This study was supported by grants from NIH NINDS R01NS102836 and the Pennsylvania Department of Health Commonwealth Universal Research Enhancement (CURE) awarded to Seena K. Ajit. We thank Dr. Bradley Nash for critical reading of the manuscript.

NOTE: All procedures were performed in compliance with the NIH Guide for the Care and Use of Laboratory Animals and approved by the Institutional Animal Care &#38; Use Committee of Drexel University College of Medicine. Timed-pregnant CD-1 mice were used for astrocytic culture, and all dams were received 15 days after impregnation. Ten-twelve weeks old C57BL/6 mice were used for in vivo uptake experiments.
1. Isolation of sEVs from RAW 264.7 macrophage cells
<ol>
	<li>Culture RAW 264.7 cells in 75 cm2 flasks in DMEM exosome-depleted medium containing 10% exosome-depleted fetal bovine serum (FBS) and 1% penicillin-streptomycin (pen-strep) for 24-48 h.</li>
	<li>Collect 300 mL of conditioned medium and centrifuge at 300 &#215; g for 10 min at 4 &#176;C.</li>
	<li>Collect the supernatant and centrifuge at 2,000 &#215; g for 20 min at 4 &#176;C.</li>
	<li>Transfer the supernatant to centrifuge tubes, centrifuge for 35 min at 12,000 &#215; g at 4 &#176;C.</li>
	<li>Collect the supernatant and filter through a 0.22 &#181;m syringe filter.</li>
	<li>Transfer to ultracentrifuge tubes and centrifuge for 80 min at 110,000 &#215; g at 4 &#176;C.</li>
	<li>Store the supernatant (exosome-depleted medium), resuspend the pellet in 2 mL of 1x phosphate-buffered saline (PBS), and centrifuge for 1 h at 110,000 &#215; g at 4 &#176;C.</li>
	<li>Resuspend the pellet in 100 &#181;L of PBS for further characterization using nanoparticle tracking analysis (NTA) and transmission electron microscopy (TEM) or in radioimmunoprecipitation assay (RIPA) buffer for western blotting.</li>
</ol>
2. Characterization of sEVs
<ol>
	<li>Nanoparticle tracking analysis (NTA) 
	NOTE: The size distribution and particle number/concentration of the purified sEVs from RAW 264.7 cells were measured by NTA.
	<ol>
		<li>Dilute the sEVs in filtered PBS to obtain 20-60 vesicles per field of view for optimal tracking.</li>
		<li>Introduce the diluted sample into a flow cell using a syringe pump with a constant flow rate.</li>
		<li>Take 3-5 videos of 30 s each. Set the shutter speed and gain, and manually focus the camera settings for the maximum number of vesicles to be visible and capable of being tracked and analyzed.</li>
		<li>Advance the samples between each recording to perform replicate measurements. Optimize the NTA post-acquisition settings and keep the settings constant between the samples.</li>
		<li>Analyze each video using the NTA software to obtain the average size and concentration of the vesicles.</li>
		<li>Carry out all NTA measurements with identical system settings for consistency.</li>
	</ol>
	</li>
	<li>Western blot
	<ol>
		<li>Quantify the total protein amounts in sEVs, cell lysates, and exosome-depleted media using a protein assay kit following the manufacturer&#39;s instructions.</li>
		<li>For cell lysate preparation, culture the RAW 264.7 cells in 75 cm2 flasks until 80-90% confluent. Detach the cells with 0.25% trypsin for 10-15 min, neutralize the trypsin with culture media, and pellet the cells by spinning at 400 &#215; g for 5 min. Resuspend the cells in fresh growth medium.</li>
		<li>Count the cells using a hemocytometer and transfer 1 &#215; 106 cells to another tube. Wash the cells with PBS twice using the same centrifugation conditions as above and add 50 &#181;L of lysis buffer (RIPA buffer with protease inhibitor cocktail added) to the cell pellet from the final spin.</li>
		<li>Vortex the cells and keep them on ice for 20 min. Subject the mixture to centrifugation at 10,000 &#215; g for 30 min at 4 &#176;C, collect the supernatant (i.e., the lysate) in fresh microcentrifuge tubes, and keep at &#8722;80 &#176;C until use.</li>
