important to use a larger barrel syringe or the pressure will be too great to push the plunger.</li>
	<li>Remove the mouse&#39;s head after injection and store at 4 &#176;C overnight allowing the silicone rubber compound to cure in the vessels.</li>
	<li>Next day, gently remove the skull of the mouse (preferably using fine forceps), extract the brain, and incubate in 4% PFA overnight at room temperature (RT).</li>
	<li>After a 24 h incubation in PFA, wash the brain in PBS 3x, place in 30% sucrose, and store at 4 &#176;C.</li>
</ol>
3. Quantitative analysis
<ol>
	<li>To capture high-quality images of the Circle of Willis in the entire brain (Figure 3):
	<ol>
		<li>Remove the brain from the sucrose solution. Thoroughly dry the brain using a paper towel to wick the surface moisture.</li>
		<li>Position the brain in a brain slicer. Place the brain slicer under a light microscope equipped with a camera.</li>
		<li>Focus the microscope to a clear, high-resolution resolution of the Circle of Willis.</li>
	</ol>
	</li>
	<li>Use image analysis software with a 10 cm ruler reference to measure approximately 1 cm and note the pixel count. Set the 1 cm scale based on this measurement, then measure the basilar artery&#39;s thickness horizontally to find its diameter.</li>
	<li>Use a standard pixel densitometry software to analyze the image. Measure (in centimeters) five separate readings of the basilar artery using an image analysis software and calculate the average to get an accurate representation of the basilar artery diameter.</li>
	<li>Use a 2-way analysis of variance (ANOVA) to compare the average change in diameter between the elastase and control group.
	<ol>
		<li>Calculate the percentage change in diameter using the formula below 21: 
		<img alt="Equation 1" src="/files/ftp_upload/66792/66792eq01.jpg" style="margin: auto;" /> 
		C = relative change 
		X1 = Initial value (Vehicle) 
		X2 = Final value (Elastase) 
		Consider P values less than 0.05 statistically significant. Perform all statistical analyses using a statistical analysis software.</li>
	</ol>
	</li>
</ol>
4. Removal of silicone rubber compound using alcohol dehydration
NOTE: It is important to dehydrate the brain to help remove the excess silicone rubber compound from the blood vessel which can potentially improve the quality of immunostaining.
<ol>
	<li>Wash the brains 3x with PBS to remove the sucrose.</li>
	<li>Place the brain in 25%, 50%, 75%, 95%, and absolute ethanol for 24 h at each concentration, for a total time of 5 days.</li>
	<li>After dehydration of tissue, place the tissue in methyl salicylate (conc. &#8804;100%) for 12-24 h. The ethanol and methyl salicylate will allow for the removal of excess silicone in the tissue.</li>
	<li>Wash the brain in PBS 3x to remove residual chemicals and leave overnight on a rocker at 4&#160;&#176;C in fresh PBS.</li>
	<li>Place the brain in sucrose and store at 4&#160;&#176;C prior to sectioning and immunostaining.</li>
</ol>
5. Immunohistochemical staining
<ol>
	<li>Sectioning of brains
	<ol>
		<li>Prior to immunosectioning, allow the brain to settle at the bottom of the tube filled with sucrose.</li>
		<li>Freeze the brain on the microtome platform using optimal cutting temperature (OCT) compound surrounded by dry ice. Slice the brain into 40 &#181;m sections using the microtome blade.</li>
		<li>Transfer each section into a 96-well plate containing cryoprotectant to prevent freezing using a fine tip paintbrush.</li>
		<li>Securely seal the 96-well plate with plastic wrap. Store the sealed plate in a -20 &#176;C freezer.</li>
	</ol>
	</li>
	<li>Immunofluorescence staining (using NeuN as an example)
	<ol>
		<li>Place the sections in a 24-well plate. Perform three washes with PBS to eliminate the cryoprotectant.</li>
		<li>To prevent non-specific binding, incubate for 1 h at RT using a blocking solution containing 2% goat serum, 0.1% Triton-X 100, and 1% BSA.</li>
		<li>Prepare all components in PBS. 
		NOTE: The primary antibody used included monoclonal mouse anti-NeuN with a 1: 500 dilution. The secondary antibody used included polyclonal goat anti-mouse 568 with a 1:200 dilution.</li>
		<li>Dilute the monoclonal mouse anti-NeuN in the blocking solution mentioned above.</li>
		<li>Add the diluted antibody to each well. Incubate overnight at 4 &#176;C, shielded from light by foil.</li>
		<li>After the primary incubation, wash the sections 3x with PBS. Dilute the polyclonal goat anti-mouse 568 in the same blocking solution.</li>
		<li>Add the diluted antibody to each well. Incubate for 2 h at room temperature, shielded from light by foil.</li>
		<li>Wash the sections 3x with PBS after the secondary incubation.</li>
		<li>Prepare for mounting by slowly transferring sections onto slides using fine paintbrushes. Allow all sections to dry.</li>
		<li>Apply 125-150 &#181;L of anti-fade mounting media once sections are dried. Gently place coverslips.</li>
		<li>Allow slides to dry for 24 h at RT in the dark, sealing edges with nail polish. Capture fluorescent images using imaging software.</li>
	</ol>
	</li>
</ol>

