The authors would like to thank Bernardo Luiz for technical assistance with filming and editing. This work was supported by Santos Dumont Institute (ISD), Brazilian Ministry of Education (MEC) and Coordena&#231;&#227;o de Aperfei&#231;oamento de Pessoal de N&#237;vel Superior (CAPES).

Animal experiments were performed in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and approved by the Santos Dumont Institute Ethics Committee (protocol 02/2015AAS).
1. Surgery preparation
<ol>
	<li>Attach each electrode array to an electrode holder compatible with the stereotaxic frame to be used.</li>
	<li>Connect one electrode holder to the stereotaxic micromanipulator and set one microwire to the interaural coordinates. Repeat this for the additional electrode arrays and holders, if necessary. 
	NOTE: The interaural coordinate of any microwire can be used to calculate the implantation coordinates for the entire array, because the relative distance between the microwires is constant. When the array has bundles with different lengths, the interaural coordinate of the longest wire is the most convenient to use for setting the interaural zero.</li>
	<li>Detach the electrode holders from the stereotaxic micromanipulator and sterilize the assemblies (electrode attached to the electrode holder)&#160;in an ultraviolet light (UV) cabinet for at least 2 h.</li>
	<li>Attach a 24 G needle to a stereotaxic probe holder, connect it to the micromanipulator and set the interaural coordinates for the tip of the needle. 
	NOTE: Prior to the surgery, the coordinates of all craniotomies must be predefined as a perimeter 200 &#181;m2 larger than the anteroposterior (AP) and mediolateral (ML) position of the array&#8217;s target implantation site. Use the 24 G needle-probe holder assembly to determine the position of the craniotomies in the skull based on the zero interaural coordinates.</li>
	<li>Detach the probe holder from the micromanipulator and sterilize the assembly in a UV cabinet for at least 2 h.</li>
	<li>Gather 6&#8722;8 titanium or stainless-steel screws. Solder a ground wire to half of them.</li>
	<li>Organize and sterilize all remaining instruments, equipment, and disposables required for the surgery.</li>
</ol>
2. Preoperative procedures
NOTE: Two adult male marmosets (Callithrix jacchus) weighing 320&#8211;370 g were used in this study. Ensure that the animal has not eaten for 6 h prior to the induction of anesthesia.
<ol>
	<li>Anesthetize the animal with an intramuscular injection of atropine (0.05 mg/kg) to reduce salivation and bronchial secretions. Check for the lack of pedal responses.</li>
	<li>After 5 min apply ketamine (10&#8722;20 mg/kg) intramuscularly.</li>
	<li>Shave the animal&#8217;s head using an electric barber clipper.</li>
	<li>Administer tramadol (2 mg/kg) intramuscularly as a general analgesic.</li>
	<li>Intubate the animal.
	<ol>
		<li>Using a mask, expose the marmoset to isoflurane in 1&#8722;2% oxygen with a flow rate of 1&#8722;5 L/min to induce deep anesthesia. When the animal is deeply anesthetized, reduce and maintain isoflurane to 1&#8722;3 L/min.</li>
		<li>Attach an elastic band to the surgical table with tape.</li>
		<li>Position the marmoset in a supine position with the head toward the technician and place the elastic band in the marmoset&#8217;s mouth behind its canines. 
		NOTE: It is best to position the head such that the dorsal surface is pointed toward the floor and its face is toward the technician.</li>
		<li>Using a cotton-tipped probe, swab dry the marmoset&#8217;s tongue, and grasp it in one hand to keep the mouth open.</li>
		<li>Spray 10% lidocaine on the tip of the endotracheal tube.</li>
		<li>Insert the uncuffed, 2.0 mm diameter endotracheal tube into the trachea until the 4.0 cm mark is at the entrance of the trachea.</li>
		<li>Attach the tube to the anesthesia assembly with the artificial ventilator&#160;set to 40 breaths/min and confirm proper expansion and contraction of the chest. 
		NOTE: At this time the isoflurane and oxygen should be delivered via the endotracheal tube, not the mask.</li>
		<li>Remove the elastic band from the marmoset&#8217;s mouth so the endotracheal tube can be taped to the jaw.</li>
	</ol>
	</li>
	<li>Position the marmoset in a prone position in line with the stereotaxic frame and fix the animal&#8217;s head into the stereotaxic frame.
	<ol>
		<li>First, insert the tip of the right ear bar into the animal&#8217;s right auditory canal.</li>
		<li>Next, insert the tip of the left ear bar into the left auditory canal.</li>
		<li>Center the animal&#8217;s head at the center of the stereotaxic frame and fix the ear bars in place.</li>
		<li>Insert the mouthpiece into the animal&#8217;s mouth and adjust its height so that it touches the animal&#8217;s palate. At the same time, position the orbital bars at the lower surface of the orbital bone.</li>
		<li>Make sure that the lower surface of the orbital bone is horizontally aligned with the center of the ear bars.</li>
	</ol>
	</li>
	<li>Connect a portable pulse oximeter to the marmoset&#8217;s hand. Ensure that the heart rate is within 154&#8722;180 beats/min (bpm) for the duration of the surgery; often a heart rate above 200 bpm implies the animal is waking up. Ensure that the oxygen saturation is above 95%. It may occasionally drop to 90% without harm. 
	NOTE: Should the heart rate drop below 154 bpm, decrease the isoflurane.</li>
	<li>Position the rectal temperature probe connected to a homeothermic heating pad into the anus, with the desired temperature set for 37 &#176;C. Tape this sensor to the tail to keep it fixed in place.</li>
	<li>Apply sterile ophthalmic lubricant to the eyes.</li>
	<li>Clean and disinfect the animal&#8217;s head with chlorhexidine and povidone iodine before covering the animal with a surgical field. 
	NOTE: Carry out all the following surgical procedures under aseptic conditions.</li>
</ol>
3. Surgery procedures
<ol>
	<li>Apply local analgesic subcutaneously (e.g., lidocaine 20 mg/mL, 0.1 mL) at the site of the intended incision. Make an incision in the midline of the scalp.</li>
	<li>Expose and prepare the surface of the skull.
	<ol>
		<li>Carefully detach the temporal muscle from the cranium. First, use a scalpel to cut the fascia at its insertion into the skull. Then, gently separate the temporal muscle from the cranium using a periosteal raspatory.</li>
		<li>Remove the periosteum from all exposed cranium using a periosteal raspatory.</li>
		<li>Control the bleeding with a sterile cotton swab, if necessary.</li>
