We like to acknowledge the National Institutes of Health and the National Institute of Neurological Disorders and Stroke for funding (R01NS079691).

The experiments were conducted in accordance with IUCAC at Duke University and University of California Los Angeles. 8 to 12-week-old male C57Bl/6J mice were used in this study. The animals were housed under controlled temperature (22 &#177; 2 &#176;C), with a 12 h light-dark cycle period and access to pelleted food and water&#160;ad libitum.Analgesia and sedation protocols are described as approved by the IUCAC but might differ from protocols used in other laboratories.
Animals may be prematurely euthanized if they experience excessive restlessness, vocalization (indicating uncontrollable pain), self-trauma, decreased or impaired mobility, physical signs of dermal infection, loss of appetite and/or weight loss greater than 10%-15% as an outcome of the survival surgery. There are no anticipated adverse effects from injection of the gel, as it is a composed of a naturally&#160;occurring polymer within the brain. Animals should be&#160;monitored daily, including weekends and holidays, and reweighed every other day. Studies from our laboratory and others&#39; have found no deficits in grooming, loss of body weight or signs/symptoms of withdrawal or chronic stress in mice after these small cortical or subcortical strokes. If animals exhibit any signs of local head wound infection they should&#160;be euthanized. Animals should also be monitored for poor grooming, wound condition, and evidence of fighting between animals (back wounds). If&#160;there is evidence of fighting or wound dehiscence, animals should&#160;first be treated after contacting the veterinarians. This often involves antibiotic in the water and/or local topical antibiotic application. If these measures fail to produce healing, animals should be&#160;euthanatized. &#160;
1. Preparation of materials and instruments
<ol>
	<li>Prepare photothrombotic dye prior to the surgery. Weigh out 15 mg of Rose Bengal into a 2 mL microcentrifuge tube. 
	CAUTION: Rose Bengal can cause serious eye damage/irritation.
	<ol>
		<li>Dissolve the Rose Bengal in 1.5 mL of sterile filtered 1x PBS to make a working concentration of 10 mg/mL.</li>
		<li>Vortex the tube until no clumps are visible, indicating the dye has fully dissolved, then cover in foil until further use. 
		NOTE: Amount of dye required will depend on the number of mice. Dye is injected intraperitoneally IP at a concentration of 10 &#181;L/g, e.g., 200 &#181;L for a 20 g mouse.</li>
	</ol>
	</li>
	<li>Turn on the heating pad to maintain the mouse body temperature during anesthesia (37 &#176;C).</li>
	<li>Prepare autoclaved scissors, forceps, cotton, vet eye ointment, surgical glue, or suture material, a blue or black surgical marker, and sterile alcohol and beta iodine wipes. Prepare bone drill with sterilized drill bit heads.</li>
	<li>Turn on the bead sterilization to 160 &#176;C to allow for sterilization of instruments between mice surgery. Set aside a 50 mL conical tube of sterile saline or 1x PBS solution for the later use in maintaining a hydrated operation area.</li>
	<li>Attach the laser to the stereotaxic device. Wearing laser safety goggles, measure the power of the laser (mW). 
	CAUTION: Use laser safety goggles whenever the laser is turned on in the procedure.
	<ol>
		<li>Adjust the current on the laser console, following the laser manufacturing instructions, until the laser power output reads 10 mW, and set the current as a pre-set on the home screen console. Then adjust the current again to establish another pre-set where the laser output power reads 40 mW. Now turn the laser off until further use. 
		NOTE: The 10 mW will be used to orient the laser prior to use.</li>
	</ol>
	</li>
	<li>Prepare a clean cage for mice after surgery. Then, place the mouse into the induction chamber with an isoflurane concentration of 4% to anesthetize it until spontaneous movement stops and its breathing comes to a slow, steady rate.</li>
	<li>Once asleep, weigh the mouse to determine how much Rose Bengal dye to inject. Then, transfer and secure the mouse to the stereotaxic device with a maintenance isoflurane concentration of 1.5%. 
	NOTE: Increasing isoflurane concentrations can dilate vessels and prevent sufficient occlusion or consistently sized strokes between mice.</li>
	<li>Apply vet eye ointment to both eyes. 
	NOTE: Watch breathing and temperature throughout the procedure and pinch toes occasionally to ensure that the mouse cannot feel anything.</li>
</ol>
2. Photothrombotic stroke model9
<ol>
	<li>Shave the fur off the mouse&#8217;s head and aseptically prepare the area by using an alcohol wipe followed by a beta iodine wipe. Repeat three times. 
	NOTE: If necessary, use an extra alcohol wipe at the end to clean off all beta iodine as it causes skin irritation which leads the mice to itch their skull after surgery.</li>
	<li>Using small operation scissors, make a lateral incision between the ears to the eyes, about 1-2 cm. Do this by pulling the skin upward with forceps to create a tent, cutting once downwards, then inserting the scissors into the opening and creating a continuous midline incision.</li>
	<li>Separate the skin and press lightly on the parietal bones with the forceps to locate the bregma. Mark the bregma with a dot using the surgical marker. 
	NOTE: Bone fragment movement highlights the frontal border of the parietal bones. The bregma is the center point at which the frontal and parietal bone fragments meet (Figure 1A).</li>
	<li>Wearing laser safety goggles, move the laser over the skull of the mouse, fixing the stereotaxic device at a 90&#176; angle. Select the 10 mW pre-set and turn on the laser.
	<ol>
		<li>Adjust the laser&#8217;s x- and y-axis until it is directly over the bregma. Reset the x and y on the digital display console, and then move the laser 1.8 mm left of the bregma. Now bring the laser as close to the skull as possible without letting it touch the skull.</li>
		<li>Move the laser to 2.2 mm left of the bregma to ensure that it can move without any obstructions. Turn off the laser.</li>
	</ol>
	</li>
	<li>Inject the appropriate amount of Rose Bengal dye intraperitoneally (IP) using a disposable 29 G needle. Immediately start a timer for 7 min to allow dye to circulate systemically. Select the pre-set for 40 mW without turning the laser on. 
	NOTE: Once comfortable with the procedure, the dye can be injected before aseptic preparation of the mouse and finished before the 7 min are over to save time.</li>
	<li>Turn on the laser (40 mW) when the 7 min timer goes off, while wearing laser safety goggles, and set a 10 min timer.
	<ol>
		<li>After 10 min, move the laser to 2.2 mm left of the bregma without turning it off, and set another 10 min timer.</li>
		<li>Once the second 10 min timer has gone off, change the laser back to 10 mW and move the laser to 2.0 mm left of the bregma. Mark this spot with the surgical marker. Then turn off the laser.</li>
		<li>Lift the laser and unlock it from the 90&#176; angle so it can be moved away from the mouse. At this point, apply the sterile saline or PBS with cotton if the surgical area is dry.</li>
	</ol>
	</li>
	<li>Drill in small bursts at the 2.0 mm mark perpendicular to the surface of the skull. 
	NOTE: Be careful to drill slowly to avoid drilling too deeply and hitting the brain. There will be a slight drop/change in resistance once the drill has gone through the skull, and drilling may cause bleeding.
	<ol>
		<li>Use cotton to absorb any excess blood. It is typical to see some blood from nicked arteries in the skull after drilling &#8211; excessive blood means the drill bit hit the brain. If this happens, record the mouse identifier, and move on.</li>
	</ol>
	</li>
	<li>Place a small wipe next to the mouse. Using the forceps, pull the skin close together to prepare for gluing. 
	NOTE: Suturing can be done here instead. Suturing typically requires only 3 sutures.
	<ol>
		<li>Using the forceps, grab the two sides of the skin and pull them together and upward. Dab off any large drops of surgical glue at the end of the tip on the wipe before applying to the mouse.</li>
		<li>Apply the surgical glue sparingly and hold the skin together with forceps for 10 s. Continue until there are no visible openings. 
		NOTE: The glue comes out quickly &#8211; do not squeeze the bottle. Avoid gluing the skin to the skull or getting glue in the eye.</li>
	</ol>
	</li>
	<li>After gluing, or suturing, place the mouse in the new cage to recover from anesthesia. It takes 5-10 min for the animal to recover, and it helps to rest half of the cage on a heating pad.</li>
</ol>
3. Sham stroke operation
<ol>
	<li>Perform all procedures identically to the operation described above in section 2 except inject 1x PBS or saline instead of Rose Bengal dye.</li>
</ol>
4. Injection of hydrogel or other biomaterials
<ol>
