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>

<ol>
	<li>Kaspar, B. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Retrograde+Viral+Delivery+of+IGF-1+Prolongs+Survival+in+a+Mouse+ALS+Model.">Retrograde Viral Delivery of IGF-1 Prolongs Survival in a Mouse ALS Model.</a> Science. 301 (5634), (2003).</li><li>Ishiyama, T., Okada, R., Nishibe, H., Mitsumoto, H., Nakayama, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Riluzole+slows+the+progression+of+neuromuscular+dysfunction+in+the+wobbler+mouse+motor+neuron+disease.">Riluzole slows the progression of neuromuscular dysfunction in the wobbler mouse motor neuron disease.</a> Brain Res. 1019 (1-2), 226-236 (2004).</li><li>Turner, B. J., Parkinson, N. J., Davies, K. E., Talbot, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Survival+motor+neuron+deficiency+enhances+progression+in+an+amyotrophic+lateral+sclerosis+mouse+model.">Survival motor neuron deficiency enhances progression in an amyotrophic lateral sclerosis mouse model.</a> Neurobiol Dis. 34 (3), 511-517 (2009).</li><li>Wegorzewska, I., Bell, S., Cairns, N. J., Miller, T. M., Baloh, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=TDP-43+mutant+transgenic+mice+develop+features+of+ALS+and+frontotemporal+lobar+degeneration.">TDP-43 mutant transgenic mice develop features of ALS and frontotemporal lobar degeneration.</a> P Natl Acad Sci USA. 106 (44), 18809-18814 (2009).</li><li>Kimura, E., Li, S., Gregorevic, P., Fall, B. M., Chamberlain, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dystrophin+delivery+to+muscles+of+mdx+mice+using+lentiviral+vectors+leads+to+myogenic+progenitor+targeting+and+stable+gene+expression.">Dystrophin delivery to muscles of mdx mice using lentiviral vectors leads to myogenic progenitor targeting and stable gene expression.</a> Mol Ther. 18 (1), 206-213 (2009).</li><li>Van Den Bosch, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+rodent+models+of+amyotrophic+lateral+sclerosis.">Genetic rodent models of amyotrophic lateral sclerosis.</a> J Biomed Biotechnol. 2011 (6), 348765–11 (2011).</li><li>Pratt, S. J. P., Shah, S. B., Ward, C. W., Inacio, M. P., Stains, J. P., Lovering, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+in+vivo+injury+on+the+neuromuscular+junction+in+healthy+and+dystrophic+muscles.">Effects of in vivo injury on the neuromuscular junction in healthy and dystrophic muscles.</a> J Physiol. 591 (Pt 2), 559-570 (2013).</li><li>Garrett, C. A., 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=DYNC1H1+mutation+alters+transport+kinetics+and+ERK1/2-cFos+signalling+in+a+mouse+model+of+distal+spinal+muscular+atrophy).">DYNC1H1 mutation alters transport kinetics and ERK1/2-cFos signalling in a mouse model of distal spinal muscular atrophy).</a> Brain. 137 (Pt 7), 1883-1893 (2014).</li><li>Bordet, T., 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=Protective+effects+of+cardiotrophin-1+adenoviral+gene+transfer+on+neuromuscular+degeneration+in+transgenic+ALS+mice).">Protective effects of cardiotrophin-1 adenoviral gene transfer on neuromuscular degeneration in transgenic ALS mice).</a> Hum Mol Gen. 10 (18), 1925-1933 (2001).</li><li>Acsadi, G., 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=Increased+survival+and+function+of+SOD1+mice+after+glial+cell-derived+neurotrophic+factor+gene+therapy.">Increased survival and function of SOD1 mice after glial cell-derived neurotrophic factor gene therapy.</a> Hum Gene Ther. 13 (9), 1047-1059 (2002).</li><li>Azzouz, M., 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=VEGF+delivery+with+retrogradely+transported+lentivector+prolongs+survival+in+a+mouse+ALS+model.">VEGF delivery with retrogradely transported lentivector prolongs survival in a mouse ALS model.</a> Nature. 429 (6990), 413-417 (2004).</li><li>Suzuki, M., 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=Direct+muscle+delivery+of+GDNF+with+human+mesenchymal+stem+cells+improves+motor+neuron+survival+and+function+in+a+rat+model+of+familial+ALS.">Direct muscle delivery of GDNF with human mesenchymal stem cells improves motor neuron survival and function in a rat model of familial ALS.</a> Mol Ther. 16 (12), 2002-2010 (2008).</li><li>Benkhelifa-Ziyyat, S., 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=Intramuscular+scAAV9-SMN+Injection+Mediates+Widespread+Gene+Delivery+to+the+Spinal+Cord+and+Decreases+Disease+Severity+in+SMA+Mice.">Intramuscular scAAV9-SMN Injection Mediates Widespread Gene Delivery to the Spinal Cord and Decreases Disease Severity in SMA Mice.