This work was supported by the Ester and Joseph Klingenstein Fund, the McKnight Endowment Fund, and NINDS 1R01NS066408-01A1.

1. Overview of Components
<ol>
	<li>A complete microdrive consists of several main components (Figure 1): a chassis that serves as the superstructure for the drive, a motor with a finely threaded output shaft, a threaded shuttle that carries the electrodes and provides a point of electrical connection, and an Omnetics (or equivalent) connector.</li>
	<li>The chassis, electrode shuttle, and electrode shuttle tubes are custom components that were designed with 3D CAD software (SolidWorks) and were machined by a local precision machinist. All other components are commercially available.</li>
	<li>Chassis and shuttle are machined from lightweight polyetherimide (PEI), while the electrode shuttle tubes are cut to size with a lathe from stainless steel hypodermic tubing (0.0293"OD x 0.00975"ID). Though these components may be cut from other materials, we have found that PEI and stainless steel have the right balance of machinability, strength, and weight for our application.</li>
	<li>To assemble the drive, a motor controller, capable of spinning the low DC-voltage stepper motor, is required. We employ a custom solution based on a three-phase motor driver IC (Faulhaber BLD05002 available from the manufacturer) in conjunction with an IC timer that provides the timestep-input. A variety of off-the-shelf solutions are also available (e.g. Faulhaber MCBL05002 or Sutter Instruments MP-285).</li>
</ol>
2. Drive Chassis Preparation and Assembly
<ol>
	<li>With fine diagonal cutters, trim the contact pins on the bottom of the Omnetics (or equivalent) connector - 1.5 mm for the rear row, 1 mm for the front row (Figure 2A). To improve the strength of the epoxy bond between the drive chassis and the connector, use a scalpel to roughen the rear surfaces of both components.</li>
	<li>Using a scalpel, cut four 15 mm long sections of polyimide tubing (0.0113" ID x 0.0133" OD) for use as wire guides. Carefully secure the chassis in a small vice with the open cavity down (Figure 2B). Use cyanoacrylate glue to attach the polyimide tubes to the chassis. They should lay flush with the rear chassis surface and be oriented parallel to each other as shown in Figure 2B.</li>
	<li>Once dry, trim the polyimide tubes to approximately 6 mm in length and remove the waste sections. The remaining tubes should be approximately 3 mm from the bottom edge of the chassis.</li>
	<li>To ensure proper clearance for mating with the headstage or cable, the connector should be mounted a few millimeters above the polyimide tubes and be parallel with the chassis surface (Figure 2B). Use epoxy (Torr Seal) to construct a small pedestal for the connector, place it on top with the shorter row of pins topmost, and add more epoxy around the connector. Do not proceed until the epoxy has completely cured.</li>
	<li>Remove the chassis from the vice, transfer to a jig that allows for connection to the motor controller (i.e., in our case, this jig is simply a rigid rod with the mating Omnetics connector affixed to one end) and orient with the front surface of the drive facing up. Guide the motor wires through the hole in the chassis and slide the motor into the groove in the base. The motor and drive shaft should be parallel to the chassis and fit snuggly against the bottom and side.</li>
	<li>Epoxy the motor to the chassis, taking care not to obstruct either the drive shaft or the space where the electrodes will be installed (see Torr Seal placement in Figure 4). Allow the epoxy to cure.</li>
	<li>Rotate the chassis so that the connector is accessible. To support the motor wires and prevent accidental breakage, add a small drop of fast-set epoxy to the wires where they emerge from the chassis. Trim, strip, and solder the wires to the appropriate contacts on the connector (Figure 2A). Test the motor to ensure smooth shaft rotation in both directions.</li>
</ol>
3. Electrode Shuttle Assembly
<ol>
	<li>To minimize the potential for motion-induced artifacts in the recordings, it is essential that the electrode shuttle fit snuggly in the chassis. Test the fit of the shuttle by threading it onto the motor shaft and running it up and down the full length of the shaft at low speed. Be careful not to wedge the shuttle in the drive and thus over-torquing the motor- it is very easy to permanently damage the plastic gearhead at this stage. If a previously spinning motor shaft ceases turning, the gearhead may be damaged.</li>
