<p class="jove_content"&gt; Diese Arbeit wurde von der intramuralen Programm des National Institute of Neurological Disorders and Stroke in der National Institutes of Health unterstützt. Wir möchten auch an Dr. George Mentis für seine früheren Beiträge zur Methode und zu den Daten in Abbildung 3 zu danken.</p

Fluoreszierende Dextrane wurden als anatomische Werkzeuge und für die Bildgebung neuronaler Aktivität 4.1 verwendet wurde. Fields et al., (2009) 4 veröffentlichte ein Protokoll für die Anwendung von Ionen-und spannungsabhängigen Farbstoffen zu axonalen Bahnen mit einem Fokus auf das Rückenmark als Modellsystem. Hier beschreiben wir eine detaillierte Verfahren für die Anwendung fluoreszierender und / oder Ionen-sensitiven Farbstoffs an den Schnittflächen ventralen Wurzeln, dorsalen Wurzeln oder neuronaler des Rückenmarks für in-vitro optophysiological und morphologischen Untersuchungen. 1. Typ-I-und Typ-II-Pipetten Starten Sie, indem Sie zwei kurze Abschnitte aus Polyethylen-Schlauch (PE90, Clay Adams Brand) über der Flamme eines Alkohols Lampe 7 bis Pipetten mit verjüngten Spitzen produzieren. Eine Pipette (Typ-I) sollte kurz sein (3-7 mm) mit einer kleinen Spitze, dicht zu halten, kann das Ziel-Root-oder Darm-Trakt (Außendurchmesser: 0,2 bis 0,4 mm; Innendurchmesser: 0,1-0,3 mm). Ziehen Sie dann eine zweite Pipette (Typ-II) mit einer längeren (8-12 cm) dünner, Welle und eine sehr feine Spitze (Aussendurchmesser: 0,2-0,3 mm, Innendurchmesser: 0,1-0,2 mm), die in einsetzbaren die Typ-I pipettieren soweit seine Spitze. Die zweite dünner Pipette verwendet zum Ansaugen und Einführung künstlicher zerebrospinaler Flüssigkeit (ACSF) und Farbstofflösung jeweils von der Typ-I-Pipette werden. 2. Positionierung von Typ-I pipettieren Sezieren und zu isolieren, die das Rückenmark 8,9 und legen Sie sie in einem Bad, mit gekühltem aCSF (~ 16 ° C) während des Ladevorgangs superfundiert. Platzieren Sie die Typ-I-Pipette auf einem Elektrodenhalter (H1/12 Elektrodenhalter, Narishige), so dass die hintere Öffnung leicht mit einer Spritze (1 ml U-100 Insulin Spritze, Becton Dickinson oder vergleichbaren) werden über einen flexiblen Schlauch verbunden (PharMed BPT, Cole-Parmer, # AY242002, 10 cm) (1A-B). 