This work was funded by the Center for Neural Communication Technology (CNCT), a P41 Resource Center funded by the National Institute of Biomedical Imaging and Bioengineering (NIBIB, P41 EB002030) and supported by the National Institutes of Health (NIH). The authors wish to thank Rio Vetter of NeuroNexus Technologies for the development of surgical techniques, and Paras Patel of the University of Michigan for compiling the list of equipment used during surgeries.

1. Pre-implant Procedures (Common)
<ol>
 <li>Shave the top of the animal&#x2019;s head from between the eyes to well behind the ears. Use electric barber&#x2019;s clippers to shave the head. Scrub head first with isopropyl alcohol, then betadine (repeat 3x). Apply eye lubricant. </li>
 <li>Mount the animal in stereotaxic ear bars. Direct the fixed tip of the right stereotaxic ear bar into the auditory canal of the animal&#x2019;s right ear by moving the head sideways against the canal. Extreme care must be taken to assure that each ear-bar tip is properly positioned in the auditory meatus. Next, push the loosened left ear bar into the auditory canal and apply slight inward pressure. Grasp the animal&#x2019;s nose and wiggle it back and forth firmly. If the animal is mounted properly the head will feel rigidly mounted. </li>
</ol>
2. Pre-implantation Surgery (Common)
<ol>
 <li>With a pair of fine scissors, cut two half circles from the midline outwards, partially removing the scalp </li>
 <li> Scrape and remove the periosteum connective tissue that adheres to the bone. Scraping with a blunt edge such as the back of a scalpel blade or the tip of a sterile cotton swab will minimize bleeding because the capillaries will be crushed rather than cut smoothly. Control bone bleeding using bone wax or a cauterizing iron. </li>
 <li>Disinfect and clean the bone surface using hydrogen peroxide. Dry the bone to make the cranial sutures more clearly visible. The Bregma and Lambda points should be noted and used to level the head. The location for craniotomy and electrode implant should be measured with respect to Bregma and can be temporarily marked with a pen or by drilling shallowly into the surface. The craniotomy location will depend on the experimental aims of the study (i.e. motor, sensory experiments) 1. </li>
 <li>Drill screwholes into the bone and place the ground and stabilizing screws. Always use stainless steel or titanium screws. Small jeweler&#x2019;s screws (for example #2-56, or #00-80) with as many threads per inch as possible are desirable. </li>
</ol>
3. Silicon Electrode Implant Procedures
<ol>
 <li>Drill the craniotomy after implanting screws. Drill four shallow pilot marks for the corners of the craniotomy. Using pilot marks as a guide drill out the perimeter of the craniotomy. Remove the center mass of bone that was not drilled using a pair of sturdy forceps. The edge of the open craniotomy may need to be cleaned with further drilling. Thin pieces of bone may be clipped with a pair of microrongeurs. Plan your surgery carefully to avoid major blood vessels such as the Sagittal Sinus, which may appear within 0.5 mm of the midline longitudinal suture 1. Puncturing the Sagittal sinus can cause extensive bleeding. Exposed dura should be kept hydrated with aCSF or Saline.</li>
 <li> To pierce the dura mater you will need to fashion a small hook using a sterile hypodermic needle of small gauge (e.g., &lt; 28 gauge). Take the bevel of the needle and press it flat against a firm surface (such as the flat part of a scalpel) tilting slightly to form a 90 degree bend at the point. Use this hook to catch the surface of the dura, and lift upwards from the brain surface. The dura can be cut using micro scissors or may be torn using lateral movements of the dura hook. Take care to avoid blood vessels on the surface of the brain when using the dura hook. You should notice CSF leaking out when dura is resected and a difference in color from the surface of the dura. </li>
 <li>Carefully attach the ground wire of the silicon electrode around the base of the ground screw, and tact in place with dental cement. The silicon electrodes should be placed carefully so that the electrodes cable leads are arched over the craniotomy. </li>
 <li>Implants will be placed using a linear motor actuator placed on a stereotaxic apparatus. To attach the electrode to the insertion device, Polyethylene glycol (PEG) is heated with a soldering iron and applied to insertion rod and the recording electrode. </li>
 <li>With the electrode secured, manually lower the tip of the electrode to the desired stereotaxic location on the surface of the brain. </li>
