demyelination and remyelination of mouse spinal cord neurons

Demyelination and Remyelination of Mouse Spinal Cord Neurons

Move the animal to a stereotactic frame, dorsal side up, elevated at the midsection with folded paper towels, to exaggerate the curvature of the spine. Then, secure the head with a tooth clamp, and fasten the arms and tail with surgical tape. Stabilizing ear bars are not necessary.
Now, use a scalpel to make a 3 centimeter midline incision, starting just below the ears and cutting in the caudal direction. Then, locate the divide between the two large adipose structures and use fine forceps in each hand to pull these apart. Spread the retractors to open the surgical field.
Under a surgical microscope, locate the prominent bony outgrowth of the T2 vertebra, characteristic of the C57 black 6 mouse strain. To view the vertebra, make a blunt dissection of the overlaying musculature with closed spring scissors. Then, feel for the hard surfaces of T3 and T4 to confirm proper anatomical location.
Now, make shallow lateral cuts in the connective tissue between T3 and T4. Be mindful that too deep a cut will pierce and damage the cord. Some bleeding is common and can be cleared with a sponge spear. The spinal cord is covered with a thick layer of dura, and will be visible if this meningeal layer was not yet cut.
If the dura remains intact, carefully remove it using gentle lateral scrapes with a 32-gauge metal needle. Avoid piercing the remaining underlying meninges, which are not as thick and are harder to see. If any cerebrospinal fluid leaks from the arachnoid, remove it with a sponge spear. Begin the injection by moving the capillary into position over the spinal cord.
The prominent caudal rostral blood vessel should not be used as a midline landmark. Instead, use the gray white matter boundaries flanking the dorsal column to estimate the midline. Now, slowly lower the capillary until the tip just barely touches the spinal cord, immediately lateral of either side of the midline. Lock the arm in place at this position.
Then, make note of the baseline position on the graded measurements of the z direction on the stereotactic arm. From this reading, subtract 1.3 millimeters, and use a quick and shallow downward motion to pierce the tissue, carefully lowering the capillary to the desired depth.
Now, using the micromanipulator, make one full rotation every 5 seconds for 2 minutes to inject a total volume of 0.5 microliters. Then, leave the capillary in place for 2 minutes. After 2 minutes, carefully retract the capillary from the tissue. This allows the solution to seep in. It could otherwise backflow.
Move the injecting apparatus away from the animal and remove the retractors. Close up the animal, by tying a single suture in the muscle adipose tissue overlaying the spinal column. Then, use a non-interrupted suture to close the skin, and apply more iodine to the incision site. Finish by placing the animal in a heated recovery chamber until it recovers. Then, return it to its cage. Post-operative care and analysis are described in the text protocol.

