All experiments were done under the supervision of the Purdue Animal Care and Use Committee and the Laboratory Animal Program at Purdue University.
This work was sponsored by the Defense Advanced Research Projects Agency (DARPA) Microsystems Technology Office (MTO), under the auspices of Dr. Jack W. Judy (jack.judy@darpa.mil) as part of the Reliable Neural Technology Program, through the Space and Naval Warfare Systems Command (SPAWAR) Systems Center (SSC) Pacific Grant No. N66001-11-1-4013.
The authors would like to thank Mikhail Slipchenko, Don Ready, Greg Richter, Aaron Taylor, and Kevin Eliceiri for sharing their microscopy expertise.

Solutions

Phosphate Buffered Saline (PBS) - in g/l; 9 g NaCl, 0.144 g KH2PO4, 0.795 g Na2HPO4, at pH 7.4
4% Formaldehyde - in ml/l; 202 ml sodium phosphate dibasic solution (0.4 M Na2HPO4), 48 ml sodium phosphate monobasic solution (0.4 M NaH2PO4), 500 ml 8% formaldehyde solution, 250 ml Milli-Q DDi water, at pH 7.4
HEPES Buffered Hank's Saline with Sodium Azide (HBHS) - in g/l; 7.5 g NaCl, 0.3 g KCl, 0.06 g KH2PO4, 0.13 g Na2HPO4, 2 g glucose, 2.4 g HEPES, 0.05 g MgCl2 6 parts H2O, 0.05 g MgSO4 7 parts H2O, 0.165 g CaCl2, 0.09 g NaN3 at pH 7.4
Wash Solution (WS) - 1% Vol/Vol, Normal Goat Serum, 0.3% Triton X-100, in HBHS with sodium azide (NaN3). Refrigerate the WS at 4 &deg;C before use in subsequent steps.
U2 Scale Solution - 4 M urea, 30% glycerol and 0.1% Triton X-100 17
PROCEDURE
All experiments were done under the supervision of the Purdue Animal Care and Use Committee and the Laboratory Animal Program at Purdue University.

