This work was supported by a National Health &#38; Medical Research Centre (NHMRC) project grant to R.M.

All experimental procedures described here complied with the Animal Care and Ethics Committee of UNSW Australia and were performed in accordance with the National Health and Medical Research Council of Australia regulations for animal experimentation. All procedures in this protocol should be performed in accordance with the requirement of the relevant Animal Care and Ethics Committee.
1. Acetylcholinesterase Histochemical Staining
<ol>
	<li>Prepare the Acetylcholinesterase reaction mixture
	<ol>
		<li>Add 290 mg of Acetylthiocholine iodide to 200 ml of 0.1 M phosphate buffer (PB). Mix with a magnetic stirrer at 900 rpm.</li>
		<li>Add 600 mg of glycine under continuous stirring.</li>
		<li>Slowly add 420 mg of copper sulphate to the reaction mixture under continuous stirring until the product is completely dissolved.</li>
	</ol>
	</li>
	<li>Remove the skin on the animal carcass (obtained through tissue sharing) by making incisions into the skin of a perfused rat or mouse, grasping the skin from the neck area and pulling it past its feet. Ensure that the fascia covering each muscle is either removed or significantly perforated to ensure ample exposure of the muscle fibres to the reaction solution.</li>
	<li>Immerse the entire body or limb of interest into the reaction mixture and incubate O/N&#160;at 4&#160;&#176;C.</li>
	<li>Wash the carcass for 2 min in distilled water. Note: At this stage, the muscles exhibit a blue coloration and the motor end plates (MEPs) can be seen as white dots.</li>
	<li>Expose the carcass to a 10% ammonium sulphide solution for 3-5 sec. 
	Note: The muscle fibres will rapidly turn brown and the MEPs will be observable as black speckled dots. If the reaction occurs too quickly, and the muscle fibres are stained too dark to distinguish between the MEPs and the muscle fibres try using lower concentrations of ammonium sulphide solution (e.g., 5%). The reaction time to get the optimal contrast between the MEPs and the muscle fibres varies from one sample to the other. Therefore it is difficult to define the exact exposure time to the reagent. The reaction should be closely monitored and stopped when the contrast is deemed adequate.</li>
	<li>Wash the carcass two times, with agitation&#160;in distilled water and&#160;gently pat the carcass dry.</li>
	<li>Photograph both the lateral and medial aspects of the limb, ensuring that the MEPs for all the muscles of interest are captured.</li>
</ol>
2. Intramuscular Injections at the Motor End Plates
<ol>
	<li>Pull glass micropipettes with a micropipette puller (the use of graded micropipettes with plungers is recommended). With the help of a dissecting microscope, break the tips of the micropipettes with a pair of forceps such that the internal diameter of the lumen of the micropipette is approximately 0.5 mm.</li>
	<li>Ensure that all surgical instruments used are freshly autoclaved and that the surgical area is sterile.</li>
	<li>Fill the micropipettes with Fluoro-Gold (5% in distilled water).</li>
	<li>Induce anaesthetise in the animal in an induction chamber with Isofluorane (4% in O2). Check for righting reflex and ensure its absence before removing it from the induction chamber. Secure a nose cone over the snout of the rat and deliver Isofluorane (2% in O2) for maintenance of anaesthesia. Pinch the animal&#8217;s toes and gently touch the animal&#8217;s eye to ensure that both the pedal withdrawal and the corneal reflexes are absent. Note: A mixture of ketamine and xylazil can also be used (80&#160;and 10 mg/kg respectively, delivered intraperitoneally). Ketamine is a Schedule 8 (&#8220;Controlled&#8221;) drug and necessary protocols associated with the purchase, use, storage and disposal of the drug will have to be adhered to.</li>
	<li>Apply eye lubricant to prevent drying of the eyes during the course of the procedure.</li>
	<li>Shave the targeted limb and use gauze to wipe the shaved area with three alternating scrubs of chlorhexidine and 70% alcohol.&#160;&#160;Transfer the animal to the surgical area.</li>
	<li>Place the animal on a clean underpad and position it appropriately to ensure good access to the targeted muscle(s).</li>
	<li>Use a pair of forceps with tooth grips to lift the skin over the targeted muscle away from the underlying musculature and make an incision in the skin with surgical scissors. Ensure that the incision is large enough to completely expose the muscle(s) of interest. Ensure that there is minimal disruption to the fascia.</li>
	<li>Use the MEP photographs to mentally transpose the location and shape of the MEP region onto the muscle(s). 
	Note: A fine felt marker could assist to reproduce the MEP region from the photograph onto the muscle(s) of interest.</li>
	<li>Perform multiple injections along the full length of the MEP region (for Fluoro-Gold, 3 to 4 injections of 1-2 &#181;l each should be sufficient). Gently wipe the muscle(s) to remove any seepage.</li>
	<li>Bring the two ends of the incised skin close together with blunt forceps and close the wound with surgical clips. Infiltrate 0.1 ml Bupivacaine (0.5% in water) (or other local anaesthetics) along the entire span of the wound.</li>
	<li>Turn the anaesthetic machine off and monitor the animal until it has is fully recovered from anaesthesia.</li>
