This study was supported by grants from the Swedish Research Council, the Swedish Cancer Society, the Swedish Foundation for Strategic Research, Knut och Alice Wallenbergs Stiftelse and the Strategic Research Programme in Stem Cells and Regenerative Medicine at Karolinska Institutet (StratRegen).

The surgical procedures hereby presented were approved by Stockholms Norra Djurf&#246;rs&#246;ksetiska N&#228;mnd and carried out in agreement with specifications provided by the research institute (Karolinska Institute, Sweden).
NOTE: Intracisternal injection can be flexibly adapted for multiple research purposes. We present below a procedure developed to efficiently label leptomeningeal cells for fate mapping based on injection of endoxifen in a transgenic mouse line carrying R26R-tdTomato12 and CreER, the latter under the Cx30 promoter13. Labelling of this population of cells may be achieved through injection of viral constructs using the same procedure outlined below. Finally, this approach can be employed for tracking cerebrospinal fluid flow, by infusion of fluorescent tracers.
1. Preparation of the Injection System
NOTE: We recommend carrying out the procedure in a suitable surgical room, and in aseptic conditions. Surgical tools can be sterilized using heat (autoclave, glass bead sterilizer) or sanitized using a high-level chemical disinfectant if they are heat-sensitive. Rinse the instruments before use when employing chemical disinfection or allow them to cool down when sanitized with heat.
<ol>
	<li>Using forceps, bend the needle of the injection syringe to 30&#176; at 3 mm from the tip. 
	NOTE: Use Hamilton syringes with a 30 G beveled needle.</li>
	<li>Prepare 1 mg/mL of endoxifen solution, diluted in 10% DMSO and backfill the syringe with the bent needle. 
	NOTE: Administer 5 &#956;L of the compound to ensure widespread exposure of meningeal cells in the adult C57Bl/6j mouse (ca. 25-30 g), although pilot experiments that test different concentrations and injection volumes may be necessary when treating animals of different ages and strains.</li>
	<li>Adjust the mouse head holder so that the mouth piece is at approximately 30&#176; from the surface of the surgical table. 
	NOTE: A stereotactic frame with three-point fixation (i.e., ears and mouth) can also be used for this procedure. In this case, however, the animal will only be fixed with the mouth piece, whereas ear bars can be extended under the animal&#39;s forelimbs and be used to support the animal&#39;s body during the procedure.</li>
</ol>
2. Induction of Anesthesia
<ol>
	<li>For injectable anesthetics, use concentrations and modes of administration recommended by the veterinary unit at the local institution.</li>
	<li>For inhalational anesthetics, such as isoflurane, prepare the administration unit according to the manufacturer&#39;s specifications.</li>
	<li>With isoflurane, set concentration of the compound at 4% for induction of anesthesia.</li>
	<li>Deliver air at a rate of 400 mL/min.</li>
	<li>Allow the anesthesia unit to fill the chamber with the anesthetic for a few minutes and place the mouse in the chamber afterwards. 
	NOTE: For the present experiments, we have used adult (&#62;2 months old and approximately 30 g) male and female transgenic mice carrying Cx30-CreER and R26R-tdTomato, and bred on a C57Bl/6j background.</li>
	<li>Monitor the animal during induction of anesthesia. The breathing pattern should slow down when the mouse is under full anesthesia.</li>
	<li>Remove the animal from the chamber and check for suppression of the paw reflex by delicately pinching the hind paws. 
	NOTE: The reflex may still be present but delayed a couple of seconds. If that is the case, place the animal back in the chamber until complete suppression of the paw reflex has been achieved.</li>
	<li>Administer analgesic (e.g., Carprofen, 5 mg/kg through subcutaneous injection) to aid post-operative pain management.</li>
</ol>
3. Positioning of the Animal for the Procedure
<ol>
