Name: minimally-invasive technique for injection into rat optic nerve
Uploaded: 2015-05-19T14:00:00.000+00:00
Duration: 7 min 34 s
Description: Watch this Scientific Journal Video about Minimally-invasive Technique for Injection into Rat Optic Nerve at JoVE.com

This study was supported by NeuralStem, Inc., and Johns Hopkins Project RESTORE.

HINWEIS: Alle tierischen Verfahren wurden von der Johns Hopkins Animal Care und Verwenden Committee genehmigt. Anästhesiegeräte erfordern jährliche Inspektion und Kalibrierung nach Bedarf. 1. Anästhesie und Lagerung <ol><li> Anästhesie. <ol><li> Führen Sie alle chirurgischen Eingriffe unter Narkose mit 2-3% Isofluran. Bestätigen angemessene Höhe der Anästhesie durch toe Prise und Atemfrequenz. Überprüfen Sie, dass die Ratte nicht in Reaktion auf eine Zehe Prise zusammenzucken. HINWEIS: Ein Zucken zeigt Anästhesie, die zu spät ist und kann mehr Anästhesie vor Beginn oder eine höhere Isofluoran Konzentration erfordern. Ein Atemfrequenz weniger als 1 Atemzug alle 2 Sek zu langsam ist, die die Narkose ist zu hoch und kann Aufhellung der Isofluoran Konzentration erfordern. </li><li> Sichern Sie das Tier wie nötig, um die Kopfbewegung während des Verfahrens zu verhindern. Geben Sie einen Tropfen von Lidocain auf der chirurgischen Augen. Verwenden Sie künstliche Tränen alle 10 min bis zur Trockenheit während der Narkose zu verhindern. Gebeneine Injektion von Buprenorphin 0,01 mg / kg SC präoperativ und dann alle 6-8 h je nach Bedarf. </li></ol></li><li> Positionierung. <ol><li> Zeigen Ratten in einen stereotaktischen Rahmen und warm halten mit einem Heizkissen. Befeuchten Sie die Kopfhaut Pelz mit Alkohol kümmert, um die Exposition der Augen zu vermeiden. Verwenden Sie sterile Instrumente und sterile Technik, um das Risiko von postoperativen Infektionen zu minimieren. </li></ol></li></ol> 2. Eye Control <ol><li> Legen Sie eine 4-0 Naht in der seitlichen Bindehaut und binden Sie es mit genügend Naht auf sanfte Traktion ermöglichen. </li></ol> 3. Dissection <ol><li> Initial Dissection. <ol><li> Machen Sie eine ~ 1 Zoll Schnitt in der Haut über der Orbital Grat mit einer Größe 10 Skalpell, wie in Abbildung 1 gezeigt. Ziehen Sie die Haut und das darunter liegende Faszie und sorgfältig sezieren entfernt die Faszie. Um starke Blutungen in das Operations vermeiden, vermeiden, schneiden die Blutgefäße während sezieren die Faszie. Verwenden Sie strategisch placed Baumwolle Tipps, um Hämostase bereitzustellen. </li></ol></li><li> Deeper Dissection. HINWEIS: Bei leichten Zug auf der Bindehaut zieht das Auge nach unten und aus der Steckdose, wird der Muskel orbitalis superior in den Blick geraten. Um den optischen Nerv freizulegen, muss dieser Muskel geschnitten werden und die retroorbitalen Fett entfernt. Das Fett kann verworfen werden und nicht nach der Injektion ersetzt. Ab diesem Zeitpunkt sollte der Sehnerv Faszien als ein Bündel des Sehnerven selbst sichtbar sein, zusammen mit Blutgefäßen in Dura verpackt (Abbildung 1). <ol><li> Machen Sie einen kleinen Schnitt in der Dura mit dem Skalpell oder eine 31 Gauge Nadel abgeschrägt, um piercing weniger traumatisch. </li></ol></li></ol> 4. Pipettieren Injector <ol><li> Ziehen einer Glasmikropipette mit einem Durchmesser von 50-100 um. Um Stabilität zu gewährleisten, montieren Sie die Mikropipette auf einem Mikromanipulator und heften sich an einer Hamilton-Spritze mit einer Infusionspumpe angeschlossen. </li><li> Ziehen Sie die Perlen (oder Stammzellen)in einem 0,5-1,0 ul Volumen in der Mikropipette zusammen mit 0,5 &amp; mgr; l Volumen der Methylenblaulösung vor und nach Retrogradation rekonstituiert. HINWEIS: Eine Infusionsrate auf 0,5 bis 2 &amp; mgr; l pro min eingestellt verhindert Trauma des Sehnervs. </li></ol> 5. Injection <ol><li> Senken Sie die Spitze der Mikropipette auf den Sehnerv direkt über der Dura geklaut. HINWEIS: Eine kleine, aber rege Bewegung der Glasspitze in den Sehnerv Ergebnisse im geringsten Schaden. Da die Infusionspumpe beginnt, sollte die Methylenblau-Farbstoff den Bereich des Sehnervs injiziert markieren. Der Farbstoff sollte innerhalb des Sehnervs ohne Austreten in den Subarachnoidalraum lokalisiert bleiben. </li><li> Folgen Sie dem zweiten Band von Methylenblau, um festzustellen, wenn die Stammzellinjektion abgeschlossen ist und schalten Sie die Infusionspumpe aus, wenn die zweite Dosis von Methylenblau eingespritzt wird. Halten Sie die Mikropipette immer noch innerhalb des Sehnerven für 2 Minuten für jeweils 1 &amp; mgr; l Volumen eingespritzt, um zu verhindern HochDruckausstoß beim Zurückziehen der Mikropipette. HINWEIS: Ein alternativer Ansatz ist, um die Pipettenspitze mit einer Luftspritze, die manuell bearbeitet werden kann angeschlossen werden. So oder so, dass kein übermäßiger Kraft, um Schäden an den Sehnerven zu minimieren. Wir empfehlen, die Injektion nicht mehr als 2 &amp; mgr; l Volumen in jedem Sehnerv. </li></ol> 6. Folgen Sie up <ol><li> Unmittelbar. <ol><li> Wenn die Injektion abgeschlossen ist, entfernen Sie die Mikropipette zusammen mit allen Baumwollspitzen verwendet werden, um die Blutstillung bereitzustellen. Naht der Haut mit 3-0 Seide und entfernen Sie den Bindehautnaht. </li><li> Halten Ratten warmen auf einem Heizkissen, bis sie aus der Narkose hervorgehen. Haben ein Tier nicht unbeaufsichtigt lassen, bis er wieder zu sich kam, um ausreichende Brustlage zu halten. Sie ein Tier, das der Operation an die Firma von anderen Tieren unterzogen wurde, bis vollständig erholt nicht zurück. </li><li> Wird das Tier Anzeichen von Schmerzen, einschließlich übermäßige Lethargie, Ataxie, oder arbeiteten breathin Ausstellerg, zu verwalten angemessenen Schmerzmitteln mit intramuskulären Buprenorphin (0,05 mg / kg) bis zu dreimal täglich. </li></ol></li><li> Langfristig. <ol><li> Für die nächsten 24-48 Stunden, beobachten die Ratte für Komplikationen der Operation, einschließlich Schwellung oder Entlassung aus der Wunde oder anderen Anzeichen von Schmerzen, wie vocalizing, gebeugt Aussehen, nicht-Pflege, oder nicht essen. Betrachten Sie Rücksprache mit einem Tierarzt oder ethische Euthanasie wie nötig. </li></ol></li></ol>

