This work was supported by the Heart and Stroke Foundation, Ontario Institute of Regenerative Medicine, Canada First Research Excellence Fund (Medicine by Design, MbD), and Connaught Innovation Award (University of Toronto).

This protocol was approved by the Animal Care Committee at the University of Toronto and adheres to the Guide to the Care and Use of Experimental Animals (2nd Edition, Canadian Council on Animal Care, 2017). For this study, wild-type (C57BL/6J) and transgenic tdTom-Cre reporter (B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J)22 mouse strains were used. Mice were 7-9 weeks old and included both males and females. Details of the reagents and equipment used are listed in the Table of Materials.
1. Endothelin-1 stroke surgery
NOTE: All surgical instruments and materials are sterilized by autoclave prior to surgery. The stereotaxic frame is sterilized with PREempt. Syringes are sterilized with PREempt, 70% alcohol, and primed with sterile PBS prior to surgery. Endothelin-1 (ET-1) is stored on ice throughout the surgery.
<ol>
	<li>Before starting the surgery, set up the recovery area by placing a clean cage on a heating pad set to 37 &#176;C. Alternatively, if a heating lamp is being used, place only half of the cage under the heat.
	<ol>
		<li>To anesthetize mice using isoflurane, follow the protocol approved by the Animal Care Committee. Place the mouse in the anesthetic induction chamber and turn on the oxygen to 1-1.5 L/min, then adjust the isoflurane vaporizer to 3%-5% to initiate anesthesia.</li>
	</ol>
	</li>
	<li>Remove the animal from the chamber once it lies on its side and can no longer stand, and place it onto a clean surface. Ensure the animal is adequately anesthetized by confirming the absence of a toe pinch reflex before and throughout the surgery.
	<ol>
		<li>Secure the nose cone extended from the anesthetic machine on the animal and adjust isoflurane to 1%-2% for maintenance. Minimize the exposure to isoflurane (5 min at 3%-5% for induction and the remaining time at 1%-2%), as the procedure can take up to 40 min to complete.</li>
	</ol>
	</li>
	<li>Use animal hair trimmers to remove fur from the neck/behind ears to the nose of the animal to expose the skin on the skull for surgery.</li>
	<li>Weigh the animal and administer the appropriate dose of pre-operative analgesic subcutaneously (Meloxicam (2.0 mg/kg)) based on the weight.</li>
	<li>Apply eye lubricant ointment on each eye of the mouse to prevent corneal desiccation during anesthesia.</li>
	<li>Sterilize the fur-clipped skin area (step 1.3) with 70% ethanol, followed by Chlorhexidine alcohol twice to form a surgical field.</li>
	<li>Transfer the mouse onto a mouse stereotaxic frame by first placing its nose in the stereotaxic nose cone, then stabilize the skull with ear bars. Keep the mouse on a heating pad set to 37 &#176;C for thermal support throughout the surgery procedure. Maintain isoflurane at 1%-2% so that the mouse displays a respiratory rate of 1 breath/s.</li>
	<li>Repeat application of eye lubrication to prevent corneal desiccation during the procedure if needed. Apply a sterile surgical drape on the body of the mouse, below the ears, to extend the size of the sterile field.</li>
	<li>Make a vertical incision parallel to the sagittal plane of the animal from behind the mid-point of the eyes to the back of the head (parietal bone) with a #10 scalpel blade. Expose the skull by gently retracting the skin towards the sides.</li>
	<li>Use a Q-tip to disrupt the fascia on the skull and allow the skull to dry to expose bregma.</li>
	<li>Set up manipulator arms with drill holder carrying high-speed stereotaxic drill with a .018&#34; drill bit.</li>
	<li>Drill 3 burr holes (.018&#34; diameter) with a high-speed stereotaxic drill at the coordinates in the right sensorimotor cortex at 2.25 mm lateral to bregma and +0.0 mm rostral, +1.0 mm rostral and -1.0 mm caudal to bregma.</li>
	<li>Replace the drill with a 26 G syringe with a 0.375&#8243; long needle.</li>
	<li>Load 1.5 &#181;L of 400 &#181;M ET-1 dissolved in sterile PBS into the syringe. To ensure good flow, release a small volume of ET-1 until a drop can be observed at the tip of the needle. Use a sterile Q-tip to wipe off the excess before performing an injection.</li>
	<li>Lower the tip of the needle past the skull in the middle burr hole (in the right sensorimotor cortex at + 0.0 anterior and 2.25 mm lateral to bregma) without puncturing the dura mater. Once aligned, lower the needle 1 mm into the brain.</li>
	<li>Inject 1 &#181;L of ET-1 by dispensing 0.1 &#181;L at a time. Push the plunger of the Hamilton syringe forward to dispense 0.1 &#181;L and wait for a minute before injecting the next 0.1 &#181;L, inject until 1 &#181;L has been dispensed.
	<ol>
		<li>After the final 0.1 &#181;L has been dispensed (0.1 &#181;L x 10 injections = 1 &#181;L total), keep the needle in place for 10 min to prevent backflow and then slowly withdraw upward to remove the syringe.</li>
	</ol>
	</li>
	<li>To ensure the needle does not become blocked during injection, release a small volume of the ET-1 until a drop can be observed at the tip of the needle. Use a sterile Q-tip to wipe off the excess before performing an injection. Sterilize syringes before injecting the next animal with PREempt, followed by 70% alcohol, and then sterile PBS.</li>
	<li>To close the wound, suture the overlying skin on the skull together using a 4-0 polysorb polyester sterile suture.</li>
	<li>Administer antibiotic ointment using a cotton swab on the sutured area to prevent post-operative infection.</li>
	<li>Remove the mouse from the stereotaxic frame and transfer it to a clean, pre-heated animal housing cage in the recovery area. Turn off the isoflurane vaporizer and oxygen.</li>
