intracranial cannular implantation in a mouse to deliver therapeutic agents to brain tumors

Intracranial Cannular Implantation in a Mouse to Deliver Therapeutic Agents to Brain Tumors

To begin, place the anesthetized mouse with a shaved head on the operation table, and gently open the bottom of the stereotactic arm using a spatula. Insert the cannula with forceps and secure it by tightening the screw on the arm until 1/2 to 2/3 of the white plastic portion and 5 millimeters of the cannula protrude from the bottom of the opening.
Insert and secure the mouse&#39;s top teeth in the bite bar of the stereotaxic apparatus. Pull the nose cone forward and tighten it, ensuring the mouse inhales isoflurane. Then mount the mouse on the warmed to stereotaxic apparatus using ear cuffs or ear bars, avoiding excessive pressure.
Disinfect the surgical site, and make a cut parallel to the skull, as described in the manuscript. Use cotton-tipped swabs to push the fascia away. Identify the landmarks bregma and lambda on the skull corresponding to anterior and posterior marks where the cranial plates meet.
To create a surface for attaching the acrylic, gently make several slits across the skull using a scalpel. Using the stereotactic arm, localize the cannula to the landmark of interest. Raise the cannula tip 1 to 2 millimeters above the skull surface and move to the desired coordinates. Using an 18-gauge needle or surgical drill, make two screw holes on the exposed skull away from where the cannula enters, ensuring enough space for the cannula.
Twist a drill through the screw hole until it attaches to the skull. With a flat-tipped screwdriver, insert and fasten two screws into the holes. Then, gently pull the screw up to ensure it is secured.
For inserting a cannula, create a hole in the skull at the identified coordinates using an 18-gauge needle or surgical drill. Using the stereotactic arm, lower the cannula to the desired DV coordinate. Add approximately 0.3 grams of acrylic resin powder and 10 to 15 drops of liquid in a porcelain 12-well plate.
Load the prepared viscous, white-colored material into a 1-milliliter syringe. Coat the skull and fill the spaces around the cannula and screw. Loosen the screw on the stereotactic arm while the cement is flexible. Gently use a spatula to release the cannula from the holder at the bottom opening, and retract the stereotactic arm away from the mouse.
Once the cement is dried, insert the dummy cannula into the guide cannula and rotate clockwise to secure it tightly. Once the procedure is complete, return the mouse to its warmed home cage for recovery.
To prepare the treatment cannula, insert its top into PKG tubing. Then, fill the treatment syringe with CAR T cell suspension and insert it through the other end of the PKG tube covering the top of the treatment cannula. After anesthetizing the mouse, fix the guide cannula at the base using forceps. Unscrew and remove the dummy cannula, allowing access to the guide cannula.
Infuse the CAR T cells for one minute and hold the treatment cannula in place for an additional minute. After removing the treatment cannula, screw back a dummy cannula tightly. Then, administer Meloxicam subcutaneously for pain control.

