We thank Yuri Shinomoto for providing animal care, training, and awake recordings. The ECoG arrays were manufactured by Cir-Tech (www.cir-tech.co.jp). Furthermore, we would like to thank Editage (www.editage.jp) for English language editing. This work was supported by the Brain Mapping by Integrated Neurotechnologies for Disease Studies (Brain/MINDS), the Japan Agency for Medical Research and development (AMED) (JP18dm0207001), the Brain Science Project of the Center for Novel Science Initiatives (CNSI), the National Institutes of Natural Sciences (NINS) (BS291004, M.K.), and by the Japan Society for the Promotion of Science (JSPS) KAKENHI (JP17H06034, M.K.).

This protocol has been performed on 6 common marmosets (4 males, 2 females; body weight = 320-470 g; age = 14-53 months). All procedures were carried out in accordance with the recommendations of the National Institutes of Health Guidelines for the Care and Use of Laboratory Animals. The protocol was approved by the RIKEN Ethical Committee (No. H28-2-221(3)). All surgical procedures were performed under anesthesia, and all efforts were made to minimize the number of animals used as well as their discomfort.
1. Preparation
<ol>
	<li>Obtain a structural magnetic resonance image (MRI) of each individual brain. This will be used to identify electrode positions through registration with a marmoset brain atlas and computer tomography (CT).</li>
	<li>Preparation of the ECoG array: prepare a customized multichannel ECoG array (Figure 1A). A 96ch ECoG array consists of two sheets with 32 and 64 electrodes. To accommodate individual differences in brain size, the ECoG array has a flexible arm. The arm can cover the temporal pole, depending on individual brain shape. Place the reference electrodes facing opposite to the ECoG electrodes and the ground electrodes facing the same direction.
	<ol>
		<li>Assemble the ECoG array with a connector case (Figure 1B) and seal gaps of connector (Figure 1C) using acrylic glue to prevent the inflow of liquid during surgery. Sterilize the array with ethylene oxide gas.</li>
	</ol>
	</li>
	<li>Prepare and sterilize instruments. 
	NOTE: All instruments used are listed in the Table of Materials.</li>
</ol>
2. Implantation of ECoG Array
NOTE: Withdraw ingestion of food and liquids greater than 4 h prior to surgery. Perform all surgical steps with aseptic technique using sterilized gloves and instruments.
<ol>
	<li>Pre-implant procedures
	<ol>
		<li>Induce anesthesia in the marmoset by intramuscular (i.m.) injection of ketamine (15 mg/kg) 5 min after i.m. atropine (0.08 mg/kg) injection.</li>
		<li>Anesthetize and maintain anesthesia using isoflurane (1-3% diluted with a mixture of oxygen/nitrous oxide)&#160;depending on the physiological state of the animal, which should be continuously monitored. Ensure that heart rate is 130-180 BPM and monitor body temperature and arterial blood oxygen saturation (SpO2) continuously to judge the animal&#8217;s condition.</li>
		<li>Shave the top of the animal&#8217;s head with clippers and a hair remover. Fully rinse hair-removal cream off the skin with wet gauze, or it will cause skin damage.</li>
		<li>Administer an antibiotic (cefovecin; 16 mg/kg s.c.), antihypertensive (furosemide; 2.0 mg/kg i.m.), and antihemorrhagic (carbazochrome sodium sulfonate hydrate; 0.2 mg/kg i.m.).</li>
		<li>Place the animal on a stereotaxic frame. At this time, apply 2% lidocaine jelly to the ear-bars and ophthalmic ointment to the eyes to prevent dryness and postoperative pain.</li>
		<li>Disinfect the surgical area with iodine solution and cover it with sterilized drapes.&#160;Apply 2% lidocaine jelly to the place of the skin incision.</li>
	</ol>
	</li>
	<li>Implantation procedures
	<ol>
		<li>Incise skin about 4 cm through the midline of the scalp with a scalpel. Detach the temporal muscle from the skull with a curette until all of the surgical area is exposed. Clean out tissues on the skull surface and stop the bleeding completely with pressure hemostasis, and with bone wax, if necessary. Wrap the edge of the skin and muscles with moistened gauze. Keep the gauze moistened during surgery.</li>
		<li>Place the frontal edge of the array onto the edge of the frontal pole. Mark a planned area for the craniotomy, slits, and holes on the skull with a sterile pencil. The craniotomy location will depend on the design of the array (Figure 2).</li>
		<li>Drill the craniotomy along mark 1, as shown in Figure 2. While drilling the bone, blow air at the cutting edge to maintain a clear view for the surgeon. Next, cut the bone all the way around mark 2, as the bone piece will still be attached to dura at the center. Lift the piece up gently from one edge and peel off the dura with a spatula. This process must be conducted slowly and carefully, or it will tear the dura easily.
		<ol>
			<li>Remove the bone tips from the bone piece and wrap the piece with moistened gauze, as this piece will be returned after implanting the array.</li>
		</ol>
		</li>
		<li>Perform craniotomy 3 and 4 as shown in Figure 2. These allow the insertion of electrodes into the orbitofrontal and occipital areas, respectively.</li>
		<li>Drill slits on mark 5 as shown in Figure 2. These slits allow examination of the array to ensure that it is properly inserted.</li>
