This work was supported by the National Natural Science Foundation of China (Grants: 82071380, 81873743).

All animal procedures were approved by the Institute of Animal Care Committee of Tongji Medical College, Huazhong University of Science and Technology, China. Adult C57BL/6 male and female mice (wild type, WT; 20-25 g; 8-10 weeks old) were used for the present study. The mice were obtained from commercial sources (see Table of Materials). Mice were housed in a specific pathogen-free (SPF) animal facility with water and food supplied ad libitum. They were kept in an alternating 12 h period of light and dark cycle in the standard conditions of 22 &#176;C temperature and relative humidity of 55%-60%.
1. LPC solution preparation
<ol>
	<li>Dissolve 25 mg of LPC powder (see Table of Materials) with 250 &#181;L of chloroform and methanol mixed solution (1:1) to make a 10% LPC solution and transfer it to a 500 &#181;L centrifuge tube. 
	NOTE: If the LPC is not completely dissolved, place the centrifuge tube in an ultrasonic cleaner and ultrasonicate at 40 kHz for ~1 h to obtain a uniform solution.</li>
	<li>Divide the solution into 3 &#181;L/tube and store at &#8722;80 &#176;C. 
	NOTE: The solution can be stored for ~2 years.</li>
	<li>Before surgery, dilute the solution (step 1.2.) with 27 &#181;L of 0.9% NaCl solution, and keep the solution in a constant temperature water bath at 37 &#176;C. 
	&#8203;NOTE: Prepare the solution just before starting the injection.</li>
</ol>
2. Surgical preparation
<ol>
	<li>Use a 32 G, 2 in needle to connect to a 5 &#181;L syringe (see Table of Materials). Ensure that the microliter syringe is unobstructed. Withdraw 5 &#181;L of LPC solution for preparation of the injection.</li>
	<li>Anesthetize the mouse in an induction chamber connected to an isoflurane vaporizer with 3% isoflurane mixed with 100% oxygen at a rate of about 0.3 L/min.</li>
	<li>Confirm the depth of anesthesia by the lack of toe-pinch reflex while the breathing is smooth.</li>
	<li>Shave the head of the mouse between the ears using an&#160;electric shaver. Apply lubricating eye drops to prevent corneal dryness during the procedure.</li>
	<li>Then, position the mouse in a stereotaxic frame (see Table of Materials) with the dorsal side up, and secure the head with a nose cone and tooth clamp. Maintain anesthesia with 1.2%-1.6% isoflurane through its nose. 
	&#8203;NOTE: Isoflurane concentration can be adjusted according to the respiratory status of the mice.</li>
</ol>
3. Surgical procedure
NOTE: Animals are placed on a heating pad during all procedures.
<ol>
	<li>Fix the mouse to the stereotaxic apparatus with bilateral ear bars. Ensure that the ear bars are level and the head is horizontal and stable.</li>
	<li>Disinfect the skin on the head&#160;by wiping several times with iodophor followed by alcohol in circular motion. Then use a scalpel&#160;to cut a small incision of about 1.5 cm along the midline of the scalp to expose the skull.</li>
	<li>Wipe the skull with a cotton swab dipped in 1% hydrogen peroxide until the bregma, lambda, and posterior fontanelle are exposed. Place the syringe on the stereotaxic apparatus. 
	NOTE: Use hydrogen peroxide with care and avoid touching the surrounding tissues.</li>
	<li>Ensure horizontal positioning of the animal&#39;s head (both front and rear and left and right).
	<ol>
		<li>Adjust the Z-axis knob of the stereotaxic frame so that the needle tip and the skull just touch without bending, then measure the Z-axis coordinate. Check the Z coordinates of bregma and posterior fontanelle.</li>
		<li>Adjust the ear bar so that the difference between the Z coordinates of bregma and posterior fontanelle is no more than 0.02 mm. Then, follow the same method to measure the Z coordinates of the corresponding positions on the left and right sides of the midline. Adjust the ear bar to ensure the left and right are at the same level.</li>
	</ol>
	</li>
	<li>Locate the corpus callosum. Set the XYZ origin to bregma. 
	NOTE: The first injection site is 1.0 mm lateral to the bregma, 2.4 mm deep, and 1.1 mm anterior. The second injection site is 1.0 mm lateral to the bregma, 2.1 mm deep, and 0.6 mm anterior. For example, the coordinate of the bregma is (0,0,0). Measure the Z coordinate of the corresponding location marked as (-1, 1.1, X) and (-1, 0.6, Y). It can be determined the first injection site coordinate of the corpus callosum is (-1, 1.1, &#8722;[X + 2.4]), and the second injection site is (-1, 0.6, &#8722;[Y + 2.1]).</li>
	<li>Once the injection site is determined, make a mark with a sterile marker on the skull and record the coordinates.</li>
	<li>Gently drill the marked site with a skull drill (see Table of Materials). Take care to avoid any bleeding.</li>
	<li>Slowly move the needle to the given coordinates and start the injection. To induce corpus callosum demyelination, inject 2 &#181;L of LPC solution (step 1.) at each injection site (step 3.5.) at a rate of 0.4 &#181;L/min.</li>
	<li>After injection, keep the needle in each site for an additional 10 min. 
	&#8203;NOTE: Ensure that the interval of two injections is no more than 20 min.</li>
	<li>Suture the skin with a 4-0 suture and wait for the animal to wake up within 10 min. Administer analgesics&#160;postoperatively in accordance with institutional animal care regulations. 
	NOTE: The recommended time for euthanasia can be determined according to the purpose of the experiment.&#160;</li>
</ol>
4. Sample extraction for focal demyelination
NOTE: For details regarding this step, please see the previously published report14.
<ol>
	<li>Dissolve neutral red dye (see Table of Materials) in a 1% solution in phosphate-buffered saline (PBS).</li>
	<li>hours before sacrificing the mice (for details, see previous reference10), inject 500 &#181;L of 1% neutral red dye in PBS by intraperitoneal injection for each mouse.</li>
	<li>Perform cardiac perfusion12 with 30 mL of 0.1% PBS precooled at 4 &#176;C.</li>
	<li>Slice the brain into 1 mm with a brain mold (see Table of Materials).</li>
	<li>Visualize the lesion stained with neutral red under the microscope and dissociate the lesion. 
	&#8203;NOTE: Remove as much surrounding normal tissue as possible to improve the accuracy of subsequent analysis. The lesion tissue can be examined with RT-PCR, electron microscopy, and western blot analyses.</li>
</ol>
5. Histological staining and immunofluorescence
<ol>
	<li>For histologic staining and immunofluorescence12, after cardiac perfusion (step 4.3.), remove the brain10, fix in 4% PFA overnight (at 4 &#176;C), and completely dehydrate in 30% sucrose.</li>
	<li>Cut into 20 mm coronal brain sections using a frozen slicer10 at constant temperature (&#8722;20 &#176;C).</li>
	<li>Use the slices for Luxol fast blue (LFB) staining, immunofluorescence, and western blot10.</li>
</ol>

