This study was supported by the Swiss National Foundation (n&#176; 3100AO-120327 and 31003A_140754/1)

Animal care was in accordance with guidelines approved by Swiss local authorities (Amt f&#252;r Landwirtschaft und Natur des Kantons Bern, Veterin&#228;rdienst, Sekretariat Tierversuche, approval Nrs. 52/11 and 35/14). These guidelines are in agreement with the European Community Directive 2010/63/EU.
Note: Work in a laminar flow hood with sterile instruments and solutions for all steps 1 - 5 including sub-steps.
1. Preparation of MEAs
Note: MEAs are composed of a glass substrate, micro-fabricated metal electrodes and a SU-8 polymer insulation layer (also see Tscherter et al. 3). For the purpose of this study, commercially available MEAs were ordered with a customized electrode array layout (Figure 1 A&#38;B). The 68 electrodes are arranged in a rectangular grid that is split into two zones by a 300 &#181;m wide groove free of electrodes, electrical leads and insulation. Each electrode is 40 x 40 &#181;m in size and they are spaced apart by 200 &#181;m from center to center. Four large ground electrodes are positioned around the recording site. They differ from other commercially available standard MEAs in their size (21 mm x 21 mm) and they have no fixed recording chamber.
<ol>
	<li>For disinfection rinse MEAs in 100% ethanol (2x), 70% ethanol (1x) and distilled water (2x) for at least 30 sec&#160;each. Let dry. Put 10 - 12 MEAs in a clean glass petri dish (150 mm x 25 mm) and close the lid.</li>
	<li>Autoclave MEAs for 20 min at 120 &#176;C. Store at RT.</li>
	<li>Prepare the coating solution (see materials list) for the MEAs and store aliquots at -20 &#176;C.</li>
	<li>Put each MEA in an individual sterile petri dish (35 mm x 10 mm).</li>
	<li>Use a cooled pipette to put 150 &#181;l chilled coating solution on top of the electrodes and close the lid of the petri dish. After about 10 min, check for air bubbles on top of the electrodes and gently remove them with a rubber-covered spatula if necessary. Let rest for 1 hr&#160;at RT.</li>
	<li>Aspirate coating solution residues, wash 1x with medium optimized for prenatal and embryonic neurons and 2x with distilled, sterile water. If MEAs are not used until the next day, keep them in distilled, sterile water at 37 &#176;C in the incubator O/N. Otherwise, let directly dry at RT.</li>
</ol>
2. Ingredients Required for the Preparation and Growth of Organotypic Cultures
<ol>
	<li>Prepare calcium-free wash solution (see materials list).</li>
	<li>Reconstitute chicken plasma in wash solution (1:1, shake gently, avoid formation of bubbles), centrifuge at 3,000 x g for about 20 min and decant the clear content into a sterile tube. Aliquot 200 &#181;l into cryotubes and store at -20 &#176;C.</li>
	<li>Reconstitute thrombin from bovine plasma accordingly (200 U/ml) and sterile-filter (0.2 &#181;m pore filter). Aliquot 200 &#181;l into crytotubes and store at -20 &#176;C.</li>
	<li>For 30 cultures prepare 100 ml of nutrient medium (see materials list).</li>
</ol>
3. Spinal Cord Tissue Dissection
Note: Procedure yields, depending on the number of embryos, spinal cord slices for about 25 - 35 co-cultures and is prepared inside a laminar flow hood under sterile conditions.
<ol>
	<li>Apply a lethal dose of pentobarbital (0.4 ml) to the mother animal by intramuscular injection. Confirm deep anesthesia by checking the pedal withdrawal reflex. Deliver E14 rat embryos by cesarean section and sacrifice by decapitation.</li>
	<li>Transfer the bodies of the embryos into a petri dish filled with sterile, chilled, wash solution.</li>
	<li>Perform one complete transversal cut with a scalpel above the hindlimbs and another one above the forelimbs and remove the limbs from the body. Next, make a cut in the frontal plane to separate viscera from the back piece containing the spinal cord.</li>
	<li>For slicing, transfer the back pieces containing the spinal cord one at a time on a mounting disk and cut at a thickness of 225 - 250 &#181;m with a tissue chopper. Put a drop of wash solution on the chopped tissue and transfer slices with a spatula in 35 mm x 10 mm petri dishes filled with sterile, chilled, wash solution.</li>
	<li>For each slice separately, dissect spinal cord away from remaining tissue. Leave dorsal root ganglia attached.</li>
	<li>Let the slices rest for 1 hr at 4 &#176;C.</li>
</ol>
4. Mounting Spinal Cord Tissue Slices on MEAs
<ol>
	<li>Warm up sterile nutrient medium in the incubator (37 &#176;C, lightly unscrewed lid for oxygenation).</li>
	<li>Position MEA with sterile tweezers with rubber-covered tips at RT in the petri dish under a stereo microscope with the electrode array in focus. Center a 6 &#181;l droplet of chicken plasma on the clean, dust-free, and sterile electrode array. Using a small spatula, carefully slide two spinal cord sections with ventral sides facing each other into the plasma droplet. Do not touch the electrodes with the spatula.</li>
	<li>Add 8 &#181;l of thrombin around the chicken plasma droplet. Using the thrombin pipette tip, carefully mix and spread the chicken plasma/thrombin mixture around the two slices. Again, do not touch the brittle electrode array directly. Just before coagulation, aspirate excess chicken plasma/thrombin.</li>
	<li>Cap the petri dish to retain high humidity while the MEA/culture assembly sits for about 1 hr in a humidified chamber inside the incubator at 37 &#176;C.</li>
	<li>Carefully add 10 &#181;l of nutrient medium to the culture chamber, cap the petri dish and put back into the incubator for about 30 min.</li>
	<li>Place each MEA/culture assembly with sterile, rubber-covered tips into a roller tube, add 3 ml of nutrient medium and tightly close the lid. Place the roller tube in the roller drum rotating at 1 - 2 rpm in the incubator in a 5% CO2-containing atmosphere at 37 &#176;C.</li>
	<li>Change half of the nutrient medium after 7 days in vitro (DIV) and afterwards 1 - 2 times&#160;per week.</li>
</ol>
5. Mechanical Lesions
<ol>
	<li>Take the MEA/culture assembly with sterile, rubber-covered tips out of the roller tube and place in a petri dish without nutrient medium under a stereo microscope with the tissue in focus. Visually verify that the two slices are fused.</li>
	<li>Hold the MEA/culture assembly steady by placing tweezers with rubber-covered tips on the MEA.</li>
	<li>Place a scalpel blade in the groove of the MEA close to the tissue slices. Hold the scalpel rather horizontally.</li>
	<li>Lift the scalpel handle up but let the scalpel blade stay in the groove of the MEA in such a way that the blade &#8220;rolls&#8221; from base to tip and thereby cuts through the tissue covering the groove.</li>
	<li>Severe any residual tissue connections with a 25 G needle tip if necessary. Work only in the area within the groove and do not touch the tender edges.</li>
	<li>Put the MEA/culture assembly back into the roller tube. Provide 3 ml of fresh nutrient medium to the cultures and place them back into the roller drum in the incubator.</li>
</ol>
6. Electrophysiological Recordings of Spontaneous Activity
<ol>
	<li>To investigate functional regeneration among the two spinal cord slices after the chosen number of DIV, mount the MEA/culture assembly in a recording chamber on a microscope and apply about 500 &#181;l extracellular solution (see materials list).</li>
	<li>Wait 10 min before the first recording to allow the system to stabilize.</li>
	<li>Record basic spontaneous activity 2x for about 10 min from each activity-detecting electrode of the MEA at RT.</li>
	<li>To ensure stable extracellular conditions, exchange extracellular solution after every recording session.</li>
	<li>To disinhibit the network, apply extracellular solution containing strychnine (1 &#956;M) and gabazine (10 &#956;M).&#160; Wait for at least 2 min before starting the recording.</li>
</ol>
7. Data Analysis
Note: For the detection of the extracellularly recorded action potentials use for each electrode a detector based on standard deviations and a subsequent discriminator. This procedure is described in detail in Tscherter et al. 3.
<ol>
	<li>Display the detected neuronal activity of each electrode in a raster plot according to standard procedures 3 (Figure 1F).</li>
	<li>Determine and display the total network activity for each slice by summing up the number of events in the according channels within a sliding window of 10 msec, shifted by 1 msec steps (Figure 1G and see Tscherter et al. 3).</li>
	<li>Detect in each slice individual bursts (clusters of activity that appear on several electrodes). Therefore, set a minimal peak activity threshold in the corresponding total network activity and define each burst start and burst end. For example, an appropriate threshold is 25% of the average burst amplitude and a burst start can be defined by being the first event in a time window of at least 5 msec, a burst end by being the last event in a time window of at least 25 msec (see Heidemann et al. 10).</li>
	<li>To quantify functional regeneration calculate the percentage of synchronized bursts between the two slices. To do this, calculate the latency between a burst in one slice and the following burst in the other slice. 
	Note: A burst pair is termed &#8220;synchronized&#8221; when the latency is smaller than the average burst length. Especially in co-cultures with a lot of activity, the possibility exists that both sides coincidentally initiate a burst in the chosen latency window. This fact has to be taken into account in the quantification of the percentage of synchronized bursts. For a detailed description of the calculations see Heidemann et al.10.</li>
</ol>
8. Fixation of the Cultures and Immunohistochemistry
<ol>
	<li>For all steps that include washing or incubation make sure to put the cultures on a tilting table to ensure proper diffusion of the solution through the tissue.</li>
	<li>Directly after the recording session is finished take the MEA/culture assembly out of the setup, put it in a petri dish (35 mm x 10 mm) and add about 2 ml of extracellular solution.</li>
	<li>To separate the slices from the surrounding cell layers that have formed during the time in vitro, take a 10 &#181;l pipette tip and circle the slices. Pay attention not to damage the cultures. It is possible to omit this step but embedding later on is easier when it is performed.</li>
	<li>Use a thin nonmetallic syringe needle (see materials list) in combination with a 1 ml syringe filled with extracellular solution to gently blow the culture off the MEA.</li>
	<li>Remove the tip of a Pasteur pipette and fit the rubber bulb on the slivered end. Use the back end to transfer the culture into fixative (4% paraformaldehyde with phosphate buffered saline (PBS, 0.1 M)). Do not use the standard tip of the Pasteur pipette for the transfer because it is too small and the cultures would be folded and probably damaged. Fix for 1 - 2 hr at 4 &#176;C. Rinse the MEA with distilled water and remove tissue residues with fingertips but do not scratch with the nail. Store MEAs in distilled water at 4&#176; C.</li>
	<li>Wash the cultures 3x in PBS for at least 10 min each. Store in PBS at 4 &#176;C or directly start with the staining procedure.</li>
	<li>Incubate the cultures for at least 90 min in blocking solution (5% goat serum, 1% bovine serum albumin, 0.3% detergent in PBS).</li>
	<li>Add the first antibody diluted in blocking solution and incubate for 3 nights at 4 &#176;C.</li>
	<li>Wash 3x in PBS for at least 30 min each.</li>
	<li>Add the second antibody diluted in PBS for 2 hr at RT.</li>
	<li>Wash 3x in PBS for at least 30 min each.</li>
	<li>Transfer the slices with a Pasteur pipette with removed tip on an object slide. Remove excess PBS and embed with mounting medium.</li>
</ol>