3. Farbauftrag <ol><li> Mit Hilfe der Spritze, zuerst ziehenaCSF in den Typ-I-Pipette (1B; 2A), gefolgt von der axonalen-Darm-Trakt oder Wurzel zu besetzen (Abb. 2B) werden. Der flexible Schlauch sollte dann von der hinteren Öffnung entfernt werden. Der Typ-II-Pipette in eine andere Spritze befestigt und in die Typ-I pipettieren Halten des axonalen Trakt oder Wurzel. Mit der Spritze an der Typ-II-Pipette absaugen aCSF, so dass der Rest nur aCSF deckt den axonalen-Darm-Trakt (Abb. 1c; 2C-D). Dann ziehen Sie den Typ-II-Pipette aus der Typ-I-Pipette. Überwachen Sie den aCSF Ebene für ein paar Minuten, um eine gute Abdichtung auf der axonalen-Darm-Trakt oder Nerven zu überprüfen (siehe Abschnitt 3 in Fehlerbehebung für &quot;gute Dichtung&quot;). </li><li> Man löst den Farbstoff in aCSF oder 0,2% Triton X-100 in doppelt destilliertem Wasser und ziehen es in die Typ-II-Pipette und achten Sie dabei keine Luftblasen enthalten. Legen Sie die Typ-II-Pipette, die den Farbstoff in den Typ-I-Pipette und in den Überlebensraum aCSF Lösung (Abb. 2E). Langsam loslassender Farbstoff in die aCSF unter leichtem Überdruck (Abb. 2F). Achten Sie darauf, um mögliche Luftblasen einzuführen oder die Verschiebung der Zielgewebe aus dem Inneren der Spitze führen. Entnehmen Sie die Typ-II-Pipette nach einer ausreichenden Menge an Farbstoff wurde veröffentlicht (3-5 ul Farbstofflösung). </li></ol> 4. Incubation Inkubieren des Gewebes in der Dunkelheit zwischen 6 und 20 Stunden, je nach der Art des Experiments. Sobald genügend Zeit für die Füllung wurde erlaubt, ziehen weg die Elektrode aus dem Gewebe verlassen das Gewebe fertig für die Bildgebung oder Histologie. Fehlerbehebung <ol><li> Wenn das Zielgewebe nicht leicht treten in der Spitze des Typ-I-Pipette oder locker ist einmal drinnen, dann wird der Durchmesser der Spitze hat die falsche Größe. Wenn dies der Fall, ziehen und verwenden Sie eine andere Pipette. </li><li> Vor Zugabe des Farbstoffs an das aCSF in der Typ-I-Pipette, um sicherzustellen, dass genug aCSF so bleibt, dass es mit der erreicht werden kannTyp-II-Pipettenspitze (2D). Andernfalls wird ein Luftspalt vorhanden ist, wenn der Farbstoff zugegeben. Wenn dies der Fall, fügen Sie nicht den Farbstoff und entfernen Sie die axonale-Darm-Trakt aus der Pipette und dann ziehen sie zurück in mit ausreichender aCSF von der Typ-II-Pipette erreicht werden. </li><li> Wenn die aCSF Ebene in den Typ-I-Pipette erhöht, nachdem sie mit dem Typ-II Pipette abgesaugt wurde, bedeutet dies, dass die Dichtung mit dem Gewebe nicht gut ist. Verwenden Sie eine andere Elektrode, da sonst der Farbstoff wird während der Etikettierung Verfahren verdünnt werden. </li><li> Wenn eine Luftblase in die Typ-I-Pipette eingeführt wird während der Injektion des Farbstoffs, die Blase abfließen kann durch Vorschieben des Typ-II-Pipettenspitze in der Nähe der Blase und sanft Anlegen eines schwachen Unterdruck unter Verwendung des beigefügten Spritze. </li></ol> 5. Repräsentative Ergebnisse Um eine Anwendung der Methode zu veranschaulichen, haben wir gleichzeitig geladen Motoneuronen und sensorischen Afferenzen spinalen Interneuronen mit drei verschiedenen dextlief-konjugierten Fluoreszenzfarbstoffe (3A). Wie in dargestellt. 