Name: adeno-associated virus-mediated transgene expression in genetically defined neurons of the spinal cord
Uploaded: 2018-05-12T13:00:00
Description: Watch this Scientific Journal Video about Adeno-associated Virus-mediated Transgene Expression in Genetically Defined Neurons of the Spinal Cord at JoVE.com

We thank Hanns Ulrich Zeilhofer for generously supporting this work. Hendrik Wildner was supported by the Olga Mayenfisch foundation. We thank Carmen Birchmeier for the Lmx1b antibody.

All animal experiments were approved by the Swiss cantonal veterinary office (Zurich) and are in accordance and compliance with all relevant regulatory and institutional guidelines.
NOTE: All materials along with respective manufacturers and/or vendors are listed in the Table of Materials.
1. Generation of Cre-dependent AAV Vectors
NOTE: A variety of Cre-dependent vectors with different promoters can be purchased (see <stro...

<ol>
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J., Dymecki, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryptic+boundaries+in+roof+plate+and+choroid+plexus+identified+by+intersectional+gene+activation.">Cryptic boundaries in roof plate and choroid plexus identified by intersectional gene activation.</a> Nat Genet. 35 (1), 70-75 (2003).</li><li>Kim, J. 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=Linking+genetically+defined+neurons+to+behavior+through+a+broadly+applicable+silencing+allele.">Linking genetically defined neurons to behavior through a broadly applicable silencing allele.</a> Neuron. 63 (3), 305-315 (2009).</li><li>Kim, J. C., Dymecki, S. 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Intraspinal injection of AAVs may become a powerful technique in a research laboratory, enabling the analysis of spinal cells with a high temporal and spatial solution. This protocol enables the transduction of the three main spinal segments innervated by sensory neurons extending their peripheral axons to the hindlimb. Transducing three segments produces robust and reproducible behavioral data. It also enables testing of a larger sensory area than possible after a single intraspinal injection. For example, the same inje...

The dorsal spinal cord is essential for information exchange between the periphery of the body and the brain. Sensory stimuli such as heat, cold, touch, or noxious stimuli are detected by specialized peripheral neurons, which convey this information to neurons of the spinal cord dorsal horn. Here, a complex network of inhibitory and excitatory interneurons modulates and eventually relays sensory information via spinal projection neurons to supraspinal sites1,2. The computations carried out by spinal inter- and projection neurons gate sensory information, thus determining which information is suppressed or relayed at which intensity. Changes in the integration of sensory stimuli, such as an altered balance between inhibition and excitation, can cause sensory dysfunctions such as hypersensitivity or allodynia (painful sensations after normally non-painful stimulation). These changes are thought to be the underlying cause of various chronic pain states3,4. Thus, spinal circuits are of high importance in sensory processing and consequently in the perception of an organism&#39;s environment and self. With the recent advent and combination of molecular, genetic, and surgical techniques that allow the precise manipulation of genetically identified spinal neuron subpopulations, scientists are now beginning to understand the underlying spinal circuits responsible for the processing of distinct sensory modalities.
Intraspinal injection of rAAV into wild-type or transgenic mice has greatly contributed to the manipulation, analysis, and understanding of the function of specific subsets of spinal neurons5,6,7,8,9,10,11. This technique allows the delivery of marker proteins (such as GFP/ GFP fusion proteins), reporter proteins (such as GCaMP), or effector proteins (such as bacterial toxins, channelrhodopsin, or pharmacogenetic receptors) in a spatially restricted manner to spinal neurons. Local injection of Cre-dependent rAAVs into transgenic mice expressing Cre recombinase in a specific subset of spinal neurons allows the specific analysis of the respective neuronal population. We have employed this technique to label, ablate, inhibit or activate spinal glycinergic neurons demonstrating that they are an essential part of the spinal gate controlling pain and itch transmission7. In these experiments, intraspinal injection of Cre-dependent rAAV into GlyT2::Cre mice enabled the selective manipulation of glycinergic neurons in the lumbar spinal cord. Thereby, simultaneous manipulation of supraspinal circuits that contain glycinergic neurons critical for the survival of the animal can be avoided.
While an intraspinal injection of rAAVs limits infection to the site of injection, viral transduction can occur not only in local neurons but also in neurons that connect to the injection site via axonal projections. The latter is often used to trace CNS areas providing neuronal input to a particular nucleus in the brain. The infection of axonal projections can, however, also be a confounding factor when a defined population of neurons shall be studied at a particular site. To address these issues, we have recently conducted a comprehensive analysis of AAV serotypes and expression cassettes to identify serotypes and promoters that can be used to either minimize or maximize retrograde transduction. In the context of this specific research in spinal circuits, we analyzed the ability of different serotypes and promoters to retrogradely transduce neurons in the dorsal root ganglia (DRG), the rostral ventromedial medulla (RVM), and the somatosensory cortex12. The technique outlined in this protocol can therefore be used either to analyze spinal neurons at the injection site or to analyze projection neurons that provide input to the injected site of the spinal cord. In the protocol described here, three injections of rAAV into the left side of the lumbar spinal cord are performed to enable transduction of neurons in the three lumbar segments (L3-L5). The L3-L5 segments receive the majority of the sensory input from the hindlimb ipsilateral to the injection site. We demonstrate that functional manipulation of genetically labeled neurons in L3-L5 is sufficient to evoke robust behavioral changes, thus providing functional evidence for the circuit function of such a genetically labeled neuron subtype.