<ol>
	<li>Dahlmann-Noor, A., Vijay, S., Jayaram, H., Limb, A., Khaw, P. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Current+approaches+and+future+prospects+for+stem+cell+rescue+and+regeneration+of+the+retina+and+optic+nerve.">Current approaches and future prospects for stem cell rescue and regeneration of the retina and optic nerve.</a> Canadian journal of ophthalmology Journal canadien d'ophtalmologie. 45 (4), 333-341 (2010).</li><li>Quigley, H. A., Iglesia, D. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stem+cells+to+replace+the+optic+nerve.">Stem cells to replace the optic nerve.</a> Eye. 18 (11), 1085-1088 (2004).</li><li>Ghaffarieh, A., Levin, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optic+nerve+disease+and+axon+pathophysiology.">Optic nerve disease and axon pathophysiology.</a> International review of neurobiology. 105, 1-17 (2012).</li><li>Fukui, Y., Hayasaka, S., Bedi, K. S., Ozaki, H. S., Takeuchi, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+study+of+the+development+of+the+optic+nerve+in+rats+reared+in+the+dark+during+early+postnatal+life.">Quantitative study of the development of the optic nerve in rats reared in the dark during early postnatal life.</a> Journal of anatomy. 174, 37-47 (1991).</li><li>Celestre, P. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Minimally+invasive+approaches+to+the+cervical+spine.">Minimally invasive approaches to the cervical spine.</a> The Orthopedic clinics of North America. 43 (1), 137-147 (2012).</li><li>Hallas, B. H., Wells, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Novel+Technique+for+Multiple+Injections+into+the+Mammalian+Optic+Nerve.">A Novel Technique for Multiple Injections into the Mammalian Optic Nerve.</a> Kopf Carrier. 54, (2001).</li><li>Harvey, A. R., Hellstrom, M., Rodger, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+therapy+and+transplantation+in+the+retinofugal+pathway.">Gene therapy and transplantation in the retinofugal pathway.</a> Progress in brain research. 175, 151-161 (2009).</li><li>Adachi-Usami, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optic+neuritis--from+diagnosis+to+optic+nerve+transplantation.">Optic neuritis--from diagnosis to optic nerve transplantation.</a> Nippon Ganka Gakkai zasshi. 104 (12), 841-857 (2000).</li><li>Slater, B. J., Vilson, F. L., Guo, Y., Weinreich, D., Hwang, S., Bernstein, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optic+nerve+inflammation+and+demyelination+in+a+rodent+model+of+nonarteritic+anterior+ischemic+optic+neuropathy.">Optic nerve inflammation and demyelination in a rodent model of nonarteritic anterior ischemic optic neuropathy.</a> Investigative ophthalmology &amp; visual science. 54 (13), 7952-7961 (2013).</li><li>Zarbin, M. A., Arlow, T., Ritch, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regenerative+nanomedicine+for+vision+restoration.">Regenerative nanomedicine for vision restoration.</a> Mayo Clinic proceedings. 88 (12), 1480-1490 (2013).</li></ol>