1. Surgery
<ol>
 <li>Various aseptic surgical methods are compatible with the presented histological method. The use of a stereotaxic frame and automated inserters are recommended to improve reproducibility and control of the microelectrode arrays (MEA) implantation angle with respect to stereotaxic planes.</li>
</ol>
Note: In this demonstration our surgical method is as follows: hold the rodent subject in stereotaxic earbars while under isoflurane anesthetic (between 1-3%) carried by medical grade oxygen. Test for the lack of a toe-pinch reflect in the rat or lack of a tail pinch reflex in the mouse to confirm that the animal is fully anesthetized. Apply eye ointment and clean the surgical site with three alternating washes of Betadine and ethanol. Wash hands, dawn surgical gloves, hairnet, facemask and gown. Inject a bolus of Lidocaine to numb the surgical area, create an incision using scissors or a scalpel, clear periosteum with bone scraper and cotton applicators, and create a craniotomy using a dental drill handpiece and drill bit. Drive a single-shank MEA into the cortex using a micromanipulator. Have a surgical assistant aid in maintaining aseptic surgery conditions and document the progress of the surgery.
<ol start="2">
 <li>Following implantation of the MEA into the brain, collect images through a surgical microscope to inform the eventual removal of the skull from around the brain. Implanted portions of the devices will be preserved in situ.</li>
</ol>
Note: The eventual collection of tissue with in situ devices relies on closely matching the plane of device insertion with the plane of tissue sectioning. Detailed notes about the device insertion plane relative to the brain are thus very important.
<ol start="3">
 <li>After device implantation, apply the silicone elastomer Kwik-Sil (WPI), around any exposed MEA shanks or cabling, and allow to cure. A sterilized plastic piece, such as a cut pipette tip, may be used to help contain the silicone elastomer in a small well around the craniotomy while curing.</li>
 <li>Apply a layer of two-part dental acrylic ("Jet Liquid" and "Jet Powder" from Lang Dental) over the Kwik-Sil and any exposed skull.</li>
</ol>
Note: In a later step, the headcap will be melted through to allow for the implant to be separated from the skull. UV-curing acrylic should be avoided over the Kwik-Sil and craniotomy, as this very hard acrylic is difficult to remove later. Visually opaque materials around the implant should also be avoided; clear Kwik-Sil provides a view of the implant during subsequent steps of this protocol.
<ol start="5">
 <li>Perform post-operative care procedures as per the local protocol of the animal welfare regulations, and return ambulatory subjects to single-housed cage-boxes.</li>
</ol>
2. Perfusion and Tissue Collection
Note: See Gage et al. for a detailed perfusion procedure walkthrough in the rat animal model19.
<ol>
 <li>Use the anesthesia and perfusion protocol approved by the institution's animal care and use committee. After deeply anesthetizing the rodent (confirmed by a lack of toe-pinch reflex in rat or tail-pinch reflex in mice) and exposing the heart, make shallow scissor cuts into the left ventricle and right atrium. Insert a blunt needle into the left ventricle and deliver room temp PBS. Deliver a total of ~10 ml PBS at ~5 ml/min in mice and ~200 ml PBS at ~100 ml/min in rats. Look for blood clearance from the liver during.</li>
 <li>Inject histologically relevant chemical labels transcardially, if so desired, by syringe delivery with care taken to avoid introducing bubbles.</li>
</ol>
Note: Vascular labels (e.g. DiI) and nucleic acid counterstain dyes (e.g. Hoechst 33342) are examples of chemical labels deliverable during perfusion. When administered properly, these histological markers should label the whole animal20.
<ol start="3">
 <li>Deliver buffered, room temperature, 4% formaldehyde transcardially to fix the animal. Avoid septum perforation across the heart, which can be evidenced by lung-nose drainage during perfusion. Look for fixation tremors in the large muscles to indicate complete perfusion. Deliver a total of ~10 ml fixative at ~5 ml/min in mice and ~200 ml fixative at ~100 ml/min in rats.</li>
 <li>Following perfusion, decapitate the animal, expose part of the brain stem or cerebellum using rongeurs or scissors, and place head in 4% formaldehyde solution for overnight at 4 &deg;C.</li>
 <li>Wash the head in three changes of PBS with one to four hour intervals between washes.</li>
 <li>Store fixed heads in HBHS containing sodium azide at 4 &deg;C for days to months.</li>
</ol>
3. Brain Removal with in situ Devices
Note: Perform this section in a fume hood while wearing appropriate personal protective equipment. Avoid tissue desiccation by returning the fixed head to HBHS solution periodically.
<ol>
 <li>Carefully dissect away skin and other tissues from around the acrylic skull cap using forceps and small scissors.</li>
 <li>While wearing heat-insensitive gloves, use a soldering iron to remove dental acrylic and expose an area of underlying clear Kwik-Sil (Figure 2a).</li>
</ol>
Note: The goal of this step is to allow micro-scissors an appropriate angle of access to positions where the implanted devices enter the brain, so that brain-implanted components may be separated from components secured to the skull.
<ol start="3">
 <li>Under a surgical microscope, cut into the silicone elastomer with curved micro-scissors, removing small pieces of Kwik-Sil with tweezers down to the position where the devices enter the brain. </li>
 <li>Continue cutting along the surface of the craniotomy with curved micro-scissors to separate device components attached to polysilicon and skull from components implanted in the brain. Again use microscope magnification and great care when cutting through the exposed device components, taking care to avoid pushing or dragging the implants in the fixed tissue.</li>
 <li>Remove bone around the headcap with care using rongeurs. Use a spatula to separate the brain with implanted devices from the skull. Store the brain in HBHS solution until ready to slice.</li>
 <li>Place the brain in a glass Petri dish filled with HBHS, and use a razor blade to remove extraneous tissue, such as spinal cord, cerebellum, or olfactory bulb, depending on whether they are relevant to the location of the device implantation. Use a brain block (Ted Pella) and razor blades to section the brain and create a flat plane closely matching the angle of the implanted devices; this is accomplished by placing the brain in an appropriately-sized brain block, orienting the brain such that any visible portion of the implant is parallel with the razor blade guide, and placing a razor blade in a blade guide at least 2 mm from the implant. Press the razor blade through the tissue. Mount this surface to a vibratome platform. Alternatively, a razor blade mounted to a micromanipulator can be used in a similarly controlled fashion as the brain block to create a plane closely matching the direction of the implant.</li>
 <li>Place ice under the vibratome platform, and, after allowing the tissue surface to briefly dry on a paper towel, adhere the brain to a vibratome using Super Glue. Glue the flat tissue plane created in the previous step is to the stage. With asymmetric tissue dimensions, orient the sample such that the widest dimension is glued down parallel with the movement direction of the vibratome blade to help avoid the blade potentially knocking the tissue over. After the glue sets (~1-2 min), add chilled (~4 &deg;C) PBS to the vibratome dish around the tissue.</li>
 <li>Collect thick tissue slices between 200-400 &mu;m, including control tissue, using the maximum vibration rate (~100 Hz) and a slow blade progression rate (&lt; 0.2 mm per second), with a blade angle of 10 &deg; from horizontal. Collect slices carefully with a brush or scoop spatula and store in HBHS in a 24 well plate at 4 &deg;C.</li>
 <li>Once the in situ device is visible to the unaided eye or a surgical microscope, collect thinner sections to approach the device in slice increments of 100 &mu;m or less.</li>
</ol>
Note: Typical silicon micro-implants are visible with the unaided eye in formaldehyde-fixed rodent cerebral cortex tissue between 300 and 500 &mu;m from the device surface.
<ol start="10">
 <li>With the in situ device now close to the surface of the tissue, collect a &gt;250 &mu;m tissue slice containing the device within. Estimation of the necessary thickness can be aided by referencing previously collected slices.</li>
</ol>
Note: Larger devices, multishank devices, and angled devices may require thicker tissue slices (&gt;500 &mu;m) to capture the device in one tissue piece; remember that both the penetration of applied labels and the microscopy imaging depth will be limited in the eventual data collection from these very thick slices. If possible, avoid striking the device with the vibratome blade, as this can cause dragging of the device through the tissue and morphological deformation.
4. Tissue Processing and Clearing
<ol>
 <li>Wash slices 3x at 5 min per wash in HBHS</li>
 <li>Incubate in sodium borohydride (5 mg NaBH4/ 1 ml of HBHS) for 30 min total (15 min each side of the slice). Flip the slices using a stainless steel or Teflon coated micro spoon and small paintbrush (Ted Pella). Slow and thoughtful movements improve the success of each slice flip. Avoid touching any areas around the implanted device with the brush. When possible, adding significantly more solution than required (&gt;2 ml) allows one to manipulate and flip the tissue slices more easily.</li>
</ol>
Note: This step reduces endogenous autofluorescence; skip this step if fluorescent labels were applied during perfusion or xFP transgenic markers are present.
<ol start="3">
 <li>Wash slices 3x at 5 min per wash in Wash Solution at room temperature.</li>
</ol>
Note: Agitation during wash and incubation steps is optional. The authors speculate that subtle jarring of the stiff implant with respect to the surrounding brain tissue may occur with agitation. However, an orbital mixer moving at a gentle rate (&lt; 60 rpm) may avoid this while increasing the movement of solutions.
<ol start="4">
 <li>Block for 2 hr in Wash Solution at room temperature (flip slices after one hour).</li>
 <li>Incubate in primary antibodies (diluted in Wash Solution) for approximately 48 hr (24 hr on each side of the slice) at 4 &deg;C.</li>
 <li>Perform 6 quick 3 min washes with Wash Solution.</li>
 <li>Perform 6, 1 hr washes (3 washes on each side of the tissue slice) in Wash Solution (store at 4 &deg;C if washing overnight).</li>
 <li>Incubate in secondary antibodies (diluted with Wash Solution) for approximately 48 hr (24 hr on each side) in 4 &deg;C.</li>
 <li>Perform 6 quick 3 min washes with Wash Solution.</li>
 <li>Perform 6, 1 hr washes (3 washes on each side) in Wash Solution; store at 4 &deg;C if washing overnight.</li>
 <li>Remove Wash Solution and add the glycerol-based "U2 - Scale" clearing solution. Allow to clear approximately 1 week for thick slices (250-500 &mu;m) or 2-3 weeks for very thick tissue sections (&gt;500 &mu;m) at 4 &deg;C.</li>
 <li>Cover plate with foil and store at 4 &deg;C.</li>
 <li>Mount in slides accommodating thick tissue sections (Figure 2b, 2c) using clearing solution as mounting media and sealing with clear nail polish. Store at 4 &deg;C away from light.</li>
</ol>
5. Panorama Imaging of Full Tissue and Device Interface
<ol>
 <li>Using longer-working distance microscope objectives (&gt; 2 mm), begin imaging with a laser scanning confocal microscope or 2-photon microscope. We will demonstrate with a Zeiss LSM 710, Carl Zeiss "Zen" software (2010), and a computer-controlled XY translating stage to image a 3D panorama around an implant.</li>
</ol>
Note: Correct for the refractive index of the clearing solution either in software, through a corrective collar on the objective, or in data processing after collection. In this demonstration we enter a refractive index value for the "U2" clearing solution of 1.4 into the Leica Zen microscope software; this setting subtly adjusts z-step intervals to account for refractive index mismatch.
<ol start="2">
 <li>In tissue near the device, roughly assess the appropriate z-axis window and data collection settings in the z-dimension for each fluorescent channel using low laser power (~0.5 to 5%) and large z-step sizes (&gt;25 &mu;m).</li>
</ol>
Note: Include additional space above and below when setting the z-axis when setting up for panoramic data collection, as the tissue may not lie entirely flat on the slide glass (~20 &mu;m each direction).
<ol start="3">
 <li>Set the objective at the x-y center of your eventual panorama; using the Zen panorama software, determine the number of collection positions required for each axis (x and y) to cover your region of interest.</li>
 <li>Reassess and finalize the z-axis collection window.</li>
 <li>Manually set the detector sensitivity and laser power to ramp appropriately with increased depth; the aim is typically to maintain approximately the same image intensity regardless of imaging depth into the tissue slice, but to also avoid high background noise.</li>
 <li>Collect each fluorescent image data series. If necessary to avoid overlapping signals, run laser lines sequentially.</li>
</ol>
Note: If available, set the software to automatically save data to the hard drive during collection.
<ol start="7">
 <li>Change the software settings to collect the "reflectance" and "transmittance" data, and use a longer, more penetrative wavelength laser (e.g. 633 nm) to capture these channels. Determine the appropriate laser power and detector sensitivity for "reflectance" and "transmittance" collection and repeat the same collection series around the implanted device.</li>
 <li>Export the data for later processing, quantification, and analysis.</li>
</ol>

