This work was supported by NIH/NINDS Grants NS26380 and NS55976, the Christopher and Dana Reeve Foundation and the Daniel Heumann Fund for Spinal Cord Research. The Drexel University College of Medicine Spinal Cord Research Center provides support for core facilities used to complete this work.

1) Preparation for microscopic surgery
<ol>
 <li>The surgical station needs to be clean and sanitized with a dilute bleach solution prior to setting out instruments and accessory materials. Instruments will be autoclaved 1 day prior to surgery and stored in a sterile container. Turn on the Hot Bead Sterilizer (Fine Science Tools) used to remove pathogens and microbial contaminants from instruments between procedures on different animals.</li>
 <li>Plug in the thermal barrier used to maintain body temperature during the surgical procedure. Set out cotton swabs and gauze pads used to control bleeding. Prepare small (&lt; 2mm3) pieces of Gel Foam to be used for micro-hemostasis and immerse in a sterile saline solution. </li>
 <li>Place rat in an induction chamber and anesthetize with Isoflurane (5% vaporized in oxygen, flow rate 2.0 L/min). Remove animal from chamber to preparation table and shave a wide area of fur from the intended surgical field. Wipe with 70% isopropyl alcohol beginning at the center of the field and radiating outward, followed by scrubbing with Polyhydroxidine Solution (Xenodine) in the same manner. Repeat this sequence of washes 3 times.</li>
 <li>Place animal on the thermal barrier, cover nose and mouth with inhalation cone, reduce Isoflurane to 3% and flow rate to 0.2 L/min. Deliver appropriate dose of antibiotic (Ampicillin) by sub-cutaneous injection and analgesic (Buprenorphine 0.05 mg/kg) by intramuscular injection. </li>
 <li>The surgeon will scrub their hands with Xenodine solution and rinse with water. A sterile surgical gown will be fitted onto the surgeon; hands will be dried with a sterile towel and covered with sterile gloves. This procedure is followed for each surgical procedure described in this text.</li>
 <li>The lid covering the sterile instruments will be removed by an assistant and instruments will be laid out on a sterile covering by the surgeon.</li>
 </ol>
2) Transection of PN to promote degeneration
 <ol>
 <li>With the animal lying on its side make a straight line incision from the head of the femur towards the knee, keeping parallel and approximately 3mm from the femur. Cut through the superficial biceps femoris muscles along the same line as the skin incision, to expose the underlying vastus lateralis muscle. </li>
 <li>Using scissors make a small cut in the vastus lateralis near the knee and extend this up towards the head of the femur. Again with scissors gently separate the vastus lateralis until the conjoined tibial and peroneal nerve branches of the sciatic nerve are visualized (running parallel to the femur).</li>
 <li>Trace these branches towards the head of the femur until the main trunk of the sciatic nerve is identified. Just below the bifurcation of the sciatic nerve use fine forceps to carefully separate connective tissue (epineurium) surrounding the tibial nerve branch (this is the larger branch).</li>
 <li>Once isolated slip a 6-0 silk ligature around the tibial nerve and tighten. Use micro scissors to transect the nerve rostral to the ligature. Pass the suture line through the vastus lateralis muscle and tie off in a knot. This will provide a line for easy location of the cut end of the tibial nerve when the nerve is to be harvested 7-10 days later.</li>
 <li>Degeneration of the tibial nerve prior to grafting provides an environment more suitable for axonal growth than does a fresh nerve preparation.</li>
 <li>Close the vastus lateralis muscle with 6-0 silk sutures. Close the biceps femoris muscle with 6-0 silk sutures. Close the skin with Michel wound clips. Turn off the flow of Isoflurane, remove the inhalation nose cone and place animal in holding cage on thermal barrier and return to home cage once it is alert and active.</li>
 </ol>
3) Removal of PN segment
<ol>
<li>Anesthetize animal, shave and clean the intended surgical field as described above (1.3). Place the animal on thermal barrier.</li>
<li>Remove wound clips and separate previous skin incision. Cut through suture of superficial muscle to expose vastus lateralis and cut through suture in this muscle.</li>
<li>Trace the suture line to cut end of tibial nerve and remove suture from nerve. Use fine forceps to dissect epineurium from a 15 mm length of the tibial nerve and cut through this distal end. Remove the nerve segment and immerse in sterile Hanks Balanced Salt Solution.</li>
<li>If this is a heterologous graft experiment euthanize the animal by intraperitoneal injection of Euthasol. If this is an autologous graft experiment, the leg muscle and skin is closed as described in 2.6) and the animal prepared for cervical spinal cord injury, as described below.</li>
</ol>
4) Cervical hemisection injury and transplantation
Day 1
<ol>
<li>Anesthetize animal, shave and clean the intended surgical field as described above (1.3). Place the animal on thermal barrier. Administer antibiotics and analgesics as above (1.4).</li>
<li>Locate the occipital notch and the prominent T2 dorsal vertebral process and make an incision the full length between these external landmarks. Cut through the midline of the superficial trapezius muscle and separate the underlying cervical rhomboideus muscles. Keep muscles apart by inserting a spreader.</li>
<li>The deep supraspinal muscles should be cut from the dorsal vertebral processes C4, C5 and C6. Using rongeurs perform a partial laminectomy of C5 to expose the entire right side and the medial portion of the left side of the spinal cord. Use the prominent dorsal vein as a landmark for the spinal cord midline.</li>
<li>Use a micro-scalpel blade or a 30 gauge needle to pierce the dura mater and extend the incision over the C5 spinal cord. Make a small incision in the underlying subarachnoid and pia mater.</li>
<li>With a pulled glass micro-cannula begin to aspirate the dorsal, right side of the spinal cord, using gentle pressure and micro-forceps to separate the tissue and facilitate its removal. Continue through intermediate and ventral portions of the spinal cord to create a 2 mm long cavity that is complete from dorsal to ventral and from midline to the lateral meninges.</li>
<li>Place saline-soaked gel foam into the cavity to achieve hemostasis.</li>
<li>Remove the segment of peripheral nerve from the Hanks solution. Gently peel the epineurium from the proximal 3mm of the nerve segment and appose this end to the rostral wall of the hemisection lesion cavity. Secure the segment with 10-0 suture through the epineurium to the dura mater, then suture the dura closed over the implanted end of the graft.</li>
<li>Sandwich the graft between 2 pieces of silastic membrane (Biobrane) for protection and easy isolation at a later time point. Place alongside of the C6, C7 and T1 vertebral processes. Do not appose the distal nerve end to spinal cord or muscle tissue.</li>
<li>Close muscles in layers with 4-0 suture and close skin incision with wound clips.</li>
<li>Place animal in holding cage on thermal barrier and return to home cage once it is alert and active.</li>
</ol>
Day 21
<ol start="11">
<li>Anesthetize animal, shave and clean the intended surgical field as described above (1.3). Place the animal on thermal barrier and administer antibiotics and analgesics as above (1.4). Cut through superficial sutures to expose silastic membrane surrounding the nerve graft. Carefully free the length of nerve from the silastic membrane using fine forceps and spring scissors and reflect from the surgical field.</li>
<li>Perform a partial laminectomy on the right side of the C7 vertebral process. Pierce the meninges as above (4.4) and by aspiration prepare a 1 mm3 dorsal quadrant lesion cavity in the C7 spinal cord. Use saline-soaked gel foam to achieve hemostasis and then replace with gel foam soaked with chondroitinase ABC. Treat the lesion site for 15 minutes, providing fresh chondroitinase-soaked gel foam every 5 minutes.</li>
<li> Cover the external length of the nerve graft with Silastic membrane. Close superficial muscles with 4-0 sutures and the skin incision with wound clips. Place animal in holding cage on thermal barrier and return to home cage once it is alert and active.</li>
