We would like to acknowledge Linus D. Sun and Jeenah Jung, who originated the use of computer-designed, 3-D printed drives in our lab for recording from mice. In addition, we would like to acknowledge all past and present Wilson Lab members for their contributions to micro-drive and tetrode technology development in the lab.

<ol>
<li>Micro-drive array overview A complete micro-drive array consists of several main components (Fig. 1):<ol>
<li>A computer designed plastic base that holds 21 independently movable micro-drives. </li>
<li>A cannula at the bottom of the base in which a set of stationary tetrode guide tubes is collected.</li>
<li>21 micro-drives that each carry one tetrode and which can be individually driven using a screw.</li>
<li>An electrode interface board for interconnection between tetrodes and pre-amplifiers.</li>
<li>A protective cone (not shown in Fig. 1) </li>
<li>In a fully assembled micro-drive array, the plastic base holds all micro-drives. Each micro-drive carries a single tetrode. The tetrode is electrically connected to the electrode interface board at the top of the micro-drive array, and passes through the stationary guide tube to exit towards the brain at the bottom of the micro-drive array. Turning the screw on each micro-drive advances or retracts the tetrode.</li>
</ol></li>
 
<li>Design of plastic base We design the base of the micro-drive assembly in a 3-D CAD software package (SolidWorks). The advantage of this in-house design process is rapid turnover and flexibility of design. In the Wilson lab we have designed many variants of this part, geared towards the specific demands of each experiment. The 3-D model file is sent to a service bureau where it is printed from a laser-curable liquid resin using stereolithography. Turnaround time is as little as 2 days. The resulting part is lightweight, reusable, and can be machined to hold a thread. In this protocol we will use a drive base that is designed with a single recording site and which holds 21 micro-drives. The SolidWorks design file is made available upon request.</li>
 
<li>Preparation of plastic base The plastic base is processed to accommodate the micro-drives and the screws to attach the electrode interface board and the protective cone. It is not possible to reliably print smooth through holes in the plastic base, so we design undersized holes, then expand them by drilling. Carefully align the drill bit with the axis of the existing hole before drilling at very low speed. A variable speed/torque hand drill works well for this task.<ol>
<li>Drill the 21 pilot holes for the micro-drive screws (the outer ring) using a #61 bit.</li>
<li>Drill the 21 holes for the the micro-drive support tubes (the inner ring) using a #65 bit. </li>
<li>Drill the holes that will be used to attach the electrode interface board and protective cone using a 1.55 mm bit. </li>
<li>Press fit a sleeve (5 mm long 20 Ga thin wall stainless steel tubing) into each of the inner ring of holes using a pair of tweezers or a pair of pliers so that it is flush with the plastic base. The sleeves ensure smooth movement of the micro-drive support tubes. Next, all holes that will hold screws are tapped. In printed plastics, thread-forming taps result in more durable threads than typical thread-cutting taps. This style of tap requires larger pilot holes. Using a hand-held pin vise to hold the tap:</li>
 
<li>Tap the holes for the micro-drive screws using a tap that matches the screw size being used (M1.2x0.25) and a dry lubricant. To help align the tap with the pilot hole, insert a length of 23 Ga tubing into the sleeve next to it.</li>
<li>Our plastic base design features a tap starter for each micro-drive screw hole to make sure the initial threads are not damaged during the threading process. After all holes are threaded, these tap starters can be easily snapped off using a razor blade.</li>
<li>Tap the holes that will hold the screws for the electrode interface board and the protective cone using a 1-72 tap. </li>
</ol></li>
 