	<li>If the shuttle does not run smoothly down the shaft, inspect the chassis surfaces and the drive screw for debris. Remove with fine forceps or compressed air, coat the threading in some light mineral oil and recheck the shuttle fit.</li>
	<li>If the shuttle is too tight, use forceps and a fine grain whetstone to remove obstructing material from the side of the shuttle. Take care to remove the material evenly.</li>
	<li>If the shuttle is too loose such that it tilts and chatters during travel, glue a small sliver of transparency film to the shuttle to fill the gap between it and the chassis surface.</li>
	<li>Once the shuttle has been properly fit, remove the shuttle from the shaft and press one of the stainless steel shuttle tubes into each of the four large holes in the shuttle. Each should be centered on the shuttle with the ends of all four tubes flush (Figure 3). Secure them with a small drop of cyanoacrylate glue applied to the base of the tube. Be very careful not to allow the glue to get on the edges of the shuttle or into the tubes.</li>
	<li>Thread the completed electrode shuttle back on to the drive shaft and bring it all the way down to the base of the threading.</li>
</ol>
4. Guide Tube and Electrode Installation
<ol>
	<li>Cut four 25 mm long sections of polyimide tubing (0.0045" ID x 0.006" OD) for use as electrode guides. In addition, cut four electrodes to 40 mm sections for use as dummy electrodes which will hold the guide tubes in position during assembly. (Note that these dummy electrodes are for assembly only and will not be used for recording, so you may reuse the same ones for each assembly.) Slide each section of tube onto on to a dummy electrode such that the electrode extends ~10 mm beyond the end of the tube.</li>
	<li>Arrange each of the polyimide tubes in the chassis as shown in Figure 4A. This will result in the ends of the electrode guides, flush with each other and aligned to the base of the motor shaft, splaying out toward the electrode shuttle and the opposite ends converging at the bottom of the drive. Mix a small amount of Kwik-Cast and apply approximately 2 mm below the top end of the electrode guides. Allow to dry.</li>
	<li>Remove the dummy electrodes and reposition the free ends of the electrode guides such that they form a tight bundle at the bottom of the drive. It may be helpful to use a fine wire to hold them in place temporarily (Figure 4B). Fix the new position with more Kwik-Cast. Allow to dry.</li>
	<li>With a sharp scalpel, trim the electrode guide tubes where they extend beyond the bottom of the microdrive. When implanted, these tubes will guide the electrodes from the bottom of the drive (resting on the skull) to the surface of the brain. Thus, the length at which the tubes should be trimmed depends on the anatomy of the implantation site (Figure 5). For songbirds, this is approximately 1.5 mm.</li>
	<li>To connect an electrode to the connector, cut a 30 mm long section of insulated platinum wire (0.003" dia.) and strip 1 mm of insulation from one end. Solder it to one of the signal pins on the connector. Bend the wire under the connector and push it through one of the wire guide tubes. Pull the wire taunt and wrap it around the guide slot in the top of the chassis.</li>
	<li>With scissors or fine wire cutters, cut an electrode to ~25 mm in length. Take care to do this without bending the electrode or damaging the tip; if either occur, the electrode cannot be used. With the electrode shuttle now at the top of the shaft, insert the electrode into a polyimide tube and pull it up through the stainless steel tube in the shuttle. Position it such that the tip of the electrode is flush with the bottom end of the polyimide tube. Trim the electrode 1 mm above the shuttle tube.</li>
	<li>With a fine set of forceps, strip the insulation off the last 2 mm of the electrode and the last 2 mm of the platinum wire. Reposition the electrode in the shuttle tube, insert the platinum wire into the tube, and pin the two together by sliding a short 1 mm section of tungsten wire (0.008" dia) into the tube. This should be a tight fit that will provide both a mechanical and an electrical connection between the electrode and the wire (Figure 3).</li>
	<li>Run the shuttle up and down the length of the drive shaft to ensure continued movement.</li>
	<li>Repeat 4.5-4.8 until all electrodes have been installed.</li>
	<li>Cut a 30 mm long section of silver wire (0.005" dia) for use as a ground wire. Strip 1 mm of insulation off one end and solder to the ground pin.</li>
</ol>
5. Final Assembly and Pre-Implantation Preparation
<ol>