	<li>Keep the cage on top of a heating pad for the duration of recovery from anesthesia. Alternatively, if using a heating lamp place the mouse on the side of the cage under the lamp so that the mouse can move to the cool side of the cage as it wakes up from anesthesia to prevent overheating. Monitor the mouse as it comes out of anesthesia in the next 5-10 min.</li>
	<li>Monitor mouse over the next 3 days (weight monitoring), provide post-operative care (moist mashed chow to ensure hydration), and analgesics (2 mg/kg meloxicam, 2x daily subcutaneously, for 2-3 days) in accordance with regulations set out by local animal care committee.</li>
</ol>
2. Adeno-Associated Virus (AAV) delivery (7 days - subacute model or 21 days - chronic model) post-stroke
CAUTION: Refer to institutional biosafety guidelines for working with AAVs. As per the guidelines at the University of Toronto, this surgery is performed in a Containment Level 2 Biosafety Cabinet.
NOTE: All surgical instruments and materials are sterilized by autoclave prior to surgery. The stereotaxic frame is sterilized with PREempt. Syringes are sterilized with PREempt, 70% alcohol, and primed with sterile PBS. AAVs are stored on ice throughout the surgery.
<ol>
	<li>Before starting the surgery, set up the recovery area by placing a clean cage on a heating pad set to 37 &#176;C. Alternatively, if a heating lamp is being used, place only half of the cage under the heat.
	<ol>
		<li>To anesthetize mice using isoflurane, place the mouse in the anesthetic induction chamber and turn on the oxygen to 1-1.5 L/min, then adjust the isoflurane vaporizer to 3%-5% to initiate anesthesia (following the instutionally approved protocols).</li>
	</ol>
	</li>
	<li>Remove the animal from the chamber once it lies on its side and can no longer stand on a clean surface. Ensure the animal is adequately anesthetized by confirming the absence of a toe pinch reflex before and throughout the surgery.
	<ol>
		<li>Secure the nose cone extended from the anesthetic machine on the animal and adjust isoflurane to 1%-2% for maintenance. Minimize the exposure to isoflurane (5 min at 3%-5% for induction and the remaining time at 1%-2%), as the procedure can take up to 40 min to complete.</li>
	</ol>
	</li>
	<li>Weigh the animal and administer the appropriate dose of pre-operative analgesic subcutaneously (Meloxicam (2.0 mg/kg)) based on the weight.</li>
	<li>Apply eye lubricant ointment on each eye of the mouse to prevent corneal desiccation from being under anesthesia.</li>
	<li>Sterilize the fur-clipped skin area with 70% ethanol, followed by Chlorhexidine alcohol twice to form a surgical field.</li>
	<li>Transfer the mouse onto a mouse stereotaxic frame by first placing its nose in the stereotaxic nose cone, then stabilize the skull with ear bars. Keep the mouse on a heating pad set to 37 &#176;C for thermal support throughout the surgery procedure. Maintain isoflurane at 1%-2% so that the mouse displays the respiratory rate of 1 breath/s.</li>
	<li>Repeat application of eye lubrication to prevent corneal desiccation during the procedure if needed. Apply a sterile surgical drape on the body of the mouse, below the ears, to extend the size of sterile field.</li>
	<li>If still present, cut sutures with a pair of surgical scissors and remove the scab with a pair of forceps. Retract the skin to expose the skull. 
	NOTE: For the chronic model it may be necessary to excise the skin with #10 scalpel blade.</li>
	<li>Use a sterile Q-tip to dry the skull and locate burr holes.</li>
	<li>Set up manipulator arms of the stereotaxic apparatus with a needle holder carrying a 26 G Syringe with a 0.375&#8243; long needle.</li>
	<li>Load 3.4 &#181;L of AAV (1 x 109 Genome Copies(GC)/&#181;L for acute study; 1 x 1010 GC/&#181;L for chronic study) into the syringe. To ensure that the flow is good, release a small volume of AAV until a drop can be observed at the tip of the needle. Use a sterile Q-tip to wipe off the excess before performing an injection.</li>
	<li>Lower the tip of the needle past the skull in the first burr hole used for ET-1 (in the right sensorimotor cortex at + 0.0 anterior and 2.25 mm lateral to bregma) without puncturing the dura mater. Once aligned, lower the needle 1 mm into the brain. (0 AP, +2.25 ML, &#8722;1.0 DV from bregma).</li>
	<li>Inject 1 &#181;L of AAV (1 x 109 GC/&#181;L for acute study; 1 x 1010 GC/&#181;L for chronic study) by dispensing 0.1 &#181;L at a time. Push the plunger of the Hamilton syringe forward to dispense 0.1 &#181;L and wait for a minute before injecting the next 0.1 &#181;L; inject until 1 &#181;L has been dispensed. After the final 0.1 &#181;L has been dispensed (0.1 &#181;L x 10 injections = 1 &#181;L total), keep the needle in place for 10 min to prevent backflow and then slowly withdraw upward to remove the syringe.</li>
	<li>To ensure the needle does not become blocked during injection, release a small volume of the AAV until a drop can be observed at the tip of the needle. Use a sterile Q-tip to wipe off the excess before performing an injection.</li>
	<li>Repeat steps 2.11-2.14 for the AAV injections into the two remaining burr holes (in the right sensorimotor cortex at +1 AP, +2.25 ML, &#8722;1.0 DV; and &#8722;1 AP, +2.25 ML, &#8722;1.0 DV mm from bregma. Sterilize syringes before injecting the next animal with PREempt, followed by 70% alcohol, and finally sterile PBS.</li>
	<li>To close the wound, suture the overlying skin on the skull together using a 4-0 polysorb polyester sterile suture.</li>
	<li>Administer antibiotic ointment by using a cotton swab on the sutured area to prevent post-operative infection.</li>
	<li>Remove the mouse from the stereotaxic frame and transfer it to a clean, pre-heated animal housing cage in the recovery area. Turn off the isoflurane vaporizer and oxygen.</li>