intracranial-cannular-implantation-mouse-to-deliver-therapeutic

Nanyang Technological University

Research

JoVE Encyclopedia of Experiments

Immunology

Immunotherapy

Model Development and Disease Study

EoE Sub Articles

eoe sub articles

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Preparing the mouse for surgery
<ol>
	<li>Anesthetize the mouse in an induction chamber with isoflurane (2-4%) at an oxygen flow rate of 1 L/min until an adequate plane of anesthesia is reached (approximately 5 min).</li>
	<li>Weigh the mouse using a scale to the nearest 0.1 g and administer subcutaneous slow-release (SR) buprenorphine (1 mg/kg) or other analgesic. 
	NOTE: SR buprenorphine provides analgesia for 72 h.</li>
	<li>Shave the fur on the top of the mouse&#39;s head using electric clippers or depilatory agents.</li>
	<li>Use a spatula to gently open the bottom of the stereotactic arm and insert the guide cannula using forceps. Tighten the screw on the arm to secure the cannula so that approximately 1/2 to 2/3 of the white plastic portion of the cannula is protruding from the bottom of the opening, along with the entire 5 mm metal length of the cannula.</li>
	<li>Insert and secure the mouse&#39;s top teeth in the bite bar of the stereotaxic apparatus. Pull the nose cone forward and tighten it, ensuring the mouse is inhaling isoflurane.</li>
	<li>Mount the mouse on the warmed stereotaxic apparatus using ear cuffs or ear bars, avoiding excessive pressure. 
	NOTE: Warmed tray should have a rectal thermometer inserted, and the warming tray should adjust to maintain a normal body temperature of the mouse during the procedure.</li>
	<li>Apply sterile ophthalmic ointment to both eyes using a cotton-tipped applicator.</li>
	<li>Wipe the surgical site with povidone-iodine on a pad or applicator, followed by an alcohol pad. Perform this step three times in total.</li>
	<li>Before beginning the procedure, perform a toe pinch with forceps to assess for adequate sedation.</li>
</ol>
2. Surgical procedure
NOTE: All aspects of the surgical procedure utilize sterilized instruments and aseptic techniques. The mice continue under anesthesia with isoflurane (2-4%) throughout the duration of the procedure, approximately 10-20 min.
<ol>
	<li>Gently pick up the scalp between the ears with forceps. Using sterile scissors, cut the lifted scalp parallel to the skull and remove an oval flap of skin (0.75-1 cm in length) to expose the skull. 
	NOTE: Scissors are preferred over a scalpel to provide a clean, oval-shaped opening and to prevent unnecessary damage to surrounding skin and tissue.</li>
	<li>Push away fascia using a scalpel or cotton-tipped swabs and a hemostatic cotton pellet to help slow excessive bleeding as needed. 
	NOTE: Using the wooden side of a sterile cotton tip can also push away fascia and help avoid excessive bleeding.</li>
	<li>Identify the landmarks bregma and lambda, the respective anterior and posterior marks on the skull where the cranial plates meet. 
	NOTE: Identification can be augmented by wiping the top of the exposed skull with hydrogen peroxide.</li>
	<li>Gently score the skull using a scalpel to create a surface for the acrylic to attach. Scoring should include multiple linear lines approximately 0.5-1 cm in length at 90&#176; angles to each other.</li>
	<li>Using the stereotaxic arm, localize the cannula to the landmark of interest (bregma or lambda). Once localized, raise the cannula tip 1-2 mm above the skull surface and move to the desired coordinates. For intratumoral injections, this uses the same A/P and M/L coordinates as the tumor placement.</li>
	<li>On the exposed skull, away from the area where the cannula will enter, make two screw holes with an 18 G needle or a surgical drill. Ensure that the holes are spaced out to include enough room for the cannula. Using a drill bit, twist through the screw holes until they catch on the skull. Insert and fasten two screws into the holes using a flat-tipped screwdriver. Then, gently pull the screws up to ensure they are secured. 
	NOTE: Do not insert the screws until they are flush with the skull, or they may damage the mouse brain underneath. Leave at least a 1-2 mm gap between the screw and the skull.</li>
	<li>Using an 18 G needle or surgical drill, drill through the skull at the identified coordinates to create a hole for the cannula to be inserted.</li>
	<li>Using the stereotactic arm, lower the cannula to the desired D/V coordinate. 
	NOTE: The D/V coordinate of cannula implantation needs to account for the projection length of the dummy and treatment cannulas, and may be more superficial than the orthotopic injection of tumor cells (Figure 1).</li>
	<li>In a porcelain 12-well plate, fill one well with acrylic resin powder (approximately 0.3 g) and 10-15 drops (approximately 0.5-0.75 mL) of acrylic resin liquid. This produces a viscous white-colored material. Draw up the mixture into a 1 mL syringe and use it to coat and cover the skull, filling in the spaces around the cannula and screws. 
	NOTE: The viscous material hardens into cement over time, so this step should be completed promptly after mixing.</li>
	<li>While the cement is still pliable, loosen the screw on the stereotactic arm and use a spatula in the opening at the bottom to gently release the cannula from the holder and slowly retract the stereotactic arm upward away from the mouse.</li>
	<li>Once the cement is completely dry, insert the dummy cannula into the guide cannula and tightly secure it by turning clockwise.</li>