	<li>Wait for 14 days between administration of intra-muscular injections and the perfusion of the animals.</li>
</ol>
3. Perfusions
<ol>
	<li>Administer a lethal dose of pentobarbitone sodium solution (e.g. Lethabarb; 150 mg/kg) to the animal by intraperitoneal injection.
	<ol>
		<li>Closely monitor the animal until a deep level of anaesthesia is reached, as confirmed by the absence of the pedal withdrawal and the corneal reflexes.</li>
	</ol>
	</li>
	<li>Position the animal on its back on a dissecting board placed over a perfusion sink.</li>
	<li>Use a pair of forceps with tooth grips to lift the skin covering the base of the sternum. Make a small skin incision with a pair of strong surgical scissors immediately below the sternum and then use the forceps to grip the xiphoid cartilage of the sternum. While holding the xiphoid cartilage, enlarge the incision on both sides of the chest cavity, from the sternum to the armpits.</li>
	<li>Cut the diaphragm to expose the heart.
	<ol>
		<li>Inject 0.ml of heparin directly into the apex of the heart to prevent coagulation of the blood.</li>
		<li>Cut the apex to allow the insertion of a cannula into the left ventricle, and then quickly clamp the cannula in place with the aid of a haemostat.</li>
		<li>Make an incision in the right atrium with a pair of fine scissors and immediately start the peristaltic pump. Perfuse with 0.1 M PB until the liquid flowing out of the atrium is nearly free of blood and the color of the liver becomes light brown. Then, perfuse with a solution of paraformaldehyde (4% in 0.1 M PB) until the entire body of the animal becomes rigid.</li>
	</ol>
	</li>
</ol>
4. Cervical Spinal Cord Dissection and Preparation for Histology
<ol>
	<li>Lay the animal carcass on its abdomen.</li>
	<li>Cut the skin at the body&#8217;s midline with a scalpel and reflect it from the base of the skull to the level of the iliac bone.</li>
	<li>Cut through and reflect the paravertebral muscles to expose the dorsal aspect of the vertebral column.</li>
	<li>Identify the bony spinous process of Atlas, the second cervical segment (i.e., C2), and remove it with a pair of surgical rongeurs or forceps to locate the corresponding underlying dorsal root.
	<ol>
		<li>Identify the right C2 root by coloring it with a permanent felt marker.</li>
		<li>Remove the next vertebrae one by one and mark the right dorsal roots with alternating colors (i.e., C2, C4, C6, and C8 can be colored in green and C3, C5, C7, T1 can be colored in blue).</li>
	</ol>
	</li>
	<li>Use a small surgical needle (e.g., 30 G gauge needle) to gently pierce through the dura covering the lower end of the spinal cord. With the bevel of the needle facing up, lift the dura away from the cord while moving the needle rostrally to make a longitudinal slit.
	<ol>
		<li>Reflect the dura.</li>
	</ol>
	</li>
	<li>Cut the spinal cord transversally into one- or two-segment blocks with a new scalpel blade.
	<ol>
		<li>Leave the spinal cord segments in situ and, for each block, carefully make a small fiducial mark on the left side of each block with the scalpel blade half way between two adjacent roots. Ensure that the fiducial mark is deep enough through the tissue as to be visible from the ventral aspect of the blocks. Make the fiducial mark at an angle of approximately 45&#176;, pointing either anteriorly or posteriorly with respect to animal anatomy, to aid in the orientation of the tissue segment following dissection.</li>
	</ol>
	</li>
	<li>Collect the individual blocks into small, clearly labelled bottles containing a solution of paraformaldehyde (4% in 0.1 M PB) and leave at RT O/N.</li>
	<li>Transfer the blocks into clean, clearly labelled bottles containing a solution of sucrose (30% in 0.1 M PB) and keep at 4 &#176;C for at least two days.</li>
	<li>Place the segments in cryo-moulds and covered them with tissue freezing medium. For longitudinal histological preparation, orient the blocks with the dorsal aspect facing up and use the fiducial mark to orient the block in the cryo-moulds.</li>
	<li>Freeze the cryo-moulds at -20&#160;&#176;C and cut with a cryostat in 50 &#181;m-thick sections. Note: The sections can be directly mounted on adhesion microscope slides and left to dry O/N. Alternatively, the sections can be floated in 48-well plates filled with 0.1 M PB and subsequently mounted using the fiducial mark to orient the tissue sections on the slides.</li>
</ol>
5. Lumbar Spinal Cord Dissection and Preparation for Histology
<ol>
	<li>Lay the animal carcass on it&#8217;s back.</li>
	<li>Perform a midline incision along the abdomen of the animal and remove the viscera. Dissect out the muscles of the posterior abdominal wall to expose the ventral aspect of the vertebral column.</li>
	<li>Locate the short caudal-most rib and its adjoining T13 vertebra where the T13 ventral root exits the bone.
	<ol>
		<li>Consecutively remove one by one the two or three vertebrae rostral to T13 (i.e., T12, T11 and, if needed, T10) with fine surgical rongeurs to follow the T13 ventral root until its point of entry in the ventral aspect of the ventral cord and mark it with a permanent felt marker.</li>
		<li>From this point on, repeat the same procedure to identify the location of the ventral root entry points for L1, L2, L3, L4, L5, L6 and S1, coloring them with alternating colors.</li>
	</ol>
	</li>
	<li>Follow the same procedure as described in Section 4.5-4.10 for lumbar spinal cord dissection.</li>
</ol>