	<li>Keep the mouse inside the cage and sterilize the outside of the cage with PREempt and 70% alcohol before placing it on top of a heating pad for recovery from anesthesia.
	<ol>
		<li>Alternatively, if using a heating lamp, place the mouse on the side of the cage under the lamp so that the mouse can move to the cool side of the cage as it wakes up from anesthesia to prevent overheating. Monitor the mouse as it comes out of anesthesia in the next 5-10 min.</li>
	</ol>
	</li>
	<li>Monitor the mouse over the next 3 days (weight monitoring), provide post-operative care (moist mashed chow to ensure hydration), and analgesics (2 mg/kg meloxicam, 2x daily subcutaneously, for 2-3 days) in accordance with regulations set out by local animal care committee.</li>
</ol>
3. Tissue preparation and dissection
<ol>
	<li>To euthanize mouse, intraperitoneally inject the mouse with Tribromoethanol (Avertin) (conc. 12.5 mg/mL, 250 mg/BW(kg)) and allow 2-3 min for complete induction. Ensure the animal is adequately anesthetized by removing the mouse from the cage and confirming the absence of a toe pinch reflex.</li>
	<li>In a fume hood, perform transcardial perfusion26 with saline followed by 4% paraformaldehyde. Spray the head of the euthanized animal with 70% ethanol until soaked.</li>
	<li>Cut off the head at the base of the skull by using a pair of large scissors. Cut open the skin along the midline of the entire skull using straight 4-1/2&#34; iris scissors. Retract the skin to expose the skull.</li>
	<li>From the opening of the spinal cavity at the neck, insert the iris scissors and cut through the skull along the sagittal and inter-frontal suture.</li>
	<li>Then insert scissors into the two eye sockets and cut through the frontal-nasal suture.</li>
	<li>Take particular care not to damage the brain. Pry open the skull plates to the left and right until the brain is exposed.</li>
	<li>Transfer the brain into a scintillation vial with 4% paraformaldehyde. Keep the brain in 4% paraformaldehyde at 4 &#176;C for 4 h.</li>
	<li>The brain is then cryoprotected by replacing 4% paraformaldehyde with 30% sucrose. Keep the brain in 30% sucrose at 4 &#176;C until the brain sinks to the bottom of the vial (~12 h).</li>
</ol>
4. Sectioning frozen brains
<ol>
	<li>Set the temperature of the cryostat chamber between -20 &#176;C to -26 &#176;C.</li>
	<li>Make a coronal cut across the cerebellum and attach the flat, caudal part of the brain onto a cryostat chuck with optimal cutting temperature compound (OCT).</li>
	<li>Place the chuck with the brain in the disk holder and align the olfactory bulb with the knife blade.</li>
	<li>Section the brain into coronal slices with a thickness of 20 &#181;m. To avoid streaks or scratch lines on sectioned brain slices, place a piece of tape on the sectioning platform to create a small gap between the anti-rolling plate and the platform. A paintbrush can be used to adjust the orientation of the section before capturing it on a positively charged glass slide.</li>
	<li>Store slides containing brain sections at -20 &#176;C.</li>
</ol>
5. Immunohistochemistry and quantification
<ol>
	<li>Process the sections for immunohistochemistry using a primary antibody against NeuN, a pan-neuronal marker, according to standard protocols22,26.</li>
	<li>Thaw cryopreserved brain sections for 5 min at room temperature. Wash the sections three times with 1x phosphate-buffered saline (PBS) for 5 min each.</li>
	<li>Incubate the sections in 0.3% Triton X-100 in PBS for 20 min at room temperature to permeabilize.</li>
	<li>Prepare the blocking solution (5% normal goat serum and 0.3% Triton X-100 in PBS) and incubate the sections in the blocking solution for 1 h at room temperature.</li>
	<li>Dilute the primary antibody, rabbit anti-NeuN, at a 1:500 ratio in the blocking solution. Incubate the sections with the primary antibody solution overnight at 4 &#176;C.</li>
	<li>Wash the sections three times with 0.3% Triton X-100 in PBS, for 5 min each wash.</li>
	<li>Dilute the secondary antibody, goat anti-rabbit 568 or 488, at a 1:1000 ratio in PBS. Incubate the sections with the secondary antibody solution for 1 h at room temperature, protected from light.</li>
	<li>Perform three washes with 0.3% Triton X-100 in PBS, for 5 min each. Incubate the sections with DAPI (1:10,000 in PBS) for 5 min at room temperature.</li>
	<li>Wash the sections three more times with PBS, for 5 min each. Mount the stained sections on a glass slide with a fluorescence mounting medium and a glass coverslip. Allow mounted slides to dry overnight.</li>
	<li>Image stained slides using a 20x objective with a confocal microscope to obtain views of Reporter+ cells around the stroke lesion in 3 different coronal sections (between the anatomical regions of the crossing of the corpus callosum anteriorly and the crossing of the anterior commissure posteriorly) for each animal. 
	NOTE: The distance of the AAV spread will vary within the hemisphere. However, based on our experience and post-tissue processing analysis of Reporter+ cells, the AAV spread generally spans approximately +3 mm to -3 mm from bregma. Reporter+ cells around the stroke lesion (as identified by the absence/sparse staining of NeuN) in the area between where the corpus callosum crosses the midline and the anterior commissure connects should be quantified.</li>
	<li>For quantification of transduced neurons, perform colocalization analysis. Manually count the number of double-positive cells (Reporter+NeuN+) and the total number of transduced (Reporter+) cells in brain slices. For results herein, &#62;3 fields of view were counted surrounding the lesion (medial, lateral, and inferior) from 3 coronal sections per mouse.</li>
</ol>