The authors have no meaningful disclosures to report.

Direct injection into the optic nerve of stem cells or other products intended to facilitate regeneration provides a convenient model compared to other means of injections into the CNS. This technique takes less time, requires less total anesthesia, avoids drilling or removing skull or bone tissue, reduces complications rates and allows for more rapid recovery following surgery. 
The most critical steps in this protocol include: 1. Adequate hemostasis in the surgical field to allow clear visua...

Der Sehnerv ist ein idealer Ort für das zentrale Nervensystem (ZNS) regenerative Forschung einschließlich ophthalmologische Erkrankungen wie Sehnervenentzündung, Glaukom und Trauma. Injektionen mit einer Vielzahl von Stammzellen wurden entweder Wirksamkeit gezeigt oder als vielversprechend bei der Ersetzung verlorener Myelin Erhöhung axonalen Zahl und / oder Prävention von degenerativen Erkrankungen. 1,2 Der menschliche Sehnerv enthält etwa 1,2 Millionen parallel Axone Weg von der Netzhaut zum chiasm mit einem Durchmesser von etwa 3,0-3,5 mm. 3 bei Erkrankungen des Menschen im Labor Modell wurde die Ratte häufig verwendet worden. Erwachsene Ratten Sehnerv enthält ungefähr 100.000 Axone innerhalb eines Durchmessers von etwa 0,5 mm. 4 eine der Hauptbeschränkungen bei der ZNS-regenerative Forschung direkten Zugriff ohne Knochen. Komplikationen und Operationsrisiken für das Tier sind höher, wenn der Schädel oder Wirbeln entfernt werden. Ähnlich zu den Vorteilenminimalinvasive Ansätze in der Wirbelsäule, 5 direkte Sehnerv Injektionen ohne Öffnen der Schädel Angebot reduziert Komplikationen und eine schnellere Erholung. Diese Technik wurde in früheren Studien verwendet worden. 6 In diesem Manuskript und begleitende Video zeigen wir eine minimal-invasives Verfahren zur Gewinnung von Stammzellen in der Ratte Sehnerv injizieren.