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The authors have no relevant disclosures to declare, financial or otherwise.

The "Device-Capture Histology" (DCHist) method demonstrated here enables the close histological assessment of morphologically preserved interactions between brain tissue and tissue implants. DCHist tissue collection requires careful separation of skull-mounted device components from components implanted in the brain. DCHist also requires collection of a thick histological tissue slice (&gt;250 &mu;m). These tissue sections, once labeled, cleared, mounted, and imaged, can provide novel insights into the implantation injury or subsequent chronic response to indwelling devices. Using advanced microscopy tools, the interface between tissue and device can be imaged and analyzed in high detail as shown in Figures 3-5.
The presented techniques in their current form rely on the ability to slowly excavate away the headcap made from two-part dental cement and Kwik-Sil down to the point of device implantation. Utilizing dental cement that can be burned away or otherwise removed and a clear silicon elastomer through which small scissors can be guided visually greatly aid in successfully separating the implanted device from its skull-mounted components. Attempting to cut the device under the skull and above the brain in order to collect it in situ is not recommended, as the skull-mounted implant will be pulled out of the brain a significant amount.
Although thinner, smaller implants, such as single shank MEAs from NeuroNexus Technologies, are most amenable to DCHist, the principles are not restricted to single device shanks or microelectrode arrays. Collection and imaging strategies are broadly applicable to multi-shank devices and larger implants such as cannulas, with the requirements being that they must be separated from any skull mount and collected within a thick tissue section. Although the authors are focused on analysis of tissue surrounding cortical implants, devices driven deeper into the brain could also be collected and imaged. The depth of implant insertion should not affect the capture of the implant in a slice provided the device does not deviate wildly from the known angle of insertion.
Limitations exist to the useful application of the DCHist method. Brain tissue sections are challenging to image through hundreds of micrometers, especially in areas of white matter, although optical clearing solutions can greatly improve imaging depths in various tissues21. To further improve imaging depth, two-photon excitation microscopy may be employed along with the optical clearing described.
Another potential limitation of the described method can be the specific fluorescent immunohistochemistry methods and antibodies employed by researchers. Passive diffusion typically drives the incorporation of these markers through fixed tissue and onto antigen binding sites. Final antibody working concentrations must be determined by researchers on a case by case basis to maximize label penetration while avoiding high levels of background labeling. Antibody labeling should not be variable between slices that are processed identically, but different antibodies may vary considerably in their ability to penetrate and tag their antigen, with some antibodies easily labeling antigens many hundreds of micrometers deep and others labeling antigens only tens of micrometers deep. We describe improving this diffusion by applying antibody labels at higher than typical concentrations, for multiple days of application, and in a solution containing dilute detergent and blocking serum. Periodically flipping the slices also promotes even labeling. Antigen retrieval steps and alternative fixation processes (e.g. glutaraldehyde, microwave, etc.) may be appropriate for specific antigens. Secondary antibody-fluorochrome conjugates may also vary in their performance labeling thick sections, though this has not been observed by the authors using Alexa Fluor labels from Invitrogen. Alternatively, transgenic animal subjects expressing fluorescent proteins in the cell types of interest may be utilized to avoid issues with antibody label penetration, as many fluorescent proteins, such as eGFP, retain their fluorescence after formaldehyde treatment, and may be immediately visualized in tissue sections.
DCHist is a powerful set of techniques to capture and analyze the impact of implanted microdevices on brain tissue. Coupling this histological protocol with in vivo assessment of electrophysiology quality and electrode impedance data22 could greatly improve our understanding of the biological sources of physiology variability and degradation. The field of implanted neural prosthetic devices in particular may benefit from the detailed DCHist imaging of the intact device/tissue interface to inform further development of biologically neutral MEA devices.