	<li>To reduce the friction of the electrodes against the guide tubes, fill the guide tubes with light mineral oil. Capillary action is sufficient to fill the tubes from a drop of oil placed at one end.</li>
	<li>To create a protective cover for the motor and electrodes, cut a 12 mm x 25 mm rectangle of transparency and fold it into a wide U-shape. Use cyanoacrylate to glue the cover to the outer surfaces of the chassis (Figure 6). Cut a 15 mm x 6 mm rectangle of transparency and use cyanoacrylate to glue it around the connector. To prevent interference with the connector, the long edge of the transparency should be flush with the top edge of the connector.</li>
	<li>Prior to implantation, sterilize the electrodes and guide tubes by lowering the extended electrodes into a 10:1 DI water-bleach solution for 10 min. Then, flush thoroughly with sterile water.</li>
	<li>A jig similar to that used in step 2.5 can be used to hold the microdrive in a stereotaxic manipulator during implantation.</li>
</ol>
6. Representative Results
This protocol requires approximately 5 hr of hands-on assembly time with an additional 6-8 hr interspersed for the epoxy and glue to dry. However, once the microdrive has been assembled the first time, it can be prepared for reuse (i.e., the electrodes, electrode guide tubes, and electrode wires can be replaced) in less than 2 hr. The quality and character of the recordings obtained by following this protocol will, of course, be in part dependent on the recording target, the choice of electrodes, and any processing performed at the headstage or further upstream of the microdrive. This aside, it is possible to obtain stable recordings from a behaving animal over a time-span of more than ten weeks. Examples of single unit recordings from the robust nucleus of the archistriatum (RA) in a behaving songbird is shown in Figure 7A; multi-unit activity from the same region is shown in Figure 7B.
<img alt="Figure 1" src="/files/ftp_upload/4314/4314fig1.jpg" /> 
Figure 1. 3D model of the assembled motorized microdrive. The microdrive consists of several major components: (i) the chassis, (ii) motor with threaded output shaft, (iii) electrode shuttle, (iv) shuttle tubes, (v) electrodes, (vi) electrode guide tubes, and (vii) connector.
<img alt="Figure 2" src="/files/ftp_upload/4314/4314fig2.jpg" /> 
Figure 2. A) Model and connection diagram for the connector. Motor wires: b, i, l. Electrode wires: c,d,e,f. Ground wire: k. All other pins available for auxiliary instrumentation. B) Photo of the connector attached to the chassis with Torr Seal epoxy. Note the position of both the connector and the underlying wire guide tubes. The cyanoacrylate glue holding the guide tubes in position dries clear and, thus, is not visible in the image.
<img alt="Figure 3" src="/files/ftp_upload/4314/4314fig3.jpg" /> 
Figure 3. Electrode shuttle assembly.
<img alt="Figure 4" src="/files/ftp_upload/4314/4314fig4.jpg" /> 
Figure 4. A) The electrode guide tubes positioned in the chassis prior to applying Kwik-Cast. B) To ensure that the electrodes exit the bottom of the drive in parallel, bring the bottom end of the guide tubes into a tight bundle before adding the last drop of the Kwik-Cast.
<img alt="Figure 5" src="/files/ftp_upload/4314/4314fig5.jpg" /> 
Figure 5. The amount that the electrode guide tubes extend beyond the bottom of the drive is specified by the anatomy of the implantation site. The diagram illustrates the dimensions that are relevant for determining the length of the guide tubes. For implanting in the songbird, 1.5 mm is sufficient.
<img alt="Figure 6" src="/files/ftp_upload/4314/4314fig6.jpg" /> 
Figure 6. The finished motorized microdrive with protective covers attached.
<img alt="Figure 7" src="/files/ftp_upload/4314/4314fig7.jpg" /> 
Figure 7. Representative recordings from the robust nucleus of the archistriatum (RA) in the behaving zebra finch. A) Single-unit activity recorded with 10M&Omega; platinum-iridium electrodes. Top: example recording made one week after implantation. Bottom: example recording from the same electrode as above, nine weeks later. B) Multi-unit activity recorded with 1M&Omega; platinum electrodes. <a href="https://www.jove.com/files/ftp_upload/4314/4314fig7large.jpg" target="_blank"> Click here to view larger figure</a>.
Additional Files: The chassis, shuttle, and electrode tubes were designed with SolidWorks 2010 CAD software. These part files are provided in both the proprietary (*.sldprt) and vendor-neutral (*.iges) formats and dimensioned production drawings are provided in PDF format.