	<li>Perform the injection of biomaterials at a user-defined time. In this experiment injections were performed between 5- and 7-days post-stroke. 2 - 6 &#181;L of hydrogel are injected (user-defined).
	<ol>
		<li>Turn on the heating pad to maintain the mouse body temperature during anesthesia (37 &#176;C).</li>
		<li>Prepare autoclaved scissors, forceps, cotton, vet eye ointment, surgical glue or suture material, and sterile alcohol and beta iodine wipes. Prepare the bone drill with sterile drill bit heads and the 25 &#181;L glass Hamilton syringes with metal 30 G needles.</li>
		<li>Turn on the bead sterilization to 160 &#176;C to allow for sterilization of instruments between mice.</li>
		<li>Attach the injection pumps to the stereotaxic device. Set the flow rate to 1 &#181;L/min and set volume to 4 &#181;L.</li>
		<li>Prepare a clean cage for the mice after surgery.</li>
	</ol>
	</li>
	<li>Place the mouse into the induction chamber with an isoflurane concentration of 4% to anesthetize it until spontaneous movement stops and its breathing comes to a slow, steady rate.
	<ol>
		<li>Once asleep, transfer and secure the mouse to the stereotaxic device with a maintenance isoflurane concentration of 1.5%.</li>
		<li>Apply the vet eye ointment to both eyes.</li>
		<li>Shave any fur that may have regrown of the mouse&#8217;s head, and aseptically prepare the area by using an alcohol wipe followed by a beta iodine wipe. Repeat three times. 
		NOTE: If necessary, use an extra alcohol wipe at the end to clean off all the beta iodine as it causes skin irritation and leads the mice to itch their skull after surgery.</li>
	</ol>
	</li>
	<li>Using the small scissors and/or forceps, reopen the skin above the brain. Use cotton with sterile saline or PBS to clean any debris from the skull. A white-yellow circle (the stroke, Figure 1B) and the burr hole from the drill will be visible. If the dead tissue is not visible, likely no stroke occurred. If the burr hole is not centered above the stroke, re-drill to center it.</li>
	<li>Orient the injection pump over the mouse and lock at 90&#176;. Clean the syringe with sterile saline or PBS solution before backloading material.
	<ol>
		<li>Once clean, disassemble the glass syringe so that there is no needle. Backload the syringe using a 25 &#181;L positive displacement pipette with the 10 &#181;L of hydrogel (or other biomaterial here with or without cells).</li>
		<li>Using the syringe plunger, push the gel all the way to the front of the syringe. Then, reassemble the syringe and continue pushing until the gel visibly exudes from the needle.</li>
	</ol>
	</li>
	<li>Place the syringe onto the pump. The back of the pump may need to be adjusted so the syringe fits. Do this by adjusting the flow rate to 30 &#181;L/min, select Withdrawal or Inject depending on the direction it needs to move, and then press Start. Hit Stop when the back of the pump is at the correct location.
	<ol>
		<li>Screw the back of the pump so the syringe is secure. If adjustments were previously made, make sure the flow rate is set back to 1 &#181;L/min and is set to inject.</li>
	</ol>
	</li>
	<li>Move the pump in the x and y direction to orient it over the hole. Move the pump down in the z direction until the needle of the syringe touches the top of the hole.</li>
	<li>Reset the z on the digital display console. Then, move the syringe down in the z direction 0.750 mm. If the syringe bends or there is resistance, the skull either healed or was not completely drilled through and needs to be re-drilled.</li>
	<li>Press Start on the pump console to inject 4 - 6 &#181;L of the hydrogel at a rate of 1 &#181;L/min.</li>
	<li>Set a 5 min timer when the pump stops injecting to allow the gel to begin crosslinking.</li>
	<li>After 5 min, slowly pull up the syringe by turning the z nob. Watch to see if any material is accidently pulled or leaking from the hole. Unlock the pump and move it away from the mouse when the needle is far enough away from the skull.</li>
	<li>Using forceps, surgical glue, or sutures, close the skin above the head. Place the mouse in the clean cage and observe its recovery. It should take about 5 &#8211; 10 min for the mouse to recover from anesthesia.</li>
</ol>
5. Perfusion and sample collection
<ol>
	<li>Anesthetize the mouse using isoflurane.</li>
	<li>Fix the animal in the supine position and open the abdominal cavity with a median cut. Optionally, clamp the descending abdominal aorta to use less fixative and PBS.</li>
	<li>Cut open the diaphragm and move up the sides of the rib cage to open the chest cavity. Insert a perfusion cannula into the left ventricle and cut an opening in the right atrium. Start to slowly perfuse with 10-20 mL of PBS or until the liquid runs semi-clear.</li>
	<li>Once clear, switch to 4% PFA for another 10 &#8211; 20 mL. Unpinning the arms allows them to contract as the PFA infuses. Once they stop moving, wait for another 30 s.</li>
	<li>Decapitate the animal and pull back the skin to expose the skull. Using blunt, thin forceps, carefully remove the pieces of the skull to expose the brain. Special care needs to be taken when removing the skull above the stroke site. Small surgical scissors can be used here to help cut away the skull. The biggest challenge is gel getting stuck to the skull and coming out of the brain as the skull is removed.</li>
	<li>Once the brain is detached, place in 10 &#8211; 15 mL of 4% PFA overnight (no longer than 24 h).</li>
	<li>Move the brains from PFA to 30% sucrose the following day. The brain is ready for cryosection once it sinks to the bottom (about 2 &#8211; 3 days). It can also be left for storage at this point.</li>
</ol>
6. Cryosectioning and staining
<ol>
	<li>Take the brain out of sucrose and place on a glass slide. Cut a small portion of the back of the brain off with a razor blade to create a flat portion to be mounted onto a chuck.
	<ol>
		<li>Place a dollop of Optimal Cutting Temperature Compound (OCT) onto the chuck, and then place the brain, flat side down, onto the chuck in the OCT. Place this specimen on dry ice until frozen or in the cryostat on the freezing block. 
		NOTE: OCT can be added to fully encompass the brain to help prevent the material from separating from the brain during sectioning.</li>
	</ol>
	</li>
	<li>Cut the brains serially on a cryostat at -20 &#176;C at 30 &#181;m thick sections across 10 gelatin coated slides. Take care when sectioning ensuring not to lose the hydrogel. If this part is difficult, place some OCT on the top of the brain over the gel (Figure 3). Store at -80 &#176;C until further use.</li>
	<li>Take slides out of the -80 &#176;C and place on a staining tray to allow them to dry. Slides that have not be frozen but just sectioned can also be stained immediately.</li>
	<li>Using a hydrophobic pen, draw a circle around the brain slices on the slide. Once dry, rehydrate the brain slices with 1x filtered PBS and place on a shaker at room temperature for 15 min. This takes about 1 mL of buffer per slide with 9 &#8211; 12 brain sections.</li>
	<li>Wash the slides with 1x filtered PBS for 5 min at room temperature, repeat three times.</li>
	<li>Block slides for 30 min to an hour with 10% donkey serum in 0.01% PBS-Triton X at room temperature on the belly dancer. This takes about 200 &#181;L per slide.</li>
	<li>Incubate the primary antibody in 10% donkey serum with 0.01% PBS-Triton X overnight at 4&#176;C on a shaker. Test primary antibodies first to determine correct concentrations. Here GFAP (1:400), Iba1 (1:250), Glut1 (1:500) (Figure 4) were used.</li>
	<li>Wash slides with 1x filtered PBS the following day for 5 min at room temperature, repeat three times.</li>
	<li>Incubate with secondary antibody in 10% donkey serum in 0.01% PBS-Triton X and DAPI for 2 h on the lab shaker at room temperature. 1:1,000 was used for all secondaries and DAPI.</li>
	<li>Wash the slides for 5 min with 1x filtered PBS.</li>
	<li>Allow sections to dry at room temperature in the dark. This can take 20 min to 4 h; placing the slides in a desiccator can speed up the process.</li>
	<li>Dehydrate slides in ascending concentrations of alcohol (1 min per solution) of 50%, 70%, 95%, 100%, and 100%.</li>
	<li>De-fat in first bath of xylene for 1 min, then second bath of xylene for 5 min.</li>
	<li>Quickly take the slides out one-by-one and add mounting medium while brain sections are wet with xylene. Quickly add a glass coverslip on top. Use a cover slip that is of correct length and thickness needed for the slide and microscope used for imaging, respectively.</li>
	<li>Get rid of any bubbles under the coverslip by pressing gently with a pipette tip in an outward motion from the center of the slide. Allow DPX to dry for 4 h to overnight in the dark at room temperature.</li>
	<li>Use a razor blade to scrape off any dried mounting medium that spilled onto the slides. Wipe down slides with lens cleaner. Slides can now be imaged with fluorescence microscopy. 
	NOTE: It is best to store slide in the dark or at 4 &#176;C until imaging is finished.</li>
</ol>