	<li>Fix the animal&#39;s head onto the head holder. To improve accessibility to the cisterna magna, position the animal&#39;s body at approximately 30&#176; from the surface of the table and the head titled downwards, to establish an angle of 120&#176; with the rest of the body and extend the back of the neck to facilitate access to the cisterna magna (Figure 1A).</li>
	<li>Add paper towels underneath the animal to support its body throughout the procedure.</li>
	<li>Secure anesthesia delivery through the mouth piece and reduce isoflurane concentration to 2.5% and air delivery to 200 ml/min.</li>
	<li>Apply ophthalmic ointment.</li>
</ol>
4. Exposure of the Cisterna Magna
<ol>
	<li>Shave the back of the animal&#39;s neck and sanitize the area with 70% ethanol and Betadine.</li>
	<li>Using surgical scissors, perform a midline incision (ca. 7 mm in length) starting at the level of the occipital bone and extending it posteriorly.</li>
	<li>Gently separate superficial connective tissue and neck muscles pulling sidewise from the midline with fine tip tweezers. This will expose the dural membrane overlaying the cisterna magna, shaped as an inverted triangle.</li>
	<li>Use sterile absorption spears or cotton swabs to control any resultant bleeding.</li>
	<li>Position a small surgical separator to maintain the neck muscles pulled aside and enable visualization of the cisterna magna throughout the procedure.</li>
</ol>
5. Intracisternal Injection
<ol>
	<li>To gain access to the cisterna magna, identify the caudal end of the occipital bone and insert the needle that was previously bent immediately underneath. 
	NOTE: There will be a sudden drop in resistance as the dural membrane is punctured. However, the tip of the needle will only penetrate slightly underneath the meningeal surface, thanks to its hooked shape.</li>
	<li>Once the dura has been perforated, allow the bent tip of the needle to penetrate underneath the dural surface by gently pulling the syringe upwards and parallel to the animal&#39;s body, in order to &#34;hook&#34; the needle to the skull (see Figure 1B). This will ensure better stability and will prevent the needle from penetrating deeper, thus avoiding the risk of damaging the underlying cerebellum or medulla.</li>
	<li>Inject the compound slowly to avoid interference with the cerebrospinal fluid&#39;s natural flow. 
	NOTE: Depending on the purpose of the experiment, the infusion rate may vary. If slow and steady infusion rate is required (e.g., when using the procedure to trace cerebrospinal fluid movement), it may be advisable to use an automatized microinfusion system in combination with the syringe.</li>
	<li>After the injection, let the needle rest in site for 1 min and carefully remove it. Use fine tip forceps to aid retraction of the needle from its hooked position. 
	NOTE: The use of a thin needle (e.g., 30 G) does not lead to substantial damage of the meningeal membrane, and consequent outflow of the cerebrospinal fluid. If larger needle sizes are required, we recommend&#160;stopping the leaking of fluid by applying pressure at the injection site using a sterile cotton tip.</li>
</ol>
6. Concluding the Procedure and Post-operative Care
<ol>
	<li>Remove the separator and allow the muscles to go back to their original position overlaying the dural membrane.</li>
	<li>Close the skin incision using a few drops of cyanoacrylate adhesive. 
	NOTE: Alternatively, use absorbable sutures (e.g., 5-0 vicryl sutures) and close the skin incision with interrupted stitches.</li>
	<li>Apply local anesthetic (e.g., 100 &#956;L of 5% lidocaine) at the incision site.</li>
	<li>Remove the animal from the holder and place in a clean cage positioned on a heating pad. Monitor the animal until it regains consciousness. 
	NOTE: The procedure is not expected to cause long-lasting pain or distress in the animal. Nevertheless, the mouse should be monitored closely for the first postoperative days and pain relief measures may be undertaken whenever deemed necessary, and in accordance with recommendations provided by the veterinary unit at your institution.</li>
</ol>