		<li>Concentrate 2 mL of exosome-depleted media to 100 &#181;L using 3 kDa-cutoff centrifugal filters before quantifying the amount of protein. Mix the sEVs with lysis buffer in a 1:1 ratio, vortex for 30 s, and incubate on ice for 15 min to quantify the amount of protein.</li>
		<li>Mix equal amounts of protein (2 &#181;g) of the sEVs, RAW 264.7 cell lysate, and exosome-depleted media with reducing sample buffer.</li>
		<li>Denature the samples at 95 &#176;C for 5 min, keep them on ice for 5 min, and spin for 2 min at 10,000 &#215; g. Load the samples on a 12% Tris-glycine protein gel&#160;and run the gel at 125 V for 45 min.</li>
		<li>Transfer the protein onto a polyvinylidene difluoride (PVDF) membrane at 25 V for 2 h.</li>
		<li>Following the transfer, block the PVDF membranes with blocking buffer (see the Table of Materials) for 1 h at room temperature.</li>
		<li>Incubate the blot with primary antibodies on a shaker overnight at 4 &#176;C. 
		NOTE: Primary antibodies used were anti-CD81 (1:1,000), anti-alpha-1,3/1,6-mannosyltransferase (ALG-2)-interacting protein X (Alix) (1:1,000), anti-Calnexin (1:1,000), and anti-glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (1:1,000).</li>
		<li>Wash the blots 3 x 15 min with 1x Tris-buffered saline, 0.1% Tween 20 (TBST), and incubate at room temperature with goat anti-mouse IgG-horseradish peroxidase (HRP)- or donkey anti-rabbit IgG-HRP-conjugated secondary antibodies (1:10,000) for 1 h on the shaker.</li>
		<li>Wash the blots 3 x 15 min with 1x TBST, and detect the proteins using an HRP substrate.</li>
		<li>Analyze the blots by enhanced chemiluminescence using a western blot imager.</li>
	</ol>
	</li>
	<li>Transmission electron microscopy (TEM)
	<ol>
		<li>Fix sEVs by resuspending them in 2% paraformaldehyde (PFA) in 0.1 M phosphate buffer (PB); vortex for 2 x 15 s.</li>
		<li>Place a drop of 10 &#181;L of sEV suspension on clean parafilm. Float the carbon-coated formvar grid on the drop with their coated side facing the suspension. Let the membranes absorb for 20 min in a dry environment.</li>
		<li>Place the grids (membrane side down) on a drop of PB to wash for 3 x 2 min.</li>
		<li>Transfer the grids to 50 &#181;L of 1% glutaraldehyde for 5 min.</li>
		<li>Wash the grids with 100 &#181;L of distilled water for 8 x 2 min.</li>
		<li>Contrast the sample by placing the grids on a drop of 1% uranyl acetate for 2 min.</li>
		<li>Embed the sample with 50 &#181;L of 0.2% uranyl acetate with 2% methylcellulose solution for 10 min on a parafilm-covered ice dish.</li>
		<li>Use stainless steel loops to hold the grids and remove excess fluid with filter paper.</li>
		<li>Air-dry the grid for 10 min while still on the loop.</li>
		<li>Observe under a transmission electron microscope at 80 kV.</li>
	</ol>
	</li>
</ol>
3. Labeling of sEVs
<ol>
	<li>Dilute 20 &#181;g of sEVs in 1 mL of diluent buffer or the same volume of PBS in 1 mL of diluent buffer for dye control.</li>
	<li>Dilute 3 &#181;L of PKH67 or PKH26 dye in 1 mL of diluent buffer and mix by pipetting.</li>
	<li>Add diluted PKH dye to the diluted sEVs and mix by pipetting. Incubate for 5 min in the dark at room temperature. For a dye control, mix the diluted dye with diluted PBS from step 3.1.</li>
	<li>Add 2 mL of 1% bovine serum albumin (BSA) in PBS to the tube with the dye and the sEV mix and to the dye control tube to absorb excess dye.</li>
	<li>Centrifuge for 1 h at 110,000 &#215; g at 4 &#176;C. Discard the supernatant, resuspend the pellet in 2 mL of PBS, and centrifuge for 1 h at 110,000 &#215; g at 4 &#176;C. Repeat the wash with PBS and resuspend the labeled sEVs or the dye control in an equal volume of PBS.</li>
	<li>Quantify the amount of total protein by the Bradford method.</li>
</ol>
4. Uptake of sEVs by Neuro-2a cells
<ol>
	<li>Place 18-mm coverslips in a 12-well plate and plate 10 &#215; 104 Neuro-2a cells in each well in a total of 1 mL of complete DMEM medium containing 10% FBS and 1% pen-strep.</li>
	<li>Change the medium to DMEM exosome-depleted medium when the cell confluency is 80-90%. Add 1, 5, or 10 &#181;g of labeled sEVs in each well for 1, 4, and 24 h for dose- and time-dependent uptake, or add an equal volume of dye control.</li>