3B, drei Klassen von Neuronen markiert sind; afferenten Neuronen (rot), Interneuronen, die in das ventrale Funiculus (grün) und Motoneuronen (blau). <img alt="1" src="/files/ftp_upload/4008/4008fig1.jpg" /> Abbildung 1. Schematische Darstellung Pipette Typ-I-und Typ-II-Montage. (A) ein Typ-I Pipette (hellblau) auf einer Elektrode Halter platziert, so dass es wieder ist in Öffnung kann einfach an dem flexiblen Elastomer-Schlauch (B) verbunden werden und in die ein Typ-II-Pipette kann eingesetzt werden (C). <img alt="2" src="/files/ftp_upload/4008/4008fig2.jpg" /> Abbildung 2. Schematische Darstellung der Ladevorgang von Motoneuronen mit fluoreszierenden Farbstoffen. (A) ein Typ-I-Pipette in der Nähe des ventralen Wurzel zu füllen angeordnet. (B) ist mit dem Saug-Pipette auf aCSF in die Pipette an der Wurzel zu ziehen, gefolgt angewendet. (C-D) Trennen Sie das flexible Elastomer-Schlauch aus dem Backend von Typ-I-Pipette und reduzieren die Menge an aCSF durch Ansaugen des Typ-II-Pipette. (E) Führen Sie die Spitze des Typ-II-Pipette mit dem gelösten Farbstoff in den verbleibenden aCSF in der Typ-I-Pipette. (EF) Langsam füllt sich die Typ-I-Pipette mit dem Farbstoff und entfernen Sie dann den Typ-II-Pipette. <img alt="Abbildung 3" src="/files/ftp_upload/4008/4008fig3.jpg" /> Abbildung 3. Anwendung des Füllvorganges zu dem isolierten Maus-Rückenmark. (A) Schematische Darstellung der drei Typ-I-Pipetten verwendet werden, um die verschiedenen neuronalen Klassen im Rückenmark zu füllen. Motoneuronen (blau; Cascade Blue Dextran) und sakralen sensorischen Afferenzen (rot; Texas-Red Dextran) wurden durch die ventralen und dorsalen Wurzeln verfüllt bzw.. Fluorescein Dextran wurde zur spinalen Interneuronen (grün), deren Axone steigen durch den ventralen Funiculus (VF) zu laden. 1 mg eines jeden Farbstoff Dextran (10.000 MW; Molecular Probes)wurde in 6 ul destilliertem Wasser mit 0,2% gelöst Triton X-100 (B). Konfokale Aufnahme eines Ausschnitts aus dem sakralen Schnur des Rückenmarks, in denen die Neuronen waren zurück-markiert, wie in A. Hinweis beschrieben, dass die markierten sensorischen Afferenzen primär ipsilateral mit ein paar kontralaterale Projektionen sind. Die markierten Motoneuronen sind ipsilateral zur gefüllten ventralen Wurzel, während die kontralaterale markierten Interneuronen an der Seite der Füllung (Pfeilspitze) sind. Kalibrierung bar 200 um.