The field of neuroprosthetic research aims to assist individuals suffering from various disabilities and disorders by bypassing diseased or damaged structures in the body through CNS interfacing devices1,2. Brain-implanted microdevices, such as microelectrode arrays (MEAs), can be used to record or stimulate brain structures, and thus allow the establishment of long-term interfaces between electronics and CNS tissue3-5. Penetrating MEAs, devices which are driven into the brain tissue, hold particular promise as bi-directional interfaces due to the close proximity within which they present electrodes to a relatively small set of nearby neurons6.
However, complex tissue responses result from the long-term implantation of penetrating MEAs, often resulting in variable and gradually degrading electrophysiological signal-to-noise ratios over days to months, and an increase in electrical impedance between electrode sites and ground7,8. The putative origins of these changes include the activation of microglia, reactive astrocytosis along the microdevices, and a loss or migration of neurons from the tissue surrounding to implanted devices9-11. A major challenge to understanding these tissue changes around chronic, penetration MEAs is the difficulty in capturing histological data of the intact tissue interface surrounding chronically implanted devices12. Histological analysis of the tissue with the device/tissue interface still present would improve upon the current device-removal histological protocols. With an undisturbed device remaining in the tissue, the biological impact of relatively subtle interactions, such as the utilization of biocompatible coatings13,14 or the electrical clearing of the electrode surface15,16, could be imaged and analyzed with respect to the implant.
Here we demonstrate a method to collect, process, and image the intact microdevice interface for detailed microscopy-based analysis of the surrounding brain tissue. In this method, the device and surrounding brain tissue are collected within a thick (&gt;250 &mu;m) tissue section using a vibratome. To improve histological label penetration into these thick slices, fluorescent histochemical and immunohistochemical labels are applied at high concentrations in solutions containing blocking serum and detergent for multiple days. An optical clearing solution is employed to improve microscopy imaging depths, and the tissue is mounted in 2-sided chambers for subsequent laser scanning confocal microscopy17. To capture the full histological interface, a computer-controlled translational stage is used during imaging to collect z-stack panoramas along the length of implants. In addition to imaging applied tissue labels, collecting laser reflectance back from implants and transmission light through the tissue both help localize the device interface in relation to surrounding tissue. Tissue prepared using this "Device-Capture Histology" (DCHist) protocol provides imaging access to the morphologically preserved tissue/device interactions, and thus improves upon previous device-removal histological protocols18.