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The authors have nothing to disclose.

MS, a chronic demyelinating disease of the CNS, is one of the most common causes of neurological dysfunction in young adults20. Clinically, approximately 60%-80% of MS patients experience the cycle of relapses and remissions before developing a secondary-progressive MS21,22, and it eventually leads to cumulative movement impairments and cognitive deficits over time23. Currently, no single experimental model covers t...

Receptor-mediated lysophospholipid signaling involves diverse physiological processes of almost all organ systems1. In the central nervous system (CNS), this signaling plays a critical role in the pathogenies of autoimmune neurological diseases such as multiple sclerosis (MS). Multiple sclerosis is a chronic immune-mediated disorder characterized by pathological demyelination and inflammatory response, causing neurologic dysfunction and cognitive impairment2,3. After continuous relapsing and remitting during the early disease, most patients eventually progress to the secondary-progressive stage, which could cause irreversible damage to the brain and resulting disability4. It is believed that the pathological hallmark of the secondary-progressive course is demyelinating plaques caused by inflammatory lesions5. Existing treatments for MS can significantly reduce the risk of relapse. However, there is still no effective therapy for long-term demyelinating damage caused by progressive MS6. Thus, a stably established and easily reproducible model is required to study preclinical therapeutics that focus on white matter degeneration.
Demyelination and remyelination are two major pathological processes in developing multiple sclerosis. Demyelination is the loss of myelin sheath around axons induced by microglia with pro-inflammatory phenotypes7, and it leads to slow conduction of nerve impulses and results in the loss of neurons and neurological disorders. Remyelination is an endogenous repair response mediated by oligodendrocytes, where disorders could lead to neurodegeneration and cognitive impairment8. The inflammatory response is crucial to the whole process, affecting both the degree of myelin damage and repair.
Therefore, a stable animal model of persistent inflammatory demyelination is meaningful for further exploration of therapeutic strategies for MS. Due to the complexity of MS, various types of animal models have been established to mimic demyelinating lesions in vivo, including experimental autoimmune encephalomyelitis (EAE), toxic-demyelinating models, cuprizone (CPZ), and lysophosphatidylcholine (LPC)9. LPC is an endogenous lysophospholipid associated with inflammation, and it could induce rapid damage with toxicity to myelin lipids, leading to focal demyelination. Based on previous reports and research10,11, a detailed protocol of two-point injection with some modifications is provided. Generally, the classic one-point LPC injection model only produces local demyelination at the injection site and is often accompanied by spontaneous remyelination12,13. However, the two-point injection LPC model can demonstrate that the LPC can directly induce demyelination in the mouse corpus callosum and cause more durable demyelination with little myelin regeneration.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >L-&#945;-Lysophosphatidylcholine from egg yolk</td>
			<td >Sigma-Aldrich</td>
			<td >L4129-25MG</td>
			<td ></td>
		</tr>
		<tr >
			<td >32 gauge Needle</td>
			<td >HAMILTON</td>
			<td >7762-05</td>
			<td></td>
		</tr>