<ol>
	<li>Avossa, D., Rosato&#8211;Siri, M., Mazzarol, F., Ballerini, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spinal+circuits+formation:+a+study+of+developmentally+regulated+markers+in+organotypic+cultures+of+embryonic+mouse+spinal+cord.">Spinal circuits formation: a study of developmentally regulated markers in organotypic cultures of embryonic mouse spinal cord.</a> Neuroscience. 122 (2), 391-405 (2003).</li><li>Cifra, A., Mazzone, G. L., Nani, F., Nistri, A., Mladinic, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Postnatal+developmental+profile+of+neurons+and+glia+in+motor+nuclei+of+the+brainstem+and+spinal+cord,+and+its+comparison+with+organotypic+slice+cultures.">Postnatal developmental profile of neurons and glia in motor nuclei of the brainstem and spinal cord, and its comparison with organotypic slice cultures.</a> Dev Neurobiol. 72 (8), 1140-1160 (2012).</li><li>Tscherter, A., Heuschkel, M. O., Renaud, P., Streit, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatiotemporal+characterization+of+rhythmic+activity+in+rat+spinal+cord+slice+cultures.">Spatiotemporal characterization of rhythmic activity in rat spinal cord slice cultures.</a> Eur. J. Neurosci. 14 (2), 179-190 (2001).</li><li>Czarnecki, A., Magloire, V., Streit, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Local+oscillations+of+spiking+activity+in+organotypic+spinal+cord+slice+cultures.">Local oscillations of spiking activity in organotypic spinal cord slice cultures.</a> Eur. J. Neurosci. 27 (8), 2076-2088 (2008).</li><li>Kirkby, L. A., Sack, G. S., Firl, A., Feller, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+role+for+correlated+spontaneous+activity+in+the+assembly+of+neural+circuits.">A role for correlated spontaneous activity in the assembly of neural circuits.</a> Neuron. 80 (5), 1129-1144 (2013).</li><li>Ballerini, L., Galante, M., Grandolfo, M., Nistri, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+rhythmic+patterns+of+activity+by+ventral+interneurones+in+rat+organotypic+spinal+slice+culture.">Generation of rhythmic patterns of activity by ventral interneurones in rat organotypic spinal slice culture.</a> J. Physiol. (Lond.). 517 ((Pt 2)), 459-475 (1999).</li><li>Hanson, M. G., Landmesser, L. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Normal+patterns+of+spontaneous+activity+are+required+for+correct+motor+axon+guidance+and+the+expression+of+specific+guidance+molecules).">Normal patterns of spontaneous activity are required for correct motor axon guidance and the expression of specific guidance molecules).</a> Neuron. 43 (5), 687-701 (2004).</li><li>Courtine, 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=Recovery+of+supraspinal+control+of+stepping+via+indirect+propriospinal+relay+connections+after+spinal+cord+injury.">Recovery of supraspinal control of stepping via indirect propriospinal relay connections after spinal cord injury.</a> Nat. Med. 14 (1), 69-74 (2008).</li><li>Bonnici, B., Kapfhammer, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spontaneous+regeneration+of+intrinsic+spinal+cord+axons+in+a+novel+spinal+cord+slice+culture+model.">Spontaneous regeneration of intrinsic spinal cord axons in a novel spinal cord slice culture model.</a> Eur. J. Neurosci. 27 (10), 2483-2492 (2008).</li><li>Heidemann, M., Streit, J., Tscherter, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+regeneration+of+intraspinal+connections+in+a+new+in+vitro+model.">Functional regeneration of intraspinal connections in a new in vitro model.</a> Neuroscience. 262, 40-52 (2014).</li><li>Nishimaru, H., Restrepo, C. E., Ryge, J., Yanagawa, Y., Kiehn, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mammalian+motor+neurons+corelease+glutamate+and+acetylcholine+at+central+synapses.">Mammalian motor neurons corelease glutamate and acetylcholine at central synapses.</a> Proc. Natl. Acad. Sci. U.S.A. 102 (14), 5245-5249 (2005).</li><li>Tsuji, 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=Proteasome+inhibition+induces+selective+motor+neuron+death+in+organotypic+slice+cultures.">Proteasome inhibition induces selective motor neuron death in organotypic slice cultures.</a> J. Neurosci. Res. 82 (4), 443-451 (2005).</li><li>Magloire, V., Streit, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intrinsic+activity+and+positive+feedback+in+motor+circuits+in+organotypic+spinal+cord+slice+cultures.">Intrinsic activity and positive feedback in motor circuits in organotypic spinal cord slice cultures.</a> Eur. J. Neurosci. 30 (8), 1487-1497 (2009).</li><li>Bahrami, F., Janahmadi, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Antibiotic+supplements+affect+electrophysiological+properties+and+excitability+of+rat+hippocampal+pyramidal+neurons+in+primary+culture.">Antibiotic supplements affect electrophysiological properties and excitability of rat hippocampal pyramidal neurons in primary culture.</a> Iran. Biomed J. 17 (2), 101-106 (2013).</li><li>Mentis, 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=Noncholinergic+excitatory+actions+of+motoneurons+in+the+neonatal+mammalian+spinal+cord.">Noncholinergic excitatory actions of motoneurons in the neonatal mammalian spinal cord.</a> Proc. Natl. Acad. Sci U.S.A.. 20 (102), 7344-7349 (2005).</li><li>Tsang, Y. M., Chiong, F., Kuznetsov, D., Kasarskis, E., Geula, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Motor+neurons+are+rich+in+non&#8211;phosphorylated+neurofilaments:+Cross&#8211;+species+comparison+and+alterations+in+ALS.">Motor neurons are rich in non&#8211;phosphorylated neurofilaments: Cross&#8211; species comparison and alterations in ALS.</a> Brain Res. 861 (45&#8211;58), (2000).</li></ol>