<table border="1">
 <tbody>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Reagents</td>
 </tr>
 <tr>
 <td>Hoechst 33342</td>
 <td>Invitrogen</td>
 <td>14533</td>
 <td>DNA marker</td>
 </tr>
 <tr>
 <td>Rabbit anti-Iba1</td>
 <td>Wako (Japan)</td>
 <td>019-19741</td>
 <td>Monocyte antibody</td>
 </tr>
 <tr>
 <td>Dental acrylic</td>
 <td>Lang Dental (various distributors)</td>
 <td>Jet Denture Repair Powder &amp; Liquid</td>
 <td>Headcap construction</td>
 </tr>
 <tr>
 <td>Kwik-Sil</td>
 <td>World Precision Instruments</td>
 <td>KWIK-SIL</td>
 <td>Covering exposed cranial implants</td>
 </tr>
 <tr>
 <td>Normal goat serum</td>
 <td>Jackson ImmunoResearch</td>
 <td>005-000-121</td>
 <td>Tissue processing</td>
 </tr>
 <tr>
 <td>Triton X-100</td>
 <td>Sigma-Aldrich</td>
 <td>X100-500ml</td>
 <td>Tissue processing</td>
 </tr>
 <tr>
 <td>Urea</td>
 <td>Sigma-Aldrich</td>
 <td>U4883</td>
 <td>Clearing solution</td>
 </tr>
 <tr>
 <td>Glycerol</td>
 <td>Sigma-Aldrich</td>
 <td>G20225</td>
 <td>Clearing solution</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Equipment</td>
 </tr>
 <tr>
 <td>Curved micro-scissors</td>
 <td>World Precision Instruments</td>
 <td>14208</td>
 <td>Cutting implant before removing brain</td>
 </tr>
 <tr>
 <td>Razor blades</td>
 <td>Ted Pella</td>
 <td>(various sizes)</td>
 <td>For use with Brain Block</td>
 </tr>
 <tr>
 <td>Acrylic Brain Matrice</td>
 <td>Ted Pella</td>
 <td>15054</td>
 <td>Brain Block to create initial plane in tissue</td>
 </tr>
 <tr>
 <td>Plastic slides</td>
 <td>Ted Pella</td>
 <td>260225</td>
 <td>Mounting tissue</td>
 </tr>
 <tr>
 <td>Cover glass</td>
 <td>Ted Pella</td>
 <td>(various sizes)</td>
 <td>Mounting tissue</td>
 </tr>
 <tr>
 <td>Electrode arrays</td>
 <td>NeuroNexus Technologies</td>
 <td>(various designs)</td>
 <td>Example MEAs</td>
 </tr>
 <tr>
 <td>Vibratome</td>
 <td>Leica</td>
 <td>VT1000 S</td>
 <td>Specific system used in presented method</td>
 </tr>
 <tr>
 <td>Vibratome blades</td>
 <td>(various suppliers)</td>
 <td></td>
 <td>Collecting slices</td>
 </tr>
 <tr>
 <td>24-well plates</td>
 <td>(various suppliers)</td>
 <td></td>
 <td>Storing and processing tissue slices</td>
 </tr>
 <tr>
 <td>Confocal microscope with motorized XY-scanning stage</td>
 <td>Carl Zeiss Microscopy</td>
 <td>Zeiss LSM 710 and 'Zen 2010' software</td>
 <td>Specific system used in presented method</td>
 </tr>
 <tr>
 <td>Soldering iron</td>
 <td>(various suppliers)</td>
 <td></td>
 <td>Excavating headcap</td>
 </tr>
 </tbody>
</table>

intact histological characterization of brain-implanted microdevices and surrounding tissue

Research into the design and utilization of brain-implanted microdevices, such as microelectrode arrays, aims to produce clinically relevant devices that interface chronically with surrounding brain tissue. Tissue surrounding these implants is thought to react to the presence of the devices over time, which includes the formation of an insulating "glial scar" around the devices. However, histological analysis of these tissue changes is typically performed after explanting the device, in a process that can disrupt the morphology of the tissue of interest.
Here we demonstrate a protocol in which cortical-implanted devices are collected intact in surrounding rodent brain tissue. We describe how, once perfused with fixative, brains are removed and sliced in such a way as to avoid explanting devices. We outline fluorescent antibody labeling and optical clearing methods useful for producing an informative, yet thick tissue section. Finally, we demonstrate the mounting and imaging of these tissue sections in order to investigate the biological interface around brain-implanted devices.

Brain-implanted MEAs can be collected in tissue slices by first separating any skull-mounted components from the brain-embedded components. Figure 2a shows the results of removing one side of a dental cement headcap and a portion of Kwik-Sil surrounding a MEA cable on a rat skull. A soldering iron was used to remove the dental cement and Kwik-Sil in a fume hood. The cabling and any non-implanted MEA structures were next cut with micro-scissors by slowly excavating through the Kwik-Sil down to the surface of the fixed brain.
After slicing, labeling, and clearing tissue, simple custom-made slides are useful for mounting the thick tissue slices (Figure 2b). A mounted brain slice containing an implanted microdevice is shown in Figure 2c. Imaging through either side of the slide can allow you to assess the interface around a device, as demonstrated in Figure 3, where microglia along the silicon backing of an MEA are visible from one side (Figure 3a) and microglia along electrode sites and traces are visible on the other side (Figure 3b).
Once mounted, tissue can be imaged using an XY translation stage. Overviews of fluorescent labels across entire tissue slices can be generated at the desired resolution (Figure 4). Detailed examination of the morphologically preserved tissue interface around implanted microdevices can be collected under high magnification for analysis of the local tissue response (Figure 5).
<img src="/files/ftp_upload/50126/50126fig1.jpg" alt="Figure 1" fo:content-width="6.5in" fo:src="/files/ftp_upload/50126/50126fig1highres.jpg"/> 
Figure 1. Overview of procedure. (a-c) Brain micro-implant surgeries are finalized with a layer of Kwik-Sil polymer immediately surrounding the device, followed by an acrylic cement covering. (d, e) Following euthanasia, the acrylic layer is burned away, and skull-mounted components of the device are separated from implanted components by cutting through the silicone elastomer. (f, g) The brain is then removed from the skull and is cut and mounted to the vibratome stage. (h-k) Tissue slices are collected, including a tissue slice containing the device. (l) Tissue can then be processed and mounted in custom chambers for detailed microscopy. <a href="https://www.jove.com/files/ftp_upload/50126/50126fig1large.jpg" target="_blank"> Click here to view larger figure</a>.
<img src="/files/ftp_upload/50126/50126fig2.jpg" alt="Figure 2"/> 
Figure 2. (a) Clear Kwik-Sil silicone elastomer surrounding an electrode-array cable (arrow) has been exposed by burning away a window in the dental cement cap on a perfused rodent head using a soldering iron. (b) To accommodate imaging into either side of a thick tissue slice, simple "slide-chambers" can be made by cutting a plastic slide and adhering coverglass to either side around the tissue and mounting solution. (c) A rat brain tissue slice with intact implant (red arrow) is shown after mounting. Either side of the tissue is quickly accessible to microscopy imaging with this setup. For scale, panel (a) is 25 mm across at the bottom, while panels (b, c) are 80 mm across at the bottom.
<img src="/files/ftp_upload/50126/50126fig3.jpg" alt="Figure 3" fo:content-width="3.5in" fo:src="/files/ftp_upload/50126/50126fig3highres.jpg"/> 
Figure 3. Maximum intensity z-projections of 40 &mu;m thick image stacks, collected around the backside (a) and front (b) of an implanted microelectrode array. Microglia (labeled with anti-Iba1 and Alexa Fluor 633, white) and laser light reflectance (red), collected from the surface of the device, were sequentially imaged from both sides of a 400 &mu;m thick tissue slice. As the brain implants are often opaque, mounting with optical access to both sides provides a clear view of labeled tissue structures "behind" the implant with respect to the microscope objective. Scale bar 50 &mu;m.
<img src="/files/ftp_upload/50126/50126fig4.jpg" alt="Figure 4"/> 
Figure 4. A labeled DCHist slice imaged using a computer-controlled stage on a laser scanning confocal microscope. (a) Cell nuclei (stained with Hoechst 33342) and (b) monocytes/microglia (labeled with anti-Iba1) were simultaneously imaged on separate channels. (c) Transmission light and reflectance were also collected, showing the location of the 4-week implant with respect to Hoechst and Iba1 data. (d) Overlay of all channels but transmission is also shown. The white rectangle area is expanded in the images appearing at right. Scale bar 1 mm.
<img src="/files/ftp_upload/50126/50126fig5.jpg" alt="Figure 5" fo:content-width="4.5in" fo:src="/files/ftp_upload/50126/50126fig5highres.jpg"/> 
Figure 5. Using a computer-controlled microscope stage and appropriate software, panoramic imaging data can be collected around the implant. Reflectance (a) and transmittance (b) allow localization of the device, while fluorescent antibody or chemical labels (c, anti-Iba1 labeling of microglia and macrophages) allow detailed imaging of tissue components along the intact tissue interface. Scale bar 200 &mu;m.

