This work was supported by the Knut and Alice Wallenberg Foundation, Hj&#228;rnfonden, Wenner Gren Foundations, and the Crafoord foundation.

All procedures were carried out in accordance with the European directive 2010/63/EU and were approved by Malm&#246;-Lund Ethical committee on Animal Research (Dnr 5.8.18-05527/2019) and conducted according to the CODEX guidelines of the Swedish Research Council.
1. Preparation
<ol>
	<li>Tracer
	<ol>
		<li>Prepare artificial CSF (126 mM NaCl, 2.5 mM KCl, 1.25 mM NaH2PO4, 2 mM MgCl2, 2 mM CaCl2, 10 mM glucose, 26 mM NaHCO3; pH 7.4)</li>
		<li>To 500 &#956;L of artificial CSF, add 10 mg of albumin from bovine serum (BSA) conjugated with Alexa Fluor 647 (BSA-647).</li>
		<li>Centrifuge at 5,000 x g for 5 min and use the supernatant.</li>
	</ol>
	</li>
	<li>Cannula
	<ol>
		<li>Attach a 1 mL syringe to the female Luer connection of the intravenous (IV) line, 3-ways tap with 10 cm extension.</li>
		<li>Attach an 18 G needle to the male end.</li>
		<li>Open the 3-way stop lock to allow for continuity from the needle to the syringe.</li>
		<li>Carefully unsheathe the needle and aspirate approximately 300 &#956;L of the saline into the IV line.</li>
		<li>Remove the needle from the saline and proceed to introduce in some air to create a small air bubble (5-10 mm) in the IV line.</li>
		<li>Place the needle into the tracer and aspirate all 500 &#956;L of the tracer. The saline in the IV line should be visibly separated by an air bubble.</li>
		<li>Discard the needle and close the 3-way stop lock.</li>
	</ol>
	</li>
	<li>Animal
	<ol>
		<li>Sedate a pig by intramuscular (i.m.) injection of tiletamine (3.75 mg/kg) and zolazepam (3.75 mg/kg) and dexmedetomidine (37.5 &#956;g/kg). Wait for it to become unconscious.</li>
		<li>Prepare an intravenous line by inserting a 20 G cannula into the ear vein. 
		&#8203;NOTE: Make sure the cannula is in the vein by injecting 5-10 mL of saline through the cannula. If the vein has been missed, this will be noticeable by small edema in the ear tissue.</li>
		<li>Intubate the pig to ensure that the breathing rate can be regulated throughout the surgery. 
		NOTE: Ensure successful intubation by applying pressure on the pig&#39;s thorax and confirm that forcibly expired air is coming out of the intubation tube.</li>
		<li>Attach the intubation tube to a ventilator set to a breath rate of 14 breaths/min.</li>
		<li>Connect a pulse oximeter and cuff to the tail to monitor the heart rate (HR), blood pressure (BP), and oxygen saturation (sats). Insert a rectal thermometer to monitor the core temperature.</li>
		<li>Prepare an IV bag of ketamine&#160;(5 mg/kg/min), midazolam (0.25 mg/kg/min), and fentanyl (2.5 &#956;g/kg/min), in saline and begin to infuse through the ear vein at approximately 2 drops/s. 
		NOTE: Throughout the surgery, the infusion rate may need to be increased or decreased based on the animal&#39;s vitals.</li>
		<li>With the pig in the prone position, palpate the back of the head and neck of the animal to locate and mark the occipital crest and spine of the first thoracic vertebrae and the base of each ear.</li>
		<li>Draw a straight line between the crest and the vertebrae along the longitudinal axis. Draw two lines from the crest to the base of each ear by following the base of the skull (Figure 1A).</li>
		<li>Check that the animal is in a deep sleep by carefully clamping the tail and watching for the absence of a tail reflex. 
		&#8203;NOTE: If the animal is still reflexive, the anesthetic infusion rate should be incrementally increased until the animal no longer exhibits a reflex.</li>
	</ol>
	</li>
</ol>
2. Surgery
NOTE: All through the surgery, it is necessary to have at least one assistant to suction the light bleeding and cauterize any severed vessels.
<ol>
	<li>Using a scalpel with a # 21 blade, make a dermal incision along the longitudinal line down to the muscle.</li>
	<li>Extend two perpendicular dermal incisions further along the shoulders, 10-15 cm in length.</li>
