We thank C. Ricard, M-C. Amoureux, A. Jaouen, and S. Meijia-Gervacio for helpful discussions; M. Hocine and C. Meunier, and the staff of the animal and PicSIL imaging facilities of the IBDML for technical support. This work was supported by an institutional grant from Centre National de la Recherche Scientifique, and by grants from the Association de Recherche sur la Scl&#233;rose en Plaque (ARSEP) (to G.R. and K.K.F), Agence Nationale de la Recherche (ANR JCJC), F&#233;d&#233;ration de Recherche sur le Cerveau (FRC) Institut de Recherche sur la Moelle Epini&#232;re (IRME) (to FD). K.K.F. was supported by an ARSEP fellowship. Imaging was performed on the PIcSIL imaging facility of the IBDML.

1. Window and Support Structure Preparation
<ol>
	<li>Window preparation - prepare several windows of various sizes (width = 1.8-2.2 mm; length = 4.5-5.2 mm) for implantation.
	<ol>
		<li>Making the modified drill burr to cut the cover glass - using a generous amount of dental cement attach a drill burr to a shaft (in our case old Bulldog clamps). Be sure that the burr is ~45&#176; relative to the shaft for easier handling. 
		Note: diamond scribes also work well for cutting cover glass.</li>
		<li>Making slide-A - using a standard microscope slide, tape a short piece of ruler paper (~2 cm) to the bottom of the slide so that you can see the ruler through the glass. Next, place a piece of tape along the edge of the 0 mm mark of the ruler to act as a fixture for the cover glass in the next step.</li>
		<li>Using the modified drill burr, slide-A, and an unmodified microscope slide (slide-B), position a piece of cover glass (no. 1.5 thickness) to the 0 mm mark on slide-A. Use slide-B to hold the cover glass in place and as a jig for the intended cut line. Cut the cover glass into a long rectangle of approximately 5 x 22 mm. Rotate the cut glass 90&#176; and cut the rectangle into smaller rectangle of ~2 x 5 mm.</li>
	</ol>
	</li>
	<li>Staple modifications - two modified staples will be implanted to support and fix the window chamber to the vertebrae (see section 2.7-8).
	<ol>
		<li>Using a standard small staple (crown = 8 mm; leg = 4 mm), bend the middle of the crown ~90&#176; so that the tips of the legs are nearly touching.</li>
		<li>Bend the tips of the legs outwards ~45&#176; at the half-way point, so that the distal half of the legs are parallel.</li>
		<li>Bend the distal half of the legs ~30&#176; upward relative to the plane of the staple. Place the modified staples into ethanol prior to surgery for sterilization. 
		Note: Staples can also be sterilized using ethylene oxide or autoclaved.</li>
	</ol>
	</li>
	<li>Paperclip modifications - prepare one modified paperclip to be implanted as part of the chamber (see sections 2.9-10) and to act as a holding clip for surgery and imaging.
	<ol>
		<li>Using a standard 25 mm paperclip, remove the small bend leaving an elongated U-shape.</li>
		<li>Using pliers or needle-drivers, reduce the diameter of the bend to ~2 mm while keeping the tips parallel.</li>
		<li>Bend the tips outward ~90&#176; at about 8-10 mm from the bend. Make any adjustments necessary so that it lies flat.</li>
	</ol>
	</li>
</ol>
2. Window Implantation
Note: All surgical and animal care procedures were conducted in accordance with the European Communities Council Directive and were approved by the National Animal Studies Committee of France (authorization no. 13,300).
<ol>
	<li>Anesthetize the mouse (Ket/Xyl; induction dose: 120 mg/kg, 12 mg/kg; supplemental dose, 40 mg/kg, 4 mg/kg), apply eye ointment to the eyes.</li>
	<li>Shave its back approximately ~2 cm wide by ~5 cm long, centered on the thoracic hump, and clean the exposed skin using iodine soap followed by iodine solution. Clean gloves by spraying with 90% ethanol.&#160;</li>
	<li>Using tissue scissors cut the dorsal skin longitudinally over vertebrae T11 to L1 (~2 cm). Using a scalpel, make bilateral longitudinal incisions between the spinous and transverse processes from T11 to L1.&#160;Note: If gloves or instruments touch the fur or skin of the animal, reclean the gloves or instrument using a cold sterilizer such as 90% ethanol or Germ-X.</li>
	<li>Suspend the mouse in the spinal fork stereotaxic - Begin by clamping the top right spinal fork into position. Grasp T11 using Adson forceps and position the T11 transverse process over the fixed spinal fork. While holding the vertebra in place, fasten the contralateral spinal fork under the contralateral transverse process.</li>
	<li>Using the same procedure as step 2.4, position L1 within the spinal fork.</li>
