PDK is supported by a CIHR operating grant (MOP 86523)

1. Surgical Technique
<ol>
<li>Experiments should be carried out using aseptic technique and following the animal use protocols of your specific institution. Instruments and materials (solutions, test substances, tracers, needles, etc.) coming into contact with living tissue must be sterile to prevent infection and adverse impacts on animal welfare and potential negative impacts on the study.</li>
</ol>
2. Anesthesia
<ol>
<li>Rats will be anaesthetized using a veterinary isoflurane vaporizer system. Use medical grade oxygen at a rate of 0.8 L/min to vaporize the isoflurane gas. Place the animal in the attached anesthesia box and dial in an isoflurane concentration of 4% until the breathing has slowed and the animal is sedate.</li>
<li>Next, switch the gas flow to the gas mask attachment for the stereotaxic frame and place the animal in the stereotaxic apparatus. Turn the isoflurane concentration down to 2% and monitor anesthesia. Larger animals (&gt;300g) may require a higher concentration of isoflurane. Anesthesia should be monitored during surgery and isoflurane dosage adjusted accordingly. Depth and rate of breathing should be constantly evaluated, and toe pinch evaluation (every 5 min) for the absence of deep pain should be performed.</li>
<li>Once the surgery is complete, turn off the isoflurane and allow the animal to breath oxygen for several minutes prior to removal from the stereotaxic device. Body temperature should be maintained by covering the animal with a surgical blanket and/or using a regulated heating blanket during surgery.</li>
</ol>
3. Syringe Preparation for Intraocular Injections
<ol>
<li>First, assemble the syringe system for performing the intraocular injections. A 10 &mu;L RN Gastight 1701 Hamilton Syringe is used for injection. Remove any needle or RN nut present on the end of the syringe.</li>
<li>Insert the PFA ferrule into the PEEK cup ferrule of the compression fitting. Then insert the complete fitting into the end of the syringe and loosely screw the RN nut over top.</li>
<li>Insert one end of the 1/16 inch PEEK tubing into the compression fitting, through the opening in the RN nut. Make sure the PEEK tubing is fully seated. Tighten the RN nut to compress the ferrule, sealing the PEEK tubing.</li>
<li>Perform Step 3.2 on both ends of the Dual RN glass coupler. Attach the free end of the PEEK tubing to one end of the Dual RN glass coupler by tightening the RN nut.</li>
<li>A pulled glass micropipette will form the needle and barrel for the injection system. Use 1.5 mm O.D. thick walled glass capillary tubing to fabricate the glass pipettes, and insert the glass pipette into the free end of the Dual RN glass coupler. Tighten the RN nut to fix the pipette in place.</li>
<li>The fine tip of pulled glass micropipettes is typically too small for performing intraocular injections. In order to create a tip of appropriate diameter, break the tip of the glass pipette under visual guidance, using a surgical microscope. The frosted end of a glass specimen slide is well suited to break the pipette. Hold the pipette at an angle and rub the tip along the glass frosting to create a break. Ideally, the final tip should be slightly bevelled. It may take a few attempts to produce a good tip, so pull multiple pipettes prior to use.</li>
<li>The injection system is hydraulic. Therefore, the syringe, PEEK tubing linker, Dual RN glass coupler and the glass micropipette must be filled with mineral oil prior to use. Using the Priming kit from Hamilton Syringe Co. (PRMKIT), fill the priming syringe with mineral oil. Use a large diameter needle to allow easy passage of the viscous oil. Replace the needle with the Hamilton 90030 needle that fits inside the barrel of the 10 &mu;L Hamilton syringe. Next, place a rubber septum over the Hamilton 90030 needle, in order to provide a seal while injecting the mineral oil.</li>
<li>Insert the Hamilton 90030 needle of the priming syringe into the back of the barrel of the 10 &mu;L Hamilton syringe of the injection system. Press the septum tightly against the end of the syringe to create a seal.</li>