		<li>The dura will now be exposed. Wash the area with saline and stop the bleeding with pressure hemostasis and a gelatin sponge, if necessary. The edge of the open craniotomy may need to be cleaned with a curette or bone rongeur.</li>
		<li>Make the slits (marked 6 in Figure 2) into which the reference electrodes are placed. Place the reference electrodes in the epidural space at the contra-lateral sensorimotor and occipital areas. The position should be determined according to specific experimental needs.</li>
		<li>Drill screw holes at four points around each stem of the connector with a 1.0 mm screw (crosses in Figure 2). To prevent damage to the dura matter, insert a spatula under the skull. These holes should be orthogonal against the skull. Then, install PEEK screws (1.4 x 2.5 mm) as anchors to fix the connector to the skull.</li>
		<li>Insert the ECoG array into the epidural space. Use flathead forceps to hold the array. 
		NOTE: The array should be inserted without bending. If the array is bent, create an appropriate space by inserting a spatula between the skull and dura. If the bending was caused by the relatively small size of the brain, cut off some of the electrodes.</li>
		<li>Fix the reference and ground electrodes with a dental acrylic. Place the reference electrodes in the epidural space and ground electrodes on the cranial surface. Both contacts should face the skull.</li>
		<li>Put the bone piece back and fix the connector and head post to the skull with dental acrylic on the screws.</li>
		<li>Suture the skin with 6-0 nylon at the forehead and rear head, and fix the skin to the sides of the connector using skin closures.</li>
	</ol>
	</li>
	<li>Post-implantation procedures
	<ol>
		<li>Remove the animal from the stereotaxic frame. Ensure that the animal is kept warm and provided with oxygen during the following steps.</li>
		<li>Immediately after surgery, inject the animal with meloxicam (0.3 mg/kg i.m.) to decrease postoperative pain. Administer an anti-inflammatory corticosteroid (dexamethasone; 2.0 mg/kg i.m.) and subcutaneous infusion (lactated Ringer&#8217;s solution; 5.0 mL), including famotidine (0.5 mg/kg) as a gastroprotectant. 
		NOTE: A concurrent use of NSAIDs with steroids has a potential for gastrointestinal side effects.</li>
		<li>After the animal has recovered (confirm by heart rate and SpO2), remove vital sign monitoring and transfer the animal into the ICU for 2-3 days.</li>
	</ol>
	</li>
</ol>
3. Postoperative Treatment
NOTE: It typically takes 5 days for animals to recover completely from the surgery.
<ol>
	<li>To prevent brain swelling, administer the anti-inflammatory corticosteroid dexamethasone (2.0 mg/kg) twice a day on the first day after surgery. Then, reduce the dose to 1.5 mg/kg twice a day on the second and third days, and 1 mg/kg twice a day on the fourth day.</li>
	<li>Administer pain relief (meloxicam; 0.1 mg/kg oral; once a day) and an antihemorrhagic (carbazochrome sodium sulfonate hydrate; 0.2 mg/kg i.m.; twice a day) for 5 days after surgery. 
	NOTE: In our case, 1-2 days after the surgery, some marmosets (3 out of 6) became less active and vomited. This may have been caused by increased intracranial pressure due to a blood clot. When marmosets presented these symptoms, we reopened the head and removed the clot under general anesthesia (alfaxalone). If there was no bending of the ECoG array during the implantation, the blood clot was likely in the space between the array and where the bone piece was returned. In this case, the blood clot can be washed away by running saline into the space using a catheter. This procedure usually leads to recovery in the animal.</li>
	<li>Identification of electrode locations
	<ol>
		<li>Around 1 week after surgery, perform a computer tomography (CT) scan of the animal&#8217;s head. 
		NOTE: This is a good opportunity to check if signals can be recorded properly. Open the connector case and remove any blood clots if they are present.</li>
		<li>Align T2-weighted MRI to stereotaxic coordinates using AFNI software18 (https://afni.nimh.nih.gov) (Figure 3A). Align the CT image to T2-weighted anatomical magnetic resonance images with AFNI (Figure 3B). Register a marmoset brain atlas to MRI (Figure 3C) with AFNI and ANTS19.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Fukushima, M., Chao, Z. C., Fujii, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Studying+brain+functions+with+mesoscopic+measurements:+Advances+in+electrocorticography+for+non-human+primates.">Studying brain functions with mesoscopic measurements: Advances in electrocorticography for non-human primates.</a> Current Opinion in Neurobiology. 32, 124-131 (2015).</li><li>Nagasaka, Y., Shimoda, K., Fujii, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multidimensional+recording+(MDR)+and+data+sharing:+an+ecological+open+research+and+educational+platform+for+neuroscience.">Multidimensional recording (MDR) and data sharing: an ecological open research and educational platform for neuroscience.</a> PLoS One. 6 (7), e22561 (2011).</li><li>Fukushima, M., 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=An+electrocorticographic+electrode+array+for+simultaneous+recording+from+medial,+lateral,+and+intrasulcal+surface+of+the+cortex+in+macaque+monkeys.">An electrocorticographic electrode array for simultaneous recording from medial, lateral, and intrasulcal surface of the cortex in macaque monkeys.