</a> Mol Ther. 21 (2), 282-290 (2013).</li><li>Azzouz, M., Hottinger, A., Paterna, J. C., Zurn, A. D., Aebischer, P., B&#252;eler, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increased+motoneuron+survival+and+improved+neuromuscular+function+in+transgenic+ALS+mice+after+intraspinal+injection+of+an+adeno-associated+virus+encoding+Bcl-2.">Increased motoneuron survival and improved neuromuscular function in transgenic ALS mice after intraspinal injection of an adeno-associated virus encoding Bcl-2.</a> Hum Mol Gen. 9 (5), (2000).</li><li>Lepore, A. C., Haenggeli, 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=Intraparenchymal+spinal+cord+delivery+of+adeno-associated+virus+IGF-1+is+protective+in+the+SOD1G93A+model+of+ALS.">Intraparenchymal spinal cord delivery of adeno-associated virus IGF-1 is protective in the SOD1G93A model of ALS.</a> Brain Res. 1185, 256-265 (2007).</li><li>Schnapp, B. J., Reese, T. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynein+is+the+motor+for+retrograde+axonal+transport+of+organelles.">Dynein is the motor for retrograde axonal transport of organelles.</a> Proc Natl Acad Sci USA. 86 (5), 1548-1552 (1989).</li><li>Vale, R. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Molecular+Motor+Toolbox+for+Intracellular+Transport.">The Molecular Motor Toolbox for Intracellular Transport.</a> Cell. 112 (4), 467-480 (2003).</li><li>Schiavo, G., Fainzilber, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+biology:+Alternative+energy+for+neuronal+motors.">Cell biology: Alternative energy for neuronal motors.</a> Nature. 495 (7440), 178-180 (2013).</li><li>Gracies, J. -. M., Lugassy, M., Weisz, D. J., Vecchio, M., Flanagan, S., Simpson, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Botulinum+Toxin+Dilution+and+Endplate+Targeting+in+Spasticity.+A+Double-Blind+Controlled+Study.">Botulinum Toxin Dilution and Endplate Targeting in Spasticity. A Double-Blind Controlled Study.</a> Arch Phys Med Rehabil. 90 (1), 9-16 (2009).</li><li>Van Campenhout, A., Molenaers, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Localization+of+the+motor+endplate+zone+in+human+skeletal+muscles+of+the+lower+limb:+anatomical+guidelines+for+injection+with+botulinum+toxin.">Localization of the motor endplate zone in human skeletal muscles of the lower limb: anatomical guidelines for injection with botulinum toxin.</a> Dev Med Child Neurol. 53 (2), 108-119 (2011).</li><li>Tosolini, A. P., Morris, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+characterization+of+the+motor+neuron+columns+supplying+the+rat+forelimb.">Spatial characterization of the motor neuron columns supplying the rat forelimb.</a> Neuroscience. 200, 19-30 (2012).</li><li>Tosolini, A. P., Mohan, R., Morris, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeting+the+full+length+of+the+motor+end+plate+regions+in+the+mouse+forelimb+increases+the+uptake+of+fluoro-gold+into+corresponding+spinal+cord+motor+neurons.">Targeting the full length of the motor end plate regions in the mouse forelimb increases the uptake of fluoro-gold into corresponding spinal cord motor neurons.</a> Front Neurol. 4, 58 (2013).</li><li>Mohan, R., Tosolini, A. P., Morris, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeting+the+motor+end+plates+in+the+mouse+hindlimb+gives+access+to+a+greater+number+of+spinal+cord+motor+neurons:+an+approach+to+maximize+retrograde+transport.">Targeting the motor end plates in the mouse hindlimb gives access to a greater number of spinal cord motor neurons: an approach to maximize retrograde transport.</a> Neuroscience. 274, 318-330 (2014).</li><li>Baumgartner, B. J., Shine, H. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuroprotection+of+spinal+motoneurons+following+targeted+transduction+with+an+adenoviral+vector+carrying+the+gene+for+glial+cell+line-derived+neurotrophic+factor.">Neuroprotection of spinal motoneurons following targeted transduction with an adenoviral vector carrying the gene for glial cell line-derived neurotrophic factor.</a> Exp Neurol. 153 (1), 102-112 (1998).</li><li>Gransee, H. M., Zhan, W. -. Z., Sieck, G. C., Mantilla, C. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+delivery+of+TrkB+receptor+to+phrenic+motoneurons+enhances+functional+recovery+of+rhythmic+phrenic+activity+after+cervical+spinal+hemisection.">Targeted delivery of TrkB receptor to phrenic motoneurons enhances functional recovery of rhythmic phrenic activity after cervical spinal hemisection.</a> PLoS ONE. 8 (5), e64755 (2013).</li></ol>