<li>Slowly depress the plunger. You will see the mineral oil pass through the glass elements of the injection system, and finally filling the glass pipette. Push all of the oil through the injection system in order to remove any air bubbles along the tract. If the filling syringe is running out of mineral oil, slowly withdraw the needle while constantly dispensing oil to prevent air bubble formation. Refill the priming syringe and inject again.</li>
<li>When the entire system is filled with mineral oil and bubble free, insert the original plunger of the 10 &mu;L Hamilton syringe. Depress the plunger until the end of its travel in order to remove additional mineral oil from the system. Wipe the glass micropipette clean and the end of the syringe clean with 70% ethanol.</li>
<li>Withdraw the plunger of the syringe to the 2 &mu;L mark on the barrel. The end of the glass micropipette will fill with air providing a buffer zone between the mineral oil and the desired fluid to be injected. The pipette barrel can now be filled by placing the end of the micropipette in the desired solution and withdrawing the plunger of the Hamilton syringe. Do not inject more than 4-5 &mu;L of solution into an adult rat eye.</li>
</ol>
4. Intraocular Injection Procedure: Targeting the Retina Globally
<ol>
<li>With the animal anesthetized and secured in the stereotaxic frame (as described in Section 2), use Alcaine eye drops to topically anesthetize the cornea. Open the eyelids with your fingers and place one drop of the anesthetic solution on the surface of the cornea.</li>
<li>Injection requires two people: one to insert the glass pipette into the vitreous chamber and maintain pipette position, and another to depress the plunger on the syringe which delivers the desired solution.</li>
<li>Under a surgical microscope, grip the glass micropipette and Dual RN glass coupler with your fingers. Spread the eyelids with the fingers of your free hand, forming a "V" shape opposite the injection site. By applying traction to the eyelids, the eye will be elevated out of the orbit, exposing the posterior surface behind the limbus. By creating a "V" shape with the fingers opposite the injection site, a cradle is formed for the eye providing stability when injecting.</li>
<li>Gently insert the tip of the glass micropipette through the conjunctiva, in an area devoid of blood vessels, and through the sclera, at a downward angle. You should feel a small "pop" when the pipette penetrates the sclera. Inserting the pipette at a slightly downward angle reduces the chance of hitting the lens when inserting the needle.</li>
<li>The person holding the syringe should now inject the 4 &mu;L of solution by depressing the plunger to the 2 &mu;L mark on the syringe. The injection should take approximately one to two seconds in total. Injecting at a very slow rate >3 seconds reduces the initial spread of the solution through the vitreous chamber. You will be able to see the injected solution flushing through the vitreous chamber under the microscope, thereby verifying the success of the procedure.</li>
<li>Hold the micropipette steady for approximately 5 seconds and then withdraw, in the same direction as the needle was inserted. The conjunctiva will flap back over the small opening, helping to seal the scleral puncture.</li>
<li>Cover the surface of the cornea with ophthalmic eye ointment (Tears Naturale P.M.) in order to prevent corneal drying during recovery, and return the animal to a recovery cage.</li>
<li>Post- surgical analgesics should be administered according to the guidelines of your animal care authorities, and animals should be carefully monitored after surgery.</li>
</ol>
5. Selectively Targeting Retinal Ganglion Cells from the Optic Nerve Stump
<ol>
<li>Applying tracers, drugs, plasmids, siRNAs or viral vectors to optic nerve stump, following axotomy, directly targets retinal ganglion cells through retrograde transport. This is accomplished by first performing an optic nerve transection according to the "Optic Nerve Transection Protocol"22.</li>
<li>There are two main methods to accomplish this procedure: using gelfoam, or direct injection into the optic nerve.</li>