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	<li>Vanlandewijck, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+molecular+atlas+of+cell+types+and+zonation+in+the+brain+vasculature.">A molecular atlas of cell types and zonation in the brain vasculature.</a> Nature. 554 (7693), 475-480 (2018).</li><li>Whish, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+inner+CSF-brain+barrier:+developmentally+controlled+access+to+the+brain+via+intercellular+junctions.">The inner CSF-brain barrier: developmentally controlled access to the brain via intercellular junctions.</a> Frontiers in Neuroscience. 9, 16 (2015).</li><li>Weller, R. O., Sharp, M. M., Christodoulides, M., Carare, R. O., Mollgard, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+meninges+as+barriers+and+facilitators+for+the+movement+of+fluid,+cells+and+pathogens+related+to+the+rodent+and+human+CNS.">The meninges as barriers and facilitators for the movement of fluid, cells and pathogens related to the rodent and human CNS.</a> Acta Neuropathologica. 135 (3), 363-385 (2018).</li><li>Choe, Y., Siegenthaler, J. A., Pleasure, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+cascade+of+morphogenic+signaling+initiated+by+the+meninges+controls+corpus+callosum+formation.">A cascade of morphogenic signaling initiated by the meninges controls corpus callosum formation.</a> Neuron. 73 (4), 698-712 (2012).</li><li>Siegenthaler, J. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Retinoic+acid+from+the+meninges+regulates+cortical+neuron+generation.">Retinoic acid from the meninges regulates cortical neuron generation.</a> Cell. 139 (3), 597-609 (2009).</li><li>Bifari, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neurogenic+Radial+Glia-like+Cells+in+Meninges+Migrate+and+Differentiate+into+Functionally+Integrated+Neurons+in+the+Neonatal+Cortex.">Neurogenic Radial Glia-like Cells in Meninges Migrate and Differentiate into Functionally Integrated Neurons in the Neonatal Cortex.</a> Cell Stem Cell. 20 (3), 360-373 (2017).</li><li>De Bock, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+angle+on+blood-CNS+interfaces:+a+role+for+connexins?.">A new angle on blood-CNS interfaces: a role for connexins?.</a> FEBS Letters. 588 (8), 1259-1270 (2014).</li><li>Ramos, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cisterna+Magna+Injection+in+Rats+to+Study+Glymphatic+Function.">Cisterna Magna Injection in Rats to Study Glymphatic Function.</a> Methods in Molecular Biology. 1938, 97-104 (2019).</li><li>Xavier, A. L. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cannula+Implantation+into+the+Cisterna+Magna+of+Rodents.">Cannula Implantation into the Cisterna Magna of Rodents.</a> Journal of Visualized Experiments. (135), (2018).</li><li>Iliff, J. 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The authors declare no competing interests.