<table><tbody><tr><td>Lewis rat</td><td>Charles River</td><td>4</td><td>Any rat strain will work.</td></tr><tr><td>Anesthesia machine</td><td>Surgivet</td><td>CDS9000</td><td>CDS 9000 Small Animal Anesthesia Machine - Pole Mount</td></tr><tr><td>Infusion pump</td><td>Stoelting</td><td>53129</td><td></td></tr><tr><td>Dissection microscope</td><td>National Optical</td><td>409-411-1105</td><td></td></tr><tr><td>Fiber-optic light source</td><td>Fisher Scientific</td><td>12-562-21</td><td></td></tr><tr><td>Dissection and Stereotaxic Instrument</td><td>Stoelting</td><td>51400</td><td></td></tr><tr><td>Pipette Puller</td><td>Kopf</td><td>750</td><td></td></tr><tr><td>Pipettes</td><td>World Precision Instruments</td><td>18150-6</td><td></td></tr><tr><td>Disposable scalpel blades</td><td>Harvard Apparatus</td><td>810-15-021</td><td></td></tr><tr><td>Iridectomy scissors</td><td>Electron Microscopy Sciences</td><td>Uniband LA-4XF</td><td></td></tr></tbody></table>

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.

Am Ende des Experiments wurden die Ratten getötet und mit 4% Paraformaldehyd perfundiert. Die Sehnerven wurden sorgfältig seziert und für Kryostat Schnitte angebracht. Abbildung 2 zeigt ein Beispiel einer Ratte ganzen Sehnerven bei geringer Leistung, bei dem Evans-Blau-Farbstoff wurde, um die Website zu visualisieren injiziert. Der Pfeil bezeichnet die genaue Lage der Injektion. Diese Präparation wurde in wenigen Minuten von der Injektion durchgeführt, wie durch die beschränkte Diffusion des Farbs...

Watch this Scientific Journal Video about Minimally-invasive Technique for Injection into Rat Optic Nerve at JoVE.com

Minimally-invasive Technique for Injection into Rat Optic Nerve

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.

Minimal-invasive Technik für die Injektion in Ratten-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 ...

minimally-invasive-technique-for-injection-into-rat-optic-nerve

Research

JoVE Journal

Neuroscience

9.5K Aufrufe.  Johns Hopkins University. 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. 

Serie: Minimally-invasive Technique for Injection into Rat Optic Nerve

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.

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.

Methoden für die experimentelle Manipulationen nach Optic Nerve Durchtrennung im Säuger-ZNS

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 ...