demyelination-and-remyelination-of-mouse-spinal-cord-neurons

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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Prepare Syringe for Injection
<ol>
	<li>Dissolve lysolecithin in a 1% solution in phosphate-buffered saline (PBS; pH 7.4) and store at -20 &#176;C in small aliquots (75 &#956;l). Thaw a vial to room temperature (RT). 
	NOTE: If lysolecithin is undissolved, sonicate the tube in an ultrasonic cleaner (40 kHz) for approximately 30 min to form a uniform solution.</li>
	<li>Handle the pre-pulled glass capillary with extreme care to avoid damaging the delicate tip. Unscrew the nut of a 10 &#956;l injecting syringe and thread it onto the flat end of the capillary, followed by two ferrules, ensuring that the mating ends of the ferrules align and the capillary is snug in the conical ferrule (Figure 1). Rinse the syringe with isopropyl alcohol and remove the plunger. Once dry, screw the needle assembly hand tight onto the injecting syringe.</li>
	<li>Attach the metal hub needle to the priming syringe. Pierce a rubber disc with the needle and slide it down to the base. Fill the priming syringe with lysolecithin. Gently depress the priming syringe until liquid is visible at the tip of the needle. This ensures air bubbles will not be introduced into the injecting syringe.</li>
	<li>Insert the metal hub needle of the priming syringe into the injecting syringe. Making a firm seal with the rubber disc, slowly depress the priming syringe until the capillary fills the tip with solution. Carefully remove the priming syringe while simultaneously depressing to fill the barrel of the injecting syringe without introducing air bubbles. Insert the plunger into the injecting syringe and ensure the solution flows from the tip of the capillary as the plunger is gently depressed. 
	NOTE: If any air bubbles are visible in the capillary or injecting syringe, the syringe preparation must be repeated from the beginning. Continuous fluid in the injecting apparatus is critical to ensure accurate injection volumes.</li>
	<li>Attach the completed injecting syringe to the arm of a stereotactic micromanipulator. This completed apparatus will be able to inject 15-20 animals before needing to be refilled.</li>
	<li>Discard the remaining lysolecithin from the priming syringe. Withdraw and depress isopropyl alcohol several times, and detach the metal hub needle. Wait several hours for the remaining isopropyl alcohol in the priming syringe to evaporate before filling again.</li>
</ol>
2. Prepare the Animal for Surgical Procedure
NOTE: This procedure is described for female C57BL/6 mice aged 8-10 weeks.
<ol>
	<li>Anesthetize the animal with an intraperitoneal injection of ketamine (200 mg/kg) and xylazine (10 mg/kg) or per institutional animal care regulations. Plan for the animal to be under anesthesia for approximately 1 hr if using injectable anesthesia.</li>
	<li>Test that the animal is deeply anesthetized by firmly pinching the foot. A properly anesthetized animal will not respond to the pinch.</li>
	<li>Using clippers, shave a 2-3 cm2 area on the dorsal side of the animal, close to the ears. Be careful not to damage the ears.</li>
	<li>Wipe the area clean with 70% ethanol applied to a gauze pad. Ensure all clipped hair has been removed from the area. Disinfect the area with iodine.</li>
	<li>Apply petroleum jelly to the eyes to prevent drying throughout the procedure.</li>
	<li>Keep the animal in a heated recovery chamber until ready to begin the procedure.</li>
</ol>
3. Perform the Surgical Procedure
NOTE: Ensure adequate aseptic technique for all steps of the procedure. This includes proper use of gloves, hairnets, masks, and drapes. All tools should be sterilized before coming in contact with the animal.
<ol>
	<li>Move the animal to a stereotactic frame, dorsal side up, elevated at the mid-section by folded paper towels to exaggerate the curvature of the spine. Fasten the arms and tail with surgical tape and secure the head with a tooth clamp. Stabilizing ear bars are not necessary for this procedure.</li>
	<li>Use a scalpel to make a 3 cm midline incision, starting just below the ears and cutting in the caudal direction.</li>
	<li>Locate the divide between the two large adipose structures and use fine forceps in each hand to pull these apart. Spread retractors to open the surgical field.</li>
	<li>Under a surgical microscope, locate the prominent outgrowth process of the thoracic vertebra 2 (T2) vertebra (NOTE: This feature is characteristic of the C57BL/6 mouse strain). Perform a blunt dissection with closed spring scissors through the overlaying musculature to better visualize T2. Using the forceps, feel for the hard surfaces of thoracic vertebra 3 (T3) and thoracic vertebra 4 (T4) to confirm proper anatomical location.</li>
	<li>Using spring scissors, make shallow lateral cuts (2-3 mm deep) of the connective tissue between T3 and T4. Due to the natural spacing between vertebrae in the upper thoracic portion of the mouse vertebral column, a laminectomy is unnecessary to reveal the spinal cord. Be mindful that a cut that is too deep will pierce and damage the cord. 
	NOTE: A small degree of bleeding is common during this step. If this occurs, hold a sponge spear into the area until the bleeding subsides (30-60 sec).</li>
	<li>Visualize the spinal cord. It will be covered with a thick layer of visible dura if this meningeal layer is not yet cut while exposing the cord. A prominent blood vessel runs caudal/rostral through the approximate midline of the spinal cord. 
	NOTE: This vasculature should not be used as a landmark for the midline. Instead, adequate lighting should reveal the grey-white matter boundaries flanking the dorsal column, which should be used to estimate the midline.</li>
	<li>If the dura remains intact, make gentle lateral scrapes with a 32 G metal needle until cleared. The goal is to remove the dura while not cutting the remaining underlying meninges, which are not as thick and harder to see. 