	<li>From the occipital crests, make dermal incisions along the line down to the base of each ear.</li>
	<li>Gripping the skin corners formed at the occipital crest with anatomical forceps, carefully separate the skin from the underlying muscle by lightly running the scalpel blade over the fascia, moving from the rostral to caudal. Once the skin has been resected following each of the five incisions, parts of the trapezius muscles should then be visible.</li>
	<li>Make a longitudinal incision with the scalpel, approximately 1 cm deep, where the trapezius comes together at the midline. 
	NOTE: When cutting through the muscles, there is an increased propensity for bleeding, so the cauterizer should be ready. If a larger vessel is severed, one person should quickly compress it with the gauze, while the other person uses the cauterizer.</li>
	<li>Using a combination of straight and curved surgical forceps, perform blunt dissection working along the longitudinal cut in the muscles. This will separate the bellies of the trapezius, as well as the underlying semispinalis capitus biventer muscle.</li>
	<li>Sever any persisting muscle fibers with a scalpel and continue blunt dissection until semispinalis capitus complexus becomes visible.</li>
	<li>Sever the origins of the trapezius and semispinalis capitus biventer muscles along the posterior aspect of the skull. Carefully separate them longitudinally with the scalpel performing blunt dissection until the semispinalis capitus complexus is fully visible.</li>
	<li>Retract the trapezius and semispinalis capitus biventer muscles using self-retaining retractors.</li>
	<li>Where the bellies of the semispinalis capitus complexus come together in the midline, make a longitudinal incision with the scalpel approximately 1 cm deep. 
	NOTE: Be aware for any additional bleeding here. Bleeding can be managed using a combination of cotton swabs and cauterization.</li>
	<li>Using surgical forceps, perform a blunt dissection working along the longitudinal cut between the muscle bellies until the dorsal aspect of the atlas (CI) is&#160;palpable.</li>
	<li>Sever the origins of the semispinalis capitus complexus muscles along the posterior aspect of the skull and separate it longitudinally from the underlying vertebrae by scalpel and blunt dissection.</li>
	<li>Retract the semispinalis capitus complexus muscles using another set of self-retaining retractors.</li>
	<li>Using a scalpel, carefully remove any remaining tissue overlying the region where the atlas meets the skull base.</li>
	<li>Placing one arm under the animal&#39;s neck and one finger at the juncture of the atlas and skull, simultaneously elevate the head and flex the neck while palpating with the finger to reveal the cisterna magna using the other hand. 
	&#8203;NOTE: The cisterna magna is recognizable when palpating as a strong elastic structure with a small amount of rebound as pressure is released with the finger.</li>
</ol>
3. Cannulation and injection
NOTE: This step also requires at least two people and is carried out with the animal&#39;s head elevated and neck flexed.
<ol>
	<li>Ensure that one person elevates and flexes the head and neck of the animal whilst the other palpates for the cisterna magna making a note of its anatomical location.</li>
	<li>Slowly and carefully introduce a 22 G cannula through the dura and into the cisterna magna at an oblique angle to the longitudinal axis. 
	NOTE: Do not insert the cannula too deep, as this can cause damage to the brain. Knowing how far to insert the cannula comes with experience in understanding how it feels for the cannula to pierce the dura. Essentially, just as the dura has been pierced, the cannula is then deep enough for a successful tracer injection. This depth is approximately 3-5 mm but will differ based on the size or age of the animal. Successful cannulation should be immediately evident through the visualization of clear, pulsatile CSF ascending the cannula. For the best outcome, it is recommended to practice several cannulations beforehand in euthanized animals to get one&#39;s understanding of dural piercing.</li>