The protocol outlined here presents a straightforward and reproducible procedure to label leptomeningeal cells for fate mapping. We use intracisternal injection of endoxifen, an active metabolite of Tamoxifen, to induce expression of tdTomato fluorescent reporter in Cx30-CreER; R26R-tdTomato mice12,13.
Compared to other protocols used for gaining access to the cerebrospinal fluid through the cisterna magna9, our...

Leptomeningeal cells are a fibroblast-like population of cells organized in a thin layer overlaying the brain and expressing genes implicated in collagen crosslinking (e.g., Dcn and Lum), and in the establishment of a brain-meningeal barrier (e.g., Cldn11)1,2. Leptomeningeal cells are implicated in a wide range of physiological functions, from strict control over the cerebrospinal fluid drainage3 to guidance of neural progenitors in the developing brain4,5. A recent study has also proposed that leptomeninges in the newborn may harbor radial glia-like cells that migrate into the brain parenchyma and develop into functional cortical neurons6.
Leptomeningeal cells are located in close proximity to surface astrocytes and share with them, as well as other parenchymal astroglia, expression of connexin-30 (Cx30)7. The surgical procedure outlined below allows widespread and specific labelling of these meningeal cells via a one-time delivery of endoxifen into the cisterna magna of transgenic mice conditionally expressing tdTomato in Cx30+ cells (i.e., using a CreER-loxP system for fate mapping). Endoxifen is an active metabolite of Tamoxifen and induces recombination of CreER-expressing cells in the same way as Tamoxifen does. It is, however, the recommended solution for topical application because it dissolves in 5-10% DMSO, instead of high concentrations of ethanol. Additionally, endoxifen does not cross the brain-meningeal barrier, thereby enabling specific recombination of leptomeningeal cells, without labelling of the underlying Cx30+ astroglial population (see Representative Results).
The technique presented here aims at manually and safely injecting the compound in the cerebrospinal fluid, via direct access to the cisterna magna. Unlike other, more invasive procedures requiring craniotomy, this approach allows to infuse compounds without causing damage to the skull or the brain parenchyma. Thus, it is not associated with the induction of inflammatory reactions triggered by activation of parenchymal glia cells. Similar to other injection strategies described before8,9,10, the present approach relies on the surgical exposure of the atlanto-occipital dural membrane covering the cisterna magna, after blunt dissection of the overlaying neck muscles. However, unlike for other procedures, we recommend the use of a needle bent at the tip, which can be stabilized against the occipital bone during administration. This will prevent the risk of the needle penetrating too deep and damaging the underlying cerebellum and medulla.
This surgical procedure is compatible with lineage tracing investigations that aim at mapping changes in cell identities and migration routes through parenchymal layers. It can also be adapted to genetic ablation studies that intend to probe the role of leptomeningeal cells in health and disease, such as their contribution to cortical development5 or the spreading of bacterial meningitis3,11. Finally, it can be utilized to track cerebrospinal fluid movement when combined with delivery of fluorescent tracers in wildtype animals.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Anesthesia unit</td>
			<td>Univentor 410</td>
			<td>8323102</td>
			<td>Complete of vaporizer, chamber, and tubing that connects to chamber and mouse head holder</td>
		</tr>
		<tr>
			<td>Anesthesia (Isoflurane)</td>
			<td>Baxter Medical AB</td>
			<td>000890</td>
			<td></td>
		</tr>
		<tr>
			<td>Betadine</td>
			<td>Sigma-Aldrich</td>
			<td>PVP1</td>
			<td></td>
		</tr>
		<tr>
			<td>Carprofen</td>
			<td>Orion Pharma AB</td>
			<td>014920</td>
			<td>Commercial name Rymadil</td>
		</tr>
		<tr>
			<td>Cyanoacrylate glue</td>
			<td>Carl Roth</td>
			<td>0258.1</td>
			<td>Use silk 5-0 sutures, in alternative</td>
		</tr>
		<tr>
			<td>Medbond Tissue Glue</td>
			<td>Stoelting</td>
			<td>50479</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma-Aldrich</td>
			<td>D2650</td>
			<td></td>
		</tr>
		<tr>
			<td>Endoxifen</td>
			<td>Sigma-Aldrich</td>
			<td>E8284</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol 70%</td>
			<td>Histolab</td>
			<td>01370</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton syringe (30G beveled needle)</td>
			<td>Hamilton</td>
			<td>80300</td>
			<td></td>
		</tr>
		<tr>
			<td>Lidocaine</td>
			<td>Aspen Nordic</td>
			<td>520455</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse head holder</td>
			<td>Narishige International</td>
			<td>SGM-4</td>
			<td>With mouth piece for inhalational anhestetics. Alternatively, use a stereotactic frame</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>Fine Science Tools</td>
			<td>15009-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Shaver</td>
			<td>Aesculap</td>
			<td>GT420</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile absorption spears</td>
			<td>Fine Science Tools</td>
			<td>18105-01</td>
			<td>Sterile cotton swabs are also a good option</td>
		</tr>
		<tr>
			<td>Surgical separator</td>
			<td>World Precision Instrument</td>
			<td>501897</td>
			<td></td>
		</tr>
		<tr>
			<td>Tweezers</td>
			<td>Dumont</td>
			<td>11251-35</td>
			<td></td>
		</tr>
		<tr>
			<td>Viscotears</td>
			<td>Bausch&#38;Lomb Nordic AB</td>
			<td>541760</td>
			<td></td>
		</tr>
	</tbody>
</table>