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 that the uptake of the transgenes by spinal cord motor neurons could be improved by the selective targeting of the motor end plates on the muscle fibres. It is important to note that, in the above-mentioned studies, the intramuscular injections of viral vectors were performed without reference to the location of the MEP region in the targeted muscles.
Throughout our analyses of the distribution of the motor end plates (MEPs) of the hindlimb and the forelimb, a few principles have emerged. First, the MEP region spans across the entire width of a muscle. Moreover, the MEPs are consistently aligned orthogonally with regards to the direction of the myofibres. Notably, the MEP region is not always located within the thickest portion of the muscle, an area often referred to as the &#8216;belly&#8217; and that is often the target of bolus intramuscular injections. It is worth noting that the current analyses were conducted on C57Bl6 mouse 22-23 and the Long Evans rat 21 and it was found that the distribution of the MEPs is conserved within each of these two strains of animals. However, whether or not the MEP distribution is conserved between different strains of animals within the same species remains to be documented.
Originally, we have documented the location of the MEP region for several muscles of the forelimb and hindlimb in order to guide small, multiple intramuscular injections of the retrograde neuronal tracer Fluoro-Gold. These MEP maps can be used to shuttle transgenes into motor neurons. In such instance, however, it is important to adjust the volumes of injections according to the type of virus used (e.g., adenovirus, lentivirus, etc.) its serotype as well as its titre. Keeping the fascia intact is a good way to keep the virus confined within the limits of the muscle(s) of interest. Another way to avoid spurious transfection of motor neurons innervating neighbour muscles is to wipe the surface of the muscle gently after the injections. These precautions might be irrelevant in some experimental designs where achieving high levels of transduction could be more relevant than containing it within the pool of motor neurons supplying the targeted muscle. Nevertheless, they are good practice for obvious safety reasons. If more than one isolated muscles need to be injected, it might be wise, if possible, to select a group of muscles that are close to each other to minimize the size of the skin incision. Rats and mice have very loose skin (i.e., not attached to the muscles), so it is relatively easy to target several muscles without having to perform huge skin incisions.
Effective gene therapy for ALS and other neuromuscular conditions using the retrograde machinery of neurons remains a challenge as, ideally, such therapy would transfer beneficial genes across the entire population of spinal cord motor neurons. However, it is not necessary to target all motor neurons to achieve sustainable levels of transduction if the therapeutic gene of interest is coding a secretory protein such as a neurotrophins. Indeed, non-transduced motor neurons in the vicinity of those that have been transduced have been shown to internalize exogenous molecules of neurotrophins through a paracrine mechanism 24. This &#8216;bystander effect&#8217; greatly enhances the efficacy of intramuscular targeting to shuttle secretory protein-encoding genes into corresponding motor neurons. On the other hand, given the fact that the loss of respiratory control is the ultimate cause of death in ALS, intramuscular injections of therapeutic viral constructs could be performed in intercostal muscles 1 or intrapleurally 25 to protect the motor neurons involved in breathing. By describing a method to easily highlight the location and organization of the MEPs on skeletal muscles, the present protocol will prove to be a valuable tool to explore novel gene therapy strategies for neuromuscular dysfunctions. This protocol also provides a minimally invasive way to preserve the integrity of the neuromuscular junction in these lower motor neuron diseases.