 <li>Using the software interface to the linear actuator, move the electrode into the desired depth within the brain tissue. </li>
 <li>Secure the recording electrode in the brain by attaching PEG from an adjacent screw to the electrode cable. You can now remove the electrode from the insertion device by adding saline to dissolve the PEG bound. Raise the insertion device away from the craniotomy and repeat for each electrode. </li>
 <li>Cover the exposed craniotomy with saline soaked Gelfoam. A silicone polymer is then used to cover the electrode cable and protect it from the dental acrylic. Position the connector in the final location, then apply dental acrylic to make a robust headcap. </li>
</ol>
4. MicroECoG Electrode Implant Procedures
<ol>
 <li>Implantation of the microECoG electrode will involve a slightly larger craniotomy. For a 5 x 5 mm device, a 6 x 6 mm craniotomy must be made. Before drilling the craniotomy, UV-curable dental acrylic is applied to the periphery of the craniotomy site while it is still dry and not in danger of touching the dura or pia. </li>
 <li>A sterile surgical drill is used with a #107 burr to drill off the surface of bone in the general shape of the craniotomy. The smaller #106 burr can then be used to drill out the rest of the bone down to a thin transparent layer. Rongeurs will then be able to lift off the remaining piece of bone. The microECoG can be implanted epidurally or subdurally. Again, keep the dura well hydrated with artificial CSF or Saline. </li>
 <li>In order to implant the microECoG, place a stereotaxic arm over the open skull, and secure the electrode to the arm using sterile tape. The electrode can now be lowered down into the craniotomy. Make sure the electrode sites are facing downward and will be making contact with the dura or pial surface. If kept moist, the electrode should slide onto the tissue and will stay in place. </li>
 <li>The ground wire is connected to the ground screw by wrapping around at least three times over and under itself. The ground screw can be any screw that has come into contact with the dura. The reference wire can also be tied to the reference screw in the same manner. </li>
 <li>Small pieces of saline soaked Gelfoam should be placed surrounding the electrode where there is dura or pia exposed. A small amount of saline soaked Gelfoam should also cover the top of the thin film electrode. UV-curable dental acrylic can be applied to the top of the Gelfoam and can be used to create a stable head cap. The acrylic is applied directly to the thin film cable covering it until the connector is reached. Be careful not to cover beyond the bottom of the connector cap. </li>
 <li>Alternatively, the implantation of the microECoG in 4.3 can be immediately followed by the implantation of a silicon microelectrode array (steps 3.2-3.6) through small holes that are manufactured in the microECoG substrate. This allows for simultaneous recording of both single units at various depths along with high resolution field recordings from the surrounding brain surface. </li>
</ol>
5. Postoperative Recovery
<ol>
 <li>After the cement has completely hardened, suture the skin tightly around the head-cap and remove the animal from the stereotaxic frame. </li>
 <li>Apply antiseptic powder or antibiotic ointment copiously around the wound. If there is any bleeding from the ears, place some antibiotic well into the ear canal. </li>
 <li>Keep the animal warm as barbiturates prevent the animal from maintaining its body heat. House each chronically implanted animal in a separate cage. It usually takes four to seven days for the animal to recover completely from the surgery. </li>
</ol>
6. Representative Results
 A successful silicon electrode implant surgery will have recording sites with impedances measuring between 500kOhm-2MOhm, and thin-film electrodes between 10kOhm-50kOhm (at 1kHz). The neural signal can be checked immediately post surgery as well. You should be able to see spikes on the implanted electrodes and see slow wave oscillations on the thin film surface electrode (Figure 1). 
<img src="/files/ftp_upload/3565/3565fig1.jpg" alt="Figure 1" /> 
Figure 1. Representative results of single unit activity and ECoG oscillations. a) Wide-band data from 6 simultaneously recorded channels of a chronic silicon electrode implanted in the neocortex. Note that spikes can often be seen across recording sites. b) ECoG oscillations from 6 channels of a 16-channel thin-film uECoG electrode array on the surface of the brain. The mean of all 16 signals were removed from each trace (common average referencing), and a 500 Hz digital low pass filter was applied.