Dolichoectasia: Vascular Dementia and Alzheimer&#39;s Disease Models

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There is no conflict of interest.

This article demonstrates an improved protocol for cerebrovascular dilatation, providing a precise and straightforward approach for elastase injection into the Cisterna magna of mice. This anatomical point serves as a direct gateway to the cerebrospinal fluid, offering a valuable avenue for the investigation of different neurological diseases. One of the main advantages of this modified technique is that injecting a single dose of elastase into the Cisterna magna of mice was able to cause and sustain dilatation of large ...

Microvascular endothelial cells that line the cerebral capillaries are the main components for forming the blood-brain barrier (BBB)1 which plays a crucial role in regulating what enters or leaves the brain circulation with the periphery. Essential nutrients needed for nervous tissue are permitted to enter the BBB, while some essential amino acids like glutamate are expelled from the brain, as high concentrations can cause permanent neuroexcitatory damage to brain tissue2. Under normal physiological conditions, the BBB limits the amount of plasma proteins like albumin3,4 and prothrombin from entering the brain since those can have detrimental effects5,6,7. Finally, the BBB protects the brain from neurotoxins that are circulating in the periphery, such as xenobiotics from food or the environment1. Overall, damage to brain tissues is irreversible, and aging that correlates with low levels of neurogenesis8 highlights the importance of the BBB in protecting and preventing any factors from accelerating the neurodegenerative process.
In dolichoectasia (or large blood vessel dilatation), a decrease in vessel elasticity is observed, which results in vessels undergoing morphological changes, thus rendering them dysfunctional9 and leading to reduced blood flow in the brain. This reduction in blood flow subsequently diminishes oxygen and glucose supply, ultimately triggering damage to the BBB through the activation of reactive astrocytes10. When the internal elastin lamina of vessels is damaged from dolichoectasia11, repeated stimulation of the vascular endothelial growth factor (VEGF) is necessary for angiogenesis. This can lead to the formation of leaky vessels and ultimately result in pathological angiogenesis, characterized by the development of defective vessels12. During pathological angiogenesis, when blood vessels become defective, a compensatory mechanism appears to restore vessel integrity by upregulating tight junction proteins. However, this process can inadvertently disrupt the BBB when the structural integrity of a blood vessel is lost13. This may occur through further disrupting the BBB and promoting the production of amyloid plaque14. Additionally, leakage from the periphery can cause neuroinflammation15, which results in neuronal degeneration and subsequent memory loss.
Structurally the protection that the BBB provides is through the tight junctions that prevent xenobiotic agents from the blood entering the brain. When permitting certain substances to enter the brain, the BBB mainly does it through two major processes, passive diffusion, or specific channels (like ion channels and transporters)1. In AD, research has demonstrated that a dysfunctional vascular system plays a significant role in the progression of the condition12,13. The formation of amyloid-beta (A&#946;) plaques and neurodegeneration can result from the breakdown of the BBB12,13 and disturbances in cerebral blood flow16. A reduction in cerebral blood flow can be seen in elderly individuals diagnosed with vascular dementia and AD17,18. Damage to the blood-brain barrier (BBB) along with a dysfunctional cerebral blood flow (CBF) can contribute to the increased production of A&#946; concentration in the brain, accompanied by the infiltration of foreign materials from the peripheral circulation19.
To investigate the pathogenesis of neurological diseases like AD and vascular dementia (VaD), models are developed to replicate the disease. In vitro models are extensively used but lack the biological environment for extensive disease modeling like mixed cell population, thus necessitating the importance of in vivo models. Mice are commonly used due to their ease of genetic manipulation in generating human-like properties (e.g., pathology) in disease. With the progression that has been made so far, there is still a continued need for improved models to emulate disease phenotypes like large vessel dilatation and their role in AD. To this end, we saw an opportunity and modified a technique that involved the injection of elastase into the Cisterna magna of mice20,21. Elastase is an enzyme that has been shown to break down elastin in connective tissue22 and in surrounding tight junctions23. The Cisterna magna was chosen as the point of injection due to it being located directly above the circle of Willis, the largest blood vessel in the brain. By injecting elastase into the Cisterna magna, we can compromise the BBB and blood vessels by breaking down the tight junctions and inducing dilatation of blood vessels (circle of Willis)24,25. Combining this technique with the use of an AD mouse model of pathology, for improved understanding of the pathogenesis for the vascular component of AD, can provide valuable insights into the complex interactions and influences between these two distinct pathologies.
Previous studies have demonstrated instances where patients display both the pathological features of AD and VaD, a condition typically referred to as mixed dementia26,27. Thus, understanding the interconnected mechanisms between both conditions can offer a more comprehensive perspective on the progression and manifestation of these neurodegenerative disorders, enhances our comprehension of the underlying mechanisms and potential therapeutic strategies. To this end, we demonstrate the application of elastase in an AD pathology mouse model (AppNL-F) to identify vascular changes.

