Modified Roller Tube Method for Precisely Localized and Repetitive Intermittent Imaging During Long-term Culture of Brain Slices in an Enclosed System

Summary

Presented here is a modified roller tube method for culturing and intermittent high-resolution imaging of rodent brain slices over many weeks with precise repositioning on photoetched coverslips. Neuronal viability and slice morphology are well maintained. Applications of this fully enclosed system using viruses for cell-type specific expression are provided.

Abstract

Cultured rodent brain slices are useful for studying the cellular and molecular behavior of neurons and glia in an environment that maintains many of their normal in vivo interactions. Slices obtained from a variety of transgenic mouse lines or use of viral vectors for expression of fluorescently tagged proteins or reporters in wild type brain slices allow for high-resolution imaging by fluorescence microscopy. Although several methods have been developed for imaging brain slices, combining slice culture with the ability to perform repetitive high-resolution imaging of specific cells in live slices over long time periods has posed problems. This is especially true when viral vectors are used for expression of exogenous proteins since this is best done in a closed system to protect users and prevent cross contamination. Simple modifications made to the roller tube brain slice culture method that allow for repetitive high-resolution imaging of slices over many weeks in an enclosed system are reported. Culturing slices on photoetched coverslips permits the use of fiducial marks to rapidly and precisely reposition the stage to image the identical field over time before and after different treatments. Examples are shown for the use of this method combined with specific neuronal staining and expression to observe changes in hippocampal slice architecture, viral-mediated neuronal expression of fluorescent proteins, and the development of cofilin pathology, which was previously observed in the hippocampus of Alzheimer's disease (AD) in response to slice treatment with oligomers of amyloid-β (Aβ) peptide.

Introduction

Primary culture of dissociated neurons from regions of rodent brain is an important tool used by researchers to observe responses to pathologically implicated stimuli. However, such studies have the disadvantage of looking at neurons in only 2D and without their glial support system. Furthermore, unless grown under conditions of very high density (640 neurons/mm² or about 16% of surface area) in which it becomes impossible to follow the random outgrowth of a dendrite or axon for more than a short distance from its cell body, hippocampal neuronal viability over 4 weeks declines significantly¹, limiting the use of dissociated cultures for extended studies of age-related pathologies. The culturing of slices prepared from rodent brain is an attractive option that overcomes these limitations by maintaining an organized cell architecture and viability for weeks or months. Conditions for maintaining many different regions of rodent brain in slice culture have been described².

Two major methods are widely used for long-term culture of brain slices: culturing on membranes at the air-liquid interface³ or culturing on coverslips in sealed tubes allowed to rotate in a roller incubator to provide aeration⁴. Slices cultured on membranes can be directly imaged with high-resolution fluorescence microscopy using an upright microscope and water immersion objectives⁵. Alternatively, slices cultured on membranes have been transferred to glass bottom dishes to achieve good resolution of dendritic spines using an inverted microscope⁶. However, both methods of imaging slices grown on membranes are open systems that require medium changes and often use antifungal and/or antibiotics to prevent or reduce contamination^5,6. Slices on a membrane at the air-medium interface maintain excellent morphology and survival, but returning to precise locations during repetitive imaging at high magnification is extremely difficult unless the experiment is following only small groups of cells expressing a fluorescent marker. Although slices grown on membranes have been used with viral-mediated expression of transgenes^5,6, biosafety protocols may require an enclosed culture system be employed for certain viral vectors that are used for expressing fluorescently tagged proteins and reporters of cell physiology. Furthermore, immersion objectives require decontamination between samples that will be followed in culture⁵. One major application of membrane interface cultures is combining high-resolution imaging with electrophysiology at single time points⁷.

The roller tube method with coverslips inside the plastic tube does not permit any electrophysiology or high-resolution imaging without removing the coverslip. Thus, this method has been most often applied to long-term studies in which post-fixation observations have been made⁸. Described here is a method that utilizes the roller tube culture technique but on drilled-out tubes with slices on coverslips that can be imaged repetitively for as long as the cultures are maintained. The enclosed system requires no medium change for imaging and utilizes photoetched coverslips to provide fiducial marks that allow imaging at high magnification, after days or weeks, the precise fields previously imaged.