<li>The gelfoam method is similar to retrograde tracing, however when delivering experimental treatments at the optic nerve it is desirable to pre-label the retinal ganglion cells by injecting retrograde tracers into the superior colliculus 1 week prior to axotomy. This is important for cell tracing because, in some instances neuronal tracers can interfere with the retrograde transport of experimental substances or vice versa.</li>
<li>Immediately after the optic nerve is cut, a small piece of gelfoam soaked in the experimental solution is placed over the transected optic nerve stump. The orbital contents are then returned to their previous location. When the eye is returned to a neutral position it is necessary to use forceps to push the piece of gelfoam down into the orbit, ensuring that it remains over the end of the optic nerve.</li>
<li>The injection technique is more effective for the delivery of substances that must gain access to the cytoplasm of the axon in order to be retrogradely transported by axoplasmic flow. This includes siRNAs and plasmids, and in some cases viral vectors. A 5-10 &mu;L Hamilton syringe is used to inject the desired solution into the transected optic nerve.</li>
<li>Grip the edge of the optic nerve with fine tip Dumont forceps. Insert the end of the needle into the optic nerve, parallel with the nerve, until the bevel is no longer visible. The inserted needle will be encased on all sides by the nerve. A needle with a short bevel is desirable.</li>
<li>Once the needle is inserted into the nerve, gently squeeze the sides of the nerve with the fine tip forceps and inject one quarter of the solution while rotating the syringe one quarter turn clockwise or counter clockwise.</li>
<li>Continue repeating Step 5.7 until you have completed a full rotation of the needle and injected the entire contents of the syringe. For best results inject a total of 10 &mu;L of solution using a gastight 10 &mu;L Hamilton syringe.</li>
<li>Remove the tip of the syringe from the nerve. Leave the excess injected fluid that has refluxed from the nerve within the orbit as this will form an additional pool for uptake by the ends of the axons.</li>
<li>Return the orbital contents to their original position, close the wound, and allow the animal to recover.</li>
<li>Post- surgical analgesics should be administered according to the guidelines of your animal care authorities, and animals should be carefully monitored after surgery.</li>
</ol>
6. Representative Results:
Intraocular injections target all cells in the retina globally, and work well for the delivery substances to injured RGCs since ganglion cells are located in the innermost neuronal layer of the retina, adjacent to the vitreous chamber. RGCs can also be directly targeted by delivering substances to the transected optic nerve stump. Fluorogold retrogradely labeled RGCs can be targeted using peptides, drugs, vectors, plasmids, or siRNAs from the nerve stump as illustrated in Figure 1. Figure 1A-C demonstrates the localization of a Cy3 labeled peptide in the somata of RGCs that were pre-labeled by injecting Fluorogold into the superior colliculus. Cy3 labeled peptides were injected into the optic nerve immediately after axotomy, and the retina was imaged alive in order to prevent the peptides from leaching out of the tissue during fixation. Figure 1D-F demonstrates the localization of Cy3 labeled siRNAs and the retrograde tracer Fluorogold in axotomized RGCs. Cy3 labeled siRNAs were injected into the optic nerve stump immediately after axotomy, and the retina was imaged alive.
<img src="/files/ftp_upload/2261/2261fig1.jpg" alt="Figure 1" /> 
Figure 1. Epifluorescence micrographs of RGCs in flatmounted retinas at 1 day after axotomy and injection of either labeled peptides or siRNAs into the optic nerve stump. (A) Fluorogold retrograde labeling in axotomized RGCs at 1 day postaxotomy (B) Cy3 fluorescence in RGCs following retrograde transport of labeled peptides, 1 day after axotomy and peptide injection into the optic nerve. (C) Overlay of (A) and (B) demonstrating the selective localization of Cy3 labeled peptides in Fluorogold labeled RGCs. (D-F) The cell bodies of Fluorogold labeled RGCs (D) are also filled with Cy3 labeled siRNAs (E) injected into the optic nerve stump 24 hours earlier, as shown in the overlay (F). Scale bar in A-C is 50 &mu;m. Scale bar in D-F is 20 &mu;m.