AAV Systems and Mouse Models to Study Ectopic Neurod1 Expression Post-Stroke

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B., 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=A+reproducible+endothelin-1+model+of+forelimb+motor+cortex+stroke+in+the+mouse.">A reproducible endothelin-1 model of forelimb motor cortex stroke in the mouse.</a> J Neurosci Methods. 233, 34-44 (2014).</li><li>Islam, R., 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=Inhibition+of+apoptosis+in+a+model+of+ischemic+stroke+leads+to+enhanced+cell+survival,+endogenous+neural+precursor+cell+activation+and+improved+functional+outcomes.">Inhibition of apoptosis in a model of ischemic stroke leads to enhanced cell survival, endogenous neural precursor cell activation and improved functional outcomes.</a> Int J Mol Sci. 25 (3), 1786 (2024).</li><li>Balkaya, M., Krober, J. M., Rex, A., Endres, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessing+post-stroke+behavior+in+mouse+models+of+focal+ischemia.">Assessing post-stroke behavior in mouse models of focal ischemia.</a> J Cereb Blood Flow Metab. 33 (3), 330-338 (2013).</li><li>Guo, Y., 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=High-titer+AAV+disrupts+cerebrovascular+integrity+and+induces+lymphocyte+infiltration+in+adult+mouse+brain.">High-titer AAV disrupts cerebrovascular integrity and induces lymphocyte infiltration in adult mouse brain.</a> Mol Ther Methods Clin Dev. 31, 101102 (2023).</li><li>Ortinski, P. I., 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=Selective+induction+of+astrocytic+gliosis+generates+deficits+in+neuronal+inhibition.">Selective induction of astrocytic gliosis generates deficits in neuronal inhibition.</a> Nat Neurosci. 13 (5), 584-591 (2010).</li><li>Jiang, M. Q., 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=Conversion+of+reactive+astrocytes+to+induced+neurons+enhances+neuronal+repair+and+functional+recovery+after+ischemic+stroke.">Conversion of reactive astrocytes to induced neurons enhances neuronal repair and functional recovery after ischemic stroke.</a> Front Aging Neurosci. 13, 612856 (2021).</li><li>Aschauer, D. F., Kreuz, S., Rumpel, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+transduction+efficiency,+tropism+and+axonal+transport+of+AAV+serotypes+1,+2,+5,+6,+8+and+9+in+the+mouse+brain.">Analysis of transduction efficiency, tropism and axonal transport of AAV serotypes 1, 2, 5, 6, 8 and 9 in the mouse brain.</a> PLoS One. 8 (9), e76310 (2013).</li><li>Su, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+specificity+of+GFAP+transgenes.">Expression specificity of GFAP transgenes.</a> Neurochem Res. 29 (11), 2075-2093 (2004).</li><li>Kofoed, R. H., 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=Efficacy+of+gene+delivery+to+the+brain+using+AAV+and+ultrasound+depends+on+serotypes+and+brain+areas.">Efficacy of gene delivery to the brain using AAV and ultrasound depends on serotypes and brain areas.</a> J Control Release. 351, 667-680 (2022).</li><li>He, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+influence+of+murine+genetic+background+in+adeno-associated+virus+transduction+of+the+mouse+brain.">The influence of murine genetic background in adeno-associated virus transduction of the mouse brain.</a> Hum Gene Ther Clin Dev. 30 (4), 169-181 (2019).</li><li>Hu, N. Y., 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=Expression+patterns+of+inducible+cre+recombinase+driven+by+differential+astrocyte-specific+promoters+in+transgenic+mouse+lines.">Expression patterns of inducible cre recombinase driven by differential astrocyte-specific promoters in transgenic mouse lines.</a> Neurosci Bull. 36 (5), 530-544 (2020).</li><li>Botterill, J. J., 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=Off-target+expression+of+cre-dependent+adeno-associated+viruses+in+wild-type+c57bl/6j+mice.">Off-target expression of cre-dependent adeno-associated viruses in wild-type c57bl/6j mice.</a> eNeuro. 8 (6), ENEURO.0363-ENEURO.0421 (2021).</li><li>Hasel, P., Rose, I. V. L., Sadick, J. S., Kim, R. D., Liddelow, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuroinflammatory+astrocyte+subtypes+in+the+mouse+brain.">Neuroinflammatory astrocyte subtypes in the mouse brain.</a> Nat Neurosci. 24 (10), 1475-1487 (2021).</li><li>Kim, R. D., 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=Temporal+and+spatial+analysis+of+astrocytes+following+stroke+identifies+novel+drivers+of+reactivity.">Temporal and spatial analysis of astrocytes following stroke identifies novel drivers of reactivity.</a> bioRxiv. , (2023).</li><li>Matias, I., Morgado, J., Gomes, F. C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Astrocyte+heterogeneity:+Impact+to+brain+aging+and+disease.">Astrocyte heterogeneity: Impact to brain aging and disease.</a> Front Aging Neurosci. 11, 59 (2019).</li><li>Miller, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Astrocyte+heterogeneity+in+the+adult+central+nervous+system.">Astrocyte heterogeneity in the adult central nervous system.</a> Front Cell Neurosci. 12, 401 (2018).</li><li>Early, A. N., Gorman, A. A., Van Eldik, L. J., Bachstetter, A. D., Morganti, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+advanced+age+upon+astrocyte-specific+responses+to+acute+traumatic+brain+injury+in+mice.">Effects of advanced age upon astrocyte-specific responses to acute traumatic brain injury in mice.</a> J Neuroinflammation. 17 (1), 115 (2020).</li><li>Donocoff, R. S., Teteloshvili, N., Chung, H., Shoulson, R., Creusot, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimization+of+tamoxifen-induced+cre+activity+and+its+effect+on+immune+cell+populations.">Optimization of tamoxifen-induced cre activity and its effect on immune cell populations.</a> Sci Rep. 10 (1), 15244 (2020).</li><li>Gresita, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Very+low+efficiency+of+direct+reprogramming+of+astrocytes+into+neurons+in+the+brains+of+young+and+aged+mice+after+cerebral+ischemia.">Very low efficiency of direct reprogramming of astrocytes into neurons in the brains of young and aged mice after cerebral ischemia.</a> Front Aging Neurosci. 11, 334 (2019).</li></ol>

The authors declare no competing interests.

This protocol details two AAV systems and mouse models for investigating the ectopic expression of Neurod1 in the context of a mild-moderate ET-1 cortical stroke model. A number of critical steps relating to the ET-1 stroke are important to consider for the reproducibility and consistency of the injury. Burr holes must be carefully drilled without puncturing the dura mater to prevent unintentional surgical injury. It is important to use consistent landmarks, in this case bregma, to ensure a similar region of int...

Stroke is a leading cause of disability worldwide1. A stroke happens when blood flow to the brain is disrupted. This can occur either through a hemorrhagic stroke (~15% of cases), where a blood vessel in the brain bursts, or through an ischemic stroke, where blood flow to the brain is blocked1. Ischemic strokes are most prevalent and account for ~85% of stroke cases1. A stroke reduces the delivery of glucose and oxygen to the brain, leading to swift neuronal cell death and impaired neural function.
Ischemic stroke leads to the loss of cells within minutes in the core of the lesion through mechanisms of excitotoxicity, oxidative and nitrosative stress, calcium dysregulation, cortical spreading depolarization, edema, disruption of the blood-brain barrier (BBB), and inflammation2,3. Cells in the peri-infarct regions undergo apoptotic cell death within hours, and for days, post-stroke4. The peri-infarct environment is exposed to pro- and anti-inflammatory signals that lead to the activation of microglia and astrocytes (&#34;reactive&#34; astrocytes), that play diverse roles after injury5,6,7,8. Reactive astrocytes undergo a number of phenotypic changes that include the upregulation of glial fibrillary acidic protein (GFAP), and the formation of a border around the lesion within days post-stroke that helps to limit the extent of the lesion as well as impacts neuroplasticity within the injured tissue8,9,10.
Direct cellular reprogramming allows for the conversion of one mature cell type into a different mature cell type without going through a pluripotent state11. Turning resident brain cells into new neurons to substitute those lost to injury presents an appealing avenue for brain restoration. Stroke is an ideal target for cell-based treatments, given the necessity for therapeutic methods to replace lost neurons, broaden the therapeutic timeframe, and ultimately facilitate functional recovery. To this end, there have been many papers demonstrating astrocyte to neuron (AtN) reprogramming in vitro and in vivo, using a variety of basic- helix-loop-helix (bHLH) neurogenic transcription factors, including Ascl1, Neurog2, Neurod1 and more recently, mutated versions of Ascl1 (Ascl1-SA6) which demonstrated improved reprogramming efficacy11,12,13,14. Important to consider are the many experimental design factors that can impact outcomes, interpretation, and comparisons between studies. Ghazale et al. demonstrated that AtN reprogramming efficiency varied between different mouse strains, AAV serotypes, and promoters when using the same bHLH transcription factor14. Other considerations, such as the injury model, brain region, viral dosing, timing of gene delivery, the reprogramming factor, and the mode of delivery, will impact the outcomes. Recent reports have demonstrated that pre-existing transduced neurons have been misinterpreted as reprogrammed neurons15,16. While the reports do not necessarily negate that AtN conversion occurs, these studies have highlighted the importance of well-controlled experiments, and several papers have put forth recommendations for the next steps in the field15,17,18,19. What is apparent from the literature is that the ectopic expression of bHLH transcription factors can improve functional outcomes in a variety of animal models of neurodegenerative disease/injury13,20,21,22,23,24.
Herein, this protocol Neurod1, a transcription factor of interest, is ectopically expressed in transduced cells13,22. The well-established endothelin-1 (ET-1) model of ischemic stroke to the sensorimotor cortex in adult mice is used. Injection of ET-1 leads to local vasoconstriction mimicking the pathophysiology of ischemic stroke, including functional impairment22,23,25. The commonly employed short-GFAP promoter (gfaABC(1)D) is used to drive the ectopic expression of Neurod1 at the injury site and perilesional region using two viral strategies: (1) AAV5-CAG-Flex::GFP+AAV5-GFAP-Neurod1-Cre delivery in wild-type mice and (2) AAV5-GFAP-Neurod1-Cre delivery in tdTom-Cre reporter mice. To further understand the potential for ectopic expression of Neurod1 in the stroke-injured brain, AAVs were delivered in the subacute phase (7 days), or in the chronic phase (21 days), post-stroke. In both paradigms, significantly more labeled neurons were found within weeks following ectopic Neurod1 expression.