<ol>
	<li>Nance, D. M., Burns, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+dextrans+as+sensitive+anterograde+neuroanatomical+tracers:+applications+and+pitfalls.">Fluorescent dextrans as sensitive anterograde neuroanatomical tracers: applications and pitfalls.</a> Brain Res. Bull. 25, 139-145 (1990).</li><li>Glover, J. C., Petursdottir, G., Jansen, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+dextran-amines+used+as+axonal+tracers+in+the+nervous+system+of+the+chicken+embryo.">Fluorescent dextran-amines used as axonal tracers in the nervous system of the chicken embryo.</a> J. Neurosci. Methods. 18, 243-254 (1986).</li><li>McPherson, D. R., McClellan, A. D., O'Donovan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+imaging+of+neuronal+activity+in+tissue+labeled+by+retrograde+transport+of+Calcium+Green+Dextran.">Optical imaging of neuronal activity in tissue labeled by retrograde transport of Calcium Green Dextran.</a> Brain Research Protocols. 1, 157-164 (1997).</li><li>Fields, D. R., Shneider, N., Mentis, G. Z., O'Donovan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+nervous+system+activity.">Imaging nervous system activity.</a> Curr. Protoc. Neurosci. Chapter 2, Unit 2.3 (2009).</li><li>O'Donovan, M. 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Methods. 70, 111-120 (1996).</li><li>Garudadri, S., Gallarda, B., Pfaff, S., Alaynick, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spinal+Cord+Electrophysiology+II:+Extracellular+Suction+Electrode+Fabrication.">Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication.</a> J. Vis. Exp. (48), e2580 (2011).</li><li>Smith, J. C., Feldman, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+brainstem-spinal+cord+preparations+for+study+of+motor+systems+for+mammalian+respiration+and+locomotion.">In vitro brainstem-spinal cord preparations for study of motor systems for mammalian respiration and locomotion.</a> J. Neurosci. Methods. 21, 321-333 (1987).</li><li>Meyer, A., Gallarda, B., Pfaff, S., Alaynick, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spinal+Cord+Electrophysiology.">Spinal Cord Electrophysiology.</a> J. Vis. Exp. (35), e1660 (2010).</li><li>Shneider, N. A., Mentis, G. Z., Schustak, J., O'Donovan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functionally+reduced+sensorimotor+connections+form+with+normal+specificity+despite+abnormal+muscle+spindle+development:+the+role+of+spindle-derived+neurotrophin+3.">Functionally reduced sensorimotor connections form with normal specificity despite abnormal muscle spindle development: the role of spindle-derived neurotrophin 3.</a> J. Neurosci. 29, 4719-4735 (2009).</li><li>Mentis, G. Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+functional+impairment+of+sensory-motor+connectivity+in+a+mouse+model+of+spinal+muscular+atrophy.">Early functional impairment of sensory-motor connectivity in a mouse model of spinal muscular atrophy.</a> Neuron. 69, 453-467 (2011).</li><li>Blivis, D., Mentis, Z., O'Donovan, J. M. G., Lev-Tov, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Studies+of+sacral+neurons+involved+in+activation+of+the+lumbar+central+pattern+generator+for+locomotion+in+the+neonatal+rodent+spinal+cord.">Studies of sacral neurons involved in activation of the lumbar central pattern generator for locomotion in the neonatal rodent spinal cord.</a> Soc. Neurosci. Abstr. 564.8, (2009).</li><li>O'Donovan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+the+spatiotemporal+organization+of+neural+activity+in+the+developing+spinal+cord.">Imaging the spatiotemporal organization of neural activity in the developing spinal cord.</a> Dev. Neurobiol. 68, 788-803 (2008).</li><li>Kasumacic, N., Glover, J. C., Perreault, M. -. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Segmental+patterns+of+vestibular-mediated+synaptic+inputs+to+axial+and+limb+motoneurons+in+the+neonatal+mouse+assessed+by+optical+recording.">Segmental patterns of vestibular-mediated synaptic inputs to axial and limb motoneurons in the neonatal mouse assessed by optical recording.</a> J. Physiol. (Lond). 588, 4905-4925 (2010).</li><li>Szokol, K., Glover, J. C., Perreault, M. -. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organization+of+functional+synaptic+connections+between+medullary+reticulospinal+neurons+and+lumbar+descending+commissural+interneurons+in+the+neonatal+mouse.">Organization of functional synaptic connections between medullary reticulospinal neurons and lumbar descending commissural interneurons in the neonatal mouse.</a> J. Neurosci. 31, 4731-4742 (2011).</li></ol>

