Name: assessment of thermal damage from robot-drilled craniotomy for cranial window surgery in mice
Uploaded: 2022-11-11T13:20:00
Description: Watch this Scientific Journal Video about Assessment of Thermal Damage from Robot-Drilled Craniotomy for Cranial Window Surgery in Mice at JoVE.com

This study was supported in part by Merit Review Awards GRANT12418820 (Capadona) and GRANTI01RX003420 (Shoffstall/Capadona), and Research Career Scientist Award # GRANT12635707 (Capadona) from the United States (US) Department of Veterans Affairs Rehabilitation Research and Development Service. Additionally, this work was also supported in part by the National Institute of Health, the National Institute of Neurological Disorders and Stroke GRANT12635723 (Capadona), and the National Institute for Biomedical Imaging and Bioengineering, T32EB004314, (Capadona/Kirsch). This material is based upon work supported by the National Science Foundation Graduate Research Fellowship under Grant No. GRANT12635723. Any opinion, findings, and conclusions or recommendations expressed in this material are those of the authors(s) and do not necessarily reflect the views of the National Science Foundation.

All procedures and animal care practices were reviewed, approved by, and performed in accordance with the Louis Stokes Cleveland Department of Veterans Affairs Medical Center Institutional Animal Care and Use Committee.
1. Surgical robot hardware setup
<ol>
	<li>Before surgery, follow the surgical robot (see Table of Materials) manual and guide to set up the hardware and software. Perform frame calibration as detailed in the manual. If the drill or frame are...

<ol>
	<li>Kilic, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+cranial+windows+for+long+term+multimodal+neurovascular+imaging+in+mice.">Chronic cranial windows for long term multimodal neurovascular imaging in mice.</a> Frontiers in Physiology. 11, 612678 (2020).</li><li>Goldey, G. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Removable+cranial+windows+for+long-term+imaging+in+awake+mice.">Removable cranial windows for long-term imaging in awake mice.</a> Nature Protocols. 9 (11), 2515-2538 (2014).</li><li>Augustinaite, S., Kuhn, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intrinsic+optical+signal+imaging+and+targeted+injections+through+a+chronic+cranial+window+of+a+head-fixed+mouse.">Intrinsic optical signal imaging and targeted injections through a chronic cranial window of a head-fixed mouse.</a> STAR Protocols. 2 (3), 100779 (2021).</li><li>Wang, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+skull-removed+chronic+cranial+window+for+ultrasound+and+photoacoustic+imaging+of+the+rodent+brain.">A skull-removed chronic cranial window for ultrasound and photoacoustic imaging of the rodent brain.</a> Frontiers in Neuroscience. 15, 673740 (2021).</li><li>Wang, Y., Xi, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+cranial+window+for+photoacoustic+imaging:+a+mini+review.">Chronic cranial window for photoacoustic imaging: a mini review.</a> Visual Computing for Industry, Biomedicine, and Art. 4 (1), 15 (2021).</li><li>Augustinaite, S., Kuhn, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+cranial+window+for+imaging+cortical+activity+in+head-fixed+mice.">Chronic cranial window for imaging cortical activity in head-fixed mice.</a> STAR Protocols. 1 (3), 100194 (2020).</li><li>Kunori, N., Takashima, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+implantable+cranial+window+using+a+collagen+membrane+for+chronic+voltage-sensitive+dye+imaging.">An implantable cranial window using a collagen membrane for chronic voltage-sensitive dye imaging.</a> Micromachines. 10 (11), 789 (2019).</li><li>Beckmann, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Longitudinal+deep-brain+imaging+in+mouse+using+visible-light+optical+coherence+tomography+through+chronic+microprism+cranial+window.">Longitudinal deep-brain imaging in mouse using visible-light optical coherence tomography through chronic microprism cranial window.</a> Biomedical Optics Express. 10 (10), 5235-5250 (2019).</li><li>Heo, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+soft,+transparent,+freely+accessible+cranial+window+for+chronic+imaging+and+electrophysiology.">A soft, transparent, freely accessible cranial window for chronic imaging and electrophysiology.</a> Scientific Reports. 6, 27818 (2016).</li><li>Holtmaat, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+neocortical+neurons+through+a+chronic+cranial+window.">Imaging neocortical neurons through a chronic cranial window.</a> Cold Spring Harbor Protocols. 2012 (6), 694-701 (2012).</li><li>Holtmaat, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=high-resolution+imaging+in+the+mouse+neocortex+through+a+chronic+cranial+window.">high-resolution imaging in the mouse neocortex through a chronic cranial window.</a> Nature Protocols. 4 (8), 1128-1144 (2009).</li><li>Sundaram, G. 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The use of EB dye and imaging is straightforward, quick, and useful for evaluating vascular damage in the brain for new methods and techniques. Whether using a surgical robot or confirming methods currently done in the lab, it is important to validate surgical methods to isolate the effects of experimental treatments vs. surgical impact and improve animal welfare. A thermocouple setup is also useful in evaluating drilling methods to ensure no heating occurs. Increases in temperature due to bone drilling have been known t...