</a> Journal of Neuroscience Methods. 233, 155-165 (2014).</li><li>Komatsu, M., Sugano, E., Tomita, H., Fujii, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Chronically+Implantable+Bidirectional+Neural+Interface+for+Non-human+Primates.">A Chronically Implantable Bidirectional Neural Interface for Non-human Primates.</a> Frontiers in Neuroscience. 11, 514 (2017).</li><li>Komatsu, M., Takaura, K., Fujii, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mismatch+negativity+in+common+marmosets:+Whole-cortical+recordings+with+multi-channel+electrocorticograms.">Mismatch negativity in common marmosets: Whole-cortical recordings with multi-channel electrocorticograms.</a> Scientific Reports. 5, 15006 (2015).</li><li>Sasaki, E., 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=Generation+of+transgenic+non-human+primates+with+germline+transmission.">Generation of transgenic non-human primates with germline transmission.</a> Nature. 459 (7246), 523-527 (2009).</li><li>Okano, H., 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=Brain/MINDS:+A+Japanese+National+Brain+Project+for+Marmoset+Neuroscience.">Brain/MINDS: A Japanese National Brain Project for Marmoset Neuroscience.</a> Neuron. 92 (3), 582-590 (2016).</li><li>de la Mothe, L. A., Blumell, S., Kajikawa, Y., Hackett, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cortical+connections+of+auditory+cortex+in+marmoset+monkeys:+lateral+belt+and+parabelt+regions.">Cortical connections of auditory cortex in marmoset monkeys: lateral belt and parabelt regions.</a> Anatomical Record. 295 (5), 800-821 (2012).</li><li>Kaas, J. H., Hackett, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subdivisions+of+auditory+cortex+and+processing+streams+in+primates.">Subdivisions of auditory cortex and processing streams in primates.</a> Proceedings of National Academy of Sciences of the United States of America. 97 (22), 11793-11799 (2000).</li><li>Ghahremani, M., Hutchison, R. M., Menon, R. S., Everling, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Frontoparietal+Functional+Connectivity+in+the+Common+Marmoset.">Frontoparietal Functional Connectivity in the Common Marmoset.</a> Cerebral Cortex. , (2016).</li><li>Belcher, A. M., 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=Functional+Connectivity+Hubs+and+Networks+in+the+Awake+Marmoset+Brain.">Functional Connectivity Hubs and Networks in the Awake Marmoset Brain.</a> Frontiers in Integrative Neuroscience. 10, 9 (2016).</li><li>Mitchell, J. F., Leopold, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+marmoset+monkey+as+a+model+for+visual+neuroscience.">The marmoset monkey as a model for visual neuroscience.</a> Neuroscience Research. 93, 20-46 (2015).</li><li>Solomon, S. G., Rosa, M. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simpler+primate+brain:+the+visual+system+of+the+marmoset+monkey.">A simpler primate brain: the visual system of the marmoset monkey.</a> Frontiers in Neural Circuits. 8, 96 (2014).</li><li>Burman, K. J., Palmer, S. M., Gamberini, M., Rosa, M. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytoarchitectonic+subdivisions+of+the+dorsolateral+frontal+cortex+of+the+marmoset+monkey+(Callithrix+jacchus),+and+their+projections+to+dorsal+visual+areas.">Cytoarchitectonic subdivisions of the dorsolateral frontal cortex of the marmoset monkey (Callithrix jacchus), and their projections to dorsal visual areas.</a> Journals of Comparative Neurology. 495 (2), 149-172 (2006).</li><li>Bakola, S., Burman, K. J., Rosa, M. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+cortical+motor+system+of+the+marmoset+monkey+(Callithrix+jacchus).">The cortical motor system of the marmoset monkey (Callithrix jacchus).</a> Neuroscience Research. 93, 72-81 (2015).</li><li>Krubitzer, L. A., Kaas, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+organization+and+connections+of+somatosensory+cortex+in+marmosets.">The organization and connections of somatosensory cortex in marmosets.</a> Journal of Neuroscience. 10 (3), 952-974 (1990).</li><li>Miller, C. T., 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=Marmosets:+A+Neuroscientific+Model+of+Human+Social+Behavior.">Marmosets: A Neuroscientific Model of Human Social Behavior.</a> Neuron. 90 (2), 219-233 (2016).</li><li>Cox, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=AFNI:+software+for+analysis+and+visualization+of+functional+magnetic+resonance+neuroimages.">AFNI: software for analysis and visualization of functional magnetic resonance neuroimages.</a> Computers and Biomedical Research. 29 (3), 162-173 (1996).</li><li>Avants, B. B., 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+reproducible+evaluation+of+ANTs+similarity+metric+performance+in+brain+image+registration.">A reproducible evaluation of ANTs similarity metric performance in brain image registration.</a> Neuroimage. 54 (3), 2033-2044 (2011).</li><li>Hashikawa, T., Nakatomi, R., Iriki, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Current+models+of+the+marmoset+brain.">Current models of the marmoset brain.</a> Neuroscience Research. 93, 116-127 (2015).</li></ol>