</ol>
5. Primary astrocytic cultures
<ol>
	<li>Anesthetize 4 postnatal pups 4 days after birth by inducing hypothermia.</li>
	<li>Collect the brains in a 60-mm Petri dish containing ice-cold Hank&#39;s Balanced Salt Solution (HBSS) supplemented with 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES).</li>
	<li>Dissect both cortical lobes and remove the meninges. Mince the tissues with a sterilized blade.</li>
	<li>Transfer the tissues to a 15 mL conical tube containing papain/deoxyribonuclease I dissociation buffer and incubate for 20 min at 37 &#176;C. Swirl every 5 min. 
	NOTE: For 4 mouse cortices, 9 mL of 7.5 U/mL papain in HBSS is activated at 37 &#176;C for at least 30 min, filtered through a 0.22 &#181;m syringe filter, and mixed with deoxyribonuclease I to a final concentration of 0.1 mg/mL.</li>
	<li>Aspirate the supernatant and add 5 mL of complete DMEM to inactivate the enzyme activity. Carefully triturate to dissociate the tissues with a 5 mL glass serological pipette and a flame-polished Pasteur pipette.</li>
	<li>Pass the cell suspension through a 40 &#181;m cell strainer and centrifuge the cells at 250 &#215; g for 5 min at 4 &#176;C. Aspirate the medium and seed the cells in 10 mL of complete DMEM in a 75 cm2 flask. Replace the supernatant medium with 15 mL of fresh DMEM medium 4 h after plating.</li>
	<li>After 14 days in vitro, transfer the flask to an orbital shaker to detach the microglia and oligodendrocytes at 320 rpm for 6 h.</li>
	<li>Trypsinize the remaining astrocytes using 5 mL of the cell dissociation enzyme (Table of Materials) for 10 min at 37 &#176;C. Add 5 mL of complete DMEM to inactivate the enzymatic action and pellet the cells at 250 &#215; g for 5 min at 4 &#176;C.</li>
	<li>Resuspend the cells in complete DMEM. Seed 5 &#215; 104 cells on 12 mm #1.5 coverslips in a 24-well plate.</li>
</ol>
6. Uptake of sEVs by astrocytes
<ol>
	<li>When astrocytes reach 80-90% confluency, change the medium to DMEM exosome-depleted medium.</li>
	<li>Add 1 &#181;g of unlabeled, labeled sEVs, an equal volume of dye control, or PBS to the cells. Use the cells for staining 1 h and 24 h after sEV treatment.</li>
</ol>
7. Immunofluorescence
<ol>
	<li>Rinse the cells with PBS 3x and fix them with 4% PFA in PB for 10 min at room temperature.</li>
	<li>Wash the fixed cells 3 x 5 min with PB and permeabilize them using 0.1% Triton X-100 in PB for 10-15 min and wash with PB 3 x 5 min.</li>
	<li>Block the cells with 5% normal goat serum (NGS) in PB for 1 h at room temperature.</li>
	<li>Incubate the cells with primary antibodies: microtubule-associated protein 2 (MAP2A, 1:500) for Neuro-2a cells or glial fibrillary acidic protein (GFAP, 1:500) for primary astrocytes in fresh 5% NGS/PB overnight at 4 &#176;C with gentle shaking.</li>
	<li>Wash 3 x 10 min with PB and add fluorophore-conjugated secondary antibodies (Goat Anti-Mouse IgG1, Alexa Fluor 594; or Goat Anti-Mouse IgG H&#38;L, Alexa Fluor 488) in 5% NGS and incubate for 2 h at room temperature on a rocker.</li>
	<li>Wash 3 x 10 min with PB and incubate with 1 &#181;g/mL of nuclear stain 4&#39;,6-diamidino-2-phenylindole (DAPI) for 10 min at room temperature. Wash the cells again 3x with PB.</li>
	<li>Mount the coverslips on #1 slides using an antifade mounting medium. Let them dry overnight in the dark, and store the prepared glass slides at 4 &#176;C until imaging on a confocal microscope.</li>
</ol>
8. In vivo uptake of sEVs
<ol>
	<li>Perform intrathecal injection of 5 &#181;g of unlabeled or labeled sEVs resuspended in 10 &#181;L of PBS, or equal volume (10 &#181;L) of dye control (as prepared in section 3) into C57BL/6 mice.</li>
	<li>After 6 and 18 h post-injection of sEVs, deeply anesthetize mice by intraperitoneal injection of 100 mg/kg body weight of ketamine and 10 mg/kg body weight of xylazine.</li>
	<li>Perform intracardial perfusion of mice with 0.9% saline to flush out blood, followed by freshly made ice-cold 4% PFA/PB.</li>