Forschung

Neurowissenschaften

Intravitreous Injection for Establishing Ocular Diseases Model

Intravitreous injection is a widely used technique in visual sciences research. It can be used to establish animal models with ocular diseases or as direct application of local treatment. This video introduces how to use simple and inexpensive tools to finish the intravitreous injection procedure. Use of a 1 ml syringe, instead of a hemilton syringe, is used. Practical tips for how to make appropriate injection needles using glass pipettes with perfect tips, and how to easily connect the syringe needle with the glass pipette tightly together, are given.
To conduct a good intravitreous injection, there are three aspects to be observed: 1) injection site should not disrupt retina structure; 2) bleeding should be avoided to reduce the risk of infection; 3) lens should be untouched to avoid traumatic cataract. In brief, the most important point is to reduce the interruption of normal ocular structure. To avoid interruption of retina, the superior nasal region of rat eye was chosen. Also, the puncture point of the needle was at the par planar, which was about 1.5 mm from the limbal region of the rat eye. A small amount of vitreous is gently pushed out through the puncture hole to reduce the intraocular pressure before injection. With the 45&deg; injection angle, it is less likely to cause traumatic cataract in the rat eye, thus avoiding related complications and influence from lenticular factors. In this operation, there was no cutting of the conjunctiva and ocular muscle, no bleeding. With quick and minor injury, a successful intravitreous injection can be done in minutes. 
The injection set outlined in this particular protocol is specific for intravitreous injection. However, the methods and materials presented here can also be used for other injection procedures in drug delivery to the brain, spinal cord or other organs in small mammals.

Intravitreous injection is a widely used technique in visual sciences research for ocular diseases or as direct application of local treatment. This video demonstrated a protocol for intravitreous injection using a 1ml syringe with glass pipette. Useful tips about avoiding massive bleeding and lens damage are given.

Intravitrealen Injektion zur Einrichtung Ocular Krankheiten Modell

Intravitreous injection is a widely used technique in visual sciences research. It can be used to establish animal models with ocular diseases or as ...

Biologie

Rat Model of Photochemically-Induced Posterior Ischemic Optic Neuropathy

Posterior Ischemic optic neuropathy (PION) is a sight-devastating disease in clinical practice. However, its pathogenesis and natural history have&#160;remained poorly understood. Recently, we developed a reliable, reproducible animal model of PION and tested the treatment effect of some neurotrophic factors in this model1. The purpose of this video is to demonstrate our photochemically induced model of posterior ischemic optic neuropathy, and to evaluate its effects with retrograde labeling of retinal ganglion cells. Following surgical exposure of the posterior optic nerve, a photosensitizing dye, erythrosin B, is intravenously injected and a laser beam is focused onto the optic nerve surface. Photochemical interaction of erythrosin B and the laser during irradiation damages the vascular endothelium, prompting microvascular occlusion mediated by platelet thrombosis and edematous compression. The resulting ischemic injury yields a gradual but pronounced retinal ganglion cell dieback, owing to a loss of axonal input - a remote, injury-induced and clinically relevant outcome. Thus, this model provides a novel platform to study the pathophysiologic course of PION,&#160;and can be further optimized for testing therapeutic approaches for optic neuropathies as well as other CNS ischemic diseases.

The goal of this protocol is to photochemically induce ischemic injury to the posterior optic nerve in rat. This model is critical to studies of the pathophysiology of posterior ischemic optic neuropathy, and therapeutic approaches for this and other optic neuropathies, as well as of other CNS ischemic diseases.

Rattenmodell der photochemisch-induzierte Posterior ischämische Optikusneuropathie

Posterior Ischemic optic neuropathy (PION) is a sight-devastating disease in clinical practice. However, its pathogenesis and natural history ...

Medizin

An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach

Subretinal injections have been successfully used in both humans and rodents to deliver therapeutic interventions of proteins, viral agents, and cells to the interphotoreceptor/subretinal compartment that has direct exposure to photoreceptors and the retinal pigment epithelium (RPE). Subretinal injections of plasminogen as well as recent preclinical and clinical trials have demonstrated safety and/or efficacy of delivering viral vectors and stem cells to individuals with advanced retinal disease. Mouse models of retinal disease, particularly hereditary retinal dystrophies, are essential for testing these therapies. The most common injection procedure in rodents is to use small transcorneal or transcleral incisions with an anterior approach to the retina. With this approach, the injection needle penetrates the neurosensory retina disrupting the underlying RPE and on insertion can easily nick the lens, causing lens opacification and impairment of noninvasive imaging. Accessing the subretinal space via a transcleral, posterior approach avoids these problems: the needle crosses the sclera approximately 0.5 mm from the optic nerve, without retinal penetration and avoids disrupting the vitreous. Collateral damage is limited to that associated with the focal sclerotomy and the effects of a transient, serous retinal detachment. The simplicity of the method minimizes ocular injury, ensures rapid retinal reattachment and recovery, and has a low failure rate. The minimal damage to the retina and RPE allows for clear assessment of the efficacy and direct effects of the therapeutic agents themselves. This manuscript describes a novel subretinal injection technique that can be used to target viral vectors, pharmacological agents, stem cells or induced pluripotent stem (iPS) cells to the subretinal space in mice with high efficacy, minimal damage, and fast recovery.