<ol>
	<li>O'Keefe, J., Dostrovsky, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+hippocampus+as+a+spatial+map.">The hippocampus as a spatial map.</a> Brain Research. 34, 171-175 (1971).</li><li>Pennartz, C. M., Berke, J. D., Graybiel, A. M., Ito, R., Lansink, C. S., van der Meer, M., Redish, A. D., Smith, K. S., Voorn, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Corticostriatal Interactions during Learning, Memory Processing, and Decision. 29, 12831-12838 (2009).</li><li>Kepecs, A., Uchida, N., Zariwala, H., Mainen, Z. 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No conflicts of interest declared.

The protocol presented here will result in a device capable of high-quality recordings with minimal motion artifacts only if proper care is taken with construction. The fit of the shuttle in the chassis if of critical importance: too tight and the risk of overloading the motor is high; too loose and the risk of significant motion artifact is high. An ideal fit will allow the shuttle to travel the entire length of the threaded shaft without tilting out of position or chattering.
	Select...

<table border="1">
	<tbody>
 <tr>
 <td>Name of item</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Chassis</td>
 <td>custom</td>
 <td></td>
 <td>Cut from PEI</td>
 </tr>
 <tr>
 <td>Electrode Shuttle</td>
 <td>custom</td>
 <td></td>
 <td>Cut from PEI</td>
 </tr>
 <tr>
 <td>Shuttle Tubes</td>
 <td>custom</td>
 <td></td>
 <td>Cut from Stainless Steel</td>
 </tr>
 <tr>
 <td>Connector</td>
 <td>Omnetics</td>
 <td>A7886-001</td>
 <td>Mates to A7877-001</td>
 </tr>
 <tr>
 <td>Motor w/ Gearhead</td>
 <td>Faulhaber</td>
 <td>0206-A-001-B-021-47:1</td>
 <td></td>
 </tr>
 <tr>
 <td>Wire Guide</td>
 <td>Small Parts, Inc</td>
 <td>SWPT-0113-12</td>
 <td></td>
 </tr>
 <tr>
 <td>Electrode Guide</td>
 <td>Small Parts, Inc</td>
 <td>SWPT-0045-12</td>
 <td></td>
 </tr>
 <tr>
 <td>10M&Omega; Pt-Ir electrodes</td>
 <td>Microprobes, Inc</td>
 <td>PI2PT310.0H3</td>
 <td></td>
 </tr>
 <tr>
 <td>Platinum Wire</td>
 <td>A-M Systems</td>
 <td>772000</td>
 <td>For electrode wires</td>
 </tr>
 <tr>
 <td>Silver Wire</td>
 <td>A-M Systems</td>
 <td>786000</td>
 <td>For ground wire</td>
 </tr>
 <tr>
 <td>Tungsten Wire</td>
 <td>A-M Systems</td>
 <td>797000</td>
 <td>For electrode pins</td>
 </tr>
 <tr>
 <td>Transparency</td>
 <td>3M</td>
 <td>AF4300</td>
 <td></td>
 </tr>
 <tr>
 <td>Torr Seal</td>
 <td>Varian</td>
 <td>9530001</td>
 <td></td>
 </tr>
 <tr>
 <td>Kwik-Cast</td>
 <td>WPI</td>
 <td>KWIK-CAST</td>
 <td></td>
 </tr>
 <tr>
 <td>Cyanoacrylate</td>
 <td>Krazy Glue</td>
 <td>KG517</td>
 <td></td>
 </tr>
 <tr>
 <td>Fast-Set Epoxy</td>
 <td>Hardman</td>
 <td>04001</td>
 <td></td>
 </tr>
 <tr>
 <td>Light Mineral Oil</td>
 <td>Sigma</td>
 <td>M5310</td>
 <td></td>
 </tr>
 <tr>
 <td>Chlorine bleach</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Diagonal cutters</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Scalpel blade</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Forceps</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Drive jig</td>
 <td>custom</td>
 <td></td>
 <td>Epoxy the mating connector to a syringe or stick</td>
 </tr>
 <tr>
 <td>Small Vice</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
	 </tbody>
 </table>