	<li>Using a scalpel (no. 23), gently scrape away the muscles between the spinous and transverse processes from T11 to L1 so that the laminae are clearly visible. For a better view of the pedicles of T11, cut the tendon projecting rostrally from the tip of the transverse process.</li>
	<li>Rinse the modified staples with PBS, and insert the tips of the staples lateral to the vertebral pedicles of T11 and L1 adjusting the distance between the tips so that they are lightly touching the pedicles. 
	Note: Use only PBS containing dexamethasone (0.1%), penicillin (1,000 U/ml), and streptomycin (1 mg/ml), to reduce postsurgical inflammation and to improve window clarity.</li>
	<li>Glue the staples in place with a small drop of cyanoacrylate at each insertion point and remove the mouse from the spinal forks.</li>
	<li>Apply a thin layer of cyanoacrylate to the cut edges of the skin and exposed muscles around the cleared vertebrae being careful to not glue the vertebrae. Bend the tips of the modified paperclip so that they enter under the protruding sections of the implanted staples and rest parallel to the long axis of the isolated vertebral column and leave in this position.</li>
	<li>Starting at the skin-muscle interface, apply several layers of dental cement to cover the entire opening except the exposed vertebrae, being sure to implant the paper clip securely within the dental cement.</li>
	<li>Once the dental cement is cured (5-15 min), attach the mouse to the &#39;clip holder&#39; by the exposed portion of the implanted paper clip. Using forceps, remove the muscle and other soft tissue between the spinous processes and laminae of the exposed vertebrae. 
	Note: Any device that can securely grasp the exposed portion of the paperclip without causing stress to the clip/chamber, and can be maneuvered to reorient the mouse throughout surgery can be used as a &#39;clip holder&#39;.</li>
	<li>Using the bone removal drill, thin the laminae of T12 and T13 bilaterally. To remove the dorsal bone, grasp the spinous process with blunt forceps and lift taking care to apply no downward force towards the spinal cord. If the bone does not easily come away, continue to thin the laminae until the spinous process can be easily removed.</li>
	<li>Using the bone removal drill, continue to shape the lateral edges of the vertebrae to form a support structure for the window. Gently remove all of the small bone fragments touching the dura mater using blunt forceps. Be sure to maintain the humidity of the dura mater using PBS.</li>
	<li>Choose one of the prepared windows that is slightly wider than the laminectomy opening and test how well it rests on the exposed bone. Ideally, the window will rest bilaterally on both vertebrae so that the middle of the window touches the dura mater (Figure&#160;1a). The window should not, however, compress the spinal cord even when pushing the window towards the spinal cord. If the window is unbalanced or rests above the dura, repeat step 2.13.</li>
	<li>Once satisfied with the laminectomy, place a small piece of autoclaved tissue-paper soaked in PBS onto the spinal cord to prevent drying of the dura and set the mouse aside.</li>
	<li>Cleaning the window - place the window in a solution of window cleaner for several seconds. Using a piece of microscope lens tissue, wipe the window dry. With the surgical microscope verify that the window is clean, and repeat the cleaning process if not. Place the window in a clean area with easy access for rapid retrieval and placement of the window in step 2.19.</li>
	<li>Retrieve the mouse and remove the tissue paper. Gently remove any debris from the dura using blunt forceps and allow the dura to dry until slightly tacky.</li>
	<li>Quickly fill the silicone (Kwik-sil) mixer tube and extrude silicone onto a clean slide until there are no bubbles emerging from the tip. Immediately administer a small bead of silicone along the midline of the exposed spinal cord.</li>
	<li>Retrieve the cleaned windows using blunt forceps and immediately place over the silicone. Using the blunt forceps, apply a small amount of pressure to the window to spread the silicone to the edges of the window and hold in place for ~1 min. 
	Note: Prior to mixing, silicone must be stored below 27 &#176;C otherwise the cure-time can increase significantly and it will not adhere to the window or dura mater leading to reduced optical clarity.</li>
	<li>Slowly release pressure from the window. If the window does not move, proceed immediately to step 2.21. If, however, the window rises when the pressure is released (i.e.&#160;the vertebrae move), proceed to step 2.21 while maintaining pressure on the window to hold it in place.</li>