<table>
	<tbody>
		<tr>
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>micropipette puller: DMZ-Universal-Electrode-Puller</td>
			<td>Zeitz</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>anesthesia unit: Oxymat3 oxygen concentrator</td>
			<td>Weinmann</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>anesthesia unit: VIP 3000 Veterinary Vaporizer</td>
			<td>Midmark</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Heat mat: Mio Star Thermocare 100</td>
			<td>Migros</td>
			<td>717614700000</td>
			<td></td>
		</tr>
		<tr>
			<td>Electric shaver</td>
			<td>Philips</td>
			<td>BT9290</td>
			<td></td>
		</tr>
		<tr>
			<td>surgical microscope (OPMI pico)</td>
			<td>Zeiss</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Small animal stereotaxic apparatus</td>
			<td>Kopf</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurostar StereoDrive (optional)</td>
			<td>Neurostar</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Model 51690 Cunningham mouse spinal adaptor</td>
			<td>Harvard Apparatus</td>
			<td>72-4811</td>
			<td></td>
		</tr>
		<tr>
			<td>PHD Ultra syringe pump with nanomite</td>
			<td>Harvard Apparatus</td>
			<td>70-3601</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton 701 RN 10 &#956;l glass microliter syringe</td>
			<td>Hamilton</td>
			<td>7635-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton Removable needle (RN) compression fitting 1 mm</td>
			<td>Hamilton</td>
			<td>55750-01</td>
			<td></td>
		</tr>
		<tr>
			<td>fine dentistry drilling apparatus: Osada success 40</td>
			<td>Osada</td>
			<td>OS-40</td>
			<td></td>
		</tr>
		<tr>
			<td>spherical cutter, 0.5mm</td>
			<td>Busch</td>
			<td>12001005B</td>
			<td></td>
		</tr>
		<tr>
			<td>electronic von Frey anesthesiometer</td>
			<td>IITC</td>
			<td>23905</td>
			<td></td>
		</tr>
		<tr>
			<td>flexible von Frey hairs</td>
			<td>IITC</td>
			<td>#7</td>
			<td></td>
		</tr>
		<tr>
			<td>LSM710 Pascal confocal microscope</td>
			<td>Zeiss</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>0.8 NA &#215; 20 Plan-apochromat objective</td>
			<td>Zeiss</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>1.3 NA &#215; 40 EC Plan-Neofluar oil-immersion objective</td>
			<td>Zeiss</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Surgical Tools</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel Handle #4, 13cm</td>
			<td>Fine Science Tools</td>
			<td>10004-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Extra Fine Bonn Scissors</td>
			<td>Fine Science Tools</td>
			<td>14084-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Adson forceps, 1 x 2 teeth, 12 cm</td>
			<td>Fine Science Tools</td>
			<td>11027-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Friedman-Pearson rongeurs, curved, 0.7 mm cup</td>
			<td>Fine Science Tools</td>
			<td>16121-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #2 laminectomy forceps</td>
			<td>Fine Science Tools</td>
			<td>11223-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Olsen-Hegar needle holders, serrated, 8.5 mm clamp length</td>
			<td>Fine Science Tools</td>
			<td>12002-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine forceps #5</td>
			<td>Fine Science Tools</td>
			<td>11254-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Consumables and Chemicals</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Thin-wall glass capillary, 1mm outside diameter</td>
			<td>World Precision Instruments</td>
			<td>TW 100-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringes (1, 5 and 20 ml)</td>
			<td>B. Braun</td>
			<td>(9166917V, 4606051V, 4606205V)</td>
			<td></td>
		</tr>
		<tr>
			<td>26G beveled needle</td>
			<td>B. Braun</td>
			<td>4665457</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile scalpel blades</td>
			<td>B. Braun</td>
			<td>BB523</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical sutures Safil Quick+ 4/0, absorbable</td>
			<td>B. Braun</td>
			<td>C1046220</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical sutures Premilene 5/0, non-absorbable</td>
			<td>B. Braun</td>
			<td>C0932191</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile PBS or saline (0.9%)</td>
			<td></td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol, 70% (disinfectant)</td>