The authors have nothing to disclose.

Several elements in the presented procedure are fundamental for the accomplishment of accurate and reproducible experiments. All steps during the preparation of the MEAs, solutions and the production of the slice cultures are performed under sterile conditions in a laminar flow hood. Antibiotics are not applied to the nutrient medium because they can affect the spontaneous activity 14. During the MEA coating, the most important part is to make sure that the extracellular matrix gel solution stays cold at all times. Thaw it in the fridge and keep it on ice for the whole procedure to ensure a maximal temperature close to 0 &#176;C. During the preparation of the slices quick isolation and sectioning, as well as paying attention to a low temperature by using ice cold dissection buffers increases the percentage of healthy cultures. Moreover, the plasma/thrombin coagulation is one of the most critical steps. First, make sure that the plasma is properly mixed with the thrombin. Second, pay special attention to removing as much residue as possible to ensure proper attachment of the slices to the MEAs. This step is extremely time-sensitive; if the time frame is too short, too much of the plasma/thrombin mixture is removed. If the time frame is too long, coagulation already happened, increasing the risk of removing the slices along with the residual plasma/thrombin.
If mechanical lesions are planned, it is important to use MEAs with an accurate design. The area where the lesion is intended needs to be free of electrodes, wires and insulation. Otherwise, the electrics of the MEA are going to be damaged and therefore, no recordings can be performed anymore.
Another crucial step concerns the size of the lesion. It has been shown previously that reconnecting two young slices after lesion takes about two days 10. For all experiments, the rule of thumb was used that the space between the slices after lesion should not be bigger than the width of the MEA groove (300 &#181;m). A bigger lesion size might result in a longer time frame for reconnection or, if the lesion is too big, no reconnection at all. To annotate, former experiments revealed that an initial placing of the two slices further away than 1 mm results in a very low connection rate.
Before the start of the recordings, an adjustment time of at least 10 min to RT, instead of 37 &#176;C of the incubator, and to the extracellular solution, instead of the nutrition medium is recommended. Close observation of the activity pattern during these initial minutes ensures that the measured data represent the bursting pattern of the culture appropriately after the recording was started.
For the detection of the activity, data acquisition and further processing, homemade hardware (recording chamber, connections to the amplifiers and amplifiers), programs in LabVIEW, C++ and IgorPro are used. Commercial MEA setups and other software packages are suited as well for these operations. Here, the signals seen by the electrodes are amplified 1001 times, and then digitized at a rate of 6 kHz.
The presented model can be a helpful tool to examine propriospinal connections because in vivo, they are difficult to study without the interference of ascending and descending fibre tracts. The co-culturing of two spinal cord slices can elegantly circumvent this issue. The cultures provide a microenvironment that can be tightly controlled and easily manipulated. On the other hand, supraspinal and sensory input is of course missing. Additionally, the process of the culture preparation represents a situation of CNS damage. Glial cells like astrocytes and microglia are known to respond to lesions with increased proliferation and therefore, the tissue is to a certain extent reorganized. Relating to the presented model the formation of a glial scar after performing mechanical lesions was not observed. One explanation could be that adaptive mechanisms of astrocytes were already triggered during the preparation process and that lesions of the cultures did not result in a further response. Also, there is no evidence for the involvement of myelin associated inhibitors, e.g., Nogo-A, in the low regeneration potential of cultures lesioned at an old age. However, it was shown that treatment with the phosphodiesterase 4 inhibitor Rolipram, starting directly after performing lesions at 23 DIV increases functional regeneration between the slices 10. These results demonstrate that functional recovery can be increased in old cultures. Noticeably, when counting the number of axons crossing the lesion site, no difference between cultures lesioned at a young age and cultures lesioned at an old age was discovered. These findings suggest that lack of axonal regeneration is not the major reason for the loss of recovery after late lesions in culture and therefore emphasize the need for a model that allows functional analysis of regeneration. Synaptic formation and synaptic plasticity could play a major role in the recovery process 10. These mechanisms gained a lot of attention during recent years and it is nowadays widely accepted that they have a major impact on the outcome after spinal cord injury. Therefore, a model that provides stable conditions to investigate these processes in isolation and in great detail can certainly help to obtain insight about functional regeneration in the spinal cord.