</ol>
Day 23
<ol start="14">
<li>Anesthetize animal, shave and clean the intended surgical field as described above (1.3). Place the animal on thermal barrier and administer antibiotics and analgesics as above (1.4). Cut through superficial sutures to expose silastic membrane surrounding the nerve graft. Do not disturb the graft but rather expose the C7 graft site and use a Hamilton syringe with pulled glass cannula to microinject 1&mu;l of chondroitinase ABC into the right side of the spinal cord approximately 1 mm distal to the apposition of graft to spinal cord.</li>
<li>Gently peel the epineurium from the distal 2mm of the nerve segment and appose this end to the C7 lesion cavity. Secure the segment with 10-0 suture through the epineurium to the dura mater.</li>
<li>Close superficial muscles with 4-0 sutures and the skin incision with wound clips. Place animal in holding cage on thermal barrier and return to home cage once it is alert and active.</li>
</ol>
5) Electrophysiological recording of action potential in graft and distal spinal cord
<ol>
<li>To determine if electrical activity is transmitted through axons that have regenerated into the PNG we must first expose and isolate the graft. Special care must be taken during this procedure to avoid physically damaging the graft and this is where the use of Silastic membrane after placement of the graft is important. Anesthetize animal, shave and clean the intended surgical field as described above (1.3). Place the animal on thermal barrier. </li>
<li>After opening the skin and separating superficial muscles, Silastic membrane strips are identified and carefully dissected free of the graft. </li>
<li>Using rongeurs remove the dorsal vertebral process immediately rostral and caudal to the PNG-spinal cord apposition sites. Take care not to disturb the graft too much during this approach. Make a slit in the dura mater at both levels.</li>
<li>Approximately 1cm rostral to the C5 graft-spinal cord apposition site insert a single wire (platinum-iridium mix) stimulating electrode into the spinal cord ipsilateral to the lesion, to a depth of approximately 1mm. Carefully lift the center portion of the graft onto a bipolar hooked electrode. Provide single stimulating pulses of 10&mu;A with 100&mu;sec duration and determine if there are action potentials traveling through axons that have regenerated into the graft.</li>
<li>Switch the connections and position of wires so that the hooked electrode becomes the stimulating electrode and the single wire is inserted into the spinal cord distal to the apposition of PNG to the spinal cord and attempt to record action potentials in the spinal cord.</li>
<li>At the end of the experiment provide a train of 1mA stimuli for 100&mu;sec duration at 50Hz, for a total of 1hr. Euthanize animals 1hr later, perfuse with 4% paraformaldehyde and process spinal cord tissue for the immunocytochemical detection of cFos in neurons that were activated by regenerated axons in the PNG.</li>
</ol>
6) Retrograde labeling of neurons that regenerate axons into the PNG and anterograde labeling of axons that regenerate back into the spinal cord
<ol>
<li>For animals not tested by electrophysiological measures it is possible to identify the source of axons that grow into the graft and the course of their termination in the spinal cord. Anesthetize animal, shave and clean the intended surgical field as described above (1.3). Place the animal on thermal barrier and expose the PNG as described in 6.1 above. </li>
<li>Use micro scissors to cut through the graft at its mid-point and strip back the epineurium from the cut ends of the graft. Reflect the cut ends away from each other and lay each on a separate sheet of parafilm.</li>
<li>Place a pledget of gel foam soaked with True Blue on the proximal cut end to label neurons that contributed an axon to the PNG. Place a pledget of gel foam soaked in biotinylated dextran amine (BDA) on the distal cut end to label axons that extend from the graft back into the spinal cord.</li>
<li>Euthanize animals 3-4d later, perfuse with 4% paraformaldehyde and process brain and spinal cord tissue sections for fluorescent microscopy for True Blue labeled neurons or for BDA histochemistry for labeled axons.</li>
</ol>
7) Thoracic complete transection injury and transplantation
This offers an alternative approach to the cervical hemisection injury model above. Here a more severe, bilateral injury is produced and segments of peripheral nerve are placed into the lesion cavity to span and rostral and caudal stumps of the injured spinal cord.
Day 1
<ol>
<li>Anesthetize animal, shave and clean the intended surgical field as described above (1.3). Place the animal on thermal barrier. Administer antibiotics and analgesics as above (1.4).</li>
<li>Identify T12 vertebral level at point of insertion of last rib. Make skin incision from this point rostral over vertebral processes T11, T10 and T9. Cut through superficial fascia through supraspinal muscles attached to these three processes.</li>
<li>Perform complete laminectomy of T10 vertebral process (overlying T12 spinal cord segment). Use a micro-scalpel blade or a 30 gauge needle to pierce the dura mater and extend the incision over the T12 spinal cord. Make a small incision in the underlying subarachnoid and pia mater.</li>
<li>With a pulled glass micro-cannula begin to aspirate the T12 spinal cord bilaterally, using gentle pressure and micro-forceps to separate the tissue and facilitate its removal. Continue through intermediate and ventral portions of the spinal cord to create a 3 mm long cavity that is complete dorsal to ventral and between the lateral meninges.</li>
<li>Use saline-soaked gel foam to achieve hemostasis and then replace with gel foam soaked with chondroitinase ABC. Treat the lesion site for 15 minutes, providing fresh chondroitinase-soaked gel foam every 5 minutes.</li>
<li>Remove the segment of peripheral nerve from the Hank s solution. Gently reflect the epineurium from the proximal 4mm of the nerve segment and cut a length to snugly fit between the rostral and caudal stumps of one side of the spinal cord. Place a second segment alongside the first grafted segment, apposed to the other side of the spinal cord. Suture the dura closed over the implanted grafts and cover dura with a small piece of silastic membrane.</li>
<li>Close muscles in layers with 4-0 suture and close skin incision with wound clips.</li>
<li>Place animal in holding cage on thermal barrier and return to home cage once it is alert and active.</li>
</ol>
Day 3
<ol start="9">
<li>Anesthetize animal, shave and clean the intended surgical field as described above (1.3). Place the animal on thermal barrier and administer antibiotics and analgesics as above (1.4). Cut through superficial sutures to expose silastic membrane overlying the graft site. Do not disturb the graft but rather expose the spinal cord caudal to the graft and use a Hamilton syringe with pulled glass cannula to microinject 1&mu;l of chondroitinase ABC into each side of the spinal cord approximately 1 mm distal to the apposition of graft to spinal cord.</li>
<li>Close superficial muscles with 4-0 sutures and the skin incision with wound clips. Place animal in holding cage on thermal barrier and return to home cage once it is alert and active. Perform electrophysiological stimulation and recording and tracer labeling as described in 5) and 6) above.</li>
</ol>
8) Representative Results
When this procedure is optimized the peripheral nerve segment will closely integrate with the host spinal cord. Ascending and descending spinal axons will enter the graft, grow in a relatively straight line parallel to the length of the graft and extend to the distal end of the graft at a rate of approximately 1 mm per day. Upon reaching the distal end, axons will penetrate the adjacent spinal cord if chondroitinase has been used to remove inhibitory proteoglycans from the surrounding scar tissue. Functional connectivity will be determined by electrophysiological and immunocytochemical detection of a response by spinal cord neurons to stimulation of the axons within the graft. Anatomical correlation for functional recovery will be assessed by tracing the number and length of axons that have extended beyond the nerve graft back into the spinal cord.

