The authors thank Mrs. Paulette McKenna and Lisa Pezzino in this study for secretarial support, and Mr. Phil Auerbach for technical support. The authors also thank Drs. Michael Ariel and Michael S. Jones (Saint Louis University School of Medicine) for introducing us to the in vitro isolated head preparation. Funding for support of this collaboration was provided by the Department of Biology (Robert S. Chase Fund), the Academic Research Committee, and the Neuroscience Program at Lafayette College. Lastly, this work is dedicated to Mr. Phil Auerbach, who passed away 28 September 2016; he decommissioned a scanning electron microscope and recognized the usefulness of its 5-axis stage for use in this protocol. His friendship and resourcefulness will be missed greatly.

NOTE: Red-eared slider turtles, both male and female, were purchased from a vendor. Turtles were housed in a warm animal suite containing two 60-gallon tubs equipped with brick islands for sunning under 250-W infrared lights. The environment was maintained on a 14/10-h light/dark cycle with the water temperature at 22 &#176;C. Lights were turned on at 6:00 am and turned off at 8:00 pm. The tanks equipped with filtering systems were cleaned weekly, and turtles were fed ad libitum every other day. The care of red-eared slider turtles and all of the following experimental procedures described here32,33 were approved by the Institutional Animal Care and Use Committee (IACUC) at Lafayette College.
1. Equipment Setup
<ol>
	<li>Prepare turtle Ringer&#39;s solution. Add the following to distilled water in this order: sodium chloride 96.5 mM (58.44 g/mol), potassium chloride 2.6 mM (74.56 g/mol), magnesium chloride 2.0 mM (203.31 g/mol), sodium bicarbonate 31.5 mM (84.01 g/mol), dextrose 20.0 mM (180.16 g/mol), concentrated hydrochloric acid to adjust the pH to 7.51, and calcium chloride 4.0 mM (110.98 g/mol) (see Table of Materials). Mix the solution while adding each salt. 
	CAUTION: Concentrated HCl is hazardous (skin, eyes, inhalation, and ingestion hazards).</li>
	<li>Make tips for the suction electrodes from 5-cm-long capillary glass (see Table of Materials), by fire-polishing to different sizes of inner diameters in order to accommodate cranial nerves of varying thickness.
	<ol>
		<li>Use a small file to etch a line across a piece of capillary glass. Place in&#160;tissue paper and break in half.</li>
		<li>Slowly roll one of the ends of the capillary glass into the flame of a Bunsen burner. Periodically examine the tip for size, smoothness, and symmetry using a dissection scope and a fiber optic light source (see Table of Materials). 
		NOTE: For turtles with head widths between 20 and 30 mm, optimal fitting inner diameter sizes typically range from 0.4 to 0.8 mm for the oculomotor nerve (nIII), 0.3 to 0.6 mm for the trochlear nerve (nIV), and 0.2 to 0.4 mm for the abducens nerve (nVI).</li>
	</ol>
	</li>
	<li>Clean and organize Rongeurs, a blunt dissection probe, microscissors, fine forceps, curved forceps, and a scalpel handle with blades (see Table of Materials) for the dissection. 
	NOTE: Sterilization of instruments is optional.</li>
</ol>
2. Anesthesia and Euthanasia
<ol>
	<li>Place the turtle in an ice bucket for 60 min to cryoanesthetize it.</li>
	<li>Euthanize the turtle by decapitation using a small animal guillotine (see Table of Materials).
	<ol>
		<li>Gently pry the jaws of the animal open with a small weighing spatula so that a hook can be inserted and turned to fit under the tip of the upper jaw.</li>
		<li>Pull with steady pressure to extend the animal&#39;s head through the guillotine. Swiftly decapitate the animal.</li>
	</ol>
	</li>
	<li>Place turtle head in a dissection dish. Have enough turtle Ringer&#8217;s solution on hand to irrigate the tissue. Oxygenize the solution with 95/5% O2/CO2 (see Table of Materials).</li>
	<li>Maintain the tissue at 4 &#176;C by placing ice around the outside of the dish.</li>
</ol>
3. Dissection
<ol>
	<li>Use the dissection scope with a fiber optic light source to carry out the dissection.</li>
	<li>Remove the lower jaw. Place a blunt dissection probe through the mouth to provide easier handling of the head. Cut the joint connecting the dentary bone to the cranium with a scalpel. Use rongeurs to pull the lower jaw away from the cranium. Use rongeurs to pull off the skin and muscles from their attachments at the dorsal and lateral regions of the cranium.</li>
	<li>Remove the vertebral column.
	<ol>
		<li>Identify the vertebral column at the caudal end of the cranium. Bend the vertebral column ventrally to expose the spinal cord. Use microscissors to snip the spinal cord. Use rongeurs to remove the vertebral column and other tissue from the cranium by pulling caudally.</li>
	</ol>
	</li>
	<li>Remove the brain from the cranium after cutting the cranial nerves.
