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. J., 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+paravascular+pathway+facilitates+CSF+flow+through+the+brain+parenchyma+and+the+clearance+of+interstitial+solutes,+including+amyloid+beta.">A paravascular pathway facilitates CSF flow through the brain parenchyma and the clearance of interstitial solutes, including amyloid beta.</a> Science Translational Medicine. 4 (147), (2012).</li><li>Coureuil, M., Lecuyer, H., Bourdoulous, S., Nassif, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+journey+into+the+brain:+insight+into+how+bacterial+pathogens+cross+blood-brain+barriers.">A journey into the brain: insight into how bacterial pathogens cross blood-brain barriers.</a> Nature Reviews Microbiology. 15 (3), 149-159 (2017).</li><li>Madisen, L., 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=Transgenic+mice+for+intersectional+targeting+of+neural+sensors+and+effectors+with+high+specificity+and+performance.">Transgenic mice for intersectional targeting of neural sensors and effectors with high specificity and performance.</a> Neuron. 85 (5), 942-958 (2015).</li><li>Slezak, 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=Transgenic+mice+for+conditional+gene+manipulation+in+astroglial+cells.">Transgenic mice for conditional gene manipulation in astroglial cells.</a> Glia. 55 (15), 1565-1576 (2007).</li><li>Hardy, S. J., Christodoulides, M., Weller, R. O., Heckels, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interactions+of+Neisseria+meningitidis+with+cells+of+the+human+meninges.">Interactions of Neisseria meningitidis with cells of the human meninges.</a> Molecular Microbiology. 36 (4), 817-829 (2000).</li><li>Colicchio, 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=The+meningococcal+ABC-Type+L-glutamate+transporter+GltT+is+necessary+for+the+development+of+experimental+meningitis+in+mice.">The meningococcal ABC-Type L-glutamate transporter GltT is necessary for the development of experimental meningitis in mice.</a> Infection and Immunity. 77 (9), 3578-3587 (2009).</li><li>Ricci, 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=Inhibition+of+matrix+metalloproteinases+attenuates+brain+damage+in+experimental+meningococcal+meningitis.">Inhibition of matrix metalloproteinases attenuates brain damage in experimental meningococcal meningitis.</a> BMC Infectious Diseases. 14, 726 (2014).</li></ol>