		<li>Clean the bone surface with hydrogen peroxide.</li>
	</ol>
	</li>
	<li>Delineate the location of the craniotomy by marking its corners with shallow burr holes into the bone surface. Then, drill out the perimeter of the craniotomy using a dental drill at maximum speed (i.e., 350,000 rpm). Add a few drops of sterile saline over the skull while drilling to prevent overheating. Measure the position of the craniotomy and the coordinates of the electrode implant with respect to the interaural coordinates.</li>
	<li>Implant screws into the skull.
	<ol>
		<li>Drill 6&#8722;8 screw holes into the cranium.</li>
		<li>Implant the screws such that each ground wire fused screw is adjacent to and in the proximity of an unaltered screw (i.e., without a ground wire attached to it).</li>
		<li>Wind each ground wire around the adjacent, unaltered screw.</li>
		<li>Add a drop of silver paint between the ground wire and each screw.</li>
	</ol>
	</li>
	<li>Remove the bone at the center of the craniotomy using forceps with a curved tip (e.g., McPherson forceps). Keep the dura mater hydrated with sterile saline.</li>
	<li>Remove the dura mater. Use a sterile hypodermic needle (25 or 26 G) with the bevel bent at approximately 90&#730; to puncture and lift the surface of the dura mater away from the brain surface. Then, cut the dura mater with microscissors. Keep the exposed brain hydrated with saline. 
	NOTE: If significant dural bleeding is observed, use cautery or sterile absorbent gelatin sponges soaked in thrombine25.</li>
	<li>Implant microelectrode arrays.
	<ol>
		<li>Attach the sterilized electrode holder and electrode array to the stereotaxic micromanipulator.</li>
		<li>Position the micromanipulator such that the electrode is at the desired anteroposterior and mediolateral coordinates.</li>
		<li>Lower the electrode array until the tip of the longest bundle touches the surface of the brain.</li>
		<li>Slowly insert the array into the brain tissue until it reaches the dorsoventral coordinates.</li>
		<li>Cover the exposed cortex with small pieces of sterile, absorbent gelatin sponges.</li>
		<li>Secure the electrode to the skull by applying dental acrylic to the exposed skull, one screw, and the electrode.</li>
		<li>Detach the electrode holder and remove it from the micromanipulator.</li>
	</ol>
	</li>
	<li>Repeat the implantation procedure from step 3.7 with the additional arrays, if necessary.</li>
	<li>Wind together and weld the ground wires of the separate arrays and screws. Use silver paint to form a bridge around the weld to ensure an electrical connection has been achieved.</li>
	<li>Using dental acrylic, make a sturdy headcap around the lateral extent of the arrays, and completely encase the ground wires and any exposed skull and screws.</li>
	<li>If necessary, insert a support bar into the headcap. This can be a sturdy plastic cylinder like those from a cotton swab. Seal it into place with the dental acrylic. 
	NOTE: This can be helpful in securing the electrophysiology cable connectors in place but may be unnecessary depending on the equipment used. In the present method, a similar support rod is affixed to the head stage such that an elastic band can robustly hold the head stages in place on the connectors.</li>
	<li>Suture the skin around the headcap.</li>
</ol>
4. Postoperative recovery
<ol>
	<li>Apply antiseptic solution (e.g., chlorhexidine) around the wound.</li>
	<li>Turn off the isoflurane supply but not the oxygen and remove the animal from the stereotaxic frame.</li>
	<li>Place the animal onto the heating pad with the oxygen maintained through the endotracheal tube.</li>
	<li>Remove the endotracheal tube when the first signs of neurogenic reflexes, such as laryngospasms, are observed.</li>
	<li>Keep supplying the oxygen with a mask until the animal presents clear signs of anesthetic recovery, such as protective reflexes, postural tone, and attempts to ambulate.</li>
	<li>Place the animal inside a clean cage in a recovery room for 24&#8722;48 h before moving the animal to its home cage. House each implanted animal individually. 
	NOTE: Because marmosets tend to climb the cage walls, use a cage with smooth walls or cover the cage walls with a smooth surface to prevent the animal from falling.</li>
	<li>In the first hour following surgery, observe the animal to watch for signs of distress or uncoordinated head contact against the side of the cage.</li>
	<li>Administer antibiotics (e.g., enrofloxacin 5 mg/kg, subcutaneously, once a day for 5&#8722;7 days), analgesics (e.g., oral tramadol 1 mg/kg, every 8 h for 3&#8722;5 days) and anti-inflammatory drugs (e.g., dexamethasone&#160;0.5&#8722;1.5 mg/kg, subcutaneously, once a day for 1&#8722;3 days). 
	NOTE: After a successful surgery, animals will be fully recovered within 3&#8722;5 days.</li>
</ol>
5. Chronic electrophysiological recordings in freely behaving marmosets
<ol>
	<li>Start the electrophysiological recording sessions at least 1 week after the surgery. 
	NOTE: Habituate the animals to the researcher and experimental environments before starting all experimental procedures for at least 1 month.</li>
	<li>At the beginning of each session, lightly anesthetize the animal using isoflurane (1&#8722;5 L/min, 1% O2). 
	NOTE: Follow the relevant institution&#8217;s guidelines regarding the sedation of small primates. If recording sessions are very frequent, habituate the animals to be handled so that cables can be connected without anesthesia.</li>
	<li>Connect the electrode arrays to a commercial neural recording system.</li>
	<li>Place the animal inside the experimental chamber. 
	NOTE: The experimental chamber used here is a cubic acrylic box (0.45 m x 0.45 m x 0.45 m) designed to evaluate the amount and pattern of spontaneous motor activity26,27.</li>
	<li>Wait for 30 min before starting the recordings to ensure the animal is fully recovered from anesthesia. 
	NOTE: Isoflurane has a rapid onset and offset action which allows for rapid sedation and awakening28. Once the isoflurane supply is turned off, the animal will start to wake up. The animal is awake when it stays in the upright position and can ambulate freely in the experimental chamber without falling. This takes less than 15 min. To ensure the absence of any sedative effects, begin the recordings 30 min after the isoflurane is discontinued.</li>
	<li>Confirm the position of the microelectrode array implants postmortem by NISSL staining after fixing and sectioning the tissue29.</li>
</ol>