<ol>
	<li>Paxinos, G. A. <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 Rat Brain in Stereotaxic Coordinates. , (2007).</li><li>Fujisawa, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Behavior-dependent+short-term+assembly+dynamics+in+the+medial+prefrontal+cortex.">Behavior-dependent short-term assembly dynamics in the medial prefrontal cortex.</a> Nat. Neurosci. 11 (7), 823-833 (2008).</li><li>Merrill, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrical+stimulation+of+excitable+tissue:+design+of+efficacious+and+safe+protocols.">Electrical stimulation of excitable tissue: design of efficacious and safe protocols.</a> J. Neurosci. Methods. 141, 171-171 (2005).</li><li>Gage, G. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Naive+coadaptive+cortical+control.">Naive coadaptive cortical control.</a> J. Neural Eng. 2, 52-52 (2005).</li><li>Marzullo, T. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+Closed-Loop+Neural+Interface+Technology+in+a+Rat+Model:+Combining+Motor+Cortex+Operant+Conditioning+With+Visual+Cortex+Microstimulation.">Development of Closed-Loop Neural Interface Technology in a Rat Model: Combining Motor Cortex Operant Conditioning With Visual Cortex Microstimulation.</a> IEEE Trans. Neur. Sys. and Rehab. Eng. 18 (2), 117-126 (2010).</li><li>Maynard, E. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Utah+intracortical+electrode+array:+a+recording+structure+for+potential+brain-computer+interfaces.">The Utah intracortical electrode array: a recording structure for potential brain-computer interfaces.</a> Electroencephalogr. Clin. Neurophysiol. 102, 228-239 (1997).</li><li>Kipke, D. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Silicon-substrate+intracortical+microelectrode+arrays+for+long-term+recording+of+neuronal+spike+activity+in+cerebral+cortex.">Silicon-substrate intracortical microelectrode arrays for long-term recording of neuronal spike activity in cerebral cortex.</a> IEEE Trans. Neural Syst. Rehab. Eng. 11 (2), 151-155 (2003).</li><li>Vetter, 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=Chronic+neural+recording+using+silicon-substrate+microelectrode+arrays+implanted+in+cerebral+cortex.">Chronic neural recording using silicon-substrate microelectrode arrays implanted in cerebral cortex.</a> IEEE Trans. Biomed. Eng. 51 (6), 896-904 (2004).</li></ol>