We apply this method to examine changes in the rodent hippocampus, a major brain region involved in memory and learning. The rodent hippocampus is often studied as a model for pathological or age-related changes observed during development of cognitive impairment⁹, such as those that occur in AD. Our method is particularly well suited to study pathological changes that develop within a single slice over time in response to environmental changes, such as increases in Aβ peptides, which is characteristic of AD⁸. One pathology associated with human and rodent AD brain is the presence of cofilin-actin aggregates and rods, the latter containing bundles of filaments in which cofilin and actin are in a 1:1 molar ratio^10,11,12. Rods have been observed in fixed slices of rat hippocampus following Aβ treatment, as well as within a live rodent brain slice expressing cofilin-GFP subjected to hypoxia⁸, and they may contribute to the synaptic dysfunction seen in AD and stroke. Here we use this new culturing method to observe the time course and distribution within slices of expressed exogenous chimeric fluorescent proteins introduced by different viruses. We then utilize the neuronal specific expression of a cofilin reporter construct to follow the development of cofilin rod and aggregate pathology in hippocampal slices in response to treatment with soluble Aβ oligomers (Aβ_o).

Protocol

Animal use follows approved breeding and animal use protocols that conform to the Animal Care and Use Guidelines of Colorado State University.

NOTE: The protocol below describes the preparation and culture method for the long-term incubation and intermittent imaging of hippocampal slices. A single hippocampal slice is attached to a specially prepared photoetched coverslip using a plasma clot, and then the coverslips are sealed onto the flat side of a drilled-out roller tube, which is maintained in a roller incubator. Plasma clots are dissolved with plasmin before viral infection for fluorescent protein expression and high-resolution imaging. A fluorescent neuronal vital dye is used to image neurons within the slices.

1. Preparation of Roller Tube Rack

1. Use the template shown in Figure 1A printed to the size shown on the scale bar. With a nail, punch small holes (large enough for a fine point marker) in the template centered on the holes.
2. Set the template on the bottom of a 15 cm tissue culture dish (nominal diameter of 14 cm) and mark the position of the holes. Repeat this on a second dish.
3. With drill bits designed for use on plastic, drill six 1.5 cm diameter holes on each dish in a hexagonal array (4.8 cm center-to-center) with hole centers 2.5 cm from the edge of the dish. Drill three holes (3 mm in diameter) 12 mm from the edge that are placed equidistantly between two of the larger holes as shown in Figure 1A.
4. With the bottoms of each dish facing each other, place a 2.5 inch long machine screw (3/16 inch diameter) with a flat washer through one of the small holes followed by a second flat washer, a piece of polyethylene tubing (spacer, 4.7 cm), another flat washer, the second tissue culture dish, another flat washer, a locking washer, and a nut.
5. Repeat step 1.4 on the other two machine screws, and tightening only loosely until all machine screws are in place. Then tighten the nuts securely.
6. Work the grommets (5/16 inch thick, 5/8 inch hole diameter) into the holes of the bottom dish to obtain the final roller tube rack (Figure 1B; shown with two tubes in place). Place a sticker on each rack with a unique number.