<p class="jove_content"&gt; Wir haben nichts zu offenbaren.</p

<p class="jove_content"&gt; Wir beschreiben hier eine einfache und kostengünstige Protokoll zur Einführung von Farbstoffen in Neuronen, Nerven und Rückenmarksbahnen. Dieses Verfahren schließt ein, dass identifizierten anatomischen Pfade einem hoch konzentrierten Farbstofflösung für die Dauer des Ladevorgangs. Dies führt zu retrograde Markierung der Ziel-Site mit wenig Hintergrund zu Mikroinjektionen, Bad und Anwendung der Elektroporation Techniken verglichen. Jedoch ist das Verfahren auf Seiten des Nervensystems, wo eine Strecke von Axon...

<table border="1"&gt;<tbody&gt;<tr&gt;<td colspan="2"<strong&gt; Material Name</strong</td&gt;<td<strong&gt; Unternehmen</strong</td&gt;<td<strong&gt; Katalog-Nummer</strong</td&gt;<td<strong&gt; Kommentare</strong</td&gt;</tr&gt;<tr&gt;<td rowspan="2"&gt; Schläuche</td&gt;<td&gt; PE90 (IDxOD: 0,034 &quot;x0.050&quot;; Wand thikness: 0,008 &quot;)</td&gt;<td&gt; Clay Adams Marke Intermedic</td&gt;<td&gt; 427421</td&gt;<td&gt; Für Typ I und Typ II Pipetten</td&gt;</tr&gt;<tr&gt;<td&gt; NSF-51 (IDxOD: 1/16x1/8; Wand thikness: 1/32)</td&gt;<td&gt; PharMed BPT, Cole-Parmer</td&gt;<td&gt; AY242002</td&gt;<td&gt; Für flexible Schläuche</td&gt;</tr&gt;<tr&gt;<td&gt; Spritze</td&gt;<td&gt; 1 ml Insulinspritze U-100</td&gt;<td&gt; Becton Dickenson</td&gt;<td&gt; 329650</td&gt;<td&gt;</td&gt;</tr&gt;<tr&gt;<td&gt; Needle</td&gt;<td&gt; 19G x1-1/2 &quot;Metal Hub Needle</td&gt;<td&gt; MONOJECT</td&gt;<td&gt; 200136</td&gt;<td&gt; Anschluss vom Typ II Pipette Spritze</td&gt;</tr&gt;<tr&gt;<td&gt;</td&gt;<td&gt; Alkohol Lampe</td&gt;<td&gt;</td&gt;<td&gt;</td&gt;<td&gt;</td&gt;</tr&gt;<tr&gt;<td&gt; Hahn</td&gt;<td&gt; Drei-Wege-Hahn mit Luer-Adapter männlich</td&gt;<td&gt; Baxter Healthcare Corp</td&gt;<td&gt; 2C6241</td&gt;<td&gt; Verwenden Sie mit Spritze</td&gt;</tr&gt;<tr&gt;<td rowspan="3"&gt; Halter</td&gt;<td&gt; H-1 Elektrodenhalter</td&gt;<td&gt; Narishige</td&gt;<td&gt; H-1/12</td&gt;<td&gt;</td&gt;</tr&gt;<tr&gt;<td&gt; Magnet-Stand</td&gt;<td&gt; Narishige</td&gt;<td&gt; GJ-8</td&gt;<td&gt;</td&gt;</tr&gt;<tr&gt;<td&gt; Mikromanipulator</td&gt;<td&gt; Narishige</td&gt;<td&gt; M-3333</td&gt;<td&gt;</td&gt;</tr&gt;<tr&gt;<td&gt; Pinzetten</td&gt;<td&gt; S &amp; T DUMONT Schweizer 00632-11</td&gt;<td&gt; DuMont</td&gt;<td&gt; JF-5 TC</td&gt;<td&gt;</td&gt;</tr&gt;</tbody&gt;</table&gt;<p class="jove_content"&gt; * Das sind Materialien vorgeschlagen. Können mit jeder kompatiblen Produkte ersetzt werden</p

retrograde loading of nerves, tracts, and spinal roots with fluorescent dyes

Retrograde Markierung von Neuronen ist ein Standard anatomische Methode, die auch 1,2 verwendet wurde, um Calcium und spannungsabhängigen Farbstoffen in Neuronen 3-6 geladen. Im Allgemeinen werden die Farbstoffe als feste Kristalle oder durch lokale Injektion mit Druck Glaspipetten aufgebracht. Dies kann jedoch zu einer Verdünnung des Farbstoffs und vermindertem Markierungsintensität führen, insbesondere wenn mehrere Stunden Farbstoffdiffusion erforderlich sind. Hier zeigen wir eine einfache und kostengünstige Technik zur Einführung fluoreszierenden und Ionen-sensitiven Farbstoffen in Neuronen ein Polyethylen Saugpipette mit dem Farbstoff-Lösung gefüllt. Dieses Verfahren bietet eine zuverlässige Möglichkeit zur Aufrechterhaltung einer hohen Konzentration des Farbstoffs in Kontakt mit Axone in der Ladevorgang.