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D., Bahr, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Growth+and+guidance+cues+for+regenerating+axons:+where+have+they+gone.">Growth and guidance cues for regenerating axons: where have they gone.</a> J Neurobiol. 59, 162-180 (2004).</li><li>Weishaupt, J. H., Bahr, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Degeneration+of+axotomized+retinal+ganglion+cells+as+a+model+for+neuronal+apoptosis+in+the+central+nervous+system+-+molecular+death+and+survival+pathways.">Degeneration of axotomized retinal ganglion cells as a model for neuronal apoptosis in the central nervous system - molecular death and survival pathways.</a> Restor. Neurol. 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No conflicts of interest declared.

Optic nerve transection is a highly reproducible model of adult CNS neuron apoptosis. The experimental manipulations demonstrated in this manuscript permit the study of the mechanisms of RGC apoptosis after injury.
Intraocular injections are useful for global targeting of the retina. This procedure requires some practice, as it is critical not to injure the lens with the tip of the glass pipette. Lens damage has been shown to cause the release of growth factors, altering cell survival and rege...

A correction to the article Methods for Experimental Manipulations after Optic Nerve Transection in the Mammalian CNS was made.

Erratum: Methods for Experimental Manipulations after Optic Nerve Transection in the Mammalian CNS

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		<th>Catalogue Number</th>
		<th>Comment</th>
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<td>Stereotaxic Frame</td>
<td>&nbsp;</td>
<td>Stoelting, Kopf, WPI</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Rat Gas Mask</td>
<td>&nbsp;</td>
<td>Stoelting, Kopf, WPI</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Anesthesia System</td>
<td>&nbsp;</td>
<td>VetEquip</td>
<td>901806</td>
<td>&nbsp;</td>
<tr>
<td>Isoflurane (PrAErrane)</td>
<td>&nbsp;</td>
<td>Baxter Corp</td>
<td>DIN 02225875</td>
<td>&nbsp;</td>
<tr>
<td>Surgical Microscope</td>
<td>&nbsp;</td>
<td>WPI, Zeiss, Leica</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Alcaine Eye Drops</td>
<td>&nbsp;</td>
<td>Alcon</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Tears Naturale P.M.</td>
<td>&nbsp;</td>
<td>Alcon</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Fine tip Dumont forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11252-00</td>
<td>&nbsp;</td>
<tr>
<td>10 &mu;l Hamilton Syringe (1701RN; 26s/2&#x201D;/2)</td>
<td>&nbsp;</td>
<td>Hamilton Syringe Co.</td>
<td>80030</td>
<td>&nbsp;</td>
<tr>
<td>1/16 inch Compression Fittings</td>
<td>&nbsp;</td>
<td>Hamilton Syringe Co.</td>
<td>55751-01</td>
<td>&nbsp;</td>
<tr>
<td>1/16 inch OD, 0.010 inch ID, PEEK Tubing</td>
<td>&nbsp;</td>
<td>Supelco, Bellefonte, PA</td>
<td>Z226661</td>
<td>&nbsp;</td>
<tr>
<td>Dual RN Glass Coupler</td>
<td>&nbsp;</td>
<td>Hamilton Syringe Co.</td>
<td>55752-01</td>
<td>&nbsp;</td>
<tr>
<td>Mineral Oil Priming Kit: includes syringe, needles, rubber septa</td>
<td>&nbsp;</td>
<td>Hamilton Syringe Co.</td>
<td>PRMKIT</td>
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methods for experimental manipulations after optic nerve transection in the mammalian cns

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. The optic nerve can be accessed within the orbit of the eye and completely transected (axotomized), cutting the axons of the entire RGC population. Optic nerve transection is a reproducible model of apoptotic neuronal cell death in the adult CNS 1-4. This model is particularly attractive because the vitreous chamber of the eye acts as a capsule for drug delivery to the retina, permitting experimental manipulations via intraocular injections. The diffusion of chemicals through the vitreous fluid ensures that they act upon the entire RGC population. Viral vectors, plasmids or short interfering RNAs (siRNAs) can also be delivered to the vitreous chamber in order to infect or transfect retinal cells 5-12. The high tropism of Adeno-Associated Virus (AAV) vectors is beneficial to target RGCs, with an infection rate approaching 90% of cells near the injection site 6, 7, 13-15. Moreover, RGCs can be selectively transfected by applying siRNAs, plasmids, or viral vectors to the cut end of the optic nerve 16-19 or injecting vectors into their target the superior colliculus 10. This allows researchers to study apoptotic mechanisms in the injured neuronal population without confounding effects on other bystander neurons or surrounding glia. RGC apoptosis has a characteristic time-course whereby cell death is delayed 3-4 days postaxotomy, after which the cells rapidly degenerate. This provides a window for experimental manipulations directed against pathways involved in apoptosis. Manipulations that directly target RGCs from the transected optic nerve stump are performed at the time of axotomy, immediately after cutting the nerve. In contrast, when substances are delivered via an intraocular route, they can be injected prior to surgery or within the first 3 days after surgery, preceding the initiation of apoptosis in axotomized RGCs. In the present article, we demonstrate several methods for experimental manipulations after optic nerve transection.