			<td></td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Iodine solution (e.g. Braunol)</td>
			<td>B. Braun</td>
			<td>18380</td>
			<td></td>
		</tr>
		<tr>
			<td>Anaesthetics&#160; (e.g. Attane isoflurane)</td>
			<td>Provet</td>
			<td>2222</td>
			<td></td>
		</tr>
		<tr>
			<td>Aldasorber</td>
			<td>Provet</td>
			<td>333526</td>
			<td></td>
		</tr>
		<tr>
			<td>analgesics (e.g. buprenorphine: temgesic)</td>
			<td>Indivior</td>
			<td>GTIN: 7680419310018</td>
			<td></td>
		</tr>
		<tr>
			<td>Ophthalmic ointment (e.g. vita-pos)</td>
			<td>Pharma medica</td>
			<td>GTIN: 4031626710635</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton swabs (e.g. from)</td>
			<td>IVF Hartmann</td>
			<td>1628100</td>
			<td></td>
		</tr>
		<tr>
			<td>Facial tissues (e.g. from)</td>
			<td>Uehlinger AG</td>
			<td>2015.10018</td>
			<td></td>
		</tr>
		<tr>
			<td>Superfrost plus microscope slides</td>
			<td>ThermoScientific</td>
			<td>J1800AMNZ</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Mice</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6J mice&#160; (wildtype)</td>
			<td>The Jackson Laboratory</td>
			<td>RRID:IMSR_JAX:000664</td>
			<td></td>
		</tr>
		<tr>
			<td>Rorbtm1.1(cre)Hze/J mice (ROR&#946;Cre)</td>
			<td>The Jackson Laboratory</td>
			<td>RRID:IMSR_JAX:023526</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J mice (R26Tom)</td>
			<td>The Jackson Laboratory</td>
			<td>RRID: IMSR_JAX:007914</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Viral vectors</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>AAV1.CB7.CI.eGFP.WPRE.rBG (AAV1.CAG.eGFP)</td>
			<td>Penn Vector Core</td>
			<td>AV-1-PV1963</td>
			<td></td>
		</tr>
		<tr>
			<td>AAV1.CAG.flex.eGFP.WPRE.bGH (AAV1.CAG.flex.eGFP)</td>
			<td>Penn Vector Core</td>
			<td>AV-1-ALL854</td>
			<td></td>
		</tr>
		<tr>
			<td>AAV1.CAG.flex.tdTomato.WPRE.bGH (AAV1.CAG.flex.tdTomato)</td>
			<td>Penn Vector Core</td>
			<td>AV-1-ALL864</td>
			<td></td>
		</tr>
		<tr>
			<td>AAV1.EF1a.flex.DTA.hGH (AAV1.EF1a.flex.DTA)</td>
			<td>Penn Vector Core</td>
			<td>Custom production</td>
			<td></td>
		</tr>
		<tr>
			<td>AAV1.hSyn.DIO.hM3D(Gq)-mCherry.hGH (AAV.flex.hM3D(Gi))</td>
			<td>Penn Vector Core</td>
			<td>Custom production</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Plasmids</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>pAAV.hSyn.flex.hM3D(Gq)-mCherry</td>
			<td>Addgene</td>
			<td>44361</td>
			<td></td>
		</tr>
		<tr>
			<td>pAAV.EF1&#945;.flex.hChR2(H134R)-eYFP</td>
			<td>Addgene</td>
			<td>20298</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Bacteria</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MDS42</td>
			<td>ScarabGenomics</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stbl3</td>
			<td>ThermoScientific</td>
			<td>C737303</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>EndoFree Plasmid Maxi Kit</td>
			<td>Quiagen</td>
			<td>12362</td>
			<td></td>
		</tr>
		<tr>
			<td>NucleoBond PC 500</td>
			<td>Machery &#38; Nagel</td>
			<td>740574</td>
			<td></td>
		</tr>
		<tr>
			<td>clozapine-N-oxide (CNO)</td>
			<td>Enzo Life Sciences</td>
			<td>BBL-NS105-0025</td>
			<td></td>
		</tr>
		<tr>
			<td>chloroquine diphosphate salt</td>
			<td>Sigma</td>
			<td>C6628</td>
			<td></td>
		</tr>
		<tr>
			<td>histamine</td>
			<td>Sigma</td>
			<td>H7125</td>
			<td></td>
		</tr>
		<tr>
			<td>Dapi</td>
			<td>Invitrogen</td>
			<td>D3571</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Antibodies (dilution)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit anti-GFP (1:1000)</td>
			<td>Molecular Probes</td>
			<td>RRID:AB_221570</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit anti-NeuN (1:3000)</td>
			<td>Abcam</td>
			<td>RRID:AB_10711153</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti-Pax2 (1 : 200)</td>
			<td>R &#38; D Systems</td>
			<td>RRID:AB_10889828</td>
			<td></td>
		</tr>
		<tr>
			<td>Guinea pig anti-Lmx1b (1 : 10 000)</td>
			<td>Dr Carmen Birchmeier</td>
			<td>Muller et al. 2002</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit anti-GFAP (1 : 1000)</td>
			<td>DakoCytomation</td>
			<td>RRID:AB_10013382</td>
			<td></td>
		</tr>
		<tr>
			<td>Secondary antibodies raised in donkey (1:800)</td>
			<td>Jackson ImmunoResearch Laboratories</td>
			<td>NA</td>
			<td></td>
		</tr>
	</tbody>
</table>