2. Preparation of Roller Tubes and Coverslips

1. Making the jig for drilling the hole in roller tubes
   1. Drill a 1.5 cm hole 8 cm deep in the center side of a 2 x 4 x 5.5 inch wooden block at an angle such that the flat side of the roller tube will be nearly parallel with the block when inserted (Figure 2A).
   2. Enlarge the hole using a round wood file to both widen and taper the hole to allow tube insertion (roller tubes are slightly larger in diameter near the cap end) (Figure 2B).
   3. Drill a 1.5 cm diameter vertical hole, 5.5 cm from the side of the block, and centered over the side hole (Figure 2C).
   4. When the side hole is tapered enough, insert a roller tube that is marked at the desired spot to center the hole for the slice and position the tube so the marked spot is centered in the 1.5 cm vertical hole.
   5. Remove the tube and measure the distance from the spot to the end of the tube. Mark this distance from the center of the hole in the jig and insert a nail to provide a stop to correctly position the tube for drilling (arrow in Figure 2C).
   6. Use a hacksaw to cut the nail off flush with the surface of the wood block to prevent injury.
   7. Add spring clips on the bottom of the jig if there is a drill press with slots that allow it to be anchored (Figure 2D black arrow). Otherwise, use C-clamps to hold the jig securely onto the drill press.
2. Using the jig described above to hold and position a flat sided 11 cm plastic culture tube with the flat side up (Figure 2A), drill a 6 mm diameter hole with the center 1.0 cm from the bottom and centered between the sides of the tube.
   NOTE: A drill bit designed for plastic should be used.
3. With a swiveling deburring tool, smooth the edges of the hole (Figure 2E) and make 4 grooves on the inside edge of the hole (Figure 2E, inset) to facilitate draining of the hole during rotation.
4. With a 12 mm hole punch, cut 12 mm diameter disks from non-toxic double sided adhesive silicon rubber sheets. Using a standard one hole paper punch (6 mm diameter), make a hole in the center of each disk.
5. Rinse the drilled tubes with 70% ethanol, air dry them in a biological safety cabinet, and sterilize the tubes and the punched adhesive discs for 40 min under the UV lamp (30 W at 70 cm average distance) in the biological safety cabinet.
6. Reposition the tubes and discs after 20 min so that all exposed surfaces are sterilized. Under sterile conditions, peel off the white backing from an adhesive disc and affix the silicone rubber to the outside of a tube, aligning the holes (Figure 2F).
   Caution: To avoid UV exposure, wear eye protection and close the cabinet before turning on the UV lamp.
7. Clean 12 mm diameter photoetched (100 center numbered 1 mm squares) German glass coverslips. Hold the coverslips gently with forceps and dip in absolute ethanol, followed by water, followed by absolute ethanol again, and finally dip the coverslips in a flame to burn off the ethanol. Allow coverslips to cool.
8. Holding the coverslips with forceps, dip into 2% 3-aminopropyltriethoxysilane in acetone for 10 s. Rinse the coverslips with ultrapure water and allow to air dry.
9. Set the coverslips on sterile filter paper inside of a biological safety cabinet and turn on the UV light. Expose each side of the coverslips for 20 min.
   Caution: To avoid UV exposure, wear eye protection and close the cabinet before turning on the UV lamp.

3. Hippocampal Slice Preparation

1. Before starting the dissection, prepare halves of double-edged razor blades for the tissue chopper. Fold the blades lengthwise carefully with fingers and snap in half.
2. Rinse the blade halves with acetone using a cotton swab to clean them, followed by rinsing in absolute ethanol and air drying. Before mounting a half blade on the tissue chopper, sterilize it by rinsing with 70% ethanol.
3. Follow protocols approved by the institutional Animal Care and Use Committee; after isoflurane anesthesia, euthanize a 4-7 day old mouse or rat pup by decapitation and remove the head with scissors or guillotine.
   NOTE: The brain slice culture protocol is independent of mouse or rat strain or genotype. Many transgenic mouse lines with different genetic backgrounds have been used.
4. Rinse the head with 70% ethanol, and place it in a 60 mm Petri dish. Until the final mounting of the brain slice onto the roller tube, all of the following steps are performed in a laminar flow hood to maintain sterility.
5. Using a #21 surgical blade, make a sagittal cut through the skin and skull. With a #5 Dumont forceps, peel back the skin and skull to expose the brain.
6. With closed forceps, gently tease out the whole brain, releasing it by pinching with the forceps through the brain stem behind the cerebellum. Place the brain in a sterile 60 mm dish containing 4 °C Gey's Balanced Salt Solution/0.5% glucose (GBSS/glucose).
7. Using a dissection microscope to visualize the brain (Figure 3A), place the brain dorsal side up and cut off the front third of the brain and the cerebellum with a surgical blade (Figure 3B).
   1. With forceps, hold the trimmed brain posterior side up and ventral side against the side of the Petri dish for stability. Gently tease away meninges around the sagittal midline and remove the midbrain tissue using a fine tipped Dumont #5 forceps (Figure 3C, dashed circle).
   2. Make two cuts along the side of the brain to spread it open (Figure 3C, dashed lines). Once the brain is placed dorsal side down and spread open, the hippocampal fissure should be visible (Figure 3D, arrow).
   3. Transfer the spread open brain to a piece of polychlorotrifluoroethylene plastic film and position it for slicing on the stage of a tissue chopper. Wet the blade with GBSS/glucose and chop the hippocampus into ~300 µm thick slices.
   4. With a transfer pipette, flush the sliced brain off the plastic film into a fresh 60 mm dish containing GBSS/glucose (Figure 3E). Gently pinch off and tease away, with fine tipped forceps, the remaining meninges and other non-hippocampal tissue (Figure 3F) from the slices (Figure 3G, H).