Watch this Scientific Journal Video about Methods for Experimental Manipulations after Optic Nerve Transection in the Mammalian CNS at JoVE.com

Methods for Experimental Manipulations after Optic Nerve Transection in the Mammalian CNS

Optic Nerve transection is a widely used model of adult CNS injury. This model is ideal for performing a number of experimental manipulations that target the retina globally or directly target the injured neuronal population of retinal ganglion cells.

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. The optic nerve can be ...

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Neuroscience

13.4K 조회수.  University of Toronto. 시신경의 절개면은 성인 CNS 손상의 널리 사용되는 모델입니다. 이 모델은 전 세계적으로 망막을 대상에 직접 망막 신경절 세포의 부상의 연결 인구를 대상으로 실험 조작의 번호를 수행하는 데 이상적입니다.

비디오: Methods for Experimental Manipulations after Optic Nerve Transection in the Mammalian CNS

Optic Nerve Transection: A Model of Adult Neuron Apoptosis in the Central Nervous System

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. 
The optic nerve can be accessed within the orbit of the eye and completely transected (axotomized), cutting the axons of the entire RGC population. Optic nerve 
transection is a reproducible model of apoptotic neuronal cell death in the adult CNS 1-4. This model is particularly attractive because the vitreous
 chamber of the eye acts as a capsule for drug delivery to the retina, permitting experimental manipulations via intraocular injections. The diffusion of chemicals 
through the vitreous fluid ensures that they act upon the entire RGC population. Moreover, RGCs can be selectively transfected by applying short interfering RNAs 
(siRNAs), plasmids, or viral vectors to the cut end of the optic nerve 5-7 or injecting vectors into their target, the superior colliculus 8. 
This allows researchers to study apoptotic mechanisms in the desired neuronal population without confounding effects on other bystander neurons or surrounding glia. 
An additional benefit is the ease and accuracy with which cell survival can be quantified after injury. The retina is a flat, layered tissue and RGCs are localized in 
the innermost layer, the ganglion cell layer. The survival of RGCs can be tracked over time by applying a fluorescent tracer (3% Fluorogold) to the cut end of the 
optic nerve at the time of axotomy, or by injecting the tracer into the superior colliculus (RGC target) one week prior to axotomy. The tracer is retrogradely transported, labeling 
the entire RGC population. Because the ganglion cell layer is a monolayer (one cell thick), RGC densities can be quantified in flat-mounted tissue, without the need 
for stereology. Optic nerve transection leads to the apoptotic death of 90% of injured RGCs within 14 days postaxotomy 9-11. RGC apoptosis has a 
characteristic time-course whereby cell death is delayed 3-4 days postaxotomy, after which the cells rapidly degenerate. This provides a time window for 
experimental manipulations directed against pathways involved in apoptosis.

Optic Nerve transection is a widely used model of adult CNS injury. Ninety percent of retinal ganglion cells (RGCs) whose axons are completely transected (axotomy) die within 14 days after axotomy. This model is easily amenable to experimental manipulations and highly reproducible.

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. 
The optic nerve can be ...

An Optic Nerve Crush Injury Murine Model to Study Retinal Ganglion Cell Survival

Injury to the optic nerve can lead to axonal degeneration, followed by a gradual death of retinal ganglion cells (RGCs), which results in irreversible vision loss. Examples of such diseases in human include traumatic optic neuropathy and optic nerve degeneration in glaucoma. It is characterized by typical changes in the optic nerve head, progressive optic nerve degeneration, and loss of retinal ganglion cells, if uncontrolled, leading to vision loss and blindness.
The optic nerve crush (ONC) injury mouse model is an important experimental disease model for traumatic optic neuropathy, glaucoma, etc. In this model, the crush injury to the optic nerve leads to gradual retinal ganglion cells apoptosis. This disease model can be used to study the general processes and mechanisms of neuronal death and survival, which is essential for the development of therapeutic measures. In addition, pharmacological and molecular approaches can be used in this model to identify and test potential therapeutic reagents to treat different types of optic neuropathy.
Here, we provide a step by step demonstration of (I) Baseline retrograde labeling of retinal ganglion cells (RGCs) at day 1, (II) Optic nerve crush injury at day 4, (III) Harvest the retinae and analyze RGC survival at day 11, and (IV) Representative result.