		<tr >
			<td >10 &#956;l syringe</td>
			<td >HAMILTON</td>
			<td >80014</td>
			<td></td>
		</tr>
		<tr >
			<td >high speed skull drill</td>
			<td >strong,korea</td>
			<td >strong204</td>
			<td></td>
		</tr>
		<tr >
			<td >drill</td>
			<td >Hager &#38; Meisinger, Germany&#160;</td>
			<td >REF 500 104 001 001 005</td>
			<td></td>
		</tr>
		<tr >
			<td >Matrx Animal Aneathesia Ventilator</td>
			<td >MIDMARK</td>
			<td >VMR</td>
			<td></td>
		</tr>
		<tr >
			<td >Portable Stereotaxic Instrument for Mouse</td>
			<td >Reward</td>
			<td >68507</td>
			<td></td>
		</tr>
		<tr >
			<td >Micro syringe</td>
			<td >Reward</td>
			<td >KDS LEGATO 130</td>
			<td></td>
		</tr>
		<tr >
			<td >Isoflurane&#160;</td>
			<td >VETEASY</td>
			<td ></td>
			<td></td>
		</tr>
		<tr >
			<td >Paraformaldehyde</td>
			<td >Servicebio</td>
			<td >G1101</td>
			<td></td>
		</tr>
		<tr >
			<td >Phosphate buffer</td>
			<td >BOSTER</td>
			<td >PYG0021</td>
			<td></td>
		</tr>
		<tr >
			<td >LuxoL fast bLue</td>
			<td >Servicebio</td>
			<td >G1030-100ML</td>
			<td></td>
		</tr>
		<tr >
			<td >Suture</td>
			<td >FUSUNPHARMA</td>
			<td >20152021225</td>
			<td></td>
		</tr>
		<tr >
			<td >Brain mold</td>
			<td >Reward</td>
			<td >68707</td>
			<td></td>
		</tr>
		<tr >
			<td >Electron microscope fixative</td>
			<td >Servicebio</td>
			<td >G1102-100ML</td>
			<td></td>
		</tr>
		<tr >
			<td >Neutral red (C.I. 50040), for microscopy Certistain</td>
			<td >Sigma-Aldrich</td>
			<td >1.01376</td>
			<td></td>
		</tr>
		<tr >
			<td >Anti-Myelin Basic Protein Antibody&#160;</td>
			<td >Millipore</td>
			<td >#AB5864</td>
			<td></td>
		</tr>
		<tr >
			<td >Anti-GST-P pAb</td>
			<td >MBL</td>
			<td >#311</td>
			<td></td>
		</tr>
		<tr >
			<td >Ki-67 Monoclonal Antibody (SolA15)</td>
			<td >Thermo Fisher&#160;Scientific</td>
			<td >14-5698-95</td>
			<td></td>
		</tr>
		<tr >
			<td >Beta Actin Monoclonal Antibody</td>
			<td >Proteintech</td>
			<td >66009-1-Ig&#160;</td>
			<td></td>
		</tr>
		<tr >
			<td >Myelin Basic Protein Polyclonal Antibody</td>
			<td >Proteintech</td>
			<td >10458-1-AP</td>
			<td></td>
		</tr>
		<tr >
			<td >OLIG2 Polyclonal Antibody</td>
			<td >Proteintech</td>
			<td >13999-1-AP</td>
			<td></td>
		</tr>
		<tr >
			<td >Alexa Fluor 488&#160;AffiniPure&#160;Donkey anti-Rabbit&#160;IgG (H+L)</td>
			<td >YEASEN</td>
			<td >34206ES60</td>
			<td></td>
		</tr>
		<tr >
			<td >Alexa Fluor 594 AffiniPure Donkey Anti-Rat IgG (H+L)&#160;</td>
			<td >YEASEN</td>
			<td >34412ES60</td>
			<td></td>
		</tr>
		<tr >
			<td >Alexa Fluor 594&#160;AffiniPure Donkey Anti-Rabbit IgG (H+L)&#160;</td>
			<td >YEASEN</td>
			<td >34212ES60</td>
			<td></td>
		</tr>
		<tr >
			<td >HRP Goat Anti-Rabbit IgG (H+L)</td>
			<td >abclonal</td>
			<td >AS014</td>
			<td></td>
		</tr>
		<tr >
			<td >HRP Goat Anti-Mouse IgG (H+L)&#160;</td>
			<td >abclonal</td>
			<td >AS003</td>
			<td></td>
		</tr>
		<tr >
			<td >Adult C57BL/6 male and female mice</td>
			<td >Hunan SJA Laboratory Animal Co. Ltd</td>
			<td ></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Two-point injection of the LPC resulted in a more durable demyelination 
LPC mainly leads to rapid damage with toxicity to myelin and cleavage of the axon integrity15. The day of injection was regarded as day 0. Mice were kept for a period of 10-28 days (10 dpi and 28 dpi). Luxol fast blue (LFB) staining10 was used to evaluate the area of demyelination in mice at these time points. In the two-point injection model, there was significant demyelination o...