	<li>Apply cyanoacrylate rostral and caudal of the window. If the cyanoacrylate does not cover the rostral and caudal edges of the window, use a toothpick to guide some of the liquid cyanoacrylate over these edges. Next, guide the glue along the lateral edges of the window (Figure&#160;1a) to cover all of the surrounding muscles. Note: The cyanoacrylate should cover the surface of the outer edges of the window. Too much cyanoacrylate can compromise optical clarity, not enough cyanoacrylate will result in air bubbles between the silicone and window.</li>
	<li>To protect the cyanoacrylate and increase structural integrity of the chamber, apply dental cement to all regions covered with cyanoacrylate (Figure 1a). As with the cyanoacrylate, start at the rostral and caudal edges and use a small wood stick to guide the dental cement along the edges of the window.</li>
</ol>
3. Postoperative Care, EAE Induction, and Window Maintenance
<ol>
	<li>Inject Dexamethasone (0.2 mg/kg) and Rimadyl (5 mg/kg) s.q. over the pelvic regions. Transfer the mouse to a warm cage (~26 &#176;C) with nesting material. Mice should be fully mobile, exploring their cage, and grooming after the effects of the anesthetic have fully subsided (usually 2-3 hr). House animals individually and make sure they have easy access to food and water for the first 1-2 days.&#160;Once the mouse begins to recover from anesthetic, inject buprenorphine (0.05 mg/kg) s.q. Following full recovery from anesthetic, the mouse should show no signs of pain.&#160;Note: Once fully recovered from anesthetic, any signs of pain or abnormal motor activity indicate that the window was not implanted correctly.</li>
	<li>Inject Dexamethasone(0.2 mg/kg) and Rimadyl (5 mg/kg) every 2 d for the first 10 d after surgery (Figure&#160;1a). To inject mice with windows, lightly sedate the animal with isoflurane in an induction chamber (~1.75% isoflurane in air for 1-2 min). Remove the mouse from the chamber and immediately inject the drugs. The mouse should be fully awake in &#60;1 min. 
	Note: Do not scruff mice with implanted windows since this can cause separation of the skin from the implant.&#160;It is possible to inject mice i.p. without sedation by allowing them to grab a cage lid with their forepaws and gently raising their hindlimbs. However, in this position the mouse will sometimes pull away causing stress to the implanted window. This can lead to bleeding between the window and spinal cord, thus compromising optical clarity.</li>
	<li>EAE can be induced at 2-3 weeks after the cessation of anti-inflammatory treatment (~3-4 weeks after surgery). Lightly sedate the mouse as described in step 3.2. Remove the mouse from the induction chamber and maintain the sedation with an isoflurane mask placed over the nose.</li>
	<li>Inject MOG35-55 peptide (75 &#181;g in Freund&#39;s adjuvant containing 800 &#181;g of mycobacterium tuberculosis) into 3 different s.q. locations (bilaterally over the femur, and at the base of the tail), and pertussis toxin (400 ng) i.p.14,15 
	Note: Mice with EAE require supplemental care compared to mice with windows only. Weigh EAE mice every day and be sure that they have easy access to food and water at all times. Supplement with a 3% agarose jelly containing to maintain animal hydration and weight. Glucose (1-3%) can also be added to the jelly for additional calories.</li>
	<li>Two days after EAE induction, repeat the sedation protocol outlined in step 3.2 and inject a second dose of pertussis toxin (400 ng) i.p.</li>
	<li>To image the spinal cord, first sedate the animal as described in step 3.2, and lightly anesthetize with ketamine/xylazine (100 mg/kg; 10 mg/kg) i.p. This will provide ~ 1 hr of imaging.</li>
	<li>For imaging sessions lasting &#60;2 hr, light anesthesia can be maintained with supplementary injections of ketamine/xylazine (50 mg/kg; 5 mg/kg) every 45-60 min. Alternatively, for longer imaging sessions, mice can be continuously sedated with isoflurane (0.75-1.25% isoflurane in humidified air). 
	Note: windows can be cleaned under a surgical microscope by gently wiping away debris with the corner of a tissue or a cotton-tipped swab. Do not use alcohol to clean the window because it can degrade some types of dental cement and it can cool the spinal cord.</li>
	<li>Once lightly anesthetized, the mouse is attached firmly to a clip holder by the exposed portion of the implanted paperclip (Figure&#160;1b).</li>
	<li>The spinal cord is aligned perpendicular to the objective lens by adjusting the positioning of the clip within the holder, adjusting the position of the holder, and adjusting the position of the stage. The dorsal surface of the spinal cord is rapidly imaged, and the dorsal vasculature is used to determine the orientation of the spinal cord.</li>
</ol>