DRK is part-owner of NeuroNexus Technologies.

There is an increasing interest in using intracortical and surface multichannel recording interfaces for researching brain function 2, providing microstimulation 3, or control signals for neuroprosthetic systems 4,5. The methods outlined in this video demonstrate how to implant chronic penetrating and surface electrode systems. While other silicon chronic electrode systems exist 6, we have focused on implanting planar electrodes developed by NeuroNexus Technologies 7,8. Techniques for the implantation of other electrode systems will vary considerably.
Of the surgical steps described, mounting the animal in the stereotaxic ear bars may be the most difficult and the most important step in determining the accuracy of placement of the implanted electrodes. Extreme care must be taken, therefore, to assume that each ear-bar tip is properly positioned in the auditory meatus. After assurance that the ear-bar tip is in the auditory meatus of the animal&#x2019;s right ear, release the animal&#x2019;s neck with your right hand while your left thumb and forefinger continue to support the head and apply pressure of the head against the ear-bar tip. With your right hand, push the loosened left ear bar into the auditory canal and apply slight inward pressure. Move the left side of the animal&#x2019;s head down and forward and all around until you can feel the left ear-bar tip slip into the auditory meatus. With your right hand, continue to apply pressure on the left ear bar. Release the animal&#x2019;s head and, with your now free left hand, tighten the set screw of the left ear bar.