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> State-of-the-Art Microwire Array Design for Chronic Neural Recordings in Behaving Animals. , (2008).</li><li>Paxinos, G., Watson, C., Petrides, M., Rosa, M., Tokuno, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Marmoset Brain in Stereotaxic Coordinates. , (2012).</li><li>Brown, M. J., Pearson, P. T., Tomson, F. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Guidelines+for+animal+surgery+in+research+and+teaching.">Guidelines for animal surgery in research and teaching.</a> American Journal of Veterinary Research. 54 (9), 1544-1559 (1993).</li><li>Flecknell, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anaesthesia+of+Animals+for+Biomedical+Research.">Anaesthesia of Animals for Biomedical Research.</a> British Journal of Anaesthesia. 71 (6), 885-894 (1993).</li><li>Kurihara, S., 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+Surgical+Procedure+for+the+Administration+of+Drugs+to+the+Inner+Ear+in+a+Non-Human+Primate+Common+Marmoset+(Callithrix+jacchus).">A Surgical Procedure for the Administration of Drugs to the Inner Ear in a Non-Human Primate Common Marmoset (Callithrix jacchus).</a> Journal of Visualized Experiments. (132), (2018).</li><li>Boer, R. A., de Vries, A. M. O., Louwerse, A. L., Sterck, E. H. 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A., Ayata, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+impact+of+anesthetics+and+hyperoxia+on+cortical+spreading+depression.">The impact of anesthetics and hyperoxia on cortical spreading depression.</a> Experimental Neurology. 212 (1), 201-206 (2008).</li><li>Ghomashchi, 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=A+low-cost,+open-source,+wireless+electrophysiology+system.">A low-cost, open-source, wireless electrophysiology system.</a> 2014 36th Annual International Conference of the IEEE Engineering in Medicine and Biology Society. , 3138-3141 (2014).</li><li>Fu, T. M., Hong, G., Viveros, R. D., Zhou, T., Lieber, C. 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The authors have nothing to disclose.

This work provides a detailed description of the procedures involved in the implantation of microelectrode recording arrays in the marmoset brain. This same protocol can be readily used when implanting electrodes, whether homemade or commercially available, in other small primates. Additionally, it can be easily adapted for other experimental ends that require precise targeting of brain structures. Therefore, this protocol is purposefully vague regarding stereotaxic coordinates and cranial drilling techniques, because th...