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The authors declare that they have no competing financial interests.

We described a method for implanting and maintaining glass windows over the exposed spinal cords of adult mice to study the dynamic progression of EAE over time. A similar glass window implantation approach has been used to study spinal cord injury models13&#160;and may also be useful for studying other spinal cord pathologies such as glioblastomas,&#160;meningitis, or the cellular physiology of the normal spinal cord. For our method, vertebral stabilization was achieved by building an exoskeleton around four ...

Multiple sclerosis is a CNS auto-immune disease that causes progressive myelin and axon damage. The standard research model for CNS autoimmune demyelinating diseases is EAE in adult rodents. However, the dynamic cellular events underlying EAE progression remain largely unexplored. One approach is to use intravital two-photon microscopy to visualize cellular populations with fluorescent markers for extended periods of time in the spinal cords of adult mice with EAE.
Intravital two-photon microscopy is a powerful way to study cellular physiology, cellular interactions, and the dynamic progression of disease in living animals with subcellular resolution1. However, a major hurdle to the approach is that it requires optical access to the region of interest. For example, cranial glass windows are a well-established methodology for repeatedly imaging the same microstructures in the brain over several weeks2,3. Adaptation of this methodology to spinal cord imaging is, however, not straightforward and it requires several technical modifications to immobilize the dorsal spinal tissue within the articulated vertebrae.
Previously, optical access to the spinal cord required surgically reexposing the spinal cord at the beginning of each imaging session and suturing the skin closed at the end of each session4-12. These repeated surgeries are traumatic for the animal, limit the number and length of imaging sessions, and increases the probability of surgery related artifacts such as inflammation and damage to the spinal cord through mechanical perturbation. To overcome these shortcomings, we recently developed a protocol to implant glass windows over the exposed spinal cords of adult fluorescent transgenic mice with traumatic spinal cord injuries for long-term in vivo two-photon microscopy experiments without repeated surgeries13. Here we present a detailed adaptation of the implantation protocol in the context of its application to study the dynamic cellular events underlying the progression of EAE in adult mice.

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	<tbody>
		<tr height="38">
			<td height="38" style="height:39px;width:220px;">Bone removal drill</td>
			<td style="width:124px;">World Precision Instruments</td>
			<td style="width:156px;">IDEAL MICRO-DRILL</td>
			<td style="width:375px;">Also sold by Harvard Apparatus. 
			Any high quality surgical bone drill will suffice.</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;width:220px;">Drill burr (#1/4 or 1/2 Carbide Round Burr)</td>
			<td style="width:124px;">World Precision Instruments</td>
			<td style="width:156px;">501860 (#1/4) 
			501860 (#1/2)</td>
			<td>Also sold by Harvard Apparatus</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Staples</td>
			<td>Lyreco</td>
			<td>5.002.567</td>
			<td style="width:375px;">Crown = 8 mm 
			Leg = 4 mm</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Paperclip</td>
			<td></td>
			<td></td>
			<td>Any standard 25 mm paperclip</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Needle drivers</td>
			<td>Fine Science Tools</td>
			<td>12003-15</td>
			<td></td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Tissue scissors</td>
			<td>Fine Science Tools</td>
			<td>14028-10</td>
			<td></td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Adson forceps</td>
			<td>Fine Science Tools</td>
			<td>11027-12</td>
			<td>1 x 2 teeth</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Blunt forceps</td>
			<td>Fine Science Tools</td>
			<td>11293-00</td>
			<td></td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Spinal forks</td>
			<td>Harvard Apparatus</td>
			<td>724813</td>
			<td></td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Dental cement</td>
			<td>GACD</td>
			<td>12-565 &#38; 12-568</td>
			<td></td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Silicone elastomer</td>
			<td style="width:124px;">World Precision Instruments</td>
			<td>KWIK-SIL</td>
			<td>Also avaiable through Coherent Scientific and Fisher Scientific</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Cyanoacrylate</td>
			<td>Eleco-EFD</td>
			<td>Cyanolit 201</td>
			<td></td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Microscope</td>
			<td>Zeiss</td>
			<td>7MP</td>
			<td style="width:375px;">Our microscope is in an upright configuration and is equipped with 5 NDD detectors.</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Objective lens</td>
			<td>Zeiss</td>
			<td style="width:156px;">W Plan-Apochromat 20x/1.0 DIC M27 75mm</td>
			<td>Working distance = 1.8 mm</td>
		</tr>
	</tbody>
</table>

implanting glass spinal cord windows in adult mice with experimental autoimmune encephalomyelitis

Experimental autoimmune encephalomyelitis (EAE) in adult rodents is the standard experimental model for studying autonomic demyelinating diseases such as multiple sclerosis. Here we present a low-cost and reproducible glass window implantation protocol that is suitable for intravital microscopy and studying the dynamics of spinal cord cytoarchitecture with subcellular resolution in live adult mice with EAE. Briefly, we surgically expose the vertebrae T12-L2 and construct a chamber around the exposed vertebrae using a combination of cyanoacrylate and dental cement. A laminectomy is performed from T13 to L1, and a thin layer of transparent silicone elastomer is applied to the dorsal surface of the exposed spinal cord. A modified glass cover slip is implanted over the exposed spinal cord taking care that the glass does not directly contact the spinal cord. To reduce the infiltration of inflammatory cells between the window and spinal cord, anti-inflammatory treatment is administered every 2 days (as recommended by ethics committee) for the first 10 days after implantation. EAE is induced only 2-3 weeks after the cessation of anti-inflammatory treatment.
Using this approach we successfully induced EAE in 87% of animals with implanted windows and, using Thy1-CFP-23 mice (blue axons in dorsal spinal cord), quantified axonal loss throughout EAE progression. Taken together, this protocol may be useful for studying the recruitment of various cell populations as well as their interaction dynamics, with subcellular resolution and for extended periods of time. This intravital imaging modality represents a valuable tool for developing therapeutic strategies to treat autoimmune demyelinating diseases such as EAE.