4. Plating Slices

1. Once slices have been obtained, place 2 µL of chicken plasma on the center of the photoetched side of a prepared coverslip. Spread the plasma slightly to achieve a 3-4 mm diameter spot.
   NOTE: The photoetched side is the top side of the coverslip when viewed through a dissection microscope such that the numbers are oriented correctly.
2. Transfer 1 brain slice with a sterile narrow-tip spatula (Figure 4A) to the plasma spot (Figure 4B). Use closed forceps to keep the slice on the spatula tip while lifting the slice from the GBSS/glucose.
3. Touch the spatula to the plasma spot on the coverslip, and with closed forceps, push the slice onto the coverslip.
4. Mix 2.5 µL of plasma with 2.5 µL of thrombin in a separate tube. Quickly place 2.5 µL of this mixture over and around the slice and pipet up and down gently to mix it (Figure 4B).
   NOTE: The plasma will clot within 10-15 s, so this must be done quickly. If slice adhesion is a problem, mix 5 µL plasma with 5 µL thrombin and use  4-5 µL on the slice, removing some after mixing so that the slice lies flat on the coverslip.
5. Remove the clear plastic covering from the exposed side of the silicone rubber adhesive previously affixed to a roller tube and place the coverslip with the brain slice onto the adhesive aligning the slice within the hole (Figure 4C).
6. To ensure adhesion, apply soft, even pressure to the coverslip with the thumb by pressing the coverslip down evenly and holding it for about 1 min while transferring it to the biological safety cabinet.
7. In a biological safety cabinet, add 0.8 mL of complete Neurobasal A culture medium (Table of Materials) to each tube (Figure 4D).
8. Flow a 5% CO₂/95% air mixture through a sterile cotton-plugged Pasteur pipette held securely by a clamp. Flush the roller tube with the gas mixture and rapidly cap the tube as it is withdrawn from around the pipette.
9. Label the tubes with the slice number and rack number. Insert tubes into a roller rack, ensuring they are geometrically balanced. If there is an odd number of tubes, add tubes to balance.
10. Place the racks in a 35 °C roller incubator with rollers turning the roller rack at about 10-13 RPH (Figure 4E). To keep the medium in the bottom of the tubes, tilt the incubator back approximately 5° by raising its front on a board.
11. Enter the slice and tube number onto a spreadsheet, which is used to record all information of slice treatments and observation dates.
12. On approximately day 6 in culture, add 1 µL (0.002 U) of active plasmin to each tube.
13. After the clots dissolve completely (usually within a few hours), remove the medium and replace it with fresh medium without plasmin. If necessary, slices can be incubated with plasmin overnight and the medium changed the next day.
14. Slices are usually incubated for at least 7-10 days before use in experiments. Aspirate medium and replace it on day 3 or 4, again on day 7, and every 7 days thereafter.

5. Preparation of Viral Vectors for Transgene Expression

NOTE: Expression of transgenes in neurons of slice cultures is achieved either by using brains from genetically engineered rodents or by introducing the transgene by infection with recombinant replication deficient viruses. Adenoviruses (AV), adeno-associated viruses (AAV), and recombinant lentivirus vectors have all been used in our hippocampal slice cultures for expression of different fluorescent protein chimeras in brain slices.