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 column of labelled motor neurons in the cervical spinal cord after an injection of retrograde neuronal tracer along the length of the motor end plate region of a mouse triceps brachii muscle. The retrogradely labelled motor neurons typically form a longitudinal column that extends across multiple segments of the spinal cord. Figure 3 illustrates the different numbers of labelled spinal cord motor neurons that was obtained through selective Fluoro-Gold targeting of the motor end plate region in triceps brachii. Maximal uptake was observed following delivery into the entire span of the MEP region, as opposed to targeting specific compartments of the MEP region.
<img alt="Figure 1" src="/files/ftp_upload/52846/52846fig1.jpg" /> 
Figure 1: View of the Lateral Forelimb of the Rat after Acetylcholinesterase Histochemical Staining for the Motor End Plates before (A) and after (B) Exposure to the Ammonium Sulphide Solution. In (A), the motor end plates are apparent as white speckled dots that form a continuous line that crosses the entire width of the muscle while the muscle fibres adopt a green-blue hue. In (B), the same forelimb is presented after immersion in the ammonium sulphide solution. After development in the ammonium sulphide solution, the motor end plates turn black over the brown muscle fibres. The arrows in both parts of the figures point to the location of the motor end plates for (1) Acromiotrapezius, (2) Spinodeltoideus and (3) Triceps Brachii.
<img alt="Figure 2" src="/files/ftp_upload/52846/52846fig2.jpg" /> 
Figure 2: Photomicrographs of 50 &#181;m Longitudinal Section through the Ventral Horn of the Spinal Cord at Cervical Levels Revealing Fluoro-Gold-labelled Motor Neurons. WM represents the white matter and GM represents the grey matter of the spinal cord. The dashed line indicates the boundary between the grey and white matter within the spinal cord section observed in the figure. The photomicrograph was taken with a 10X objective under the DAPI filter whereas the inset was taken with the 20X objective under the same filter. In both panels, Fluoro-Gold labelling can be observed in the motor neuron cytoplasm as well as proximal axon/dendritic processes. Figure as originally published in Tosolini et al. (2013) Targeting the Full Length of the Motor End Plate Regions in the Mouse Forelimb Increases the Uptake of Fluoro-Gold into Corresponding Spinal Cord Motor Neurons Front. Neurol. 4:58. doi: 10.3389/fneur.2013.00058
<img alt="Figure 3" src="/files/ftp_upload/52846/52846fig3.jpg" /> 
Figure 3: Selective Fluoro-Gold Targeting of the Motor End Plate Region in the Triceps Brachii and the Resulting Labelling in the Spinal Cord Motor Neurons. (A) is a schematic representation of the location of the motor end plates on triceps brachii fibres. The three different sections of the MEP region that were targeted individually are represented by the green, red and blue dots. The green and red dots represent the anterior and posterior halves of the entire MEP region respectively, whilst the blue dots represent the location of the bolus injection of FG into the belly of the muscle. The double-headed arrow orientates the muscle in an anterior-posterior axis. (B) is a composite diagram illustrating the labelling obtained following selective targeting of the different sections of the MEP region of the triceps brachii, as indicated in (A). Each dot in (B) represents a single labelled motor neuron. The black column is representative of typical labelling observed following injection of FG into the entire span of the MEP region of the muscle. The red motor neuron column is obtained following FG injections into the posterior half of the muscle, whilst the green motor neuron column is obtained following FG injections into the anterior half of the muscle. Injections restricted to the &#8220;belly&#8221; of the muscle yielded motor neurons in only a small number of spinal cord segments, as demonstrated by the blue column. Figure as originally published in Tosolini et al. (2013) Targeting the Full Length of the Motor End Plate Regions in the Mouse Forelimb Increases the Uptake of Fluoro-Gold into Corresponding Spinal Cord Motor Neurons Front. Neurol. 4:58. doi: 10.3389/fneur.2013.00058

Watch this Scientific Journal Video about Intramuscular Injections Along the Motor End Plates: A Minimally Invasive Approach to Shuttle Tracers Directly into Motor Neurons at JoVE.com