Watch this Scientific Journal Video about Intact Histological Characterization of Brain-implanted Microdevices and Surrounding Tissue at JoVE.com

Intact Histological Characterization of Brain-implanted Microdevices and Surrounding Tissue

Here we present a histological method for capturing, labeling, optically clearing, and imaging the intact brain tissue interface around chronically implanted microdevices in rodent brain tissue. Results from the techniques comprising this method are useful for understanding the impact of various penetrating brain-implants on their surrounding tissue.

Research  into the design and utilization of brain-implanted microdevices, such as  microelectrode arrays, aims to produce clinically relevant devices ...

intact-histological-characterization-brain-implanted-microdevices

Research

JoVE Journal

Neuroscience

16.8K Views.  Purdue University. Here we present a histological method for capturing, labeling, optically clearing, and imaging the intact brain tissue interface around chronically implanted microdevices in rodent brain tissue. Results from the techniques comprising this method are useful for understanding the impact of various penetrating brain-implants on their surrounding tissue.

Video: Intact Histological Characterization of Brain-implanted Microdevices and Surrounding Tissue

Scalable Fluidic Injector Arrays for Viral Targeting of Intact 3-D Brain Circuits

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted in neurological and psychiatric disorders--would be greatly facilitated by a technology for rapidly targeting genes to complex 3-dimensional neural circuits, enabling fast creation of "circuit-level transgenics." We have recently developed methods in which viruses encoding for light-sensitive proteins can sensitize specific cell types to millisecond-timescale activation and silencing in the intact brain. We here present the design and implementation of an injector array capable of delivering viruses (or other fluids) to dozens of defined points within the 3-dimensional structure of the brain (Figure. 1A, 1B). The injector array comprises one or more displacement pumps that each drive a set of syringes, each of which feeds into a polyimide/fused-silica capillary via a high-pressure-tolerant connector. The capillaries are sized, and then inserted into, desired locations specified by custom-milling a stereotactic positioning board, thus allowing viruses or other reagents to be delivered to the desired set of brain regions. To use the device, the surgeon first fills the fluidic subsystem entirely with oil, backfills the capillaries with the virus, inserts the device into the brain, and infuses reagents slowly (&lt;0.1 microliters/min). The parallel nature of the injector array facilitates rapid, accurate, and robust labeling of entire neural circuits with viral payloads such as optical sensitizers to enable light-activation and silencing of defined brain circuits. Along with other technologies, such as optical fiber arrays for light delivery to desired sets of brain regions, we hope to create a toolbox that enables the systematic probing of causal neural functions in the intact brain. This technology may not only open up such systematic approaches to circuit-focused neuroscience in mammals, and facilitate labeling of brain regions in large animals such as non-human primates, but may also open up a clinical translational path for cell-specific optical control prosthetics, whose precision may enable improved treatment of intractable brain disorders. Finally, such devices as described here may facilitate precisely-timed fluidic delivery of other payloads, such as stem cells and pharmacological agents, to 3-dimensional structures, in an easily user-customizable fashion.

Controlling and analyzing neural circuits in vivo would be facilitated by a technology for delivery of viruses and other reagents to desired 3-dimensional sets of brain regions. We demonstrate customized fluidic injector array fabrication, and delivery of virally-encoded optical sensitizers, enabling optical manipulation of complex brain circuits.

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted ...