Cranial windows have become ubiquitously used throughout the fields of neuroscience, neural engineering, and biology to allow for direct visualization and imaging of the cortex in living animals1,2,3,4,5,6,7,8,9,10,11. The powerful combination of transgenic mice and multiphoton imaging has provided extremely valuable insights into circuit activity and other biological insights in the in vivo brain12,13,14,15,16,17,18. Miniature microscopes mounted on the skull have further extended these capabilities to enable recordings in awake, freely moving animals19. The process of creating a cranial window requires power-drilling to thin or completely remove the cranial bone to produce large enough craniotomies to secure a transparent piece of glass over the cortex20. Polydimethylsiloxane (PDMS) and other polymers have also been tested as cranial window materials9,21. Ultimately, the ideal cranial window is one that does not alter or interfere with normal endogenous activity underneath. However, it is commonly accepted that cranial window drilling aggravates underlying tissue, leading to damage to the brain, disruption of the environment, and effecting meninges to the point of occluding multiphoton imaging depth22. The resulting neuroinflammation has a wide array of effects ranging from permeability of the blood-brain barrier (BBB), to activation and recruitment of glial cells around the implant site23. Therefore, characterizing safer and more reproducible cranial window drilling methods is crucial for consistent imaging quality and reducing confounding factors.
While care is taken to minimize trauma to the underlying tissue, the act of drilling the bone has the potential to cause both thermal and mechanical perturbations to the brain24,25. Mechanical trauma from accidental drill penetration into the dura may further induce varying degrees of cortical injury24. In a study by Shoffstall et al.25, the heat from bone-drilling resulted in an increased BBB permeability, as indicated by the presence of Evans Blue (EB) dye in the brain parenchyma25. EB dye, injected intravenously, binds to circulating albumin in the bloodstream and therefore does not normally cross a healthy BBB in appreciable concentrations. As a result, EB dye is commonly used as a sensitive marker of BBB permeability26,27. While their study did not directly measure the impact of the BBB permeability on subsequent biological sequelae under study, prior studies have correlated BBB permeability to an increased neuroinflammatory response to chronically implanted microelectrodes and alterations in motor function28.
Depending on the goals of the study, the magnitude of thermal and mechanical damage may contribute a source of experimental error, negatively affecting the rigor and reproducibility of the study. There are dozens of cited methods for producing cranial windows, each using different drilling equipment, speeds, techniques, and users1,2,3,4,5,6,7,8,9,10,11. Shoffstall et al.25 reported that the observed variation in the heating outcomes was attributed to variability in the drill&#39;s applied force, feed rate, and angle of application, among other aspects that cannot be controlled for when drilling by hand25. There is a belief that automated drilling systems and other stereotaxic equipment can improve reproducibility and outcome consistency, but published method studies have not rigorously evaluated temperature or BBB permeability as one of the outcomes. Therefore, there is a need for more reproducible and consistently applied methods to produce cranial windows, as well as methods rigorously applied to assess the impact of cranial window drilling on underlying neural tissue.
The focus of this study is to determine and develop consistent and safe drilling methods for cranial windows. The size of the craniotomy for cranial window installation is significantly larger than standard craniotomies for brain implanted microelectrodes. Such craniotomies cannot be completed with a single burr hole when using standard equipment, thereby introducing more inter-surgeon technique variability when performed by hand20. Surgical drilling robots have been introduced to the field, but have not been widely adopted1,6,29. Automation of drilling offers control over variables contributing to observed trial-to-trial variation, suggesting that use of the equipment can reduce inter- and intra-surgeon effects. This is of particular interest given the added difficulty of the larger craniotomy needed for cranial window placement. While one could assume there to be clear benefits to the control provided by automating the drilling, there has been little assessment of the implementation of these equipment. Although visible lesions have not been observed5, the higher sensitivity test using EB is desired.
