We would like to acknowledge Jun Yamamoto for his design and construction of the automatic, motorized tetrode twister device depicted in the video. 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.

1. Fabrication of tetrodes
<ol>
<li>Begin the construction of one tetrode by obtaining a 50 cm long, polyimide insulated, ni-chrome wire with a 12.5 micron diameter core. Fold the wire in half. Run your fingers along the pair to make them stick together. Make sure the pair of wires have good contact throughout their entire length.</li>
<li>Fold the pair again by holding the two ends together. Make sure that the loop formed on one end is not kinked. Cut the four wires on the non-looped end so that the four tips are aligned.</li>
<li>For the next step, you will need a modified alligator clip, a motorized turning device, and a horizontal bar above the turning device. The alligator clip is modified by gluing a plastic bar to the base of the clip. Where the wire tips are exposed, clamp the four wires together with the modified alligator clip. Hang the loop of wires over the horizontal bar. Place the alligator clip into the motorized stage.</li>
<li>Apply 80 clock-wise twists followed by 40 counter clock-wise twists to the wire bundle over the course of approximately three minutes. These parameters can be varied according to your needs.</li>
<li>After tetrode twisting is completed, fuse the wires together by heating from three different angles with a heat gun (420&deg; C or 790&deg; F), using medium-low flow. For each angle, begin 1-2 cm below where the wire bundle splits, run the heat gun down and up at a 2 cm distance from the wires for about 5 seconds. Use caution as high temperatures will completely melt the insulation and lead to short circuits. </li>
<li>Now that the wires have fused together, remove the tetrode from the twisting apparatus by gently lifting the alligator clip to relieve tension on the tetrode, and cut the tetrode near the alligator clip. At the other end, cut the loop such that there are four non-bonded strands of wire of equal length.</li>
<li>Next, separate the individual strands at the top by gently bending the wires with a soft tipped tweezer. </li>
<li>The tetrode is now ready for loading into the micro-drive array. Make 21 to 25 more tetrodes and store them in a dust free box until it is time for the tetrode loading process.</li>
</ol>
2. Loading the tetrodes and ground wires into the micro-drive array
<ol>
<li>To proceed, you will need a complete micro-drive array. If you have not yet built one, refer to the video 'Micro-drive array for chronic in-vivo recording: drive fabrication.' In our design plan, the tetrode will be attached at one end to connector hardware, and will run through the polyimide carrier tube in the micro-drive such that the electrode tips extend below the base of the micro-drive array.</li>
<li>Before beginning, build a drive holder that can be attached to the connector board on one end, and on the other end, can be clamped by a panavise. In this example, the drive holder is a mill-max connector glued to one end of an X-ACTO knife handle (Figure 1c).</li>
<li>Holding the tetrode with soft tipped tweezers and using a stereoscope, push the tip of the tetrode into the polyimide carrier tube of one of the micro-drives. Push the tetrode into the tube until the individual electrode wires are close to the connector board at the top of the drive array. Be careful not to kink or bend the wires, as that will cause the tetrode to enter the brain at an angle, rather than perpendicular, and will weaken the integrity of the tetrode.</li>
<li>Gently feed the other end of the four wires into their respective holes in the electrode interface board using soft tipped tweezers. Again, avoid kinking or bending the wires. With all four wires in place, push the gold pins into their holes with a pair of pliers that have a shortened lower jaw. When the pins are pushed into the hole, they will strip the wire insulation and create an electrical connection.</li>
<li>For later reference, keep track of the mapping between micro-drives and pin position on the electrode interface board. </li>
<li>Continue loading a total of 18 tetrodes. The three remaining micro-drives on the array will be used to house the reference electrodes. Each reference electrode is made of a tetrode that uses only one of the four unbonded wire ends. To attach the reference electrodes, follow the loading procedure but attach only one of the four electrodes to the respective reference electrode pin on the connector board. </li>