Watch this Scientific Journal Video about Retrograde Loading of Nerves, Tracts, and Spinal Roots with Fluorescent Dyes at JoVE.com

Retrograde Loading of Nerves, Tracts, and Spinal Roots with Fluorescent Dyes

Wir beschreiben ein einfaches und kostengünstiges Verfahren zum Einführen hohe Konzentration von fluoreszierenden und Calcium-sensitiven Farbstoffen in Neuronen oder neuronaler ein Polyethylen Saugpipette.

Retrograde Loading von Nerven, Tracts, und Spinalwurzeln mit Fluoreszenzfarbstoffen

Retrograde labeling of neurons is a standard anatomical method1,2 that has also been used to load calcium and voltage-sensitive dyes into neurons3-6. ...

retrograde-loading-nerves-tracts-spinal-roots-with-fluorescent

Research

JoVE Journal

Neuroscience

12.2K Aufrufe.  National Institute of Neurological Disorders and Stroke, National Institutes of Health. Wir beschreiben ein einfaches und kostengünstiges Verfahren zum Einführen hohe Konzentration von fluoreszierenden und Calcium-sensitiven Farbstoffen in Neuronen oder neuronaler ein Polyethylen Saugpipette.

Serie: Retrograde Loading of Nerves, Tracts, and Spinal Roots with Fluorescent Dyes

Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v), process that can be followed by monitoring the fluorescence intensity decrease (destaining).

Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.

FM dyes have been of invaluable help in the understanding of synaptic dynamics. FMs are normally followed under the fluorescent microscope during different stimulation conditions. However, photoconversion of FM dyes combined with electron microscopy allows the visualization of distinct synaptic vesicle pools, among other ultrastructure components, in synaptic boutons.

Studium synaptischer Vesikel Pools mit Photokonversion Styrylfarbstoffe

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The ...

Forschung

Neurowissenschaften

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of its convenient genetics and fully sequenced genome. Additionally, the larval nervous system is an ideal model system to study mechanisms of axonal transport as the larval segmental nerves contain bundles of axons with their cell bodies located within the brain and their nerve terminals ending along the length of the body. Here we describe the procedure for visualization of synaptic vesicle proteins within larval segmental nerves. If done correctly, all components of the nervous system, along with associated tissues such as muscles and NMJs, remain intact, undamaged, and ready to be visualized. 3rd instar larvae carrying various mutations are dissected, fixed, incubated with synaptic vesicle antibodies, visualized and compared to wild type larvae. This procedure can be adapted for several different synaptic or neuronal antibodies and changes in the distribution of a variety of proteins can be easily observed within larval segmental nerves.

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster larvae provide an ideal model system to investigate the mechanisms of axonal transport within larval segmental nerves. Using this procedure, 3rd instar larvae carrying various mutations can be compared to wild type larvae.

Visualisierung von Larven segmentalen Nerven in 3 Rd Instar Drosophila Larven Vorbereitungen

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of ...

Optical Imaging of Neurons in the Crab Stomatogastric Ganglion with Voltage-sensitive Dyes

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous activity of participating neurons leads to the emergence of the integral functionality of the network. Here we present the methodology of application of this technique to identified pattern generating neurons in the crab stomatogastric ganglion. We demonstrate the loading of these neurons with the fluorescent voltage-sensitive dye Di-8-ANEPPQ and we show how to image the activity of dye loaded neurons using the MiCAM02 high speed and high resolution CCD camera imaging system. We demonstrate the analysis of the recorded imaging data using the BVAna imaging software associated with the MiCAM02 imaging system. The simultaneous voltage-sensitive dye imaging of the detailed activity of multiple neurons in the crab stomatogastric ganglion applied together with traditional electrophysiology techniques (intracellular and extracellular recordings) opens radically new opportunities for the understanding of how central pattern generator neural networks work.

Here we present the methodology for fast and high resolution fluorescent voltage-sensitive dye imaging of detailed activity of neurons in the crab stomatogastric ganglion.