<li>Insertion of electrode guide tubes <ol> 
<li>To make the cannula that holds all the stationary electrode guide tubes, cut a 1.5cm-long piece of 13 Ga stainless steel tubing and roughen the surface with a dremel grinding wheel to enhance adhesion by the dental acrylic.</li>
<li>Insert this cannula into the hole at the bottom of the plastic base, verify that it is co-linear with the vertical axis of the drive, and secure it with dental acrylic or epoxy. Leave 5 to 7 mm of the cannula exposed to facilitate surgical implantation.</li>
<li>Cut 21 stationary electrode guide tubes (flexible polyimide tubing, inner diameter: 0.0071", outer diameter: 0.0116") with a razor blade to a length of 6&nbsp; cm. </li>
<li>Feed the guide tubes one at a time into the sleeves at the top of the drive base and all the way through the collector cannula at the bottom. Visually check for kinks or excessive bending and replace tubes if necessary. To facilitate the insertion and parallel alignment of the tetrode guide tubes, one can start with first feeding the entire bundle of 21 guide tubes up into the collector cannula from below. Then, one at a time, remove a guide tube from the bundle and feed it from the top of the plastic base through the hole that it left behind in the bundle. </li>
<li>Push the guide tubes down until they extend only 1-2 mm from the top of the plastic base. Apply a small amount of thin cyanoacrylate glue to the tubes as they exit the collector cannula, being careful not to let any glue flow up into the end of the guide tubes.</li>
<li>Use a fresh razor blade to cut off the excess from the bundle of guide tubes at the bottom of the collector cannula.</li>
 </ol> </li>
<li>Fabrication of micro-drives Each micro-drive consists of a custom-machined screw and a hollow support tube (Fig. 2). The screw and support tube are attached together such that a lip on the screw is embedded in acrylic, leaving the screw to rotate freely (Fig. 2B). The screw is threaded into the plastic base, and the support tube is then driven up or down when the screw is turned, advancing or retracting the tetrode that it carries. For ease of adjustment on the behaving animal, the head of the screw is in the shape of a half-cylinder. Make a custom screwdriver for this screw: <ol>
<li>Cut 5 cm of 15 Ga stainless steel tubing.</li>
<li>Take one of the custom screws and grind off the protruding lip.</li>
<li>Insert the screw into the tubing until the top of the screw is flush with the end of the tubing. </li>
<li>Crush the tubing around the screw threads repeatedly with a pair of locking pliers so that the screw cannot rotate relative to the cannula. </li>
<li>Use a pin vise to hold the custom screw driver. To fabricate the micro-drives we use a 3-D printed plastic mold with small wells (3 mm x 5 mm, 2 mm deep). Each well has two holes in the bottom surface, spaced 2 mm apart, for the support tube and the screw. Alternatively, molds can be fabricated from an engineering plastic like Delrin, using a drill press to create the well and holes.</li>
 
<li>Prepare the mold by clearing the hole for the support cannula using a 23 Ga needle and tapping the hole for the screw as described for the plastic base, above.</li>
<li>Line the inside of the well with teflon lubricant or a thin film of petroleum jelly to allow for release of the cured dental acrylic from the mold.. </li>
<li>Apply teflon lubricant to a screw and insert it into the mold using the custom made screwdriver until the threads are just below the bottom of the well. </li>
<li>Roughen the top 2-3 mm of a 14 mm-long support tube (23 Ga stainless steel tubing) with a &nbsp;grinding wheel to enhance adhesion of the dental acrylic. Insert it into the mold until the top of the cannula is halfway between the top of the well and the top of the screw.</li>
<li>Pour dental acrylic into the well. Agitating motion is used to ensure that the dental acrylic flows into all the spaces around the screw and support tube. Remove any air bubbles with a small needle. </li>
<li>Wait until the dental acrylic is completely cured (15-30 minutes) and then remove the micro-drive from the mold by turning the screw counter-clock wise. </li>
<li>It is important to do a quality check of the micro-drives. Inspect the cured dental acrylic for cracks and/or air bubbles, which are usually caused by dental acrylic that was too thick or too thin. Also, make sure the screw and the support tube are parallel to each other and that the screw turns smoothly. Screws and support tubes of micro-drives that do not meet these criteria can be cracked free from the dental acrylic and recycled. </li>
<li>Repeat steps 5.6 through 5.11 until you have 21 micro-drives. </li>
</ol></li>
 