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The authors declare that they have no competing financial interests.

Intramuscular targeting and subsequent retrograde transfer of therapeutic transgenes to the corresponding &#945; motor neurons for the experimental treatment of neuromuscular condition is not a new strategy. For instance, this delivery method has been used to delay neuromuscular degeneration at different stages of the ALS progression in SOD1 mice and rats 1,9-12 as well as in mice with SMA 13. Whilst promising, the efficacy of these gene therapy scenarios has been limited. In this regard, we propose...

The loss of control of voluntary movement that results from neurological conditions such as motor neuron disease and spinal- as well as Duchenne muscular atrophy is a debilitating condition that has high and long lasting impact on the every day life of affected individuals. Over the last decade, research efforts aiming to stop or at least delay the deleterious effects of these neuromuscular diseases has been a priority for many clinicians and scientist around the world. In this regard, the recent generation of animal models that mimic these neuromuscular diseases has been instrumental in obtaining fundamental insights into the physiological mechanisms underlying the development and progression of these conditions 1-13. Treatment of these neuromuscular disease requires direct access to the spinal cord and can be achieved by spinal cord injections 14,15. Recent advances in gene therapy have also targeted the striated muscles of the upper and lower limbs to shuttle therapeutic genes to the corresponding &#945; motor neurons that are located within the ventral horn of the spinal cord 1,9-13. However, this once promising strategy has failed to improve the outcome of these neurological conditions. While it is fair to conclude that these poor outcomes could be, at least partly, be attributable to the low efficacy of these protective genes, one cannot exclude the low efficacy of these gene delivery methods.
Motor end plates (MEPs) are specialized regions of the skeletal myofibres that are indented by the axon terminals of large peripheral motor fibres originating from &#945; motor neurons. Together, the peripheral nerve fibre endings and the MEPs form the neuromuscular junction, i.e., the site where synaptic impulses are triggered by the anterograde release of the neurotransmitter, acetylcholine. Importantly, the relationship between peripheral nerve fibres and the MEPs is bi-directional, although different motors are responsible for the transport of molecules and organelles towards as well as away from the neuron somata 16-18. In light of these anatomical considerations, MEPs appear to be the targets of choice for the delivery and subsequent retrograde transport of genetic material to the corresponding motor neurons. In this context, it is not surprising that the success of motor neuron transduction greatly depends on the distance between the intramuscular injection of viral vectors and the muscle&#8217;s MEPs 19-20. Surprisingly, however, the exact location of the MEP zones on the myofibres of the laboratory rat and mouse, the two species of choice to model neuromuscular diseases, were not available until recently.
We have produced comprehensive maps of the MEP region for several forelimb muscles in the rat and the mouse 21-22. More recently, we have shown the details of the organization of the MEP region for several muscles of the mouse hindlimb 23 and we are currently analysing the features of the MEPs on the rat hindlimb. In our hands, intramuscular injections of retrograde tracers directed to the entire MEP zones in these muscles gave rise to more labelled motor neurons that are spanning more spinal cord segments than previously reported. Here we present the protocol that has been developed over the last few years to reveal the location of the MEPs on the external surface as well as throughout the depth of hindlimb and forelimb muscles in both the mouse and the rat.