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 ...

intramuscular-injections-along-motor-end-plates-minimally-invasive

Universidad San Sebastian

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

Targeting of Deep Brain Structures with Microinjections for Delivery of Drugs, Viral Vectors, or Cell Transplants

Microinjections into the brain parenchyma are important procedures to deliver drugs, viral vectors or cell transplants.  The brain lesion that an injecting needle produces during its trajectory is a major concern especially in the mouse brain for not only the brain is small but also sometimes multiple injections are needed.  We show here a method to produce glass capillary needles with a 50-&mu;m lumen which significantly reduces the brain damage and allows a precise targeting into the rodent brain. This method allows a delivery of small volumes (from 20 to 100 nl), reduces bleeding risks, and minimizes passive diffusion of drugs into the brain parenchyma. By using different size of capillary glass tubes, or changing the needle lumen, several types of substances and cells can be injected. Microinjections with a glass capillary tube represent a significant improvement in injection techniques and deep brain targeting with minimal collateral damage in small rodents.

In this article, we show a method to make glass capillary needles with a 50-&mu;m lumen. This technique significantly reduces the brain damage, minimizes passive diffusion of drugs and allows a precise targeting into the rodent brain.

Microinjections into the brain parenchyma are important procedures to deliver drugs, viral vectors or cell transplants.  The brain lesion that an ...

Delivery of Therapeutic Agents Through Intracerebroventricular (ICV) and Intravenous (IV) Injection in Mice

Despite the protective role that blood brain barrier plays in shielding the brain, it limits the access to the central nervous system (CNS) which most often results in failure of potential therapeutics designed for neurodegenerative disorders 1,2. Neurodegenerative diseases such as Spinal Muscular Atrophy (SMA), in which the lower motor neurons are affected, can benefit greatly from introducing the therapeutic agents into the CNS. The purpose of this video is to demonstrate two different injection paradigms to deliver therapeutic materials into neonatal mice soon after birth. One of these methods is injecting directly into cerebral lateral ventricles (Intracerebroventricular) which results in delivery of materials into the CNS through the cerebrospinal fluid 3,4. The second method is a temporal vein injection (intravenous) that can introduce different therapeutics into the circulatory system, leading to systemic delivery including the CNS 5. Widespread transduction of the CNS is achievable if an appropriate viral vector and viral serotype is utilized. Visualization and utilization of the temporal vein for injection is feasible up to postnatal day 6. However, if the delivered material is intended to reach the CNS, these injections should take place while the blood brain barrier is more permeable due to its immature status, preferably prior to postnatal day 2. The fully developed blood brain barrier greatly limits the effectiveness of intravenous delivery. Both delivery systems are simple and effective once the surgical aptitude is achieved. They do not require any extensive surgical devices and can be performed by a single person. However, these techniques are not without challenges. The small size of postnatal day 2 pups and the subsequent small target areas can make the injections difficult to perform and initially challenging to replicate.

Delivery of Therapeutic Agents Through Intracerebroventricular ICV and Intravenous IV Injection in Mice

This article demonstrates two very different methods of injection: 1) into the brain (intracerebroventricular) and 2) systemic (intravenous) to introduce the therapeutic agents into the central nervous system of neonatal mice.

Despite the protective role that blood brain barrier plays in shielding the brain, it limits the access to the central nervous system (CNS) which most ...

Medicine

Intraspinal Cell Transplantation for Targeting Cervical Ventral Horn in Amyotrophic Lateral Sclerosis and Traumatic Spinal Cord Injury