Using chronically implanted glass windows in combination with intravital two-photon microscopy and EAE models may be a useful tool to understand the cellular mechanisms underlying autoimmune demyelinating disease pathology. Using our implantation method we found that most windows were clear at the time of EAE induction, and remained clear throughout EAE progression (Figure&#160;1c). Using this approach, we found that windows remained clear for the entire postimplantation period (average ~68 d&#177;7 s.e....

Watch this Scientific Journal Video about Implanting Glass Spinal Cord Windows in Adult Mice with Experimental Autoimmune Encephalomyelitis at JoVE.com

Implanting Glass Spinal Cord Windows in Adult Mice with Experimental Autoimmune Encephalomyelitis

We describe a method for implanting and maintaining glass windows over the exposed spinal cords of adult mice for studying experimental autoimmune encephalomyelitis. These windows provide chronic optical access to the spinal cord for monitoring cell dynamics at the subcellular level in live animals using two-photon microscopy.

Experimental autoimmune encephalomyelitis (EAE) in adult rodents is the standard experimental model for studying autonomic demyelinating diseases such as ...

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Research

JoVE Journal

Medicine

7.4K Views.  Aix Marseille University. We describe a method for implanting and maintaining glass windows over the exposed spinal cords of adult mice for studying experimental autoimmune encephalomyelitis. These windows provide chronic optical access to the spinal cord for monitoring cell dynamics at the subcellular level in live animals using two-photon microscopy.

Video: Implanting Glass Spinal Cord Windows in Adult Mice with Experimental Autoimmune Encephalomyelitis

Modeling Stroke in Mice - Middle Cerebral Artery Occlusion with the Filament Model

Stroke is among the most frequent causes of death and adult disability, especially in highly developed countries. However, treatment options to date are very limited. To meet the need for novel therapeutic approaches, experimental stroke research frequently employs rodent models of focal cerebral ischaemia. Most researchers use permanent or transient occlusion of the middle cerebral artery (MCA) in mice or rats.
Proximal occlusion of the middle cerebral artery (MCA) via the intraluminal suture technique (so called filament or suture model) is probably the most frequently used model in experimental stroke research. The intraluminal MCAO model offers the advantage of inducing reproducible transient or permanent ischaemia of the MCA territory in a relatively non-invasive manner. Intraluminal approaches interrupt the blood flow of the entire territory of this artery. Filament occlusion thus arrests flow proximal to the lenticulo-striate arteries, which supply the basal ganglia. Filament occlusion of the MCA results in reproducible lesions in the cortex and striatum and can be either permanent or transient. In contrast, models inducing distal (to the branching of the lenticulo-striate arteries) MCA occlusion typically spare the striatum and primarily involve the neocortex. In addition these models do require craniectomy. In the model demonstrated in this article, a silicon coated filament is introduced into the common carotid artery and advanced along the internal carotid artery into the Circle of Willis, where it blocks the origin of the middle cerebral artery. In patients, occlusions of the middle cerebral artery are among the most common causes of ischaemic stroke. Since varying ischemic intervals can be chosen freely in this model depending on the time point of reperfusion, ischaemic lesions with varying degrees of severity can be produced. Reperfusion by removal of the occluding filament at least partially models the restoration of blood flow after spontaneous or therapeutic (tPA) lysis of a thromboembolic clot in humans.
In this video we will present the basic technique as well as the major pitfalls and confounders which may limit the predictive value of this model.

Filamentous occlusion of the Middle cerebral artery is a common model for studying ischemic stroke in mice.

Stroke is among the most frequent causes of death and adult disability, especially in highly developed countries. However, treatment options to date are ...