<table><tbody><tr><td>Fluoro-Gold</td><td>Fluorochrome, LLC</td><td>Nil</td><td>Diluted to 5%</td></tr><tr><td>Drummond PCR Micropipets 1-10 &amp;micro;l</td><td>Drummond Scientific</td><td>5-000-1001-X10</td><td>accompanied with plungers</td></tr><tr><td>Acetylthiocholine Iodide</td><td>Sigma Life Science</td><td>A5751-25G</td><td></td></tr><tr><td>Copper(II) Sulfate Anhydrous</td><td>Sigma-Aldrich</td><td>61230-500G-F</td><td></td></tr><tr><td>Tissue-Tek O.C.T Compound</td><td>Sakura Finetek</td><td>25608-930</td><td></td></tr><tr><td>Glycine</td><td>Ajax Finechem</td><td>1083-500G</td><td></td></tr><tr><td>Dextran, Tetramethylrhodamine and biotin</td><td>Life Technologies</td><td>D-3312</td><td>Diliuted in distilled water</td></tr><tr><td>Isothesia</td><td>Provet</td><td>ISOF00</td><td>1000 mg/g Isoflurane inhalation vapour</td></tr><tr><td>Autoclip 9 mm Wound Clips</td><td>Texas Scientific Instruments, LLC</td><td>205016</td><td></td></tr><tr><td>Lethabarb Enthanasia Injection</td><td>Virbac (Australia) Pty Ltd.</td><td>LETHA450</td><td></td></tr><tr><td>Formaldehyde Solution</td><td>Ajax Finechem</td><td>A809-2.5L PL</td><td></td></tr><tr><td>SuperFrost Plus glass slides</td><td>Menzel-Glaser</td><td>J1800AMNZ</td><td></td></tr><tr><td>Ammonium Sulphide</td><td>Sigma-Aldrich</td><td>A1952</td><td>Diluted to 10%</td></tr><tr><td>Marcain Spinal 0.5% (Bupivacaine hydrochloride)</td><td>Astrazenca</td><td></td><td>Diluted to 0.25%</td></tr></tbody></table>

intramuscular injections along the motor end plates: a minimally invasive approach to shuttle tracers directly into motor neurons

Diseases affecting the integrity of spinal cord motor neurons are amongst the most debilitating neurological conditions. Over the last decades, the development of several animal models of these neuromuscular disorders has provided the scientific community with different therapeutic scenarios aimed at delaying or reversing the progression of these conditions. By taking advantage of the retrograde machinery of neurons, one of these approaches has been to target skeletal muscles in order to shuttle therapeutic genes into corresponding spinal cord motor neurons. Although once promising, the success of such gene delivery approach has been hampered by the sub-optimal number of transduced motor neurons it has so far shown to yield. Motor end plates (MEPs) are highly specialized regions on the skeletal musculature that are in direct synaptic contact to the spinal cord &#945; motor neurons. In this regard, it is important to note that, so far, the efforts to retrogradely transfer genes into motor neurons were made without reference to the location of the MEP region in the targeted muscles. Here, we describe a simple protocol 1) to reveal the exact location of the MEPs on the surface of skeletal muscles and 2) to use this information to guide the intramuscular delivery and subsequent optimal retrograde transport of retrograde tracers into motor neurons. We hope to utilize the results from these tracing experiments in further studies into investigating retrograde transport of therapeutic genes to spinal cord motor neurons through the targeting of MEPs.

Acetylcholinesterase histochemical staining reveals the location of the motor end plates across the width of the muscles. Figure 1 illustrates the results of such staining performed on a whole rat forelimb. It is suggested to optimise the concentration of the ammonium sulphide solution (e.g., 5-7% instead of 10%) as well as the time the specimen is immersed in the solution if the non-specific background staining on the muscles fibres is too excessive. Figure 2 illustrates a colu...

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Intramuscular Injections Along the Motor End Plates: A Minimally Invasive Approach to Shuttle Tracers Directly into Motor Neurons

The efficacy of intramuscular uptake and retrograde transport of molecules to corresponding motor neurons depends on the location of the injection sites with respect to the motor end plates (MEPs). Here, we describe how to locate MEPs on skeletal muscles to optimise retrograde transport of tracers into motor neurons.