Organotypic slice cultures of the central nervous system (CNS) have been used extensively to study various physiological and pathological phenomena reaching from neuronal development to neurodegeneration. Compared to dissociated cell cultures, organotypic slices offer the advantage of more complexity with an intact cytoarchitecture and local synaptic circuitry. At the same time, the tissue can be analyzed in an isolated way without the need to implicate the intricacy of the whole in vivo context. Moreover, easy and repeated access to the cells of choice is warranted, the extracellular environment can be precisely controlled and usually, in vitro models are less costly than their in vivo counterparts. In recent years, various studies presented striking similarities between the developmental profiles of long term cultures versus the corresponding living animals 1,2. It has been shown that neuronal circuits of various CNS regions express spontaneous network activity during development and that organotypic slices partially maintain this phenomenon 3&#8211;7. Isolated spinal cord preparations have been particularly used to investigate rhythm generation 6 and the formation of neuronal circuits 1.
A way to record the spontaneous activity of organotypic slices is to culture them on top of multi-electrode arrays (MEAs). These devices contain multiple electrodes that can monitor the extracellular potentials generated by action potentials from numerous cells simultaneously (multi-unit recording). The high temporal resolution of the MEA recordings can be used to reconstruct the precise activity dynamics within a given neuronal circuit. Additionally, in contrast to patch-clamping the technique is non-invasive, which permits long-term measurements and results in the fact that neurons are not affected in their behavior.
For the purpose of this study, the combination of organotypic spinal cord co-cultures and multisite recordings was used to investigate functional regeneration within the spinal cord. Since adult higher vertebrates have limited potential to recover from damage of the spinal cord, different strategies have been studied to promote regeneration after spinal cord injury. So far, the corticospinal tract has usually been the model system of choice for such investigations. These experiments are typically time-consuming, costly and require large animal cohorts due to high variability within single groups. Moreover, evidence suggests that propriospinal connections (neurons that are entirely confined within the spinal cord) can play a pivotal role in the recovery process following spinal cord lesions 8. These fibers are difficult to study in vivo without the interference of ascending and descending fiber tracts. Bonnici and Kapfhammer 9 used longitudinal spinal cord slice cultures of postnatal mice as an alternative approach. After performing mechanical lesions they analyzed semi-quantitatively the morphological regeneration of axons between spinal cord segments. They observed less but not none axons crossing the lesion site in cultures cut at a mature age. In contrast, cultures that were lesioned at a younger stage displayed a high amount of axonal regeneration 5 - 7 days after the damage. However, proof for functional connections was not presented.
Therefore, the development of a representative in vitro model of propriospinal fibers that allows the investigation of functional regeneration can be a valuable tool to extend our knowledge about regenerative processes in the spinal cord.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Planar multielectrode array</td>
			<td>Qwane Biosciences</td>
			<td></td>
			<td>custom-made according to the design of our lab</td>
		</tr>
		<tr>
			<td>Nutrient medium</td>
			<td></td>
			<td></td>
			<td>For 100 ml</td>
		</tr>
		<tr>
			<td>Dulbeccos modified Eagle&#39;s medium</td>
			<td>Gibco</td>
			<td>31966-021</td>
			<td>80 ml</td>
		</tr>
		<tr>
			<td>Horse serum</td>
			<td>Gibco</td>
			<td>26050-070</td>
			<td>10 ml</td>
		</tr>
		<tr>
			<td>distilled, sterile water</td>
			<td></td>
			<td></td>
			<td>10 ml&#160;</td>
		</tr>
		<tr>
			<td>Nerve growth factor-7S [5ng/mL]</td>
			<td>Sigma-Aldrich</td>
			<td>N0513</td>
			<td>200 &#181;l</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>reconstituted in wash solution with 1% BSA</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Coating solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Extracellular matrix gel</td>
			<td>BD Biosciences</td>
			<td>356230</td>
			<td>Must stay cold at all times; dilute 1:50 with medium optimized for prenatal and embryonic neurons</td>
		</tr>
		<tr>
			<td>medium optimized for prenatal and embryonic neurons</td>
			<td>Gibco</td>
			<td>21103-049</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Wash solution</td>
			<td></td>
			<td></td>
			<td>[mg/L]</td>
		</tr>
		<tr>
			<td>MgCl2-6H2O</td>
			<td></td>
			<td></td>
			<td align="right">100</td>
		</tr>
		<tr>
			<td>MgSO4-7H2O</td>
			<td></td>
			<td></td>
			<td align="right">100</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td></td>
			<td></td>
			<td align="right">400</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td></td>
			<td></td>
			<td align="right">60</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td></td>
			<td></td>
			<td align="right">8000</td>
		</tr>
		<tr>
			<td>Na2HPO4 anhydrous</td>
			<td></td>
			<td></td>
			<td align="right">48</td>
		</tr>
		<tr>
			<td>D-Glucose (Dextrose)</td>
			<td></td>
			<td></td>
			<td align="right">1000</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>According to the protocol on http://www.lifetechnologies.com/ch/en/home/technical-resources/media-formulation.152.html</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Extracellular solution (pH 7.4)</td>
			<td></td>
			<td></td>
			<td>[mM]</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td></td>
			<td></td>
			<td>145</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td></td>
			<td></td>
			<td>4</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td></td>
			<td></td>
			<td>1</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td></td>
			<td></td>
			<td>2</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td></td>
			<td></td>
			<td>5</td>
		</tr>
		<tr>
			<td>Na-pyruvate</td>
			<td></td>
			<td></td>
			<td>2</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td></td>
			<td></td>
			<td>5</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Blocking Solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Detergent: Triton X-100</td>
			<td></td>
			<td></td>
			<td>0.3%, harmful if swallowed</td>
		</tr>
		<tr>
			<td>Bovine serum albumin</td>
			<td>Sigma-Aldrich</td>
			<td>A9418</td>
			<td>1%</td>
		</tr>
		<tr>
			<td>Goat serum</td>
			<td>Gibco</td>
			<td>16210-064</td>
			<td align="right">5%</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Chicken Plasma</td>
			<td>Sigma-Aldrich</td>
			<td>P2366</td>
			<td>Lyophilized, reconstitute with wash solution</td>
		</tr>
		<tr>
			<td>Gabazine</td>
			<td>Sigma-Aldrich</td>
			<td>SR-95531</td>
			<td></td>
		</tr>
		<tr>
			<td>Mecamylamine hydrochloride</td>
			<td>Tocris</td>
			<td>2843</td>
			<td>toxic if swallowed</td>
		</tr>
		<tr>
			<td>Micropipette</td>
			<td>World Precision Instruments, Inc.</td>
			<td>MF28G-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td></td>
			<td></td>
			<td>harmful, 4%</td>
		</tr>
		<tr>
			<td>SMI-31</td>
			<td>Covance</td>
			<td>SMI-31P</td>
			<td></td>
		</tr>
		<tr>
			<td>SMI-32</td>
			<td>Covance</td>
			<td>SMI-32R-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Strychnine</td>
			<td>Sigma-Aldrich</td>
			<td>S0532</td>
			<td>toxic</td>
		</tr>
		<tr>
			<td>Thrombin</td>
			<td>Merck Millipore</td>
			<td>1123740001</td>
			<td>irritant, sensitizing, reconstitute with wash solution</td>
		</tr>
		<tr>
			<td>Tissue culture flat tube 10 (= roller tube)</td>
			<td>Techno Plastic Products AG</td>
			<td>91243</td>
			<td></td>
		</tr>
	</tbody>
</table>