Physiological, Morphological and Neurochemical Characterization of Neurons Modulated by Movement

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor functions. Most investigations of sensorimotor mechanisms rely on either examination of neurons while an animal is static1,2 or record extracellular neuronal activity during a movement.3,4 While these studies have provided the fundamental background for sensorimotor function, they either do not evaluate functional information which occurs during a movement or are limited in their ability to fully characterize the anatomy, physiology and neurochemical phenotype of the neuron. A technique is shown here which allows extensive characterization of individual neurons during an in vivo movement. This technique can be used not only to study primary afferent neurons but also to characterize motoneurons and sensorimotor interneurons. Initially the response of a single neuron is recorded using electrophysiological methods during various movements of the mandible followed by determination of the receptive field for the neuron. A neuronal tracer is then intracellularly injected into the neuron and the brain is processed so that the neuron can be visualized with light, electron or confocal microscopy (Fig. 1). The detailed morphology of the characterized neuron is then reconstructed so that neuronal morphology can be correlated with the physiological response of the neuron (Figs. 2,3). In this communication important key details and tips for successful implementation of this technique are provided. Valuable additional information can be determined for the neuron under study by combining this method with other techniques. Retrograde neuronal labeling can be used to determine neurons with which the labeled neuron synapses; thus allowing detailed determination of neuronal circuitry. Immunocytochemistry can be combined with this method to examine neurotransmitters within the labeled neuron and to determine the chemical phenotypes of neurons with which the labeled neuron synapses. The labeled neuron can also be processed for electron microscopy to determine the ultrastructural features and microcircuitry of the labeled neuron. Overall this technique is a powerful method to thoroughly characterize neurons during in vivo movement thus allowing substantial insight into the role of the neuron in sensorimotor function.

A technique is described to quantify the in vivo physiological response of mammalian neurons during movement and correlate the physiology of the neuron with neuronal morphology, neurochemical phenotype and synaptic microcircuitry.

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor ...

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Electrophysiological Characterization of GFP-Expressing Cell Populations in the Intact Retina

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack conspicuous morphological features or show a low population density. This applies particularly to retinal amacrine cells, an exceptionally multiform class of interneurons that comprise roughly 30 subtypes in mammals1. Though being a crucial part of the visual processing by shaping the retinal output2, most of these subtypes have not been studied up to now in a functional context because encountering these cells with a recording electrode is a rare event.
Recently, a multitude of transgenic mouse lines is available that express fluorescent markers like green fluorescent protein (GFP) under the control of promoters for membrane receptors or enzymes that are specific to only a subset of neurons in a given tissue3,4. These pre-labeled cells are therefore accessible to directed microelectrode targeting under microscopic control, permitting the systematic study of their physiological properties in situ. However, excitation of fluorescent markers is accompanied by the risk of phototoxicity for the living tissue. In the retina, this approach is additionally hampered by the problem that excitation light causes appropriate stimulation of the photoreceptors, thus inflicting photopigment bleaching and transferring the retinal circuits into a light-adapted condition. These drawbacks are overcome by using infrared excitation delivered by a mode-locked laser in short pulses of the femtosecond range. Two-photon excitation provides energy sufficient for fluorophore excitation and at the same time restricts the excitation to a small tissue volume minimizing the hazards of photodamage5. Also, it leaves the retina responsive to visual stimuli since infrared light (&gt;850 nm) is only poorly absorbed by photopigments6.
In this article we demonstrate the use of a transgenic mouse retina to attain electrophysiological in situ recordings from GFP-expressing cells that are visually targeted by two-photon excitation. The retina is prepared and maintained in darkness and can be subjected to optical stimuli which are projected through the condenser of the microscope (Figure 1). Patch-clamp recording of light responses can be combined with dye filling to reveal the morphology and to check for gap junction-mediated dye coupling to neighboring cells, so that the target cell can by studied on different experimental levels.

This article depicts the recording of individual cells from fluorescently tagged neuronal populations in the intact mouse retina. By using two-photon infrared excitation transgenetically labeled cells were targeted for patch-clamp recording to study their light responses, receptive field properties, and morphology.

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack ...

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

3D Modeling of the Lateral Ventricles and Histological Characterization of Periventricular Tissue in Humans and Mouse

The ventricular system carries and circulates cerebral spinal fluid (CSF) and facilitates clearance of solutes and toxins from the brain. The functional units of the ventricles are ciliated epithelial cells termed ependymal cells, which line the ventricles and through ciliary action are capable of generating laminar flow of CSF at the ventricle surface. This monolayer of ependymal cells also provides barrier and filtration functions that promote exchange between brain interstitial fluids (ISF) and circulating CSF. Biochemical changes in the brain are thereby reflected in the composition of the CSF and destruction of the ependyma can disrupt the delicate balance of CSF and ISF exchange. In humans there is a strong correlation between lateral ventricle expansion and aging. Age-associated ventriculomegaly can occur even in the absence of dementia or obstruction of CSF flow. The exact cause and progression of ventriculomegaly is often unknown; however, enlarged ventricles can show regional and, often, extensive loss of ependymal cell coverage with ventricle surface astrogliosis and associated periventricular edema replacing the functional ependymal cell monolayer. Using MRI scans together with postmortem human brain tissue, we describe how to prepare, image and compile 3D renderings of lateral ventricle volumes, calculate lateral ventricle volumes, and characterize periventricular tissue through immunohistochemical analysis of en face lateral ventricle wall tissue preparations. Corresponding analyses of mouse brain tissue are also presented supporting the use of mouse models as a means to evaluate changes to the lateral ventricles and periventricular tissue found in human aging and disease. Together, these protocols allow investigations into the cause and effect of ventriculomegaly and highlight techniques to study ventricular system health and its important barrier and filtration functions within the brain.

Using MRI scans (human), 3D imaging software, and immunohistological analysis, we document changes to the brain&#8217;s lateral ventricles. Longitudinal 3D mapping of lateral ventricle volume changes and characterization of periventricular cellular changes that occur in the human brain due to aging or disease are then modeled in mice.