	<li>Dissect the spinal cord and fix in 4% PFA/PB at 4 &#176;C for 24 h. Cryoprotect the tissues in 30% sucrose in PB at 4 &#176;C for 24 h or until the tissues sink. Store the tissues at 4 &#176;C until immunohistochemistry.</li>
</ol>
9. Immunohistochemistry
<ol>
	<li>Embed L4-L5 spinal cord in O.C.T compound. Freeze on dry ice until completely solidified.</li>
	<li>Section the tissues at 30 &#181;m (cross-sectionally for the spinal cord) using a cryostat, and collect the sections in a 24-well plate containing PB. Wash the sections 3 x 5 min with 0.3% Triton in PB.</li>
	<li>Block non-specific binding sites with 5% NGS in 0.3% Triton/PB for 2 h at room temperature.</li>
	<li>Dilute primary antibodies: Anti-MAP2A (1:500), GFAP (1:1,000), Iba1 for microglia (1:2,000) with 5% NGS in 0.3% Triton/PB, and incubate the sections overnight at 4 &#176;C on a shaker.</li>
	<li>Wash the sections 3 x 5 min with 0.3% Triton/PB, and add secondary antibodies (Donkey Anti-Rabbit IgG Alexa Fluor 488, 1:500, or Goat Anti-Mouse IgG Alexa Fluor 488, 1:500) in 5% NGS/PB for 2 h at room temperature.</li>
	<li>Wash 3 x 5 min with PB, and incubate the sections in 1 &#181;g/mL of DAPI for 10 min at room temperature. Wash the sections 3 x 5 min with PB.</li>
	<li>Mount the sections on a clean adhesive slide (Table of Materials) with a fine paintbrush under a light microscope.</li>
	<li>Wet the coverslip with mounting medium. Cure overnight in the dark at room temperature.</li>
	<li>Image under a confocal microscope with the respective lasers.</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Confounding+factors+in+vesicle+uptake+studies+using+fluorescent+lipophilic+membrane+dyes.">Confounding factors in vesicle uptake studies using fluorescent lipophilic membrane dyes.</a> Journal of Extracellular Vesicles. 6 (1), 1388731 (2017).</li><li>Pu&#382;ar Dominku&#353;, P., 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=PKH26+labeling+of+extracellular+vesicles:+Characterization+and+cellular+internalization+of+contaminating+PKH26+nanoparticles.">PKH26 labeling of extracellular vesicles: Characterization and cellular internalization of contaminating PKH26 nanoparticles.</a> Biochimica et Biophysica Acta. Biomembranes. 1860 (6), 1350-1361 (2018).</li><li>Dehghani, M., Gulvin, S. M., Flax, J., Gaborski, T. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Systematic+evaluation+of+PKH+labelling+on+extracellular+vesicle+size+by+nanoparticle+tracking+analysis.">Systematic evaluation of PKH labelling on extracellular vesicle size by nanoparticle tracking analysis.</a> Scientific Reports. 10 (1), 9533 (2020).</li><li>Shimomura, 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=New+lipophilic+fluorescent+dyes+for+labeling+extracellular+vesicles:+characterization+and+monitoring+of+cellular+uptake.">New lipophilic fluorescent dyes for labeling extracellular vesicles: characterization and monitoring of cellular uptake.</a> Bioconjugate Chemistry. 32 (4), 680-684 (2021).</li><li>Yuan, 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=Macrophage+exosomes+as+natural+nanocarriers+for+protein+delivery+to+inflamed+brain.">Macrophage exosomes as natural nanocarriers for protein delivery to inflamed brain.</a> Biomaterials. 142, 1-12 (2017).</li><li>Polanco, J. C., Li, C., Durisic, N., Sullivan, R., G&#246;tz, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exosomes+taken+up+by+neurons+hijack+the+endosomal+pathway+to+spread+to+interconnected+neurons.">Exosomes taken up by neurons hijack the endosomal pathway to spread to interconnected neurons.</a> Acta Neuropathologica Communications. 6 (1), 10 (2018).</li><li>Jurgielewicz, B. J., Yao, Y., Stice, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinetics+and+specificity+of+HEK293T+extracellular+vesicle+uptake+using+imaging+flow+cytometry.">Kinetics and specificity of HEK293T extracellular vesicle uptake using imaging flow cytometry.</a> Nanoscale Research Letters. 15 (1), 170 (2020).</li></ol>