investigating functional regeneration in organotypic spinal cord co-cultures grown on multi-electrode arrays

Adult higher vertebrates have a limited potential to recover from spinal cord injury. Recently, evidence emerged that propriospinal connections are a promising target for intervention to improve functional regeneration. So far, no in vitro model exists that grants the possibility to examine functional recovery of propriospinal fibers. Therefore, a representative model that is based on two organotypic spinal cord sections of embryonic rat, cultured next to each other on multi-electrode arrays (MEAs) was developed. These slices grow and, within a few days in vitro, fuse along the sides facing each other. The design of the used MEAs permits the performance of lesions with a scalpel blade through this fusion site without inflicting damage on the MEAs. The slices show spontaneous activity, usually organized in network activity bursts, and spatial and temporal activity parameters such as the location of burst origins, speed and direction of their propagation and latencies between bursts can be characterized. Using these features, it is also possible to assess functional connection of the slices by calculating the amount of synchronized bursts between the two sides. Furthermore, the slices can be morphologically analyzed by performing immunohistochemical stainings after the recordings. 
Several advantages of the used techniques are combined in this model: the slices largely preserve the original tissue architecture with intact local synaptic circuitry, the tissue is easily and repeatedly accessible and neuronal activity can be detected simultaneously and non-invasively in a large number of spots at high temporal resolution. These features allow the investigation of functional regeneration of intraspinal connections in isolation in vitro in a sophisticated and efficient way.