Respiratory compromise due to phrenic motor neuron loss is a debilitating consequence of a large proportion of human traumatic spinal cord injury (SCI) cases 1 and is the ultimate cause of death in patients with the motor neuron disorder, amyotrophic laterals sclerosis (ALS) 2.
ALS is a devastating neurological disorder that is characterized by relatively rapid degeneration of upper and lower motor neurons. Patients ultimately succumb to the disease on average 2-5 years following diagnosis because of respiratory paralysis due to loss of phrenic motor neuron innnervation of the diaphragm 3. The vast majority of cases are sporadic, while 10% are of the familial form. Approximately twenty percent of familial cases are linked to various point mutations in the Cu/Zn superoxide dismutase 1 (SOD1) gene on chromosome 21 4. Transgenic mice 4,5 and rats 6 carrying mutant human SOD1 genes (G93A, G37R, G86R, G85R) have been generated, and, despite the existence of other animal models of motor neuron loss, are currently the most highly used models of the disease.
Spinal cord injury (SCI) is a heterogeneous set of conditions resulting from physical trauma to the spinal cord, with functional outcome varying according to the type, location and severity of the injury 7. Nevertheless, approximately half of human SCI cases affect cervical regions, resulting in debilitating respiratory dysfunction due to phrenic motor neuron loss and injury to descending bulbospinal respiratory axons 1. A number of animal models of SCI have been developed, with the most commonly used and clinically-relevant being the contusion 8.
Transplantation of various classes of neural precursor cells (NPCs) is a promising therapeutic strategy for treatment of traumatic CNS injuries and neurodegeneration, including ALS and SCI, because of the ability to replace lost or dysfunctional CNS cell types, provide neuroprotection, and deliver gene factors of interest 9.
Animal models of both ALS and SCI can model many clinically-relevant aspects of these diseases, including phrenic motor neuron loss and consequent respiratory compromise 10,11. In order to evaluate the efficacy of NPC-based strategies on respiratory function in these animal models of ALS and SCI, cellular interventions must be specifically directed to regions containing therapeutically relevant targets such as phrenic motor neurons. We provide a detailed protocol for multi-segmental, intraspinal transplantation of NPCs into the cervical spinal cord ventral gray matter of neurodegenerative models such as SOD1G93A mice and rats, as well as spinal cord injured rats and mice 11.

Neural precursor transplantation is a promising strategy for protecting and/or replacing lost/dysfunctional cervical phrenic motor neurons in spinal cord injury (SCI) and the motor neuron disorder, amyotrophic laterals sclerosis (ALS). We provide a protocol for cell delivery to cervical spinal cord ventral horn in rodent models of ALS and SCI.

Respiratory compromise due to phrenic motor neuron loss is a debilitating consequence of a large proportion of human traumatic spinal cord injury (SCI) ...

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 ...

Ultrasound-guided Transthoracic Intramyocardial Injection in Mice

Murine models of cardiovascular disease are important for investigating pathophysiological mechanisms and exploring potential regenerative therapies. Experiments involving myocardial injection are currently performed by direct surgical access through a thoracotomy. While convenient when performed at the time of another experimental manipulation such as coronary artery ligation, the need for an invasive procedure for intramyocardial delivery limits potential experimental designs. With ever improving ultrasound resolution and advanced noninvasive imaging modalities, it is now feasible to routinely perform ultrasound-guided, percutaneous intramyocardial injection. This modality efficiently and reliably delivers agents to a targeted region of myocardium. Advantages of this technique include the avoidance of surgical morbidity, the facility to target regions of myocardium selectively under ultrasound guidance, and the opportunity to deliver injectate to the myocardium at multiple, predetermined time intervals. With practiced technique, complications from intramyocardial injection are rare, and mice quickly return to normal activity on recovery from anesthetic. Following the steps outlined in this protocol, the operator with basic echocardiography experience can quickly become competent in this versatile, minimally invasive technique.

Echocardiography-guided percutaneous intramyocardial injection represents an efficient, reliable, and targetable modality for the delivery of gene transfer agents or cells into the murine heart. Following the steps outlined in this protocol, the operator can quickly become competent in this versatile, minimally invasive technique.

Murine models of cardiovascular disease are important for investigating pathophysiological mechanisms and exploring potential regenerative therapies. ...

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

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 ...

Nerve Stimulator-guided Injection of Autologous Stem Cells Near the Equine Left Recurrent Laryngeal Nerve