1. Prepare replication deficient AV for expressing the RNA of interest according to methods described elsewhere^13,14. Titer the viruses for infectious U/mL by the serial dilution method using an antibody to a virally expressed protein as described¹⁴. To observe cofilin aggregates and cofilin-actin rod formation, utilize the cofilin-R21Q-mRFP cDNA (plasmid #51279)¹⁶.
   NOTE: The synapsin 1 promoter is an excellent choice for neuronal specific expression¹⁵, whereas the cytomegalovirus promoter is useful for driving high expression levels in many cell types¹³.
2. Prepare AAV by co-transfection of transfer plasmid containing the gene of interest and a rep/cap plasmid, with or without a helper plasmid, into HEK293 packaging cells, which supply the viral E1 gene, as previously described^17,18.
   NOTE: Recombinant AAV can also be made for targeted insertion into the host cell genome¹⁹. For the transfer plasmid, we use human cofilin 1 with a C-terminal mRFP1 fluorescence protein tag (plasmid #50856) cloned into a synapsin promoter-containing AAV plasmid downstream from the calcium sensor GCaMP5G²⁰. A piece of DNA encoding the P2A self-cleaving peptide sequence is inserted by PCR between GCaMP5G and cofilin-RFP during the preparation of the transfer plasmid to provide expression of both proteins from a single AAV transcript²¹.
3. Prepare recombinant lentivirus vectors by co-transfection of transfer plasmid containing the gene or cDNA of interest and integration signals, along with a third-generation lentivirus packaging mixture that divides the viral gag, pol, rev, and vsv-g genes onto three separate plasmids^22,23.
4. For the transfer plasmid, use a single step cloning system²⁴ to assemble the synapsin promoter and cofilin-R21Q-mRFP cDNA (from plasmid #51279) into pLKO.1-GFP (plasmid #30323), with the synapsin promoter and cofilin-R21Q-mRFP replacing the hPGK promoter and GFP cDNA, respectively.
5. Transfect the final plasmid into HEK293T cells by calcium phosphate as previously described²³.
6. Collect medium from four 10 cm dishes, concentrate to 500 µL using 150K-cutoff centrifugal concentrators, and store the final lentivirus at -80 °C in small aliquots after quick freezing in liquid nitrogen. Thaw an aliquot only once for infecting cells.
7. Determine empirically the volume of each virus type prepared to achieve the degree of expression desired by setting up a number of different slice cultures to follow the expression of the transgenes after infection with various volumes of virus.
   NOTE: Typically, 1-10 µL of virus is used per slice.

6. Slice Treatments

1. Infecting slices with virus
   1. Working in a biological safety cabinet approved for virus work at the biological safety level appropriate for the vector, mix an aliquot of the virus (usually 1-10 µL) with 0.8 mL of complete medium.
   2. Aspirate the medium from the slice using a sterile Pasteur pipette into a collection trap containing bleach. A secondary trap is always used between the first trap and the vacuum source.
   3. Replace this medium with the virus-containing aliquot prepared above, return the culture tubes to a rack, and place in the incubator.
   4. After 2-5 days of incubating the slices with virus in the biological safety cabinet, remove the virus-containing medium with a sterile transfer pipette and place it into a bottle containing an approved antiviral agent to kill virus.
2. Staining of neurons with vital dye
   1. Prepare and store aliquots of fluorescent neuronal vital dye²⁵ by quick freezing 4 µL of aliquots in liquid nitrogen at a concentration of 100 µM and store these at -20 °C. Do not freeze/thaw the dye more than once.
   2. To label neurons for visualization by fluorescence microscopy, thaw one aliquot of the neuronal vital dye and dilute to 4 mL in complete Neurobasal medium (final dye concentration is 100 nM).
   3. Remove the medium from slices by aspiration (or with a transfer pipette if the medium contains virus) and replace it with 0.8 mL of the medium containing 100 nM neuronal vital dye. Return the slices to the roller apparatus in the incubator.
   4. After incubating the slices for 2 h, aspirate the dye-containing medium and replace it with 0.8 mL of fresh complete medium in a biological safety cabinet.
      NOTE: Labeling of neurons in slices with vital dye requires several hours of incubation. The first images are usually taken 24 h after dye treatment. Although neurons are specifically labeled, there is background fluorescence that declines over 2-3 days to give better neuronal imaging. Intensity of the vital dye declines after 72 h.
   5. To follow changes in slice morphology over time, relabel the slices every 7 days.