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L., Anandakumaran, P. N., Varga, B. V., Morshead, C. M., Nagy, A., Shoichet, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+maturation+of+human+iPSC-derived+neuroepithelial+cells+influences+transplant+survival+in+the+stroke-injured+rat+brain.">In vitro maturation of human iPSC-derived neuroepithelial cells influences transplant survival in the stroke-injured rat brain.</a> Tissue Engineering Part A. 24, 351-360 (2018).</li><li>Lee, I. 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=Delayed+epidural+transplantation+of+human+induced+pluripotent+stem+cell-derived+neural+progenitors+enhances+functional+recovery+after+stroke.">Delayed epidural transplantation of human induced pluripotent stem cell-derived neural progenitors enhances functional recovery after stroke.</a> Science Reports. 7, 1943 (2017).</li><li>Somaa, F. 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=Peptide-based+scaffolds+support+human+cortical+progenitor+graft+integration+to+reduce+atrophy+and+promote+functional+repair+in+a+model+of+stroke.">Peptide-based scaffolds support human cortical progenitor graft integration to reduce atrophy and promote functional repair in a model of stroke.</a> Cell Reports. 20, 1964-1977 (2017).</li><li>Griffin, D. R., Weaver, W. M., Scumpia, P., Di Carlo, D., Segura, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Accelerated+wound+healing+by+injectable+microporous+gel+scaffolds+assembled+from+annealed+building+blocks.">Accelerated wound healing by injectable microporous gel scaffolds assembled from annealed building blocks.</a> Nature Materials. 14 (7), 737-744 (2016).</li><li>Sideris, E. Y., Aaron, C. J., Carmichael, S. T., Segura, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hyaluronic+acid+particle+hydrogels+decrease+cerebral+atrophy+and+promote+pro-reparative+astrocyte/axonal+infiltration+in+the+core+after+ischemic+stroke.">Hyaluronic acid particle hydrogels decrease cerebral atrophy and promote pro-reparative astrocyte/axonal infiltration in the core after ischemic stroke.</a> bioRxiv. , (2019).</li><li>Yu, 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=Combinated+Transplantation+of+Neural+Stem+Cells+and+Collagen+Type+I+Promote+Functional+Recovery+After+Cerebral+Ischemia+in+Rats.">Combinated Transplantation of Neural Stem Cells and Collagen Type I Promote Functional Recovery After Cerebral Ischemia in Rats.</a> The Anatomical Record. 293 (5), 911-917 (2010).</li><li>Jin, K., 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=Transplantation+of+Human+Neural+Precursor+Cells+in+Matrigel+Scaffolding+Improves+Outcome+from+Focal+Cerebral+Ischemia+after+Delayed+Postischemic+Treatment+in+Rats.">Transplantation of Human Neural Precursor Cells in Matrigel Scaffolding Improves Outcome from Focal Cerebral Ischemia after Delayed Postischemic Treatment in Rats.</a> Journal of Cerebral Blood Flow &amp; Metabolism. 30 (3), 534-544 (2009).</li><li>Gelderblom, 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=Temporal+and+spatial+dynamics+of+cerebral+immune+cell+accumulation+in+stroke.">Temporal and spatial dynamics of cerebral immune cell accumulation in stroke.</a> Stroke. 40 (5), 1849-1857 (2009).</li><li>Wei, L., Erinjeri, J. P., Rovainen, C. M., Woolsey, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Collateral+growth+and+angiogenesis+around+cortical+stroke.">Collateral growth and angiogenesis around cortical stroke.</a> Stroke. 32 (9), 2179-2184 (2001).</li><li>Zhang, 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=Activated+Neural+Stem+Cells+Contribute+to+Stroke-Induced+Neurogenesis+and+Neuroblast+Migration+toward+the+Infarct+Boundary+in+Adult+Rats.">Activated Neural Stem Cells Contribute to Stroke-Induced Neurogenesis and Neuroblast Migration toward the Infarct Boundary in Adult Rats.</a> Journal of Cerebral Blood Flow &amp; Metabolism. 24 (4), 441-448 (2004).</li></ol>

The authors declare that TS is a founder in Tempo Therapeutics, which seeks to commercialize MAP hydrogels.

Here we demonstrate an easily reproducible, minimally invasive, permanent stroke model and describe how to inject a biomaterial into the infarct five days after stroke. The use of the photothrombotic dye Rose Bengal and a 520 nm collimated laser connected to the stereotaxic device gives us the ability to position the stroke at the motor cortex of the mouse with enhanced precision. Five days after stroke, the location of the infarct is visible by eye at the center of irradiation, 2.0 mm medio-lateral to the bregma. Hydrog...