The authors have no conflicts of interest to disclose.

In this protocol, we showed the labeling of sEVs with PKH dyes and the visualization of their uptake in the spinal cord. PKH lipophilic fluorescent dyes are widely used for labeling cells by flow cytometry and fluorescent microscopy3,5,6,12,24,25. Due to their relatively long half-life and low cytotoxicity, PKH dyes can be us...

Small extracellular vesicles (sEVs) are nanosized, membrane-derived vesicles with a size range of 50-150 nm. They originate from multi-vesicular bodies (MVBs) and are released from cells upon fusion of the MVBs with the plasma membrane. sEVs contain miRNAs, mRNAs, proteins, and bioactive lipids, and these molecules are transferred between cells in the form of cell-to-cell communication. sEVs can be internalized by recipient cells by a variety of endocytic pathways, and this capture of sEVs by recipient cells is mediated by the recognition of surface molecules on both EVs and the target cells1.
sEVs have gained interest due to their capacity to trigger molecular and phenotypic changes in acceptor cells, their utility as a therapeutic agent, and their potential as carriers for cargo molecules or pharmacological agents. Due to their small size, the imaging and tracking of sEVs can be challenging, especially for in vivo studies and clinical settings. Therefore, many methods have been developed to label and image sEVs to assist their biodistribution and tracking in vitro and in vivo2.
The most common technique to study sEV biodistribution and target cell interactions involves labeling them with fluorescent dye molecules3,4,5,6,7. EVs were initially labeled with cell membrane dyes that were commonly used to image cells. These fluorescent dyes generally stain the lipid bilayer or proteins of interest on sEVs. Several lipophilic dyes display a strong fluorescent signal when incorporated into the cytosol, including DiR (1,1&#8242;-dioctadecyl-3,3,3&#8242;,3&#8242;-tetramethylindotricarbocyanine iodide), DiL (1, 1&#8242;-dioctadecyl-3, 3, 3&#8242;, 3&#8242;-tetramethyl indocarbocyanine perchlorate), and DiD (1, 1&#8242;-dioctadecyl-3, 3, 3&#8242;, 3&#8242;-tetramethyl indocarbocyanine 4-chlorobenzenesulfonate salt)8,9,10,11.
Other lipophilic dyes, such as PKH67 and PKH26, have a highly fluorescent polar head group and a long aliphatic hydrocarbon tail that readily intercalates into any lipid structure and leads to long-term dye retention and stable fluorescence12. PKH dyes can also label EVs, which allows the study of EV properties in vivo13. Many other dyes have been used to observe exosomes using fluorescence microscopy and flow cytometry, including lipid-labeling dyes14 and cell-permeable dyes such as carboxyfluorescein diacetate succinimidyl ester (CFDA-SE)15,16 and calcein acetoxymethyl (AM) ester17.
Studies of sEV-mediated crosstalk between different cells in the CNS have provided important insights on the pathogenesis of neuroinflammatory and neurodegenerative diseases18. For example, sEVs from neurons can spread beta-amyloid peptides and phosphorylated tau proteins and aid in the pathogenesis of Alzheimer&#39;s disease19. Additionally, EVs derived from erythrocytes contain large amounts of alpha-synuclein and can cross the blood-brain barrier and contribute to Parkinson&#39;s pathology20. The ability of sEVs to cross physiological barriers21 and transfer their biomolecules to target cells makes them convenient tools to deliver therapeutic drugs to the CNS22.
Visualizing sEV uptake by myriad CNS cells in the spinal cord will enable both mechanistic studies and the evaluation of the therapeutic benefits of exogenously administered sEVs from various cellular sources. This paper describes the methodology to label sEVs derived from macrophages and image their uptake in vitro and in vivo in the lumbar spinal cord by neurons, microglia, and astrocytes to qualitatively confirm sEV delivery by visualization.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Amicon Ultra 0.5 mL centrifugal filters</td>
			<td>MilliporeSigma</td>
			<td>Z677094</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-Alix Antibody</td>
			<td>Abcam</td>
			<td>ab186429</td>
			<td>1:1000</td>
		</tr>
		<tr>
			<td>Anti-Calnexin Antibody</td>
			<td>Abcam</td>
			<td>Ab10286</td>
			<td>1:1000</td>
		</tr>
		<tr>
			<td>Anti-CD81 Antibody</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-166029</td>
			<td>1:1000</td>
		</tr>
		<tr>
			<td>Anti-GAPDH Monoclonal Antibody (14C10)</td>
			<td>Cell Signaling Technology</td>
			<td>2118</td>
			<td>1:1000</td>
		</tr>
		<tr>
			<td>Anti-Glial Fibrillary Acidic Protein Antibody</td>
			<td>Sigma-Aldrich</td>
			<td>MAB360</td>
			<td>1:500 for IF; 1:1000 for IHC</td>
		</tr>
		<tr>
			<td>Anti-Iba1 Antibody</td>
			<td>Wako</td>
			<td>019-19741</td>
			<td>1:2000</td>
		</tr>
		<tr>
			<td>Anti-MAP2A Antibody</td>
			<td>Sigma-Aldrich</td>
			<td>MAB378</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>VWR</td>
			<td>0332</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Strainer, 40 &#956;m</td>
			<td>VWR</td>
			<td>15-1040-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge Tubes</td>
			<td>Thermo Scientific</td>
			<td>3118-0050</td>
			<td>12,000 x g</td>
		</tr>
		<tr>
			<td>Coverslip, 12-mm, #1.5</td>
			<td>Electron Microscopy Sciences</td>
			<td>72230-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslip, 18-mm, #1.5</td>