Common marmosets (Callithrix jacchus) are gaining recognition as an important model organism in many fields of research, including neuroscience. These new-world primates represent an important complementary animal model to both rodents and other non-human primates (NHPs), such as the rhesus macaque. Like rodents, these animals are small, easy to manipulate, and relatively economical to care for and breed1,2,3,4, as compared with larger NHPs. Furthermore, these animals have a propensity for twinning and high fecundity relative to other NHPs1,2,3. Another advantage the marmoset has over many other primates is that modern molecular biology tools3,4,5,6,7 and a sequenced genome2,3,4,5,8 have been used to genetically modify them. Both knock-in animals using lentivirus5, and knock-out animals using zinc-finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENS)7, have yielded viable founder animals.
An advantage in relation to rodents is that marmosets, as primates, are phylogenetically closer to humans3,5,6,9,10,11. Like humans, marmosets are diurnal animals that depend on a highly developed visual system to guide much of their behavior10. Further, marmosets exhibit behavioral complexity, including a wide range of social behaviors such as the use of different vocalizations3, allowing researchers to address questions not possible in other species. From a neuroscientific perspective, marmosets have lissencephaly brains, unlike the more commonly used rhesus macaque9. Furthermore, marmosets have a central nervous system similar to humans, including a more highly developed prefrontal cortex9. Together, all these characteristics position marmosets as a valuable model to study brain function in health and disease.
A common method for studying brain function involves implanting electrodes in anatomically specific locations by means of stereotaxic neurosurgery. This allows for chronical recording of the neural activity in different target areas in awake and freely behaving animals12,13. Stereotaxic neurosurgery is an indispensable technique used in many lines of research, as it allows precise targeting of neuroanatomical regions. Compared to macaque and rodent literature, there are fewer published studies describing the stereotaxic neurosurgery specific to the marmoset, and they tend to provide sparse detail of the steps involved in the surgery. Moreover, those with greater detail mainly focus on procedures for electrophysiology recording in head-restrained animals14,15,16,17.
In order to facilitate wider adoption of marmosets as a model organism in neuroscience research, the present method defines specific steps necessary for a successful stereotaxic neurosurgery in this species. In addition to implantation of recording arrays, as detailed in the present method, the same technique can be adapted for many other experimental ends, including implantation of stimulating electrodes for the treatment of diseases18 or causally driving circuit behavior19; implantation of guide cannulas for extraction and quantification of neurotransmitters20, injections of reagents, including those for inducing disease models12 or for circuit tracing studies15; ablation of discrete tissue regions21; implantation of optrodes for optogenetic studies22; implantation of optical windows for cortical microscopic analysis23; and implantation of electrocorticographic (ECoG) arrays24. Thus, the overall goal of this procedure is to outline the surgical steps involved in the implantation of microelectrode arrays for chronic electrophysiological recordings in freely behaving marmosets.