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 approach ensures a safe manual administration thanks to the use of a bent needle that can be stabilized to the occipital bone of the skull. Once the dural membrane of the cisterna magna is perforated, there is a sudden drop in resistance. At this point, other protocols recommend lowering the needle to a maximum of 1-2 mm from the dural surface and manually keeping it in a steady position throughout the procedure9. As opposed to a straight needle, the hooked needle is secured to the skull and cannot penetrate deeper in the tissue, thus eliminating the risk of damaging the underlying cerebellum or medulla. Our hooked system allows for a safer administration of solution, particularly when using a slow rate of infusion.
The procedure outlined here is not expected to cause long-lasting discomfort to the operated animal. Care must be taken, however, when administering large volumes of solution. A fast delivery rate may lead to alterations in the intracranial pressure and the development of neurological symptoms in the mouse. We suggest injecting volumes up to 5-10 &#956;L to avoid this risk or assemble the syringe onto a micromanipulator that has control over the delivery rate. This is particularly important when adapting this procedure to the study of the cerebrospinal fluid movement. It is recommended to avoid manual injection and use a slow rate of infusion (e.g., 1 &#956;L/min) to prevent excessive perturbance of the physiological flow. Furthermore, the present protocol is designed to perform a single intracisternal injection, which efficiently labels leptomeningeal cells. We recommend considering ethical specifications, as well as the animal&#39;s ability to withstand multiple surgical procedures, should the study require repeated administration of compounds.
In addition to endoxifen administration, the technique outlined here can be combined with delivery of viruses carrying reporter genes under a leptomeningeal cell-specific promoter. Furthermore, the present delivery system can be utilized for acute tracing of the cerebrospinal fluid flow10. For this purpose, fluorescent tracers such as Cell Tracker (ca. 700 Da) or Dextran Fluorescin (ca. 3000 Da) can be delivered through the cisterna magna, and the syringe may be mounted onto a micromanipulator to enable control over the rate of infusion of the compound. This may be important in order to avoiding excessive disturbance of the natural cerebrospinal fluid movement in tracing experiments.
Leptomeningeal cells express claudin-11 and other proteins associated with tight junctions, which contribute to the establishment of a blood-cerebrospinal fluid barrier in the subarachnoid space and to the homeostatic control of fluid and nutrients circulation3. The approach outlined here may be combined with conditional ablation of genes implicated in the junctional control of the barrier to probe their putative role in maintaining strictly regulated cerebrospinal fluid composition. Additionally, cells from the leptomeninges play a role in development, where they provide extrinsic signals that contribute to the generation of cortical neurons5 and the formation of callosal connections4. Our method can also be adapted to gain further insight into the role of the leptomeninges in correct cortical development and axonal pathfinding. Finally, bacteria such as Neisseria meningitidis have been shown to attach to human leptomeningeal cells14, and animal models for the disease have been developed to study bacterial invasion and resulting neurological damage15,16, although surface ligands responsible for the infection are yet to be fully determined. Selective recombination of leptomeningeal cells achieved with our technique could aid identification of the adhesion sites used by bacteria to infect the subarachnoid space. Of note, the protocol hereby presented may require modifications to account for additional ethical and safety requirements necessary to carry out procedures that entail bacterial infections.
In conclusion, the intracisternal injection herein described represents a simple and well-tolerated surgical approach that offers the opportunity to investigate a wide range of leptomeningeal and cerebrospinal fluid functions, when combined with gene-editing approaches or infusion of labelled molecules.

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 approximately 120&#176;, thus, allowing the back of its neck to be stretched (Figure 1A). The atlanto-occipital portion of dural membrane is then exposed through blunt dissection of the neck muscles, thus gaining access to the underlying cisterna magna. A safe manual injection is performed with a needle bent to approximately 30&#176; at 3 mm from the tip. This allows the syringe to be held against the occipital bone of the cranium, thus improving stability during administration (Figure 1B). Taking advantage of the physiological movement of the cerebrospinal fluid, the endoxifen solution is distributed throughout the subarachnoid space to efficiently recombine leptomeningeal cells overlaying olfactory bulbs, cortex, and cerebellum (Figure 1C). As demonstrated in Figure 1, the solution does not cross the brain-meningeal barrier and does not come in contact with astroglial cells of the parenchyma, as opposed to systemic administration through oral gavage (2 mg/mL per day, on five consecutive days; Figure 1C-E).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61009/61009fig1.jpg" /> 
Figure 1: Specific labelling of leptomeningeal cells via intracisternal injection of endoxifen. Panel A and B illustrate the procedure developed for intracisternal administration. In order to gain access to the cisterna magna, the back of the neck should be stretched. The anesthetized animal is, therefore, positioned at an approximate angle of 120&#176; between the body and the head, which is tilted downwards (A). The hooked needle allows to secure the syringe to the skull and to safely proceed with a manual administration of the solution (B). Panel C displays a sagittal section of the brain after intracisternal administration of endoxifen in a transgenic mouse model carrying CreER under the Cx30 promoter and inducible expression of tdTomato fluorescent reporter. The asterisk (*) marks the injection site and demonstrates the absence of operative damage following the procedure. Endoxifen selectively induces genetic recombination and reporter gene expression in cells in the meningeal layer overlaying the olfactory bulb (Ob), cortex (Ctx), and cerebellum (Cb). Only a few astrocytes in the midbrain (arrowhead) become recombined after intracisternal delivery of endoxifen. Panels D-F are magnifications of the boxed area in C in animals that were treated with vehicle solution (D), subjected to intracisternal injection of endoxifen (E), or treated systemically through oral gavage (F). Whereas intracisternal administration specifically labels cells of the leptomeninges, systemic delivery also leads to recombination of Cx30-expressing astrocytes throughout the cortical layers (L1 to L6). Panel G illustrates recombination of leptomeningeal cells (asterisk), identified through Pdgfra reactivity, after intracisternal injection. By contrast, surface (arrowhead) and parenchymal (arrow) astrocytes expressing Gfap remain unlabeled. Scale bars =&#160;1,000 &#956;m&#160;(C),&#160;150 &#956;m (D-F), and&#160;40 &#956;m (G). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61009/61009fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Neuroscience