design and assembly of an ultra-light motorized microdrive for chronic neural recordings in small animals

The ability to chronically record from populations of neurons in freely behaving animals has proven an invaluable tool for dissecting the function of neural circuits underlying a variety of natural behaviors, including navigation1 , decision making 2,3, and the generation of complex motor sequences4,5,6. Advances in precision machining has allowed for the fabrication of light-weight devices suitable for chronic recordings in small animals, such as mice and songbirds. The ability to adjust the electrode position with small remotely controlled motors has further increased the recording yield in various behavioral contexts by reducing animal handling.6,7
 Here we describe a protocol to build an ultra-light motorized microdrive for long-term chronic recordings in small animals. Our design evolved from an earlier published version7, and has been adapted for ease-of use and cost-effectiveness to be more practical and accessible to a wide array of researchers. This proven design 8,9,10,11 allows for fine, remote positioning of electrodes over a range of ~ 5 mm and weighs less than 750 mg when fully assembled. We present the complete protocol for how to build and assemble these drives, including 3D CAD drawings for all custom microdrive components.

Watch this Scientific Journal Video about Design and Assembly of an Ultra-light Motorized Microdrive for Chronic Neural Recordings in Small Animals at JoVE.com

Design and Assembly of an Ultra-light Motorized Microdrive for Chronic Neural Recordings in Small Animals

The design, fabrication and assembly of an ultra-light motorized microdrive is described. The device provides a cost-effective and easy-to-use solution for chronic recordings of single units in small behaving animals.

The ability to chronically record from populations of neurons in freely behaving animals has proven an invaluable tool for dissecting the function of ...

design-assembly-an-ultra-light-motorized-microdrive-for-chronic

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JoVE Journal

Neuroscience

12.0K ビュー.  Harvard University. 超軽量電動マイクロドライブの設計、製造、組立が記載されている。デバイスは、小型動物行動における単一ユニットの慢性的な録音のための費用対効果と使いやすいソリューションを提供します。

ビデオ: Design and Assembly of an Ultra-light Motorized Microdrive for Chronic Neural Recordings in Small Animals

Electrode Fabrication and Implantation in Aplysia californica for Multi-channel Neural and Muscular Recordings in Intact, Freely Behaving Animals