	NOTE: The release of cerebrospinal fluid indicates a breach of the arachnoid, and while this can occur without mechanical damage to the tissue, the accumulated cerebrospinal fluid should be removed with a sponge spear to visualize the surface of the cord better.</li>
	<li>Move the injecting syringe in place and slowly lower it until the tip of the capillary just barely touches the spinal cord immediately lateral of either side of the midline. Lock the arm in place.</li>
	<li>Use the graded measurements of the Z-direction stereotactic arm to make a baseline position measurement. From this reading, subtract 1.3 mm. Use a quick and shallow downward motion to pierce the tissue and carefully lower the capillary until the new measurement is reached. Optional: if desired, lesions in the dorsal column can be produced by the same piercing motion at the midline and a depth of 0.3 mm. 
	NOTE: These values are specific for injecting between T3 and T4. If opting to perform the injection at any other location in the spinal cord, these values must be derived from any available mouse brain atlas.</li>
	<li>Use the micromanipulator to depress lysolecithin into the ventral spinal cord white matter. Make one rotation of the micromanipulator every 5 sec for 2 min, resulting in a final volume of 0.5 &#956;l. Leave the capillary in place for an additional 2 min to prevent backflow of solution, and then carefully remove the capillary.</li>
	<li>Tie a single suture in the muscle/adipose tissue overlaying the spinal column. Use a non-interrupted suture to close the skin. Apply more iodine to the incision site.</li>
	<li>Place the animal in a heated recovery chamber until it recovers, then return it to its cage. Apply analgesics post-operatively as per institutional animal care regulations. Additional post-operative care is usually not necessary as the animals are fully ambulatory and capable of self-feeding and drinking as soon as they recover from anesthesia.</li>
	<li>Repeat the procedure for the remaining animals. 
	NOTE: With proficiency, the operation can be completed in 10-15 min per animal, particularly with the help of a second person tying sutures. The same glass capillary can be used for approximately 15-20 surgeries before the tip becomes blunt and should be replaced. Control surgeries can be performed identically as described, with an injection of PBS into the spinal cord instead of lysolecithin. We do not recommend cleaning the capillaries for future use.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Spring scissors</td>
			<td>Fine Science Tools</td>
			<td>15004-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Fine Science Tools</td>
			<td>11254-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Retractor</td>
			<td>Fine Science Tools</td>
			<td>17003-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Clippers</td>
			<td>Philips</td>
			<td>QG3330</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating recovery chamber</td>
			<td>Peco Services</td>
			<td>V1200</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical tape</td>
			<td>3M</td>
			<td>1527-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel handle</td>
			<td>Fine Science Tools</td>
			<td>10003-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel blade</td>
			<td>Feather</td>
			<td>No. 15</td>
			<td></td>
		</tr>
		<tr>
			<td>Sponge spear</td>
			<td>Beaver Visitec</td>
			<td>581089</td>
			<td></td>
		</tr>
		<tr>
			<td>5-0 Vicryl sutures</td>
			<td>Ethicon</td>
			<td>J511G</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved ToughCut spring scissors</td>
			<td>Fine Science Tools</td>
			<td>15123-12</td>
			<td></td>
		</tr>
		<tr>
			<td>32 gauge metal needle</td>
			<td>BD</td>
			<td>305106</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle holders</td>
			<td>Fine Science Tools</td>
			<td>12002-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Angled forceps</td>
			<td>Fine Science Tools</td>
			<td>11251-35</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton tipped applicator</td>
			<td>Puritan</td>
			<td>806-WC</td>
			<td></td>
		</tr>
		<tr>
			<td>Gauze pads</td>
			<td>Safe Cross First Aid</td>
			<td>3763</td>
			<td></td>
		</tr>
		<tr>
			<td>10 &#956;L syringe</td>
			<td>Hamilton</td>
			<td>7635-01</td>
			<td>Make sure to purchase the microliter, not gastight syringe</td>
		</tr>
		<tr>
			<td>Compression fitting</td>
			<td>Hamilton</td>
			<td>55750-01</td>
			<td>Contains the 2 ferrules and removable nut, but the nut that comes with the 10 &#956;L syringe is a tighter fit</td>
		</tr>
		<tr>
			<td>Priming kit</td>
			<td>Hamilton</td>
			<td>PRMKIT</td>
			<td>Contains the priming syringe, removable hub needle and the rubber discs</td>
		</tr>
		<tr>
			<td>Pre-pulled glass capillaries</td>
			<td>WPI</td>
			<td>TIP10TW1 (pack of 10)</td>
			<td>Contains capillaries with 10 &#956;m inner diameter. 30 &#956;m inner diameter also work (TIP30TW1)</td>
		</tr>
		<tr>
			<td>Stereotactic frame</td>
			<td>David Kopf instruments</td>
			<td>Model 900</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasonic cleaner</td>
			<td>Fisher Scientific</td>
			<td>FS-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope slides</td>
			<td>VWR</td>
			<td>48311-703</td>
			<td></td>
		</tr>
		<tr>
			<td>Bright field microscope</td>
			<td>Olympus</td>
			<td>BX51</td>
			<td></td>
		</tr>
		<tr>
			<td>Lysophosphatidylchoine</td>
			<td>Sigma</td>
			<td>L1381</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Oxoid</td>
			<td>BR0014</td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropyl alcohol</td>
			<td>Sigma</td>
			<td>109827</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>CDMV</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>CDMV</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Iodine</td>
			<td>West Penetone</td>
			<td>2021</td>
			<td></td>
		</tr>
		<tr>
			<td>Vaseline petroleum jelly</td>
			<td>VWR</td>
			<td>CA05971</td>
			<td></td>
		</tr>
	</tbody>
</table>