adeno-associated virus-mediated transgene expression in genetically defined neurons of the spinal cord

Selective manipulation of spinal neuronal subpopulations has mainly been achieved by two different methods: 1) Intersectional genetics, whereby double or triple transgenic mice are generated in order to achieve selective expression of a reporter or effector gene (e.g., from the Rosa26 locus) in the desired spinal population. 2) Intraspinal injection of Cre-dependent recombinant adeno-associated virus (rAAV); here Cre-dependent AAV vectors coding for the reporter or effector gene of choice are injected into the spinal cord of mice expressing Cre recombinase in the desired neuronal subpopulation. This protocol describes how to generate Cre-dependent rAAV vectors and how to transduce neurons in the dorsal horn of the lumbar spinal cord segments L3-L5 with rAAVs. As the lumbar spinal segments L3-L5 are innervated by those peripheral sensory neurons that transmit sensory information from the hindlimbs, spontaneous behavior and responses to sensory tests applied to the hindlimb ipsilateral to the injection side can be analyzed in order to interrogate the function of the manipulated neurons in sensory processing. We provide examples of how this technique can be used to analyze genetically defined subsets of spinal neurons. The main advantages of virus-mediated transgene expression in Cre transgenic mice compared to classical reporter mouse-induced transgene expression are the following: 1) Different Cre-dependent rAAVs encoding various reporter or effector proteins can be injected into a single Cre transgenic line, thus overcoming the need to create several multiple transgenic mouse lines. 2) Intraspinal injection limits manipulation of Cre-expressing cells to the injection site and to the time after injection. The main disadvantages are: 1) Reporter gene expression from rAAVs is more variable. 2) Surgery is required to transduce the spinal neurons of interest. Which of the two methods is more appropriate depends on the neuron population and research question to be addressed.