Recording from key nerves and muscles of Aplysia during feeding behavior allows us to study the patterns of neural control in an intact animal. Simultaneously recording from multiple nerves and muscles gives us precise information about the timing of neural activity. Previous recording methods have worked for two electrodes, but the study of additional nerves or muscles required combining and averaging the recordings of multiple animals, which made it difficult to determine fine details of timing and phasing, because of variability from response to response, and from animal to animal. Implanting four individual electrodes has a very low success rate due to the formation of adhesions that prevent animals from performing normal feeding movements. We developed a new method of electrode fabrication that reduces the bulk of the electrodes inside the animal allowing for normal feeding movements. Using a combination of glues to attach the electrodes results in a more reliable insulation of the electrode which lasts longer, making it possible to record for periods as long as a week. The fabrication technique that we describe could be extended to incorporate several additional electrodes, and would be applicable to vertebrate animals.

Electrode Fabrication and Implantation in Aplysia californica for Multi-channel Neural and Muscular Recordings in Intact, Freely Behaving Animals

A technique is described for implanting four in vivo electrodes to monitor the neuromuscular control of feeding behavior in Aplysia californica.

Recording from key nerves and muscles of Aplysia during feeding behavior allows us to study the patterns of neural control in an intact animal.  ...

A Thin-skull Window Technique for Chronic Two-photon In vivo Imaging of Murine Microglia in Models of Neuroinflammation

Traditionally in neuroscience, in vivo two photon imaging of the murine central nervous system has either involved the use of open-skull1,2 or thinned-skull 3 preparations. While the open-skull technique is very versatile, it is not optimal for studying microglia because it is invasive and can cause microglial activation. Even though the thinned-skull approach is minimally invasive, the repeated re-thinning of skull required for chronic imaging increases the risks of tissue injury and microglial activation and allows for a limited number of imaging sessions. Here we present a chronic thin-skull window method for monitoring murine microglia in vivo over an extended period of time using two-photon microscopy. We demonstrate how to prepare a stable, accessible, thinned-skull cortical window (TSCW) with an apposed glass coverslip that remains translucent over the course of three weeks of intermittent observation. This TSCW preparation is far more immunologically inert with respect to microglial activation than open craniotomy or repeated skull thinning and allows an arbitrary number of imaging sessions during a time period of weeks. We prepare TSCW in CX3CR1 GFP/+ mice 4 to visualize microglia with enhanced green fluorescent protein to &le;150 &mu;m beneath the pial surface. We also show that this preparation can be used in conjunction with stereotactic brain injections of the HIV-1 neurotoxic protein Tat, adjacent to the TSCW, which is capable of inducing durable microgliosis. Therefore, this method is extremely useful for examining changes in microglial morphology and motility over time in the living brain in models of HIV Associated Neurocognitive Disorder (HAND) and other neurodegenerative diseases with a neuroinflammatory component.

A Thin-skull Window Technique for Chronic Two-photon In vivo Imaging of Murine Microglia in Models of Neuroinflammation

We describe a method for repeatedly visualizing murine microglia and circulating monocytes in vivo over hours, days or weeks using transcranial two-photon microscopy. We demonstrate how to prepare a thinned-skull window that allows intermittent observation of quiescent microglia that can be activated by adjacent stereotactic injection of the HIV-1 regulatory protein Tat.

Traditionally in neuroscience, in vivo two photon imaging of the murine central nervous system has either involved the use of open-skull1,2 or ...

Combining Behavioral Endocrinology and Experimental Economics: Testosterone and Social Decision Making