This protocol shows how to retrogradely label retinal ganglion cells, and how to subsequently make an optic nerve crush injury in order to analyze retinal ganglion cell survival and apoptosis. It is an experimental disease model for different types of optic neuropathy, including glaucoma.

Injury to the optic nerve can lead to axonal degeneration, followed by a gradual death of retinal ganglion cells (RGCs), which results in irreversible ...

Transplantation of Olfactory Ensheathing Cells to Evaluate Functional Recovery after Peripheral Nerve Injury

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory system is characterized by its ability to give rise to new neurons even in adult animals. This particular ability is partly due to the presence of OECs which create a favorable microenvironment for neurogenesis. This property of OECs has been used for cellular transplantation such as in spinal cord injury models. Although the peripheral nervous system has a greater capacity to regenerate after nerve injury than the central nervous system, complete sections induce misrouting during axonal regrowth in particular after facial of laryngeal nerve transection. Specifically, full sectioning of the recurrent laryngeal nerve (RLN) induces aberrant axonal regrowth resulting in synkinesis of the vocal cords. In this specific model, we showed that OECs transplantation efficiently increases axonal regrowth.
OECs are constituted of several subpopulations present in both the olfactory mucosa (OM-OECs) and the olfactory bulbs (OB-OECs). We present here a model of cellular transplantation based on the use of these different subpopulations of OECs in a RLN injury model. Using this paradigm, primary cultures of OB-OECs and OM-OECs were transplanted in Matrigel after section and anastomosis of the RLN. Two months after surgery, we evaluated transplanted animals by complementary analyses based on videolaryngoscopy, electromyography (EMG), and histological studies. First, videolaryngoscopy allowed us to evaluate laryngeal functions, in particular muscular cocontractions phenomena. Then, EMG analyses demonstrated richness and synchronization of muscular activities. Finally, histological studies based on toluidine blue staining allowed the quantification of the number and profile of myelinated fibers.
All together, we describe here how to isolate, culture, identify and transplant OECs from OM and OB after RLN section-anastomosis and how to evaluate and analyze the efficiency of these transplanted cells on axonal regrowth and laryngeal functions.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth of the primary olfactory neurons. This specific property can be used for cellular transplantation. We present here a model of cellular transplantation based on the use of OECs in a laryngeal nerve injury model.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory ...

Motor Nerve Transection and Time-lapse Imaging of Glial Cell Behaviors in Live Zebrafish

The nervous system is often described as a hard-wired component of the body even though it is a considerably fluid organ system that reacts to external stimuli in a consistent, stereotyped manner, while maintaining incredible flexibility and plasticity. Unlike the central nervous system (CNS), the peripheral nervous system (PNS) is capable of significant repair, but we have only just begun to understand the cellular and molecular mechanisms that govern this phenomenon. Using zebrafish as a model system, we have the unprecedented opportunity to couple regenerative studies with in vivo imaging and genetic manipulation. Peripheral nerves are composed of axons surrounded by layers of glia and connective tissue. Axons are ensheathed by myelinating or non-myelinating Schwann cells, which are in turn wrapped into a fascicle by a cellular sheath called the perineurium. Following an injury, adult peripheral nerves have the remarkable capacity to remove damaged axonal debris and re-innervate targets. To investigate the roles of all peripheral glia in PNS regeneration, we describe here an axon transection assay that uses a commercially available nitrogen-pumped dye laser to axotomize motor nerves in live transgenic zebrafish. We further describe the methods to couple these experiments to time-lapse imaging of injured and control nerves. This experimental paradigm can be used to not only assess the role that glia play in nerve regeneration, but can also be the platform for elucidating the molecular mechanisms that govern nervous system repair.

Although the peripheral nervous system (PNS) is capable of significant repair after injury, little is known about the cellular and molecular mechanisms that govern this phenomenon. Using live, transgenic zebrafish and a reproducible nerve transection assay, we can study dynamic glial cell behaviors during nerve degeneration and regeneration.