			<td>Electron Microscopy Sciences</td>
			<td>72222-01</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma-Aldrich</td>
			<td>D9542-1MG</td>
			<td>1 &#181;g/mL</td>
		</tr>
		<tr>
			<td>DC Protein Assay</td>
			<td>Bio-Rad</td>
			<td>500-0116</td>
			<td></td>
		</tr>
		<tr>
			<td>Deoxyribonuclease I (DNAse I)</td>
			<td>MilliporeSigma</td>
			<td>D4513-1VL</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey Anti-Rabbit IgG H&#38;L (HRP)</td>
			<td>Abcam</td>
			<td>ab16284</td>
			<td>1:10000</td>
		</tr>
		<tr>
			<td>Donkey Anti-Rabbit IgG H&#38;L, Alexa Fluor 488</td>
			<td>Invitrogen</td>
			<td>A-21206</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>Double Frosted Microscope Slides, #1</td>
			<td>Thermo Scientific</td>
			<td>12-552-5</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS without Calcium and Magnesium</td>
			<td>Corning</td>
			<td>21-031-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium (DMEM)</td>
			<td>Corning</td>
			<td>10-013-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Exosome-Depleted Fetal Bovine Serum</td>
			<td>Gibco</td>
			<td>A27208-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>Corning</td>
			<td>35-011-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>FluorChem M imaging system</td>
			<td>ProteinSimple</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>FV3000 Confocal Microscope</td>
			<td>Olympus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Goat Anti-Mouse IgG H&#38;L (HRP)</td>
			<td>Abcam</td>
			<td>ab6789</td>
			<td>1:10000</td>
		</tr>
		<tr>
			<td>Goat Anti-Mouse IgG H&#38;L, Alexa Fluor 488</td>
			<td>Invitrogen</td>
			<td>A-11001</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>Goat Anti-Mouse IgG1, Alexa Fluor 594</td>
			<td>Invitrogen</td>
			<td>A-21125</td>
			<td></td>
		</tr>
		<tr>
			<td>Hank&#39;s Balanced Salt Solution (HBSS)</td>
			<td>VWR</td>
			<td>02-0121</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Gibco</td>
			<td>15630080</td>
			<td></td>
		</tr>
		<tr>
			<td>HRP Substrate</td>
			<td>Thermo Scientific</td>
			<td>34094</td>
			<td></td>
		</tr>
		<tr>
			<td>Intercept blocking buffer, TBS</td>
			<td>LI-COR Biosciences</td>
			<td>927-60001</td>
			<td></td>
		</tr>
		<tr>
			<td>Laemmli SDS Sample Buffer</td>
			<td>Alfa Aesar</td>
			<td>AAJ61337AC</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro Cover Glass, #1</td>
			<td>VWR</td>
			<td>48404-454</td>
			<td></td>
		</tr>
		<tr>
			<td>Microm HM550</td>
			<td>Thermo Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NanoSight NS300 system</td>
			<td>Malvern Panalytical</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NanoSight NTA 3.2 software</td>
			<td>Malvern Panalytical</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Neuro-2a Cell Line</td>
			<td>ATCC</td>
			<td>CCL-131</td>
			<td></td>
		</tr>
		<tr>
			<td>Normal Goat Serum</td>
			<td>Vector Laboratories</td>
			<td>S-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>O.C.T Compound</td>
			<td>Sakura Finetek</td>
			<td>4583</td>
			<td></td>
		</tr>
		<tr>
			<td>Papain</td>
			<td>Worthington Biochemical Corporation</td>
			<td>NC9597281</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>19210</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Gibco</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr>
			<td>PKH26</td>
			<td>Sigma-Aldrich</td>
			<td>MINI26-1KT</td>
			<td></td>
		</tr>
		<tr>
			<td>PKH67</td>
			<td>Sigma-Aldrich</td>
			<td>MINI67-1KT</td>
			<td></td>
		</tr>
		<tr>
			<td>Protease Inhibitor Cocktail</td>
			<td>Thermo Scientific</td>
			<td>1862209</td>
			<td></td>
		</tr>
		<tr>
			<td>PVDF Transfer Membrane</td>
			<td>MDI</td>
			<td>SVFX8302XXXX101</td>
			<td></td>
		</tr>
		<tr>
			<td>RAW 267.4 Cell Line</td>
			<td>ATCC</td>
			<td>TIB-71</td>
			<td></td>
		</tr>
		<tr>
			<td>RIPA Buffer</td>
			<td>Sigma-Aldrich</td>
			<td>R0278</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>AMRESCO</td>
			<td>0241-2.5KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Superfrost Plus Gold Slides</td>
			<td>Thermo Scientific</td>
			<td>15-188-48</td>
			<td>adhesive slides</td>
		</tr>
		<tr>
			<td>T-75 Flasks</td>
			<td>Corning</td>
			<td>431464U</td>
			<td></td>
		</tr>
		<tr>
			<td>Tecnai 12 Digital Transmission Electron Microscope</td>
			<td>FEI Company</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>TEM Grids</td>
			<td>Electron Microscopy Sciences</td>
			<td>FSF300-cu</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris-Glycine Protein Gel, 12%</td>
			<td>Invitrogen</td>
			<td>XP00120BOX</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris-Glycine SDS Running Buffer</td>
			<td>Invitrogen</td>
			<td>LC26755</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris-Glycine Transfer Buffer</td>
			<td>Invitrogen</td>
			<td>LC3675</td>
			<td></td>
		</tr>
		<tr>
			<td>TrypLE Express</td>
			<td></td>
			<td>cell dissociation enzyme</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Acros Organics</td>
			<td>327371000</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin, 0.25%</td>
			<td>Corning</td>
			<td>25-053-CL</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween 20</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ultracentrifuge Tubes</td>
			<td>Beckman</td>
			<td>344058</td>
			<td>110,000 x g</td>
		</tr>
	</tbody>
</table>