MK is applying for a patent on whole-cortical ECoG array she have used in this protocol (No. 2018-210975).

For successful implantation, animals should be provided with adequate nutrition before and after surgery. Short operating time is also important to optimize the animal&#39;s recovery. Preparations should be finished at least one day before surgery. To reduce operating time, previous craniotomy training with electrode array insertion in terminated animals for other experimental purposes is recommended.&#160;Table 1 shows an example of the time course for this protocol.
We modif...

Cognition requires the coordination of neural ensembles across widespread brain networks, particularly the neocortex that is well-developed in humans and believed to be involved in higher cognitive behaviors. However, how the neocortex achieves this cognitive behavior is an unsolved issue in the neuroscience field. Recent development of thin, flexible electrocorticographic (ECoG) electrodes enables conduction of stable recordings from large-scale cortical activity1. Fujii and colleagues have developed a whole-cortical ECoG array for macaque monkeys2,3. The array continuously covers almost the entire lateral cortex, from the occipital pole to the temporal and frontal poles, and captures whole-cortical neural activity in one shot. We have further developed this system for application in the common marmoset4,5, a small, new-world monkey with genetic manipulability6,7. This animal has several advantages compared to other species. The visual, auditory, somatosensory, motor, and frontal cortical areas of this species have been previously mapped and reported to have basic homologous organization to the same areas in humans and macaques8,9,10,11,12,13,14,15,16. Their brains are smooth, and most lateral cortical areas are exposed to the surface of the cortex, which is harder to access with ECoG in macaques. Based on these features, the marmoset is suitable for electrocorticographic studies. Furthermore, marmosets exhibit social behaviors and have been proposed to serve as a candidate model of human social behaviors17.
This protocol describes an epidural implantation procedure of the ECoG array on the whole lateral surface of the cortex in a common marmoset. It provides an opportunity to monitor large-scale cortical activity for primate cortical neuroscience, including sensory, motor, higher cognitive, and social domains.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Beaker (100 cc)</td>
			<td></td>
			<td></td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Cotton ball</td>
			<td></td>
			<td></td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Absorption triangles</td>
			<td>Fine Science Tools Inc.</td>
			<td>18105-03</td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Cotton swab with fine tip</td>
			<td>Clean Cross Co., Ltd.</td>
			<td>HUBY340 BB-013</td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Gauze</td>
			<td></td>
			<td></td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Towel forceps</td>
			<td></td>
			<td></td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Scalpel handle</td>
			<td></td>
			<td></td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Needle Holder</td>
			<td></td>
			<td></td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Iris Scissor</td>
			<td></td>
			<td></td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Micro-Mosquito Forceps</td>
			<td></td>
			<td></td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Adson, 1x2 teeth</td>
			<td></td>
			<td></td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Bone Curette</td>
			<td></td>
			<td></td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Micro spatura</td>
			<td>Fine Science Tools Inc.</td>
			<td>10091-12</td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Needle Holders, 12.5cm, Curved, Smooth Jaws</td>
			<td>World Precision Instruments</td>
			<td>14132</td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Vessel Dilator, 12cm, 0.1mm tip</td>
			<td>Fine Science Tools Inc.</td>
			<td>18131-12</td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Vessel Dilator, 12cm, 0.2 mm tip</td>
			<td>Fine Science Tools Inc.</td>
			<td>18132-12</td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Fine-tipped rongeur</td>
			<td>Fine Science Tools Inc.</td>
			<td>16221-14</td>
			<td>Outocrave</td>
		</tr>
		<tr>
			<td>Manipurator of a stereotaxic frame</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Wrench for the manipurator</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Hand-made fixture for the connector</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Silicon cup for dental acril</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Silicon cup hlder</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Paintbrush</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Pencil</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Micro screw, 1.4 mm x 2.0 mm</td>
			<td>Nippon Chemical Screw Co., Ltd.</td>
			<td>PEEK/MPH-M1.4-L2</td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Screw driver for the micro screw</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Micromotor handpiece of a drill</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Stainless steel burr, 1.4 mm</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Stainless steel burr, 1.0 mm</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Drill bit, 1.2 mm</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
		<tr>
			<td>Rubber air blower</td>
			<td></td>
			<td></td>
			<td>Gas sterilization</td>
		</tr>
	</tbody>
</table>