	<li>Retract the needle from the cannula and place a cap on the lock.</li>
	<li>First, apply superglue and an accelerator where the cannula enters the tissue, followed by the application of the dental cement. Wait for 5 min for the cement to harden.</li>
	<li>Carefully remove the cap from the cannula and attach the male end of the previously prepared IV line tap with 10 cm extension, with the tracer, to the cannula.</li>
	<li>Slowly inject the tracer by hand or using a micro-infusion pump at a rate of 100 &#956;L/min. Remove the 3-ways IV line tap with 10 cm extension and replace it with the cap. Tracer should now be visible pulsating at the base of the cannula (Supplementary Video 1). 
	NOTE: If injecting by hand, do this until the tracer is just still visible in the cannula shaft, approximately 1-2 mm above where the dental cement is covering the shaft.</li>
	<li>Following injection, place sandbags under the neck to maintain some flexion. The head may then be released, and the animal is left in a resting prone position.</li>
	<li>Release the self-retaining retractors and place muscles as they lay before. Bring the skin together over the muscles using surgical towel clamps.</li>
	<li>Cover towel clamps and incision with gauze and then a blanket to limit heat loss.</li>
	<li>Allow the tracer to circulate for the desired time before euthanizing the animal by i.v. Pentobarbital injection (140 mg/kg). Confirm euthanasia by the absence of heart sounds upon auscultation with a stethoscope.</li>
</ol>
4. Brain extraction and processing
<ol>
	<li>Using a scalpel with 20-blade, extend the longitudinal dermal incision from the occipital crest to approximately 7 cm above the nose.</li>
	<li>Reflect the skin overlying the dorsal aspect of the skull using the scalpel. 
	NOTE: There are several ways to cut and remove the dorsal aspect of the pig skull on an animal-by-animal basis. What follows is the procedure that has worked most often for this experiment.</li>
	<li>Using a handheld compact saw, make a coronal cut in the skull, approximately 3 cm above the two large veins seen exiting the skull. Extend to two further vertical cuts from the coronal cuts and two more further cuts to bring the vertical cuts together in the midline. 
	NOTE: Maintain a firm grip of the saw when making the skull bone cuts as it will tend to pull away upon the first contact with the bone or tissue, which can lead to a severe injury.</li>
	<li>Ensure the skull cuts are through the entire thickness of the bone by following up with a hammer and narrow chisel (10 mm) to each of the cuts.</li>
	<li>Using the hammer, finally knock a wide chisel (25-30 mm) into the coronal cut. With one person supporting the head, ensure that the other person applies leverage on the chisel to wince open the dorsal skull.</li>
	<li>Once the dorsal skull fragment has been removed, dissect out the overlying dura mater using curved surgical scissors.</li>
	<li>Use a spatula to severe the spinal cord from the cerebellum at the rostral aspect. Then proceed to guide the spatula under the brain from the front, severing the olfactory bulbs, pituitary gland, and cranial nerves.</li>
	<li>Place the spatula behind the cerebellum and apply a fair amount of pressure to dislodge the brain from the cranial cavity, carefully lifting it out once loose.</li>
	<li>Immediately fix the whole brain by tissue immersion in 4% paraformaldehyde overnight. 
	NOTE: After this step, it is possible to carry out the whole brain imaging using a stereoscope (Figure 1E).</li>
	<li>The next day, make coronal slices of the brain using a salmon knife and fix the slices overnight by tissue immersion in 4% paraformaldehyde.</li>
	<li>Finally, place the slices in 0.01% azide in PBS for long-term storage.</li>
</ol>