An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach

Subretinal injections are the most common technique for delivering large therapeutic agents such as proteins and viral vectors to photoreceptors and the retinal pigment epithelium. An alternative method in mice that successfully targets the subretinal space with minimal collateral damage and fast recovery times is described here.

Eine Alternative und Validated Injektionsmethode für den Zugriff auf den subretinalen Raum<i&gt; über</i&gt; A Transcleral dorsalen Zugang

Subretinal injections have been successfully used in both humans and rodents to deliver therapeutic interventions of proteins, viral agents, and cells to ...

The Rodent Model of Nonarteritic Anterior Ischemic Optic Neuropathy (rNAION)

Nonarteritic anterior ischemic optic neuropathy (NAION) is a focal ischemic lesion of the optic nerve that affects 1/700 individuals throughout their lifetime. NAION results in optic nerve edema, selective loss of the retinal ganglion cell neurons (RGCs) and atrophy of the optic nerve. A rodent model of NAION that expresses most NAION features and sequelae has been developed, which is applicable to both rats and mice. This model utilizes a focal laser application of 532 nm wavelength to illuminate a photoactive dye, Rose Bengal (RB), to cause capillary damage and leakage at the targeted anterior optic nerve (the laminar region). After rNAION induction, there is an early optic nerve ischemia, optic nerve edema, and intraneural inflammation, followed by selective RGC and axonal loss. Since the optic nerve is a CNS white matter tract, the rNAION model is applicable to mechanistic studies of selective white matter ischemia, as well as neuroprotective analyses and short and long-term mechanisms of glial and neuronal response to ischemia.

The Rodent Model of Nonarteritic Anterior Ischemic Optic Neuropathy rNAION

The following report describes how to replicate the rodent model of nonarteritic anterior ischemic optic neuropathy (rNAION), using the appropriate dye, contact lens and laser parameters. We also reveal the appropriate steps for evaluating the rNAION lesion in vivo.

Das Nagetier-Modell der nonarteritic anteriore ischämische Optikusneuropathie (rNAION)

Nonarteritic anterior ischemic optic neuropathy (NAION) is a focal ischemic lesion of the optic nerve that affects 1/700 individuals throughout their ...

Ultrahigh Resolution Mouse Optical Coherence Tomography to Aid Intraocular Injection in Retinal Gene Therapy Research

HR-SD-OCT is utilized to monitor the progression of photoreceptor degeneration in live mouse models, assess the delivery of therapeutic agents into the subretinal space, and to evaluate toxicity and efficacy in vivo. HR-SD-OCT uses near infrared light (800-880 nm) and has optics specifically designed for the unique optics of the mouse eye with sub-2-micron axial resolution. Transgenic mouse models of outer retinal (photoreceptor) degeneration and controls were imaged to assess the disease progression. Pulled glass microneedles were used to deliver sub retinal injections of adeno-associated virus (AAV) or nanoparticles (NP) via a trans-scleral and trans-choroidal approach. Careful positioning of the needle into the subretinal space was required prior to a calibrated pressure injection, which delivers fluid into the sub retinal space. Real time subretinal surgery was conducted on our retinal imaging system (RIS). HR-SD-OCT demonstrated progressive uniform retinal degeneration due to expression of a toxic mutant human mutant rhodopsin (P347S) (RHOP347S) transgene in mice. HR-SD-OCT allows rigorous quantification of all the retinal layers. Outer nuclear layer (ONL) thickness and photoreceptor outer segment length (OSL) measurements correlate with photoreceptor vitality, degeneration, or rescue. The RIS delivery system allows real-time visualization of subretinal injections in neonatal (~P10-14) or adult mice, and HR-SD-OCT immediately determines success of delivery and maps areal extent. HR-SD-OCT is a powerful tool that can evaluate the success of subretinal surgery in mice, in addition to measuring vitality of photoreceptors in vivo. HR-SD-OCT can also be used to identify uniform animal cohorts to evaluate the extent of retinal degeneration, toxicity, and therapeutic rescue in preclinical gene therapy research studies.