<li>Final assembly of micro-drive array<ol> 
<li>Lower each micro-drive into the plastic drive base. Each micro-drive support tube should move smoothly both inside its sleeve and over its stationary guide tube. Check to make sure that the guide tubes do not buckle as the micro-drive is being lowered. </li>
<li>With the micro-drives completely lowered, add a thin layer of dental acrylic to the interior of the drive base to fix the electrode guide tubes in place. </li>
<li>Note that failure to lower the micro drives all the way before adding the dental acrylic will result in wicking of the acrylic all the way up the sleeves and obstruction of micro-drive movement. </li>
<li>Label the micro-drives 1-21. </li>
<li>Turn the drive array upside down and take a picture of the bundle of guide tubes. This picture will be used to map the location of the guide tube corresponding to each micro-drive. </li>
<li>Raise all the micro-drives by several mm.</li>
<li>Insert a polyimide carrier tube (0.005") into each guide tube from the bottom of the drive base. Let the carrier tube extend 1-2 mm from the top of the fully lowered micro-drive and and record on the photograph the identity of the corresponding micro-drive.</li>
<li>Glue the carrier tube to the micro-drive support tube using 5-minute epoxy or cyanoacrylate glue. Note that glue or epoxy that is too thin will flow down the support cannula and obstruct free motion of the micro-drive or, worse, fix it to the guide cannula. If this happens, that particular micro-drive unit can not be used. </li>
<li>Fully lower all the micro-drives. Cut all the carrier tubes off flush at the bottom of the collector cannula using a fresh razor blade.</li>
<li>Finally, mount the Neuralynx electrode interface board to the drive base using two screws (size: 1-72, length: 3/16"). </li>
</ol></li>
</ol>
At this point the drive array is ready to be loaded with tetrodes. Fabrication of tetrodes, how to load them into the micro-drives, and how to prepare the drive array for surgical implantation are the subjects of a subsequent video article.
<img src="/files/ftp_upload/1094/Figure1.png" alt="Figure 1" /> Figure 1. Model of finished micro-drive array. A micro-drive array consists of several major components: a plastic drive base; a collector cannula from which the tetrodes enter the brain; 21 micro-drives that each drive a single tetrode; an electrode interface board that connects to pre-amplifiers.
<img src="/files/ftp_upload/1094/Figure2.png" alt="Figure 2" /> Figure 2. Custom micro-drive screws. A. Technical drawings of the custom screw design. The screw features a threaded part (M1.2x0.25; 1.2 mm diameter, 0.25 mm / turn), a half-cylinder head that matches our custom made screw driver and a smooth part with lip (see detail). B. Model of complete micro-drive. The support tube and custom screw are connected with dental cement. The lip on the screw is embedded in the dental cement and allows for free rotation while driving the support tube up or down.
Additional Files
We designed the printed drive base, the protective cone and the mold for construction of the micro-drives in SolidWorks 2008 3D CAD software:
<ul style="list-style-type:none;">
<li>Download the file:<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/1094/DriveBase.SLDPRT"> DriveBase.sldprt</a> which contains models of the drive base and protective cone and cap.</li>
<li>Download the file:<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/1094/MicroDriveMold.SLDPRT"> MicroDriveMold.sldprt</a> which contains model of the micro-drive mold</li>
</ul>
The 3D printing service bureau can not handle native Solidworks files directly, but rather requires the design files in stereolithography CAD format (STL). For people who do not have access to Solidworks software, but still would like to use the printed parts used in the video article we include the designs in STL format as well:
<ul style="list-style-type:none;">
<li>Download the file:<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/1094/DriveBase.STL"> DriveBase.stl</a> which contains model of drive base</li>
<li>Download the file:<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/1094/MicroDriveMold.STL"> MicroDriveMold.stl</a> which contains model of micro-drive mold</li>
<li>Download the file:<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/1094/ProtectionCap.STL"> ProtectionCap.stl</a> which contains model of cap for protective cone</li>
<li>Download the file:<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/1094/ProtectionCone.STL"> ProtectionCone.stl</a> which contains model of protective cone</li>
</ul>
Fixes to these files, and additional models, are tracked on <a href="https://github.com/wilsonlab/CAD-Files/tree/master/stl">github</a>.

<ol>
	<li>Yamamoto, J., Wilson, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Large-scale+chronically+implantable+precision+motorized+microdrive+array+for+freely+behaving+animals.">Large-scale chronically implantable precision motorized microdrive array for freely behaving animals.</a> J. Neurophysiol. 100 (4), 2430-2440 (2008).</li></ol>

This protocol describes the general features of the micro-drive building process. It has been successfully modified to produce drive arrays with multiple recording targets or smaller arrays for recordings in mice. In addition to recording from the hippocampus, many researchers use these drives to target cortical and sub-cortical structures. The use of longer screws may be required in order to record from deeper brain structures. With extensive modification to the drive base, motorized drives my be used in lieu of manually-turned ones, allowing more precise, remote tetrode adjustment (Yamamoto and Wilson, 2008). Customized stationary guide cannula arrangements have been used in place of the circular main cannula to achieve spatial control over the distribution of tetrodes in the cortex and hippocampus.