Optical Imaging von Neuronen im Crab stomatogastrischen Ganglion mit spannungsabhängigen Farbstoffen

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

Retrograde Fluoreszenzmarkierung Ermöglicht Gezielte Extracellular Einzel-Einheit Aufnahme von identifizierten Neuronen<em&gt; In vivo</em

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Promotion of Survival and Differentiation of Neural Stem Cells with Fibrin and Growth Factor Cocktails after Severe Spinal Cord Injury

Neural stem cells (NSCs) can self-renew and differentiate into neurons and glia. Transplanted NSCs can replace lost neurons and glia after spinal cord injury (SCI), and can form functional relays to re-connect spinal cord segments above and below a lesion. Previous studies grafting neural stem cells have been limited by incomplete graft survival within the spinal cord lesion cavity. Further, tracking of graft cell survival, differentiation, and process extension had not been optimized. Finally, in previous studies, cultured rat NSCs were typically reported to differentiate into glia when grafted to the injured spinal cord, rather than neurons, unless fate was driven to a specific cell type. To address these issues, we developed new methods to improve the survival, integration and differentiation of NSCs to sites of even severe SCI. NSCs were freshly isolated from embryonic day 14 spinal cord (E14) from a stable transgenic Fischer 344 rat line expressing green fluorescent protein (GFP) and were embedded into a fibrin matrix containing growth factors; this formulation aimed to retain grafted cells in the lesion cavity and support cell survival. NSCs in the fibrin/growth factor cocktail were implanted two weeks after thoracic level-3 (T3) complete spinal cord transections, thereby avoiding peak periods of inflammation. Resulting grafts completely filled the lesion cavity and differentiated into both neurons, which extended axons into the host spinal cord over remarkably long distances, and glia. Grafts of cultured human NSCs expressing GFP resulted in similar findings. Thus, methods are defined for improving neural stem cell grafting, survival and analysis of in vivo findings.

Fibrin matrices containing growth factors were used to retain grafted neural stem cells into sites of complete spinal cord transection. Grafted cells completely filled the lesion cavity and differentiated into multiple neural cell types, including neurons that extended axons into host spinal cord over long distances.

Förderung von Überleben und die Differenzierung neuraler Stammzellen mit Fibrin und Growth Factor Cocktails nach schweren Rückenmarksverletzungen

Neural stem cells (NSCs) can self-renew and differentiate into neurons and glia. Transplanted NSCs can replace lost neurons and glia after spinal cord ...

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs survival and regeneration experiments. Many studies have been performed in mammalian animals to research RGCs survival after optic nerve injury. However, retrograde labeling of RGCs in adult zebrafish has not yet been reported, though some alternative methods can count cell numbers in retinal ganglion cell layers (RGCL). Considering the small size of the adult zebrafish skull and the high risk of death after drilling on the skull, we open the skull with the help of acid-etching and seal the hole with a light curing bond, which could significantly improve the survival rate. After absorbing the dyes for 5 days, almost all the RGCs are labeled. As this method does not need to transect the optic nerve, it is irreplaceable in the research of RGCs survival after optic nerve crush in adult zebrafish. Here, we introduce this method step by step and provide representative results.

We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

Retrograde Markierung von retinalen Ganglienzellen im erwachsenen Zebrafisch mit Fluoreszenzfarbstoffen

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs ...

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

In-vivo-Elektrophysiologische Messungen an Maus Ischiasnerven

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

Imaging Membrane Potential with Two Types of Genetically Encoded Fluorescent Voltage Sensors

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate goal is to image neuronal activity in vivo, one must be able to image activity of a single cell to ensure successful in vivo preparations. This procedure will describe how to image membrane potential in a single cell to provide a foundation to eventually image in vivo. Here we describe methods for imaging GEVIs consisting of a voltage-sensing domain fused to either a single fluorescent protein (FP) or two fluorescent proteins capable of F&#246;rster resonance energy transfer (FRET) in vitro. Using an image splitter enables the projection of images created by two different wavelengths onto the same charge-coupled device (CCD) camera simultaneously. The image splitter positions a second filter cube in the light path. This second filter cube consists of a dichroic and two emission filters to separate the donor and acceptor fluorescent wavelengths depending on the FPs of the GEVI. This setup enables the simultaneous recording of both the acceptor and donor fluorescent partners while the membrane potential is manipulated via whole cell patch clamp configuration. When using a GEVI consisting of a single FP, the second filter cube can be removed allowing the mirrors in the image splitter to project a single image onto the CCD camera.