In order to illustrate the expression levels that can be obtained by the intraspinal injection of rAAV encoding a marker protein, we first injected AAV1.CAG.eGFP into the lumbar spinal cord of wild-type mice. Three injections spaced approximately 1 mm apart produced a nearly continuous infection of lumbar spinal segments L3 to L5 (Figure 1A-C). Virus injection at a depth of 300 &#181;m from the spinal surface leads to predominant infection of...

Watch this Scientific Journal Video about Adeno-associated Virus-mediated Transgene Expression in Genetically Defined Neurons of the Spinal Cord at JoVE.com

Adeno-associated Virus-mediated Transgene Expression in Genetically Defined Neurons of the Spinal Cord

Intraspinal injection of recombinase dependent recombinant adeno-associated virus (rAAV) can be used to manipulate any genetically labelled cell type in the spinal cord. Here we describe how to transduce neurons in the dorsal horn of the lumbar spinal cord. This technique enables functional interrogation of the manipulated neuron subtype.

Selective manipulation of spinal neuronal subpopulations has mainly been achieved by two different methods: 1) Intersectional genetics, whereby double or ...

The overall goal of intraspinal delivery of recombinant adeno-associated viruses is to enable the histological labeling and functional manipulation of genetically defined neurons in the dorsal spinal cord. This method can help answer key questions in the pain field, such as which neuronal circuits are required for the perception of different pain modalities. The main advantage of this technique is that genetically labeled neurons can be manipulated in a specially restricted and temporally controlled fashion.Locate the most caudal rib pair by palpating along the vertebral column of the anesthetized mouse. Then, make a 1.5 to 2.5 centimeter longitudinal cut into the skin, starting from just rostral to the most caudal rib pair. Lift the skin with forceps and detach it from the underlying muscle with scissors.Throughout the surgery, keep the exposed tissues moist with sterile 0.9%sodium chloride. Using fine forceps and small scissors, make an incision into the next to thin membranous layer, right next to the midline, and cut it away from the spinous processes. Several anatomical landmark can aid in identifying the correct vertebrae to target.Here, we describe three alternatives. Palpate along the vertebral column. The most caudal rib pair is located just rostral to the T13 vertebrae and the pelvic bone at hip level is at the level of the L6 vertebrae.Alternatively, pull back the skin towards the tail to expose the iliac crest. At the same level, the most caudal pair of visible intetransverse ligaments joins the L6 spinal process. Count backwards in a caudal to rostral direction to identify the vertebrae of interest.Otherwise, locate the spot where the tendon along the side of the vertebral column is whitest and most medial. The T13 vertebrae is located just rostral. The lumbar spinal cord segment L4 is located within this vertebrae.Next, place the animal onto a cushion of rolled up tissues to elevate it to the spinal clamps of the stereotaxic frame. Align the clamps adjacent to the target vertebrae. Fix one clamp in position, then hold the vertebral column with Adson forceps while fixing the second clamp.Confirm proper clamping by carefully pressing on the targeted vertebrae. Then, remove the paraspinous muscle above the vertebrae of interest. First, make an incision just medial to the tendons parallel to the vertebral column.Then, make perpendicular incisions rostral and caudal to the target vertebrae. Next, use rongeurs to tear or cut away the paraspinous muscle above the vertebrae of interest. Use forceps as needed to remove tissue remaining on the vertebrae or above the dura in the intravertebral space.The dorsal blood vessel should now be visible, marking the midline of the spinal cord. For unilateral injections, perform a partial laminectomy using a fine dentist drill equipped with a 0.5 millimeter spherical cutter to very carefully drill a hole into the