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			<td>23 G catheter</td>
			<td>University Medstore</td>
			<td>2546-CABD305145</td>
			<td>Needed for perfusion&#160; (https://www.uoftmedstore.com/index.sz)</td>
		</tr>
		<tr>
			<td>Absolute ethanol</td>
			<td>University Medstore</td>
			<td>https://www.uoftmedstore.com/index.sz</td>
			<td>For removing the microfil</td>
		</tr>
		<tr>
			<td>Betadine scrub</td>
			<td>#</td>
			<td>https://www.pittsborofeed.com/products/betadine-surgical-scrub</td>
			<td>Sterilization</td>
		</tr>
		<tr>
			<td>Betadine solution</td>
			<td>Amazon</td>
			<td>https://www.amazon.ca/Povidone-Iodine-10-Topical-Solution-100ml/dp/B09DTKJGHW</td>
			<td>Sterilization</td>
		</tr>
		<tr>
			<td>Bupivacaine</td>
			<td>Provided by animal facility</td>
			<td>N/A</td>
			<td>Analgesic</td>
		</tr>
		<tr>
			<td>Clippers</td>
			<td>BrainTree Scientific Inc</td>
			<td>CLP-41590</td>
			<td>Shave fur</td>
		</tr>
		<tr>
			<td>Cotton Q-tip</td>
			<td>University Medstore</td>
			<td>1962</td>
			<td>For surgery (https://www.uoftmedstore.com/index.sz)</td>
		</tr>
		<tr>
			<td>Elastase</td>
			<td>Sigma-aldrich</td>
			<td>&#160;E7885</td>
			<td>Used for the dilatation of blood vessel</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>University Medstore</td>
			<td>39752-P016-EAAN</td>
			<td>Sterilization (https://www.uoftmedstore.com/index.sz)</td>
		</tr>
		<tr>
			<td>Goat anti-mouse 568</td>
			<td>Invitrogen</td>
			<td>A11004</td>
			<td>For staining mature neurons</td>
		</tr>
		<tr>
			<td>Graphpad prism 10</td>
			<td>Graphpad prism 10</td>
			<td>https://www.graphpad.com/</td>
			<td>Statistical analysis software</td>
		</tr>
		<tr>
			<td>Hamilton syringe</td>
			<td>Sigma-aldrich</td>
			<td>28614-U</td>
			<td>Injection elastase</td>
		</tr>
		<tr>
			<td>Heat pad</td>
			<td>Amazon</td>
			<td>https://www.amazon.ca/iPower-Temperature-Controller-Terrarium-Amphibians/dp/B08L4DBFFZ</td>
			<td>Maintain body temperature</td>
		</tr>
		<tr>
			<td>ImageJ software</td>
			<td>Fiji Imagej software</td>
			<td>imagej.net (USA)</td>
			<td>Image analysis software</td>
		</tr>
		<tr>
			<td>Induction chamber</td>
			<td>Provided by animal facility</td>
			<td>N/A</td>
			<td>Anesthesia induction</td>
		</tr>
		<tr>
			<td>Metacam</td>
			<td>Provided by animal facility</td>
			<td>N/A</td>
			<td>Analgesic</td>
		</tr>
		<tr>
			<td>Methyl salicylate</td>
			<td>Sigma-aldrich</td>
			<td>M6752</td>
			<td>For removing the microfil</td>
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		<tr>
			<td>Microfil</td>
			<td>Flow Tech, Carver, Massachusetts</td>
			<td>https://www.flowtech-inc.com/order/</td>
			<td>&#160;Dye (yellow)</td>
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		<tr>
			<td>Mouse monoclonal anti-NeuN</td>
			<td>Millipore Sigma</td>
			<td>MAB377</td>
			<td>For staining mature neurons</td>
		</tr>
		<tr>
			<td>Olympus VS200 slide scanner and VSI software.</td>
			<td>Olympus Life Science</td>
			<td>https://www.olympus-lifescience.com/en/downloads/detail-iframe/?0[downloads][id]=847254104</td>
			<td>Imaging software</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>University Medstore</td>
			<td>PAR070.1</td>
			<td>For protein fixation&#160; (https://www.uoftmedstore.com/index.sz)</td>
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			<td>Perfusion pump</td>
			<td>VWR International</td>
			<td>https://pr.vwr.com/store/product/4787969/vwr-variable-speed-peristaltic-pumps</td>
			<td>Needed for perfusion</td>
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		<tr>
			<td>Scalpel</td>
			<td>University Medstore</td>
			<td>2580-M90-10</td>
			<td>For surgery (https://www.uoftmedstore.com/index.sz)</td>
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		<tr>
			<td>Stereotaxic</td>
			<td>Provided by animal facility</td>
			<td>N/A</td>
			<td>So secure the animal for surgery</td>
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		<tr>
			<td>Surgical scissor</td>
			<td>University Medstore</td>
			<td>22751-A9-240</td>
			<td>For surgery (https://www.uoftmedstore.com/index.sz)</td>
		</tr>
		<tr>
			<td>Surgical tape</td>
			<td>University Medstore</td>
			<td>https://www.amazon.ca/3M-Micropore-Tape-1530-2-Rolls/dp/B0082A9GS2</td>
			<td>Secure the animal on the diaper</td>
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		<tr>
			<td>Sutures</td>
			<td>University Medstore</td>
			<td>2297-VS881</td>
			<td>For surgery (https://www.uoftmedstore.com/index.sz)</td>
		</tr>
		<tr>
			<td>X2 tweezers</td>
			<td>University Medstore</td>
			<td>7731-A10-612</td>
			<td>For surgery (https://www.uoftmedstore.com/index.sz)</td>
		</tr>
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a visual approach for inducing dolichoectasia in mice to model large vessel-mediated cerebrovascular dysfunction