Stroke is the leading cause of disability and the fifth-leading cause of death in the United States1. Approximately 87% of all strokes are ischemic, while a majority of the remaining 13% are hemorrhagic2. An ischemic stroke is defined as the blockage of blood flow in an artery to the surrounding tissue. This occlusion results in oxygen deprivation and subsequent necrosis that often leads to permanent disability in surviving patients. While there has been a decrease in the mortality rate of stroke3, its prevalence is expected to increase to 3.4 million people by 20304. This increase in disabled survivors and consequent economic burden has led to a push for stroke research that focuses on mechanisms of neural repair. Following stroke there is an inflammatory period that leads to the formation of a scar that prevents the necrotic region from expanding. The region surrounding the necrotic core is termed &#8220;peri-infarct&#8221; and there is a strong evidence that the plasticity in this region, which includes increased angiogenesis, neurogenesis, and axonal sprouting, is directly linked to the observed recovery in animal models and humans5. Since there are no in vitro models that can properly replicate the complex interactions following stroke, animal models are essential for stroke research.
There are several in vivo models that can be used to produce ischemic stroke. One of the most common models used in mice is middle cerebral artery occlusion, or MCAo, through distal or proximal (via intra-arterial filament) occlusion. The proximal model, also known as filament MCAo (fMCAo), typically results in large ischemic strokes that encompass anywhere from 5% to 50% of the cerebral hemisphere, dependent upon number of factors6. In these models, a suture or filament is advanced from the internal carotid artery into the base of the middle cerebral artery (MCA) and kept in place for a specific period of time. This method for occlusion, which can be made temporary or permanent, produces an infarct that is centered in the striatum and may or may not involve overlying cortex6. The resulting stroke size is highly variable, and imaging techniques, such as laser doppler, are required to confirm the effectiveness of the procedure in each mouse. Intra-arterial or intra-luminal filament occlusion that lasts greater than 30 minutes produces strokes at the larger end of the size range. Some investigators have focused on shorter filament occlusion times, which require substantial experimental focus and lab validation7. Filament MCAo models in mice follow similar stages of cell death, ischemic progression, and formation of a peri-infarct region as seen in human stroke cases; however, the larger strokes more closely resemble the disease state of malignant cerebral infarctions, which are less common, less treatable human strokes6. Meanwhile, distal MCA occlusion requires a more involved surgery and craniectomy. In this model, the distal part of the MCA that runs along the surface of the brain is directly occluded with a suture tie or cauterization. In some variations of the technique, the carotid arteries are unilaterally or transiently-bilaterally occluded. A benefit of the distal MCAo is that it produces a cortical-based stroke that is less variable in size than the filament model. However, the distal model produces poorer behavioral output due to transection of the external carotid artery (ECA), which is also a concern with fMCAo6.
An alternative stroke model that is known for being less invasive is the photothrombotic (PT) model. The PT model results in a well-defined location of ischemia and is associated with a high survival rate8. The technique relies on a photosensitive dye injected intraperitoneally that allows for the intravascular photo-oxidation simply by irradiating the desired tissue with a light or laser9. Upon excitation, oxygen radicals are formed that cause endothelial damage, which activates platelet aggregation and clot formation in the irradiated area8,9. The tight control over stroke size and location, as well as high reproducibility of the PT model, makes it ideal for studying biomaterials. While precision is made possible using a laser and stereotaxic coordinates, there are some disadvantages that may make this model less ideal for few studies. Unlike the fMCAo model, the PT stroke model cannot be reperfused. Therefore, materials for investigating neuroprotective agents specific to damages following reperfusion or the mechanisms following reperfusion would not be useful here8. Additionally, due to the microvascular insult of the PT model, relatively small ischemic penumbra is seen. Instead, local vasogenic edema occurs, which is uncharacteristic for human stroke, making this model undesirable for preclinical drug studies focused on the peri-infarct area6,8.
The overall goal of biomaterial strategies in stroke is to either deliver bioactive agents or to act as a surrogate extracellular matrix for brain tissue growth. One strategy that we will explore using our methods is to deliver hydrogel directly into the stroke core, as opposed to the peri-infarct tissue where many current cell therapies are delivered10. The rationale for this approach is that delivery into the necrotic tissue found in the core will avoid disrupting the surrounding healthy or recovering tissue. We assume diffusion of any active agents included within the biomaterial will be able to reach the peri-infarct from the core, especially since we find that delivery of hydrogel biomaterials reduces the thickness of the glial scar11. This is important since the peri-infarct region has been shown to exhibit neuroplasticity after stroke, making it an attractive target. Furthermore, the delivery of a surrogate matrix to the stroke core can be loaded with angiogenic12 or neurogenic13 factors to guide the formation of new tissue, as well as cells for the delivery14. Cell delivery is greatly enhanced by using a matrix because it protects cells from the harsh injection forces and local environment present during delivery, as well as encourages differentiation and engraftment15.
These injectable therapeutic biomaterials have clinical relevance in stroke applications, as there are currently no medical therapies that stimulate neuronal recovery after stroke. The underlying neural circuits involved in recovery lie in brain tissue that is adjacent to the stroke core16, while the stroke core itself is devoid of viable neural tissue. We anticipate that delivering a biomaterial into the necrotic stroke core has potential to stimulate the adjacent tissue toward regenerative processes through a number of mechanisms previously mentioned, including depot release of growth factors13, stimulation of tissue in-growth and promotion of recovering brain tissue development11,12, alteration of immune responses17, and delivery of stem cell-derived therapeutics14,18. However, to effectively study the possibility of these applications, a consistent and reproducible method for inducing stroke and injecting biomaterials is needed. The PT stroke model uses techniques that offer precise control over the orientation and location of the stroke. A laser attached to the stereotaxic device guides orientations, and pumps attached to the stereotaxic devices control the injection rate of the material without the need for additional forms of imaging. Therefore, we have chosen to describe the methods for performing a PT stroke in the motor cortexes of mice and for injecting biomaterials into the stroke core. Here, we use microporous annealed particle (MAP) hydrogels as the biomaterial for injection with no added cells or growth factors. Additionally, we explain how to successfully retrieve the brain with intact biomaterial, and we discuss immunochemistry assays used to analyze the stroke outcome with and without injection of biomaterials.