middle of the target side of the vertebrae. Remove any remaining bone fragments with a 26 gauge beveled needle to expose the spinal cord.Use the needle to perforate the dura in the intravertebral spaces rostral and caudal to the target vertebrae, approximately 300 microns lateral to the dorsal blood vessel. Then, perforate the dura underneath the drilled hole. At this point, cerebrospinal fluid should escape from the holes and the spinal cord should be slightly bulging out.Prepare the injection syringe by first mounting the glass capillary onto a microliter syringe. Fasten the nut tightly to ensure a tight and secure fit. Fill the microliter syringe with sterile distilled water, then depress the plunger.If the water is not easily expelled from the tip, the micropipette is likely blocked and should be exchanged. Mount the syringe onto a micromanipulator connected to an electronically controlled microinjector. Then, use the microinjector to draw up about one microliter of air to separate the water and virus.Next, pipette a 2.5 microliter droplet of diluted virus solution onto a piece of paraffin film. Carefully move the tip of the micropipette into the droplet using the sterotaxic frame and draw it up into the micropipette. Then, press dispense until a drop of virus solution emerges at the tip.Retract the syringe and use a pen to mark the micropipette with a scale to monitor the dispensed volume. Use the stereotaxic arm to move the tip of the micropipette over one of the holes in the dura and then down until a slight depression appears on the surface of the exposed spinal cord. To target the injection to the spinal dorsal horn, move the tip down to a depth of 500 microns in 100 micron increments and then up 200 microns to allow for mechanical stabilization of the tissue.The final depth of the tip is 300 microns. After programming the pump to inject 300 nanoliters at an injection speed of 50 nanoliters per minute, press the start button to begin the infusion. After the injection is complete, check the scale on the micropipette to see whether the virus level has dropped.Leave the micropipette in place for an additional three minutes to allow the pressure to equilibrate. After three minutes, slowly retract the micropipette and then repeat the procedure for the other two injection sites. After the final injection, remove the spinal clamps and the cushion below the mouse.Close the incision layers with interrupted stitches, suturing the superficial tissue layers with absorbable sutures and the skin with non-absorbable sutures. Apply iodine disinfectant on the sutured wound. Terminate the anesthesia and leave the animal on the heat mat until it recovers before returning it to its home cage.Spinal tissue at the injection site is to be examined at the end of the experiment. This is necessary to verify successful injection and transduction and to exclude excessive tissue damage as a result of the surgery. To illustrate the expression levels that can be obtained by intraspinal injection of recombinant AAV, a virus encoding the eGFP fluorescent protein was injected into the lumbar spinal cord of wild-type mice.Three injections spaced approximately one millimeter apart produced a nearly continuous infection of lumbar spinal segments L3 to L5.Virus injection at a depth of 300 microns from the spinal surface leads to predominant infection of cells in the spinal cord dorsal horn. However, infected cells could also be found in the ventral horn. A cre dependent flex vector expressing eGFP was injected into the spinal cord of GlyT2:Cre transgenic mice.This resulted in more restricted eGFP expression reflecting the distribution of GLYT2 positive neurons. Following this protocol will allow targeting of three consecutive spine segments. We found that this is sufficient to obtain robust behavior results.This technique will allow studying the function of specific neuron and cell populations and sensory pathways.

adeno-associated-virus-mediated-transgene-expression-genetically

Research

JoVE Journal

Neuroscience

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 ...