The blood-brain (BBB) is a crucial system that regulates selective brain circulation with the periphery, as an example, allowing necessary nutrients to enter and expel excessive amino acids or toxins from the brain. To model how the BBB can be compromised in diseases like vascular dementia (VaD) or Alzheimer's disease (AD), researchers developed novel methods to model vessel dilatation. A compromised BBB in these disease states can be detrimental and result in the dysregulation of the BBB leading to untoward and pathological consequences impacting brain function. We were able to modify an existing technique that enabled us to inject directly into the Cisterna magna (CM) to induce dilatation of blood vessels using elastase, and disrupt the tight junctions (TJ) of the BBB. With this method, we were able to see various metrics of success over previous techniques, including consistent blood vessel dilatation, reduced mortality or improved recovery, and improving the fill/opacifying agent, a silicone rubber compound, delivery for labeling blood vessels for dilatation analysis. This modified minimally invasive method has had promising results, with a 19%-32% increase in sustained dilatation of large blood vessels in mice from 2 weeks to 3 months post-injection. This improvement contrasts with previous studies, which showed increased dilatation only at the 2 week mark. Additional data suggests sustained expansion even after 9.5 months. This increase was confirmed by comparing the diameter of blood vessels of the elastase and the vehicle-injected group. Overall, this technique is valuable for studying pathological disorders that affect the central nervous system (CNS) using animal models.