	<ol>
		<li>Starting at the foramen magnum, use rongeurs to cut two incisions on the dorsal cranium. Make cuts superficial to avoid damaging the brain beneath.</li>
		<li>Use rongeurs to carefully pull off the dorsal cranium. Use microscissors to remove the meninx to expose the rest of the brain. Remove enough meninx until the olfactory bulbs, in the anterior cranial cavity, can be identified (see Figure 1A). Continue to irrigate the brain with turtle Ringer&#39;s solution, as necessary.</li>
		<li>Use curved forceps to gently pull the cerebrum caudally and produce slight tension on the cranial nerves. Carefully cut away and remove the olfactory bulbs and cerebrum with curved forceps.</li>
		<li>Use microscissors to gently push the midbrain toward the midline to expose the cranial nerves; nIII, about 0.6 mm, can be seen in front of nIV, and the diameter of nIV will be slightly less than nIII. Cut nIII and nIV where they attach to the midbrain (see Figure 1B). Repeat this on the other side.</li>
		<li>Cut the left and right optic nerve (nII) with microscissors. Then tilt the brainstem to one side. Observe nVI emerging from the ventral surface near the junction of the pons and the medulla (see Figure 1C); the diameter of nVI is small and approximately 0.3 mm. Cut both the left and right nVI.</li>
		<li>Remove the remaining parts of the brainstem from the turtle with fine forceps and microscissors. Once the cranium is empty, examine the cranial cavity floor. Identify nIII, nIV, and nVI.</li>
	</ol>
	</li>
	<li>Remove the upper and lower eye-lids with fine forceps and microscissors.</li>
</ol>
4. Calibration of Eye Movements
<ol>
	<li>Use a rigid table (see Table of Materials) to support the placement of the gimbal and other instruments. Place the turtle head into the gimbal chuck so that the dorsal surface of the head is parallel to the horizon using a small bubble-level resting across the skull. Roughly position one of the eyes at the center of the gimbal&#39;s horizontal and vertical rotations.</li>
	<li>Place the infrared camera, fitted with an infrared light emitting diode (LED), which is part of the video-based eye tracking system (see Table of Materials), at a viewing distance of approximately 12 cm from the turtle&#39;s eye.
	<ol>
		<li>Angle the camera 45 degrees above the line of sight of the eye. The LED should be at the 11 o&#8217;clock position when looking at the camera lens. Center the LED along the optical axis of the eye. The camera will be slightly off axis (as viewed from above the eye).</li>
		<li>Adjust the distance of the camera from the eye so that the camera view is maximally filled by the eyeball. Ensure that the corners of the eyes (canthi) are at the edges of the horizontal view.</li>
	</ol>
	</li>
	<li>Connect the camera to the video-based eye tracking system to process the data. Split the signal to a DVD-recorder to capture the raw video. Focus the camera to obtain a clear image of the eye. Take care to fine-position the eye at the center of the camera view using the three degrees of linear adjustment (x, y, z) provided with the gimbal.</li>
	<li>Detect the dark pupil by setting the threshold and contrast appropriately using the program provided with the video-based eye tracking system.
	<ol>
		<li>Using the mouse, click on the &#34;Video&#34; menu and under &#34;Mode&#34; select &#34;High Precision&#34; to capture images at a sampling rate of 30 Hz (resolution of 640 pixels x 480 lines). Also under &#34;Video&#34;, select &#34;Dark Pupil&#34; for &#34;Pupil Type&#34; and &#34;Ellipse (rotated ellipse)&#34; for &#34;Pupil Segmentation Method&#34;.</li>
		<li>In the &#34;EyeCamera&#34; window, click on the &#34;Pupil Search Area Adjustment&#34; icon (small vertical rectangle with a dot in the center). Use the mouse to drag out a rectangle that limits an area around the pupil. Avoid dark areas that could be confused with the pupil.</li>
		<li>In the &#34;Controls&#34; window, confirm that boxes for &#34;Auto Image&#34; and &#34;Positive-Lock Threshold-Tracking&#34; are checked. Click on &#34;Auto-Threshold&#34; to optimize the density of scanning, which will show as green dots over the dark pupil.</li>
	</ol>
	</li>
	<li>Calibrate the video-display of the video-based eye tracking program to the rotations of the gimbal 12.5 &#176; (+/-) around its horizontal axis and 10 &#176; (+/-) around its vertical axis.