Diseases affecting the integrity of spinal cord motor neurons are amongst the most debilitating neurological conditions. Over the last decades, the ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

12.5K Views.  University of New South Wales. The efficacy of intramuscular uptake and retrograde transport of molecules to corresponding motor neurons depends on the location of the injection sites with respect to the motor end plates (MEPs). Here, we describe how to locate MEPs on skeletal muscles to optimise retrograde transport of tracers into motor neurons.

Video: Intramuscular Injections Along the Motor End Plates: A Minimally Invasive Approach to Shuttle Tracers Directly into Motor Neurons

Subcutaneous Administration of Muscarinic Antagonists and Triple-Immunostaining of the Levator Auris Longus Muscle in Mice

Hind limb muscles of rodents, such as gastrocnemius and tibialis anterior, are frequently used for in vivo pharmacological studies of the signals essential for the formation and maintenance of mammalian NMJs. However, drug penetration into these muscles after subcutaneous or intramuscular administration is often incomplete or uneven and many NMJs can remain unaffected. Although systemic administration with devices such as mini-pumps can improve the spatiotemporal effects, the invasive nature of this approach can cause confounding inflammatory responses and/or direct muscle damage. Moreover, complete analysis of the NMJs in a hind limb muscle is challenging because it requires time-consuming serial sectioning and extensive immunostaining. 
The mouse LAL is a thin, flat sheet of muscle located superficially on the dorsum of the neck. It is a fast-twitch muscle that functions to move the pinna. It contains rostral and caudal portions that originate from the midline of the cranium and extend laterally to the cartilaginous portion of each pinna. The muscle is supplied by a branch of the facial nerve that projects caudally as it exits the stylomastoid foramen. We and others have found LAL to be a convenient preparation that offers advantages for the investigation of both short and long-term in vivo effects of drugs on NMJs and muscles. First, its superficial location facilitates multiple local applications of drugs under light anesthesia. Second, its thinness (2-3 layers of muscle fibers) permits visualization and analysis of almost all the NMJs within the muscle. Third, the ease of dissecting it with its nerve intact together with the pattern of its innervation permits supplementary electrophysiological analysis in vitro9,5. Last, and perhaps most importantly, a small applied volume (&tilde;50&mu;l) easily covers the entire muscle surface, provides a uniform and prolonged exposure of all its NMJs to the drug and eliminates the need for a systemic approach1,8.

We describe procedures for repeated administration of inhibitors of muscarinic signaling to the levator auris longus (LAL) muscle of young adult mice and for subsequent immunostaining of its neuromuscular junctions (NMJs) in wholemounts. The LAL muscle has unique advantages for revealing in vivo pharmacological effects on NMJs.

Hind limb muscles of rodents, such as gastrocnemius and tibialis anterior, are frequently used for in vivo  pharmacological studies of the signals ...

Revealing Neural Circuit Topography in Multi-Color

Neural circuits are organized into functional topographic maps. In order to visualize complex circuit architecture we developed an approach to reliably label the global patterning of multiple topographic projections. The cerebellum is an ideal model to study the orderly arrangement of neural circuits. For example, the compartmental organization of spinocerebellar mossy fibers has proven to be an indispensable system for studying mossy fiber patterning. We recently showed that wheat germ agglutinin (WGA) conjugated to Alexa 555 and 488 can be used for tracing spinocerebellar mossy fiber projections in developing and adult mice (Reeber et al. 2011). We found three major properties that make the WGA-Alexa tracers desirable tools for labeling neural projections. First, Alexa fluorophores are intense and their brightness allows for wholemount imaging directly after tracing. Second, WGA-Alexa tracers label the entire trajectory of developing and adult neural projections. Third, WGA-Alexa tracers are rapidly transported in both retrograde and anterograde directions. Here, we describe in detail how to prepare the tracers and other required tools, how to perform the surgery for spinocerebellar tracing and how best to image traced projections in three dimensions. In summary, we provide a step-by-step tracing protocol that will be useful for deciphering the organization and connectivity of functional maps not only in the cerebellum but also in the cortex, brainstem, and spinal cord.

We provide a practical guide for delivering tracers in vivo and use the spinocerebellar pathway as a model system to demonstrate essential steps for successful neuronal circuit analysis in mice. We describe in detail our versatile tracing protocol that exploits wheat germ agglutinin (WGA) conjugated to Alexa fluorophores.