Behavioral endocrinological research in humans as well as in animals suggests that testosterone plays a key role in social interactions. Studies in rodents have shown a direct link between testosterone and aggressive behavior1 and folk wisdom adapts these findings to humans, suggesting that testosterone induces antisocial, egoistic or even aggressive behavior2. However, many researchers doubt a direct testosterone-aggression link in humans, arguing instead that testosterone is primarily involved in status-related behavior3,4. As a high status can also be achieved by aggressive and antisocial means it can be difficult to distinguish between anti-social and status seeking behavior.
We therefore set up an experimental environment, in which status can only be achieved by prosocial means. In a double-blind and placebo-controlled experiment, we administered a single sublingual dose of 0.5 mg of testosterone (with a hydroxypropyl-&beta;-cyclodextrin carrier) to 121 women and investigated their social interaction behavior in an economic bargaining paradigm. Real monetary incentives are at stake in this paradigm; every player A receives a certain amount of money and has to make an offer to another player B on how to share the money. If B accepts, she gets what was offered and player A keeps the rest. If B refuses the offer, nobody gets anything. A status seeking player A is expected to avoid being rejected by behaving in a prosocial way, i.e. by making higher offers.
The results show that if expectations about the hormone are controlled for, testosterone administration leads to a significant increase in fair bargaining offers compared to placebo. The role of expectations is reflected in the fact that subjects who report that they believe to have received testosterone make lower offers than those who say they believe that they were treated with a placebo. These findings suggest that the experimental economics approach is sensitive for detecting neurobiological effects as subtle as those achieved by administration of hormones. Moreover, the findings point towards the importance of both psychosocial as well as neuroendocrine factors in determining the influence of testosterone on human social behavior.

The procedure described in this protocol shows that testosterone administration and folk beliefs about testosterone may be associated with directly opposed social behaviors.

Behavioral endocrinological research in humans as well as in animals suggests that testosterone plays a key role in social interactions. Studies in ...

High-density EEG Recordings of the Freely Moving Mice using Polyimide-based Microelectrode

Electroencephalogram (EEG) indicates the averaged electrical activity of the neuronal populations on a large-scale level. It is widely utilized as a noninvasive brain monitoring tool in cognitive neuroscience as well as a diagnostic tool for epilepsy and sleep disorders in neurology. However, the underlying mechanism of EEG rhythm generation is still under the veil. Recently introduced polyimide-based microelectrode (PBM-array) for high resolution mouse EEG1 is one of the trials to answer the neurophysiological questions on EEG signals based on a rich genetic resource that the mouse model contains for the analysis of complex EEG generation process. This application of nanofabricated PBM-array to mouse skull is an efficient tool for collecting large-scale brain activity of transgenic mice and accommodates to identify the neural correlates to certain EEG rhythms in conjunction with behavior. However its ultra-thin thickness and bifurcated structure cause a trouble in handling and implantation of PBM-array. In the presented video, the preparation and surgery steps for the implantation of PBM-array on a mouse skull are described step by step. Handling and surgery tips to help researchers succeed in implantation are also provided.

In this article, we described the surgery procedure and handling tips for implantation of ultra-thin polyimide-based microelectrode array (PBM-array) on the mouse skull for acquisition of high-density encephalography (EEG) in a mouse model.

Electroencephalogram (EEG) indicates the averaged electrical activity of the neuronal populations on a large-scale level. It is widely utilized as a ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges.&#160;Methods for performing this technique and the equipment necessary are described.&#160;Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell ...

Design and Fabrication of Ultralight Weight, Adjustable Multi-electrode Probes for Electrophysiological Recordings in Mice

The number of physiological investigations in the mouse, mus musculus, has experienced a recent surge, paralleling the growth in methods of genetic targeting for microcircuit dissection and disease modeling. The introduction of optogenetics, for example, has allowed for bidirectional manipulation of genetically-identified neurons, at an unprecedented temporal resolution. To capitalize on these tools and gain insight into dynamic interactions among brain microcircuits, it is essential that one has the ability to record from ensembles of neurons deep within the brain of this small rodent, in both head-fixed and freely behaving preparations. To record from deep structures and distinct cell layers requires a preparation that allows precise advancement of electrodes towards desired brain regions. To record neural ensembles, it is necessary that each electrode be independently movable, allowing the experimenter to resolve individual cells while leaving neighboring electrodes undisturbed. To do both in a freely behaving mouse requires an electrode drive that is lightweight, resilient, and highly customizable for targeting specific brain structures.
A technique for designing and fabricating miniature, ultralight weight, microdrive electrode arrays that are individually customizable and easily assembled from commercially available parts is presented. These devices are easily scalable and can be customized to the structure being targeted; it has been used successfully to record from thalamic and cortical regions in a freely behaving animal during natural behavior.