	<li>Once the procedure is complete, place the mouse back in its warmed home cage to carefully monitor, ensuring adequate recovery and recording any post-procedure observations, including that the mouse has fully regained consciousness, before returning to the colony. 
	NOTE: It is generally recommended to place only half of the cage on a heating pad to allow for the animal to move to the cooler side to avoid overheating.</li>
	<li>Administer additional analgesics as needed if the mice display behaviors indicative of pain post-operatively, such as meloxicam 5 mg/kg subcutaneously delivered once daily for up to 3 days.</li>
</ol>
3. Preparing chimeric antigen receptor (CAR) T cells
<ol>
	<li>Measure the pre-transfected CAR T cell concentration using a cell counter.</li>
	<li>Centrifuge pre-transfected T cells at 200 x g for 5 min at room temperature (RT).</li>
	<li>Aspirate the supernatant using a sterile Pasteur pipet on a vacuum aspiration system and resuspend the pellet in phosphatae buffered saline (PBS) to a desired concentration. Typical delivery volumes are 2-5 &#181;L. Typical cell doses are 0.5-5 x 106 cells.</li>
</ol>
4. CAR T cell infusions
<ol>
	<li>Prepare the treatment cannula by feeding the top through a small piece of PKG tubing.</li>
	<li>Fill the treatment syringe with the CAR T cell suspension and insert it through the other end of the PKG tubing, enough to cover the top of the treatment cannula.</li>
	<li>Anesthetize the mouse with isoflurane (2-4%) at an oxygen flow rate of 1 L/min.</li>
	<li>Stabilize the guide cannula using forceps at the base, and then carefully unscrew and remove the dummy cannula, allowing for access into the guide cannula. 
	NOTE: A stereotaxic setup is not necessary, but can be used to stabilize the head for treatment.</li>
	<li>Infuse the CAR T cells for 1 min and hold the treatment cannula in place for an additional 1 min following the cessation of infusion.</li>
	<li>Remove the treatment cannula and screw the dummy cannula tightly back in place.</li>
	<li>Administer subcutaneous meloxicam (5 mg/kg) for optional pain control.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>18 G needles</td>
			<td>BD</td>
			<td>511097</td>
			<td>1 1/2 inch metal hub</td>
		</tr>
		<tr>
			<td>Acrylic resin liquid</td>
			<td>Lang Dental</td>
			<td>B1323</td>
			<td></td>
		</tr>
		<tr>
			<td>Acrylic resin powder</td>
			<td>Lang Dental</td>
			<td>B1323</td>
			<td></td>
		</tr>
		<tr>
			<td>Alcohol wipes</td>
			<td>BD</td>
			<td>326895</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge 5240</td>
			<td>Eppendorf</td>
			<td>5420000040</td>
			<td>Centrifuge</td>
		</tr>
		<tr>
			<td>Cotton tipped swabs</td>
			<td>Puritan</td>
			<td>826-WC</td>
			<td>Handle Width = 2.11 mm (0.083), Head Width = 1.27 mm (0.050), Handle Length = 147.62 mm (5.812), Overall Length = 152.4 mm (6), Head Length = 12.7 mm (0.500)</td>
		</tr>
		<tr>
			<td>Drill bit holder</td>
			<td>P1 Technologies</td>
			<td>DH-1</td>
			<td>Drill bit holder for D56-D70</td>
		</tr>
		<tr>
			<td>Drill bit</td>
			<td>P1 Technologies</td>
			<td>D58</td>
			<td>1.07 mm</td>
		</tr>
		<tr>
			<td>Dummy cannula</td>
			<td>P1 Technologies</td>
			<td>C315DCS-5/SPC</td>
			<td>Configuration: Small cap; &#160;Length: Cut 5.00 mm below pedestal; Projection: 0.50 mm</td>
		</tr>
		<tr>
			<td>Flat tip screwdriver</td>
			<td>P1 Technologies</td>
			<td>SD-80</td>
			<td>Screwdriver</td>
		</tr>
		<tr>
			<td>Graefe forceps</td>
			<td>Fine Science Tools</td>
			<td>11051-10</td>
			<td>Forceps</td>
		</tr>
		<tr>
			<td>Guide cannula</td>
			<td>P1 Technologies</td>
			<td>C315GS-5/SPC</td>
			<td>Configuration: 5.00 mm pedestal height; Length: Cut 5.00 mm below pedestal</td>
		</tr>
		<tr>
			<td>Hemostatic cotton pellets with racemic epinephrine</td>
			<td>Pascal</td>
			<td>1151602</td>
			<td></td>
		</tr>
		<tr>
			<td>MOXI Z Mini automated cell counter Kit</td>
			<td>Moxi</td>
			<td>MXZ001</td>
			<td>Cell counter</td>
		</tr>
		<tr>
			<td>NOD scid gamma (NSG) mice</td>
			<td>Jackson Laboratory</td>
			<td>5557</td>
			<td>6 to 12-week-old males and females</td>
		</tr>
		<tr>
			<td>Pasteur pipet</td>
			<td>VWR</td>
			<td>14673-043</td>
			<td></td>
		</tr>
		<tr>
			<td>PKG tubing</td>
			<td>P1 Technologies</td>
			<td>C313CT</td>
			<td>Diameter: 0.58 mm x 1.27 mm</td>
		</tr>
		<tr>
			<td>Porcelain 12 well plate</td>
			<td>Flinn Scientific</td>
			<td>AP6064</td>
			<td></td>
		</tr>
		<tr>
			<td>Povidone iodine</td>
			<td>Medline</td>
			<td>MDS093943</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>World Precision Instrument</td>
			<td>50-822-457</td>
			<td>Disposable Scalpel, no.10, sterile, 10/box, Plastic Handle with 6&#34; Ruler</td>
		</tr>
		<tr>
			<td>Screws</td>
			<td>P1 Technologies</td>
			<td>0-80 X 3/32</td>
			<td>2.4 mm</td>
		</tr>
		<tr>
			<td>Stereotaxic Frame</td>
			<td>David Kopf Instruments</td>
			<td>940</td>
			<td>Model 940 Small Animal Stereotaxic Instrument with Digital Display Console</td>
		</tr>
		<tr>
			<td>Student fine scissors</td>
			<td>Fine Science Tools</td>
			<td>91460-12</td>
			<td>Scissors</td>
		</tr>
		<tr>
			<td>Treatment cannula</td>
			<td>P1 Technologies</td>
			<td>C315IS-5/SPC</td>
			<td>33GA; Configuration: Standard internal; Length: Cut 5.00 mm below pedestal; Projection: 0.50 mm</td>
		</tr>
		<tr>
			<td>Treatment syringes</td>
			<td>Hamilton</td>
			<td>87908</td>
			<td>5 &#181;L, Model 75 Cemented Needle Special (SN) Syringe, 75SN/22/0.5&#34;/PT3</td>
		</tr>
		<tr>
			<td>Vactrap XL</td>
			<td>Foxx Life Sciences</td>
			<td>305-4401-FLS</td>
			<td>Vacuum System</td>
		</tr>
	</tbody>
</table>