Neural circuits are organized into functional topographic maps. In order to visualize complex circuit architecture we developed an approach to reliably ...

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

The spinal motoneuron has long been a good model system for studying neural function because it is a neuron of the central nervous system with the unique properties of (1) having readily identifiable targets (the muscle fibers) and therefore having a very well-known function (to control muscle contraction); (2) being the convergent target of many spinal and descending networks, hence the name of "final common pathway"; and (3) having a large soma which makes it possible to penetrate them with sharp intracellular electrodes. Furthermore, when studied in vivo, it is possible to record simultaneously the electrical activity of the motoneurons and the force developed by their muscle targets. Performing intracellular recordings of motoneurons in vivo therefore put the experimentalist in the unique position of being able to study, at the same time, all the compartments of the "motor unit" (the name given to the motoneuron, its axon, and the muscle fibers it innervates1): the inputs impinging on the motoneuron, the electrophysiological properties of the motoneuron, and the impact of these properties on the physiological function of the motoneurons, i.e. the force produced by its motor unit. However, this approach is very challenging because the preparation cannot be paralyzed and thus the mechanical stability for the intracellular recording is reduced. Thus, this kind of experiments has only been achieved in cats and in rats. However, the study of spinal motor systems could make a formidable leap if it was possible to perform similar experiments in normal and genetically modified mice.
For technical reasons, the study of the spinal networks in mice has mostly been limited to neonatal in vitro preparations, where the motoneurons and the spinal networks are immature, the motoneurons are separated from their targets, and when studied in slices, the motoneurons are separated from most of their inputs. Until recently, only a few groups had managed to perform intracellular recordings of motoneurons in vivo2-4 , including our team who published a new preparation which allowed us to obtain very stable recordings of motoneurons in vivo in adult mice5,6. However, these recordings were obtained in paralyzed animals, i.e. without the possibility to record the force output of these motoneurons. Here we present an extension of this original preparation in which we were able to obtain simultaneous recordings of the electrophysiological properties of the motoneurons and of the force developed by their motor unit. This is an important achievement, as it allows us to identify the different types of motoneurons based on their force profile, and thereby revealing their function. Coupled with genetic models disturbing spinal segmental circuitry7-9, or reproducting human disease10,11, we expect this technique to be an essential tool for the study of spinal motor system.

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

This new method permits the simultaneous intracellular recording of a single adult mouse motoneuron and the measurement of the force produced by its muscle fibers. The combined investigation of the electrical and mechanical properties of motor units in normal and genetically modified animals is a breakthrough for the study of the neuromuscular system.

The  spinal motoneuron has long been a good model system for studying neural  function because it is a neuron of the central nervous system with the ...

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.
This extracellular technique has advantages. First, extracellular electrodes can stimulate and record neurons through the sheath5, so it does not need to be removed. Thus, neurons will be healthier in extracellular experiments than in intracellular ones. Second, if ganglia are rotated by appropriate pinning of the sheath, extracellular electrodes can access neurons on both sides of the ganglion, which makes it easier and more efficient to identify multiple neurons in the same preparation. Third, extracellular electrodes do not need to penetrate cells, and thus can be easily moved back and forth among neurons, causing less damage to them. This is especially useful when one tries to record multiple neurons during repeating motor patterns that may only persist for minutes. Fourth, extracellular electrodes are more flexible than intracellular ones during muscle movements. Intracellular electrodes may pull out and damage neurons during muscle contractions. In contrast, since extracellular electrodes are gently pressed onto the sheath above neurons, they usually stay above the same neuron during muscle contractions, and thus can be used in more intact preparations.
To uniquely identify motor neurons for a motor pool (in particular, the I1/I3 muscle in Aplysia) using extracellular electrodes, one can use features that do not require intracellular measurements as criteria: soma size and location, axonal projection, and muscle innervation4,6,7. For the particular motor pool used to illustrate the technique, we recorded from buccal nerves 2 and 3 to measure axonal projections, and measured the contraction forces of the I1/I3 muscle to determine the pattern of muscle innervation for the individual motor neurons.
We demonstrate the complete process of first identifying motor neurons using muscle innervation, then characterizing their timing during motor patterns, creating a simplified diagnostic method for rapid identification. The simplified and more rapid diagnostic method is superior for more intact preparations, e.g. in the suspended buccal mass preparation8 or in vivo9. This process can also be applied in other motor pools10,11,12 in Aplysia or in other animal systems2,3,13,14.

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.