<table border="1">
 <tbody>
 <tr>
 <td>Equipment</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>Bone Screw - #2-56 x 3/16, 				phillips pan head, 18-8 S/S</td>
 <td>BoltDepot</td>
 <td>7650</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Bone Screw - #2-56 x 1/8, phillips 				pan head, 18-8 S/S</td>
 <td>BoltDepot</td>
 <td>9617</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Large Oxygen Tank - Size K</td>
 <td>Cryogenic Gases</td>
 <td>OXKME</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Small Oxygen Tank - Size E</td>
 <td>Cryogenic Gases</td>
 <td>OXYE-AL</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Vannas-Tubingen Spring Scissors - 				2.5mm Blade, Angled</td>
 <td>Fine Science Tools</td>
 <td>15002-08</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Dumont #5 Forceps, Teflon</td>
 <td>Fine Science Tools</td>
 <td>11626-11</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Dumont #5 Forceps, Dumostar</td>
 <td>Fine Science Tools</td>
 <td> 11295-10</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Student Vannas Spring Scissors </td>
 <td>Fine Science Tools</td>
 <td>91500-09</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Adson Forceps</td>
 <td>Fine Science Tools</td>
 <td>11006-12</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Friendman-Pearson Ronqeur, Curved</td>
 <td>Fine Science Tools</td>
 <td>16221-14</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Iris Scissors - ToughCut</td>
 <td>Fine Science Tools</td>
 <td>14058-11</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Vannas-Tubingen Spring Scissors - 				5mm Blade, Straight</td>
 <td>Fine Science Tools</td>
 <td>15003-08</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td> Scalpel Handle</td>
 <td>Fine Science Tools</td>
 <td>10003-12</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td> Plastic Sterilization Box</td>
 <td>Fine Science Tools</td>
 <td>20810-02</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Stainless Steel Ruler</td>
 <td>Fine Science Tools</td>
 <td>30086-15</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Probe &amp; Spatula - Straight 				14cm</td>
 <td>Fine Science Tools</td>
 <td>10090-13</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Hemostat Straight</td>
 <td>Fine Science Tools</td>
 <td>13002-10</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Hemostat Curved</td>
 <td>Fine Science Tools</td>
 <td>13003-10</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr valign="bottom">
 <td>Micro Drill Stainless Steel Burrs 				- 0.7mm</td>
 <td>Fine Science Tools</td>
 <td>19008-07</td>
 <td>10/Pack</td>
 </tr>
 <tr valign="bottom">
 <td>Micro Drill Stainless Steel Burrs 				- 0.5mm</td>
 <td>Fine Science Tools</td>
 <td>19007-05</td>
 <td>10/Pack</td>
 </tr>
 <tr valign="bottom">
 <td>Self-Tapping Bone Screws - 1.17mm 				(diam) - 4.7mm (length)</td>
 <td>Fine Science Tools</td>
 <td>19010-00</td>
 <td>100/Pack</td>
 </tr>
 <tr valign="bottom">
 <td>Artificial 				Tears</td>
 <td>Fisher</td>
 <td>NC9860842</td>
 <td>Each</td>
 </tr>
 <tr valign="bottom">
 <td>Betadine</td>
 <td>Fisher</td>
 <td>19-066452</td>
 <td>Bottle</td>
 </tr>
 <tr valign="bottom">
 <td>F-AIR Carbon 				Filters</td>
 <td>Fisher</td>
 <td>NC9112250 </td>
 <td>Canister</td>
 </tr>
 <tr valign="bottom">
 <td>Applicator 				Cotton Tipped, Non Sterile</td>
 <td>Materiel 				Services</td>
 <td>1104</td>
 <td>10Pack/Case</td>
 </tr>
 <tr valign="bottom">
 <td>Gauze 2&quot; 				x 2&quot; </td>
 <td>Materiel 				Services</td>
 <td>1630</td>
 <td>25Pack/Case</td>
 </tr>
 <tr valign="bottom">
 <td>Needle tip, 				23 GA x 1&quot; </td>
 <td>Materiel 				Services</td>
 <td>39412</td>
 <td>Sold 				by Case</td>
 </tr>
 <tr valign="bottom">
 <td>Needle tip, 				27 GA x 1.25&quot; </td>
 <td>Materiel 				Services</td>
 <td>25251</td>
 <td>&nbsp;</td>
 </tr>
 <tr valign="bottom">
 <td>Needle tip, 				30 GA x 0.5&quot; </td>
 <td>Materiel 				Services</td>
 <td>22023</td>
 <td>10Box/Case</td>
 </tr>
 <tr valign="bottom">
 <td>Ointment, 				Triple Antibiotic 0.6g</td>
 <td>Materiel 				Services</td>
 <td>2528</td>
 <td>&nbsp;</td>
 </tr>
 <tr valign="bottom">
 <td>Pouch, Self 				Seal Sterilization Pouches, 3.5&quot; x 8&quot;</td>
 <td>Materiel 				Services</td>
 <td>1023</td>
 <td>10Box/Case</td>
 </tr>
 <tr valign="bottom">
 <td>Ringers 				Solution 1000ml </td>
 <td>Materiel 				Services</td>
 <td>5263</td>
 <td>12Bag/Case</td>
 </tr>
 <tr valign="bottom">
 <td>Scalpel 				Blade disposable sterile #15</td>
 <td>Materiel 				Services</td>
 <td>1975</td>
 <td>3Box/Case</td>
 </tr>
 <tr valign="bottom">
 <td>Suture, 				Nylon w/P-3 Needle, 5-0, 18in.</td>
 <td>Materiel 				Services</td>
 <td>4618</td>
 <td>12/Box</td>
 </tr>
 <tr valign="bottom">
 <td>Underpad, 				Disposable 23&quot; x 36&quot;</td>
 <td>Materiel 				Services</td>
 <td>2545</td>
 <td>15Pack/Case</td>
 </tr>
 <tr valign="bottom">
 <td>Lubricant, 				Jelly Surgical Sterile</td>
 <td>Materiel 				Services</td>
 <td>2538</td>
 <td>12Box/Case</td>
 </tr>
 <tr valign="bottom">
 <td>Absorbable 				Foam Gel 2cmx6cmx7mm</td>
 <td>Materiel 				Services</td>
 <td>3107</td>
 <td>&nbsp;</td>
 </tr>
 <tr valign="bottom">
 <td>Six Piece Set-Screw Screwdriver 				Set </td>
 <td>National Jewelers 				Supplies</td>
 <td>ETSCR98001</td>
 <td>&nbsp;</td>
 </tr>
 <tr valign="bottom">
 <td>Kwik Sil 				Adhesive</td>
 <td>WPI</td>
 <td>KWIK-SIL</td>
 <td>&nbsp;</td>
 </tr>
 <tr valign="bottom">
 <td>Kwik Sil 				Tubes</td>
 <td>WPI</td>
 <td>600009</td>
 <td>&nbsp;</td>
 </tr>
 <tr valign="top">
 <td>Chronic silicon electrodes 16 to 				64 sites</td>
 <td>NeuroNexus 				Technologies</td>
 <td>Varies on Probe 				Design</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr valign="top">
 <td>Chronic thin film surface 				electrodes (microECoG) 16 to 32 sites</td>
 <td>NeuroNexus 				Technologies</td>
 <td>Varies on Probe 				Design</td>
 <td>&nbsp;
 </td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Surgical Implantation of Chronic Neural Electrodes for Recording Single Unit Activity and Electrocorticographic Signals at JoVE.com