chronic implantation of whole-cortical electrocorticographic array in the common marmoset

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent development of thin, flexible ECoG electrodes has enabled conduction of stable recordings of large-scale cortical activity. We have developed a whole-cortical ECoG array for the common marmoset. The array continuously covers almost the entire lateral surface of cortical hemisphere, from the occipital pole to the temporal and frontal poles, and it captures whole-cortical neural activity in one shot. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain. Marmosets have two advantages regarding ECoG recordings, one being the homologous organization of anatomical structures in humans and macaques, including frontal, parietal, and temporal complexes. The other advantage is that the marmoset brain is lissencephalic and contains a large number of complexes, which are more difficult to access in macaques with ECoG, that are exposed to the brain surface.These features allow direct access to most cortical areas beneath the surface of the brain. This system provides an opportunity to investigate global cortical information processing with high resolutions at a sub-millisecond order in time and millimeter order in space.

The whole-cortical ECoG array can simultaneously capture neuronal activity from the entirety of a hemisphere. Figure 4 shows examples of auditory evoked potentials (AEPs) from multiple auditory areas in an awake marmoset. ECoG recordings were conducted in passive listening conditions. Each marmoset was exposed to auditory stimuli, which consisted of randomized pure tones with 20 types of frequency. Then, we calculated AEPs by averaging ECoGs aligned with onse...

Watch this Scientific Journal Video about Chronic Implantation of Whole-cortical Electrocorticographic Array in the Common Marmoset at JoVE.com

Chronic Implantation of Whole-cortical Electrocorticographic Array in the Common Marmoset

We have developed a whole-cortical electrocorticographic array for the common marmoset that continuously covers almost the entire lateral surface of cortex, from the occipital pole to the temporal and frontal poles. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain.

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent ...

chronic-implantation-whole-cortical-electrocorticographic-array

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

9.5K Views.  RIKEN Center for Brain Science. We have developed a whole-cortical electrocorticographic array for the common marmoset that continuously covers almost the entire lateral surface of cortex, from the occipital pole to the temporal and frontal poles. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain.

Video: Chronic Implantation of Whole-cortical Electrocorticographic Array in the Common Marmoset

Modeling Neural Immune Signaling of Episodic and Chronic Migraine Using Spreading Depression In Vitro

Migraine and its transformation to chronic migraine are healthcare burdens in need of improved treatment options. We seek to define how neural immune signaling modulates the susceptibility to migraine, modeled in vitro using spreading depression (SD), as a means to develop novel therapeutic targets for episodic and chronic migraine. SD is the likely cause of migraine aura and migraine pain. It is a paroxysmal loss of neuronal function triggered by initially increased neuronal activity, which slowly propagates within susceptible brain regions. Normal brain function is exquisitely sensitive to, and relies on, coincident low-level immune signaling. Thus, neural immune signaling likely affects electrical activity of SD, and therefore migraine. Pain perception studies of SD in whole animals are fraught with difficulties, but whole animals are well suited to examine systems biology aspects of migraine since SD activates trigeminal nociceptive pathways. However, whole animal studies alone cannot be used to decipher the cellular and neural circuit mechanisms of SD. Instead, in vitro preparations where environmental conditions can be controlled are necessary. Here, it is important to recognize limitations of acute slices and distinct advantages of hippocampal slice cultures. Acute brain slices cannot reveal subtle changes in immune signaling since preparing the slices alone triggers: pro-inflammatory changes that last days, epileptiform behavior due to high levels of oxygen tension needed to vitalize the slices, and irreversible cell injury at anoxic slice centers. 
In contrast, we examine immune signaling in mature hippocampal slice cultures since the cultures closely parallel their in vivo counterpart with mature trisynaptic function; show quiescent astrocytes, microglia, and cytokine levels; and SD is easily induced in an unanesthetized preparation. Furthermore, the slices are long-lived and SD can be induced on consecutive days without injury, making this preparation the sole means to-date capable of modeling the neuroimmune consequences of chronic SD, and thus perhaps chronic migraine. We use electrophysiological techniques and non-invasive imaging to measure neuronal cell and circuit functions coincident with SD. Neural immune gene expression variables are measured with qPCR screening, qPCR arrays, and, importantly, use of cDNA preamplification for detection of ultra-low level targets such as interferon-gamma using whole, regional, or specific cell enhanced (via laser dissection microscopy) sampling. Cytokine cascade signaling is further assessed with multiplexed phosphoprotein related targets with gene expression and phosphoprotein changes confirmed via cell-specific immunostaining. Pharmacological and siRNA strategies are used to mimic and modulate SD immune signaling.