The  nervous system is often described as a hard-wired component of the body even  though it is a considerably fluid organ system that reacts to external ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

In vivo Imaging of Optic Nerve Fiber Integrity by Contrast-Enhanced MRI in Mice

The rodent visual system encompasses retinal ganglion cells and their axons that form the optic nerve to enter thalamic and midbrain centers, and postsynaptic projections to the visual cortex. Based on its distinct anatomical structure and convenient accessibility, it has become the favored structure for studies on neuronal survival, axonal regeneration, and synaptic plasticity. Recent advancements in MR imaging have enabled the in vivo visualization of the retino-tectal part of this projection using manganese mediated contrast enhancement (MEMRI). Here, we present a MEMRI protocol for illustration of the visual projection in mice, by which resolutions of (200 &#181;m)3 can be achieved using common 3 Tesla scanners. We demonstrate how intravitreal injection of a single dosage of 15 nmol MnCl2 leads to a saturated enhancement of the intact projection within 24 hr. With exception of the retina, changes in signal intensity are independent of coincided visual stimulation or physiological aging. We further apply this technique to longitudinally monitor axonal degeneration in response to acute optic nerve injury, a paradigm by which Mn2+ transport completely arrests at the lesion site. Conversely, active Mn2+ transport is quantitatively proportionate to the viability, number, and electrical activity of axon fibers. For such an analysis, we exemplify Mn2+ transport kinetics along the visual path in a transgenic mouse model (NF-&#954;B p50KO) displaying spontaneous atrophy of sensory, including visual, projections. In these mice, MEMRI indicates reduced but not delayed Mn2+ transport as compared to wild type mice, thus revealing signs of structural and/or functional impairments by NF-&#954;B mutations.
In summary, MEMRI conveniently bridges in vivo assays and post mortem histology for the characterization of nerve fiber integrity and activity. It is highly useful for longitudinal studies on axonal degeneration and regeneration, and investigations of mutant mice for genuine or inducible phenotypes.

In vivo Imaging of Optic Nerve Fiber Integrity by Contrast-Enhanced MRI in Mice

This video illustrates a method, using a clinical 3 T scanner, for contrast-enhanced MR imaging of the na&#239;ve mouse visual projection and for repetitive and longitudinal in vivo studies of optic nerve degeneration associated with acute optic nerve crush injury and chronic optic nerve degeneration in knock-out mice (p50KO).

The rodent visual system encompasses retinal ganglion cells and their axons that form the optic nerve to enter thalamic and midbrain centers, and ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

Minimally-invasive Technique for Injection into Rat Optic Nerve

The rat optic nerve is a useful model for stem cell regeneration research. Direct injection into the rat optic nerve allows delivery into the central nervous system in a minimally-invasive surgery without bone removal. This technique describes an approach to visualization and direct injection of the optic nerve following minor fascial dissection from the orbital ridge, using a conjunctival traction suture to gently pull the eye down and out. Representative examples of an injected optic nerve show successful injection of dyed beads.

Direct injection into the rat optic nerve is useful for regenerative research. We demonstrate a minimally-invasive technique for direct injection into a rat optic nerve that does not involve opening the skull. Using this method, surgical complications are minimized and recovery is more rapid.

The rat optic nerve is a useful model for stem cell regeneration research. Direct injection into the rat optic nerve allows delivery into the central ...

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 ...

Exposure of the Pig CNS for Histological Analysis: A Manual for Decapitation, Skull Opening, and Brain Removal

Pigs have become increasingly popular in large-animal translational neuroscience research as an economically and ethically feasible substitute to non-human primates. The large brain size of the pig allows the use of conventional clinical brain imagers and the direct use and testing of neurosurgical procedures and equipment from the human clinic. Further macroscopic and histological analysis, however, requires postmortem exposure of the pig central nervous system (CNS) and subsequent brain removal. This is not an easy task, as the pig CNS is encapsulated by a thick, bony skull and spinal column. The goal of this paper and instructional video is to describe how to expose and remove the postmortem pig brain and the pituitary gland in an intact state, suitable for subsequent macroscopic and histological analysis.

The goal of this paper and instructional video is to describe how to expose and remove the postmortem pig brain and pituitary gland in an intact state, suitable for subsequent macroscopic and histological analysis.