<table>
	<tbody>
		<tr>
			<td>10% Normal Goat Serum</td>
			<td>VWR</td>
			<td>100504-028</td>
			<td>For blocking buffer</td>
		</tr>
		<tr>
			<td>2-ply alcohol pre pad, Sterile, Medium</td>
			<td>Medline</td>
			<td>MDS090735</td>
			<td></td>
		</tr>
		<tr>
			<td>25uL Hamilton Syringe 702RN, no needle</td>
			<td>Fishcer Scientific</td>
			<td>14824663</td>
			<td>Syringes used to inject biomaterials</td>
		</tr>
		<tr>
			<td>25uL Positive displacement pipette</td>
			<td>Gilson</td>
			<td>M-25</td>
			<td></td>
		</tr>
		<tr>
			<td>2x Beam Expander, 400-650nm</td>
			<td>Thorlabs</td>
			<td>GBE02-A</td>
			<td>Laser beam expander</td>
		</tr>
		<tr>
			<td>Adjustable Stage Platform</td>
			<td>Kopf Instruments</td>
			<td>901</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-Glucose Transporter GLUT1 antibody, rabbit</td>
			<td>Abcam</td>
			<td>ab113435</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-Iba1 Antibody, goat</td>
			<td>abcam</td>
			<td>ab5076</td>
			<td></td>
		</tr>
		<tr>
			<td>BD Vacutainer Safety-Lok Blood Collection Sets. 25G, 12&#34;</td>
			<td>Medsupply</td>
			<td>367294</td>
			<td>For perfusions</td>
		</tr>
		<tr>
			<td>BKF12- Matte Black Aluminum Foil</td>
			<td>Thorlabs</td>
			<td>BKF12</td>
			<td>To cover anything that is reflective when using laser.</td>
		</tr>
		<tr>
			<td>Bone Iris Mini Scissors - 3-1/2&#34;</td>
			<td>Sklar surgical instruments</td>
			<td>64-2035</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6 Mice</td>
			<td>Jackson Laboratory</td>
			<td>000664</td>
			<td>8-12 weeks of age</td>
		</tr>
		<tr>
			<td>Cage Assembly Rob</td>
			<td>Thorlabs</td>
			<td>ER3-P4</td>
			<td>3&#34; Long, diameter 6mm, 4 pack - for attaching laser to sterotax</td>
		</tr>
		<tr>
			<td>Carbon Steel Burrs -0.5mm Diameter</td>
			<td>Fine Science Tools</td>
			<td>19007-05</td>
			<td>For creating burr hole</td>
		</tr>
		<tr>
			<td>Chromium(III) potassium sulfate dodecahydrate</td>
			<td>VWR</td>
			<td>EM1.01036.0250</td>
			<td></td>
		</tr>
		<tr>
			<td>Compact Controller for pigtailed lasers</td>
			<td>Thorlabs</td>
			<td>CLD1010LP</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton Swabs</td>
			<td>VWR</td>
			<td>89031-288</td>
			<td></td>
		</tr>
		<tr>
			<td>CP 25 pipette tips</td>
			<td>Gilson</td>
			<td>F148012</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey anti-goat IgG H&#38;L (488)</td>
			<td>abcam</td>
			<td>ab150129</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey Anti-rabbit IgG H&#38;L (647)</td>
			<td>abcam</td>
			<td>ab150075</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey Anti-rat IgG H&#38;L (555)</td>
			<td>abcam</td>
			<td>ab150154</td>
			<td></td>
		</tr>
		<tr>
			<td>EMS DPX Mountant</td>
			<td>Elecron Microscopy Sciences</td>
			<td>13512</td>
			<td>Mounting solution for slides</td>
		</tr>
		<tr>
			<td>EMS Gelatin Powder Type A 300 Bloom</td>
			<td>Electron Microscopy Sciences</td>
			<td>16564</td>
			<td>For gelatin coating slides</td>
		</tr>
		<tr>
			<td>EMS Paraformaldehyde, Granular</td>
			<td>VWR</td>
			<td>100504-162</td>
			<td>For making 45 PFA</td>
		</tr>
		<tr>
			<td>ESD Worstation kit</td>
			<td>Elmstat</td>
			<td>WSKK5324SB</td>
			<td>Need for setting up the laser</td>
		</tr>
		<tr>
			<td>Fiber Bench Wall Plate, unthreaded</td>
			<td>Thorlabs</td>
			<td>HCA3</td>
			<td>Need for connecting laser to Kopf shaft</td>
		</tr>
		<tr>
			<td>FiberPort</td>
			<td>Thorlabs</td>
			<td>PAF-X-5-A</td>
			<td>FC/APC&#38; APC, f=4.6mm, 350-700nm, diameter 0.75mm</td>
		</tr>
		<tr>
			<td>Fine Scissors - straight/sharp-blunt/10cm</td>
			<td>Fine Science Tools</td>
			<td>14028-10</td>
			<td></td>
		</tr>
		<tr>
			<td>GFAP Antibody, rat</td>
			<td>Thermo Fisher Scientific</td>
			<td>13-0300</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating Plate</td>
			<td>Kopf Instruments</td>
			<td>HP-4M</td>
			<td></td>
		</tr>
		<tr>
			<td>ImmEdge Hydrophobic Barrier Pen</td>
			<td>Vector Laboratories</td>
			<td>H-4000</td>
			<td>For staining slides</td>
		</tr>
		<tr>
			<td>IMPAC 6-Integrated Multi Patient Anesthesia Center</td>
			<td>VetEquip</td>
			<td>901808</td>
			<td></td>
		</tr>
		<tr>
			<td>Iodine Prep Pads</td>
			<td>Medx Supple</td>
			<td>MED MDS093917H</td>
			<td></td>
		</tr>
		<tr>
			<td>Jewlers Forceps #5</td>
			<td>GFS chemicals</td>
			<td>46085</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser Safety Glasses</td>
			<td>Thorlabs</td>
			<td>LG10B</td>
			<td>Amber Lenses, 35% Visible light (googles versions available too)</td>
		</tr>
		<tr>
			<td>M27-1084 Powerful LED Dual Goose-neck</td>
			<td>United Scope</td>
			<td>LED-11C</td>
			<td></td>
		</tr>
		<tr>
			<td>Medical USP Grade Oxygen</td>
			<td>Airgas</td>
			<td>OX USP250</td>
			<td></td>
		</tr>
		<tr>
			<td>Miltex Adson Dressing Forceps, Disecting-grade</td>
			<td>Intefra Miltex</td>
			<td>V96-118</td>
			<td></td>
		</tr>
		<tr>
			<td>Mini Cord/Cordless Small Animal Trimmer</td>
			<td>Harvard aparatus</td>
			<td>72-6110</td>
			<td></td>
		</tr>
		<tr>
			<td>Mini-pump variable flow</td>
			<td>Thomas Scientific</td>
			<td>70730-064</td>
			<td>Pump for perfusions</td>
		</tr>
		<tr>
			<td>Mouse Brain Matrices, Coronal Slices, 1mm</td>
			<td>Kent Scientific</td>
			<td>RBMA-200c</td>
			<td>For TTC slices</td>
		</tr>
		<tr>
			<td>Mouse Gas Anethesia Head Holder</td>
			<td>Kopf Instruments</td>
			<td>923-B</td>
			<td></td>
		</tr>
		<tr>
			<td>Nanojet Control Box</td>
			<td>Chemyx</td>
			<td>10050</td>
			<td></td>
		</tr>
		<tr>
			<td>Nanojet pump header</td>
			<td>Chemxy</td>
			<td>10051</td>
			<td>Attach to stereotaxic device for injecting biomaterial</td>
		</tr>
		<tr>
			<td>Needle RN 30G PT STY 3, 0.5 inch</td>
			<td>Fishcer Scientific</td>
			<td>NC9459562</td>
			<td></td>
		</tr>
		<tr>
			<td>Non-rupture ear bars 60&#186;</td>
			<td>Kopf Instruments</td>
			<td>922</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS buffer pH 7.4</td>
			<td>VWR</td>
			<td>97062 338</td>
			<td></td>
		</tr>
		<tr>
			<td>Pigtaled laser 520 nm, 100mW, 5G Pin</td>
			<td>Thorlabs</td>
			<td>LP520-MF100</td>
			<td></td>
		</tr>
		<tr>
			<td>Positive charge glass slides</td>
			<td>Hareta</td>
			<td>AHS90-WH</td>
			<td></td>
		</tr>
		<tr>
			<td>Power engergy meter</td>
			<td>Thorlabs</td>
			<td>PM100D</td>
			<td>Used to measure your mW laser output</td>
		</tr>
		<tr>
			<td>Puralube Vet Ointment</td>
			<td>Dr. Foster Smith</td>
			<td>9N-76855</td>
			<td></td>
		</tr>
		<tr>
			<td>Rectal Probe Mouse</td>
			<td>kopf Instruments</td>
			<td>Ret-3-ISO</td>
			<td></td>
		</tr>
		<tr>
			<td>Rose Bengal Dye 95%</td>
			<td>Sigma-Aldrich</td>
			<td>330000-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>Shaft Modified 8-32 threaded hole 1/2&#34; depth</td>
			<td>Kopf Instruments</td>
			<td>1770-02</td>
			<td>For connecting laser to sterotaxic device</td>
		</tr>
		<tr>
			<td>Slim photodiode power sensor</td>
			<td>Thorlabs</td>
			<td>S130VC</td>
			<td>Used with power energy meter</td>
		</tr>
		<tr>
			<td>SM1-Threaed 30 mm Cage Plate 0.35&#34; thick 2 Retaining</td>
			<td>Thorlabs</td>
			<td>CP02</td>
			<td>For connecting laser expander</td>
		</tr>
		<tr>
			<td>Sol-M U-100 Insuline syringe with 1/2 unit markings 0.5 mL</td>
			<td>VWR</td>
			<td>10002-726</td>
			<td>To inject rose bengal</td>
		</tr>
		<tr>
			<td>StainTray Slide Staining System</td>
			<td>Simport Scientific</td>
			<td>M920-2</td>
			<td>For staining slides</td>
		</tr>
		<tr>
			<td>Sterotaxic device</td>
			<td>Kopf Instruments</td>
			<td>940</td>
			<td>Small Animal Stereotaxic Instrument</td>
		</tr>
		<tr>
			<td>Student Adson Forceps -1x2 teeth</td>
			<td>Fine Science Tools</td>
			<td>91127-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Student Fine Forceps - straight/broad Shanks</td>
			<td>Fine Science Tools</td>
			<td>91113-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature Controller</td>
			<td>Kopf Instruments</td>
			<td>TCAT-2LV</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue-Tek OCT compound</td>
			<td>VWR</td>
			<td>25608-930</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>VWR</td>
			<td>97063-864</td>
			<td></td>
		</tr>
		<tr>
			<td>Upper Bracket Clamp</td>
			<td>Kopf Instruments</td>
			<td>1770-c</td>
			<td>For connecting laser to sterotaxic device</td>
		</tr>
		<tr>
			<td>Vetbond Tissue Adhessive 3mL</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-361931</td>
			<td></td>
		</tr>
		<tr>
			<td>Vogue Professional My Manicurist</td>
			<td>Bargin Source</td>
			<td>6400</td>
			<td>For Burrs</td>
		</tr>
		<tr>
			<td>VWR Bead Sterilizers</td>
			<td>VWR</td>
			<td>75999-328</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Tek OCT compound</td>
			<td>Sakura</td>
			<td>4583</td>
			<td>For tissue embeding</td>
		</tr>
	</tbody>
</table>