7. Slice Imaging

1. View slices on an inverted microscope. For brightest fluorescence imaging, at 24 h before imaging, exchange culture medium with complete Neurobasal A medium without the Phenol Red pH indicator.
   NOTE: For experiments reported here, slices are viewed on an inverted spinning disc confocal fluorescence microscope equipped with a linear encoded x, y stage with piezo z control and a sensitive high-resolution digital camera.
2. Transfer the tube with the slice culture to be imaged from the roller tube apparatus to the custom-made tube holder (Figure 5A), which is placed in the stage adapter (Figure 5B), to keep the coverslip perpendicular to the objective and maintain the slice in the same orientation during repetitive imaging sessions over long intervals.
3. Push the slider on the stage adapter tight against the tube to hold the tube in position (Figure 5B). Maintain slice temperature at 35 °C during imaging by use of heating strips and a thermoregulatory controller built into the custom-made stage adapter (Figure 5C).
   NOTE: Details of the tube holder, stage adapter, and heater can be accessed at: https://vpr.colostate.edu/min/custom-machining/roller-tube-holder/.
4. Using a low power (e.g., 4X) objective and bright field transillumination, focus on the photoetched grid pattern (Figure 6A) under the slice. For the initial imaging session, quickly scan around the slice to locate and record the number for the various regions where higher magnification imaging is desired (e.g., cornu ammonis (CA)1, CA3, dentate gyrus (DG), etc.).
5. Move the stage to the first grid square containing a region of interest. Switch to the 20X air objective and locate a fiducial mark (e.g., tip of an etched number) (Figure 6B).
6. Switch to a higher power objective (40X or 60X), localize the fiducial mark, and then record the x and y position of the stage. Move the stage to find the field(s) of interest nearby and record their x and y offsets from the fiducial mark (Figure 6B, arrow). Repeat in other grid areas if desired.
   NOTE: These off-set positions from the fiducial mark allow consistent pinpointing of the same coverslip location when the slice is imaged, even though the original x, y setting of the fiducial mark changes when the tube or stage adapter are removed and replaced.
7. Capture an image Z-stack within each selected field using either the microscope objective control or a piezo stage control, if available.
   NOTE: Image planes are usually obtained at intervals of 0.5 µm to 2 µm, depending on the size and desired resolution of the image features. Building a quality 3D image requires that image features extend over multiple planes of acquisition, and so to visualize smaller features, smaller intervals are required between planes.
8. Keep the total imaging time for each slice as short as possible.
   NOTE: Most imaging sessions reported here were under 18 min/slice. However, we have imaged some slices ten or more times, and even as long as 40 min in a single session, without apparent harm in the long-term survival of the slice.

Results

To determine how accurately fiducial marks can be utilized to reimage the same cells within the same fields over time, we examined slices grown on photoetched coverslips (Figure 6A). Neurons were visualized by staining with a vital dye (100 nM for 2 h; does not stain non-neuronal cells), which disappears from neurons over time without harming the cells²⁵. We identified a fiducial mark in a single grid square (