The ventricular system carries and circulates cerebral spinal fluid (CSF) and facilitates clearance of solutes and toxins from the brain. The functional ...

Electroencephalographic, Heart Rate, and Galvanic Skin Response Assessment for an Advertising Perception Study: Application to Antismoking Public Service Announcements

The evaluation of advertising, products, and packaging is traditionally performed through methods based on self-reports and focus groups, but these approaches often appear poorly accurate in scientific terms. Neuroscience is increasingly applied to the investigation of the neurophysiological bases of the perception of and reaction to commercial stimuli to support traditional marketing methods. In this context, a particular sector or marketing is represented by public service announcements (PSAs). The objective of this protocol is to apply electroencephalography (EEG) and autonomic signal analysis to study responses to selected antismoking PSAs. Two EEG indices were employed: the frontal alpha band EEG asymmetry (the Approach Withdrawal (AW) index) and the frontal theta (effort index). Furthermore, the autonomic Emotional Index (EI) was calculated, as derived from the Galvanic Skin Response (GSR) and Heart Rate (HR) signals. The present protocol describes a series of operational and computational steps required to properly estimate, through the aforementioned indices, the emotional and cerebral reaction of a group of subjects towards a selected number of antismoking PSAs. In particular, a campaign characterized by a symbolic communication style (classified as "awarded" on the basis of the prizes received by specialized committees) obtained the highest approach values, as estimated by the AW index. A spot and an image belonging to the same PSA campaign based on the "fear arousing appeal" and with a narrative/experiential communication style (classified as "effective" on the basis of the economical/health-related improvements promoted) reported the lowest and highest effort values, respectively. This is probably due to the complexity of the storytelling (spot) and to the immediateness of the image (a lady who underwent a tracheotomy). Finally, the same "effective" campaign showed the highest EI values, possibly because of the empathy induced by the testimonial and the explicitness of the message.

The following protocol describes a series of operational and computational steps required to properly estimate the emotional and cerebral reaction of a group of subjects towards a selected number of Public Service Announcements (PSAs) against smoking, aired in the USA and Europe during the period between 1998 and 2015.

The evaluation of advertising, products, and packaging is traditionally performed through methods based on self-reports and focus groups, but these ...

A Rat Model of Mild Intrauterine Hypoperfusion with Microcoil Stenosis

Intrauterine hypoperfusion/ischemia is one of the major causes of intrauterine/fetal growth restriction, preterm birth, and low birth weight. Most studies of this phenomenon have been performed in either models with severe intrauterine ischemia or models with gradient degree of intrauterine hypoperfusion. No study has been performed in a model on uniform mild intrauterine hypoperfusion (MIUH). Two models have been used for studies of MIUH: a model based on suture ligation of either side of the arterial arcade formed with the uterine and ovarian arteries, and a transient model based on clipping the bilateral ovarian arteries and aorta having patency. Those two rodent models of MIUH have some limitations, e.g., not all fetuses are subjected to MIUH, depending on their position in the uterine horn. In our MIUH model, all fetuses are subjected to a comparable level of intrauterine hypoperfusion. MIUH was achieved by mild stenosis of all four arteries feeding the uterus, i.e., the bilateral uterine and ovarian arteries.
Arterial stenosis was induced by metal microcoils wrapped around the feeding arteries. Producing arterial stenosis with microcoils allowed us to control, optimize, and reproduce decreased blood flow with very little inter-animal variability and a low mortality rate, thus enabling accurate evaluation. When microcoils with an inner diameter of 0.24 mm were used, the blood flow in both the placenta and fetus was mildly decreased (approximately 30% from the pre-stenosis level in the placenta). The offspring of our MIUH model clearly demonstrates long-lasting alterations in neurological, neuroanatomical and behavioral test results.

Mild intrauterine hypoperfusion was produced by artery stenosis with metal microcoils wrapped around the uterine and ovarian arteries in rats at embryonic day 17. This procedure produced prenatal hypoperfusion and intrauterine growth restriction.

Intrauterine hypoperfusion/ischemia is one of the major causes of intrauterine/fetal growth restriction, preterm birth, and low birth weight. Most studies ...

Characterization and Isolation of Mouse Primary Microglia by Density Gradient Centrifugation

Microglia, the resident immune cells in the brain, are the first responders to inflammation or injury in the central nervous system. Recent research has revealed microglia to be dynamic, capable of assuming both pro-inflammatory and anti-inflammatory phenotypes. Both M1 (pro-inflammatory) and M2 (pro-reparative) phenotypes play an important role in neuroinflammatory conditions such as perinatal brain injury, and exhibit differing functions in response to certain environmental stimuli. The modulation of microglial activation has been noted to confer neuroprotection thus suggesting microglia may have therapeutic potential in brain injury. However, more research is required to better understand the role of microglia in disease, and this protocol facilitates that. The protocol described below combines a density gradient centrifugation process to reduce cellular debris, with magnetic separation, producing a highly pure sample of primary microglial cells that can be used for in vitro experimentation, without the need for 2-3 weeks culturing. Additionally, the characterization steps yield robust functional data about microglia, aiding studies to better our understanding of the polarization and priming of these cells, which has strong implications in the field of regenerative medicine.

A protocol for the isolation of primary microglia from murine brains is presented. This technique aids in furthering the current understanding of neurological conditions. Density gradient centrifugation and magnetic separation are combined to produce sufficient yield of a highly pure sample. Furthermore, we outline the steps for characterization of microglia.

Microglia, the resident immune cells in the brain, are the first responders to inflammation or injury in the central nervous system. Recent research has ...