uptake of fluorescent labeled small extracellular vesicles in vitro and in spinal cord

Small extracellular vesicles (sEVs) are 50-150 nm vesicles secreted by all cells and present in bodily fluids. sEVs transfer biomolecules such as RNA, proteins, and lipids from donor to acceptor cells, making them key signaling mediators between cells. In the central nervous system (CNS), sEVs can mediate intercellular signaling, including neuroimmune interactions. sEV functions can be studied by tracking the uptake of labeled sEVs in recipient cells both in vitro and in vivo. This paper describes the labeling of sEVs from the conditioned media of RAW 264.7 macrophage cells using a PKH membrane dye. It shows the uptake of different concentrations of labeled sEVs at multiple time points by Neuro-2a cells and primary astrocytes in vitro. Also shown is the uptake of sEVs delivered intrathecally in mouse spinal cord neurons, astrocytes, and microglia visualized by confocal microscopy. The representative results demonstrate time-dependent variation in the uptake of sEVs by different cells, which can help confirm successful sEVs delivery into the spinal cord.

After the isolation of sEVs from RAW 264.7 conditioned media via centrifugation, NTA was used to determine the concentration and size distribution of the purified sEVs. The average mean size of RAW 264.7-derived sEVs was 140 nm, and the peak particle size was 121.8 nm, confirming that most detectable particles in the light scattering measurement fell within the size range of exosomes or sEVs at 50-150 nm (Figure 1A). As suggested in the minimal information for studies of extracellular vesicl...

Watch this Scientific Journal Video about Uptake of Fluorescent Labeled Small Extracellular Vesicles In Vitro and in Spinal Cord at JoVE.com

Uptake of Fluorescent Labeled Small Extracellular Vesicles In Vitro and in Spinal Cord

We describe a protocol to label macrophage-derived small extracellular vesicles with PKH dyes and observe their uptake in vitro and in the spinal cord after intrathecal delivery.