<table>
	<tbody>
		<tr>
			<td>Equipments</td>
			<td></td>
			<td></td>
			<td></td>
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		<tr>
			<td>683 Small Animal Ventilator</td>
			<td>Harvard Apparatus, Inc.</td>
			<td>55-0000</td>
			<td></td>
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			<td>Anesthesia Assembly</td>
			<td>BRASMED</td>
			<td>COLIBRI</td>
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			<td>Barber Clippers</td>
			<td>Mundial</td>
			<td>HC-SERIES</td>
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			<td>Dental Drill</td>
			<td>Norgen</td>
			<td>B07-201-M1KG</td>
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		<tr>
			<td>Homeothermic Heating Pad and Monitor</td>
			<td>Harvard Apparatus, Inc.</td>
			<td>50-7212</td>
			<td></td>
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		<tr>
			<td>Marmoset Stereotaxic Frame</td>
			<td>Narishige Scientific Instrument Lab</td>
			<td>SR-6C-HT</td>
			<td></td>
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		<tr>
			<td>Patient Monitor and Pulse Oximeter</td>
			<td>Bionet Co., Ltd</td>
			<td>BM3</td>
			<td></td>
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		<tr>
			<td>Stereotaxic Micromanipulator</td>
			<td>Narishige Scientific Instrument Lab</td>
			<td>SM-15R</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Microscope</td>
			<td>Opto</td>
			<td>SM PLUS IBZ</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
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		<tr>
			<td>Instruments</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Allis tissue forceps</td>
			<td>Sklar</td>
			<td>36-2275</td>
			<td></td>
		</tr>
		<tr>
			<td>Alm Retractor, rounded point, 4x4 teeth</td>
			<td>Rhosse</td>
			<td>RH11078</td>
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			<td>Angled McPherson Forceps</td>
			<td>Oftalmologiabr</td>
			<td>11301A</td>
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			<td>Curved Surgial Scissors</td>
			<td>Harvard Apparatus, Inc.</td>
			<td>72-8422</td>
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		<tr>
			<td>Curved Tissue Forceps</td>
			<td>Sklar</td>
			<td>47-1186</td>
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		<tr>
			<td>Delicate Dissection forceps</td>
			<td>WPI</td>
			<td>WP5015</td>
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			<td>Dental Drill Bit</td>
			<td>Microdont</td>
			<td>ISO.806.314.001.524.010</td>
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			<td>Essring Tissue Forceps</td>
			<td>Sklar</td>
			<td>19-2460</td>
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		<tr>
			<td>FG 1/4 Dental Drill Bit</td>
			<td>Microdont</td>
			<td>ISO.700.314.001.006.005</td>
			<td></td>
		</tr>
		<tr>
			<td>Halsey Needle Holder</td>
			<td>WPI</td>
			<td>15926-G</td>
			<td></td>
		</tr>
		<tr>
			<td>Halstead Mosquito forceps</td>
			<td>WPI</td>
			<td>503724-12</td>
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		<tr>
			<td>Hemostatic Forceps, Straight</td>
			<td>Sklar</td>
			<td>17-1260</td>
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			<td>Jewler Forceps</td>
			<td>Sklar</td>
			<td>66-7436</td>
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			<td>McPherson-Vannas Optathalmic microscissor, 3 mm point</td>
			<td>Argos Instrumental</td>
			<td>ARGOS-4004</td>
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			<td>Pereosteal Raspatory</td>
			<td>Golgran</td>
			<td>38-1</td>
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			<td>Scalpal Handle</td>
			<td>Harvard Apparatus, Inc.</td>
			<td>72-8354</td>
			<td></td>
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		<tr>
			<td>Screwdrivers</td>
			<td>Eurotool</td>
			<td>SCR-830.00</td>
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			<td>Sodering Iron</td>
			<td>Hikari</td>
			<td>21K006</td>
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			<td>Surgical Scissor</td>
			<td>Harvard Apparatus, Inc.</td>
			<td>72-8400</td>
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			<td>Toothed forceps</td>
			<td>WPI</td>
			<td>501266-G</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Disposables/Single Use</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1 ml sterile syringe with 26 G needle</td>
			<td>Descarpack</td>
			<td>7898283812785</td>
			<td></td>
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		<tr>
			<td>130 cm x 140 cm surgical field, presterilized</td>
			<td>ProtDesc</td>
			<td>7898467276344</td>
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			<td>24G Needle, presterilized</td>
			<td>Descarpack</td>
			<td>7898283812846</td>
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		<tr>
			<td>50 cm x 50 cm surgical field, presterilized</td>
			<td>Esterili-med</td>
			<td>110100236</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton Tipped Probes, Presterilized</td>
			<td>Jiangsu Suyun Medical Materials Co. LTD</td>
			<td>23007</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton tipped Qutips</td>
			<td>Higie Topp</td>
			<td>7898095296063</td>
			<td></td>
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		<tr>
			<td>Electrode Array</td>
			<td></td>
			<td></td>
			<td>Home made</td>
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		<tr>
			<td>Endotracheal tube without cuff, internal diameter 2.0 mm, outer diameter 2.9 mm</td>
			<td>Solidor</td>
			<td>7898913077201</td>
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			<td>Tinned copper wire, 0.15 mm diameter</td>
			<td></td>
			<td></td>
			<td></td>
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		<tr>
			<td>M1.4x3 Stainless steel screws</td>
			<td>USMICROSCREW</td>
			<td>M14-30M-SS-P</td>
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			<td>Medical Tape</td>
			<td>Missner</td>
			<td>7896544910102</td>
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			<td>Nylon surgical sutures</td>
			<td>Shalon</td>
			<td>N540CTI25</td>
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			<td>Scalpal Blade, presterilized</td>
			<td>AdvantiVe</td>
			<td>1037</td>
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			<td>solder</td>
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			<td>Sterile Saline 0.9%</td>
			<td>Isofarma</td>
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			<td>Sterile Surgical Gloves</td>
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			<td>Sterile Surgical Gown</td>
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			<td>Surgical Gauze, 15 cm x 26 cm presterilized</td>
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			<td>Gelfoam</td>
			<td>Pfizer</td>
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			<td></td>
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			<td>0.25mg/ml Atropine</td>
			<td>Isofarma</td>
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			<td>10% Lidocaine Spray</td>
			<td>Produtos Qu&#237;micos Farmac&#234;uticos Ltda.</td>
			<td>7896676405644</td>
			<td></td>
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			<td>2.5% Enrofloxacino veterinary antibiotic</td>
			<td>Chemitec</td>
			<td>0137-02</td>
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		<tr>
			<td>Dexametasona Veterinary Anti inflammatory</td>
			<td>MSD</td>
			<td>R06177091A-00-15</td>
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			<td>Hydrogen Peroxide</td>
			<td>Farmax</td>
			<td>7896902211537</td>
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			<td>Isoflourane</td>
			<td>BioChimico</td>
			<td>7897406113068</td>
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		<tr>
			<td>Jet Acrylic polymerization solution</td>
			<td>Artigos Odontol&#243;gicos Cl&#225;ssico</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Jet Auto Polymerizing Acrylic</td>
			<td>Artigos Odontol&#243;gicos Cl&#225;ssico</td>
			<td></td>
			<td></td>
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		<tr>
			<td>Ketamine 10%</td>
			<td>Syntec</td>
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			<td>Lidocaine and Phenylephrine 1.8 ml local anesthetic</td>
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			<td>Povidone-Iodine solutiom</td>
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			<td>Riohex 2% surgical Soap</td>
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			<td>Silver Paint</td>
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			<td>05002-AB</td>
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		<tr>
			<td>Tramadol chloride 50 mg/ml</td>
			<td>Uni&#227;o Qu&#237;mica</td>
			<td>7896006245452</td>
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		<tr>
			<td>Refresh gel (polyacrylic acid)</td>
			<td>Allergan</td>
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</table>

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.