<ol>
	<li>Houle, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Demonstration+of+the+potential+for+chronically+injured+neurons+to+regenerate+axons+into+intraspinal+peripheral+nerve+grafts.">Demonstration of the potential for chronically injured neurons to regenerate axons into intraspinal peripheral nerve grafts.</a> Exp. Neurol. 113, 1-9 (1991).</li><li>Ye, J. H., Houle, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Treatment+of+the+chronically+injured+spinal+cord+with+neurotrophic+factors+can+promote+axonal+regeneration+from+supraspinal+neurons.">Treatment of the chronically injured spinal cord with neurotrophic factors can promote axonal regeneration from supraspinal neurons.</a> Exp. Neurol. 143, 70-81 (1997).</li><li>Houle, J. D., Tom, V., Mayes, D., Wagoner, G., Phillips, N., Silver, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Combining+an+autologous+peripheral+nerve+bridge+and+matrix+modification+by+chondroitinase+allows+robust+functional+regeneration+across+a+hemisection+lesion+of+the+adult+rat+spinal+cord.">Combining an autologous peripheral nerve bridge and matrix modification by chondroitinase allows robust functional regeneration across a hemisection lesion of the adult rat spinal cord.</a> J. Neurosci. 26, 7405-7415 (2006).</li><li>Tom, V., Houle, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intraspinal+microinjection+of+chondroitinase+ABC+following+injury+promotes+axonal+regeneration+out+of+a+peripheral+nerve+graft.">Intraspinal microinjection of chondroitinase ABC following injury promotes axonal regeneration out of a peripheral nerve graft.</a> Exp. Neurol. 211, 315-319 (2008).</li></ol>