induction of leptomeningeal cells modification via intracisternal injection

The protocol outlined here describes how to safely and manually inject solutions through the cisterna magna while eliminating the risk of damage to the underlying parenchyma. Previously published protocols recommend using straight needles that should be lowered to a maximum of 1-2 mm from the dural surface. The sudden drop in resistance once the dural membrane has been punctured makes it difficult to maintain the needle in a steady position. Our method, instead, employs a needle bent at the tip that can be stabilized against the occipital bone of the skull, thus preventing the syringe from penetrating into the tissue after perforation of the dural membrane. The procedure is straightforward, reproducible, and does not cause long-lasting discomfort in the operated animals. We describe the intracisternal injection strategy in the context of genetic fate mapping of vascular leptomeningeal cells. The same technique can, furthermore, be utilized to address a wide range of research questions, such as probing the role of leptomeninges in neurodevelopment and the spreading of bacterial meningitis, through genetic ablation of genes putatively implicated in these phenomena. Additionally, the procedure can be combined with an automatized infusion system for a constant delivery and used for tracking cerebrospinal fluid movement via injection of fluorescently labelled molecules.

Intracisternal injection of endoxifen in transgenic mice expressing CreER under the Cx30 promoter13 and an inducible fluorescent reporter allows for specific recombination of leptomeningeal cells without labelling the neighboring Cx30-expressing surface and parenchymal astrocytes in the cortex (Figure 1). In order to gain access to the cisterna magna, the anesthetized animal is positioned with its body and its head at an angle of appro...

Watch this Scientific Journal Video about Induction of Leptomeningeal Cells Modification Via Intracisternal Injection at JoVE.com

Induction of Leptomeningeal Cells Modification Via Intracisternal Injection

We describe an intracisternal injection that employs a needle bent at the tip that can be stabilized to the skull, thus eliminating the risk of damage to the underlying parenchyma. The approach can be used for genetic fate mapping and manipulations of leptomeningeal cells and for tracking cerebrospinal fluid movement.

The protocol outlined here describes how to safely and manually inject solutions through the cisterna magna while eliminating the risk of damage to the ...

induction-leptomeningeal-cells-modification-via-intracisternal

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7.9K Views.  Karolinska Institute. We describe an intracisternal injection that employs a needle bent at the tip that can be stabilized to the skull, thus eliminating the risk of damage to the underlying parenchyma. The approach can be used for genetic fate mapping and manipulations of leptomeningeal cells and for tracking cerebrospinal fluid movement.

Video: Induction of Leptomeningeal Cells Modification Via Intracisternal Injection

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Designing, Packaging, and Delivery of High Titer CRISPR Retro and Lentiviruses via Stereotaxic Injection

Replication defective lentiviruses or retroviruses are capable of stably integrating transgenes into the genome of an infected host cell. This technique has been widely used to encode fluorescent proteins, opto- or chemo-genetic controllers of cell activity, or heterologous expression of human genes in model organisms. These viruses have also successfully been used to deliver recombinases to relevant target sites in transgenic animals, or even deliver small hairpin or micro RNAs in order to manipulate gene expression. While these techniques have been fruitful, they rely on transgenic animals (recombinases) or frequently lack high efficacy and specificity (shRNA/miRNA). In contrast, the CRISPR/Cas system uses an exogenous Cas nuclease which targets specific sites in an organism's genome via an exogenous guide RNA in order to induce double stranded breaks in DNA. These breaks are then repaired by non-homologous end joining (NHEJ), producing insertion and deletion (indel) mutations that can result in deleterious missense or nonsense mutations. This manuscript provides detailed methods for the design, production, injection, and validation of single lenti/retro virus particles that can stably transduce neurons to express a fluorescent reporter, Cas9, and sgRNAs to knockout genes in a model organism.

The CRISPR/Cas9 system offers the potential to make targeted genome editing accessible and affordable to the scientific community. This protocol is intended to demonstrate how to create viruses that will knockout a gene of interest using the CRISPR/Cas9 system, and then inject them stereotaxically into the adult mouse brain.

Replication defective lentiviruses or retroviruses are capable of stably integrating transgenes into the genome of an infected host cell. This technique ...