Discussion

The roller tube method described here allows for long-term culturing and high-resolution live imaging of sliced brain tissue. One major issue with the slice technique as applied here is in the mounting and maintenance of slices. Coverslip coatings that support slice adhesion, promote slice thinning by enhancing the outgrowth of neurites and migration of cells out of the slice; thus, we avoided the use of these substrates. The insertion of amino groups onto the glass by treatment with 3-aminopropyltriethoxysilane improved...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Bottoms from 15 cm culture dishes	VWR Scientific	25384-326
Phillips Head Machine Screws (#10-32)	Ace Hardware		2.5" long and 3/16" in diameter
Flat Washers #10	ACE Hardware
Machine Screw Nuts (#10-32)	ACE Hardware
Rubber Grommets	ACE Hardware		5/16", thick; 5/8", hole diameter; 1.125", OD
Polyethylene tubing (5/16"; OD, 3/16"; ID)	ACE Hardware		Cut to 1.8" length
Lock Washer #10	ACE Hardware
Drill Press, 5 speed	Ace Hardware	ProTech Model 1201
Nunclon Delta Flat-Sided Tubes	VWR	62407-076
Drill bits, 3 mm, 6 mm and 15 mm	Ace Hardware	Diablo freud brand	Drill bits for cutting plastic.
Drill bits for wood, 1.5 cm and 1 mm	Ace Hardware
Wood file, 1/4" round	Ace Harware
Spring clips, 16 mm snap holder	Ace Hardware
Swivel Head Deburring Tool, 5"	Ace Hardware	26307
Adhesive Silicone Sheet (Secure Seal)	Grace Bio-Labs	666581	0.5 mm Thickness
6 mm hole punch	Office Max
12 mm hole punch	thepunchbunch.com
70% Ethanol
Phototeched Coverslips, 12 mm diameter	Bellco Glass, Inc.	1916-91012
Bunsen Burner
Absolute Ethanol
Nanopure Water
3-aminopropyltriethoxylane	Sigma-Aldrich	A3648
Acetone	Sigma-Aldrich	179124
#5 Dumont Forceps	Fine Science Tools	11251-30
McIlwain Tissue Chopper	Ted Pella, Inc.	10180
Double Edge Razor Blades	Ted Pella, Inc.	121-6
Whatman Filter Paper	VWR	28450-182	Cut into 5.8 cm diameter circles
Poly-chloro-trifluoro-ethylene (Aclar)	Ted Pella, Inc.	10501-10	Cut into 5.8 cm diameter circles
#21 Surgical Blade	VWR Scientific	25860-144
#5 Dumont Forceps	Fine Science Tools	11251-30
Spatula, stainless with tapered end	VWR	82027-518
Gey's Balanced Salt Solution	Sigma-Aldrich	G9779
Glucose	ThermoFisher Scientific	15023-021	25% (w/v) Solution, 0.2 mm filter sterilized
Chicken Plasma	Cocalico Biologicals	30-0300-5L	Rehydrate in sterile water, centrifuge at 2500 x g 30 min at 4 °C, quick freeze aliquots in liquid nitrogen and store at  -80 °C.
Thrombin, Topical (Bovine)	Pfizer	Thrombin-JMI	Quick freeze aliquots in liquid nitrogen at 1,000 international units/mL in diluent provided and store at -80°C. Use at 250 units/mL.
Cell Roller System	Bellco Biotech	SciERA
Roller Incubator	Forma	Model 3956
N21-MAX	ThermoFisher Scientific	AR008
Pen/Strep (100X)	ThermoFisher Scientific	15140122
200 mM Glutamine	ThermoFisher Scientific	25030081
Glucose	ThermoFisher Scientific	15023-021	25% (w/v) Solution, 0.2 mm filter sterilized
Neurobasal A	ThermoFisher Scientific	10888-022	Complete Medium: 48 mL Neurobasal A, 1 mL N21-MAX, 0.625 mL 200 mM Glutamine, 0.180 mL 25% Glucose, 0.250 mL 100x pen/strep.
Third generation lentivirus packaging	Life Technologies	K4975-00
159 K cutoff centrifugal filters (Centricon)	EMD Millipore
Lentiviral cloning system (InFusion)	Clonetech
Plasmids 30323, 50856, 51279	Addgene
Neuronal cell viability dye (NeuO)	Stemcell technologies	1801	Thaw once and quick freeze in 4 µL aliquots. Store at -20 °C
Inverted microscope	Olympus	IX83
Microscope objectives	Olympus	air: 4X, 20; oil: 40X, 60X,
Spinning disc confocal system	Yokagawa	CSU22
Microscope EMCCD camera	Photometrics	Cascade II
Linear encoded (x,y), piezo z flat top stage	ASI
Microscope lasers and integration	Intelligent Imaging Innovations
HEK293T cells	American Type Culture Collection	CRL-3216
Human Plasmin	Sigma Aldrich	P1867	0.002 U/mL in 0.1% bovine serum albumin (0.2 mm filter sterilized), quick freeze in liquid nitrogen and store at -80 °C.