The purpose of this study was to describe a stereotaxic neurosurgical procedure for implantation of microelectrode arrays for electrophysiological recordings in the common marmoset. A typical surgery (from anesthesia induction to anesthesia recovery) will last for approximately 5&#8722;7 h, depending on the number of arrays implanted. Here, two arrays were symmetrically implanted, one in each brain hemisphere. Each array contained 32 stainless steel microwires arranged in seven bundles targeting several structures of the...

Watch this Scientific Journal Video about Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset Callithrix jacchus at JoVE.com

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).

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 ...

stereotaxic-surgery-for-implantation-microelectrode-arrays-common

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Neuroscience

9.2K Views.  Santos Dumont Institute. This work presents a protocol to perform a stereotaxic, neurosurgical implantation of 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).

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

Rodent Stereotaxic Surgery and Animal Welfare Outcome Improvements for Behavioral Neuroscience

Stereotaxic surgery for the implantation of cannulae into specific brain regions has for many decades been a very successful experimental technique to investigate the effects of locally manipulated neurotransmitter and signaling pathways in awake, behaving animals. Moreover, the stereotaxic implantation of electrodes for electrophysiological stimulation and recording studies has been instrumental to our current understanding of neuroplasticity and brain networks in behaving animals. Ever-increasing knowledge about optimizing surgical techniques in rodents1-4, public awareness concerning animal welfare issues and stringent legislation (e.g., the 2010 European Union Directive on the use of laboratory animals5) prompted us to refine these surgical procedures, particularly with respect to implementing new procedures for oxygen supplementation and the continuous monitoring of blood oxygenation and heart rate levels during the surgery as well as introducing a standardized protocol for post-surgical care. Our observations indicate that these modifications resulted in an increased survival rate and an improvement in the general condition of the animals after surgery (e.g. less weight loss and a more active animal). This video presentation will show the general procedures involved in this type of stereotaxic surgery with special attention to our several modifications. We will illustrate these surgical procedures in rats, but it is also possible to perform this type of surgery in mice or other small laboratory animals by using special adaptors for the stereotaxic apparatus6.

Stereotaxic surgery on rodents allows for targeted administration of drugs or electrical stimulation and recordings in awake, behaving animals. In this video presentation we will demonstrate recent procedural refinements to this long-standing procedure that successfully improved survival rate and reduced post-surgical weight loss.

Stereotaxic surgery for the implantation of cannulae into specific brain regions has for many decades been a very successful experimental technique to ...

Stereotaxic Surgery for Excitotoxic Lesion of Specific Brain Areas in the Adult Rat

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the function of various brain regions for behavior or other experimental outcomes is to implement a localized ablation of function. In humans, patient populations with localized brain lesions are often studied for deficits, in hopes of revealing the underlying function of the damaged area. In rodents, one can experimentally induce lesions of specific brain regions.
Lesion can be accomplished in several ways. Electrolytic lesions can cause localized damage but will damage a variety of cell types as well as traversing fibers from other brain regions that happen to be near the lesion site. Inducible genetic techniques using cell-type specific promoters may also enable site-specific targeting. These techniques are complex and not always practical depending on the target brain area. Excitotoxic lesion using stereotaxic surgery, by contrast, is one of the most reliable and practical methods of lesioning excitatory neurons without damaging local glial cells or traversing fibers.
Here, we present a protocol for stereotaxic infusion of the excitotoxin, N-methyl-D-aspartate (NMDA), into the basolateral amygdala complex. Using anatomical indications, we apply stereotaxic coordinates to determine the location of our target brain region and lower an injection needle in place just above the target. We then infuse our excitotoxin into the brain, resulting in excitotoxic death of nearby neurons. While our experimental subject of choice is a rat, the same methods can be applied to other mammals, with the appropriate adjustments in equipment and coordinates.
This method can be used on a variety of brain regions, including the basolateral amygdala1-6, other amygdala nuclei6, 7, hippocampus8, entorhinal cortex9 and prefrontal cortex10. It can also be used to infuse biological compounds such as viral vectors1, 11. The basic stereotaxic technique could also be adapted for implantation of more permanent osmotic pumps, allowing more prolonged exposure to a compound of interest.

Targeted ablation of specific brain region(s) by infusion of an excitotoxin using stereotaxic coordinates is described. This technique could also be adapted for infusion of other chemicals into the rat brain.

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the ...