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

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 ...

surgical-implantation-chronic-neural-electrodes-for-recording-single

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

47.2K Views.  University of Michigan. 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.

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

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Recording Human Electrocorticographic (ECoG) Signals for Neuroscientific Research and Real-time Functional Cortical Mapping

Neuroimaging studies of human cognitive, sensory, and motor processes are usually based on noninvasive techniques such as electroencephalography (EEG), magnetoencephalography or functional magnetic-resonance imaging. These techniques have either inherently low temporal or low spatial resolution, and suffer from low signal-to-noise ratio and/or poor high-frequency sensitivity. Thus, they are suboptimal for exploring the short-lived spatio-temporal dynamics of many of the underlying brain processes. In contrast, the invasive technique of electrocorticography (ECoG) provides brain signals that have an exceptionally high signal-to-noise ratio, less susceptibility to artifacts than EEG, and a high spatial and temporal resolution (i.e., &lt;1 cm/&lt;1 millisecond, respectively). ECoG involves measurement of electrical brain signals using electrodes that are implanted subdurally on the surface of the brain. Recent studies have shown that ECoG amplitudes in certain frequency bands carry substantial information about task-related activity, such as motor execution and planning1, auditory processing2 and visual-spatial attention3. Most of this information is captured in the high gamma range (around 70-110 Hz). Thus, gamma activity has been proposed as a robust and general indicator of local cortical function1-5. ECoG can also reveal functional connectivity and resolve finer task-related spatial-temporal dynamics, thereby advancing our understanding of large-scale cortical processes. It has especially proven useful for advancing brain-computer interfacing (BCI) technology for decoding a user's intentions to enhance or improve communication6 and control7. Nevertheless, human ECoG data are often hard to obtain because of the risks and limitations of the invasive procedures involved, and the need to record within the constraints of clinical settings. Still, clinical monitoring to localize epileptic foci offers a unique and valuable opportunity to collect human ECoG data. We describe our methods for collecting recording ECoG, and demonstrate how to use these signals for important real-time applications such as clinical mapping and brain-computer interfacing. Our example uses the BCI2000 software platform8,9 and the SIGFRIED10 method, an application for real-time mapping of brain functions. This procedure yields information that clinicians can subsequently use to guide the complex and laborious process of functional mapping by electrical stimulation.
Prerequisites and Planning:
Patients with drug-resistant partial epilepsy may be candidates for resective surgery of an epileptic focus to minimize the frequency of seizures. Prior to resection, the patients undergo monitoring using subdural electrodes for two purposes: first, to localize the epileptic focus, and second, to identify nearby critical brain areas (i.e., eloquent cortex) where resection could result in long-term functional deficits. To implant electrodes, a craniotomy is performed to open the skull. Then, electrode grids and/or strips are placed on the cortex, usually beneath the dura. A typical grid has a set of 8 x 8 platinum-iridium electrodes of 4 mm diameter (2.3 mm exposed surface) embedded in silicon with an inter-electrode distance of 1cm. A strip typically contains 4 or 6 such electrodes in a single line. The locations for these grids/strips are planned by a team of neurologists and neurosurgeons, and are based on previous EEG monitoring, on a structural MRI of the patient's brain, and on relevant factors of the patient's history. Continuous recording over a period of 5-12 days serves to localize epileptic foci, and electrical stimulation via the implanted electrodes allows clinicians to map eloquent cortex. At the end of the monitoring period, explantation of the electrodes and therapeutic resection are performed together in one procedure.
In addition to its primary clinical purpose, invasive monitoring also provides a unique opportunity to acquire human ECoG data for neuroscientific research. The decision to include a prospective patient in the research is based on the planned location of their electrodes, on the patient's performance scores on neuropsychological assessments, and on their informed consent, which is predicated on their understanding that participation in research is optional and is not related to their treatment. As with all research involving human subjects, the research protocol must be approved by the hospital's institutional review board. The decision to perform individual experimental tasks is made day-by-day, and is contingent on the patient's endurance and willingness to participate. Some or all of the experiments may be prevented by problems with the clinical state of the patient, such as post-operative facial swelling, temporary aphasia, frequent seizures, post-ictal fatigue and confusion, and more general pain or discomfort.
At the Epilepsy Monitoring Unit at Albany Medical Center in Albany, New York, clinical monitoring is implemented around the clock using a 192-channel Nihon-Kohden Neurofax monitoring system. Research recordings are made in collaboration with the Wadsworth Center of the New York State Department of Health in Albany. Signals from the ECoG electrodes are fed simultaneously to the research and the clinical systems via splitter connectors. To ensure that the clinical and research systems do not interfere with each other, the two systems typically use separate grounds. In fact, an epidural strip of electrodes is sometimes implanted to provide a ground for the clinical system. Whether research or clinical recording system, the grounding electrode is chosen to be distant from the predicted epileptic focus and from cortical areas of interest for the research. Our research system consists of eight synchronized 16-channel g.USBamp amplifier/digitizer units (g.tec, Graz, Austria). These were chosen because they are safety-rated and FDA-approved for invasive recordings, they have a very low noise-floor in the high-frequency range in which the signals of interest are found, and they come with an SDK that allows them to be integrated with custom-written research software. In order to capture the high-gamma signal accurately, we acquire signals at 1200Hz sampling rate-considerably higher than that of the typical EEG experiment or that of many clinical monitoring systems. A built-in low-pass filter automatically prevents aliasing of signals higher than the digitizer can capture. The patient's eye gaze is tracked using a monitor with a built-in Tobii T-60 eye-tracking system (Tobii Tech., Stockholm, Sweden). Additional accessories such as joystick, bluetooth Wiimote (Nintendo Co.), data-glove (5th Dimension Technologies), keyboard, microphone, headphones, or video camera are connected depending on the requirements of the particular experiment.
Data collection, stimulus presentation, synchronization with the different input/output accessories, and real-time analysis and visualization are accomplished using our BCI2000 software8,9. BCI2000 is a freely available general-purpose software system for real-time biosignal data acquisition, processing and feedback. It includes an array of pre-built modules that can be flexibly configured for many different purposes, and that can be extended by researchers' own code in C++, MATLAB or Python. BCI2000 consists of four modules that communicate with each other via a network-capable protocol: a Source module that handles the acquisition of brain signals from one of 19 different hardware systems from different manufacturers; a Signal Processing module that extracts relevant ECoG features and translates them into output signals; an Application module that delivers stimuli and feedback to the subject; and the Operator module that provides a graphical interface to the investigator.
A number of different experiments may be conducted with any given patient. The priority of experiments will be determined by the location of the particular patient's electrodes. However, we usually begin our experimentation using the SIGFRIED (SIGnal modeling For Realtime Identification and Event Detection) mapping method, which detects and displays significant task-related activity in real time. The resulting functional map allows us to further tailor subsequent experimental protocols and may also prove as a useful starting point for traditional mapping by electrocortical stimulation (ECS).
Although ECS mapping remains the gold standard for predicting the clinical outcome of resection, the process of ECS mapping is time consuming and also has other problems, such as after-discharges or seizures. Thus, a passive functional mapping technique may prove valuable in providing an initial estimate of the locus of eloquent cortex, which may then be confirmed and refined by ECS. The results from our passive SIGFRIED mapping technique have been shown to exhibit substantial concurrence with the results derived using ECS mapping10.
The protocol described in this paper establishes a general methodology for gathering human ECoG data, before proceeding to illustrate how experiments can be initiated using the BCI2000 software platform. Finally, as a specific example, we describe how to perform passive functional mapping using the BCI2000-based SIGFRIED system.

