1Weldon School of Biomedical Engineering, Purdue University, 2Department of Biological Sciences, Purdue University
Woolley, A. J., Desai, H. A., Gaire, J., Ready, A. L., Otto, K. J. Intact Histological Characterization of Brain-implanted Microdevices and Surrounding Tissue. J. Vis. Exp. (72), e50126, doi:10.3791/50126 (2013).
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.
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 (>250 μm) 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.
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 °C before use in subsequent steps.
U2 Scale Solution - 4 M urea, 30% glycerol and 0.1% Triton X-100 17
All experiments were done under the supervision of the Purdue Animal Care and Use Committee and the Laboratory Animal Program at Purdue University.
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.
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.
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.
2. Perfusion and Tissue Collection
Note: See Gage et al. for a detailed perfusion procedure walkthrough in the rat animal model19.
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.
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.
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.
Note: Typical silicon micro-implants are visible with the unaided eye in formaldehyde-fixed rodent cerebral cortex tissue between 300 and 500 μm from the device surface.
Note: Larger devices, multishank devices, and angled devices may require thicker tissue slices (>500 μm) 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
Note: This step reduces endogenous autofluorescence; skip this step if fluorescent labels were applied during perfusion or xFP transgenic markers are present.
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 (< 60 rpm) may avoid this while increasing the movement of solutions.
5. Panorama Imaging of Full Tissue and Device Interface
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.
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 μm each direction).
Note: If available, set the software to automatically save data to the hard drive during collection.
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).
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. Click here to view larger figure.
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.
Figure 3. Maximum intensity z-projections of 40 μm 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 μm 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 μm.
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.
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 μm.
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 (>250 μm). 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 authors have no relevant disclosures to declare, financial or otherwise.
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 (firstname.lastname@example.org) 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.
|Hoechst 33342||Invitrogen||14533||DNA marker|
|Rabbit anti-Iba1||Wako (Japan)||019-19741||Monocyte antibody|
|Dental acrylic||Lang Dental (various distributors)||Jet Denture Repair Powder & Liquid||Headcap construction|
|Kwik-Sil||World Precision Instruments||KWIK-SIL||Covering exposed cranial implants|
|Normal goat serum||Jackson ImmunoResearch||005-000-121||Tissue processing|
|Triton X-100||Sigma-Aldrich||X100-500ml||Tissue processing|
|Curved micro-scissors||World Precision Instruments||14208||Cutting implant before removing brain|
|Razor blades||Ted Pella||(various sizes)||For use with Brain Block|
|Acrylic Brain Matrice||Ted Pella||15054||Brain Block to create initial plane in tissue|
|Plastic slides||Ted Pella||260225||Mounting tissue|
|Cover glass||Ted Pella||(various sizes)||Mounting tissue|
|Electrode arrays||NeuroNexus Technologies||(various designs)||Example MEAs|
|Vibratome||Leica||VT1000 S||Specific system used in presented method|
|Vibratome blades||(various suppliers)||Collecting slices|
|24-well plates||(various suppliers)||Storing and processing tissue slices|
|Confocal microscope with motorized XY-scanning stage||Carl Zeiss Microscopy||Zeiss LSM 710 and ’Zen 2010’ software||Specific system used in presented method|
|Soldering iron||(various suppliers)||Excavating headcap|