To investigate the potential for functional recovery of the co-cultures derived from the spinal cord of E14 rat embryos, lesions were performed in a time window of 8 to 28 DIV. 2 to 3 weeks later, the spontaneous neuronal activity was recorded with the MEAs (Figure 1 C&#38;D). The extracellularly recorded action potentials were identified offline for each individual electrode (Figure 1E). From these data raster plots were generated (Figure 1F), followed by network activity plots to visualize the total activity separately per side (Figure 1G). In each slice the spontaneous activity is usually organized in bursts. If the slices are functionally connected, these bursts can propagate from one slice to the other. Therefore, the amount of synchronized bursts between the two slices was calculated to quantitatively measure functional regeneration. For a detailed description of how the transformation into the different plots is performed and how the formula for calculating the percentage of synchronized activity was derived, please see Heidemann et al. 10.
In all experiments, the activity was first recorded under standard conditions. In a second step, synaptic inhibition was blocked by adding strychnine (1 &#181;M) and gabazine (10 &#181;M) to the extracellular bath solution. This disinhibition leads usually to a more regular activity pattern with prolonged bursts and burst intervals, which facilitates the definition of bursts and thus the determination of burst synchronization between the slices.
Cultures lesioned at a young age (7 - 9 DIV) displayed a high amount of synchronized activity 2-3 weeks after lesion whereas cultures lesioned at later stages (&#62;19 DIV) showed a distinct reduction in the ability to regenerate (Figure 2 A - F). These findings indicate that the regeneration ability of functional connections in spinal cord slice cultures decreases from a high level, representing the embryonic state in vivo, to a low level, representing the further developed state in vivo, within three weeks in culture.
What kinds of connections are actually involved in burst synchronization between the two slices? Besides propriospinal connections, the fibers could also arise from motoneurons or dorsal root ganglion neurons. It has been shown previously that dorsal root ganglion neurons are not relevant for the functional connectivity between the spinal cord slices 10. However, the involvement of motoneurons remained a possibility. Since motoneurons are known to form cholinergic connections to spinal cord neurons through nicotinic receptors 13, the activity of spinal cord co-cultures was recorded at 8 DIV, a time when the cultures show highly synchronized activity 10, and the nicotinic antagonist Mecamylamine (MEC, 100 &#181;M) was added to the extracellular bath solution. A participation of motoneurons in the connection between the two slices would result in a decreased percentage of synchronized activity. However, this was not the case (MEC: 91.0 &#177; 4.5%, n = 7; control: 93.6 &#177; 4.6%, n = 7, p &#62;0.05, Wilcoxon matched pairs test). These results suggest that cholinergic synapses do not contribute to the functional connection between slices. However, since motoneurons have also been shown to release glutamate at synapses within the CNS 11, 15, these experiments do not entirely exclude their involvement.
To identify motoneurons immunohistochemical stainings against the non-phosphorylated neurofilament H with SMI-32 were performed. They revealed several labelled large cell bodies typical for motoneurons, distributed over the slices (Figure 3A). The SMI-32 antibody has been reported to label the cell bodies of motoneurons 12 and this finding holds also true for the used cultures 13. Also, stainings of neuronal cell bodies in general, e.g., against b-III-tubulin or NeuN as well as glial cell bodies, e.g., astrocytes with GFAP antibody are in line with other reports and match the morphological descriptions of these cell types (Figure 3 B - D). Nevertheless, the labelling of axons with the SMI-32 antibody is concerning (Figure 3E). Against expectations, a huge network of fibers appears when using this antibody and it looks similar to the SMI-31 staining that labels phosphorylated neurofilament H (Figure 3F). One interpretation of this result could be that the developmental regulation of neurofilament phosphorylation/dephosphorylation is not as tight in the used cultures compared to the in vivo situation. A mixed occurrence of phosphorylated and non-phosphorylated neurofilaments in the same axon could explain this finding. Another possibility is that the SMI-32 labelled axons arise from projection neurons. Tsang and colleagues 16 have shown that e.g., Clark&#8217;s column and intermediolateral cell column neurons are stained by SMI-32 besides motoneurons. Pronounced neurite proliferation of such neurons could also explain the abundance of SMI-32 positive fibers.
<img alt="Figure 1" src="/files/ftp_upload/53121/53121fig1.jpg" /> 
Figure 1.&#160;Display and Analysis of Spontaneous Activity.&#160;(A) Diagram of a MEA. The platinum covered electrodes are depicted in black, the transparent wires in red and the groove in the middle of the MEA in yellow. (B) Close-up of the electrode array located in the center of the MEA. The diagrams are kindly provided by Dr. M. Heuschkel. (C) Bright-field image of an 8 DIV old culture. The slices have grown and fused along the sides facing each other. The yellow bar represents the electrode- and insulation-free groove of the MEA. Scale bar = 400 &#181;m (D) Timeline of experiments. Two spinal cord slices of E14 rat embryos are placed next to each other on MEAs. Within a few days, the slices grow and fuse along the sides facing each other. In a time frame of 8 - 28 DIV, complete lesions are performed through the fusion site. Two to three weeks later the spontaneous activity is recorded and the cultures are fixed for immunohistochemical stainings. (E) Spontaneous activity traces of each individual electrode of a 23 DIV old culture. For clearer visualization, only every second trace is illustrated. Orange traces depict activity that has been recorded from the slice on the right side, blue traces from the left side. Most of the bursts are synchronized between the two. The arrow points to a burst occurring in the left slice that only partially propagated to the right slice. The activity in the right slice however did not reach the chosen threshold of at least 25% of the averaged maximal peak activity of the according side and therefore, is not detected as a burst. Magnifications on the right depict the last synchronized burst pair. (F) Raster plot of the activity shown in (E). (G) Network activity plot with defined bursts (bars below baseline) of the activity shown in (E). Adapted from Heidemann et al. 10. <a href="https://www.jove.com/files/ftp_upload/53121/53121fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53121/53121fig2.jpg" /> 
Figure 2.&#160;Synchronized versus Not Synchronized Activity.&#160;(A, B) Raster plots of cultures lesioned at a young age (at 7 - 9 DIV), showing synchronized activity under standard conditions (A) and under disinhibition (B). (C, D) Raster plots of cultures lesioned at an old age (at&#160;&#62;19 DIV), showing not synchronized activity under standard conditions (C) and under disinhibition (D). (E, F) Average percentage of synchronized activity plotted for each individual lesion day recorded under standard conditions (E) and under disinhibition (F). Recordings were taken 2 - 3 weeks after the day of lesion. Adapted from Heidemann et al. 10. <a href="https://www.jove.com/files/ftp_upload/53121/53121fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53121/53121fig3.jpg" /> 
Figure 3.&#160;Immunohistochemical Characterization.&#160;(A) Staining of motoneuronal cell bodies with SMI-32. (B) &#946;-III tubulin labeling of mature neuronal cell bodies and their processes. (C) Identification of astrocytes with GFAP. (D) Labeling of mature neuronal cell body with NeuN. (E) SMI-32 staining of non-phosphorylated neurofilament H. (F) Phosphorylated neurofilament H identified by SMI-31. Adapted from Heidemann et al. 10. Note that with both, SMI-31 and SMI-32, a large network of fibers is visible in the cultures. Bars A-D = 20 &#181;m, H&#38;F = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/53121/53121fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Investigating Functional Regeneration in Organotypic Spinal Cord Co-cultures Grown on Multi-electrode Arrays at JoVE.com

Investigating Functional Regeneration in Organotypic Spinal Cord Co-cultures Grown on Multi-electrode Arrays

Here we present a protocol that is based on spinal cord slices cultured on multi-electrode arrays to study functional regeneration of propriospinal connections in vitro.

Adult higher vertebrates have a limited potential to recover from spinal cord injury. Recently, evidence emerged that propriospinal connections are a ...

investigating-functional-regeneration-organotypic-spinal-cord-co

Research

JoVE Journal

Neuroscience

9.0K Views.  University of Bern. Here we present a protocol that is based on spinal cord slices cultured on multi-electrode arrays to study functional regeneration of propriospinal connections in vitro.