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

Intracranial Injection of Adeno-associated Viral Vectors

Intracranial injection of viral vectors engineered to express a fluorescent protein is a versatile labeling technique for visualization of specific subsets of cells in different brain regions both in vivo and in brain sections. Unlike the injection of fluorescent dyes, viral labeling offers targeting of individual cell types and is less expensive and time consuming than establishing transgenic mouse lines. In this technique, an adeno-associated viral (AAV) vector is injected intracranially using stereotaxic coordinates, a micropipette and an automated pump for precise delivery of AAV to the desired area with minimal damage to the surrounding tissue. Injection parameters can be tailored to individual experiments by adjusting the animal age at injection, injection location, volume of injection, rate of injection, AAV serotype and the promoter driving gene expression. Depending on the conditions chosen, virally-induced transgene expression can allow visualization of groups of cells, individual cells or fine cellular processes, down to the level of dendritic spines. The experiment shown here depicts an injection of double-stranded AAV expressing green fluorescent protein for the labeling of neurons and glia in the mouse primary visual cortex.

Here we present the intracranial injection of AAV vectors for fluorescent labeling of neurons and glia in the visual cortex.

Intracranial injection of viral vectors engineered to express a fluorescent protein is a versatile labeling technique for visualization of specific ...

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 ...

Efficient Derivation of Human Neuronal Progenitors and Neurons from Pluripotent Human Embryonic Stem Cells with Small Molecule Induction

There is a large unfulfilled need for a clinically-suitable human neuronal cell source for repair or regeneration of the damaged central nervous system (CNS) structure and circuitry in today's healthcare industry. Cell-based therapies hold great promise to restore the lost nerve tissue and function for CNS disorders. However, cell therapies based on CNS-derived neural stem cells have encountered supply restriction and difficulty to use in the clinical setting due to their limited expansion ability in culture and failing plasticity after extensive passaging1-3. Despite some beneficial outcomes, the CNS-derived human neural stem cells (hNSCs) appear to exert their therapeutic effects primarily by their non-neuronal progenies through producing trophic and neuroprotective molecules to rescue the endogenous cells1-3. Alternatively, pluripotent human embryonic stem cells (hESCs) proffer cures for a wide range of neurological disorders by supplying the diversity of human neuronal cell types in the developing CNS for regeneration1,4-7. However, how to channel the wide differentiation potential of pluripotent hESCs efficiently and predictably to a desired phenotype has been a major challenge for both developmental study and clinical translation. Conventional approaches rely on multi-lineage inclination of pluripotent cells through spontaneous germ layer differentiation, resulting in inefficient and uncontrollable lineage-commitment that is often followed by phenotypic heterogeneity and instability, hence, a high risk of tumorigenicity7-10. In addition, undefined foreign/animal biological supplements and/or feeders that have typically been used for the isolation, expansion, and differentiation of hESCs may make direct use of such cell-specialized grafts in patients problematic11-13. To overcome these obstacles, we have resolved the elements of a defined culture system necessary and sufficient for sustaining the epiblast pluripotence of hESCs, serving as a platform for de novo derivation of clinically-suitable hESCs and effectively directing such hESCs uniformly towards clinically-relevant lineages by small molecules14 (please see a schematic in Fig. 1). Retinoic acid (RA) does not induce neuronal differentiation of undifferentiated hESCs maintained on feeders1, 14. And unlike mouse ESCs, treating hESC-differentiated embryoid bodies (EBs) only slightly increases the low yield of neurons1, 14, 15. However, after screening a variety of small molecules and growth factors, we found that such defined conditions rendered retinoic acid (RA) sufficient to induce the specification of neuroectoderm direct from pluripotent hESCs that further progressed to neuroblasts that generated human neuronal progenitors and neurons in the developing CNS with high efficiency (Fig. 2). We defined conditions for induction of neuroblasts direct from pluripotent hESCs without an intervening multi-lineage embryoid body stage, enabling well-controlled efficient derivation of a large supply of human neuronal cells across the spectrum of developmental stages for cell-based therapeutics.