In animals with large  identified neurons (e.g. mollusks), analysis of motor pools is done using  intracellular techniques1,2,3,4. Recently,  we developed ...

Transsynaptic Tracing from Peripheral Targets with Pseudorabies Virus Followed by Cholera Toxin and Biotinylated Dextran Amines Double Labeling

Transsynaptic tracing has become a powerful tool used to analyze central efferents that regulate peripheral targets through multi-synaptic circuits. This approach has been most extensively used in the brain by utilizing the swine pathogen pseudorabies virus (PRV)1. PRV does not infect great apes, including humans, so it is most commonly used in studies on small mammals, especially rodents. The pseudorabies strain PRV152 expresses the enhanced green fluorescent protein (eGFP) reporter gene and only crosses functional synapses retrogradely through the hierarchical sequence of synaptic connections away from the infection site2,3. Other PRV strains have distinct microbiological properties and may be transported in both directions (PRV-Becker and PRV-Kaplan)4,5&#160;.&#160;This protocol will deal exclusively with PRV152. By delivering the virus at a peripheral site, such as muscle, it is possible to limit the entry of the virus into the brain through a specific set of neurons. The resulting pattern of eGFP signal throughout the brain then resolves the neurons that are connected to the initially infected cells. As the distributed nature of transsynaptic tracing with pseudorabies virus makes interpreting specific connections within an identified network difficult, we present a sensitive and reliable method employing biotinylated dextran amines (BDA) and cholera toxin subunit b (CTb) for confirming the connections between cells identified using PRV152. Immunochemical detection of BDA and CTb with peroxidase and DAB (3, 3&#39;-diaminobenzidine) was chosen because they are effective at revealing cellular processes including distal dendrites6-11.

Transsynaptic tracing has become a powerful tool for analyzing central efferents regulating peripheral targets through multi-synaptic circuits. Here we present a protocol that exploits the transsynaptic pseudorabies virus to identify and localize a functional brain circuit, followed by classical tract tracing techniques to validate specific connections in the circuit between identified groups of neurons.

Transsynaptic tracing has become a powerful tool used to analyze central efferents that regulate peripheral targets through multi-synaptic circuits. This ...

Implantation of Miniosmotic Pumps and Delivery of Tract Tracers to Study Brain Reorganization in Pathophysiological Conditions

Pharmacological treatment in animal models of cerebral disease imposes the problem of repeated injection protocols that may induce stress in animals and result in impermanent tissue levels of the drug. Additionally, drug delivery to the brain is delicate due to the blood brain barrier (BBB), thus significantly reducing intracerebral concentrations of selective drugs after systemic administration. Therefore, a system that allows both constant drug delivery without peak levels and circumvention of the BBB is in order to achieve sufficiently high intracerebral concentrations of drugs that are impermeable to the BBB. In this context, miniosmotic pumps represent an ideal system for constant drug delivery at a fixed known rate that eludes the problem of daily injection stress in animals and that may also be used for direct brain delivery of drugs. Here, we describe a method for miniosmotic pump implantation and post operatory care that should be given to animals in order to successfully apply this technique. We embed the aforementioned experimental paradigm in standard procedures that are used for studying neuroplasticity within the brain of C57BL6 mice. Thus, we exposed animals to 30 min brain infarct and implanted with miniosmotic pumps connected to the skull via a cannula in order to deliver a pro-plasticity drug. Behavioral testing was done during 30 days of treatment. After removal the animals received injections of anterograde tract tracers to analyze neuronal plasticity in the chronic phase of recovery. Results indicated that neuroprotection by the delivered drug was accompanied with increase in motor fibers crossing the midline of the brain at target structures. The results affirm the value of these techniques for drug administration and brain plasticity studies in modern neuroscience.

In order to study brain reorganization under pathological conditions we used miniosmotic pumps for direct protein delivery into the brain circumventing the blood brain barrier. Tract tracers are then injected to study alterations in brain connectivity under the influence of the protein.

Pharmacological treatment in animal models of cerebral disease imposes the problem of repeated injection protocols that may induce stress in animals and ...