Recurrent laryngeal neuropathy (RLN) commonly affects horses and is characterized by abnormal respiratory sounds and exercise intolerance. The recurrent laryngeal nerve shows lesions of demyelination. The benefit of applying stem cells to demyelinated nerves has been demonstrated in various animal models. The aim of the study was to test the feasibility and safety of a peri-neuronal injection of autologous muscle-derived mesenchymal stem cells to the left recurrent laryngeal nerve in healthy horses by using an electrical nerve stimulator.
Muscle-derived stems cell are obtained from five healthy Standardbred horses by sampling 20 mg of muscle tissue with a semi-automatic 14 G biopsy needle from the triceps muscle. Movements of the larynx are monitored via upper-airway video endoscopy. The left recurrent laryngeal nerve is approached with an insulated nerve block needle. Nerve stimulation is applied, starting at 2 mA, and the successful abduction of the left arytenoid is monitored. The stimulation intensity is reduced progressively. When a loss of the motor response is observed at 0.5 mA, 107 autologous muscle-derived stem cells are injected. Two examiners, who are blinded to the time point, score the laryngeal function of the horses prior to the treatment and at day 1, day 7, and day 28 after the injection of the cells. In a sixth horse, 1 mL of 2% lidocaine is injected to further confirm the correct positioning of the needle. This leads to a temporary paralysis of the left arytenoid cartilage.
This study proves that the recurrent laryngeal nerve can be approached with the help of an electrical nerve stimulator and that the electrical stimulation of the nerve is well tolerated by the horses. No modification of the laryngeal function was observed in any of the horses after the injection of the stem cells. Further studies should be conducted to describe the effects of a peri-neuronal injection of autologous muscle-derived mesenchymal stem cells to horses suffering from RLN.

Here, we present a protocol to inject autologous muscle-derived stem cells in the proximity to the recurrent laryngeal nerve with the help of an electrical nerve stimulator. This novel technique may become useful for the treatment of equine recurrent laryngeal neuropathy.

Recurrent laryngeal neuropathy (RLN) commonly affects horses and is characterized by abnormal respiratory sounds and exercise intolerance. The recurrent ...

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

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

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

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

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

Biology

Electroporation and Nerve Injury in Mouse Levator Auris Longus Muscle

Combined In Vivo Electroporation and Short&#45;Term Reinnervation of the Cranial Levator Auris Longus Skeletal Muscle

The neuromuscular junction (NMJ) is the peripheral synapse controlling the contraction of skeletal muscle fibers to allow the coordinated movement of many organisms. At the NMJ, a presynaptic motor axon terminal contacts a muscle postsynaptic domain and is covered by terminal Schwann cells. The integrity and function of the NMJ is compromised under several conditions, including aging, neuromuscular diseases, and traumatic injuries. To analyze the potential contribution of muscle-derived proteins to NMJ maintenance and regeneration, an in vivo gene transfer strategy has been combined with the denervation of the cranial levator auris longus (LAL) muscle after mechanical nerve injury. Previous findings showed that the forced expression of control fluorescent proteins does not alter NMJ organization or neurotransmission. This procedure aims to describe a detailed method of in vivo electroporation of the LAL muscle followed by transection or crushing of the specific branch of the facial nerve innervating cranial muscles, leading to NMJ denervation and reinnervation, respectively. The combination of these experimental strategies in the convenient LAL muscle constitutes an efficient method to study the potential contribution of muscle protein overexpression or silencing in the context of short-term NMJ reinnervation.

Author Spotlight: Regenerative Roles of Muscle Proteins at Neuromuscular Junction Post-Nerve Injury

Here, we present a protocol that combines in vivo electroporation and denervation of the cranial levator auris longus (LAL) muscle. This procedure enables the study of the potential role of muscle-derived proteins in the regeneration of the neuromuscular synapse.

The neuromuscular junction (NMJ) is the peripheral synapse controlling the contraction of skeletal muscle fibers to allow the coordinated movement of many ...

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

Summary

Explore More Videos

Chapters in this video

Targeting of Deep Brain Structures with Microinjections for Delivery of Drugs, Viral Vectors, or Cell Transplants

Delivery of Therapeutic Agents Through Intracerebroventricular (ICV) and Intravenous (IV) Injection in Mice

Intraspinal Cell Transplantation for Targeting Cervical Ventral Horn in Amyotrophic Lateral Sclerosis and Traumatic Spinal Cord Injury

Revealing Neural Circuit Topography in Multi-Color

Ultrasound-guided Transthoracic Intramyocardial Injection in Mice

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

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

Nerve Stimulator-guided Injection of Autologous Stem Cells Near the Equine Left Recurrent Laryngeal Nerve

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

Combined In Vivo Electroporation and Short-Term Reinnervation of the Cranial Levator Auris Longus Skeletal Muscle