We have established a protocol for induction of neuroblasts direct from pluripotent human embryonic stem cells maintained under defined conditions with small molecules, which enables derivation of a large supply of human neuronal progenitors and neuronal cell types in the developing CNS for neural repair.

There is a large unfulfilled need for a clinically-suitable human neuronal cell source for repair or regeneration of the damaged central nervous system ...

Expansion of Embryonic and Adult Neural Stem Cells by In Utero Electroporation or Viral Stereotaxic Injection

Somatic stem cells can divide to generate additional stem cells (expansion) or more differentiated cell types (differentiation), which is fundamental for tissue formation during embryonic development and tissue homeostasis during adulthood 1. Currently, great efforts are invested towards controlling the switch of somatic stem cells from expansion to differentiation because this is thought to be fundamental for developing novel strategies for regenerative medicine 1,2. However, a major challenge in the study and use of somatic stem cell is that their expansion has been proven very difficult to control.
Here we describe a system that allows the control of neural stem/progenitor cell (altogether referred to as NSC) expansion in the mouse embryonic cortex or the adult hippocampus by manipulating the expression of the cdk4/cyclinD1 complex, a major regulator of the G1 phase of the cell cycle and somatic stem cell differentiation 3,4. Specifically, two different approaches are described by which the cdk4/cyclinD1 complex is overexpressed in NSC in vivo. By the first approach, overexpression of the cell cycle regulators is obtained by injecting plasmids encoding for cdk4/cyclinD1 in the lumen of the mouse telencephalon followed by in utero electroporation to deliver them to NSC of the lateral cortex, thus, triggering episomal expression of the transgenes 5-8. By the second approach, highly concentrated HIV-derived viruses are stereotaxically injected in the dentate gyrus of the adult mouse hippocampus, thus, triggering constitutive expression of the cell cycle regulators after integration of the viral construct in the genome of infected cells 9. Both approaches, whose basic principles were already described by other video protocols 10-14, were here optimized to i) reduce tissue damage, ii) target wide portions of very specific brain regions, iii) obtain high numbers of manipulated cells within each region, and iv) trigger high expression levels of the transgenes within each cell. Transient overexpression of the transgenes using the two approaches is obtained by different means i.e. by natural dilution of the electroporated plasmids due to cell division or tamoxifen administration in Cre-expressing NSC infected with viruses carrying cdk4/cyclinD1 flanked by loxP sites, respectively 9,15.
These methods provide a very powerful platform to acutely and tissue-specifically manipulate the expression of any gene in the mouse brain. In particular, by manipulating the expression of the cdk4/cyclinD1 complex, our system allows the temporal control of NSC expansion and their switch to differentiation, thus, ultimately increasing the number of neurons generated in the mammalian brain. Our approach may be critically important for basic research and using somatic stem cells for therapy of the mammalian central nervous system while providing a better understanding of i) stem cell contribution to tissue formation during development, ii) tissue homeostasis during adulthood, iii) the role of adult neurogenesis in cognitive functions, and perhaps, iv) better using somatic stem cells in models of neurodegenerative diseases.