We were successful in locating the Cisterna magna beneath the occipital region of the skull after carefully positioning the mouse on the stereotaxic frame and dissecting the muscles. This anatomical structure, resembling an inverted triangle and highlighted in yellow, is situated beneath the base of the skull (Figure 1). To ensure precision and prevent any damage to the brain tissue, 1-2 mm of the Hamilton syringe bevel was gently inserted into the Cisterna magna.
<p class="jove_content"...

Read this Scientific Article Protocol about A Visual Approach for Inducing Dolichoectasia in Mice to Model Large Vessel-Mediated Cerebrovascular Dysfunction at JoVE.com

Author Spotlight: Modeling Vascular Contributions to Alzheimer's Disease in Transgenic Mice

We demonstrate chemically inducing large blood vessel dilatation in mice as a model for investigating cerebrovascular dysfunction, which can be used for vascular dementia and Alzheimer's disease modeling. We also demonstrate visualizing the vasculature by injecting silicone rubber compound and providing clear visual guidance for measuring changes in blood vessel size.

A Visual Approach for Inducing Dolichoectasia in Mice to Model Large Vessel-Mediated Cerebrovascular Dysfunction

The blood-brain (BBB) is a crucial system that regulates selective brain circulation with the periphery, as an example, allowing necessary nutrients to ...

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Research

JoVE Journal

Neuroscience

501 Views.  Centre for Addiction and Mental Health. We demonstrate chemically inducing large blood vessel dilatation in mice as a model for investigating cerebrovascular dysfunction, which can be used for vascular dementia and Alzheimer's disease modeling. We also demonstrate visualizing the vasculature by injecting silicone rubber compound and providing clear visual guidance for measuring changes in blood vessel size.

Video: Author Spotlight: Modeling Vascular Contributions to Alzheimer's Disease in Transgenic Mice

Double Fluorescence in situ Hybridization in Fresh Brain Sections

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh frozen brain sections. Our group has successfully used this approach to study gene co-regulation. More specifically, we have used this dFISH method to explore the anatomical organization, neurochemical properties, and the impact of sensory experience in central sensory circuits, at single cell resolution. This protocol has been validated in brain tissue from mice, rats and songbirds but is expected to be easily adaptable to other vertebrate species, as well as to an array of non-neural tissues. In this film we provide a detailed demonstration of the main steps of this procedure.

Double Fluorescence in situ Hybridization in Fresh Brain Sections

This protocol involves a non-radioactive in-situ hybridization procedure that enables the simultaneous identification of two transcript species, at a single cell resolution, in thin sections of the vertebrate brain.

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh ...

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% ...

Inducing Dendritic Growth in Cultured Sympathetic Neurons

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs 4-6. Altered patterns of dendritic growth and plasticity are associated with impaired neurobehavioral function in experimental models 7, and are thought to contribute to clinical symptoms observed in both neurodevelopmental disorders 8-10 and neurodegenerative diseases 11-13. Such observations underscore the functional importance of precisely regulating dendritic morphology, and suggest that identifying mechanisms that control dendritic growth will not only advance understanding of how neuronal connectivity is regulated during normal development, but may also provide insight on novel therapeutic strategies for diverse neurological diseases.
Mechanistic studies of dendritic growth would be greatly facilitated by the availability of a model system that allows neurons to be experimentally switched from a state in which they do not extend dendrites to one in which they elaborate a dendritic arbor comparable to that of their in vivo counterparts. Primary cultures of sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rodents provide such a model. When cultured in defined medium in the absence of serum and ganglionic glial cells, sympathetic neurons extend a single process which is axonal, and this unipolar state persists for weeks to months in culture 14,15. However, the addition of either bone morphogenetic protein-7 (BMP-7) 16,17 or Matrigel 18 to the culture medium triggers these neurons to extend multiple processes that meet the morphologic, biochemical and functional criteria for dendrites. Sympathetic neurons dissociated from the SCG of perinatal rodents and grown under defined conditions are a homogenous population of neurons 19 that respond uniformly to the dendrite-promoting activity of Matrigel, BMP-7 and other BMPs of the decapentaplegic (dpp) and 60A subfamilies 17,18,20,21. Importantly, Matrigel- and BMP-induced dendrite formation occurs in the absence of changes in cell survival or axonal growth 17,18.
Here, we describe how to set up dissociated cultures of sympathetic neurons derived from the SCG of perinatal rats so that they are responsive to the selective dendrite-promoting activity of Matrigel or BMPs.