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			<td >#77 Drill Bit (.018&#8221;)</td>
			<td >David Kopf Instruments</td>
			<td >8177</td>
			<td ></td>
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			<td >AAV5-GFAP(0.7)-mNeurod1-2A-iCre</td>
			<td >Vector&#160;Biolabs</td>
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			<td >Absorbable Suture with Needle Polysorb&#8482; Polyester CV-15 3/8 Circle Taper Point Needle Size 4 - 0 Braided</td>
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			<td >Anti-NeuN Antibody (rabbit)&#160;</td>
			<td >Millipore Sigma</td>
			<td >ABN78&#160;</td>
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			<td >C57BL/6 Mice</td>
			<td >Charles River</td>
			<td >027</td>
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			<td >Chlorhexidine Solution</td>
			<td >Partnar</td>
			<td >PCH-020</td>
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			<td >Contec&#8482;&#160;PREempt&#8482; RTU Disinfectant Solution</td>
			<td >Fisher Scientific</td>
			<td >29-636-6212</td>
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			<td >Cryostat</td>
			<td >Thermo Scientific&#160;</td>
			<td >HM525 NX&#160;</td>
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			<td >Endothelin 1</td>
			<td >Millipore Sigma</td>
			<td >05-23-3800-0.5MG</td>
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			<td >Feather Safety Razor Microtome Blades</td>
			<td >Feather</td>
			<td >12-631P</td>
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			<td >Fisherbrand&#160;Cover Glasses: Rectangles</td>
			<td >Fisher Scientific</td>
			<td >12-545M&#160;</td>
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			<td >Fluorescence Mounting Medium</td>
			<td >Agilent Technologies</td>
			<td >S3023&#160;</td>
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			<td >Hamilton Syringe</td>
			<td >Hamilton Company</td>
			<td >7634-01</td>
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			<td >High Speed Stereotaxic Drill</td>
			<td >David Kopf Instruments</td>
			<td >1474</td>
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			<td >Metacam Solution (Meloxicam)</td>
			<td >Boehringer Ingelheim</td>
			<td ></td>
			<td></td>
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			<td >O.C.T Compound</td>
			<td >Fisher Scientific</td>
			<td >23-730-571</td>
			<td></td>
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			<td >pAAV2/5-GFAP-iCre</td>
			<td >Vector Builder</td>
			<td >P190924-1001suq</td>
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			<td >Polyderm Ointment USP</td>
			<td >TARO</td>
			<td >2181908</td>
			<td></td>
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			<td >SOMNI Scientific&#8482;&#160;The Animal Temperature Heating Pad</td>
			<td >Fisher Scientific</td>
			<td >04-777-177</td>
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			<td >Stereotaxic Instruments</td>
			<td >David Kopf Instruments</td>
			<td >Model 902</td>
			<td></td>
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			<td >Superfrost Plus Microscope Slides, white</td>
			<td >Fisherbrand</td>
			<td >12-550-15</td>
			<td></td>
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			<td >tdTomato Reporter Mice&#160;</td>
			<td >The Jackson Laboratory</td>
			<td >007914</td>
			<td></td>
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			<td >V-1 Tabletop Laboratory Animal Anesthesia System</td>
			<td >VetEquip</td>
			<td >901806</td>
			<td></td>
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aav systems and mouse models for investigating ectopic expression of neurod1 in transduced cells at subacute and chronic times post-ischemic stroke

Ectopic expression of neurogenic factors in vivo has emerged as a promising approach for replacing lost neurons in disease models. The use of neural basic helix-loop-helix (bHLH) transcription factors via non-propagating virus-like particle systems, including retrovirus, lentivirus, and adeno-associated virus (AAV), has been extensively reported. For in vivo experiments, AAVs are increasingly used due to their low pathogenicity and potential for translatability. This protocol describes two AAV systems for investigating the ectopic expression of transcription factors in transduced cells post-ischemic stroke. In both systems, Neurod1 expression is controlled by the short GFAP (gfaABC(1)D) promoter, which is upregulated in reactive astrocytes post-stroke as well as in endogenous neurons when combined with neurogenic factor expression. In the ischemic stroke model described, focal ischemia is induced by injecting endothelin-1 (ET-1) into the motor cortex of mice, creating a lesion surrounded by reactive GFAP-expressing astrocytes and surviving neurons. Intracerebral injections of AAV are performed to ectopically induce the expression of Neurod1 in the subacute (7 days) and chronic (21 days) phases post-stroke. Within weeks following AAV injection, a significantly higher number of neurons among transduced cells are identified in mice ectopically expressing Neurod1 compared to mice receiving AAV control viruses. The AAV-based strategies used replicated observed outcomes of increased numbers of neurons expressing the reporter gene in a model of mild-to-moderate cortical stroke. This protocol establishes a standard platform for exploring the effects of ectopic expression of transcription factors delivered with AAV-based systems, contributing to the understanding of neurogenic factor expression in the context of stroke.