injection of hydrogel biomaterial scaffolds to the brain after stroke

Stroke is the leading cause of disability and the fifth-leading cause of death in the United States. Approximately 87% of all strokes are ischemic strokes and are defined as the sudden blockage of a vessel supplying blood to the brain. Within minutes of the blockage, cells begin to die and result in irreparable tissue damage. Current therapeutic treatments focus on clot removal or lysis to allow for the reperfusion and prevent more severe brain damage. Although transient brain plasticity may salvage some of the damaged tissue over time, significant fractions of patients are left with neurological deficits that will never resolve. There is a lack of therapeutic options to treat neurological deficits caused by stroke, emphasizing the need to develop new strategies to treat this growing patient population. Injectable biomaterials are currently being designed to enhance brain plasticity and improve endogenous repair through the delivery of active agents or stem cells. One method to test these approaches is to utilize a rodent stroke model, inject the biomaterial into the stroke core, and assess repair. Knowing the precise location of the stroke core is imperative for the accurate treatment after stroke, therefore, a stroke model that results in a predictable stroke location is preferable to avoid the need for imaging prior to injection. The following protocol will cover how to induce a photothrombotic stroke, how to inject a hydrogel in a controlled and precise manner, and how to extract and cryosection the brain while keeping the biomaterial intact. In addition, we will highlight how these same hydrogel materials can be used for the co-delivery of stem cells. This protocol can be generalized to the use of other injectable biomaterials into the stroke core.

The aim of this method was to demonstrate how to inject biomaterials into the brain after stroke. A photothrombotic model with rose bengal and a 520 nm laser was used for controlled orientation of the stroke lesion in both size and location. Five days after stroke the infarct could be visualized during surgery (Figure 1B) and by TTC and imaging IHC stained slides (Figure 2). An increase in laser diameter with a 2x lens lead to a visual increase in the stroke les...

Watch this Scientific Journal Video about Injection of Hydrogel Biomaterial Scaffolds to The Brain After Stroke at JoVE.com

Injection of Hydrogel Biomaterial Scaffolds to The Brain After Stroke

Stroke is a global issue with minimal treatment options and no current clinical therapy for regenerating the lost brain tissue. Here we describe methods for creating precise photothrombotic stroke in the motor cortex of rodents and subsequent injection of hydrogel biomaterials to study their effects on tissue regeneration after stroke.

Stroke is the leading cause of disability and the fifth-leading cause of death in the United States. Approximately 87% of all strokes are ischemic strokes ...

injection-of-hydrogel-biomaterial-scaffolds-to-the-brain-after-stroke

Research

JoVE Journal

Biology

4.9K Views.  Duke University. Stroke is a global issue with minimal treatment options and no current clinical therapy for regenerating the lost brain tissue. Here we describe methods for creating precise photothrombotic stroke in the motor cortex of rodents and subsequent injection of hydrogel biomaterials to study their effects on tissue regeneration after stroke.

Video: Injection of Hydrogel Biomaterial Scaffolds to The Brain After Stroke

Intravitreous Injection for Establishing Ocular Diseases Model

Intravitreous injection is a widely used technique in visual sciences research. It can be used to establish animal models with ocular diseases or as direct application of local treatment. This video introduces how to use simple and inexpensive tools to finish the intravitreous injection procedure. Use of a 1 ml syringe, instead of a hemilton syringe, is used. Practical tips for how to make appropriate injection needles using glass pipettes with perfect tips, and how to easily connect the syringe needle with the glass pipette tightly together, are given.
To conduct a good intravitreous injection, there are three aspects to be observed: 1) injection site should not disrupt retina structure; 2) bleeding should be avoided to reduce the risk of infection; 3) lens should be untouched to avoid traumatic cataract. In brief, the most important point is to reduce the interruption of normal ocular structure. To avoid interruption of retina, the superior nasal region of rat eye was chosen. Also, the puncture point of the needle was at the par planar, which was about 1.5 mm from the limbal region of the rat eye. A small amount of vitreous is gently pushed out through the puncture hole to reduce the intraocular pressure before injection. With the 45&deg; injection angle, it is less likely to cause traumatic cataract in the rat eye, thus avoiding related complications and influence from lenticular factors. In this operation, there was no cutting of the conjunctiva and ocular muscle, no bleeding. With quick and minor injury, a successful intravitreous injection can be done in minutes. 
The injection set outlined in this particular protocol is specific for intravitreous injection. However, the methods and materials presented here can also be used for other injection procedures in drug delivery to the brain, spinal cord or other organs in small mammals.

Intravitreous injection is a widely used technique in visual sciences research for ocular diseases or as direct application of local treatment. This video demonstrated a protocol for intravitreous injection using a 1ml syringe with glass pipette. Useful tips about avoiding massive bleeding and lens damage are given.

Intravitreous injection is a widely used technique in visual sciences research. It can be used to establish animal models with ocular diseases or as ...