Microelectrode Guided Implantation of Electrodes into the Subthalamic Nucleus of Rats for Long-term Deep Brain Stimulation

Deep brain stimulation (DBS) is a widely used and effective therapy for several neurologic disorders, such as idiopathic Parkinson&#8217;s disease, dystonia or tremor. DBS is based on the delivery of electrical stimuli to specific deep anatomic structures of the central nervous system. However, the mechanisms underlying the effect of DBS remain enigmatic. This has led to an interest in investigating the impact of DBS in animal models, especially in rats. As DBS is a long-term therapy, research should be focused on molecular-genetic changes of neural circuits that occur several weeks after DBS. Long-term DBS in rats is challenging because the rats move around in their cage, which causes problems in keeping in place the wire leading from the head of the animal to the stimulator. Furthermore, target structures for stimulation in the rat brain are small and therefore electrodes cannot easily be placed at the required position. Thus, a set-up for long-lasting stimulation of rats using platinum/iridium electrodes with an impedance of about 1 M&#937; was developed for this study. An electrode with these specifications allows for not only adequate stimulation but also recording of deep brain structures to identify the target area for DBS. In our set-up, an electrode with a plug for the wire was embedded in dental cement with four anchoring screws secured onto the skull. The wire from the plug to the stimulator was protected by a stainless-steel spring. A swivel was connected to the circuit to prevent the wire from becoming tangled. Overall, this stimulation set-up offers a high degree of free mobility for the rat and enables the head plug, as well as the wire connection between the plug and the stimulator, to retain long-lasting strength.

A method for implanting electrodes into the subthalamic nucleus (STN) of rats is described. Better localization of the STN was achieved by using a microrecording system. Furthermore, a stimulation set-up is presented that is characterized by long-lasting connections between the head of the animal and the stimulator.

Deep brain stimulation (DBS) is a widely used and effective therapy for several neurologic disorders, such as idiopathic Parkinson&#8217;s disease, ...

Stereotaxic Surgery for Genetic Manipulation in Striatal Cells of Neonatal Mouse Brains

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, they may function to regulate the developmental process and/or physiological function in neonatal brains. To investigate neurobiological functions of specific genes in the brain, it is essential to inactivate genes in the brain. Here, we describe a simple stereotaxic method to inactivate gene expression in the striatum of transgenic mice at neonatal time windows. AAV-eGFP-Cre viruses were microinjected into the striatum of Ai14 reporter gene mice at postnatal day (P) 2 by stereotaxic brain surgery. The tdTomato reporter gene expression was detected in P14 striatum, suggesting a successful Cre-loxP mediated DNA recombination in AAV-transduced striatal cells. We further validated this technique by microinjecting AAV-eGFP-Cre viruses into P2Foxp2fl/fl mice. Double labeling of GFP and Foxp2 showed that GFP-positive cells lacked Foxp2 immunoreactivity in P9 striatum, suggesting the loss of Foxp2 protein in AAV-eGFP-Cre transduced striatal cells. Taken together, these results demonstrate an effective genetic deletion by stereotaxically microinjected AAV-eGFP-Cre viruses in specific neuronal populations in the neonatal brains of floxed transgenic mice. In conclusion, our stereotaxic technique provides an easy and simple platform for genetic manipulation in neonatal mouse brains. The technique can not only be used to delete genes in specific regions of neonatal brains, but it also can be used to inject pharmacological drugs, neuronal tracers, genetically modified optogenetics and chemogenetics proteins, neuronal activity indicators and other reagents into the striatum of neonatal mouse brains.

We describe a protocol of stereotaxic surgery with a homemade head-fixed device for microinjecting reagents into the striatum of neonatal mouse brains. This technique allows genetic manipulation in neuronal cells of specific regions of neonatal mouse brains.

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, ...

Recording and Modulation of Epileptiform Activity in Rodent Brain Slices Coupled to Microelectrode Arrays

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) is a promising approach when pharmacological treatment fails or neurosurgery is not recommended. Acute brain slices coupled to microelectrode arrays (MEAs) represent a valuable tool to study neuronal network interactions and their modulation by electrical stimulation. As compared to conventional extracellular recording techniques, they provide the added advantages of a greater number of observation points and a known inter-electrode distance, which allow studying the propagation path and speed of electrophysiological signals. However, tissue oxygenation may be greatly impaired during MEA recording, requiring a high perfusion rate, which comes at the cost of decreased signal-to-noise ratio and higher oscillations in the experimental temperature. Electrical stimulation further stresses the brain tissue, making it difficult to pursue prolonged recording/stimulation epochs. Moreover, electrical modulation of brain slice activity needs to target specific structures/pathways within the brain slice, requiring that electrode mapping be easily and quickly performed live during the experiment. Here, we illustrate how to perform the recording and electrical modulation of 4-aminopyridine (4AP)-induced epileptiform activity in rodent brain slices using planar MEAs. We show that the brain tissue obtained from mice outperforms rat brain tissue and is thus better suited for MEA experiments. This protocol guarantees the generation and maintenance of a stable epileptiform pattern that faithfully reproduces the electrophysiological features observed with conventional field potential recording, persists for several hours, and outlasts sustained electrical stimulation for prolonged epochs. Tissue viability throughout the experiment is achieved thanks to the use of a small-volume custom recording chamber allowing for laminar flow and quick solution exchange even at low (1 mL/min) perfusion rates. Quick MEA mapping for real-time monitoring and selection of stimulating electrodes is performed by a custom graphic user interface (GUI).