Uptake of Fluorescent Labeled Small Extracellular Vesicles In Vitro and in Spinal Cord

Small extracellular vesicles (sEVs) are 50-150 nm vesicles secreted by all cells and present in bodily fluids. sEVs transfer biomolecules such as RNA, ...

uptake-fluorescent-labeled-small-extracellular-vesicles-vitro-spinal

Research

JoVE Journal

Neuroscience

3.4K Views.  Drexel University College of Medicine. We describe a protocol to label macrophage-derived small extracellular vesicles with PKH dyes and observe their uptake in vitro and in the spinal cord after intrathecal delivery.

Video: Uptake of Fluorescent Labeled Small Extracellular Vesicles In Vitro and in Spinal Cord

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 ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

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 ...

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g., calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability.

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal ...

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 ...

Laminectomy and Spinal Cord Window Implantation in the Mouse

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated digital vaporizer is utilized to achieve a stable plane of anesthesia at a low-flow rate of isoflurane. A single vertebral spine is removed, and a commercially available cover-glass is overlaid on a thin agarose bed. A 3D-printed plastic backplate is then affixed to the adjacent vertebral spines using tissue adhesive and dental cement. A stabilization platform is used to reduce motion artifact from respiration and heartbeat. This rapid and clamp-free method is well-suited for acute multi-photon fluorescence microscopy. Representative data are included for an application of this technique to two-photon microscopy of the spinal cord vasculature in transgenic mice expressing eGFP:Claudin-5 &#8212; a tight junction protein.

This protocol describes implantation of a glass window onto the spinal cord of a mouse to facilitate visualization by intravital microscopy.

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated ...

Preparation of Rhythmically-active In Vitro Neonatal Rodent Brainstem-spinal Cord and Thin Slice

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a signal driving the rhythmic contraction of inspiratory muscles. Rhythmic neural activity generated in the pBC and carried to other neuronal pools to drive the musculature of breathing may be studied using various approaches, including en bloc nerve recordings and transverse slice recordings. However, previously published methods have not extensively described the brainstem-spinal cord dissection process in a transparent and reproducible manner for future studies. Here, we present a comprehensive overview of a method used to reproducibly cut rhythmically-active brainstem slices containing the necessary and sufficient neuronal circuitry for generating and transmitting inspiratory drive. This work builds upon previous brainstem-spinal cord electrophysiology protocols to enhance the likelihood of reliably obtaining viable and rhythmically-active slices for recording neuronal output from the pBC, hypoglossal premotor neurons (XII pMN), and hypoglossal motor neurons (XII MN). The work presented expands upon previous published methods by providing detailed, step-by-step illustrations of the dissection, from whole rat pup, to in vitro slice containing the XII rootlets.

This protocol both visually communicates the brainstem-spinal cord preparation and clarifies the preparation of brainstem transverse slices in a comprehensive step-by-step manner. It was designed to increase reproducibility and enhance the likelihood of obtaining viable, long lasting, rhythmically-active slices for recording neural output from the respiratory regions of the brainstem.

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a ...

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 ...

A Minimally Invasive, Fast Spinal Cord Lateral Hemisection Technique for Modeling Open Spinal Cord Injuries in Rats

Open spinal cord injury techniques modeling laceration-like injuries are time-consuming and invasive because they involve laminectomy. This new technique eliminates laminectomy by removing two spinous processes and lifting, then tilting the caudal vertebral arch. The surgical area opens up without the need for laminectomy. Lateral hemisection is then performed with direct visible control under a microscope. The trauma is minimized, requiring only a small bone wound. 
This technique has several advantages: it is faster and, therefore, less of a burden for the animal, and the bone wound is smaller. Because the laminectomy is eliminated, there is less chance for unwanted injury to the spinal cord, and there are no bone splinters that can cause problems (bone splinters embedded in the spinal cord can cause swelling and secondary damage). The vertebral canal remains intact. The main limitation is that the hemisection can only be performed in the intervertebral spaces. 
The results show that this technique can be performed much faster than the traditional surgical approach, using laminectomy (11 min vs. 35 min). This technique can be useful for researchers working with animal models of open spinal cord injury as it is widely adaptable and does not require any additional specialized instrumentation.

Here, we describe a new, fast technique modeling open spinal cord injury in rats that eliminates laminectomy. Lateral hemisection is performed while viewing through a microscope. The technique is versatile and can also be used in the cervical, thoracic, and lumbar regions of the spinal cord of other animals.

Open spinal cord injury techniques modeling laceration-like injuries are time-consuming and invasive because they involve laminectomy. This new technique ...