We describe a protocol for using bone morphogenetic protein-7 (BMP-7) or Matrigel to selectively induce dendritic growth in primary sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rats.

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

The Use of Trace Eyeblink Classical Conditioning to Assess Hippocampal Dysfunction in a Rat Model of Fetal Alcohol Spectrum Disorders

Neonatal rats were administered a relatively high concentration of ethyl alcohol (11.9% v/v) during postnatal days 4-9, a time when the fetal brain undergoes rapid organizational change and is similar to accelerated brain changes that occur during the third trimester in humans. This model of fetal alcohol spectrum disorders (FASDs) produces severe brain damage, mimicking the amount and pattern of binge-drinking that occurs in some pregnant alcoholic mothers. We describe the use of trace eyeblink classical conditioning (ECC), a higher-order variant of associative learning, to assess long-term hippocampal dysfunction that is typically seen in alcohol-exposed adult offspring. At 90 days of age, rodents were surgically prepared with recording and stimulating electrodes, which measured electromyographic (EMG) blink activity from the left eyelid muscle and delivered mild shock posterior to the left eye, respectively. After a 5 day recovery period, they underwent 6 sessions of trace ECC to determine associative learning differences between alcohol-exposed and control rats. Trace ECC is one of many possible ECC procedures that can be easily modified using the same equipment and software, so that different neural systems can be assessed. ECC procedures in general, can be used as diagnostic tools for detecting neural pathology in different brain systems and different conditions that insult the brain.

Trace eyeblink classical conditioning (ECC) was used to assess hippocampal-dependent associative learning in adult rats that were administered a high concentration (11.9% v/v) of alcohol during early neonatal brain development. In general, ECC procedures are sound diagnostic tools for detecting brain dysfunction across many psychological and biomedical settings.

Neonatal rats were administered a relatively high concentration of ethyl alcohol (11.9% v/v) during postnatal days 4-9, a time when the fetal brain ...

Partial Optic Nerve Transection in Rats: A Model Established with a New Operative Approach to Assess Secondary Degeneration of Retinal Ganglion Cells

Previous studies have shown that the secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. Partial optic nerve transection is considered a useful and reproducible model. Compared with other optic nerve injury models used commonly for assessing secondary degeneration, e.g. complete optic nerve transection and optic nerve crush models, the partial optic nerve transection model is superior as it distinguishes primary from secondary degeneration in situ. Therefore, it serves as an excellent tool for evaluating secondary degeneration. This study describes a novel operative approach of partial optic nerve transection by directly accessing the area of the retrobulbar optic nerve through the orbital lateral wall of the eyeball. Moreover, we present a newly designed, low cost surgical instrument to assist with transection. As demonstrated by the representative results in distinguishing the boundary of primary and secondary injury areas, the new approach and instrument ensures high efficiency and stability of the model by providing adequate space for surgical operation. This in turn makes it easy to separate the meningeal sheath and ophthalmic vessels from the optic nerve before transection. An additional benefit is that this space-saving operative approach improves the investigators&#39; ability to administer drugs, carriers, or selective RGC tracers to the stump of the partially transected optic nerve, allowing the exploration of mechanisms behind secondary injury in RGCs, in a new way.

Secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. This study describes an innovative operative approach for partial optic nerve transection. The use of this space-saving operative approach extends the model's application range, and allows exploration of secondary injury mechanisms in RGCs in a new way.

Previous studies have shown that the secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. Partial optic nerve transection ...

A Simple Approach to Induce Experimental Autoimmune Neuritis in C57BL/6 Mice for Functional and Neuropathological Assessments

Experimental autoimmune neuritis (EAN) is a well-appreciated experimental model of autoimmune peripheral demyelinating diseases. EAN disease is induced by immunizing mice with neurogenic peptides to direct an inflammatory attack toward components of the peripheral nervous system (PNS). Recent advances have enabled the induction of EAN in the relatively resistant C57BL/6 mouse line using myelin protein zero (P0)106-125 or P0180-199 peptides delivered in adjuvant combined with the injection of pertussis toxin. The ability to induce EAN in the C57BL/6 strain allows for the use of the numerous genetic tools that exist on this mouse background, and thus allows the sophisticated study of disease pathogenesis and interrogation of the mechanistic action of novel therapeutics in combination with transgenic approaches. In this study, we demonstrate a simple approach to successfully induce EAN using the P0180-199 peptide in C57BL/6 mice. We also outline a protocol for the assessment of functional deficits that occur in this model, accompanied by an array of neuropathological features. Thus, this model is a powerful experimental model to study the pathogenesis of human peripheral demyelinating neuropathies, and to determine the efficacy of potential therapies that aim to promote myelin repair and protect against nerve damage in autoimmune neuritis.