An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach

Subretinal injections have been successfully used in both humans and rodents to deliver therapeutic interventions of proteins, viral agents, and cells to the interphotoreceptor/subretinal compartment that has direct exposure to photoreceptors and the retinal pigment epithelium (RPE). Subretinal injections of plasminogen as well as recent preclinical and clinical trials have demonstrated safety and/or efficacy of delivering viral vectors and stem cells to individuals with advanced retinal disease. Mouse models of retinal disease, particularly hereditary retinal dystrophies, are essential for testing these therapies. The most common injection procedure in rodents is to use small transcorneal or transcleral incisions with an anterior approach to the retina. With this approach, the injection needle penetrates the neurosensory retina disrupting the underlying RPE and on insertion can easily nick the lens, causing lens opacification and impairment of noninvasive imaging. Accessing the subretinal space via a transcleral, posterior approach avoids these problems: the needle crosses the sclera approximately 0.5 mm from the optic nerve, without retinal penetration and avoids disrupting the vitreous. Collateral damage is limited to that associated with the focal sclerotomy and the effects of a transient, serous retinal detachment. The simplicity of the method minimizes ocular injury, ensures rapid retinal reattachment and recovery, and has a low failure rate. The minimal damage to the retina and RPE allows for clear assessment of the efficacy and direct effects of the therapeutic agents themselves. This manuscript describes a novel subretinal injection technique that can be used to target viral vectors, pharmacological agents, stem cells or induced pluripotent stem (iPS) cells to the subretinal space in mice with high efficacy, minimal damage, and fast recovery.

An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach

Subretinal injections are the most common technique for delivering large therapeutic agents such as proteins and viral vectors to photoreceptors and the retinal pigment epithelium. An alternative method in mice that successfully targets the subretinal space with minimal collateral damage and fast recovery times is described here.

Subretinal injections have been successfully used in both humans and rodents to deliver therapeutic interventions of proteins, viral agents, and cells to ...

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 ...

Brain Death Induction in Mice Using Intra-Arterial Blood Pressure Monitoring and Ventilation via Tracheostomy

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain death (BD) still represents the major source for solid transplants. Unfortunately, the irreversible loss of brain function is known to induce multiple pathophysiological changes, including hemodynamic as well as hormonal modifications, finally leading to a systemic inflammatory response. Models that allow a systematic investigation of these effects in vivo are scarce. We present a murine model of BD induction, which could aid investigations into the devastating effects of BD on allograft quality. After implementing intra-arterial blood pressure measurement via the common carotid artery and reliable ventilation via a tracheostomy, BD is induced by steadily increasing intracranial pressure using a balloon catheter. Four hours after BD induction, organs may be harvested for analysis or for further transplantation procedures. Our strategy enables the comprehensive analysis of donor BD in a murine model, therefore allowing an in-depth understanding of BD-related effects in solid organ transplantation and potentially paving the way to optimized organ preconditioning.

We present a murine model of brain death induction in order to evaluate the influence of its pathophysiological effects on organs as well as on consecutive grafts in the context of solid organ transplantation.

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain ...

Shuttle Box Assay as an Associative Learning Tool for Cognitive Assessment in Learning and Memory Studies using Adult Zebrafish

Cognitive deficits, including impaired learning and memory, are a primary symptom of various developmental and age-related neurodegenerative diseases and traumatic brain injury (TBI). Zebrafish are an important neuroscience model due to their transparency during development and robust regenerative capabilities following neurotrauma. While various cognitive tests exist in zebrafish, most of the cognitive assessments that are rapid examine non-associative learning. At the same time, associative-learning assays often require multiple days or weeks. Here, we describe a rapid associative-learning test that utilizes an adverse stimulus (electric shock) and requires minimal preparation time. The shuttle box assay, presented here, is simple, ideal for novice investigators, and requires minimal equipment. We demonstrate that, following TBI, this shuttle box test reproducibly assesses cognitive deficit and recovery from young to old zebrafish. Additionally, the assay is adaptable to examine either immediate or delayed memory. We demonstrate that both a single TBI and repeated TBI events negatively affect learning and immediate memory but not delayed memory. We, therefore, conclude that the shuttle box assay reproducibly tracks the progression and recovery of cognitive impairment.