Here we demonstrate a novel approach to using high resolution spectral-domain optical coherence tomography (HR-SD-OCT) to assist delivery of gene therapy agents into the subretinal space, assess its areal coverage, and characterize photoreceptor vitality.

Höchste Auflösung Maus optische Kohärenztomographie, intraokulare Injektion in die retinale Gentherapie Forschung unterstützen

HR-SD-OCT is utilized to monitor the progression of photoreceptor degeneration in live mouse models, assess the delivery of therapeutic agents into the ...

Sheath-Preserving Optic Nerve Transection in Rats to Assess Axon Regeneration and Interventions Targeting the Retinal Ganglion Cell Axon

Retinal ganglion cell (RGC) axons converge at the optic nerve head to convey visual information from the retina to the brain. Pathologies such as glaucoma, trauma, and ischemic optic neuropathies injure RGC axons, disrupt transmission of visual stimuli, and cause vision loss. Animal models simulating RGC axon injury include optic nerve crush and transection paradigms. Each of these models has inherent advantages and disadvantages. An optic nerve crush is generally less severe than a transection and can be used to assay axon regeneration across the lesion site. However, differences in crush force and duration can affect tissue responses, resulting in variable reproducibility and lesion completeness. With optic nerve transection, there is a severe and reproducible injury that completely lesions all axons. However, transecting the optic nerve dramatically alters the blood brain barrier by violating the optic nerve sheath, exposing the optic nerve to the peripheral environment. Moreover, regeneration beyond a transection site cannot be assessed without reapposing the cut nerve ends. Furthermore, distinct degenerative changes and cellular pathways are activated by either a crush or transection injury.
The method described here incorporates the advantages of both optic nerve crush and transection models while mitigating the disadvantages. Hydrostatic pressure delivered into the optic nerve by microinjection completely transects the optic nerve while maintaining the integrity of the optic nerve sheath. The transected optic nerve ends are reapposed to allow for axon regeneration assays. A potential limitation of this method is the inability to visualize the complete transection, a potential source of variability. However, visual confirmation that the visible portion of the optic nerve has been transected is indicative of a complete optic nerve transection with 90-95% success. This method could be applied to assess axon regeneration promoting strategies in a transection model or investigate interventions that target the axonal compartments.

This protocol describes an optic nerve transection that preserves the optic nerve sheath in rats. Hydrostatic pressure from microinjections into the optic nerve generate a complete transection, allowing for sutureless reapposition of the transected optic nerve ends and direct targeting of the axonal compartment in a transection model.

Retinal ganglion cell (RGC) axons converge at the optic nerve head to convey visual information from the retina to the brain. Pathologies such as ...

Direct Vagus Nerve Injection Protocol For Rats

There is a relative abundance of strategies and methodologies to facilitate drug delivery to the central nervous system. However, drug delivery directly to the peripheral nervous system is less common, with fewer detailed methods publications available to aid researchers. Here, we describe a direct nerve injection method for peripheral nervous system drug delivery, using the vagus nerve as a model nerve. This method can be used in the treatment of autonomic nervous system disorders through targeting of the left vagus nerve, although this general injection method can be extrapolated to injection of other nerves with minor modification. This method explains all critical steps involved in the procedure involving microsurgery in anesthetized adult rats under a dissecting microscope. The use of a tracking dye is described to facilitate the monitoring of injection fidelity in real time. Illustrations of successful and failed injections are provided. If carried out properly, direct vagus nerve injections can be conducted in a safe manner that is well-tolerated by the rat without post-delivery complications. For example, once the surgeons were trained in this method, six out of six rats were successfully injected without any complications. This method of direct nerve injection for preclinical rat studies is capable of delivering agents (including but not limited to gene therapy) to peripheral nerves.