Here, BBB permeability is measured using a commercially available surgical drilling robot with corresponding software, which allows for programming of stereotaxic coordinates, craniotomy planning/mapping, and a selection of drilling styles (&#34;point-by-point&#34; vs &#34;horizontal&#34;), referring to the routed path of the drill bit. Initially, eight &#34;seed&#34; points are drilled (Figure 1A), outlining the cranial window. From here, the space in between the seeds is cut out using either the &#34;point-by-point&#34; or &#34;horizontal&#34; drill method. &#34;Point-by-point&#34; performs vertical pilot hole cuts (similar to a CNC drill press), while &#34;horizontal&#34; performs horizontal cuts along the circumference of the cranial window that outline the hole (similar to a CNC router). The result for both methods are a piece of skull that can be removed to reveal the cranial window. To isolate damage from drilling, the cranial window is not physically removed, so as to avoid any additional damage. A combination of EB dye coupled with fluorescent imaging is used to measure BBB permeability after performing craniotomies in mice, and an inserted thermocouple is used to directly measure temperature of the brain surface during drilling (Figure 1B,C). Previous observations indicated that pulsed drilling on/off with 2 s intervals was sufficient to mitigate drill heating25, and therefore is incorporated into the experimental approach for the surgical robot.
The intent of the presented work is to demonstrate methods of assessing thermal damage from craniotomy drilling. While the methods are presented in the context of automated drilling, such methods can be applied to manual drilling schemes as well. These methods can be used to validate the use of equipment and/or drilling schemes before adopting as a standard procedure.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64188/64188fig01.jpg" /> 
Figure 1: Experimental pipeline schematic. Schematic demonstrating the process animals underwent for EB quantification post-cranial window procedure. (A) Schematic setup of the mouse with the stereotaxic frame and surgical robot drill. An example cranial window is shown over the motor cortex with seed points (green) and edge points (blue). (B) The perfusion setup includes injecting 1x Phosphate Buffered Saline (PBS) throughout the animal to remove any blood, followed by extraction of the brain. (C) The brain is then put into the EB fluorescent imaging system chamber to conduct fluorescent imaging on the Evans Blue dye. <a href="https://www.jove.com/files/ftp_upload/64188/64188fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1x Phosphate Buffered Saline 
			Type: Reagent</td>
			<td>VWR</td>
			<td>MRGF-6235</td>
			<td>For Evans Blue dilution</td>
		</tr>
		<tr>
			<td>Aura Software 
			Type: Tool</td>
			<td>Spectral Instruments Imaging</td>
			<td></td>
			<td>Open access imaging processing software for Lumina imaging sytems</td>
		</tr>
		<tr>
			<td>Buprenorphine 
			Type: Drug</td>
			<td>Sourced from Animal Facility</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Carbide Drill Bit, 0.6mm (Robot Drill) 
			Type: Tool</td>
			<td>Stoelting</td>
			<td>58640-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Carprofen 
			Type: Drug</td>
			<td>Sourced from Animal Facility</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cefazolin 
			Type: Drug</td>
			<td>Sourced from Animal Facility</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Evans Blue Dye 
			Type: Reagent</td>
			<td>Millipore Sigma</td>
			<td>E2129</td>
			<td>Reconstituted in 1x phosphate-buffered saline</td>
		</tr>
		<tr>
			<td>Isoflurane 
			Type: Drug</td>
			<td>Sourced from Animal Facility</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>IVIS Lumina II 
			Type: Tool</td>
			<td>Perkin Elmer</td>
			<td>CLS136334</td>
			<td>IVIS Lumina III currently in place of Lumina II on the market</td>
		</tr>
		<tr>
			<td>Jenco Linearizing Thermometer 
			Type: Tool</td>
			<td>Jenco</td>
			<td>765JF</td>
			<td>For Thermocouple setup</td>
		</tr>
		<tr>
			<td>Ketamine 
			Type: Drug</td>
			<td>Sourced from Animal Facility</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>LivingImage 
			Type: Tool</td>
			<td>Perkin Elmer</td>
			<td></td>
			<td>Software for IVIS Lumina III</td>
		</tr>
		<tr>
			<td>Marcaine 
			Type: Drug</td>
			<td>Sourced from Animal Facility</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Neurostar Software 
			Type: Tool</td>
			<td>Stoelting</td>
			<td></td>
			<td>Comes with surgical robot purchase</td>
		</tr>
		<tr>
			<td>Physiosuite with MouseSTAT&#174; Pulse Oximeter &#38; Heart Rate Monitor 
			Type: Tool</td>
			<td>Kent Scientific</td>
			<td>PS-03</td>
			<td>Used to monitor vitals</td>
		</tr>
		<tr>
			<td>PrismPlus mice 
			Type: Animal</td>
			<td>Jackson Labortory</td>
			<td>031478, RRID:IMSR_JAX:031478, Male, ~8 months old</td>
			<td>Animals used for the study</td>
		</tr>
		<tr>
			<td>Stoelting Drill and Injection Robot for Motorized Stereotaxic Instruments 
			Type: Tool</td>
			<td>Stoelting</td>
			<td>58640</td>
			<td>Main robotic drill with stereotaxic frame</td>
		</tr>
		<tr>
			<td>Thermocouple 
			Type: Tool</td>
			<td>TC Direct</td>
			<td>206-557</td>
			<td>For Thermocouple setup</td>
		</tr>
		<tr>
			<td>USB-6008 Multifunction I/O DAQ 
			Type: Tool</td>
			<td>National Instruments</td>
			<td>USB-6008</td>
			<td>For Thermocouple setup</td>
		</tr>
		<tr>
			<td>Xylazine 
			Type: Drug</td>
			<td>Sourced from Animal Facility</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

