Injection of Hydrogel Biomaterial Scaffolds to The Brain After Stroke

Summary

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.

Abstract

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.

Introduction

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 “peri-infarct” 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 area^8,9. 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 area^6,8.

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 development^11,12, alteration of immune responses¹⁷, and delivery of stem cell-derived therapeutics^14,18. 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.

Protocol

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 ± 2 °C), with a 12 h light-dark cycle period and access to pelleted food and water 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 occurring polymer within the brain. Animals should be monitored daily, including weekends and holidays, and reweighed every other day. Studies from our laboratory and others' 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 be euthanized. Animals should also be monitored for poor grooming, wound condition, and evidence of fighting between animals (back wounds). If there is evidence of fighting or wound dehiscence, animals should 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 euthanatized.  

1. Preparation of materials and instruments

1. 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.
   1. Dissolve the Rose Bengal in 1.5 mL of sterile filtered 1x PBS to make a working concentration of 10 mg/mL.
   2. 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 µL/g, e.g., 200 µL for a 20 g mouse.
2. Turn on the heating pad to maintain the mouse body temperature during anesthesia (37 °C).
3. 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.
4. Turn on the bead sterilization to 160 °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.
5. 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.
   1. 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.
6. 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.
7. 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.
8. 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.

2. Photothrombotic stroke model⁹

1. Shave the fur off the mouse’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.
2. 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.
3. 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).
4. Wearing laser safety goggles, move the laser over the skull of the mouse, fixing the stereotaxic device at a 90° angle. Select the 10 mW pre-set and turn on the laser.
   1. Adjust the laser’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.
   2. Move the laser to 2.2 mm left of the bregma to ensure that it can move without any obstructions. Turn off the laser.
5. 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.
6. Turn on the laser (40 mW) when the 7 min timer goes off, while wearing laser safety goggles, and set a 10 min timer.
   1. After 10 min, move the laser to 2.2 mm left of the bregma without turning it off, and set another 10 min timer.
   2. 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.
   3. Lift the laser and unlock it from the 90° 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.
7. 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.
   1. Use cotton to absorb any excess blood. It is typical to see some blood from nicked arteries in the skull after drilling – excessive blood means the drill bit hit the brain. If this happens, record the mouse identifier, and move on.
8. 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.
   1. 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.
   2. 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 – do not squeeze the bottle. Avoid gluing the skin to the skull or getting glue in the eye.
9. 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.

3. Sham stroke operation

1. Perform all procedures identically to the operation described above in section 2 except inject 1x PBS or saline instead of Rose Bengal dye.

4. Injection of hydrogel or other biomaterials

1. 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 µL of hydrogel are injected (user-defined).
   1. Turn on the heating pad to maintain the mouse body temperature during anesthesia (37 °C).
   2. 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 µL glass Hamilton syringes with metal 30 G needles.
   3. Turn on the bead sterilization to 160 °C to allow for sterilization of instruments between mice.
   4. Attach the injection pumps to the stereotaxic device. Set the flow rate to 1 µL/min and set volume to 4 µL.
   5. Prepare a clean cage for the mice after surgery.
2. 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.
   1. Once asleep, transfer and secure the mouse to the stereotaxic device with a maintenance isoflurane concentration of 1.5%.
   2. Apply the vet eye ointment to both eyes.
   3. Shave any fur that may have regrown of the mouse’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.
3. 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.
4. Orient the injection pump over the mouse and lock at 90°. Clean the syringe with sterile saline or PBS solution before backloading material.
   1. Once clean, disassemble the glass syringe so that there is no needle. Backload the syringe using a 25 µL positive displacement pipette with the 10 µL of hydrogel (or other biomaterial here with or without cells).
   2. 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.
5. 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 µL/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.
   1. 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 µL/min and is set to inject.
6. 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.
7. 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.
8. Press Start on the pump console to inject 4 - 6 µL of the hydrogel at a rate of 1 µL/min.
9. Set a 5 min timer when the pump stops injecting to allow the gel to begin crosslinking.
10. 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.
11. 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 – 10 min for the mouse to recover from anesthesia.

5. Perfusion and sample collection

1. Anesthetize the mouse using isoflurane.
2. 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.
3. 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.
4. Once clear, switch to 4% PFA for another 10 – 20 mL. Unpinning the arms allows them to contract as the PFA infuses. Once they stop moving, wait for another 30 s.
5. 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.
6. Once the brain is detached, place in 10 – 15 mL of 4% PFA overnight (no longer than 24 h).
7. 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 – 3 days). It can also be left for storage at this point.

6. Cryosectioning and staining

1. 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.
   1. 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.
2. Cut the brains serially on a cryostat at -20 °C at 30 µm 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 °C until further use.
3. Take slides out of the -80 °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.
4. 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 – 12 brain sections.
5. Wash the slides with 1x filtered PBS for 5 min at room temperature, repeat three times.
6. 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 µL per slide.
7. Incubate the primary antibody in 10% donkey serum with 0.01% PBS-Triton X overnight at 4°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.
8. Wash slides with 1x filtered PBS the following day for 5 min at room temperature, repeat three times.
9. 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.
10. Wash the slides for 5 min with 1x filtered PBS.
11. 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.
12. Dehydrate slides in ascending concentrations of alcohol (1 min per solution) of 50%, 70%, 95%, 100%, and 100%.
13. De-fat in first bath of xylene for 1 min, then second bath of xylene for 5 min.
14. 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.
15. 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.
16. 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 °C until imaging is finished.

Results

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...

Discussion

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...

Materials

List of materials used in this article

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