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

Viral Tracing of Genetically Defined Neural Circuitry

Classical methods for studying neuronal circuits are fairly low throughput. Transsynaptic viruses, particularly the pseudorabies (PRV) and rabies virus (RABV), and more recently vesicular stomatitis virus (VSV), for studying circuitry, is becoming increasingly popular. These higher throughput methods use viruses that transmit between neurons in either the anterograde or retrograde direction.
Recently, a modified RABV for monosynaptic retrograde tracing was developed. (Figure 1A). In this method, the glycoprotein (G) gene is deleted from the viral genome, and resupplied only in targeted neurons. Infection specificity is achieved by substituting a chimeric G, composed of the extracellular domain of the ASLV-A glycoprotein and the cytoplasmic domain of the RABV-G (A/RG), for the normal RABV-G1. This chimeric G specifically infects cells expressing the TVA receptor1. The gene encoding TVA can been delivered by various methods2-8. Following RABV-G infection of a TVA-expressing neuron, the RABV can transmit to other, synaptically connected neurons in a retrograde direction by nature of its own G which was co-delivered with the TVA receptor. This technique labels a relatively large number of inputs (5-10%)2 onto a defined cell type, providing a sampling of all of the inputs onto a defined starter cell type.
We recently modified this technique to use VSV as a transsynaptic tracer9. VSV has several advantages, including the rapidity of gene expression. Here we detail a new viral tracing system using VSV useful for probing microcircuitry with increased resolution. While the original published strategies by Wickersham et al.4 and Beier et al.9 permit labeling of any neurons that project onto initially-infected TVA-expressing-cells, here VSV was engineered to transmit only to TVA-expressing cells (Figure 1B). The virus is first pseudotyped with RABV-G to permit infection of neurons downstream of TVA-expressing neurons. After infecting this first population of cells, the virus released can only infect TVA-expressing cells. Because the transsynaptic viral spread is limited to TVA-expressing cells, presence of absence of connectivity from defined cell types can be explored with high resolution. An experimental flow chart of these experiments is shown in Figure 2. Here we show a model circuit, that of direction-selectivity in the mouse retina. We examine the connectivity of starburst amacrine cells (SACs) to retinal ganglion cells (RGCs).

A method of tracing synaptically connected neurons is described. We use TVA specificity of an upstream cell to probe whether a cell population of interest receives synaptic input from genetically defined cell types.

Classical methods for studying neuronal circuits are fairly low throughput. Transsynaptic viruses, particularly the pseudorabies (PRV) and rabies virus ...

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 ...

RNA-Seq Analysis of Differential Gene Expression in Electroporated Chick Embryonic Spinal Cord

In ovo electroporation of the chick neural tube is a fast and inexpensive method for identification of gene function during neural development. Genome wide analysis of differentially expressed transcripts after such an experimental manipulation has the potential to uncover an almost complete picture of the downstream effects caused by the transfected construct. This work describes a simple method for comparing transcriptomes from samples of transfected embryonic spinal cords comprising all steps between electroporation and identification of differentially expressed transcripts. The first stage consists of guidelines for electroporation and instructions for dissection of transfected spinal cord halves from HH23 embryos in ribonuclease-free environment and extraction of high-quality RNA samples suitable for transcriptome sequencing. The next stage is that of bioinformatic analysis with general guidelines for filtering and comparison of RNA-Seq datasets in the Galaxy public server, which eliminates the need of a local computational structure for small to medium scale experiments. The representative results show that the dissection methods generate high quality RNA samples and that the transcriptomes obtained from two control samples are essentially the same, an important requirement for detection of differential expression genes in experimental samples. Furthermore, one example is provided where experimental overexpression of a DNA construct can be visually verified after comparison with control samples. The application of this method may be a powerful tool to facilitate new discoveries on the function of neural factors involved in spinal cord early development.

This video shows the basic steps for performing whole transcriptome analysis on dissected chick embryonic spinal cord samples after transfection with in ovo electroporation.

In ovo electroporation of the chick neural tube is a fast and inexpensive method for identification of gene function during neural development. Genome ...