Acute and Chronic Tactile Sensory Testing after Spinal Cord Injury in Rats

Spinal cord injury (SCI) impairs sensory systems causing allodynia1-8. To identify cellular and molecular causes of allodynia, sensitive and valid sensory testing in rat SCI models is needed. However, until recently, no single testing approach had been validated for SCI so that standardized methods have not been implemented across labs. Additionally, available testing methods could not be implemented acutely or when severe motor impairments existed, preventing studies of the development of SCI-induced allodynia3. Here we present two validated sensory testing methods using von Frey Hair (VFH) monofilaments which quantify changes in tactile sensory thresholds after SCI4-5. One test is the well-established Up-Down test which demonstrates high sensitivity and specificity across different SCI severities when tested chronically5. The other test is a newly-developed dorsal VFH test that can be applied acutely after SCI when allodynia develops, prior to motor recovery4-5. Each VFH monofilament applies a calibrated force when touched to the skin of the hind paw until it bends. In the up-down method, alternating VFHs of higher or lower forces are used on the plantar L5 dermatome to delineate flexor withdrawal thresholds. Successively higher forces are applied until withdrawal occurs then lower force VFHs are used until withdrawal ceases. The tactile threshold reflects the force required to elicit withdrawal in 50% of the stimuli. For the new test, each VFH is applied to the dorsal L5 dermatome of the paw while the rat is supported by the examiner. The VFH stimulation occurs in ascending order of force until at least 2 of 3 applications at a given force produces paw withdrawal. Tactile sensory threshold is the lowest force to elicit withdrawal 66% of the time. Acclimation, testing and scoring procedures are described. Aberrant trials that require a retest and typical trials are defined. Animal use was approved by Ohio State University Animal Care and Use Committee.

We describe two tactile sensory testing methods for acute or chronic periods of spinal cord injury in rats. These validated procedures can detect the development and maintenance of allodynia-like sensations.

Spinal cord injury (SCI) impairs sensory systems causing allodynia1-8. To identify cellular and molecular causes of allodynia, sensitive and valid sensory ...

Controlled Cervical Laceration Injury in Mice

Use of genetically modified mice enhances our understanding of molecular mechanisms underlying several neurological disorders such as a spinal cord injury (SCI). Freehand manual control used to produce a laceration model of SCI creates inconsistent injuries often associated with a crush or contusion component and, therefore, a novel technique was developed. Our model of cervical laceration SCI has resolved inherent difficulties with the freehand method by incorporating 1) cervical vertebral stabilization by vertebral facet fixation, 2) enhanced spinal cord exposure, and 3) creation of a reproducible laceration of the spinal cord using an oscillating blade with an accuracy of &plusmn;0.01 mm in depth without associated contusion. Compared to the standard methods of creating a SCI laceration such as freehand use of a scalpel or scissors, our method has produced a consistent lesion. This method is useful for studies on axonal regeneration of corticospinal, rubrospinal, and dorsal ascending tracts.

A novel technique to create a reproducible in vivo model of cervical spinal cord laceration injury in the mouse is described. This technique is based on spine stabilization by fixation of the cervical facets and laceration of the spinal cord using an oscillating blade with an accuracy of &plusmn;0.01 mm.

Use of genetically modified mice enhances our  understanding of molecular mechanisms underlying several neurological disorders  such as a spinal cord ...

A Contusion Model of Severe Spinal Cord Injury in Rats

The translational potential of novel treatments should be investigated in severe spinal cord injury (SCI) contusion models. A detailed methodology is described to obtain a consistent model of severe SCI. Use of a stereotactic frame and computer controlled impactor allows for creation of reproducible injury. Hypothermia and urinary tract infection pose significant challenges in the post-operative period. Careful monitoring of animals with daily weight recording and bladder expression allows for early detection of post-operative complications. The functional results of this contusion model are equivalent to transection models. The contusion model can be utilized to evaluate the efficacy of both neuroprotective and neuroregenerative approaches.

A contusion model of severe spinal cord injury is described. Detailed pre-operative, operative and post-operative steps are described to obtain a consistent model.

The translational potential of novel treatments should be investigated in severe spinal cord injury (SCI) contusion models. A detailed methodology is ...

Evaluation of Respiratory Muscle Activation Using Respiratory Motor Control Assessment (RMCA) in Individuals with Chronic Spinal Cord Injury

During breathing, activation of respiratory muscles is coordinated by integrated input from the brain, brainstem, and spinal cord. When this coordination is disrupted by spinal cord injury (SCI), control of respiratory muscles innervated below the injury level is compromised1,2 leading to respiratory muscle dysfunction and pulmonary complications. These conditions are among the leading causes of death in patients with SCI3. Standard pulmonary function tests that assess respiratory motor function include spirometrical and maximum airway pressure outcomes: Forced Vital Capacity (FVC), Forced Expiratory Volume in one second (FEV1), Maximal Inspiratory Pressure (PImax) and Maximal Expiratory Pressure (PEmax)4,5. These values provide indirect measurements of respiratory muscle performance6. In clinical practice and research, a surface electromyography (sEMG) recorded from respiratory muscles can be used to assess respiratory motor function and help to diagnose neuromuscular pathology. However, variability in the sEMG amplitude inhibits efforts to develop objective and direct measures of respiratory motor function6. Based on a multi-muscle sEMG approach to characterize motor control of limb muscles7, known as the voluntary response index (VRI)8, we developed an analytical tool to characterize respiratory motor control directly from sEMG data recorded from multiple respiratory muscles during the voluntary respiratory tasks. We have termed this the Respiratory Motor Control Assessment (RMCA)9. This vector analysis method quantifies the amount and distribution of activity across muscles and presents it in the form of an index that relates the degree to which sEMG output within a test-subject resembles that from a group of healthy (non-injured) controls. The resulting index value has been shown to have high face validity, sensitivity and specificity9-11. We showed previously9 that the RMCA outcomes significantly correlate with levels of SCI and pulmonary function measures. We are presenting here the method to quantitatively compare post-spinal cord injury respiratory multi-muscle activation patterns to those of healthy individuals.