Laser-guided Neuronal Tracing In Brain Explants

We present a technique which combines an in vitro tracer injection protocol, which uses a series of electrical and pressure pulses to increase dye uptake through electroporation in brain explants with targeted laser illumination and matching filter goggles during the procedure. The described technique of in vitro electroporation by itself yields relatively good visual control for targetting certain areas of the brain. By combining it with laser excitation of fluorescent genetic markers and their read-out through band-passing filter goggles, which can pick up the emissions of the genetically labeled cells/nuclei and the fluorescent tracing dye, a researcher can substantially increase the accuracy of injections by finding the area of interest and controlling for the dye-spread/uptake in the injection area much more efficiently. In addition, the laser illumination technique allows to study the functionality of a given neurocircuit by providing information about the type of neurons projecting to a certain area in cases where the GFP expression is linked to the type of transmitter expressed by a subpopulation of neurons.

We describe a technique to label neurons and their processes via anterograde or retrograde tracer injections into brain nuclei using an in vitro preparation. We modified an existing method of in vitro tracer electroporation by taking advantage of fluorescently labeled mouse mutants and basic optical equipment in order to increase labeling accuracy.

We present a technique which combines an in vitro tracer injection protocol, which uses a series of electrical and pressure pulses to increase dye uptake ...

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.

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

Direct Injection of a Lentiviral Vector Highlights Multiple Motor Pathways in the Rat Spinal Cord

Introducing proteins of interest into cells in the nervous system is challenging due to innate biological barriers that limit access to most molecules. Injection directly into spinal cord tissue bypasses these barriers, providing access to cell bodies or synapses where molecules can be incorporated. Combining viral vector technology with this method allows for introduction of target genes into nervous tissue for the purpose of gene therapy or tract tracing. Here a virus engineered for highly efficient retrograde transport (HiRet) is introduced at the synapses of propriospinal interneurons (PNs) to encourage specific transport to neurons in the spinal cord and brainstem nuclei. Targeting PNs takes advantage of the numerous connections they receive from motor pathways such as the rubrospinal and reticulospinal tracts, as well as their interconnection with each other throughout spinal cord segments. Representative tracing using the HiRet vector with constitutively active green fluorescent protein (GFP) shows high fidelity details of cell bodies, axons and dendritic arbors in thoracic PNs and in reticulospinal neurons in the pontine reticular formation. HiRet incorporates well into brainstem pathways and PNs but shows age dependent integration into corticospinal tract neurons. In summary, spinal cord injection using viral vectors is a suitable method for introduction of proteins of interest into neurons of targeted tracts.

This protocol demonstrates injection of a retrogradely transportable viral vector into rat spinal cord tissue. The vector is taken up at the synapse and transported to the cell body of target neurons. This model is suitable for retrograde tracing of important spinal pathways or targeting cells for gene therapy applications.

Introducing proteins of interest into cells in the nervous system is challenging due to innate biological barriers that limit access to most molecules. ...

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal transport have devastating consequences for individual neurons and their networks, and contribute to a plethora of neurological disorders. As many of these conditions involve both cell autonomous and non-autonomous mechanisms, and often display a spectrum of pathology across neuronal subtypes, methods to accurately identify and analyze neuronal subsets are imperative.
This paper details protocols to assess in vivo axonal transport of signaling endosomes and mitochondria in sciatic nerves of anesthetized mice. Stepwise instructions are provided to 1) distinguish motor from sensory neurons in vivo, in situ, and ex vivo by using mice that selectively express fluorescent proteins within cholinergic motor neurons; and 2) separately or concurrently assess in vivo axonal transport of signaling endosomes and mitochondria. These complementary intravital approaches facilitate the simultaneous imaging of different cargoes in distinct peripheral nerve axons to quantitatively monitor axonal transport in health and disease.

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Using transgenic fluorescent mice, detailed protocols are described to assess in vivo axonal transport of signaling endosomes and mitochondria within motor and sensory axons of the intact sciatic nerve in live animals.

Intramuscular Injections Along the Motor End Plates: A Minimally Invasive Approach to Shuttle Tracers Directly into Motor Neurons

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Subcutaneous Administration of Muscarinic Antagonists and Triple-Immunostaining of the Levator Auris Longus Muscle in Mice

Revealing Neural Circuit Topography in Multi-Color

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

Transsynaptic Tracing from Peripheral Targets with Pseudorabies Virus Followed by Cholera Toxin and Biotinylated Dextran Amines Double Labeling

Implantation of Miniosmotic Pumps and Delivery of Tract Tracers to Study Brain Reorganization in Pathophysiological Conditions

Laser-guided Neuronal Tracing In Brain Explants

Retrograde Neuroanatomical Tracing of Phrenic Motor Neurons in Mice

Direct Injection of a Lentiviral Vector Highlights Multiple Motor Pathways in the Rat Spinal Cord

Expanding the Toolkit for In Vivo Imaging of Axonal Transport