Modeling Neural Immune Signaling of Episodic and Chronic Migraine Using Spreading Depression In Vitro

Migraine and its transformation to chronic migraine are immense healthcare burdens in need of improved treatment options. We seek to define how neural immune signaling modulates the susceptibility to migraine, modeled in vitro using spreading depression in hippocampal slice cultures, as a means to develop novel therapeutic targets.

Migraine and its transformation to chronic migraine are healthcare burdens in need of improved treatment options.  We seek to define how neural immune ...

Surgical Implantation of Chronic Neural Electrodes for Recording Single Unit Activity and Electrocorticographic Signals

The success of long-term electrophysiological recordings often depends on the quality of the implantation surgery. Here we provide useful information for surgeons who are learning the process of implanting electrode systems. We demonstrate the implantation procedure of both a penetrating and a surface electrode. The surgical process is described from start to finish, including detailed descriptions of each step throughout the procedure. It should also be noted that this video guide is focused towards procedures conducted in rodent models and other small animal models. Modifications of the described procedures are feasible for other animal models.

We provide useful information for surgeons who are learning the process of implanting chronic neural recording electrodes. Techniques for both penetrating and surface electrode systems are described in a rodent animal model.

The success of long-term electrophysiological recordings often depends on the quality of the implantation surgery.  Here we provide useful information for ...

Fiber-optic Implantation for Chronic Optogenetic Stimulation of Brain Tissue

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard for analyzing patterns of synaptic connectivity, but paired electrophysiological recordings can be both cumbersome and experimentally limiting. The development of optogenetics has introduced an elegant method to stimulate neurons and circuits, both in vitro1 and in vivo2,3. By exploiting cell-type specific promoter activity to drive opsin expression in discrete neuronal populations, one can precisely stimulate genetically defined neuronal subtypes in distinct circuits4-6. Well described methods to stimulate neurons, including electrical stimulation and/or pharmacological manipulations, are often cell-type indiscriminate, invasive, and can damage surrounding tissues. These limitations could alter normal synaptic function and/or circuit behavior. In addition, due to the nature of the manipulation, the current methods are often acute and terminal. Optogenetics affords the ability to stimulate neurons in a relatively innocuous manner, and in genetically targeted neurons. The majority of studies involving in vivo optogenetics currently use a optical fiber guided through an implanted cannula6,7; however, limitations of this method include damaged brain tissue with repeated insertion of an optical fiber, and potential breakage of the fiber inside the cannula. Given the burgeoning field of optogenetics, a more reliable method of chronic stimulation is necessary to facilitate long-term studies with minimal collateral tissue damage. Here we provide our modified protocol as a video article to complement the method effectively and elegantly described in Sparta et al.8 for the fabrication of a fiber optic implant and its permanent fixation onto the cranium of anesthetized mice, as well as the assembly of the fiber optic coupler connecting the implant to a light source. The implant, connected with optical fibers to a solid-state laser, allows for an efficient method to chronically photostimulate functional neuronal circuitry with less tissue damage9 using small, detachable, tethers. Permanent fixation of the fiber optic implants provides consistent, long-term in vivo optogenetic studies of neuronal circuits in awake, behaving mice10 with minimal tissue damage.

The development of optogenetics now provides the means to precisely stimulate genetically defined neurons and circuits, both in vitro and in vivo. Here we describe the assembly and implantation of a fiber optic for chronic photostimulation of brain tissue.

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard ...