<table>
<tbody>
	<tr>
<td>3D-printed plastic parts</td>
<td>Material</td>
<td>American Precision Prototyping (www.approto.com)</td>
<td>&nbsp;</td>
<td>Parts are printed by stereolithography (SLA) using DSM 14120 'ABS-like', DSM 11120 'Watershed' or Accura 55 resins. Request high resolution, no sandblasting.</td>
</tr>
<tr>
<td>stainless steel tubing</td>
<td>Material</td>
<td>Small Parts (www.smallparts.com)</td>
<td>&nbsp;</td>
<td>Cut to custom length and deburred. Sizes: 13 Ga (1pc @ 10 mm), 15 Ga (1pc @ 50 mm), 20 Ga thin wall (21pcs @ 5 mm), 23 Ga (21pcs @ 14 mm) </td>
</tr>
<tr>
<td>Dental cement powder</td>
<td>Material</td>
<td>A-M Systems (www.a-msystems.com)</td>
<td>525000</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Dental cement solvent</td>
<td>Material</td>
<td>A-M Systems</td>
<td>526000</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Electrode interface board</td>
<td>Material</td>
<td>Neuralynx (www.neuralynx.com)</td>
<td>EIB-81</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>micro-drive screws</td>
<td>Material</td>
<td>Advanced Machining and Tooling (www.amtmfg.com)</td>
<td>&nbsp;</td>
<td>Custom made screws with M1.2x0.25 threads (1.2mm major diameter; 0.25 mm thread pitch) </td>
</tr>
<tr>
<td>'Tap-Ease' tapping lubricant </td>
<td>Tool</td>
<td>AGS Company</td>
<td>TA-2</td>
<td>Available from MSC Industrial Supply (www.mscdirect.com)</td>
</tr>
<tr>
<td>Polyimide tubing </td>
<td>Material</td>
<td>IWG High Performance Conductors (www.iwghpc.com)</td>
<td>&nbsp;</td>
<td>Sizes:
30 Ga (outer diam: 0.0071", inner diam: 0.0116") 
5 mil (nominal outer diam: 0.0055", wall: 0.0010") </td>
</tr>
<tr>
<td>Cordless drill/driver </td>
<td>Tool</td>
<td>DeWalt (www.dewalt.com)</td>
<td>DC750KA </td>
<td>Any drill/driver with low-speed setting will do.</td>
</tr>
<tr>
<td>Variable-Speed Rotary Tool Kit with Flex-Shaft </td>
<td>Tool</td>
<td>Dremel (www.dremel.com)</td>
<td>3956-02</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Razor blades </td>
<td>Tool</td>
<td>VWR (www.vwr.com)</td>
<td>55411-050</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Watch glass </td>
<td>Tool</td>
<td>VWR</td>
<td>66112-107</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Tap for custom screws</td>
<td>Tool</td>
<td>Balax</td>
<td>01302</td>
<td>M1.2x0.25 Thredfloer thread-forming tap. Available from MSC Industrial Supply.</td>
</tr>
<tr>
<td>Taps</td>
<td>Tool</td>
<td>Small Parts</td>
<td>HSBT-0172, HSBT-0080 </td>
<td>1-72, 0-80 bottoming taps</td>
</tr>
<tr>
<td>Pin vise</td>
<td>Tool</td>
<td>Vargus</td>
<td>3SHC1--I </td>
<td>To hold taps. Available from MSC Industrial Supply</td>
</tr>
<tr>
<td>Drill bits</td>
<td>Tool</td>
<td>Small Parts</td>
<td>&nbsp;</td>
<td>Sizes: 65, 61, 1.55mm</td>
</tr>
<tr>
<td>Benchtop vise</td>
<td>Tool</td>
<td>Panavise</td>
<td>301</td>
<td>Available from DigiKey (www.digikey.com).</td>
</tr>
<tr>
<td>No.2 Dumont forceps</td>
<td>Tool</td>
<td>Stoelting (www.stoeltingco.com)</td>
<td>52100-27</td>
<td>Used for inserting 20 Ga thin wall sleeves.</td>
</tr>
<tr>
<td>5-minute epoxy</td>
<td>Tool</td>
<td>Allied Electronics (www.alliedelec.com)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Accu-Tek Carbofib Tip Tweezers</td>
<td>Tool</td>
<td>Aven (www.aveninc.com)</td>
<td>18768 (Pattern 304)</td>
<td>Used for handling polyimide tubing.</td>
</tr>
<tr>
<td>Micro Dissecting Scissors</td>
<td>Tool</td>
<td>Biomedical Research Instruments (www.biomedinstr.com)</td>
<td>25-1000</td>
<td>Used for cutting polyimide carrier tubes</td>
</tr>
<tr>
<td>Xcelite 378M Pliers</td>
<td>Tool</td>
<td>Newark</td>
<td>96F8903</td>
<td>Used for handling stainless steel tubing and for construction of custom screwdriver.</td>
</tr>
<tr>
<td>mini pin vise</td>
<td>Tool</td>
<td>MSC Industrial Supply (www.mscdirect.com)</td>
<td>00920314 </td>
<td>Used as a handle for custom made screwdriver.</td>
</tr>		
</tbody>
</table>