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The authors have nothing to disclose.

Herein, is described, a detailed protocol to perform the direct cannulation of the cisterna magna in pigs, including the necessary preparation, surgical procedure, tracer infusion and extraction of the brain. This requires someone with experience and certification for working with large animals. If carried out correctly, this allows for the delivery of desired molecules with surety directly into the CSF, after which a series of different advanced light imaging modalities can be used to explore CSF distribution and glymph...

Cerebrospinal fluid (CSF) is an ultrafiltrate of blood which is found within and around the central nervous system (CNS)1,2. Apart from giving buoyancy to the brain or absorbing damaging mechanical forces, CSF also plays a pivotal role in clearing metabolic waste from the CNS3. Waste clearance is facilitated by the recently characterized glymphatic system which permits the convective flow of CSF through the brain parenchyma via perivascular spaces (PVS), which encircle penetrating arteries3,4,5. This process has been shown to be dependent on aquaporin-4 (AQP4), a water channel expressed primarily on the astrocytic endfeet, bound to the PVS4,6. The study of the glymphatic system is achieved by both in vivo and ex vivo imaging, using either advanced light microscopy or magnetic resonance imaging (MRI), following the introduction of a fluorescent/radioactive tracer or contrast agent into the CSF7,8,9,10,11.
An effective way to introduce a tracer into the CSF without incurring damage to the brain parenchyma is through cisterna magna cannulation (CMc)12,13. A large majority of all glymphatic studies, thus far have been carried out in rodents and avoided in higher mammals because of the invasiveness of CMc coupled to the practical simplicity of working with a small mammal. Additionally, the thin skulls of mice permit in vivo imaging without the need for a cranial window and subsequently allow for an uncomplicated brain extraction11,14. Experiments carried out in humans have yielded a valuable macroscopic in vivo data on the glymphatic function, but relied on intrathecal tracer injections in the distal lumbar spine and, furthermore, utilize MRI which does not yield sufficient resolution to capture the microanatomy the glymphatic system7,15,16. Understanding the architecture and extent of the glymphatic system in higher mammals is essential for its translation to humans. In order to facilitate glymphatic translation to humans, it is important to apply techniques that are carried out in rodents to higher mammals so as to allow for direct comparisons of the glymphatic system across species of increasing cognition and brain complexity17. Pig and human brains are gyrencephalic, possessing a folded neuroarchitecture, while rodent brains are lissencephalic, thereby having substantial difference among each other. In terms of the overall size, pig brains are, also, more comparable to humans, being 10-15 times smaller than the human brain, while mouse brains are 3,000 times smaller18. By better understanding the glymphatic system in large mammals, it may be possible to utilize the human glymphatic system for future therapeutic intervention in conditions such as stroke, traumatic brain injury and neurodegeneration. Direct CMc in pigs in vivo is a method that allows for the high-resolution light microscopy of the glymphatic system in a higher mammal. Furthermore, due to the size of the pigs used, it is possible to apply monitoring systems similar to those used in human surgeries making it feasible to tightly document and regulate vital functions in order to assess how these contribute to the glymphatic function.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.01% azide in PBS</td>
			<td>Sigmaaldrich</td>
			<td>S2002</td>
			<td></td>
		</tr>
		<tr>
			<td>18G needle</td>
			<td>Mediq</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1ml Syringe</td>
			<td>FischerSci</td>
			<td>15849152</td>
			<td></td>
		</tr>
		<tr>
			<td>20G cannula</td>
			<td>Mediq</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>22G cannula</td>
			<td>Mediq</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>4% paraformaldehyde</td>
			<td>Sigmaaldrich</td>
			<td>P6148</td>
			<td></td>
		</tr>
		<tr>
			<td>Anatomical forceps</td>
			<td>NA</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine serum albumin Alexa-Fluor 647 Conjugate</td>
			<td>ThermoFischer</td>
			<td>A34785</td>
			<td>2 vials (10mg)</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigmaaldrich</td>
			<td>C1016</td>
			<td></td>
		</tr>
		<tr>
			<td>Chisel</td>
			<td>ClasOhlson</td>
			<td>40-8870</td>
			<td></td>
		</tr>
		<tr>
			<td>Dental cement</td>
			<td>Agnthos</td>
			<td>7508</td>
			<td></td>
		</tr>
		<tr>
			<td>compact saw</td>
			<td>ClasOhlson</td>
			<td>40-9517</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigmaaldrich</td>
			<td>G8270</td>
			<td></td>
		</tr>
		<tr>
			<td>Hammer</td>
			<td>ClasOhlson</td>
			<td>40-7694</td>
			<td></td>
		</tr>
		<tr>
			<td>Insta-Set CA Accelerator</td>
			<td>BSI-Inc</td>
			<td>BSI-151</td>
			<td></td>
		</tr>
		<tr>
			<td>IV line TAP, 3-WAYS with 10cm extension</td>
			<td>Bbraun</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigmaaldrich</td>
			<td>P9333</td>
			<td></td>
		</tr>
		<tr>
			<td>Marker pen</td>
			<td>NA</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Sigmaaldrich</td>
			<td>M8266</td>
			<td></td>
		</tr>
		<tr>
			<td>MilliQ water</td>
			<td>NA</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCL</td>
			<td>Sigmaaldrich</td>
			<td>S7653</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Sigmaaldrich</td>
			<td>S8282</td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Sigmaaldrich</td>
			<td>S5761</td>
			<td></td>
		</tr>
		<tr>
			<td>No. 20 scalpel blade</td>
			<td>Agnthos</td>
			<td>BB520</td>
			<td></td>
		</tr>
		<tr>
			<td>No. 21 Scalpel blade</td>
			<td>Agnthos</td>
			<td>BB521</td>
			<td></td>
		</tr>
		<tr>
			<td>No. 4 Scalpel handle</td>
			<td>Agnthos</td>
			<td>10004-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Saline</td>
			<td>Mediq</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Salmon knife</td>
			<td>Fiskers</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Self-retaining retractors</td>
			<td>NA</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Superglue</td>
			<td>NA</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical curved scissors</td>
			<td>NA</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical forceps</td>
			<td>NA</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical towel clamps</td>
			<td>NA</td>
			<td>NA</td>
			<td></td>
		</tr>
	</tbody>
</table>