<ol>
 <li>For reproducibility of the experiment it is important that the level of spinal cord injury be consistent from animal to animal. Therefore it is critical that the appropriate vertebral process for laminectomy be identified. Because we are using a hemisection or transection lesion model for this study there is no ambiguity about the size of the lesion or whether specific spinal tracts were injured or spared.</li>
 <li>It is necessary that the peripheral nerve be pre-degenerated before transplantation because this initiates digestion...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>10-0 silk suture</td>
<td>&nbsp;</td>
<td>ArosSurgical</td>
<td>T5A10N10</td>
<td>&nbsp;</td>
<tr>
<td>6-0 silk suture</td>
<td>&nbsp;</td>
<td>McKesson</td>
<td>2693</td>
<td>&nbsp;</td>
<tr>
<td>Ampicillin</td>
<td>&nbsp;</td>
<td>McKesson</td>
<td>483549</td>
<td>&nbsp;</td>
<tr>
<td>Antibody to cFos</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>F7799</td>
<td>&nbsp;</td>
<tr>
<td>Biotinylated dextran Amine</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>D7135</td>
<td>&nbsp;</td>
<tr>
<td>Buprenorphin (.3mg/ml)</td>
<td>&nbsp;</td>
<td>McKesson</td>
<td>12496075701</td>
<td>&nbsp;</td>
<tr>
<td>Chondroitinase ABC</td>
<td>&nbsp;</td>
<td>Associates of Cape Cod</td>
<td>100332-1A</td>
<td>&nbsp;</td>
<tr>
<td>Euthasol</td>
<td>&nbsp;</td>
<td>Webster Veterinary</td>
<td>07-805-9296</td>
<td>&nbsp;</td>
<tr>
<td>Hanks Balanced Salt Solution</td>
<td>&nbsp;</td>
<td>Cellgro</td>
<td>21-021-CV</td>
<td>&nbsp;</td>
<tr>
<td>Isoflurane</td>
<td>&nbsp;</td>
<td>Henry Schein</td>
<td>209-1966</td>
<td>&nbsp;</td>
<tr>
<td>Michel Wound Clips</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>12040-02</td>
<td>&nbsp;</td>
<tr>
<td>Neurotrace Kit</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>N7167</td>
<td>&nbsp;</td>
<tr>
<td>True Blue</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>T5891</td>
<td>&nbsp;</td>
<tr>
<td>Xenodine</td>
<td>&nbsp;</td>
<td>Webster Veterinary</td>
<td>92201</td>
<td>&nbsp;</td>
<tr>
<td>Hot Bead Sterilizer</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Forced Exercise Wheel</td>
<td>&nbsp;</td>
<td>Lafayette Instruments</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>TreadScan System</td>
<td>&nbsp;</td>
<td>Clever System</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Infinite Horizon Impact Device</td>
<td>&nbsp;</td>
<td>Precision Systems and Instrumentation</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Magnetic Stimulation Device</td>
<td>&nbsp;</td>
<td>MagStim Inc.</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