micro-drive array for chronic in vivo recording: drive fabrication

Chronic recording of large populations of neurons is a valuable technique for studying the function of neuronal circuits in awake behaving rats. Lightweight recording devices carrying a high density array of tetrodes allow for the simultaneous monitoring of the activity of tens to hundreds of individual neurons. Here we describe a protocol for the fabrication of a micro-drive array with twenty one independently movable micro-drives. This device has been used successfully to record from hippocampal and cortical neurons in our lab. We show how to prepare a custom designed, 3-D printed plastic base that will hold the micro-drives. We demonstrate how to construct the individual micro-drives and how to assemble the complete micro-drive array. Further preparation of the drive array for surgical implantation, such as the fabrication of tetrodes, loading of tetrodes into the drive array and gold-plating, is covered in a subsequent video article.

Watch this Scientific Journal Video about Micro-drive Array for Chronic in vivo Recording: Drive Fabrication at JoVE.com

Micro-drive Array for Chronic in vivo Recording: Drive Fabrication

In this protocol we demonstrate how to fabricate a micro-drive array for chronic electrophysiological recordings in rats.

Micro-drive Array for Chronic in vivo Recording: Drive Fabrication

Chronic recording of large populations of neurons is a valuable technique for studying the function of neuronal circuits in awake behaving rats. ...

micro-drive-array-for-chronic-in-vivo-recording-drive-fabrication

Nanyang Technological University

Research

JoVE Journal

Biology

25.2K Views.  MIT - Massachusetts Institute of Technology. In this protocol we demonstrate how to fabricate a micro-drive array for chronic electrophysiological recordings in rats.

Video: Micro-drive Array for Chronic in vivo Recording: Drive Fabrication

Population Replacement Strategies for Controlling Vector Populations and the Use of Wolbachia pipientis for Genetic Drive

In this video, Jason Rasgon discusses population replacement strategies to control vector-borne diseases such as malaria and dengue. "Population replacement" is the replacement of wild vector populations (that are competent to transmit pathogens) with those that are not competent to transmit pathogens. There are several theoretical strategies to accomplish this. One is to exploit the maternally-inherited symbiotic bacteria Wolbachia pipientis. Wolbachia is a widespread reproductive parasite that spreads in a selfish manner at the extent of its host's fitness. Jason Rasgon discusses, in detail, the basic biology of this bacterial symbiont and various ways to use it for control of vector-borne diseases.

In this interview, Jason Rasgon explains the concept of genetic drive and the characteristics of an effective gene drive system. The use of the endosymbiotic bacterium, Wolbachia pipientis, as a means to spread genes through mosquito populations, is hypothesized.

In this video, Jason Rasgon discusses population replacement strategies to control vector-borne diseases such as malaria and dengue. "Population ...