direct cannula implantation in the cisterna magna of pigs

The glymphatic system is a waste clearance system in the brain that relies on the flow of cerebrospinal fluid (CSF) in astrocyte-bound perivascular spaces and has been implicated in the clearance of neurotoxic peptides such as amyloid-beta. Impaired glymphatic function exacerbates disease pathology in animal models of neurodegenerative diseases, such as Alzheimer's, which highlights the importance of understanding this clearance system. The glymphatic system is often studied by cisterna magna cannulations (CMc), where tracers are delivered directly into the cerebrospinal fluid (CSF). Most studies, however, have been carried out in rodents. Here, we demonstrate an adaptation of the CMc technique in pigs. Using CMc in pigs, the glymphatic system can be studied at a high optical resolution in gyrencephalic brains and in doing so bridges the knowledge gap between rodent and human glymphatics.

Once the pig is unconscious, it is palpated, and its surface anatomy is marked, starting at the occipital crest (OC) and working towards the thoracic vertebrae (TV) and each ear base (EB). It is along these lines that the dermal incisions are made (Figure 1A). The three muscle layers including trapezius, semispinalis capitus biventer and semispinalis capitus complexus are resected and held open by two sets of self-retaining retractors to expose the cisterna magna (CM) (F...

Watch this Scientific Journal Video about Direct Cannula Implantation in the Cisterna Magna of Pigs at JoVE.com

Direct Cannula Implantation in the Cisterna Magna of Pigs

This article presents a step-by-step protocol for the direct cannula implantation in the cisterna magna of pigs.

The glymphatic system is a waste clearance system in the brain that relies on the flow of cerebrospinal fluid (CSF) in astrocyte-bound perivascular spaces ...

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3.6K Views.  Lund University. This article presents a step-by-step protocol for the direct cannula implantation in the cisterna magna of pigs. 