Microfabrication of Chip-sized Scaffolds for Three-dimensional Cell cultivation

Using microfabrication technologies is a prerequisite to create scaffolds of reproducible geometry and constant quality for three-dimensional cell cultivation. These technologies offer a wide spectrum of advantages not only for manufacturing but also for different applications. The size and shape of formed cell clusters can be influenced by the exact and reproducible architecture of the microfabricated scaffold and, therefore, the diffusion path length of nutrients and gases can be controlled.1 This is unquestionably a useful tool to prevent apoptosis and necrosis of cells due to an insufficient nutrient and gas supply or removal of cellular metabolites.
Our polymer chip, called CellChip, has the outer dimensions of 2 x 2 cm with a central microstructured area. This area is subdivided into an array of up to 1156 microcontainers with a typical dimension of 300 m edge length for the cubic design (cp- or cf-chip) or of 300 m diameter and depth for the round design (r-chip).2
So far, hot embossing or micro injection moulding (in combination with subsequent laborious machining of the parts) was used for the fabrication of the microstructured chips. Basically, micro injection moulding is one of the only polymer based replication techniques that, up to now, is capable for mass production of polymer microstructures.3 However, both techniques have certain unwanted limitations due to the processing of a viscous polymer melt with the generation of very thin walls or integrated through holes. In case of the CellChip, thin bottom layers are necessary to perforate the polymer and provide small pores of defined size to supply cells with culture medium e.g. by microfluidic perfusion of the containers.
In order to overcome these limitations and to reduce the manufacturing costs we have developed a new microtechnical approach on the basis of a down-scaled thermoforming process. For the manufacturing of highly porous and thin walled polymer chips, we use a combination of heavy ion irradiation, microthermoforming and track etching. In this so called "SMART" process (Substrate Modification And Replication by Thermoforming) thin polymer films are irradiated with energetic heavy projectiles of several hundred MeV introducing so-called "latent tracks" Subsequently, the film in a rubber elastic state is formed into three dimensional parts without modifying or annealing the tracks. After the forming process, selective chemical etching finally converts the tracks into cylindrical pores of adjustable diameter.

We present two processes for the microfabrication of porous polymer chips for three-dimensional cell cultivation. The first one is hot embossing combined with a solvent vapour welding process. The second one uses a recently developed microthermoforming process combined with ion track technology leading to a significant simplification of manufacture.

Using microfabrication technologies is a prerequisite to create scaffolds of reproducible geometry and constant quality for three-dimensional cell ...

Zebrafish Brain Ventricle Injection

Proper brain ventricle formation during embryonic brain development is required for normal brain function.  Brain ventricles are the highly conserved cavities within the brain that are filled with cerebrospinal fluid.  In zebrafish, after neural tube formation, the neuroepithelium undergoes a series of constrictions and folds while it fills with fluid resulting in brain ventricle formation.  In order to understand the process of ventricle formation, and the neuroepithelial shape changes that occur at the same time, we needed a way to visualize the ventricle space in comparison to the brain tissue.  However, the nature of transparent zebrafish embryos makes it difficult to differentiate the tissue from the ventricle space.  Therefore, we developed a brain ventricle injection technique where the ventricle space is filled with a fluorescent dye and imaged by brightfield and fluorescent microscopy.  The brightfield and the fluorescent images are then processed and superimposed in Photoshop.  This technique allows for visualization of the ventricle space with the fluorescent dye, in comparison to the shape of the neuroepithelium in the brightfield image.  Brain ventricle injection in zebrafish can be employed from 18 hours post fertilization through early larval stages.  We have used this technique extensively in our studies of brain ventricle formation and morphogenesis as well as in characterizing brain morphogenesis mutants (1-3).

After neural tube formation, the neuroepithelium constricts and folds while the tube fills with embryonic cerebrospinal fluid (eCSF) to form the embryonic brain ventricles. We developed this ventricle injection technique to better visualize the fluid filled space in contrast to the neuroepithelial shape in a live embryo.

Proper brain ventricle formation during embryonic brain development is required for normal brain function.  Brain ventricles are the highly conserved ...

The Gateway to the Brain: Dissecting the Primate Eye

The visual system in humans is considered the gateway to the world and plays a principal role in the plethora of sensory, perceptual and cognitive processes. It is therefore not surprising that quality of vision is tied to quality of life . Despite widespread clinical and basic research surrounding the causes of visual disorders, many forms of visual impairments, such as retinitis pigmentosa and macular degeneration, lack effective treatments. Non-human primates have the closest general features of eye development to that of humans. Not only do they have a similar vascular anatomy, but amongst other mammals, primates have the unique characteristic of having a region in the temporal retina specialized for high visual acuity, the fovea1. Here we describe a general technique for dissecting the primate retina to provide tissue for retinal histology, immunohistochemistry, laser capture microdissection, as well as light and electron microscopy. With the extended use of the non-human primate as a translational model, our hope is that improved understanding of the retina will provide insights into effective approaches towards attenuating or reversing the negative impact of visual disorders on the quality of life of affected individuals.

The non-human primate is an important translational species for our understanding of development and aging. The anatomical organization of the primate retina may provide important insights into normal and pathological conditions in humans.

The visual system in humans is considered the  gateway  to the world and plays a principal role in the plethora of sensory, perceptual and cognitive ...

Mapping the After-effects of Theta Burst Stimulation on the Human Auditory Cortex with Functional Imaging

Auditory cortex pertains to the processing of sound, which is at the basis of speech or music-related processing1. However, despite considerable recent progress, the functional properties and lateralization of the human auditory cortex are far from being fully understood. Transcranial Magnetic Stimulation (TMS) is a non-invasive technique that can transiently or lastingly modulate cortical excitability via the application of localized magnetic field pulses, and represents a unique method of exploring plasticity and connectivity. It has only recently begun to be applied to understand auditory cortical function 2. 
An important issue in using TMS is that the physiological consequences of the stimulation are difficult to establish. Although many TMS studies make the implicit assumption that the area targeted by the coil is the area affected, this need not be the case, particularly for complex cognitive functions which depend on interactions across many brain regions 3. One solution to this problem is to combine TMS with functional Magnetic resonance imaging (fMRI). The idea here is that fMRI will provide an index of changes in brain activity associated with TMS. Thus, fMRI would give an independent means of assessing which areas are affected by TMS and how they are modulated 4. In addition, fMRI allows the assessment of functional connectivity, which represents a measure of the temporal coupling between distant regions. It can thus be useful not only to measure the net activity modulation induced by TMS in given locations, but also the degree to which the network properties are affected by TMS, via any observed changes in functional connectivity.
Different approaches exist to combine TMS and functional imaging according to the temporal order of the methods. Functional MRI can be applied before, during, after, or both before and after TMS. Recently, some studies interleaved TMS and fMRI in order to provide online mapping of the functional changes induced by TMS 5-7. However, this online combination has many technical problems, including the static artifacts resulting from the presence of the TMS coil in the scanner room, or the effects of TMS pulses on the process of MR image formation. But more importantly, the loud acoustic noise induced by TMS (increased compared with standard use because of the resonance of the scanner bore) and the increased TMS coil vibrations (caused by the strong mechanical forces due to the static magnetic field of the MR scanner) constitute a crucial problem when studying auditory processing. 
This is one reason why fMRI was carried out before and after TMS in the present study. Similar approaches have been used to target the motor cortex 8,9, premotor cortex 10, primary somatosensory cortex 11,12 and language-related areas 13, but so far no combined TMS-fMRI study has investigated the auditory cortex. The purpose of this article is to provide details concerning the protocol and considerations necessary to successfully combine these two neuroscientific tools to investigate auditory processing. 
Previously we showed that repetitive TMS (rTMS) at high and low frequencies (resp. 10 Hz and 1 Hz) applied over the auditory cortex modulated response time (RT) in a melody discrimination task 2. We also showed that RT modulation was correlated with functional connectivity in the auditory network assessed using fMRI: the higher the functional connectivity between left and right auditory cortices during task performance, the higher the facilitatory effect (i.e. decreased RT) observed with rTMS. However those findings were mainly correlational, as fMRI was performed before rTMS. Here, fMRI was carried out before and immediately after TMS to provide direct measures of the functional organization of the auditory cortex, and more specifically of the plastic reorganization of the auditory neural network occurring after the neural intervention provided by TMS. 
Combined fMRI and TMS applied over the auditory cortex should enable a better understanding of brain mechanisms of auditory processing, providing physiological information about functional effects of TMS. This knowledge could be useful for many cognitive neuroscience applications, as well as for optimizing therapeutic applications of TMS, particularly in auditory-related disorders.