Implantation of Chronic Silicon Probes and Recording of Hippocampal Place Cells in an Enriched Treadmill Apparatus

An important requisite for understanding brain function is the identification of behavior and cell activity correlates. Silicon probes are advanced electrodes for large-scale electrophysiological recording of neuronal activity, but the procedures for their chronic implantation are still underdeveloped. The activity of hippocampal place cells is known to correlate with an animal&#39;s position in the environment, but the underlying mechanisms are still unclear. To investigate place cells, here we describe a set of techniques which range from the fabrication of devices for chronic silicon probe implants to the monitoring of place field activity in a cue-enriched treadmill apparatus. A micro-drive and a hat are built by fitting and fastening together 3D-printed plastic parts. A silicon probe is mounted on the micro-drive, cleaned, and coated with dye. A first surgery is performed to fix the hat on the skull of a mouse. Small landmarks are fabricated and attached to the belt of a treadmill. The mouse is trained to run head-fixed on the treadmill. A second surgery is performed to implant the silicon probe in the hippocampus, following which broadband electrophysiological signals are recorded. Finally, the silicon probe is recovered and cleaned for reuse. The analysis of place cell activity in the treadmill reveals a diversity of place field mechanisms, outlining the benefit of the approach.

We describe the diverse steps to implant chronic silicon probes and to record place cells in mice that are running head-fixed on a cue-enriched treadmill apparatus.

An important requisite for understanding brain function is the identification of behavior and cell activity correlates. Silicon probes are advanced ...

Laminectomy and Spinal Cord Window Implantation in the Mouse

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated digital vaporizer is utilized to achieve a stable plane of anesthesia at a low-flow rate of isoflurane. A single vertebral spine is removed, and a commercially available cover-glass is overlaid on a thin agarose bed. A 3D-printed plastic backplate is then affixed to the adjacent vertebral spines using tissue adhesive and dental cement. A stabilization platform is used to reduce motion artifact from respiration and heartbeat. This rapid and clamp-free method is well-suited for acute multi-photon fluorescence microscopy. Representative data are included for an application of this technique to two-photon microscopy of the spinal cord vasculature in transgenic mice expressing eGFP:Claudin-5 &#8212; a tight junction protein.

This protocol describes implantation of a glass window onto the spinal cord of a mouse to facilitate visualization by intravital microscopy.

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated ...

Chronic Implantation of Multiple Flexible Polymer Electrode Arrays

Simultaneous recordings from large populations of individual neurons across distributed brain regions over months to years will enable new avenues of scientific and clinical development. The use of flexible polymer electrode arrays can support long-lasting recording, but the same mechanical properties that allow for longevity of recording make multiple insertions and integration into a chronic implant a challenge. Here is a methodology by which multiple polymer electrode arrays can be targeted to a relatively spatially unconstrained set of brain areas.
The method utilizes thin-film polymer devices, selected for their biocompatibility and capability to achieve long-term and stable&#160;electrophysiologic recording interfaces. The resultant implant allows accurate and flexible targeting of anatomically distant regions, physical stability for months, and robustness to electrical noise. The methodology supports up to sixteen serially inserted devices across eight different anatomic targets. As previously demonstrated, the methodology is capable of recording from 1024 channels. Of these, the 512 channels in this demonstration used for single neuron recording yielded 375 single units distributed across six recording sites. Importantly, this method also can record single units for at least 160 days.
This implantation strategy, including temporarily bracing each device with a retractable silicon insertion shuttle, involves tethering of devices at their target depths to a skull-adhered plastic base piece that is custom-designed for each set of recording targets, and stabilization/protection of the devices within a silicone-filled, custom-designed plastic case. Also covered is the preparation of devices for implantation, and design principles that should guide adaptation to different combinations of brain areas or array designs.

Described below is a method for implantation of multiple polymer electrode arrays across anatomically distant brain regions for chronic electrophysiological recording in freely moving rats. Preparation and surgical implantation are described in detail, with emphasis on design principles to guide adaptation of these methods for use in other species.

Simultaneous recordings from large populations of individual neurons across distributed brain regions over months to years will enable new avenues of ...

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset (Callithrix jacchus)

Marmosets (Callithrix jacchus) are small non-human primates that are gaining popularity in biomedical and preclinical research, including the neurosciences. Phylogenetically, these animals are much closer to humans than rodents. They also display complex behaviors, including a wide range of vocalizations and social interactions. Here, an effective stereotaxic neurosurgical procedure for implantation of recording electrode arrays in the common marmoset is described. This protocol also details the pre- and postoperative steps of animal care that are required to successfully perform such a surgery. Finally, this protocol shows an example of local field potential and spike activity recordings in a freely behaving marmoset 1 week after the surgical procedure. Overall, this method provides an opportunity to study the brain function in awake and freely behaving marmosets. The same protocol can be readily used by researchers working with other small primates. In addition, it can be easily modified to allow other studies requiring implants, such as stimulating electrodes, microinjections, implantation of optrodes or guide cannulas, or ablation of discrete tissue regions.