<li>If you are planning multi-site recordings, it may be worthwhile to plan out the connections from the tetrodes to the connector board ahead of time. This should minimize the possibility of adjusting the wrong tetrode during experiments.</li>
<li>Once tetrode loading is completed, make two ground wires, one for the animal and one for the protective cone. Cut one 6' long insulated steel wire (text: 0.005' diameter) and remove 3 mm of insulation, using metal tweezers, from each end of the wire. Then cut a 4' long insulated steel wire (text: 0.005' diameter). Remove 3 mm of insulation at one end and 1 cm at the other end. </li>
<li>Route the 6' ground wire through the hole on the side of the micro-drive array and up towards the interface board. Connect the exposed wire to the electrode interface board with a gold pin at the designated ground hole. This wire will connect to the animal's skull.</li>
<li> Route the 4' ground wire parallel to the previously installed ground wire. Connect the 3 mm stripped end to the electrode interface board using a gold pin at another designated ground hole. This wire will later be connected to a protective cone, which will act as a small Faraday cage to reduce noise pickup. We are now finished connecting wires and can proceed with refining our tetrodes.</li>
</ol>
3. Setting the length of the tetrodes
<ol>
<li>Our goal in this process is to cut all the tetrodes so that they are slightly longer than the depth of the target brain structure. In our example, the micro-drives are designed for a maximum travel of 5-6 mm, sufficient to reach many neocortical areas and the dorsal hippocampus of an adult rat. First, lower all the micro-drives so that the tetrodes are maximally exposed. Simultaneously cut all the tetrodes to lengths that are approximately 5 mm longer than the desired final length using a pair of sharp fine scissors. </li>
<li>For the final cut, prepare a pair of serrated scissors in a panavise. Clamp one handle with a slight angle downwards and with serrations facing upwards, while leaving the other handle dangling.</li>
<li>Now completely withdraw all the tetrodes into their guide cannula by turning the micro-drives up.</li>
<li>Using a stereoscope, fully extend one tetrode out from its cannula. Using a ruler, position the serrated scissors at the desired distance from the guide cannula. Slowly and gently cut the tetrode with one smooth movement. After this tetrode has been cut, withdraw the micro-drive up by 3-4mm. Repeat this process for each tetrode until all have been cut. At this point you are ready for gold plating. </li>
</ol>
4. Gold plating of tetrodes and quality check
Gold plating the nichrome tetrode wires is critical for long term stable recordings. It prevents corrosion and improves biocompatibility.
<ol>
<li>Setup the impedance meter in external mode and the current generator in DC check mode with audio output (Figure 1d). Touch the two leads of the current generator to the six possible pairs of channels within each tetrode, and use the audio feedback to determine if shorts exist between pairs. If a short exists, re-cut the tetrode with serrated scissors and test again. If the problem persists, discard the tetrode and replace it.</li>
<li>When no shorts exist within all the tetrodes, move all the micro-drives down, extending all the tetrodes as far out as possible. </li>
<li>Modify the equipment configuration for checking electrode impedance (Figure 1e). Now dip the tips of all the tetrodes into a bath of standard gold-plating solution. The gold bath is electrically connected to the positive lead of the impedance meter. Check the impedance of the electrode by touching the negative lead of the impedance meter to the corresponding pin on the connector board.</li>
<li>Measure and record the impedance of every wire on the micro-drive array. Normal values range from 1-3 MOhm. </li>
<li>On the current generator, set the current between 1-3 &mu;A. Next, securely attach the negative lead of the impedance meter to the pin on the connector board. Place the equipment into the electroplating configuration (Figure 1f). On the impedance meter, quickly switch from normal mode, to bypass mode, and back. While in bypass mode, current will pass through the tetrode and gold solution, and gold will be plated onto the electrode tip. The impedance after each round of plating should be less than before. Repeat the current pulse if the impedance did not drop below 1 MOhm. A conservative range of acceptable impedances are 250kOhm to 350 kOhm. If the impedance drops below 200 kOhm, this may indicate a short circuit.</li>