This report outlines a simple approach to successfully induce experimental autoimmune neuritis (EAN) using the myelin protein zero (P0)180-199 peptide in combination with Freund&#39;s complete adjuvant and pertussis toxin. We present a sophisticated paradigm capable of accurately assessing the extent of functional deficits and neuropathology that occur in this EAN.

Experimental autoimmune neuritis (EAN) is a well-appreciated experimental model of autoimmune peripheral demyelinating diseases. EAN disease is induced by ...

Single Synapse Indicators of Glutamate Release and Uptake in Acute Brain Slices from Normal and Huntington Mice

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other neurodegenerative disorders, this independence might be compromised due to insufficient glutamate clearance and the resulting spill-in and spill-out effects. Altered astrocytic coverage of the presynaptic terminals and/or dendritic spines as well as a reduced size of glutamate transporter clusters at glutamate release sites have been implicated in the pathogenesis of diseases resulting in symptoms of dys-/hyperkinesia. However, the mechanisms leading to the dysfunction of glutamatergic synapses in HD are not well understood. Improving and applying synapse imaging we have obtained data shedding new light on the mechanisms impeding the initiation of movements. Here, we describe the principle elements of a relatively inexpensive approach to achieve single synapse resolution by using the new genetically encoded ultrafast glutamate sensor iGluu, wide-field optics, a scientific CMOS (sCMOS) camera, a 473 nm laser and a laser positioning system to evaluate the state of corticostriatal synapses in acute slices from age appropriate healthy or diseased mice. Glutamate transients were constructed from single or multiple pixels to obtain estimates of i) glutamate release based on the maximal elevation of the glutamate concentration [Glu] next to the active zone and ii) glutamate uptake as reflected in the time constant of decay (TauD) of the perisynaptic [Glu]. Differences in the resting bouton size and contrasting patterns of short-term plasticity served as criteria for the identification of corticostriatal terminals as belonging to the intratelencephalic (IT) or the pyramidal tract (PT) pathway. Using these methods, we discovered that in symptomatic HD mice ~40% of PT-type corticostriatal synapses exhibited insufficient glutamate clearance, suggesting that these synapses might be at risk to excitotoxic damage. The results underline the usefulness of TauD as a biomarker of dysfunctional synapses in Huntington mice with a hypokinetic phenotype.

We present a protocol to evaluate the balance between glutamate release and clearance at single corticostriatal glutamatergic synapses in acute slices from adult mice. This protocol uses the fluorescent sensor iGluu for glutamate detection, a sCMOS camera for signal acquisition and a device for focal laser illumination.

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other ...

How to Obtain Reliable Visual Event-related Potentials in Newborns

The present study discusses the characteristics of visual event-related potentials (VEPs) and outlines methodological steps for obtaining reliable measurements in newborns. Obtaining high-quality, reliable VEPs is crucial for the early detection of abnormal development of the central nervous system in at-risk newborns, and for implementing successful early interventions. Recommendations are based on a previous study which showed that when post-conceptional age, polysomnography-identified sleep stages, and light-emitting diodes (LEDs) googles as the luminous source are controlled, no more than 4 repetitions of VEP averages are required to obtain replicable recordings, variability decreases, and reliable VEPs can be obtained. By controlling for these sources of variability and using statistical analyses, we were able to clearly and reliably identify the amplitude and latency of three main components (NII, PII and NIII) present in 100% of newborns (n = 20) during active sleep. Recording VEPs during awake states, quiet sleep and transitional sleep is not recommended because VEP morphology may differ significantly from one average to the next, leading to the risk of misleading clinical prognoses. Moreover, it is easier to obtain VEPs during active sleep because this state can be clearly and reliably identified at this stage of development, sleep cycles are short enough to allow measurements to be taken in a reasonable time, and the method does not require new o expensive equipment.

Several important points for obtaining high-quality reliable visual evoked potentials (VEPs) in newborns while minimizing variability and the risk of misleading prognoses are presented.