combining peripheral nerve grafting and matrix modulation to repair the injured rat spinal cord

Traumatic injury to the spinal cord (SCI) causes death of neurons, disruption of motor and sensory nerve fiber (axon) pathways and disruption of communication with the brain. One of the goals of our research is to promote axon regeneration to restore connectivity across the lesion site. To accomplish this we developed a peripheral nerve (PN) grafting technique where segments of sciatic nerve are either placed directly between the damaged ends of the spinal cord or are used to form a bridge across the lesion. There are several advantages to this approach compared to transplantation of other neural tissues; regenerating axons can be directed towards a specific target area, the number and source of regenerating axons is easily determined by tracing techniques, the graft can be used for electrophysiological experiments to measure functional recovery associated with axons in the graft, and it is possible to use an autologous nerve to reduce the possibility of graft rejection. In our lab we have performed both autologous (donor and recipient are the same animal) and heterologous (donor and recipient are different animals) grafts with comparable results. This approach has been used successfully in both acute and chronic injury situations. Regenerated axons that reach the distal end of the PN graft often fail to extend back into the spinal cord, so we use microinjections of chondroitinase to degrade inhibitory molecules associated with the scar tissue surrounding the area of SCI. At the same time we have found that providing exogenous growth and trophic molecules encourages longer distance axonal regrowth into the spinal cord. Several months after transplantation we perform a variety of anatomical, behavioral and electrophysiological tests to evaluate the recovery of function in our spinal cord injured animals. This experimental approach has been used successfully in several spinal cord injury models, at different levels of injury and in different species (mouse, rat and cat). Importantly, the peripheral nerve grafting approach is effective in promoting regeneration by acute and chronically injured neurons.