<li>If after repeated pulses the impedance does not drop sufficiently, the electrode tip may be obstructed. Re-cut the tetrode, check for shorts, and repeat the gold-plating process. If re-cutting doesn't help, replace the tetrode and start again. </li>
<li>After plating all tetrodes, dip the tetrode tips briefly in ethanol, and let them dry for 3 to 5 minutes.</li>
<li>Test for short-circuits within tetrodes using the current generator with the tetrode tips exposed to air. If a DC connection exists between two or more electrodes of a tetrode, re-cut the tetrode, and plate with gold again.</li>
<li>After gold-plating of all the tetrodes is complete, add a drop of medium thickness cyano-acrylate glue to the interface between each tetrode and its polyimide tubing. This will secure the tetrode to its micro-drive and maintain the length of the tetrode. </li>
<li>As a final step, move all the tetrodes into the drive by turning all the screws up.</li>
</ol>5. Attaching the protective cone
The protective cone's purpose is to shield micro-drives and the exposed tetrode wire from the environment. It also provides support for handling during an experiment and reduces electrical noise. We use a 3-D printed plastic cone (Reference Part I: drive fabrication). However, it is possible to make the cone from other materials, like a bent piece of soda can.
<ol>
<li>Take the plastic cone and fix some aluminum foil tape to the inner or outer surface.</li>
<li>Next, drill a small hole (1.5 mm in diameter) through the cone and loosely attach a screw with a washer and nut inside the cone. This will serve as an electrical passage for the ground wire. We recommend using an 0-80 size screw and washer combination.</li>
<li>First, insert and partially turn three 1-72 size screws (3/16 inches in length) into the sides of the cone. With the drive array suspended, insert the cone over the drive. Make sure both ground wires extend from the bottom of the drive array. Wrap the exposed part of the 4' ground wire around one of the 1-72 screws several times and fasten the screw tightly. Secure the cone to the drive base by tightening the remaining two cone screws.</li>
<li>Lastly, take the 1 cm stripped end of the short ground wire and wind it around the screw, in between the foil tape and washer. Secure the ground by tightening the screw. Now we have a micro-drive array that is at last ready for surgical implantation.</li>
</ol>6. Representative results
<ol>
 <li>Micro-drive array production is now complete. Your micro-drive array is loaded with 18 tetrodes and 3 reference channels. It is encased in a protective cone and cap for durability. The finished micro-drive array with protective cone weighs 20g to 25g on average.</li>
 <li>The finished drive array is implanted using standard surgical procedure. As you can see, the drive is attached to a preamplifier chip and a bundle of wires that carry the signal to data acquisition hardware and software.</li>
</ol>
<img src="/files/ftp_upload/1098/CompositeFigure1_highqualityn.jpg" alt="figure 1" /> Figure 1: A) Schematic of tetrode twisting configuration. A horizontal bar suspends the unbonded electrode wires of the tetrode above the automated tetrode twisting device. B) Setup for tetrode impedance measurement and electrode plating. When checking for short circuits, the circuit is different: the black wire is attached to another electrode channel on the connector board. C) A close-up of a custom made micro-drive array holder tool. The cylindrical bar is an X-Acto knife handle. The connector is a 26-pin, double row, Mill-max connector. The two pieces are bonded together using dental acrylic. D) Configuration for short checking. Channel X and Channel Y are any two different channels of the four tetrode channels. E) Configuration for measuring electrode impedance. The current source is 'off'. The impedance meter is set to 'on' and 'test'. F) Configuration for gold plating of electrode tips. The current source is set to 'on', audio is 'on', mode is 'unipolar', DC Test is 'on', current is set to 2 microamps.