Recording Human Electrocorticographic ECoG Signals for Neuroscientific Research and Real-time Functional Cortical Mapping

We present a method for collecting electrocorticographic signals for research purposes from humans who are undergoing invasive epilepsy monitoring. We show how to use the BCI2000 software platform for data collection, signal processing and stimulus presentation. Specifically, we demonstrate SIGFRIED, a BCI2000-based tool for real-time functional brain mapping.

Neuroimaging studies of human cognitive, sensory, and motor processes are usually based on noninvasive techniques such as electroencephalography (EEG), ...

Optical Recording of Suprathreshold Neural Activity with Single-cell and Single-spike Resolution

Signaling of information in the vertebrate central nervous system is often carried by populations of neurons rather than individual neurons. Also propagation of suprathreshold spiking activity involves populations of neurons. Empirical studies addressing cortical function directly thus require recordings from populations of neurons with high resolution. Here we describe an optical method and a deconvolution algorithm to record neural activity from up to 100 neurons with single-cell and single-spike resolution. This method relies on detection of the transient increases in intracellular somatic calcium concentration associated with suprathreshold electrical spikes (action potentials) in cortical neurons. High temporal resolution of the optical recordings is achieved by a fast random-access scanning technique using acousto-optical deflectors (AODs)1. Two-photon excitation of the calcium-sensitive dye results in high spatial resolution in opaque brain tissue2. Reconstruction of spikes from the fluorescence calcium recordings is achieved by a maximum-likelihood method. Simultaneous electrophysiological and optical recordings indicate that our method reliably detects spikes (&gt;97% spike detection efficiency), has a low rate of false positive spike detection (&lt; 0.003 spikes/sec), and a high temporal precision (about 3 msec) 3. This optical method of spike detection can be used to record neural activity in vitro and in anesthetized animals in vivo3,4.

Understanding the function of the vertebrate central nervous system requires recordings from many neurons because cortical function arises on the level of populations of neurons. Here we describe an optical method to record suprathreshold neural activity with single-cell and single-spike resolution, dithered random-access scanning. This method records somatic fluorescence calcium signals from up to 100 neurons with high temporal resolution. A maximum-likelihood algorithm deconvolves the underlying suprathreshold neural activity from the somatic fluorescence calcium signals. This method reliably detects spikes with high detection efficiency and a low rate of false positives and can be used to study neural populations in vitro and in vivo.

Signaling of information in the vertebrate central nervous system is often carried by populations of neurons rather than individual neurons. Also ...

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 ...

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

A Method for Systematic Electrochemical and Electrophysiological Evaluation of Neural Recording Electrodes

New materials and designs for neural implants are typically tested separately, with a demonstration of performance but without reference to other implant characteristics.&#160;This precludes a rational selection of a particular implant as optimal for a particular application and the development of new materials based on the most critical performance parameters.&#160;This article develops a protocol for in vitro and in vivo testing of neural recording electrodes.&#160;Recommended parameters for electrochemical and electrophysiological testing are documented with the key steps and potential issues discussed.&#160;This method eliminates or reduces the impact of many systematic errors present in simpler in vivo testing paradigms, especially variations in electrode/neuron distance and between animal models.&#160;The result is a strong correlation between the critical in vitro and in vivo responses, such as impedance and signal-to-noise ratio.&#160;This protocol can easily be adapted to test other electrode materials and designs.&#160;The in vitro techniques can be expanded to any other nondestructive method to determine further important performance indicators.&#160;The principles used for the surgical approach in the auditory pathway can also be modified to other neural regions or tissue.