Video: Investigating Functional Regeneration in Organotypic Spinal Cord Co-cultures Grown on Multi-electrode Arrays

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Multi-electrode Array Recordings of Neuronal Avalanches in Organotypic Cultures

The cortex is spontaneously active, even in the absence of any particular input or motor output. During development, this activity is important for the migration and differentiation of cortex cell types and the formation of neuronal connections1. In the mature animal, ongoing activity reflects the past and the present state of an animal into which sensory stimuli are seamlessly integrated to compute future actions. Thus, a clear understanding of the organization of ongoing i.e. spontaneous activity is a prerequisite to understand cortex function. 
Numerous recording techniques revealed that ongoing activity in cortex is comprised of many neurons whose individual activities transiently sum to larger events that can be detected in the local field potential (LFP) with extracellular microelectrodes, or in the electroencephalogram (EEG), the magnetoencephalogram (MEG), and the BOLD signal from functional magnetic resonance imaging (fMRI). The LFP is currently the method of choice when studying neuronal population activity with high temporal and spatial resolution at the mesoscopic scale (several thousands of neurons). At the extracellular microelectrode, locally synchronized activities of spatially neighbored neurons result in rapid deflections in the LFP up to several hundreds of microvolts. When using an array of microelectrodes, the organizations of such deflections can be conveniently monitored in space and time. 
Neuronal avalanches describe the scale-invariant spatiotemporal organization of ongoing neuronal activity in the brain2,3. They are specific to the superficial layers of cortex as established in vitro4,5, in vivo in the anesthetized rat 6, and in the awake monkey7. Importantly, both theoretical and empirical studies2,8-10 suggest that neuronal avalanches indicate an exquisitely balanced critical state dynamics of cortex that optimizes information transfer and information processing.
In order to study the mechanisms of neuronal avalanche development, maintenance, and regulation, in vitro preparations are highly beneficial, as they allow for stable recordings of avalanche activity under precisely controlled conditions. The current protocol describes how to study neuronal avalanches in vitro by taking advantage of superficial layer development in organotypic cortex cultures, i.e. slice cultures, grown on planar, integrated microelectrode arrays (MEA; see also 11-14).

A robust way to study neuronal avalanches, i.e. scale-invariant spatio-temporal activity bursts, indicative of critical state dynamics in cortex. Avalanches emerge spontaneously in developing superficial layers of cultured cortex which allows for long-term measurements of the activity with planar integrated multi-electrode arrays (MEA) under precisely controlled conditions.

The cortex is spontaneously active, even in the absence of any particular input or motor output.  During development, this activity is important for the ...

Spinal Cord Transection in the Larval Zebrafish

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

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

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

Spiral Ganglion Neuron Explant Culture and Electrophysiology on Multi Electrode Arrays

Spiral ganglion neurons (SGNs) participate in the physiological process of hearing by relaying signals from sensory hair cells to the cochlear nucleus in the brain stem. Loss of hair cells is a major cause of sensory hearing loss. Prosthetic devices such as cochlear implants function by bypassing lost hair cells and directly stimulating SGNs electrically, allowing for restoration of hearing in deaf patients. The performance of these devices depends on the functionality of SGNs, the implantation procedure and on the distance between the electrodes and the auditory neurons. 
We hypothesized, that reducing the distance between the SGNs and the electrode array of the implant would allow for improved stimulation and frequency resolution, with the best results in a gapless position. Currently we lack in vitro culture systems to study, modify and optimize the interaction between auditory neurons and electrode arrays and characterize their electrophysiological response. To address these issues, we developed an in vitro bioassay using SGN cultures on a planar multi electrode array (MEA). With this method we were able to perform extracellular recording of the basal and electrically induced activity of a population of spiral ganglion neurons. We were also able to optimize stimulation protocols and analyze the response to electrical stimuli as a function of the electrode distance. This platform could also be used to optimize electrode features such as surface coatings.

We present a protocol to culture primary murine spiral ganglion neuron explants on multi electrode arrays to study neuronal response profiles and optimize stimulation parameters. Such studies aim to improve the neuron-electrode interface of cochlear implants to benefit hearing in patients as well as the energy consumption of the device.

Spiral ganglion neurons (SGNs) participate in the physiological process of hearing by relaying signals from sensory hair cells to the cochlear nucleus in ...

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

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

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

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

Automated Gait Analysis to Assess Functional Recovery in Rodents with Peripheral Nerve or Spinal Cord Contusion Injury

Peripheral and central nerve injuries are mostly studied in rodents, especially rats, given the fact that these animal models are both cost-effective and a lot of comparative data has been published in the literature. This includes a multitude of assessment methods to study functional recovery following nerve injury and repair. Besides evaluation of nerve regeneration by means of histology, electrophysiology, and other in vivo and in vitro assessment techniques, functional recovery is the most important criterion to determine the degree of neural regeneration. Automated gait analysis allows recording of a vast quantity of gait-related parameters such as Paw Print Area and Paw Swing Speed as well as measures of inter-limb coordination. Additionally, the method provides digital data of the rats&#8217; paws after neuronal damage and during nerve regeneration, adding to our understanding of how peripheral and central nervous injuries affect their locomotor behavior. Besides the predominantly used sciatic nerve injury model, other models of peripheral nerve injury such as the femoral nerve can be studied by means of this method. In addition to injuries of the peripheral nervous systems, lesions of the central nervous system, e.g., spinal cord contusion can be evaluated. Valid and reproducible data assessment is strongly dependent on meticulous adjustment of the hard- and software settings prior to data acquisition. Additionally, proper training of the experimental animals is of crucial importance. This work aims to illustrate the use of computerized automated gait analysis to assess functional recovery in different animal models of peripheral nerve injury as well as spinal cord contusion injury. It also emphasizes the method&#8217;s limitations, e.g., evaluation of nerve regeneration in rats with sciatic nerve neurotmesis due to limited functional recovery. Therefore, this protocol is thought to help researchers interested in peripheral and central nervous injuries to assess functional recovery in rodent models.

Automated gait analysis is a feasible tool to evaluate functional recovery in rodent models of peripheral nerve injury and spinal cord contusion injury. While it requires only one setup to assess locomotor function in various experimental models, meticulous hard- and soft-ware adjustment and training of the animals is highly important.

Peripheral and central nerve injuries are mostly studied in rodents, especially rats, given the fact that these animal models are both cost-effective and ...

Electromagnetic Controlled Closed-Head Model of Mild Traumatic Brain Injury in Mice

Highly reproducible animal models of traumatic brain injury (TBI), with well-defined pathologies, are needed for testing therapeutic interventions and understanding the mechanisms of how a TBI alters brain function. The availability of multiple animal models of TBI is necessary to model the different aspects and severities of TBI seen in people. This manuscript describes the use of a midline closed head injury (CHI) to develop a mouse model of mild TBI. The model is considered mild because it does not produce structural brain lesions based on neuroimaging or gross neuronal loss. However, a single impact creates enough pathology that cognitive impairment is measurable at least 1 month after injury. A step-by-step protocol to induce a CHI in mice using a stereotaxically guided electromagnetic impactor is defined in the paper. The benefits of the mild midline CHI model include the reproducibility of the injury-induced changes with low mortality. The model has been temporally characterized up to 1 year after the injury for neuroimaging, neurochemical, neuropathological, and behavioral changes. The model is complementary to open skull models of controlled cortical impact using the same impactor device. Thus, labs can model both mild diffuse TBI and focal moderate-to-severe TBI with the same impactor.