assessment of thermal damage from robot-drilled craniotomy for cranial window surgery in mice

Cranial window surgery allows for the imaging of brain tissue in live mice with the use of multiphoton or other intravital imaging techniques. However, when performing any craniotomy by hand, there is often thermal damage to brain tissue, which is inherently variable surgery-to-surgery and may be dependent on individual surgeon technique. Implementing a surgical robot can standardize surgery and lead to a decrease in thermal damage associated with surgery. In this study, three methods of robotic drilling were tested to evaluate thermal damage: horizontal, point-by-point, and pulsed point-by-point. Horizontal drilling utilizes a continuous drilling schematic, while point-by-point drills several holes encompassing the cranial window. Pulsed point-by-point adds a &#34;2 s on, 2 s off&#34; drilling scheme to allow for cooling in between drilling. Fluorescent imaging of Evans Blue (EB) dye injected intravenously measures damage to brain tissue, while a thermocouple placed under the drilling site measures thermal damage. Thermocouple results indicate a significant decrease in temperature change in the pulsed point-by-point (6.90 &#176;C &#177; 1.35 &#176;C) group compared to the horizontal (16.66 &#176;C &#177; 2.08 &#176;C) and point-by-point (18.69 &#176;C &#177; 1.75 &#176;C) groups. Similarly, the pulsed point-by-point group also showed significantly less EB presence after cranial window drilling compared to the horizontal method, indicating less damage to blood vessels in the brain. Thus, a pulsed point-by-point drilling method appears to be the optimal scheme for reducing thermal damage. A robotic drill is a useful tool to help minimize training, variability, and reduce thermal damage. With the expanding use of multiphoton imaging across research labs, it is important to improve the rigor and reproducibility of results. The methods addressed here will help inform others of how to better use these surgical robots to further advance the field.

Thermal evaluation 
Potential for thermal damage was evaluated by measuring the change in temperature from baseline due to drilling using horizontal (Figure 2A), point-by-point (Figure 2B), and pulsed point-by-point (Figure 2C) methods. Figure 2D displays the experimental setup for obtaining thermal data. A sample size of N = 4 cranial windows was used for thermal evaluation. Horiz...

Watch this Scientific Journal Video about Assessment of Thermal Damage from Robot-Drilled Craniotomy for Cranial Window Surgery in Mice at JoVE.com

Assessment of Thermal Damage from Robot-Drilled Craniotomy for Cranial Window Surgery in Mice

Cranial windows have become a ubiquitously implemented surgical technique to allow for intravital imaging in transgenic mice. This protocol describes the use of a surgical robot that performs semi-automated bone drilling of cranial windows and can help reduce surgeon-to-surgeon variability and partially mitigate thermal blood-brain barrier damage.

Cranial window surgery allows for the imaging of brain tissue in live mice with the use of multiphoton or other intravital imaging techniques. However, ...