Auditory processing is the basis of speech and music-related processing. Transcranial Magnetic Stimulation (TMS) has been used successfully to study cognitive, sensory and motor systems but has rarely been applied to audition. Here we investigated TMS combined with functional Magnetic Resonance Imaging to understand the functional organization of auditory cortex.

Auditory cortex pertains to the processing of sound, which is at the basis of speech or music-related processing1. However, despite considerable recent ...

Intraductal Injection for Localized Drug Delivery to the Mouse Mammary Gland

Herein we describe a protocol to deliver various reagents to the mouse mammary gland via intraductal injections. Localized drug delivery and knock-down of genes within the mammary epithelium has been difficult to achieve due to the lack of appropriate targeting molecules that are independent of developmental stages such as pregnancy and lactation. Herein, we describe a technique for localized delivery of reagents to the mammary gland at any stage in adulthood via intraductal injection into the nipples of mice. The injections can be performed on live mice, under anesthesia, and allow for a non-invasive and localized drug delivery to the mammary gland. Furthermore, the injections can be repeated over several months without damaging the nipple. Vital dyes such as Evans Blue are very helpful to learn the technique. Upon intraductal injection of the blue dye, the entire ductal tree becomes visible to the eye. Furthermore, fluorescently labeled reagents also allow for visualization and distribution within the mammary gland. This technique is adaptable for a variety of compounds including siRNA, chemotherapeutic agents, and small molecules.

A protocol for the non-invasive intraductal delivery of aqueous reagents to the mouse mammary gland is described. The method takes advantage of localized injection into the nipples of mammary glands targeting mammary ducts specifically. This technique is adaptable for a variety of compounds including siRNA, chemotherapeutic agents and small molecules.

Herein we describe a protocol to deliver various reagents to the mouse mammary gland via intraductal injections. Localized drug delivery and knock-down of ...

Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow GUVs must be modified in order to form GUVs containing functional transmembrane proteins. This article describes two dehydration-rehydration methods &#8212; electroformation and gel-assisted swelling &#8212; to form GUVs containing the voltage-gated potassium channel, KvAP. In both methods, a solution of protein-containing small unilamellar vesicles is partially dehydrated to form a stack of membranes, which is then allowed to swell in a rehydration buffer. For the electroformation method, the film is deposited on platinum electrodes so that an AC field can be applied during film rehydration. In contrast, the gel-assisted swelling method uses an agarose gel substrate to enhance film rehydration. Both methods can produce GUVs in low (e.g., 5 mM) and physiological (e.g., 100 mM) salt concentrations. The resulting GUVs are characterized via fluorescence microscopy, and the function of reconstituted channels measured using the inside-out patch-clamp configuration. While swelling in the presence of an alternating electric field (electroformation) gives a high yield of defect-free GUVs, the gel-assisted swelling method produces a more homogeneous protein distribution and requires no special equipment.

The reconstitution of the transmembrane protein, KvAP, into giant unilamellar vesicles (GUVs) is demonstrated for two dehydration-rehydration methods &#8212; electroformation, and gel-assisted swelling. In both methods, small unilamellar vesicles containing the protein are fused together to form GUVs that can then be studied by fluorescence microscopy and patch-clamp electrophysiology.

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow ...

Sampling Blood from the Lateral Tail Vein of the Rat

Blood samples are commonly obtained in many experimental contexts to measure targets of interest, including hormones, immune factors, growth factors, proteins, and glucose, yet the composition of the blood is dynamically regulated and easily perturbed. One factor that can change the blood composition is the stress response triggered by the sampling procedure, which can contribute to variability in the measures of interest. Here we describe a procedure for blood sampling from the lateral tail vein in the rat. This procedure offers significant advantages over other more commonly used techniques. It permits rapid sampling with minimal pain or invasiveness, without anesthesia or analgesia. Additionally, it can be used to obtain large volume samples (upwards of 1 ml in some rats), and it may be used repeatedly across experimental days. By minimizing the stress response and pain resulting from blood sampling, measures can more accurately reflect the true basal state of the animal, with minimal influence from the sampling procedure itself.

Blood samples are useful for assessing biomarkers of physiological states or disease in vivo. Here we describe the methodology to sample blood from the lateral tail vein in the rat. This method provides rapid samples with minimal pain and invasiveness.

Blood samples are commonly obtained in many experimental contexts to measure targets of interest, including hormones, immune factors, growth factors, ...

Adipo-Clear: A Tissue Clearing Method for Three-Dimensional Imaging of Adipose Tissue

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte precursors, immune cells, fibroblasts, blood vessels, and nerve projections. Although the molecular control of cell type specification and how these cells interact have been increasingly delineated, a more comprehensive understanding of these adipose-resident cells can be achieved by visualizing their distribution and architecture throughout the whole tissue. Existing immunohistochemistry and immunofluorescence approaches to analyze adipose histology rely on thin paraffin-embedded sections. However, thin sections capture only a small portion of tissue; as a result, the conclusions can be biased by what portion of tissue is analyzed. We have therefore developed an adipose tissue clearing technique, Adipo-Clear, to permit comprehensive three-dimensional visualization of molecular and cellular patterns in whole adipose tissues. Adipo-Clear was adapted from iDISCO/iDISCO+, with specific modifications made to completely remove the lipid stored in the tissue while preserving native tissue morphology. In combination with light-sheet fluorescence microscopy, we demonstrate here the use of the Adipo-Clear method to obtain high-resolution volumetric images&#160;of an entire adipose tissue.

Due to the high lipid content, adipose tissue has been challenging to visualize using traditional histological methods. Adipo-Clear is a tissue clearing technique that allows robust labeling and high-resolution volumetric fluorescent imaging of adipose tissue. Here, we describe the methods for sample preparation, pretreatment, staining, clearing, and mounting for imaging.

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte ...

A High Output Method to Isolate Cerebral Pericytes from Mouse

In recent years, cerebral pericytes have become the focus of extensive research in vascular biology and pathology. The importance of pericytes in blood brain barrier formation and physiology is now demonstrated but its molecular basis remains largely unknown. As the pathophysiological role of cerebral pericytes in neurological disorders is intriguing and of great importance, the in vitro models are not only sufficiently appropriate but also able to incorporate different techniques for these studies. Several methods have been proposed as in vitro models for the extraction of cerebral pericytes, although an antibiotic-free protocol with high output is desirable. Most importantly, a method that has increased output per extraction reduces the usage of more animals.
Here, we propose a simple and efficient method for extracting cerebral pericytes with sufficiently high output. The mouse brain tissue homogenate is mixed with a BSA-dextran solution for the separation of the tissue debris and microvascular pellet. We propose a three-step separation followed by filtration to obtain a microvessel rich filtrate. With this method, the quantity of microvascular fragments obtained from 10 mice is sufficient to seed 9 wells (9.6 cm2 each) of a 6-well plate. Most interestingly with this protocol, the user can obtain 27 pericyte rich wells (9.6 cm2 each) in passage 2. The purity of the pericyte cultures are confirmed with the expression of classical pericyte markers: NG2, PDGFR-&#946; and CD146. This method demonstrates an efficient and feasible in vitro tool for physiological and pathophysiological studies on pericytes.

We present a protocol for the extraction of murine cerebral pericytes. Based on an antibiotic-free enrichment oriented pericyte extraction, this protocol is a valuable tool for in vitro studies providing high purity and high yield, thus decreasing the number of experimental animals used.

In recent years, cerebral pericytes have become the focus of extensive research in vascular biology and pathology. The importance of pericytes in blood ...