To examine the cellular outcomes of short GFAP (gfaABC(1)D) promoter driven, ectopic Neurod1 expression in wild-type (C57BL/6J) and in transgenic tdTom-Cre reporter strains (specifically, B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J)22 two AAV5-based systems were utilized in ET-1 stroke injured mice (Figure 1A-C). In a subacute model of stroke, Neurod1 was&#160;packaged in the Cre-Flex AAV system and deliver...

AAV Systems and Mouse Models for Investigating Ectopic Expression of Neurod1 in Transduced Cells at Subacute and Chronic Times Post-Ischemic Stroke

This protocol describes the viral-mediated ectopic expression of Neurod1 following cortical ischemic stroke. Neurod1 is delivered (1) using the Cre-Flex AAV system in wild-type mice during the subacute phase post-stroke (7 days) and (2) using a single AAV vector in conditional reporter mice during the chronic phase post-stroke (21 days).

Ectopic expression of neurogenic factors in vivo has emerged as a promising approach for replacing lost neurons in disease models. The use of neural basic ...

aav-systems-mouse-models-for-investigating-ectopic-expression-neurod1

Research

JoVE Journal

Neuroscience

344 Views.  University of Toronto. This protocol describes the viral-mediated ectopic expression of Neurod1 following cortical ischemic stroke. Neurod1 is delivered (1) using the Cre-Flex AAV system in wild-type mice during the subacute phase post-stroke (7 days) and (2) using a single AAV vector in conditional reporter mice during the chronic phase post-stroke (21 days).

Video: AAV Systems and Mouse Models for Investigating Ectopic Expression of Neurod1 in Transduced Cells at Subacute and Chronic Times Post-Ischemic Stroke

Mouse Models of Periventricular Leukomalacia

We describe a protocol for establishing mouse models of periventricular leukomalacia (PVL). PVL is the predominant form of brain injury in premature infants and the most common antecedent of cerebral palsy. PVL is characterized by periventricular white matter damage with prominent oligodendroglial injury. Hypoxia/ischemia with or without systemic infection/inflammation are the primary causes of PVL. We use P6 mice to create models of neonatal brain injury by the induction of hypoxia/ischemia with or without systemic infection/inflammation with unilateral carotid ligation followed by exposure to hypoxia with or without injection of the endotoxin lipopolysaccharide (LPS). Immunohistochemistry of myelin basic protein (MBP) or O1 and electron microscopic examination show prominent myelin loss in cerebral white matter with additional damage to the hippocampus and thalamus. Establishment of mouse models of PVL will greatly facilitate the study of disease pathogenesis using available transgenic mouse strains, conduction of drug trials in a relatively high throughput manner to identify candidate therapeutic agents, and testing of stem cell transplantation using immunodeficiency mouse strains.

We established mouse models of periventricular leukomalacia (PVL), the predominant brain injury in premature infants characterized by periventricular white matter lesions. Hypoxia/ischemia with/without systemic infection are the primary causes of PVL. Unilateral carotid ligation and hypoxia exposure with/without lipopolysaccharide injection creates PVL-like lesions in P6 mice.

We describe a protocol for establishing mouse models of periventricular leukomalacia (PVL). PVL is the predominant form of brain injury in premature ...

Combining Behavioral Endocrinology and Experimental Economics: Testosterone and Social Decision Making

Behavioral endocrinological research in humans as well as in animals suggests that testosterone plays a key role in social interactions. Studies in rodents have shown a direct link between testosterone and aggressive behavior1 and folk wisdom adapts these findings to humans, suggesting that testosterone induces antisocial, egoistic or even aggressive behavior2. However, many researchers doubt a direct testosterone-aggression link in humans, arguing instead that testosterone is primarily involved in status-related behavior3,4. As a high status can also be achieved by aggressive and antisocial means it can be difficult to distinguish between anti-social and status seeking behavior.
We therefore set up an experimental environment, in which status can only be achieved by prosocial means. In a double-blind and placebo-controlled experiment, we administered a single sublingual dose of 0.5 mg of testosterone (with a hydroxypropyl-&beta;-cyclodextrin carrier) to 121 women and investigated their social interaction behavior in an economic bargaining paradigm. Real monetary incentives are at stake in this paradigm; every player A receives a certain amount of money and has to make an offer to another player B on how to share the money. If B accepts, she gets what was offered and player A keeps the rest. If B refuses the offer, nobody gets anything. A status seeking player A is expected to avoid being rejected by behaving in a prosocial way, i.e. by making higher offers.
The results show that if expectations about the hormone are controlled for, testosterone administration leads to a significant increase in fair bargaining offers compared to placebo. The role of expectations is reflected in the fact that subjects who report that they believe to have received testosterone make lower offers than those who say they believe that they were treated with a placebo. These findings suggest that the experimental economics approach is sensitive for detecting neurobiological effects as subtle as those achieved by administration of hormones. Moreover, the findings point towards the importance of both psychosocial as well as neuroendocrine factors in determining the influence of testosterone on human social behavior.

The procedure described in this protocol shows that testosterone administration and folk beliefs about testosterone may be associated with directly opposed social behaviors.

Behavioral endocrinological research in humans as well as in animals suggests that testosterone plays a key role in social interactions. Studies in ...

Sequential Photo-bleaching to Delineate Single Schwann Cells at the Neuromuscular Junction