Watch this Scientific Journal Video about A Stably Established Two-Point Injection of Lysophosphatidylcholine-Induced Focal Demyelination Model in Mice at JoVE.com

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

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

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

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3.4K Views.  Huazhong University of Science and Technology. The present protocol describes a two-point injection of lysophosphatidylcholine via a stereotaxic frame to generate a stable and reproducible demyelination model in mice.

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

Autologous Blood Injection to Model Spontaneous Intracerebral Hemorrhage in Mice

Investigation of the pathophysiology of injury after intracerebral
hemorrhage (ICH) requires a reproducible animal model. While ICH
accounts for 10-15% of all strokes, there remains no specific
effective therapy. The autologous blood injection model in mice
involves the stereotaxic injection of arterial blood into the basal
ganglia mimicking a spontaneous hypertensive hemorrhage in man. The
response to hemorrhage can then be studied in vivo and the
neurobehavioral deficits quantified, allowing for description of the
ensuing pathology and the testing of potential therapeutic agents. The
procedure described in this protocol uses the double injection
technique to minimize risk of blood reflux up the needle track, no
anticoagulants in the pumping system, and eliminates all dead space
and expandable tubing in the system.

The autologous blood injection model of intracerebral hemorrhage in mice described in this protocol uses the double injection technique to minimize risk of blood reflux up the needle track, no anticoagulants in the pumping system, and eliminates all dead space and expandable tubing in the system.

Investigation of the pathophysiology of injury after intracerebral
hemorrhage (ICH) requires a reproducible animal model.  While ICH
accounts for 10-15% ...