<ol>
	<li>Gray, C. M., Maldonado, P. E., Wilson, M., McNaughton, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tetrodes+markedly+improve+the+reliability+and+yield+of+multiple+single-unit+isolation+from+multi-unit+recordings+in+cat+striate+cortex.">Tetrodes markedly improve the reliability and yield of multiple single-unit isolation from multi-unit recordings in cat striate cortex.</a> J. Neurosci. Methods. 63 (1-2), 43-54 (1995).</li></ol>

This protocol is the second installment of a two part protocol: &ldquo;Micro-drive array for chronic in-vivo recording.&rdquo; The combination of the micro-drive array and tetrode is a powerful tool for obtaining simultaneous recordings of many neurons in an awake, behaving animal preparation. Hopefully, we have provided you with all the necessary information for you will need to begin or improve electrophysiological recordings in your own laboratory.
The motorized tetrode twister device wa...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Panavise</td>
<td>Tool</td>
<td>Panavise</td>
<td>Model 301</td>
<td>Holds either micro-drive array or serrated scissors</td>
</tr>
<tr>
<td>Modified Xcelite 378M Pliers</td>
<td>Tool</td>
<td>Newark</td>
<td>96F8903</td>
<td>For inserting connector board pins. To make, remove 2mm from one tip of the pliers.</td>
</tr>
<tr>
<td>Slow Cure Glue</td>
<td>Material</td>
<td>Great Planes (Champaign, IL)</td>
<td>GPMR6015</td>
<td>To glue the tetrode to the micro-drive</td>
</tr>
<tr>
<td>Gold Plating Solution</td>
<td>Material</td>
<td>Neuralynx</td>
<td>Sifco Selectron Gold NC</td>
<td>Contains gold particles for electroplating</td>
</tr>
<tr>
<td>Current Generator</td>
<td>Tool</td>
<td>World Precision Instruments</td>
<td>A365D-B</td>
<td>A.K.A. Stimulus Isolator. Used to electroplate gold solution onto electrode tips.</td>
</tr>
<tr>
<td>Impedance Meter</td>
<td>Tool</td>
<td>BAK Electronics</td>
<td>IMP-1 / 6662 / 2788</td>
<td>To check impedance of electrodes</td>
</tr>
<tr>
<td>Hot Air Gun</td>
<td>Tool</td>
<td>Steinel (Germany)</td>
<td>HG 3002 LCD</td>
<td>To fuse electrodes together to make tetrode</td>
</tr>
<tr>
<td>Accu-Tek Carbofib Tip Tweezers</td>
<td>Tool</td>
<td>Aven (Ann Arbor, MI)</td>
<td>18768 (Pattern 304)</td>
<td>For handling tetrodes</td>
</tr>
<tr>
<td>Alligator Clip</td>
<td>Tool</td>
<td>Fine Science Tools</td>
<td>18050-35</td>
<td>A.K.A. Bulldog serrefine. To clip the tetrodes together during twisting</td>
</tr>
<tr>
<td>Micro Dissecting Scissors</td>
<td>Tool</td>
<td>Biomedical Research Instruments</td>
<td>25-1000</td>
<td>For general cutting of tetrode/electrode wire</td>
</tr>
<tr>
<td>Iris Toughcut Serrated Scissors</td>
<td>Tool</td>
<td>Fine Science Tools</td>
<td>14058-09</td>
<td>For final cut of tetrodes</td>
</tr>
<tr>
<td>Teflon Coated Stainless Steel Wire</td>
<td>Material</td>
<td>A-M Systems (Carlsborg, WA)</td>
<td>790500</td>
<td>Ground wire</td>
</tr>
<tr>
<td>12.5 micron nichrome wire</td>
<td>Material</td>
<td>Kanthal Palm Coast</td>
<td>Rediohm-800 1/4 HARD PAC COATING</td>
<td>Polyimide coated micro-wire for making tetrodes.</td>
</tr>
<tr>
<td>Tetrode Twister</td>
<td>Tool</td>
<td>Neuralynx</td>
<td>Tetrode Assembly Station</td>
<td>Increases the speed and consistency of tetrode fabrication.