We illustrate how to perform recording and electrical modulation of 4-aminopyridine-induced epileptiform activity in rodent brain slices using microelectrode arrays. A custom recording chamber maintains tissue viability throughout prolonged experimental sessions. Live electrode mapping and selection of stimulating pairs are performed by a custom graphical user interface.

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) ...

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.

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 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 ...

Construction and Implementation of Carbon Fiber Microelectrode Arrays for Chronic and Acute In Vivo Recordings

Multichannel electrode arrays offer insight into the working brain and serve to elucidate neural processes at the single-cell and circuit levels. Development of these tools is crucial for understanding complex behaviors and cognition and for advancing clinical applications. However, it remains a challenge to densely record from cell populations stably and continuously over long time periods. Many popular electrodes, such as tetrodes and silicon arrays, feature large cross-diameters that produce damage upon insertion and elicit chronic reactive tissue responses associated with neuronal death, hindering the recording of stable, continuous neural activity. In addition, most wire bundles exhibit broad spacing between channels, precluding simultaneous recording from a large number of cells clustered in a small area. The carbon fiber microelectrode arrays described in this protocol offer an accessible solution to these concerns. The study provides a detailed method for fabricating carbon fiber microelectrode arrays that can be used for both acute and chronic recordings in vivo. The physical properties of these electrodes make them ideal for stable and continuous long-term recordings at high cell densities, enabling the researcher to make robust, unambiguous recordings from single units across months.

Construction and Implementation of Carbon Fiber Microelectrode Arrays for Chronic and Acute In Vivo Recordings

This protocol describes a procedure for constructing carbon fiber microelectrode arrays for chronic and acute in vivo electrophysiological recordings in mouse (Mus musculus) and ferret (Mustela putorius furo) from multiple brain regions. Each step, following the purchase of raw carbon fibers to microelectrode array implantation, is described in detail, with emphasis on microelectrode array construction.

Multichannel electrode arrays offer insight into the working brain and serve to elucidate neural processes at the single-cell and circuit levels. ...

Recording Network Activity in Spinal Nociceptive Circuits Using Microelectrode Arrays

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an increasingly detailed view of the circuits underpinning spinal pain processing. However, the effects of these connections for broader network activity in the DH remain less well understood because most studies focus on the activity of single neurons and small microcircuits. Alternatively, the use of microelectrode arrays (MEAs), which can monitor electrical activity across many cells, provides high spatial and temporal resolution of neural activity. Here, the use of MEAs with mouse spinal cord slices to study DH activity induced by chemically stimulating DH circuits with 4-aminopyridine (4-AP) is described. The resulting rhythmic activity is restricted to the superficial DH, stable over time, blocked by tetrodotoxin, and can be investigated in different slice orientations. Together, this preparation provides a platform to investigate DH circuit activity in tissue from na&#239;ve animals, animal models of chronic pain, and mice with genetically altered nociceptive function. Furthermore, MEA recordings in 4-AP-stimulated spinal cord slices can be used as a rapid screening tool to assess the capacity of novel antinociceptive compounds to disrupt activity in the spinal cord DH.

The combined use of microelectrode array technology and 4-aminopyridine-induced chemical stimulation for investigating network-level nociceptive activity in the spinal cord dorsal horn is outlined.

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an ...

Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging

A miniature fluorescence microscope (miniscope) is a potent tool for in vivo calcium imaging from freely behaving animals. It offers several advantages over conventional multi-photon calcium imaging systems: (1) compact; (2) light-weighted; (3) affordable; and (4) allows recording from freely behaving animals. This protocol describes brain surgeries for deep brain in vivo calcium imaging using a custom-developed miniscope recording system. The preparation procedure consists of three steps, including (1) stereotaxically injecting the virus at the desired brain region of a mouse brain to label a specific subgroup of neurons with genetically encoded calcium sensor; (2) implantation of gradient-index (GRIN) lens that can relay calcium image from deep brain region to the miniscope system; and (3) affixing the miniscope holder over the mouse skull where miniscope can be attached later. To perform in vivo calcium imaging, the miniscope is fastened onto the holder, and neuronal calcium images are collected along with simultaneous behavior recordings. The present surgery protocol is compatible with any commercial or custom-built single-photon and two-photon imaging systems for deep brain in vivo calcium imaging.

Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging

Miniscope in vivo calcium imaging is a powerful technique to study neuronal dynamics and microcircuits in freely behaving mice. This protocol describes performing brain surgeries to achieve good in vivo calcium imaging using a miniscope.

A miniature fluorescence microscope (miniscope) is a potent tool for in vivo calcium imaging from freely behaving animals. It offers several advantages ...