Source: Keough, M. B., et al. <a href="https://app.jove.com/v/52679">Experimental Demyelination and Remyelination of Murine Spinal Cord by Focal Injection of Lysolecithin.</a> J. Vis. Exp. (2015)
This video demonstrates the method for inducing demyelination in mouse spinal cord neurons using lysolecithin injections, followed by remyelination through the migration and differentiation of oligodendrocyte precursor cells. The process impairs and then gradually restores neuronal signal transmission.

<img alt="Figure 1" src="/files/ftp_upload/23116/23116_Figure1.jpg" /> 
Figure 1. Assembly of the injecting syringe. (A) The nut of the injecting syringe is threaded onto the flat end of the glass capillary, followed by the two ferrules such that their mating ends interlock. Once the capillary is firmly snug in the conical ferrule, the assembly is screwed hand-tight onto the end of the injecting syringe. (B) Piece the center of a rubber disc with the metal hub needle attached to the priming syringe and slide it down to the base. (C) Withdraw lysolecithin solution into the priming syringe. (D) Gently depress the priming syringe until the first drop of lysolecithin is visible at the tip of the needle. (E) Insert the priming syringe into the barrel of the injecting syringe, making a firm seal with the rubber disc. Gently depress the solution until it runs to the end of the capillary. Carefully withdraw the priming syringe while maintaining pressure on the plunger to remove the metal hub needle without introducing air bubbles into the injecting syringe.

Source: Keough, M. B., et al. Experimental Demyelination and Remyelination of Murine Spinal Cord by Focal Injection of Lysolecithin. J. Vis. Exp. (2015)
...

Begin with an anesthetized mouse positioned on a stereotactic frame, with an incision exposing a region of the spinal cord.
Take the syringe containing lysolecithin, a detergent. Position it over the exposed area and touch one side of the midline.
Inject lysolecithin into the spinal cord&#39;s white matter, targeting the neurons with myelinated axons, which facilitate faster signal transmission.
Keep the syringe in place briefly to prevent backflow, then remove it.
Suture the incision, apply an antiseptic to prevent infection, and allow the mouse to recover.
Injected lysolecithin disrupts the myelin sheath around the neurons, impairing axonal signal transmission.
This process also triggers inflammation, recruiting immune cells to clear the myelin debris, causing localized demyelination of axons, a neuropathological condition.
Over time, oligodendrocyte precursor cells migrate to this site and differentiate into oligodendrocytes.
Later, these oligodendrocytes regenerate the myelin sheath around the demyelinated neurons and gradually restore the signal transmission.

15 vistas. Demyelination and Remyelination of Mouse Spinal Cord Neurons

Video: Demyelination and Remyelination of Mouse Spinal Cord Neurons - Experimento

Rat Model of Widespread Cerebral Cortical Demyelination Induced by an Intracerebral Injection of Pro-Inflammatory Cytokines

Multiple sclerosis (MS) is the most common immune-mediated disease of the central nervous system (CNS) and progressively leads to physical disability and death, caused by white matter lesions in the spinal cord and cerebellum, as well as by demyelination in grey matter. Whilst conventional models of experimental allergic encephalomyelitis are suitable for the investigation of the cell-mediated inflammation in the spinal and cerebellar white matter, they fail to address grey matter pathologies. Here, we present the experimental protocol for a novel rat model of cortical demyelination allowing the investigation of the pathological and molecular mechanisms leading to cortical lesions. The demyelination is induced by an immunization with low-dose myelin oligodendrocyte glycoprotein (MOG) in an incomplete Freund&#39;s adjuvant followed by a catheter-mediated intracerebral delivery of pro-inflammatory cytokines. The catheter, moreover, enables multiple rounds of demyelination without causing injection-induced trauma, as well as the intracerebral delivery of potential therapeutic drugs undergoing a preclinical investigation. The method is also ethically favorable as animal pain and distress or disability are controlled and relatively minimal. The expected timeframe for the implementation of the entire protocol is around 8 - 10 weeks.