Different electrode coatings affect neural recording performance through changes to electrochemical, chemical and mechanical properties. Comparison of electrodes in vitro is relatively simple, however comparison of in vivo response is typically complicated by variations in electrode/neuron distance and between animals. This article provides a robust method to compare neural recording electrodes.

New materials and designs for neural implants are typically tested separately, with a demonstration of performance but without reference to other implant ...

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral cortex. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex using a method developed by Lima and colleagues1. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. This technique &#8211; termed &#8220;PINP&#8221;, or Photostimulation-assisted Identification of Neuronal Populations &#8211; can be implemented with standard extracellular recording equipment. It can serve as an inexpensive and accessible alternative to calcium imaging or visually-guided patching, for the purpose of targeting extracellular recordings to genetically-identified cells. Here we provide a set of guidelines for optimizing the method in everyday practice. We refined our strategy specifically for targeting parvalbumin-positive (PV+) cells, but have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons.

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching.

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral ...

Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and awake animals. Therefore, methods allowing the manipulation of synaptic systems in awake animals are required in order to determine the contribution of synaptic inputs to neuronal processing unaffected by anesthetics. Here, we present methodology for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. By combining these procedures, we performed microiontophoretic injections of gabazine to selectively block GABAA receptors in neurons of the inferior colliculus of head-restrained mice. Gabazine successfully modified neural response properties such as the frequency response area and stimulus-specific adaptation. Thus, we demonstrate that our methods are suitable for recording single-unit activity and for dissecting the role of specific neurotransmitter receptors in auditory processing.
The main limitation of the described procedure is the relatively short recording time (~3 hr), which is determined by the level of habituation of the animal to the recording sessions. On the other hand, multiple recording sessions can be performed in the same animal. The advantage of this technique over other experimental procedures used to manipulate the level of neurotransmission or neuromodulation (such as systemic injections or the use of optogenetic models), is that the drug effect is confined to the local synaptic inputs to the target neuron. In addition, the custom-manufacture of electrodes allows adjustment of specific parameters according to the neural structure and type of neuron of interest (such as the tip resistance for improving the signal-to-noise ratio of the recordings).

We present methods for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. This technique allows the detailed analysis of putative local synaptic inputs to the neuron of interest.

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and ...

Fabrication of High Contact-Density, Flat-Interface Nerve Electrodes for Recording and Stimulation Applications

Many attempts have been made to manufacture multi-contact nerve cuff electrodes that are safe, robust and reliable for long term neuroprosthetic applications. This protocol describes a fabrication technique of a modified cylindrical nerve cuff electrode to meet these criteria. Minimum computer-aided design and manufacturing (CAD and CAM) skills are necessary to consistently produce cuffs with high precision (contact placement 0.51 &#177; 0.04 mm) and various cuff sizes. The precision in spatially distributing the contacts and the ability to retain a predefined geometry accomplished with this design are two criteria essential to optimize the cuff&#39;s interface for selective recording and stimulation. The presented design also maximizes the flexibility in the longitudinal direction while maintaining sufficient rigidity in the transverse direction to reshape the nerve by using materials with different elasticities. The expansion of the cuff&#39;s cross sectional area as a result of increasing the pressure inside the cuff was observed to be 25% at 67 mm Hg. This test demonstrates the flexibility of the cuff and its response to nerve swelling post-implant. The stability of the contacts&#39; interface and recording quality were also examined with contacts&#39; impedance and signal-to-noise ratio metrics from a chronically implanted cuff (7.5 months), and observed to be 2.55 &#177; 0.25 k&#8486; and 5.10 &#177; 0.81 dB respectively.

This article provides a detailed description on the fabrication process of a high contact-density flat interface nerve electrode (FINE). This electrode is optimized for recording and stimulating neural activity selectively within peripheral nerves.

Many attempts have been made to manufacture multi-contact nerve cuff electrodes that are safe, robust and reliable for long term neuroprosthetic ...

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 ...

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

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Chapters in this video

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Recording Human Electrocorticographic (ECoG) Signals for Neuroscientific Research and Real-time Functional Cortical Mapping

Optical Recording of Suprathreshold Neural Activity with Single-cell and Single-spike Resolution

Fiber-optic Implantation for Chronic Optogenetic Stimulation of Brain Tissue

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

A Method for Systematic Electrochemical and Electrophysiological Evaluation of Neural Recording Electrodes

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice

Fabrication of High Contact-Density, Flat-Interface Nerve Electrodes for Recording and Stimulation Applications

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