Video: Direct Cannula Implantation in the Cisterna Magna of Pigs

Electrode Positioning and Montage in Transcranial Direct Current Stimulation

Transcranial direct current stimulation (tDCS) is a technique that has been intensively investigated in the past decade as this method offers a non-invasive and safe alternative to change cortical excitability2. The effects of one session of tDCS can last for several minutes, and its effects depend on polarity of stimulation, such as that cathodal stimulation induces a decrease in cortical excitability, and anodal stimulation induces an increase in cortical excitability that may last beyond the duration of stimulation6. These effects have been explored in cognitive neuroscience and also clinically in a variety of neuropsychiatric disorders &#x2013; especially when applied over several consecutive sessions4. One area that has been attracting attention of neuroscientists and clinicians is the use of tDCS for modulation of pain-related neural networks3,5. Modulation of two main cortical areas in pain research has been explored: primary motor cortex and dorsolateral prefrontal cortex7. Due to the critical role of electrode montage, in this article, we show different alternatives for electrode placement for tDCS clinical trials on pain; discussing advantages and disadvantages of each method of stimulation.

Transcranial direct current stimulation (tDCS) is an established technique to modulate cortical excitability1,2. It has been used as an investigative tool in neuroscience due to its effects on cortical plasticity, easy operation, and safe profile. One area that tDCS has been showing encouraging results is pain alleviation 3-5.

Transcranial direct current stimulation (tDCS) is a technique that has been intensively investigated in the past decade as this method offers a ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Multimodal Imaging of Stem Cell Implantation in the Central Nervous System of Mice

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, both in preclinical and clinical studies. However, to date results regarding functional outcome and/or tissue regeneration following stem cell transplantation are quite diverse. Generally, a clinical benefit is observed without profound understanding of the underlying mechanism(s)1. Therefore, multiple efforts have led to the development of different molecular imaging modalities to monitor stem cell grafting with the ultimate aim to accurately evaluate survival, fate and physiology of grafted stem cells and/or their micro-environment. Changes observed in one or more parameters determined by molecular imaging might be related to the observed clinical effect. In this context, our studies focus on the combined use of bioluminescence imaging (BLI), magnetic resonance imaging (MRI) and histological analysis to evaluate stem cell grafting.
BLI is commonly used to non-invasively perform cell tracking and monitor cell survival in time following transplantation2-7, based on a biochemical reaction where cells expressing the Luciferase-reporter gene are able to emit light following interaction with its substrate (e.g. D-luciferin)8, 9. MRI on the other hand is a non-invasive technique which is clinically applicable10 and can be used to precisely locate cellular grafts with very high resolution11-15, although its sensitivity highly depends on the contrast generated after cell labeling with an MRI contrast agent. Finally, post-mortem histological analysis is the method of choice to validate research results obtained with non-invasive techniques with highest resolution and sensitivity. Moreover end-point histological analysis allows us to perform detailed phenotypic analysis of grafted cells and/or the surrounding tissue, based on the use of fluorescent reporter proteins and/or direct cell labeling with specific antibodies.
In summary, we here visually demonstrate the complementarities of BLI, MRI and histology to unravel different stem cell- and/or environment-associated characteristics following stem cell grafting in the CNS of mice. As an example, bone marrow-derived stromal cells, genetically engineered to express the enhanced Green Fluorescent Protein (eGFP) and firefly Luciferase (fLuc), and labeled with blue fluorescent micron-sized iron oxide particles (MPIOs), will be grafted in the CNS of immune-competent mice and outcome will be monitored by BLI, MRI and histology (Figure 1).

This article describes an optimized sequence of events for multimodal imaging of cellular grafts in rodent brain using: (i) in vivo bioluminescence and magnetic resonance imaging, and (ii) post mortem histological analysis. Combining these imaging modalities on a single animal allows cellular graft evaluation with high resolution, sensitivity and specificity.