</td>
</tr>
<tr>
<td>Silver Paint</td>
<td>Material</td>
<td>GC Electronics</td>
<td>22-023</td>
<td>For connecting the micro-electrode wires of the tetrode to the connector pins (see Discussion)</td>
</tr>
<tr>
<td>X-Acto Knife Handle</td>
<td>Material</td>
<td>MSC Direct</td>
<td>X3001</td>
<td>For constructing a micro-drive holding device.</td>
</tr>
<tr>
<td>26 Pin, 2 Row Connector</td>
<td>Material</td>
<td>Mill-max</td>
<td>852-10-026-30-001000</td>
<td>For constructing a micro-drive holding device.</td>
</tr>		</tbody>
		</table>
		</div>

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.



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

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.

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 ...

micro-drive-array-for-chronic-in-vivo-recording-tetrode-assembly

Research

JoVE Journal

Biology

33.1K Views.  MIT - Massachusetts Institute of Technology. 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.

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

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 ...

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.

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.

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

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 ...

Assembly, Loading, and Alignment of an Analytical Ultracentrifuge Sample Cell

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed centrifugation. When properly planned and executed, an AUC sedimentation velocity or sedimentation equilibrium experiment can reveal a great deal about a protein in regards to size and shape, sample purity, sedimentation coefficient, oligomerization states and protein-protein interactions.
This technique, however, requires a rigorous level of technical attention. Sample cells hold a sectored center piece sandwiched between two window assemblies. They are sealed with a torque pressure of around 120-140 in/lbs. Reference buffer and sample are loaded into the centerpiece sectors and then after sealing, the cells are precisely aligned into a titanium rotor so that the optical detection systems scan both sample and reference buffer in the same radial path midline through each centerpiece sector while rotating at speeds of up to 60, 000 rpm and under very high vacuum
Not only is proper sample cell assembly critical, sample cell components are very expensive and must be properly cared for to ensure they are in optimum working condition in order to avoid leaks and breakage during experiments. Handle windows carefully, for even the slightest crack or scratch can lead to breakage in the centrifuge. The contact between centerpiece and windows must be as tight as possible; i.e. no Newton s rings should be visible after torque pressure is applied. Dust, lint, scratches and oils on either the windows or the centerpiece all compromise this contact and can very easily lead to leaking of solutions from one sector to another or leaking out of the centerpiece all together. Not only are precious samples lost, leaking of solutions during an experiment will cause an imbalance of pressure in the cell that often leads to broken windows and centerpieces. In addition, plug gaskets and housing plugs must be securely in place to avoid solutions being pulled out of the centerpiece sector through the loading holes by the high vacuum in the centrifuge chamber. Window liners and gaskets must be free of breaks and cracks that could cause movement resulting in broken windows.
This video will demonstrate our procedures of sample cell assembly, torque, loading and rotor alignment to help minimize component damage, solution leaking and breakage during the perfect AUC experiment.

The analytical ultracentrifuge (AUC) sample cell holds sample and reference buffer and during experiments and is exposed to high vacuum and rotor speeds up to 60,000 rpm. This video will demonstrate the rigorous attention to detail necessary for assembly, loading and alignment of this very important component of an AUC experiment.

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed ...

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, ...

Ambulatory ECG Recording in Mice

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although the surface ECG is utilized for short-term measurements of waveform intervals, it is not practical for long-term studies of heart rate variability or the capture of rare episodes of arrhythmias. Implantable ECG telemeters offer the advantages of simple surgical implantation, long-term recording of electrograms in ambulatory mice, and scalability with simultaneous recordings of multiple animals. Here, we present a step-by-step guide to the implantation of telemeters for ambulatory ECG recording in mice. Careful adherence to aseptic technique is required for favorable survival results with the possibility of implantation and recording from weeks to months. Thus, implantable ECG telemetry is a valuable tool for detection of critical information on cardiac electrophysiology in ambulatory animal models such as the mouse. 

Telemetric ECG has emerged as an essential tool in evaluating animal models for cardiac arrhythmias and sudden cardiac death. Here, we present a stepwise guide to telemetric ECG recordings for application in long-term ambulatory ECG monitoring in mice.

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although ...

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 ...