We present a protocol for direct vagus nerve injection in rats, enabling drug delivery directly into the nerve without post-injection complications. This method applies to preclinical neurological studies involving autonomic nervous system manipulation. It can be used for direct nerve injection for other nerves in rats and other species, with necessary modifications.

Direktes Vagusnerv-Injektionsprotokoll für Ratten

There is a relative abundance of strategies and methodologies to facilitate drug delivery to the central nervous system. However, drug delivery directly ...

Transconjunctival Approach for Injection into the Rat Optic Nerve

The rat serves as an important model for studying disorders of the optic nerve, including heritable, traumatic, neoplastic, and autoimmune conditions. Accessing the rat optic nerve for experimental manipulations and injections is challenging due to the small size of the orbit and the vascularity of the surrounding orbital tissues. Previous techniques have involved a cutaneous incision, which carries a higher risk of infection and requires wound closure. This study aims to describe a unique, less invasive, and potentially more efficient approach to accessing the rat optic nerve. A transconjunctival incision, along with a rat spinal cord hook, is used to isolate the optic nerve for experimental injections. In these experiments, an adenoviral vector is injected into the optic nerve with the goal of using this technique for direct gene therapy delivery in the future. This technique can also be applied by investigators requiring access to the rat optic nerve for other experimental studies, including drug delivery and optic nerve injury research.

This protocol describes a unique technique using a transconjunctival approach to access the rat optic nerve for injections.

Transkonjunktivaler Zugang zur Injektion in den Sehnerv der Ratte

The rat serves as an important model for studying disorders of the optic nerve, including heritable, traumatic, neoplastic, and autoimmune conditions. ...

<meta charset="utf8"></head><body>Optimierte intrakamerale Injektionstechnik für Ratten zur Reduzierung von Gewebeschäden

Intracameral Injection in Rats with Low Risk of Adverse Effects

Intracameral injection is a standard administration routine in ophthalmology. The application of intracameral injection in rodents for research is challenging due to the limiting dimensions and anatomy of the eye, including the small aqueous humor volume, the lens curvature, and lens thickness. Potential damage during intracameral injections introduces adverse effects and experimental variability. This protocol describes a procedure for intracameral injection in rats, allowing precision and reproducibility.
Sprague-Dawley rats were used as experimental models. Since the lens position in rats protrudes into the anterior chamber, injecting from the periphery, as done in humans, is unfavorable. Therefore, an incision is created in the central corneal region using a 31 gauge 0.8 mm stiletto blade to form a self-sealing tunnel into the anterior chamber. An incision at&#160;an angle close to the flat allows to create a long tunnel, which minimizes the loss of aqueous humor and shallowing of the anterior chamber. A 34 gauge nanoneedle is inserted into the tunnel for injection. This enables penetration with minimal friction resistance and avoids touching the lens. Injection of trypan-blue allows&#160;visualization&#160;by slit microscopy the presence of the dye in the anterior chamber and exclude leakage. Bioavailability to the corneal endothelial layer is demonstrated by injection of Hoechst dye, which stained the nuclei of corneal endothelial cells after injection.
In conclusion, this protocol implements a procedure for accurate intracameral injection in rats. This procedure may be used for intracameral delivery of various drugs and compounds in experimental rat models, increasing the efficiency and reproducibility of ophthalmic research.

<meta charset="utf8"></head><body>Autor im Rampenlicht: Ein neuartiges Protokoll für intrakamerale Injektionen zur Verbesserung der Präzision in der Nagetier-Ophthalmologie

This protocol describes a technique for intracameral injection in rats using a central corneal incision and a long tunnel into the anterior chamber. This injection method minimizes the risk of inducing inadvertent tissue damage and thereby improves precision and reproducibility.

Minimal-invasive Technik für die Injektion in Ratten-Optic Nerve

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