A method for imaging changes in membrane potential using genetically encoded voltage indicators is described.

Imaging Membrane Potential mit zwei Arten von genetisch kodierten Fluorescent Spannungssensoren

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate ...

Retrograde Neuroanatomical Tracing of Phrenic Motor Neurons in Mice

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic nerves innervating the respiratory diaphragm. In spinal cord slices, phrenic motor neurons cannot be identified from other motor neurons on morphological or biochemical criteria. We provide the description of procedures for visualizing phrenic motor neuron cell bodies in mice, following intrapleural injections of cholera toxin subunit beta (CTB) conjugated to a fluorophore. This fluorescent neuroanatomical tracer has the ability to be caught up at the diaphragm neuromuscular junction, be carried retrogradely along the phrenic axons and reach the phrenic cell bodies. Two methodological approaches of intrapleural CTB delivery are compared: transdiaphragmatic versus transthoracic injections. Both approaches are successful and result in similar number of CTB-labeled phrenic motor neurons. In conclusion, these techniques can be applied to visualize or quantify the phrenic motor neurons in various experimental studies such as those focused on the diaphragm-phrenic circuitry.

Here, we describe a protocol for identifying phrenic motor neurons in mice after intrapleural delivery of fluorophore conjugated cholera toxin subunit beta. Two techniques are compared to inject the pleural cavity: transdiaphragmatic versus transthoracic approaches.

Retrograde neuroanatomischen Tracing der Phrenicus Motoneuronen bei Mäusen

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic ...

Activity-based Training on a Treadmill with Spinal Cord Injured Wistar Rats

Spinal cord injury (SCI) results in lasting deficits that include both mobility and a multitude of autonomic-related dysfunctions. Locomotor training (LT) on a treadmill is widely used as a rehabilitation tool in the SCI population with many benefits and improvements to daily life. We utilize this method of activity-based task-specific training (ABT) in rodents after SCI to both elucidate the mechanisms behind such improvements and to enhance and improve upon existing clinical rehabilitation protocols. Our current goal is to determine the mechanisms underlying ABT-induced improvements in urinary, bowel, and sexual function in SCI rats after a moderate to severe level of contusion. After securing each individual animal in a custom-made adjustable vest, they are secured to a versatile body weight support mechanism, lowered to a modified three-lane treadmill and assisted in step-training for 58 minutes, once a day for 10 weeks. This setup allows for the training of both quadrupedal and forelimb-only animals, alongside two different non-trained groups. Quadrupedal-trained animals with body weight support are aided by a technician present to assist in stepping with proper hind limb placement as necessary, while forelimb-only trained animals are raised at the caudal end to ensure no hind limb contact with the treadmill and no weight-bearing. One non-trained SCI group of animals is placed in a harness and rests next to the treadmill, while the other control SCI group remains in its home cage in the training room nearby. This paradigm allows for the training of multiple SCI animals at once, thus making it more time-efficient in addition to ensuring that our pre-clinical animal model mimics the clinical representation as close as possible, particularly with respect to the body weight support with manual assistance.

This protocol demonstrates our model of activity-based locomotor treadmill training for rats with spinal cord injury (SCI). Included is both quadrupedal and forelimb-only groups, in addition to two distinct types of non-trained control groups. Investigators are able to assess training effects on SCI rats using this protocol.

Activity-based Training auf einem Laufband mit Rückenmark verletzt Wistar-Ratten

Spinal cord injury (SCI) results in lasting deficits that include both mobility and a multitude of autonomic-related dysfunctions. Locomotor training (LT) ...

Retrograde Loading von Nerven, Tracts, und Spinalwurzeln mit Fluoreszenzfarbstoffen

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