This protocol provides a method for evaluating vascular damage as a result of cranial window drilling. A surgical robot offers improved consistencies across procedures while reducing the amount of training to learn the technique. During this procedure, we utilized cranial injections which can be very challenging.We recommend extensive training for those new to the technique. To begin, set up the system as described in the manuscript and perform frame calibration following the surgical robot manual. Navigate to the surgical software and create a new project by selecting start with a clean project.Then set the subject as mouse at the top to designate the drilling coordinate atlas to be used. Next, select start new project, and click on planning in the bottom left corner to navigate to the drilling coordinate planning screen. Create the drilling scheme for the cranial window technique to be performed by clicking anywhere on the stereotaxic atlas.Use bregma as the reference and input the coordinates for the motor cortex as described in the manuscript. Press Enter on the keyboard to update the selected coordinates. Click store target to save these coordinates and input an appropriate name.Then click on the move button in the bottom left to navigate back to the main drilling screen. Click on tools, then project, and save as option to reuse this template project for later projects, as it will automatically retain the drilling coordinates. First, administer subcutaneous injections of antibiotics, cefazolin, analgesic, carprofen, and buprenorphine to the anesthetized mouse.Next, place the mouse on a heating pad and maintain anesthesia with 0.5%to 2%isoflurane via inhalation through a nose cone. Mount the animal on the surgical robot stereotaxic frame using supplied ear bars and routinely assess vitals and anesthetic depth via toe pinch. Then scrub the surgical area with chlorhexidine gluconate and 70%isopropanol for sterilization.And place a sterile plastic wrap over the mouse and stereotactic frame to maintain sterility during surgery. Using a scalpel blade, perform a one-inch incision on the midline of the skull, starting at the back of the eyes, dry out and clean the skull using 3%hydrogen peroxide with cotton tipped applicators. To prepare the tail for easy injection, wipe down with alcohol.Using the thumb and forefinger, bend the tail to expose the tail vein on top of the bend of the tail. Insert the syringe parallel with the vein, and slowly inject the volume of Evans blue. Once injected, wait for five minutes to allow Evans blue to circulate.A successful injection is immediately verified as the mouse's extremities and surgical window turn blue. Once the skull is prepared for drilling, navigate back to the surgical software and click on tools. Then select project, followed by new, and select a template project from the dropdown menu to choose a template project.Select same protocol elements. Click on planning, then select drill parameters to carry over to this new project and click start new project. Next, correct the drill and frame to account for the tilt and scaling of the mouse skull of the current animal by clicking tools, and select correct for tilt and scaling to open the corrections screen.At the top of the screen, ensure that the drill is active by clicking on the light red drill button After correction has been performed, come out of the correction window by clicking close in the bottom middle of the screen and navigate to the drilling screen by clicking tools and then selecting drill to begin the drilling procedure. Confirm the drill site by selecting the craniotomy shape window at the top of the screen, followed by viewing the drilling target details in the dropdown below. To carry out manual pulse drilling, turn off the auto-stop feature by unchecking the checkbox next to the auto-stop option in the drill menu, which allow for controlling when the drill is off for the pulsing.In the drill menu, select 100 micrometers as the drill depth advancement. Open up the drill site menu and navigate to the first drill site. Use the advance button or manual controls to lower the drill until it touches the skull and press set surface once the drill bit reaches the skull.Once ready, click advance to commence drilling. And after the drill has advanced 100 micrometers, press Escape twice to stop the drill. After two seconds, repeat this cycle for the depth of the skull.For each seed and edge point during drilling, ensure to set dura using the button in the drill menu once the dura has been reached. Measure the changes in temperature of the skull and brain using a thermocouple in combination with the three different drilling schemes connected to a data acquisition system, that allows for the measurement to be read into MATLAB. Mount a cadaver mouse onto the stereotaxic frame and robotic drill set up.Manually drill a small hole approximately two millimeter away from where the cranial window will be made into the side of the skull, which allows for thermocouple to be slid into position under where drilling of the cranial window occurs. Begin the drilling process for each of the three schemes as demonstrated previously, resulting in spikes due to temperature change, indicating heating occurring near the brain as the drill goes through the skull. Potential for thermal damage was evaluated by measuring the change in temperature from baseline due to drilling using horizontal, point by point, and pulsed point by point methods.Both the horizontal and point by point drilling schemes show non-significant differences in thermal changes. However, changing to a pulsed point by point method resulted in significantly less heating of the brain than both horizontal and point by point drilling. The horizontal edge drilling took 300 seconds, while point by point edge drilling took 200 seconds.The pulse method took the longest with seed and edge drilling, taking approximately 500 seconds each. To monitor the relationship between the drilling scheme and vascular damage, Evans blue fluorescent imaging was performed. The drilling via horizontal and point by point cutting was seen to be damaging to the vasculature in the brain.In comparison to the control groups, the pulsed point by point method had less localized damage at the seed and edge point, but still had visible Evans blue presence within the cranial window. When manually drilling it is crucial to check the craniotomy after advancing the drill to ensure the dura is not breached. In addition to cranial windows, a surgical robot can be used for standard craniotomies to improve surgical outcomes.

Research

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Neuroscience

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