Evaluation of Respiratory Muscle Activation Using Respiratory Motor Control Assessment RMCA in Individuals with Chronic Spinal Cord Injury

The purpose of this publication is to present our original work on a multi-muscle surface electromyographic approach to quantitatively characterize respiratory muscle activation patterns in individuals with chronic spinal cord injury using vector-based analysis.

During breathing, activation of respiratory muscles is coordinated by integrated input from the brain, brainstem, and spinal cord. When this coordination ...

Controlled Cortical Impact Model for Traumatic Brain Injury

Every year over a million Americans suffer a traumatic brain injury (TBI). Combined with the incidence of TBIs worldwide, the physical, emotional, social, and economical effects are staggering. Therefore, further research into the effects of TBI and effective treatments is necessary. The controlled cortical impact (CCI) model induces traumatic brain injuries ranging from mild to severe. This method uses a rigid impactor to deliver mechanical energy to an intact dura exposed following a craniectomy. Impact is made under precise parameters at a set velocity to achieve a pre-determined deformation depth. Although other TBI models, such as weight drop and fluid percussion, exist, CCI is more accurate, easier to control, and most importantly, produces traumatic brain injuries similar to those seen in humans. However, no TBI model is currently able to reproduce pathological changes identical to those seen in human patients. The CCI model allows investigation into the short-term and long-term effects of TBI, such as neuronal death, memory deficits, and cerebral edema, as well as potential therapeutic treatments for TBI.

Traumatic brain injuries (TBIs) remain a serious health problem. Using the controlled cortical impact surgery model, research on the effects of TBI and possible treatment methods may be performed.

Every year over a million Americans suffer a traumatic brain injury (TBI).  Combined with the incidence of TBIs worldwide, the physical, emotional, ...

A Radio-telemetric System to Monitor Cardiovascular Function in Rats with Spinal Cord Transection and Embryonic Neural Stem Cell Grafts

High thoracic or cervical spinal cord injury (SCI) can lead to cardiovascular dysfunction. To monitor cardiovascular parameters, we implanted a catheter connected to a radio transmitter into the femoral artery of rats that underwent a T4 spinal cord transection with or without grafting of embryonic brainstem-derived neural stem cells expressing green fluorescent protein. Compared to other methods such as cannula insertion or tail-cuff, telemetry is advantageous to continuously monitor blood pressure and heart rate in freely moving animals. It is also capable of long term multiple data acquisitions. In spinal cord injured rats, basal cardiovascular data under unrestrained condition and autonomic dysreflexia in response to colorectal distension were successfully recorded. In addition, cardiovascular parameters before and after SCI can be compared in the same rat if a transmitter is implanted before a spinal cord transection. One limitation of the described telemetry procedure is that implantation in the femoral artery may influence the blood supply to the ipsilateral hindlimb.

We present a protocol for using a radio-telemetric system to record cardiovascular parameters in T4 spinal cord transected rats eight weeks after embryonic brainstem neural stem cell grafting into the lesion site. Telemetry is an advanced technique to accurately evaluate cardiovascular function in conscious freely moving spinal cord injured rats.

High thoracic or cervical spinal cord injury (SCI) can lead to cardiovascular dysfunction. To monitor cardiovascular parameters, we implanted a catheter ...

Synergetic Use of Neural Precursor Cells and Self-assembling Peptides in Experimental Cervical Spinal Cord Injury

Spinal cord injuries (SCI) cause serious neurological impairment and psychological, economic, and social consequences for patients and their families. Clinically, more than 50% of SCI affect the cervical spine1. As a consequence of the primary injury, a cascade of secondary mechanisms including inflammation, apoptosis, and demyelination occur finally leading to tissue scarring and development of intramedullary cavities2,3. Both represent physical and chemical barriers to cell transplantation, integration, and regeneration. Therefore, shaping the inhibitory environment and bridging cavities to create a supportive milieu for cell transplantation and regeneration is a promising therapeutic target4. Here, a contusion/compression model of cervical SCI using an aneurysm clip is described. This model is more clinically relevant than other experimental models, since complete transection or ruptures of the cord are rare. Also in comparison to the weight drop model, which in particular damage the dorsum columns, circumferential compression of the spinal cord appears advantageous. Clip closing force and duration can be adjusted to achieve different injury severity. A ring spring facilitates precise calibration and constancy of clip force. Under physiological conditions, synthetic self-assembling peptides (SAP) self-assemble into nanofibers and thus, are appealing for application in SCI5. They can be injected directly into the lesion minimizing damage to the cord. SAPs are biocompatible structures erecting scaffolds to bridge intramedullary cavities and thus, equip the damaged cord for regenerative treatments. K2(QL)6K2 (QL6) is a novel SAP introduced by Dong et al.6 In comparison to other peptides, QL6 self-assembles into &#946;-sheets at neutral pH6.14 days after SCI, after the acute stage, SAPs are injected into the center of the lesion and neural precursor cells (NPC) are injected into adjacent dorsal columns. In order to support cell survival, transplantation is combined with continuous subdural administration of growth factors by osmotic micro pumps for 7 days.