Inhibitory Synapse Formation in a Co-culture Model Incorporating GABAergic Medium Spiny Neurons and HEK293 Cells Stably Expressing GABAA Receptors

Inhibitory neurons act in the central nervous system to regulate the dynamics and spatio-temporal co-ordination of neuronal networks. GABA (&#947;-aminobutyric acid) is the predominant inhibitory neurotransmitter in the brain. It is released from the presynaptic terminals of inhibitory neurons within highly specialized intercellular junctions known as synapses, where it binds to GABAA receptors (GABAARs) present at the plasma membrane of the synapse-receiving, postsynaptic neurons. Activation of these GABA-gated ion channels leads to influx of chloride resulting in postsynaptic potential changes that decrease the probability that these neurons will generate action potentials. 
During development, diverse types of inhibitory neurons with distinct morphological, electrophysiological and neurochemical characteristics have the ability to recognize their target neurons and form synapses which incorporate specific GABAARs subtypes. This principle of selective innervation of neuronal targets raises the question as to how the appropriate synaptic partners identify each other. 
To elucidate the underlying molecular mechanisms, a novel in vitro co-culture model system was established, in which medium spiny GABAergic neurons, a highly homogenous population of neurons isolated from the embryonic striatum, were cultured with stably transfected HEK293 cell lines that express different GABAAR subtypes. Synapses form rapidly, efficiently and selectively in this system, and are easily accessible for quantification. Our results indicate that various GABAAR subtypes differ in their ability to promote synapse formation, suggesting that this reduced in vitro model system can be used to reproduce, at least in part, the in vivo conditions required for the recognition of the appropriate synaptic partners and formation of specific synapses. Here the protocols for culturing the medium spiny neurons and generating HEK293 cells lines expressing GABAARs are first described, followed by detailed instructions on how to combine these two cell types in co-culture and analyze the formation of synaptic contacts.

Inhibitory Synapse Formation in a Co-culture Model Incorporating GABAergic Medium Spiny Neurons and HEK293 Cells Stably Expressing GABAA Receptors

The molecular mechanisms that co-ordinate the formation of inhibitory GABAergic synapses during ontogeny are largely unknown. To study these processes,we have developed a co-culture model system which incorporates embryonic medium spiny GABAergic neurons cultured together with stably transfected human embryonic kidney 293 (HEK293) cells expressing functional GABAA receptors.

Inhibitory neurons act in the central nervous system to regulate the dynamics and spatio-temporal co-ordination of neuronal networks. GABA ...

Experimental Demyelination and Remyelination of Murine Spinal Cord by Focal Injection of Lysolecithin

Multiple sclerosis is an inflammatory demyelinating disease of the central nervous system characterized by plaque formation containing lost oligodendrocytes, myelin, axons, and neurons. Remyelination is an endogenous repair mechanism whereby new myelin is produced subsequent to proliferation, recruitment, and differentiation of oligodendrocyte precursor cells into myelin-forming oligodendrocytes, and is necessary to protect axons from further damage. Currently, all therapeutics for the treatment of multiple sclerosis target the aberrant immune component of the disease, which reduce inflammatory relapses but do not prevent progression to irreversible neurological decline. It is therefore imperative that remyelination-promoting strategies be developed which may delay disease progression and perhaps reverse neurological symptoms. Several animal models of demyelination exist, including experimental autoimmune encephalomyelitis and curprizone; however, there are limitations in their use for studying remyelination. A more robust approach is the focal injection of toxins into the central nervous system, including the detergent lysolecithin into the spinal cord white matter of rodents. In this protocol, we demonstrate that the surgical procedure involved in injecting lysolecithin into the ventral white matter of mice is fast, cost-effective, and requires no additional materials than those commercially available. This procedure is important not only for studying the normal events involved in the remyelination process, but also as a pre-clinical tool for screening candidate remyelination-promoting therapeutics.

Demyelinating diseases can be modeled in animals by focal application of lysolecithin into the CNS. A single injection of lysolecithin into mouse spinal cord produces a lesion that spontaneously repairs over time. The goal is to study factors involved in de- and remyelination, and to test agents for enhancing repair.

Multiple sclerosis is an inflammatory demyelinating disease of the central nervous system characterized by plaque formation containing lost ...

Visual Evoked Potential Recording in a Rat Model of Experimental Optic Nerve Demyelination

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to monitor the disease course in the follow-up period. Changes in the VEP parameters closely correlate with pathological damage in the optic nerve. This protocol provides a detailed description about the rodent model of optic nerve microinjection, in which a partial demyelination lesion is produced in the optic nerve. VEP recording techniques are also discussed. Using skull implanted electrodes, we are able to acquire reproducible intra-session and between-session VEP traces. VEPs can be recorded on individual animals over a period of time to assess the functional changes in the optic nerve longitudinally. The optic nerve demyelination model, in conjunction with the VEP recording protocol, provides a tool to investigate the disease processes associated with demyelination and remyelination, and can potentially be employed to evaluate the effects of new remyelinating drugs or neuroprotective therapies.