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. Few studies have examined gene transfer to the central auditory system. The dorsal cochlear nucleus (DCN) of the brainstem, which contains second order neurons of the auditory pathway, is a potential site for gene transfer. In this protocol, a technique for direct and maximal exposure of the murine DCN via a posterior fossa approach is demonstrated. This approach allows for either acute or survival surgery. Following direct visualization of the DCN, a host of experiments are possible, including injection of opsins into the cochlear nucleus and subsequent stimulation by an optical fiber coupled to a blue light laser.&#160;Other neurophysiology experiments, such as electrical stimulation and neural injector tracings are also feasible.&#160;The level of visualization and the duration of stimulation achievable make this approach applicable to a wide range of experiments.

The goal of this protocol is to outline a surgical approach to provide direct access to the dorsal cochlear nucleus in a murine model.

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. ...

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 ...

Laminectomy and Spinal Cord Window Implantation in the Mouse

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated digital vaporizer is utilized to achieve a stable plane of anesthesia at a low-flow rate of isoflurane. A single vertebral spine is removed, and a commercially available cover-glass is overlaid on a thin agarose bed. A 3D-printed plastic backplate is then affixed to the adjacent vertebral spines using tissue adhesive and dental cement. A stabilization platform is used to reduce motion artifact from respiration and heartbeat. This rapid and clamp-free method is well-suited for acute multi-photon fluorescence microscopy. Representative data are included for an application of this technique to two-photon microscopy of the spinal cord vasculature in transgenic mice expressing eGFP:Claudin-5 &#8212; a tight junction protein.

This protocol describes implantation of a glass window onto the spinal cord of a mouse to facilitate visualization by intravital microscopy.

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated ...

Chronic Implantation of Whole-cortical Electrocorticographic Array in the Common Marmoset

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent development of thin, flexible ECoG electrodes has enabled conduction of stable recordings of large-scale cortical activity. We have developed a whole-cortical ECoG array for the common marmoset. The array continuously covers almost the entire lateral surface of cortical hemisphere, from the occipital pole to the temporal and frontal poles, and it captures whole-cortical neural activity in one shot. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain. Marmosets have two advantages regarding ECoG recordings, one being the homologous organization of anatomical structures in humans and macaques, including frontal, parietal, and temporal complexes. The other advantage is that the marmoset brain is lissencephalic and contains a large number of complexes, which are more difficult to access in macaques with ECoG, that are exposed to the brain surface.These features allow direct access to most cortical areas beneath the surface of the brain. This system provides an opportunity to investigate global cortical information processing with high resolutions at a sub-millisecond order in time and millimeter order in space.

We have developed a whole-cortical electrocorticographic array for the common marmoset that continuously covers almost the entire lateral surface of cortex, from the occipital pole to the temporal and frontal poles. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain.

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent ...

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset (Callithrix jacchus)

Marmosets (Callithrix jacchus) are small non-human primates that are gaining popularity in biomedical and preclinical research, including the neurosciences. Phylogenetically, these animals are much closer to humans than rodents. They also display complex behaviors, including a wide range of vocalizations and social interactions. Here, an effective stereotaxic neurosurgical procedure for implantation of recording electrode arrays in the common marmoset is described. This protocol also details the pre- and postoperative steps of animal care that are required to successfully perform such a surgery. Finally, this protocol shows an example of local field potential and spike activity recordings in a freely behaving marmoset 1 week after the surgical procedure. Overall, this method provides an opportunity to study the brain function in awake and freely behaving marmosets. The same protocol can be readily used by researchers working with other small primates. In addition, it can be easily modified to allow other studies requiring implants, such as stimulating electrodes, microinjections, implantation of optrodes or guide cannulas, or ablation of discrete tissue regions.

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset Callithrix jacchus

This work presents a protocol to perform a stereotaxic, neurosurgical implantation of&#160;microelectrode arrays in the common marmoset. This method specifically enables electrophysiological recordings in freely behaving animals but can be easily adapted to any other similar neurosurgical intervention in this species (e.g., cannula for drug administration or electrodes for brain stimulation).

Marmosets (Callithrix jacchus) are small non-human primates that are gaining popularity in biomedical and preclinical research, including the ...

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.

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