Treating cervical spinal cord injury with both self-assembling peptides (SAP) and neural precursor cells (NPC), together with growth factors, is a promising approach to promote regeneration and recovery. A contusion/compression aneurysm clip rat model of cervical SCI and combined treatment involving SAP injection and NPC transplantation is established.

Spinal cord injuries (SCI) cause serious neurological impairment and psychological, economic, and social consequences for patients and their families. ...

Neural Stem Cell Transplantation in Experimental Contusive Model of Spinal Cord Injury

Spinal cord injury is a devastating clinical condition, characterized by a complex of neurological dysfunctions. Animal models of spinal cord injury can be used both to investigate the biological responses to injury and to test potential therapies. Contusion or compression injury delivered to the surgically exposed spinal cord are the most widely used models of the pathology. In this report the experimental contusion is performed by using the Infinite Horizon (IH) Impactor device, which allows the creation of a reproducible injury animal model through definition of specific injury parameters. Stem cell transplantation is commonly considered a potentially useful strategy for curing this debilitating condition. Numerous studies have evaluated the effects of transplanting a variety of stem cells. Here we demonstrate an adapted method for spinal cord injury followed by tail vein injection of cells in CD1 mice. In short, we provide procedures for: i) cell labeling with a vital tracer, ii) pre-operative care of mice, iii) execution of a contusive spinal cord injury, and iv) intravenous administration of post mortem neural precursors. This contusion model can be utilized to evaluate the efficacy and safety of stem cell transplantation in a regenerative medicine approach.

Spinal cord injury is a traumatic condition that causes severe morbidity and high mortality. In this work we describe in detail a contusion model of spinal cord injury in mice followed by a transplantation of neural stem cells.

Spinal cord injury is a devastating clinical condition, characterized by a complex of neurological dysfunctions. Animal models of spinal cord injury can ...

Contrast Enhanced Ultrasound Imaging for Assessment of Spinal Cord Blood Flow in Experimental Spinal Cord Injury

Reduced spinal cord blood flow (SCBF) (i.e., ischemia) plays a key role in traumatic spinal cord injury (SCI) pathophysiology and is accordingly an important target for neuroprotective therapies. Although several techniques have been described to assess SCBF, they all have significant limitations. To overcome the latter, we propose the use of real-time contrast enhanced ultrasound imaging (CEU). Here we describe the application of this technique in a rat contusion model of SCI. A jugular catheter is first implanted for the repeated injection of contrast agent, a sodium chloride solution of sulphur hexafluoride encapsulated microbubbles. The spine is then stabilized with a custom-made 3D-frame and the spinal cord dura mater is exposed by a laminectomy at ThIX-ThXII. The ultrasound probe is then positioned at the posterior aspect of the dura mater (coated with ultrasound gel). To assess baseline SCBF, a single intravenous injection (400 &#181;l) of contrast agent is applied to record its passage through the intact spinal cord microvasculature. A weight-drop device is subsequently used to generate a reproducible experimental contusion model of SCI. Contrast agent is re-injected 15 min following the injury to assess post-SCI SCBF changes. CEU allows for real time and in-vivo assessment of SCBF changes following SCI. In the uninjured animal, ultrasound imaging showed uneven blood flow along the intact spinal cord. Furthermore, 15 min post-SCI, there was critical ischemia at the level of the epicenter while SCBF remained preserved in the more remote intact areas. In the regions adjacent to the epicenter (both rostral and caudal), SCBF was significantly reduced. This corresponds to the previously described &#8220;ischemic penumbra zone&#8221;. This tool is of major interest for assessing the effects of therapies aimed at limiting ischemia and the resulting tissue necrosis subsequent to SCI.

Contrast Enhanced Ultrasound imaging is a reliable in-vivo tool for quantifying spinal cord blood flow in an experimental rat spinal cord injury model. This paper contains a comprehensive protocol for application of this technique in association with a contusion model of thoracic spinal cord injury.

Reduced spinal cord blood flow (SCBF) (i.e., ischemia) plays a key role in traumatic spinal cord injury (SCI) pathophysiology and is accordingly an ...