Watch this Scientific Journal Video about Combining Peripheral Nerve Grafting and Matrix Modulation to Repair the Injured Rat Spinal Cord at JoVE.com

Combining Peripheral Nerve Grafting and Matrix Modulation to Repair the Injured Rat Spinal Cord

Traumatic injury to the spinal cord disrupts communication with the brain. To restore lost connectivity we utilize a peripheral nerve graft to provide a substratum for regenerating fibers in combination with neurotrophic factors and matrix-modulating enzymes to remove inhibitory molecules to promote long distance growth.

Traumatic injury to the spinal cord (SCI) causes death of neurons, disruption of motor and sensory nerve fiber (axon) pathways and disruption of ...

combining-peripheral-nerve-grafting-matrix-modulation-to-repair

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16.3K 次观看.  Drexel University College of Medicine. 脊髓外伤，会扰乱大脑的沟通。要恢复失去的连接，我们利用了周围神经移植提供一个底层与神经营养因子和基质调节酶，再生纤维结合，以消除抑制分子，促进长途增长。

视频: Combining Peripheral Nerve Grafting and Matrix Modulation to Repair the Injured Rat Spinal Cord

Spinal Cord Electrophysiology

The neonatal mouse spinal cord is a model for studying the development of neural circuitries and locomotor movement. We demonstrate the spinal cord dissection and preparation of recording bath artificial cerebrospinal fluid used for locomotor studies. Once dissected, the spinal cord ventral nerve roots can be attached to a recording electrode to record the electrophysiologic signals of the central pattern generating circuitry within the lumbar cord.

A demonstration of the isolation of neonatal mouse spinal cord for electrophysiologic studies.

The neonatal mouse spinal cord is a model for studying the development of neural circuitries and locomotor movement. We demonstrate the spinal cord ...

Using the optokinetic response to study visual function of zebrafish

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the visual functions of larval zebrafish1-5. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adult zabrafish. Here, we introduce how to measure the OKR of adult zebrafish with our simple custom-built apparatus using a new protocol which is established in our lab. Both our apparatus and step-by-step procedure of OKR in adult zebrafish are illustrated in this video. In addition, the measurements of the larval OKR, as well as the optomotor response (OMR) test of adult zebrafish, are also demonstrated in this video. This OKR assay of adult zebrafish in our experiment may last for up to 4 hours. Such OKR test applied in adult fish will benefit to visual function investigation more efficiently when the adult fish vision system is manipulated.
Su-Qi Zou and Wu Yin contributed equally to this paper.

Optokinetic response has been widely used to assess the visual functions of larval zebrafish. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adults1-5. Here, we introduce how to measure the OKR of adult zebrafish using a new protocol which is established in our lab.

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the ...

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter implantation. The technical limitation of this surgical technique lies in the finesse of the hands. The injection is rapid, especially for a trained experimenter, and since tissue disruption with this technique is minimal, repeated injections are possible; moreover immune reaction to foreign tools (e.g. catheter) does not occur, thereby giving a better and more specific read out of spinal cord modulation. Since the application of the substance is largely limited to the target region of the spinal cord, drugs do not need to be applied in large dosages, and more importantly unwanted effects on other tissue, as observed with a systemic delivery, could be circumvented1,2. Moreover, we combine this technique with in vivo transfection of nucleic acid with the help of polyethylenimine (PEI) reagent3, which provides tremendous versatility for studying spinal functions via delivery of pharmacological agents as well as gene, RNA, and protein modulators.

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a simple and rapid technique of intrathecal needle puncture for a localized transfection of siRNA in the lumbar spinal cord in mouse under short lasting light anesthesia.

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter ...

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix biology, there are still major gaps related to how the matrix is formed, how physiological and pathological remodeling differ, and most importantly how matrix dynamics might be manipulated to promote healing and inhibit fibrosis. There is currently no treatment option for controlling, preventing, or reversing cardiac fibrosis. Part of the reason is likely the sheer complexity of cardiac scar formation, such as occurs after myocardial infarction to immediately replace dead or dying cardiomyocytes. The extracellular matrix itself participates in remodeling by activating resident cells and also by helping to guide infiltrating cells to the defunct lesion. The matrix is also a storage locker of sorts for matricellular proteins that are crucial to normal matrix turnover, as well as fibrotic signaling. The matrix has additionally been demonstrated to play an electromechanical role in cardiac tissue. Most techniques for assessing fibrosis are not qualitative in nature, but rather provide quantitative results that are useful for comparing two groups but that do not provide information related to the underlying matrix structure. Highlighted here is a technique for visualizing cardiac matrix ultrastructure. Scanning electron microscopy of decellularized heart tissue reveals striking differences in structure that might otherwise be missed using traditional quantitative research methods.