Sequential photo-bleaching provides a non-invasive way to label individual SCs at the NMJ. The NMJ is the largest synapse of the mammalian nervous system and has served as guiding model to study synaptic structure and function. In mouse NMJs motor axon terminals form pretzel-like contact sites with muscle fibers. The motor axon and its terminal are sheathed by SCs. Over the past decades, several transgenic mice have been generated to visualize motor neurons and SCs, for example Thy1-XFP1 and Plp-GFP mice2, respectively.
Along motor axons, myelinating axonal SCs are arranged in non-overlapping internodes, separated by nodes of Ranvier, to enable saltatory action potential propagation. In contrast, terminal SCs at the synapse are specialized glial cells, which monitor and promote neurotransmission, digest debris and guide regenerating axons. NMJs are tightly covered by up to half a dozen non-myelinating terminal SCs - these, however, cannot be individually resolved by light microscopy, as they are in direct membrane contact3.
Several approaches exist to individually visualize terminal SCs. None of these are flawless, though. For instance, dye filling, where single cells are impaled with a dye-filled microelectrode, requires destroying a labelled cell before filling a second one. This is not compatible with subsequent time-lapse recordings3. Multi-spectral "Brainbow" labeling of SCs has been achieved by using combinatorial expression of fluorescent proteins4. However, this technique requires combining several transgenes and is limited by the expression pattern of the promoters used. In the future, expression of "photo-switchable" proteins in SCs might be yet another alternative5. Here we present sequential photo-bleaching, where single cells are bleached, and their image obtained by subtraction. We believe that this approach - due to its ease and versatility - represents a lasting addition to the neuroscientist's technology palette, especially as it can be used in vivo and transferred to others cell types, anatomical sites or species6.
In the following protocol, we detail the application of sequential bleaching and subsequent confocal time-lapse microscopy to terminal SCs in triangularis sterni muscle explants. This thin, superficial and easily dissected nerve-muscle preparation7,8 has proven useful for studies of NMJ development, physiology and pathology9. Finally, we explain how the triangularis sterni muscle is prepared after fixation to perform correlated high-resolution confocal imaging, immunohistochemistry or ultrastructural examinations.

Visualizing individual cells in densely packed tissues, such as terminal Schwann cells (SCs) at neuromuscular junctions (NMJs), is challenging. "Sequential photo-bleaching" allows delineating single terminal SCs, for instance in the triangularis sterni muscle explant, a convenient nerve-muscle preparation, where sequential bleaching can be combined with time-lapse imaging and post-hoc immunostainings.

Sequential photo-bleaching provides a non-invasive way to label individual SCs at the NMJ. The NMJ is the largest synapse of the mammalian nervous system ...

Isolation and Culture of Mouse Cortical Astrocytes

Astrocytes are an abundant cell type in the mammalian brain, yet much remains to be learned about their molecular and functional characteristics. In vitro astrocyte cell culture systems can be used to study the biological functions of these glial cells in detail. This video protocol shows how to obtain pure astrocytes by isolation and culture of mixed cortical cells of mouse pups. The method is based on the absence of viable neurons and the separation of astrocytes, oligodendrocytes and microglia, the three main glial cell populations of the central nervous system, in culture. Representative images during the first days of culture demonstrate the presence of a mixed cell population and indicate the timepoint, when astrocytes become confluent and should be separated from microglia and oligodendrocytes. Moreover, we demonstrate purity and astrocytic morphology of cultured astrocytes using immunocytochemical stainings for well established and newly described astrocyte markers. This culture system can be easily used to obtain pure mouse astrocytes and astrocyte-conditioned medium for studying various aspects of astrocyte biology.

Astrocytes have been recognized to be versatile cells participating in fundamental biological processes that are essential for normal brain development and function, and central nervous system repair. Here we present a rapid procedure to obtain pure mouse astrocyte cultures to study the biology of this major class of central nervous system cells.

Astrocytes  are an abundant cell type in the mammalian brain, yet much remains to be  learned about their molecular and functional characteristics. In ...

Lateral Chronic Cranial Window Preparation Enables In Vivo Observation Following Distal Middle Cerebral Artery Occlusion in Mice

Focal cerebral ischemia (i.e., ischemic stroke) may cause major brain injury, leading to a severe loss of neuronal function and consequently to a host of motor and cognitive disabilities. Its high prevalence poses a serious health burden, as stroke is among the principal causes of long-term disability and death worldwide1. Recovery of neuronal function is, in most cases, only partial. So far, treatment options are very limited, in particular due to the narrow time window for thrombolysis2,3. Determining methods to accelerate recovery from stroke remains a prime medical goal; however, this has been hampered by insufficient mechanistic insights into the recovery process. Experimental stroke researchers frequently employ rodent models of focal cerebral ischemia. Beyond the acute phase, stroke research is increasingly focused on the sub-acute and chronic phase following cerebral ischemia. Most stroke researchers apply permanent or transient occlusion of the MCA in mice or rats. In patients, occlusions of the MCA are among the most frequent causes of ischemic stroke4. Besides proximal occlusion of the MCA using the filament model, surgical occlusion of the distal MCA is probably the most frequently used model in experimental stroke research5. Occlusion of a distal (to the branching of the lenticulo-striate arteries) MCA branch typically spares the striatum and primarily affects the neocortex. Vessel occlusion can be permanent or transient. High reproducibility of lesion volume and very low mortality rates with respect to the long-term outcome are the main advantages of this model. Here, we demonstrate how to perform a chronic cranial window (CW) preparation lateral to the sagittal sinus, and afterwards how to surgically induce a distal stroke underneath the window using a craniotomy approach. This approach can be applied for sequential imaging of acute and chronic changes following ischemia via epi-illuminating, confocal laser scanning, and two-photon intravital microscopy.

Lateral Chronic Cranial Window Preparation Enables In Vivo Observation Following Distal Middle Cerebral Artery Occlusion in Mice

Surgical occlusion of a distal middle cerebral artery branch (MCAo) is a frequently used model in experimental stroke research. This manuscript describes the basic technique of permanent MCAo, combined with the insertion of a lateral cranial window, which offers the opportunity for longitudinal intravital microscopy in mice.

Focal cerebral ischemia (i.e., ischemic stroke) may cause major brain injury, leading to a severe loss of neuronal function and consequently to a host of ...

Evaluation of Bioenergetic Function in Cerebral Vascular Endothelial Cells

The integrity of the blood-brain-barrier (BBB) is critical to prevent brain injury. Cerebral vascular endothelial (CVE) cells are one of the cell types that comprise the BBB; these cells have a very high-energy demand, which requires optimal mitochondrial function. In the case of disease or injury, the mitochondrial function in these cells can be altered, resulting in disease or&#160;the opening of the BBB. In this manuscript, we introduce a method to measure mitochondrial function in CVE cells by using whole, intact cells and a bioanalyzer. A mito-stress assay is used to challenge the cells that have been perturbed, either physically or chemically, and evaluate their bioenergetic function. Additionally, this method also provides a useful way to screen new therapeutics that have direct effects on mitochondrial function. We have optimized the cell density necessary to yield oxygen consumption rates that allow for the calculation of a variety of mitochondrial parameters, including ATP production, maximal respiration, and spare capacity. We also show the sensitivity of the assay by demonstrating that the introduction of the&#160;microRNA, miR-34a, leads to a pronounced and detectable decrease in mitochondrial activity. While the data shown in this paper is optimized for the bEnd.3 cell line, we have also optimized the protocol for primary CVE cells, further suggesting the utility in preclinical and clinical models.