The protocol presented here allows the reproduction of a widespread grey matter demyelination of both cortical hemispheres in adult male Dark Agouti rats. The method comprises of intracerebral implantation of a catheter, subclinical immunization against myelin oligodendrocyte glycoprotein, and intracerebral injection of a pro-inflammatory cytokine mixture through the implanted catheter.

Modelo de rata de desmielinización cortical cerebral generalizada inducida por una inyección intracerebral de citoquinas proinflamatorias

Multiple sclerosis (MS) is the most common immune-mediated disease of the central nervous system (CNS) and progressively leads to physical disability and ...

Investigación

JoVE Journal

Neurociencias

Positron Emission Tomography Imaging for In Vivo Measuring of Myelin Content in the Lysolecithin Rat Model of Multiple Sclerosis

Multiple sclerosis (MS) is a neuroinflammatory disease with expanding axonal and neuronal degeneration and demyelination in the central nervous system, leading to motor dysfunctions, psychical disability, and cognitive impairment during MS progression. Positron emission tomography (PET) is an imaging technique able to quantify in vivo cellular and molecular alterations.
Radiotracers with affinity to intact myelin can be used for in vivo imaging of myelin content changes over time. It is possible to detect either an increase or decrease in myelin content, what means this imaging technique can detect demyelination and remyelination processes of the central nervous system. In this protocol we demonstrate how to use PET imaging to detect myelin changes in the lysolecithin rat model, which is a model of focal demyelination lesion (induced by stereotactic injection) (i.e., a model of multiple sclerosis disease). 11C-PIB PET imaging was performed at baseline, and 1 week and 4 weeks after stereotaxic injection of lysolecithin 1% in the right striatum (4 &#181;L) and corpus callosum (3 &#181;L) of the rat brain, allowing quantification of focal demyelination (injection site after 1 week) and the remyelination process (injection site at 4 weeks).
Myelin PET imaging is an interesting tool for monitoring in vivo changes in myelin content which could be useful for monitoring demyelinating disease progression and therapeutic response.

Positron Emission Tomography Imaging for In Vivo Measuring of Myelin Content in the Lysolecithin Rat Model of Multiple Sclerosis

This protocol has the aim of monitoring in vivo myelin changes (demyelination and remyelination) by positron emission tomography (PET) imaging in an animal model of multiple sclerosis.

Imágenes de tomografía por emisión de Positron para la medición in vivo del contenido de mielina en el modelo de rata de lysolecitina de esclerosis múltiple

Multiple sclerosis (MS) is a neuroinflammatory disease with expanding axonal and neuronal degeneration and demyelination in the central nervous system, ...

A Stably Established Two-Point Injection of Lysophosphatidylcholine-Induced Focal Demyelination Model in Mice

Receptor-mediated lysophospholipid signaling contributes to the pathophysiology of diverse neurological diseases, especially multiple sclerosis (MS). Lysophosphatidylcholine (LPC) is an endogenous lysophospholipid associated with inflammation, and it could induce rapid damage with toxicity to myelin lipids, leading to focal demyelination. Here, a detailed protocol is presented for stereotactic two-point LPC injection that could directly cause severe demyelination and replicate the experimental demyelination injury quickly and stably in mice by surgical procedure. Thus, this model is highly relevant to demyelination diseases, especially MS, and it can contribute to the related advancing clinically-relevant research. Also, immunofluorescence and Luxol fast blue staining methods were used to depict the time course of demyelination in the corpus callosum of mice injected with LPC. In addition, the behavioral method was used to evaluate the cognitive function of mice after modeling. Overall, the two-point injection of lysophosphatidylcholine via a stereotaxic frame is a stable and reproducible method to generate a demyelination model in mice for further study.

The present protocol describes a two-point injection of lysophosphatidylcholine via a stereotaxic frame to generate a stable and reproducible demyelination model in mice.

Una inyección de dos puntos establecida de manera estable de modelo de desmielinización focal inducida por lisofosfatidilcolina en ratones

Receptor-mediated lysophospholipid signaling contributes to the pathophysiology of diverse neurological diseases, especially multiple sclerosis (MS). ...

Demyelination and Remyelination of Mouse Spinal Cord Neurons

Transcripción

Demyelination and Remyelination of Mouse Spinal Cord Neurons