Shown here is a method for visualizing extracellular matrix ultrastructure in decellularized cardiac tissues.

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix ...

Cystometric and External Urethral Sphincter Measurements in Awake Rats with Implanted Catheter and Electrodes Allowing for Repeated Measurements

Lower urinary tract function is mainly assessed by means of cystometric bladder function analysis in rodents. Conventional cystometries are usually performed as terminal analysis under urethane anesthesia. It is well known that anesthetic drugs can influence bladder function. Hence, the aim of this technique is to perform cystometric measurements of the urinary bladder and external urethral sphincter in lightly restrained awake rats. For this purpose, a bladder catheter is implanted into the bladder dome. Subsequently, two electrodes are implanted bilateral to the external urethral sphincter and a ground electrode is sutured to a non-responsive skeletal muscle. The bladder catheter and the three electrodes are finally tunneled subcutaneously to the neck region and affixed to a harness. With this technique, the lower urinary tract can be measured at multiple time points in the same animal to assess lower urinary tract function. The main application of this technique is the follow-up of simultaneous urinary bladder and external urethral sphincter function in awake healthy rats and after induction of a disease or injury. Moreover, subsequent lower urinary tract monitoring can be performed during evaluation of the disease/injury and to monitor treatment efficacy.

This protocol first describes the surgical procedure of the permanent implantation of a urinary bladder catheter combined with external urethral sphincter electrodes, and second, the measurement of the function of the urinary bladder and external urethral sphincter in implanted awake animals.

Lower urinary tract function is mainly assessed by means of cystometric bladder function analysis in rodents. Conventional cystometries are usually ...

Sensitive Measurement of Mitophagy by Flow Cytometry Using the pH-dependent Fluorescent Reporter mt-Keima

Mitophagy is a process of selective removal of damaged or unnecessary mitochondria using autophagy machinery. Close links have been found between defective mitophagy and various human diseases, including neurodegenerative diseases, cancer, and metabolic diseases. In addition, recent studies have shown that mitophagy is involved in normal cellular processes, such as differentiation and development. However, the precise role of and molecular mechanisms underlying mitophagy require further study. Therefore, it is critical to develop a robust and convenient method for measuring changes in mitophagy activity. Here, we describe a detailed protocol for quantitatively assessing mitophagy activity through flow cytometry using the mitochondria-targeted fluorescent protein Keima (mt-Keima). This flow cytometry assay can analyze mitophagy activity more rapidly and sensitively than conventional microscopy- or immunoblotting-based methods. This protocol can be applied to analyze mitophagy activity in various cell types.

Mitophagy, the selective degradation of mitochondria, has been implicated in mitochondrial homeostasis and is deregulated in various human diseases. However, convenient experimental methods for measuring mitophagy activity are very limited. Here, we provide a sensitive assay for measuring mitophagy activity using flow cytometry.

Mitophagy is a process of selective removal of damaged or unnecessary mitochondria using autophagy machinery. Close links have been found between ...

Adipo-Clear: A Tissue Clearing Method for Three-Dimensional Imaging of Adipose Tissue

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte precursors, immune cells, fibroblasts, blood vessels, and nerve projections. Although the molecular control of cell type specification and how these cells interact have been increasingly delineated, a more comprehensive understanding of these adipose-resident cells can be achieved by visualizing their distribution and architecture throughout the whole tissue. Existing immunohistochemistry and immunofluorescence approaches to analyze adipose histology rely on thin paraffin-embedded sections. However, thin sections capture only a small portion of tissue; as a result, the conclusions can be biased by what portion of tissue is analyzed. We have therefore developed an adipose tissue clearing technique, Adipo-Clear, to permit comprehensive three-dimensional visualization of molecular and cellular patterns in whole adipose tissues. Adipo-Clear was adapted from iDISCO/iDISCO+, with specific modifications made to completely remove the lipid stored in the tissue while preserving native tissue morphology. In combination with light-sheet fluorescence microscopy, we demonstrate here the use of the Adipo-Clear method to obtain high-resolution volumetric images&#160;of an entire adipose tissue.