Focal demyelination is induced in the optic nerve using lysolecithin microinjection. Visual evoked potentials are recorded via skull electrodes implanted over the visual cortex to examine the signal conduction along the visual pathway in vivo. This protocol details the surgical procedures underlying electrode implantation and optic nerve microinjection.

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to ...

Partial Optic Nerve Transection in Rats: A Model Established with a New Operative Approach to Assess Secondary Degeneration of Retinal Ganglion Cells

Previous studies have shown that the secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. Partial optic nerve transection is considered a useful and reproducible model. Compared with other optic nerve injury models used commonly for assessing secondary degeneration, e.g. complete optic nerve transection and optic nerve crush models, the partial optic nerve transection model is superior as it distinguishes primary from secondary degeneration in situ. Therefore, it serves as an excellent tool for evaluating secondary degeneration. This study describes a novel operative approach of partial optic nerve transection by directly accessing the area of the retrobulbar optic nerve through the orbital lateral wall of the eyeball. Moreover, we present a newly designed, low cost surgical instrument to assist with transection. As demonstrated by the representative results in distinguishing the boundary of primary and secondary injury areas, the new approach and instrument ensures high efficiency and stability of the model by providing adequate space for surgical operation. This in turn makes it easy to separate the meningeal sheath and ophthalmic vessels from the optic nerve before transection. An additional benefit is that this space-saving operative approach improves the investigators&#39; ability to administer drugs, carriers, or selective RGC tracers to the stump of the partially transected optic nerve, allowing the exploration of mechanisms behind secondary injury in RGCs, in a new way.

Secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. This study describes an innovative operative approach for partial optic nerve transection. The use of this space-saving operative approach extends the model's application range, and allows exploration of secondary injury mechanisms in RGCs in a new way.

Previous studies have shown that the secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. Partial optic nerve transection ...

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.

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

Direct Intrathecal Injection of Recombinant Adeno-associated Viruses in Adult Mice

Intrathecal (IT) injection of adeno-associated virus (AAV) has drawn considerable interest in CNS gene therapy by virtue of its safety, noninvasiveness, and excellent transduction efficacy in the CNS.&#160;Previous studies have demonstrated the therapeutic potency of AAV-delivered gene therapy in neurodegenerative disorders by IT administration. However, high rates of unpredictable failure due to the technical limitation of IT administration in small animals have been reported. Here, we established a scoring system to indicate the success extent of lumbar puncture in small animals by adding 1% lidocaine hydrochloride into the injection solution. We further show that the extent of transient weakness following injection can predict the transduction efficiency of AAV. Thus, this IT injection method can be used to optimize therapeutic trials in mouse models of CNS diseases that afflict wide regions of the CNS.

Here we present a direct intrathecal injection technique using 1% lidocaine hydrochloride in a viral solution to ensure efficient adeno-associated virus delivery to small animals and establish a scoring system to predict transduction efficiency in the central nervous system according to the degree of transient weakness induced by lidocaine.

Intrathecal (IT) injection of adeno-associated virus (AAV) has drawn considerable interest in CNS gene therapy by virtue of its safety, noninvasiveness, ...

Double Direct Injection of Blood into the Cisterna Magna as a Model of Subarachnoid Hemorrhage