Application of Light-cured Dental Adhesive Resin for Mounting Electrodes or Microdialysis Probes in Chronic Experiments

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these acrylics do not bond to the bone, screws have been used as anchors. However, in small experimental animals like finches or mouse, their craniums are very fragile and can not successfully hold the anchors. In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium. Therefore, anchor screws are not required and surgical field can be reduced considerably. Past experiences show that the bonding effect maintains more than 2 months. Conventional resin's window of time when the materials are pliable and workable is a few minutes. However, the window of working time for these dental adhesives is significantly wider and adjustable.

In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium.

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these ...

Immunohistochemistry on Paraffin Sections of Mouse Epidermis Using Fluorescent Antibodies

In the epidermis, immunohistochemistry is an efficient means of localizing specific proteins to their relative expression compartment; namely the basal, suprabasal, and stratum corneum layers. The precise localization within the epidermis of a particular protein lends clues toward its functional role within the epidermis. In this chapter, we describe a reliable method for immunolocalization within the epidermis modified for both frozen and paraffin sections that we use very routinely in our laboratory. Paraffin sections generally provide much better morphology, hence, superior results and photographs; however, not all antibodies will work with the harsh fixation and treatment involved in their processing. Therefore, the protocol for frozen sectioning is also included. Within paraffin sectioning, two fixation protocols are described (Bouin's and paraformaldehyde); the choice of fixative will be directly related to the antibody specifications and may require another fixing method.

We describe a reliable method for immunolocalization within the epidermis modified for both frozen and paraffin sections that we use very routinely in our laboratory.

In the epidermis, immunohistochemistry is an efficient means of localizing specific proteins to their relative expression compartment; namely the basal, ...

A Craniotomy Surgery Procedure for Chronic Brain Imaging

Imaging techniques are becoming increasingly important in the study brain function. Among them, two-photon laser scanning microscopy has emerged as an extremely useful method, because it allows the study of the live intact brain. With appropriate preparations, this technique allows the observation of the same cortical area chronically, from minutes to months. In this video, we show a preparation for chronic in vivo imaging of the brain using two-photon microscopy. This technique was initially pioneered by Dr. Karel Svoboda, who is now a Howard Hughes Medical Institute Investigator at Janelia Farm. Preparations like the one shown here can be used for imaging of neocortical structure (e.g., dendritic and axonal dynamics),  to record neuronal activity using calcium-sensitive dyes, to image cortical blood flow dynamics, or for intrinsic optical imaging studies. Deep imaging of the neocortex is possible with optimal cranial window surgeries. Operating under the most sterile conditions possible to avoid infections, together with using extreme care to do not damage the dura mater during the surgery, will result in successful and long-lasting glass-covered cranial windows.


This video and protocol demonstrate how to implant a glass-covered cranial window in rodents. These preparations can be used for chronic in vivo two-photon imaging of the neocortex over time scales of months. It may also be used for other types of imaging, including optical intrinsic signal imaging.

Imaging techniques are becoming increasingly important in the study brain function. Among them, two-photon laser scanning microscopy has emerged as an ...

Micro-drive Array for Chronic in vivo Recording: Tetrode Assembly

The tetrode, a bundle of four electrodes, has proven to be a valuable tool for the simultaneous recording of multiple neurons in-vivo.  The differential amplitude of action potential signatures over the channels of a tetrode allows for the isolation of single-unit activity from multi-unit signals.  The ability to precisely control the stereotaxic location and depth of the tetrode is critical for studying coordinated neural activity across brain regions.  In combination with a micro-drive array, it is possible to achieve precise placement and stable control of many tetrodes over the course of days to weeks.  In this protocol, we demonstrate how to fabricate and condition tetrodes using basic tools and materials, install the tetrodes into a multi-drive tetrode array for chronic in-vivo recording in the rat, make ground wire connections to the micro-drive array, and attach a protective cone onto the micro-drive array in order to protect the tetrodes from physical contact with the environment.



Micro-drive Array for Chronic in vivo Recording: Tetrode Assembly

In this protocol we demonstrate how to fabricate and condition tetrodes for use with a micro-drive array, which was designed for chronic electrophysiological recordings in rats. In addition, we illustrate the final stages of micro-drive array construction, which includes installing ground wires and a protective cone.

The tetrode, a bundle of four electrodes, has proven to be a valuable tool for the simultaneous recording of multiple neurons in-vivo.  The differential ...