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset Callithrix jacchus

This work presents a protocol to perform a stereotaxic, neurosurgical implantation of&#160;microelectrode arrays in the common marmoset. This method specifically enables electrophysiological recordings in freely behaving animals but can be easily adapted to any other similar neurosurgical intervention in this species (e.g., cannula for drug administration or electrodes for brain stimulation).

Marmosets (Callithrix jacchus) are small non-human primates that are gaining popularity in biomedical and preclinical research, including the ...

Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. Several recent studies have demonstrated the effectiveness of chronic VNS in enhancing central nervous system plasticity to improve motor rehabilitation, extinction learning, and sensory discrimination. Construction of chronically implantable devices for use in such studies is challenging due to rats&#8217; small size, and typical protocols require extensive training of personnel and time-consuming microfabrication methods. Alternatively, commercially available implantable cuff electrodes can be purchased at a significantly higher cost. In this protocol, we present a simple, low-cost method for construction of small, chronically implantable peripheral nerve cuff electrodes for use in rats. We validate the short and long-term reliability of our cuff electrodes by demonstrating that VNS in ketamine/xylazine anesthetized rats produces decreases in breathing rate consistent with activation of the Hering-Breuer reflex, both at the time of implantation and up to 10 weeks after device implantation. We further demonstrate the suitability of the cuff electrodes for use in chronic stimulation studies by pairing VNS with skilled lever press performance to induce motor cortical map plasticity.

Existing approaches for constructing chronically implantable peripheral nerve cuff electrodes for use in small rodents often require specialized equipment and/or highly trained personnel. In this protocol we demonstrate a simple, low-cost approach for fabricating chronically implantable cuff electrodes, and demonstrate their effectiveness for vagus nerve stimulation (VNS) in rats.

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. ...

Electrocorticographic Recording of Cerebral Cortex Areas Manipulated Using an Adeno-Associated Virus Targeting Cofilin in Mice

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. Adeno-associated viruses (AAVs) are increasingly used to improve understanding of brain circuits and their functions. The AAV-mediated manipulation of specific cell populations and/or of precise molecular components has been tremendously useful to identify new sleep regulatory circuits/molecules and key proteins contributing to the adverse effects of sleep loss. For instance, inhibiting activity of the filamentous actin-severing protein&#160;cofilin&#160;using AAV prevents sleep deprivation-induced memory impairment. Here, a protocol is&#160;described that combines the manipulation of cofilin function in a cerebral cortex area with the recording of ECoG activity to examine whether cortical cofilin modulates the wakefulness and sleep ECoG signals. AAV injection is performed during the same surgical procedure as the implantation of ECoG and electromyographic (EMG) electrodes in adult male and female mice. Mice are anesthetized, and their heads are shaved. After skin cleaning and incision, stereotaxic coordinates of the motor cortex are determined, and the skull is pierced at this location. A cannula prefilled with an AAV expressing cofilinS3D, an inactive form of cofilin, is slowly positioned in the cortical tissue. After AAV infusion, gold-covered screws (ECoG electrodes) are screwed through the skull and cemented to the skull with gold wires inserted in the neck muscles (EMG electrodes). The animals are allowed three weeks to recover and to ensure sufficient expression of cofilinS3D. The infected area and cell type are verified using immunohistochemistry, and the ECoG is analyzed using visual identification of vigilance states and spectral analysis. In summary, this combined methodological approach allows the investigation of the precise contribution of molecular components regulating neuronal morphology and connectivity to the regulation of synchronized cerebral cortex activity during wakefulness and sleep.

This article describes a protocol for the manipulation of molecular targets in the cerebral cortex using adeno-associated viruses and for monitoring the effects of this manipulation during wakefulness and sleep using electrocorticographic recordings.

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. ...

Direct Cannula Implantation in the Cisterna Magna of Pigs

The glymphatic system is a waste clearance system in the brain that relies on the flow of cerebrospinal fluid (CSF) in astrocyte-bound perivascular spaces and has been implicated in the clearance of neurotoxic peptides such as amyloid-beta. Impaired glymphatic function exacerbates disease pathology in animal models of neurodegenerative diseases, such as Alzheimer's, which highlights the importance of understanding this clearance system. The glymphatic system is often studied by cisterna magna cannulations (CMc), where tracers are delivered directly into the cerebrospinal fluid (CSF). Most studies, however, have been carried out in rodents. Here, we demonstrate an adaptation of the CMc technique in pigs. Using CMc in pigs, the glymphatic system can be studied at a high optical resolution in gyrencephalic brains and in doing so bridges the knowledge gap between rodent and human glymphatics.

This article presents a step-by-step protocol for the direct cannula implantation in the cisterna magna of pigs.

The glymphatic system is a waste clearance system in the brain that relies on the flow of cerebrospinal fluid (CSF) in astrocyte-bound perivascular spaces ...