Among strokes, subarachnoid hemorrhage (SAH) consecutive to the rupture of a cerebral arterial aneurysm represents 5-9% but is responsible for about 30% of the total stroke-related mortality with an important morbidity in terms of neurological outcome. A delayed cerebral vasospasm (CVS) may occur most often in association with a delayed cerebral ischemia. Different animal models of SAH are now being used including endovascular perforation and direct injection of blood into the cisterna magna or even the prechiasmatic cistern, each exhibiting distinct advantages and disadvantages. In this article, a standardized mouse model of SAH by double direct injection of determined volumes of autologous whole blood into the cisterna magna is presented. Briefly, mice were weighed and then anesthetized by isoflurane inhalation. Then, the animal was placed in a reclining position on a heated blanket maintaining a rectal temperature of 37 &#176;C and positioned in a stereotactic frame with a cervical bend of about 30&#176;. Once in place, the tip of an elongated glass micropipette filled with the homologous arterial blood taken from carotid artery of another mouse of the same age and gender (C57Bl/6J) was positioned at a right angle in contact with the atlanto-occipital membrane by means of a micromanipulator. Then 60 &#181;L of blood was injected in the cisterna magna followed by a 30&#176; downward tilt of the animal for 2 minutes. The second infusion of 30 &#181;L of blood into the cisterna magna was performed 24 h after the first one. The individual follow-up of each animal is carried out daily (careful evaluation of weight and well-being). This procedure allows a predictable and highly reproducible distribution of blood, likely accompanied by intracranial pressure elevation that can be mimicked by an equivalent injection of an artificial cerebral spinal fluid (CSF), and represents an acute to mild-model of SAH inducing low mortality.

We described in this protocol a standardized subarachnoid hemorrhage (SAH) mouse model by a double injection of autologous whole-blood into the cisterna magna. The high degree of standardization of the double-injection procedure represents a middle-to-acute model of SAH with relative safety regarding mortality.

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

Modeling Stroke in Mice: Focal Cortical Lesions by Photothrombosis

Stroke is a leading cause of death and acquired adult disability in developed countries. Despite extensive investigation for novel therapeutic strategies, there remain limited therapeutic options for stroke patients. Therefore, more research is needed for pathophysiological pathways such as post-stroke inflammation, angiogenesis, neuronal plasticity, and regeneration. Given the inability of in vitro models to reproduce the complexity of the brain, experimental stroke models are essential for the analysis and subsequent evaluation of novel drug targets for these mechanisms. In addition, detailed standardized models for all procedures are urgently needed to overcome the so-called replication crisis. As an effort within the ImmunoStroke research consortium, a standardized photothrombotic mouse model using an intraperitoneal injection of Rose Bengal and the illumination of the intact skull with a 561 nm laser is described. This model allows the performance of stroke in mice with allocation to any cortical region of the brain without invasive surgery; thus, enabling the study of stroke in various areas of the brain. In this video, the surgical methods of stroke induction in the photothrombotic model along with&#160;histological analysis are demonstrated.

Described here is the photothrombotic stroke model, where a stroke is produced through the intact skull by inducing permanent microvascular occlusion using laser illumination after administration of a photosensitive dye.

Stroke is a leading cause of death and acquired adult disability in developed countries. Despite extensive investigation for novel therapeutic strategies, ...

Modeling Neonatal Intraventricular Hemorrhage Through Intraventricular Injection of Hemoglobin

Neonatal intraventricular hemorrhage (IVH) is a common consequence of premature birth and leads to brain injury, posthemorrhagic hydrocephalus (PHH), and lifelong neurological deficits. While PHH can be treated by temporary and permanent cerebrospinal fluid (CSF) diversion procedures (ventricular reservoir and ventriculoperitoneal shunt, respectively), there are no pharmacological strategies to prevent or treat IVH-induced brain injury and hydrocephalus. Animal models are needed to better understand the pathophysiology of IVH and test pharmacological treatments. While there are existing models of neonatal IVH, those that reliably result in hydrocephalus are often limited by the necessity for large-volume injections, which may complicate modeling of the pathology or introduce variability in the clinical phenotype observed.
Recent clinical studies have implicated hemoglobin and ferritin in causing ventricular enlargement after IVH. Here, we develop a straightforward animal model that mimics the clinical phenotype of PHH utilizing small-volume intraventricular injections of the blood breakdown product hemoglobin. In addition to reliably inducing ventricular enlargement and hydrocephalus, this model results in white matter injury, inflammation, and immune cell infiltration in periventricular and white matter regions. This paper describes this clinically relevant, simple method for modeling IVH-PHH in neonatal rats using intraventricular injection and presents methods for quantifying ventricle size post injection.

We present a model of neonatal intraventricular hemorrhage using rat pups that mimics the pathology seen in humans.

Neonatal intraventricular hemorrhage (IVH) is a common consequence of premature birth and leads to brain injury, posthemorrhagic hydrocephalus (PHH), and ...