Source: Harvey, K. et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/64886">Intracranial Cannula Implantation for Serial Locoregional Chimeric Antigen Receptor (CAR) T Cell Infusions in Mice.</a> J. Vis. Exp. (2023)
This video presents a procedure for intracranial cannula implantation in a brain tumor mouse model, to facilitate the administration of an immunotherapeutic agent. The delivery of glypican-2-specific chimeric antigen receptor T cells in the tumor microenvironment triggers T cell activation, resulting in tumor lysis and tumor regression.

<img alt="Figure 1" src="/files/ftp_upload/22232/22232_Figure1.jpg" /> 
Figure 1: Guide cannula with projection dummy and treatment cannulas. (A) Guide cannula with 0.5 mm projection and 2 mm projection dummy cannulas in place. (B) Guide cannula with 0.5 mm projection and 2 mm projection treatment cannulas in place.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Begin with an anesthetized mouse with a brain tumor, secured on a stereotaxic apparatus, carrying a guide cannula.
Disinfect and then dissect the scalp. Remove the subcutaneous tissue to expose the skull.
Scratch the skull surface to make it uneven. Anchor the screws to support the cannula.
Drill a hole into the skull at a precise location to reach the tumor.
Descend the guide cannula through the hole to insert it into the tumor.
Apply an acrylic resin. The uneven skull surface provides a grip for the adhesive to secure the guide cannula.
Allow the mouse to recover.
Re-anesthetize and secure the mouse.
Take a treatment cannula attached to a syringe containing a therapeutic agent.
Position the treatment cannula over the guide cannula.
Inject the therapeutic agent and allow it to flow into the tumor.

183 vues. Implantation d’un canule intracrânienne chez une souris pour administrer des agents thérapeutiques aux tumeurs cérébrales

Vidéo: Implantation d’un canule intracrânienne chez une souris pour administrer des agents thérapeutiques aux tumeurs cérébrales - Expérience

Cannula Implantation into the Cisterna Magna of Rodents

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

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

Implantation de la canule dans la Magna de Cisterna de rongeurs

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

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Neurosciences

Intracranial Pharmacotherapy and Pain Assays in Rodents

Pain is a salient sensory experience with affective and cognitive dimensions. However, central mechanisms for pain remain poorly understood, hindering the development of effective therapeutics. Intracranial pharmacology presents an important tool for understanding the molecular and cellular mechanisms of pain in the brain, as well as for novel treatments. Here we present a protocol that integrates intracranial pharmacology with pain behavior testing. Specifically, we show how to infuse analgesic drugs into a select brain region, which may be responsible for pain modulation. Furthermore, to determine the effect of the candidate drug in the central nerve system, pain assays are performed after intracranial treatment. Our results demonstrate that intracranial administration of analgesic drugs in a targeted region can provide relief of pain in rodents. Thus, our protocol successfully demonstrates that intracranial pharmacology, combined with pain behavior testing, can be a powerful tool for the study of pain mechanisms in the brain.

Here we present a protocol to perform intracranial pharmacological experiments followed by pain behavior assays in rodents. This protocol allows researchers to deliver molecular and cellular targets in the brain, for pharmacologic agents in the treatment of pain.

Pharmacothérapie intracrânienne et dosages de la douleur chez les rongeurs

Pain is a salient sensory experience with affective and cognitive dimensions. However, central mechanisms for pain remain poorly understood, hindering the ...

Direct Cannula Implantation in the Cisterna Magna of Pigs

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

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

Implantation directe de canules dans la Cisterna Magna des porcs

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

Intracranial Cannular Implantation in a Mouse to Deliver Therapeutic Agents to Brain Tumors

Transcription

Intracranial Cannular Implantation in a Mouse to Deliver Therapeutic Agents to Brain Tumors