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

In vivo-like Organotypic Murine Retinal Wholemount Culture

Targeted ablations of genes and analysis of animal models is the classical strategy for enrolling specific retinal gene function. However, transgenic, retina-specific or conditional knockout mouse models often display early lethality or suffer from severe malformations, preventing an analysis beyond embryonic or early postnatal stages.
Primary cell culture is an alternative to investigate the effects of exogenously applied recombinant factors, overexpression of genes or siRNA-mediated gene knockdown in a controlled environment. Dissociated cell culture has the advantage that the endogenous signals reaching the target cells are reduced, thereby facilitating the identification of exogenously triggered effects after pharmacological manipulation. However, important cell-cell interactions are initially destroyed by enzymatic digestion or mechanical dissociation, even if re-aggregated retinospheroid cultures1 are used.
By contrast, organotypic retinal wholemount cultures provide a system close to the physiological in vivo situation with neuronal interactions and connections still preserved2-5.
In this video article we provide a step by step demonstration of (1) the establishment of in vivo-like organotypic retinal wholemount cultures including dissection peculiarities of embryonic, postnatal and adult murine eyes and (2) a dissociation and cytospin procedure for analysis of neuronal apoptosis and retinal cell proliferation in organotypic wholemounts, e.g. after culture in the presence of exogenously applied recombinant factors.

In vivo-like Organotypic Murine Retinal Wholemount Culture

This video article demonstrates the establishment of organotypic retinal wholemount cultures and a cytospin procedure for analysis of exogenously induced effects. Organotypic retinal wholemount cultures mimic the in vivo situation and significantly facilitate the accessibility of murine retinas for experimental manipulations while circumventing the disadvantages of classical murine animal models.

Targeted ablations of genes and analysis of animal models is the classical strategy for enrolling specific retinal gene function. However, transgenic, ...

One-channel Cell-attached Patch-clamp Recording

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate depends on properties intrinsic to each channel protein as well as the mechanism by which it is generated and controlled and represents an important area of current research. Information about the operational dynamics of ion channel proteins can be obtained by observing long stretches of current produced by a single molecule. Described here is a protocol for obtaining one-channel cell-attached patch-clamp current recordings for a ligand gated ion channel, the NMDA receptor, expressed heterologously in HEK293 cells or natively in cortical neurons. Also provided are instructions on how to adapt the method to other ion channels of interest by presenting the example of the mechano-sensitive channel PIEZO1. This method can provide data regarding the channel&#8217;s conductance properties and the temporal sequence of open-closed conformations that make up the channel&#8217;s activation mechanism, thus helping to understand their functions in health and disease.

Described here is a procedure for obtaining long stretches of current recording from one ion channel with the cell-attached patch-clamp technique. This method allows for observing, in real time, the pattern of open-close channel conformations that underlie the biological signal. These data inform about channel properties in undisturbed biological membranes.

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate ...

Array Comparative Genomic Hybridization (Array CGH) for Detection of Genomic Copy Number Variants

Array CGH for the detection of genomic copy number variants has replaced G-banded karyotype analysis. This paper describes the technology and its application in a clinical diagnostic service laboratory. DNA extracted from a patient&#8217;s sample (blood, saliva or other tissue types) is labeled with a fluorochrome (either cyanine 5 or cyanine 3). A reference DNA sample is labeled with the opposite fluorochrome. There follows a cleanup step to remove unincorporated nucleotides before the labeled DNAs are mixed and resuspended in a hybridization buffer and applied to an array comprising ~60,000 oligonucleotide probes from loci across the genome, with high probe density in clinically important areas. Following hybridization, the arrays are washed, then scanned and the resulting images are analyzed to measure the red and green fluorescence for each probe. Software is used to assess the quality of each probe measurement, calculate the ratio of red to green fluorescence and detect potential copy number variants.

Array Comparative Genomic Hybridization Array CGH for Detection of Genomic Copy Number Variants

Array CGH for the detection of genomic copy number variants has replaced G-banded karyotype analysis. This paper describes the technology and its application in a diagnostic service laboratory.

Array CGH for the detection of genomic copy number variants has replaced G-banded karyotype analysis. This paper describes the technology and its ...

In Vivo Microinjection and Electroporation of Mouse Testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

In Vivo Microinjection and Electroporation of Mouse Testis

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...