Measuring DNA Damage and Repair in Mouse Splenocytes After Chronic In Vivo Exposure to Very Low Doses of Beta- and Gamma-Radiation

Low dose radiation exposure may produce a variety of biological effects that are different in quantity and quality from the effects produced by high radiation doses. Addressing questions related to environmental, occupational and public health safety in a proper and scientifically justified manner heavily relies on the ability to accurately measure the biological effects of low dose pollutants, such as ionizing radiation and chemical substances. DNA damage and repair are the most important early indicators of health risks due to their potential long term consequences, such as cancer. Here we describe a protocol to study the effect of chronic in vivo exposure to low doses of &#947;- and &#946;-radiation on DNA damage and repair in mouse spleen cells. Using a commonly accepted marker of DNA double-strand breaks, phosphorylated histone H2AX called &#947;H2AX, we demonstrate how it can be used to evaluate not only the levels of DNA damage, but also changes in the DNA repair capacity potentially produced by low dose in vivo exposures. Flow cytometry allows fast, accurate and reliable measurement of immunofluorescently labeled &#947;H2AX in a large number of samples. DNA double-strand break repair can be evaluated by exposing extracted splenocytes to a challenging dose of 2 Gy to produce a sufficient number of DNA breaks to trigger repair and by measuring the induced (1 hr post-irradiation) and residual DNA damage (24 hrs post-irradiation). Residual DNA damage would be indicative of incomplete repair and the risk of long-term genomic instability and cancer. Combined with other assays and end-points that can easily be measured in such in vivo studies (e.g., chromosomal aberrations, micronuclei frequencies in bone marrow reticulocytes, gene expression, etc.), this approach allows an accurate and contextual evaluation of the biological effects of low level stressors.

Measuring DNA Damage and Repair in Mouse Splenocytes After Chronic In Vivo Exposure to Very Low Doses of Beta- and Gamma-Radiation

A protocol to evaluate changes in DNA damage levels and DNA repair capacity that may be induced by chronic in vivo low dose irradiation in mouse spleen lymphocytes, by measuring phosphorylated histone H2AX, a marker of DNA double-strand breaks, using flow cytometry is presented.

Low dose radiation exposure may produce a variety of biological effects that are different in quantity and quality from the effects produced by high ...

The Assembly and Application of 'Shear Rings': A Novel Endothelial Model for Orbital, Unidirectional and Periodic Fluid Flow and Shear Stress

Deviations from normal levels and patterns of vascular fluid shear play important roles in vascular physiology and pathophysiology by inducing adaptive as well as pathological changes in endothelial phenotype and gene expression. In particular, maladaptive effects of periodic, unidirectional flow induced shear stress can trigger a variety of effects on several vascular cell types, particularly endothelial cells. While by now endothelial cells from diverse anatomic origins have been cultured, in-depth analyses of their responses to fluid shear have been hampered by the relative complexity of shear models (e.g., parallel plate flow chamber, cone and plate flow model). While these all represent excellent approaches, such models are technically complicated and suffer from drawbacks including relatively lengthy and complex setup time, low surface areas, requirements for pumps and pressurization often requiring sealants and gaskets, creating challenges to both maintenance of sterility and an inability to run multiple experiments. However, if higher throughput models of flow and shear were available, greater progress on vascular endothelial shear responses, particularly periodic shear research at the molecular level, might be more rapidly advanced. Here, we describe the construction and use of shear rings: a novel, simple-to-assemble, and inexpensive tissue culture model with a relatively large surface area that easily allows for a high number of experimental replicates in unidirectional, periodic shear stress studies on endothelial cells.

The Assembly and Application of &#039;Shear Rings&#039;: A Novel Endothelial Model for Orbital, Unidirectional and Periodic Fluid Flow and Shear Stress

Different levels and patterns of fluid shear are known to modulate endothelial gene expression, phenotype and susceptibility to disease. We discuss the assembly and use of 'shear rings': a model that produces unidirectional, periodic shear stress patterns. Shear rings are simple to assemble, economical and can produce high cell yields.

Deviations from normal levels and patterns of vascular fluid shear play important roles in vascular physiology and pathophysiology by inducing adaptive as ...