Understanding the neural substrates of behavior requires brain circuit ensemble recording. Because of its genetic tractability, the mouse offers a model for circuit dissection and disease mimicry. Here, a method of designing and fabricating miniaturized probes is described that is suitable for targeting deep brain structure in the mouse.

The number of physiological investigations in the mouse, mus musculus, has experienced a recent surge, paralleling the growth in methods of genetic ...

Simultaneous Recordings of Cortical Local Field Potentials, Electrocardiogram, Electromyogram, and Breathing Rhythm from a Freely Moving Rat

Monitoring the physiological dynamics of the brain and peripheral tissues is necessary for addressing a number of questions about how the brain controls body functions and internal organ rhythms when animals are exposed to emotional challenges and changes in their living environments. In general experiments, signals from different organs, such as the brain and the heart, are recorded by independent recording systems that require multiple recording devices and different procedures for processing the data files. This study describes a new method that can simultaneously monitor electrical biosignals, including tens of local field potentials in multiple brain regions, electrocardiograms that represent the cardiac rhythm, electromyograms that represent awake/sleep-related muscle contraction, and breathing signals, in a freely moving rat. The recording configuration of this method is based on a conventional micro-drive array for cortical local field potential recordings in which tens of electrodes are accommodated, and the signals obtained from these electrodes are integrated into a single electrical board mounted on the animal's head. Here, this recording system was improved so that signals from the peripheral organs are also transferred to an electrical interface board. In a single surgery, electrodes are first separately implanted into the appropriate body parts and the target brain areas. The open ends of all of these electrodes are then soldered to individual channels of the electrical board above the animal's head so that all of the signals can be integrated into the single electrical board. Connecting this board to a recording device allows for the collection of all of the signals into a single device, which reduces experimental costs and simplifies data processing, because all data can be handled in the same data file. This technique will aid the understanding of the neurophysiological correlates of the associations between central and peripheral organs.

This study introduces a method for the simultaneous recording of local field potentials in the brain, electrocardiograms, electromyograms, and breathing signals of a freely moving rat. This technique, which reduces experimental costs and simplifies data analysis, will contribute to the understanding of the interactions between the brain and peripheral organs.

Monitoring the physiological dynamics of the brain and peripheral tissues is necessary for addressing a number of questions about how the brain controls ...

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 ...

Rabies Necropsy Techniques in Large and Small Animals

The New York State Department of Health (NYSDOH) Rabies Laboratory receives between 6,000 to 9,000 specimens annually and performs rabies testing for the entire state, with the exception of New York City. The Rabies laboratory necropsies a variety of animals ranging in size from bats to bovids. Most of these specimens are animals exhibiting neurological signs, however, less than 10% actually test positive for rabies; implying trauma, lesions or other infectious agents as the cause of these symptoms. Due to the risk of aerosolizing undiagnosed infectious agents, the Rabies Laboratory does not use power tools or saws. Three necropsy techniques will be presented for animals whose skulls are impenetrable with scissors. The laboratory has implemented these techniques to decrease potential exposure to infectious agents, eliminate unnecessary manipulation of the specimen and reduce processing time. The advantages of a preferred technique opposed to another is subject to the trained individual processing the specimen.

The goal of this protocol is to demonstrate safe necropsy techniques in small and large animals to obtain satisfactory tissue samples for rabies testing.

The New York State Department of Health (NYSDOH) Rabies Laboratory receives between 6,000 to 9,000 specimens annually and performs rabies testing for the ...