Learning and memory are potent metrics in studying either developmental, disease-dependent, or environmentally induced cognitive impairments. Most cognitive assessments require specialized equipment and extensive time commitments. However, the shuttle box assay is an associative learning tool that utilizes a conventional gel box for rapid and reliable assessment of adult zebrafish cognition.

Cognitive deficits, including impaired learning and memory, are a primary symptom of various developmental and age-related neurodegenerative diseases and ...

Organotypic Cultures of Adult Human Cortex as an Ex vivo Model for Human Stem Cell Transplantation and Validation

Neurodegenerative disorders are common and heterogeneous in terms of their symptoms and cellular affectation, making their study complicated due to the lack of proper animal models that fully mimic human diseases and the poor availability of post-mortem human brain tissue. Adult human nervous tissue culture offers the possibility to study different aspects of neurological disorders. Molecular, cellular, and biochemical mechanisms could be easily addressed in this system, as well as testing and validating drugs or different treatments, such as cell-based therapies. This method combines long-term organotypic cultures of the adult human cortex, obtained from epileptic patients undergoing resective surgery, and ex vivo intracortical transplantation of induced pluripotent stem cell-derived cortical progenitors. This method will allow the study of cell survival, neuronal differentiation, the formation of synaptic inputs and outputs, and the electrophysiological properties of human-derived cells after transplantation into intact adult human cortical tissue. This approach is an important step prior to the development of a 3D human disease modeling platform that will bring basic research closer to the clinical translation of stem cell-based therapies for patients with different neurological disorders and allow the development of new tools for reconstructing damaged neural circuits.

Organotypic Cultures of Adult Human Cortex as an Ex vivo Model for Human Stem Cell Transplantation and Validation

This protocol describes long-term organotypic cultures of adult human cortex combined with ex vivo intracortical transplantation of induced pluripotent stem cell-derived cortical progenitors, which present a novel methodology to further test stem cell-based therapies for human neurodegenerative disorders.

Neurodegenerative disorders are common and heterogeneous in terms of their symptoms and cellular affectation, making their study complicated due to the ...

Establishing In Vitro LLEC Cultures Using Magnetic Cell Selection

Harvest and Primary Culture of Leptomeningeal Lymphatic Endothelial Cells

Leptomeningeal lymphatic endothelial cells (LLECs) are a recently discovered intracranial cellular population with a unique distribution clearly distinct from peripheral lymphatic endothelial cells. Their cellular function and clinical implications remain largely unknown. Consequently, the availability of a supply of LLECs is essential for conducting functional research in vitro. However, there is currently no existing protocol for harvesting and culturing LLECs in vitro.
This study successfully harvested LLECs using a multi-step protocol, which included coating the flask with fibronectin, dissecting the leptomeninges with the assistance of a microscope, enzymatically digesting the leptomeninges to prepare a single-cell suspension, inducing the expansion of LLECs with vascular endothelial growth factor-C (VEGF-C), and selecting lymphatic vessel hyaluronic receptor-1 (LYVE-1) positive cells through magnetic-activated cell sorting (MACS). This process ultimately led to the establishment of a primary culture. The purity of the LLECs was confirmed through immunofluorescence staining and flow cytometric analysis, with a purity level exceeding 95%. This multi-step protocol has demonstrated reproducibility and feasibility, which will greatly facilitate the exploration of the cellular function and clinical implications of LLECs.

Author Spotlight: Unveiling Cellular Functions and Potential Clinical Implications of Leptomeningeal Lymphatic Endothelial Cells 

Leptomeningeal lymphatic endothelial cells (LLECs), a recently identified intracranial cell type, have poorly understood functions. This study presents a reproducible protocol for harvesting LLECs from mice and establishing in vitro primary cultures. This protocol is designed to enable researchers to delve into the cellular functions and potential clinical implications of LLECs.

Leptomeningeal lymphatic endothelial cells (LLECs) are a recently discovered intracranial cellular population with a unique distribution clearly distinct ...