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing organisms, combined with the embryonic optical clarity, served as initial compelling attributes of this model. Over the past two decades, the success of this model has been further propelled by its amenability to large-scale mutagenesis screens and by the ease of transgenesis. More recently, gene-editing approaches have extended the power of the model.
For neurodevelopmental studies, the zebrafish embryo and larva provide a model to which multiple methods can be applied. Here, we focus on methods that allow the study of an essential property of neurons, electrical excitability. Our preparation for the electrophysiological study of zebrafish spinal neurons involves the use of veterinarian suture glue to secure the preparation to a recording chamber. Alternative methods for recording from zebrafish embryos and larvae involve the attachment of the preparation to the chamber using a fine tungsten pin1,2,3,4,5. A tungsten pin is most often used to mount the preparation in a lateral orientation, although it has been used to mount larvae dorsal-side up4. The suture glue has been used to mount embryos and larvae in both orientations. Using the glue, a minimal dissection can be performed, allowing access to spinal neurons without the use of an enzymatic treatment, thereby avoiding any resultant damage. However, for larvae, it is necessary to apply a brief enzyme treatment to remove the muscle tissue surrounding the spinal cord. The methods described here have been used to study the intrinsic electrical properties of motor neurons, interneurons, and sensory neurons at several developmental stages6,7,8,9.

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

This manuscript describes methods for electrophysiological recordings from spinal neurons of zebrafish embryos and larvae. The preparation maintains neurons in situ and often involves minimum dissection. These methods allow for the electrophysiological study of a variety of spinal neurons, from the initial electrical excitability acquisition through the early larval stages.

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing ...

Spinal Cord Neurons Isolation and Culture from Neonatal Mice

We present a protocol for the isolation and culture of spinal cord neurons. The neurons are obtained from neonatal C57BL/6 mice and are isolated on postnatal day 1-3. A mouse litter, usually 4-10 pups born from one breeding pair, is gathered for one experiment, and spinal cords are collected individually from each mouse after euthanasia with isoflurane. The spinal column is dissected out and then the spinal cord is released from the column. The spinal cords are then minced to increase the surface area of delivery for an enzymatic protease that allows for the neurons and other cells to be released from the tissue. Trituration is then used to release the cells into solution. This solution is subsequently fractionated in a density gradient to separate the various cells in solution, allowing for neurons to be isolated. Approximately 1-2.5 x 106 neurons can be isolated from one litter group. The neurons are then seeded onto wells coated with adhesive factors that allow for proper growth and maturation. The neurons take approximately 7 days to reach maturity in the growth and culture medium and can be used thereafter for treatment and analysis.

This study presents a technique for the isolation of neurons from WT neonatal mice. It requires the careful dissection of the spinal cord from the neonatal mouse, followed by the separation of neurons from the spinal cord tissue through mechanical and enzymatic cleavage.

We present a protocol for the isolation and culture of spinal cord neurons. The neurons are obtained from neonatal C57BL/6 mice and are isolated on ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to intracranial delivery, intra-arterial administration of NSCs is less invasive and produces a more diffuse distribution of NSCs within the brain parenchyma. Further, intra-arterial delivery allows the first-pass effect in the brain circulation, lessening the potential for trapping of cells in peripheral organs, such as liver and spleen, a complication associated with peripheral injections. Here, we detail the methodology, in both mice and rats, for delivery of NSCs through the common carotid artery (mouse) or external carotid artery (rat) to the ipsilateral hemisphere after an ischemic stroke. Using GFP-labeled NSCs, we illustrate the widespread distribution achieved throughout the rodent ipsilateral hemisphere at 1 d, 1 week and 4 weeks after postischemic delivery, with a higher density in or near the ischemic injury site. In addition to long-term survival, we show evidence of differentiation of GFP-labeled cells at 4 weeks. The intra-arterial delivery approach described here for NSCs can also be used for administration of therapeutic compounds, and thus has broad applicability to varied CNS injury and disease models across multiple species.

A method for delivering neural stem cells, adaptable for injecting solutions or suspensions, through the common carotid artery (mouse) or external carotid artery (rat) after ischemic stroke is reported. Injected cells are distributed broadly throughout the brain parenchyma and can be detected up to 30 d after delivery.

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to ...

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 ...