The protocol describes mild traumatic brain injury in a mouse model. In particular, a step-by-step protocol to induce a mild midline closed head injury and the characterization of the animal model is fully explained.

Highly reproducible animal models of traumatic brain injury (TBI), with well-defined pathologies, are needed for testing therapeutic interventions and ...

Long-Term Engraftment of h-SC-NES Cells in Spinal Cord Organotypic Cultures

Long-Term Mouse Spinal Cord Organotypic Slice Culture as a Platform for Validating Cell Transplantation in Spinal Cord Injury

Resolutive cures for spinal cord injuries (SCIs) are still lacking, due to the complex pathophysiology. One of the most promising regenerative approaches is based on stem cell transplantation to replace lost tissue and promote functional recovery. This approach should be further explored better in vitro and ex vivo for safety and efficacy before proceeding with more expensive and time-consuming animal testing. In this work, we show the establishment of a long-term platform based on mouse spinal cord (SC) organotypic slices transplanted with human neural stem cells to test cellular replacement therapies for SCIs.
Standard SC organotypic cultures are maintained for around 2 or 3 weeks in vitro. Here, we describe an optimized protocol for long-term maintenance (&#8805;30 days) for up to 90 days. The medium used for long-term culturing of SC slices was also optimized for transplanting neural stem cells into the organotypic model. Human SC-derived neuroepithelial stem (h-SC-NES) cells carrying a green fluorescent protein (GFP) reporter were transplanted into mouse SC slices. Thirty days after the transplant, cells still show GFP expression and a low apoptotic rate, suggesting that the optimized environment sustained their survival and integration inside the tissue. This protocol represents a robust reference for efficiently testing cell replacement therapies in the SC tissue. This platform will allow researchers to perform an ex vivo pre-screening of different cell transplantation therapies, helping them to choose the most appropriate strategy before proceeding with in vivo experiments.

Author Spotlight: Long-Term Spinal Cord Slice Culture for Advancing Spinal Cord Regeneration Therapies

In this paper, we provide a reproducible method to generate and maintain long-term spinal cord organotypic slices transplanted with neural stem cells as an ex vivo model for testing cellular replacement therapies.

Resolutive cures for spinal cord injuries (SCIs) are still lacking, due to the complex pathophysiology. One of the most promising regenerative approaches ...

Standardized Evaluation of DIAm EMG Post-C2SH in Rats to Minimize Variability

Assessing Functional Recovery of Eupneic Diaphragm Activity Following Unilateral Cervical Spinal Cord Hemisection in Rats

Following cSCI, activation of the DIAm can be impacted depending on the extent of the injury. The present manuscript describes a unilateral C2 hemisection (C2SH) model of cSCI that disrupts eupneic ipsilateral diaphragm (iDIAm) electromyographic (EMG) activity during breathing in rats. To evaluate recovery of DIAm motor control, the extent of deficit due to C2SH must first be clearly established. By verifying a complete initial loss of iDIAm EMG during breathing, subsequent recovery can be classified as either absent or present, and the extent of recovery can be estimated using the EMG amplitude. Additionally, by measuring the continued absence of iDIAm EMG activity during breathing after the acute spinal shock period following C2SH, the success of the initial C2SH may be validated. Measuring contralateral diaphragm (cDIAm) EMG activity can provide information about the compensatory effects of C2SH, which also reflects neuroplasticity. Moreover, DIAm EMG recordings from awake animals can provide vital physiological information about the motor control of the DIAm after C2SH. This article describes a method for a rigorous, reproducible, and reliable C2SH model of cSCI in rats, which is an excellent platform for studying respiratory neuroplasticity, compensatory cDIAm activity, and therapeutic strategies and pharmaceuticals.

Author Spotlight: Neuromotor Control and Recovery of Diaphragm Function Following Cervical Spinal Hemisection in Rats

Respiratory complications are the leading cause of death in individuals with cervical spinal cord injury (cSCI). Animal models of cSCI are essential for mechanistic evaluations and pre-clinical studies. Here, we introduce a reproducible method to assess functional recovery of diaphragm muscle (DIAm) activity following unilateral C2 spinal hemisection (C2SH) in rats.

Following cSCI, activation of the DIAm can be impacted depending on the extent of the injury. The present manuscript describes a unilateral C2 hemisection ...

Spinal Cord Surgery and Optical Shank Implantation in Rodents

Implantation of Optoelectronic Devices in the Rodent Spinal Cord

Neuromodulation can provide diagnostic, modulatory, and therapeutic applications. While extensive work has been conducted in the brain, modulation of the spinal cord remains relatively unexplored. The inherently delicate and mobile spinal cord tissue imposes constraints that make the precise implantation of neural probes challenging. Despite recent advances in neuromodulation devices, particularly flexible bioelectronics, opportunities to expand their use in the spinal cord have been limited by the surgical complexities of device implantation. Here, we provide a series of surgical protocols tailored specifically for the implantation of a custom-made optoelectronic device that interfaces with the spinal cord in rodents. The steps to place and anchor an optical shank on a specific segment of the spinal cord via two different surgical implantation methods are detailed here. These methods are optimized for a diverse range of devices and applications, which may or may not require direct contact with the spinal cord for optical stimulation. To elucidate the methodology, the vertebral anatomy is referenced first to identify prominent landmarks before making a skin incision. The surgical steps to secure an optical shank over the cervical spine in rodents are demonstrated. Procedures are then outlined for securing the optoelectronic device connected to the optical shank in a subcutaneous space away from the spinal cord, minimizing unnecessary direct contact. Behavioral studies comparing animals receiving the implants to those undergoing sham surgeries indicate that the optical shanks did not adversely affect hindlimb or forelimb function seven days post-implantation. The present work broadens the neuromodulation toolkit for use in future studies aimed at investigating various spinal cord interventions.

Author Spotlight: Innovative Methodology for Implanting and Securing Neural Probes in the Rodent Spinal Cord

This protocol details a surgical procedure for performing spinal cord surgery and for implanting and securing an optical shank over the spinal cord in rodents.

Neuromodulation can provide diagnostic, modulatory, and therapeutic applications. While extensive work has been conducted in the brain, modulation of the ...