Expansion of Embryonic and Adult Neural Stem Cells by In Utero Electroporation or Viral Stereotaxic Injection

Controlling the expansion of somatic stem cells is a major factor hampering their study and use in therapy. Here we describe a system to temporally control neural stem cells expansion during development and adulthood, which can be used to increase the number of neurons generated in the mouse brain.

Somatic stem cells can divide to generate additional stem cells (expansion) or more differentiated cell types (differentiation), which is fundamental for ...

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 ...

Induction of Complete Transection-Type Spinal Cord Injury in Mice

Spinal cord injury (SCI) largely leads to irreversible and permanent loss of function, most commonly as a result of trauma. Several treatment options, such as cell transplantation methods, are being researched to overcome the debilitating disabilities arising from SCI. Most pre-clinical animal trials are conducted in rodent models of SCI. While rat models of SCI have been widely used, mouse models have received less attention, even though mouse models can have significant advantages over rat models. The small size of mice equates to lower animal maintenance costs than for rats, and the availability of numerous transgenic mouse models is advantageous for many types of studies. Inducing repeatable and precise injury in the animals is the primary challenge for SCI research, which in small rodents requires high-precision surgery. The transection-type injury model has been a commonly used injury model over the last decade for transplantation-based therapeutic research, however a standardized method for inducing a complete transection-type injury in mice does not exist. We have developed a surgical protocol for inducing a complete transection type injury in C57BL/6 mice at thoracic vertebral level 10 (T10). The procedure uses a small tip drill instead of rongeurs to precisely remove the lamina, after which a thin blade with rounded cutting edge is used to induce the spinal cord transection. This method leads to reproducible transection-type injury in small rodents with minimal collateral muscle and bone damage and therefore minimizes confounding factors, specifically where behavioral functional outcomes are analyzed.

This protocol describes how to create a precise laminectomy for induction of stable transection-type spinal cord injury in the mouse model, with minimal collateral damage for spinal cord injury research.

Spinal cord injury (SCI) largely leads to irreversible and permanent loss of function, most commonly as a result of trauma. Several treatment options, ...

A Scalable Model to Study the Effects of Blunt-Force Injury in Adult Zebrafish

Blunt-force traumatic brain injuries (TBI) are the most common form of head trauma, which spans a range of severities and results in complex and heterogenous secondary effects. While there is no mechanism to replace or regenerate the lost neurons following a TBI in humans, zebrafish possess the ability to regenerate neurons throughout their body, including the brain. To examine the breadth of pathologies exhibited in zebrafish following a blunt-force TBI and to study the mechanisms underlying the subsequent neuronal regenerative response, we modified the commonly used rodent Marmarou weight drop for the use in adult zebrafish. Our simple blunt-force TBI model is scalable, inducing a mild, moderate, or severe TBI, and recapitulates many of the phenotypes observed following human TBI, such as contact- and post-traumatic seizures, edema, subdural and intracerebral hematomas, and cognitive impairments, each displayed in an injury severity-dependent manner. TBI sequelae, which begin to appear within minutes of the injury, subside and return to near undamaged control levels within 7 days post-injury. The regenerative process begins as early as 48 hours post-injury (hpi), with the peak cell proliferation observed by 60 hpi. Thus, our zebrafish blunt-force TBI model produces characteristic primary and secondary injury TBI pathologies similar to human TBI, which allows for investigating disease onset and progression, along with the mechanisms of neuronal regeneration that is unique to zebrafish.

We modified the Marmarou weight drop model for adult zebrafish to examine a breadth of pathologies following blunt-force traumatic brain injury (TBI) and the mechanisms underlying subsequent neuronal regeneration. This blunt-force TBI model is scalable, induces a mild, moderate, or severe TBI, and recapitulates injury heterogeneity observed in human TBI.