Endothelial cell mitochondria are critical to maintain blood-brain-barrier integrity. We introduce a protocol to measure bioenergetic function in cerebral vascular endothelial cells.

The integrity of the blood-brain-barrier (BBB) is critical to prevent brain injury. Cerebral vascular endothelial (CVE) cells are one of the cell types ...

In Vivo Targeted Expression of Optogenetic Proteins Using Silk/AAV Films

The quest to understand how neural circuits process information in order to drive behavioral output has been greatly aided by recently-developed optical methods for manipulating and monitoring the activity of neurons in vivo. These types of experiments rely on two main components: 1) implantable devices that provide optical access to the brain, and 2) light-sensitive proteins that change neuronal excitability or provide a readout of neuronal activity. There are a number of ways to express light-sensitive proteins, but stereotaxic injection of viral vectors is currently the most flexible approach because expression can be controlled with genetic, anatomical, and temporal precision. Despite the great utility of viral vectors, delivering the virus to the site of optical implants poses numerous challenges. Stereotaxic virus injections are demanding surgeries that increase surgical time, increase the cost of studies, and pose a risk to the animal's health. The surrounding tissue can be physically damaged by the injection syringe, and by immunogenic inflammation caused by the abrupt delivery of a bolus of high-titer virus. Aligning injections with optical implants is especially difficult when targeting small regions deep in the brain. To overcome these challenges, we describe a method for coating multiple types of optical implants with films composed of silk fibroin and Adeno-associated viral (AAV) vectors. Fibroin, a polymer derived from the cocoon of Bombyx mori, can encapsulate and protect biomolecules and can be processed into forms ranging from soluble films to ceramics. When implanted into the brain, silk/AAV coatings release virus at the interface between optical elements and the surrounding brain, driving expression precisely where it is needed. This method is easily implemented and promises to greatly facilitate in vivo studies of neural circuit function.

In Vivo Targeted Expression of Optogenetic Proteins Using Silk/AAV Films

Here, we present a method for delivering viral expression vectors into the brain using silk fibroin films. This method allows targeted delivery of expression vectors using silk/AAV coated optical fibers, tapered optical fibers, and cranial windows.

The quest to understand how neural circuits process information in order to drive behavioral output has been greatly aided by recently-developed optical ...

Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. Several recent studies have demonstrated the effectiveness of chronic VNS in enhancing central nervous system plasticity to improve motor rehabilitation, extinction learning, and sensory discrimination. Construction of chronically implantable devices for use in such studies is challenging due to rats&#8217; small size, and typical protocols require extensive training of personnel and time-consuming microfabrication methods. Alternatively, commercially available implantable cuff electrodes can be purchased at a significantly higher cost. In this protocol, we present a simple, low-cost method for construction of small, chronically implantable peripheral nerve cuff electrodes for use in rats. We validate the short and long-term reliability of our cuff electrodes by demonstrating that VNS in ketamine/xylazine anesthetized rats produces decreases in breathing rate consistent with activation of the Hering-Breuer reflex, both at the time of implantation and up to 10 weeks after device implantation. We further demonstrate the suitability of the cuff electrodes for use in chronic stimulation studies by pairing VNS with skilled lever press performance to induce motor cortical map plasticity.

Existing approaches for constructing chronically implantable peripheral nerve cuff electrodes for use in small rodents often require specialized equipment and/or highly trained personnel. In this protocol we demonstrate a simple, low-cost approach for fabricating chronically implantable cuff electrodes, and demonstrate their effectiveness for vagus nerve stimulation (VNS) in rats.

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. ...

Modeling Stroke in Mice: Focal Cortical Lesions by Photothrombosis

Stroke is a leading cause of death and acquired adult disability in developed countries. Despite extensive investigation for novel therapeutic strategies, there remain limited therapeutic options for stroke patients. Therefore, more research is needed for pathophysiological pathways such as post-stroke inflammation, angiogenesis, neuronal plasticity, and regeneration. Given the inability of in vitro models to reproduce the complexity of the brain, experimental stroke models are essential for the analysis and subsequent evaluation of novel drug targets for these mechanisms. In addition, detailed standardized models for all procedures are urgently needed to overcome the so-called replication crisis. As an effort within the ImmunoStroke research consortium, a standardized photothrombotic mouse model using an intraperitoneal injection of Rose Bengal and the illumination of the intact skull with a 561 nm laser is described. This model allows the performance of stroke in mice with allocation to any cortical region of the brain without invasive surgery; thus, enabling the study of stroke in various areas of the brain. In this video, the surgical methods of stroke induction in the photothrombotic model along with&#160;histological analysis are demonstrated.

Described here is the photothrombotic stroke model, where a stroke is produced through the intact skull by inducing permanent microvascular occlusion using laser illumination after administration of a photosensitive dye.

Stroke is a leading cause of death and acquired adult disability in developed countries. Despite extensive investigation for novel therapeutic strategies, ...

Modeling Stroke in Mice: Transient Middle Cerebral Artery Occlusion via the External Carotid Artery

Stroke is the third most common cause of mortality and the leading cause of acquired adult disability in developed countries. To date, therapeutic options are limited to a small proportion of stroke patients within the first hours after stroke. Novel therapeutic strategies are being extensively investigated, especially to prolong the therapeutic time window. These current investigations include the study of important pathophysiological pathways after stroke, such as post-stroke inflammation, angiogenesis, neuronal plasticity, and regeneration. Over the last decade, there has been increasing concern about the poor reproducibility of experimental results and scientific findings among independent research groups. To overcome the so-called &#8220;replication crisis&#8221;, detailed standardized models for all procedures are urgently needed. As an effort within the &#8220;ImmunoStroke&#8221; research consortium (https://immunostroke.de/), a standardized mouse model of transient middle cerebral artery occlusion (MCAo) is proposed. This model allows the complete restoration of the blood flow upon removal of the filament, simulating the therapeutic or spontaneous clot lysis that occurs in a large proportion of human strokes. The surgical procedure of this &#8220;filament&#8221; stroke model and tools for its functional analysis are demonstrated in the accompanying video.

Different models of middle cerebral artery occlusion (MCAo) are used in experimental stroke research. Here, an experimental stroke model of transient MCAo via the external carotid artery (ECA) is described, which aims to mimic human stroke, in which the cerebrovascular thrombus is removed due to spontaneous clot lysis or therapy.

Stroke is the third most common cause of mortality and the leading cause of acquired adult disability in developed countries. To date, therapeutic options ...