Due to the high lipid content, adipose tissue has been challenging to visualize using traditional histological methods. Adipo-Clear is a tissue clearing technique that allows robust labeling and high-resolution volumetric fluorescent imaging of adipose tissue. Here, we describe the methods for sample preparation, pretreatment, staining, clearing, and mounting for imaging.

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte ...

Assessment of Vascular Tone Responsiveness using Isolated Mesenteric Arteries with a Focus on Modulation by Perivascular Adipose Tissues

Altered vascular tone responsiveness to pathophysiological stimuli contributes to the development of a wide range of cardiovascular and metabolic diseases. Endothelial dysfunction represents a major culprit for the reduced vasodilatation and enhanced vasoconstriction of arteries. Adipose (fat) tissues surrounding the arteries play important roles in the regulation of endothelium-dependent relaxation and/or contraction of the vascular smooth muscle cells. The cross-talks between the endothelium and perivascular adipose tissues can be assessed ex vivo using mounted blood vessels by a wire myography system. However, optimal settings should be established for arteries derived from animals of different species, ages, genetic backgrounds and/or pathophysiological conditions.

The protocol describes the use of wire myography to evaluate the transmural isometric tension of mesenteric arteries isolated from mice, with special consideration of the modulation by factors released from endothelial cells and perivascular adipose tissues.

Altered vascular tone responsiveness to pathophysiological stimuli contributes to the development of a wide range of cardiovascular and metabolic ...

A Single Cell Dissociation Approach for Molecular Analysis of Urinary Bladder in the Mouse Following Spinal Cord Injury

We describe the implementation of spinal cord injury in mice to elicit detrusor-sphincter dyssynergia, a functional bladder outlet obstruction, and subsequent bladder wall remodeling. To facilitate assessment of the cellular composition of the bladder wall in non-injured control and spinal cord injured mice, we developed an optimized dissociation protocol that supports high cell viability and enables the detection of discrete subpopulations by flow cytometry.
Spinal cord injury is created by complete transection of the thoracic spinal cord. At the time of tissue harvest, the animal is perfused with phosphate-buffered saline under deep anesthesia and bladders are harvested into Tyrode&#8217;s buffer. Tissues are minced prior to incubation in digestion buffer that has been optimized based on the collagen content of mouse bladder as determined by interrogation of publicly available gene expression databases. Following generation of a single cell suspension, material is analyzed by flow cytometry for assessment of cell viability, cell number and specific subpopulations. We demonstrate that the method yields cell populations with greater than 90% viability, and robust representation of cells of mesenchymal and epithelial origin. This method will enable accurate downstream analysis of discrete cell types in mouse bladder and potentially other organs.

The goal of this protocol is to apply an optimized tissue dissociation protocol to a mouse model of spinal cord injury and validate the approach for single cell analysis by flow cytometry.

We describe the implementation of spinal cord injury in mice to elicit detrusor-sphincter dyssynergia, a functional bladder outlet obstruction, and ...

An Adipocyte Cell Culture Model to Study the Impact of Protein and Micro-RNA Modulation on Adipocyte Function

Alteration of adipocyte function contributes to the pathogenesis of metabolic diseases including Type 2 diabetes and insulin resistance. This highlights the need to better understand the molecular mechanism involved in adipocyte dysfunction to develop new therapies against obesity-related diseases. Modulating the expression of proteins and micro-RNAs in adipocytes remains highly challenging. This paper describes a protocol to differentiate murine fibroblasts into mature adipocytes and to modulate the expression of proteins and micro-RNAs in mature adipocytes through reverse-transfection using small-interfering RNA (siRNA) and micro-RNA mimicking (miR mimic) oligonucleotides. This reverse-transfection protocol involves the incubation of the transfection reagent and the oligonucleotides to form a complex in the cell culture plate to which the mature adipocytes are added. The adipocytes are then allowed to reattach to the adherent plate surface in the presence of the oligonucleotides/transfection reagent complex. Functional analyses such as the study of insulin signaling, glucose uptake, lipogenesis, and lipolysis can be performed on the transfected 3T3-L1 mature adipocytes to study the impact of protein or micro-RNA manipulation on adipocyte function.

Presented here is a protocol to deliver oligonucleotides such as small-interfering RNA (siRNA), micro-RNA mimics (miRs), or anti-micro-RNA (anti-miR) into mature adipocytes to modulate protein and micro-RNA expression.

Alteration of adipocyte function contributes to the pathogenesis of metabolic diseases including Type 2 diabetes and insulin resistance. This highlights ...