	<ol>
		<li>In the &#34;Controls&#34; window, click on &#34;Display&#34;. Check the boxes under both &#34;Gaze&#34; and &#34;Stim&#34; for &#34;Gaze Point&#34;, &#34;Calib Region&#34;, and &#34;Geometry Grid&#34;. After checking the boxes under &#34;Geometry Grid&#34;, a window will pop up and say, &#34;The stimulus display geometry must be measured before the geometry grid can be displayed. Do you wish to do this now?&#34; Select &#34;Y&#34; for yes.</li>
		<li>Using the mouse, click on the &#34;Windows&#34; menu and select &#34;Stimulus&#34;. The &#34;Stimulus&#34; window will open showing a vertical and horizontal line crossing at the center of the display. Measure the lengths of the lines to the nearest mm. Press &#34;Esc&#34; on the keyboard to exit the &#34;Stimulus&#34; window.</li>
		<li>Using the mouse, click on the &#34;Stimulus&#34; menu and select &#34;Geometry Setup&#34;. Input the lengths of the lines that were just measured. Adjust the viewing distance so that the degrees/line equal 25 &#176; for the horizontal line and 20 &#176; for the vertical line. Click on the &#34;Store&#34; button and close the window.</li>
		<li>In the &#34;EyeSpace&#34; window, select the number of calibration &#34;Data Point&#34; to be &#34;9&#34;. With the turtle&#39;s eye positioned at the center, click on the center data point and click the &#34;Re-present&#34; button. 
		NOTE: The &#34;Stimulus&#34; window will open, and &#34;Get Ready&#34; will appear at the center of the screen. A box will appear at the center location and then disappear. On its disappearance, the &#34;Stimulus&#34; window will close. The center position should now be calibrated.</li>
		<li>Repeat the procedure by rotating the gimbal right/left, +12.5 &#176;/-12.5 &#176;, and up/down +10 &#176;/-10 &#176; to calibrate the remaining data points.</li>
	</ol>
	</li>
	<li>To calibrate torsional rotation, use the template fitting algorithm provided with the video-based eye tracking program. The algorithm sets a zero position based on markings of the iris and computes an angle of rotation when the markings become offset from the centroid of the pupil.
	<ol>
		<li>Using the mouse pointer, click on the &#34;Windows&#34; menu and select &#34;Torsion&#34;. Click the &#34;START&#34; button in the &#34;Torsion&#34; window. In the &#34;EyeCamera&#34; window, an arc will appear over the image of the eye.</li>
		<li>Adjust the radius, the angle, and the length of the arc using the sliders at a location where irregular markings are present in the iris. Check the boxes for &#34;Real-time graphics&#34; and &#34;Auto-Set after adjust&#34;. If necessary, adjust the brightness and contrast in the Controls window and re-threshold the dark pupil (see step 4). Click the &#34;Set Template&#34; button.</li>
	</ol>
	</li>
	<li>Place a ruler in the same focal plane as the pupil and record the width of the full camera view. The value will be used later to determine the actual width of the pupil.</li>
</ol>
5. Positioning of Suction Electrode on Cranial Nerve to Evoke Eye Movements
<ol>
	<li>Carefully place a pin reference electrode into the connective or muscle tissue remaining on the head.</li>
	<li>Place the suction electrode (see Table of Materials) on the cranial nerve using a micromanipulator and dissection scope mounted on a boom. Use a fiber optic light source to view and guide the placement.
	<ol>
		<li>Match the size of a nerve to a capillary glass tip. Trial and error is necessary to obtain a best fit around the diameter of a nerve (see step 1.2 for size recommendations). Place the glass tip onto the suction electrode. Fill the suction electrode with Ringer&#39;s solution and adjust the volume within the syringe to about half of its capacity.</li>
		<li>Carefully move the glass tip of the electrode using the micromanipulator to a position above the cut-end of the selected nerve. Ensure that Ringer&#39;s solution fills the cranium and the tip is below its surface. If necessary, use modeling clay to dam up locations where the Ringer&#39;s solution is leaking out of the cranium.</li>
		<li>Pull back on the plunger of the syringe. 
		NOTE: The vacuum will draw the nerve into the end of the capillary tip. A good fit is indicated by the nerve remaining within the tip with little or no additional vacuum applied.</li>
	</ol>
	</li>
</ol>
6. Stimulation of Cranial Nerve and Analysis of Eye Movements
<ol>
	<li>Use a general purpose nerve/muscle stimulator with a current isolation device (see Table of Materials) to stimulate the cranial nerve via the suction electrode.
	<ol>
		<li>Connect the suction electrode to the current isolation device using a cable. Connect the lead from the pin reference electrode to the ground connection of the isolation device.</li>
		<li>Select parameters of the currents using the dials and switches on the stimulator and isolation device. Use a range of currents from 1 to 100 &#181;A, with frequency of 10 to 400 Hz. Use 1- or 2-ms pulses in trains lasting 100, 500, or 1,000 ms.</li>
	</ol>
	</li>
	<li>Record the timing of stimulations. 
	NOTE: Transistor-transistor-logic (TTL) pulses are synchronized with the deliveries of the currents from the stimulator and communicate in real-time via a cable to input channels of the video-based eye tracking system. A software module provided with the video-based eye tracking program controls the communication.
	<ol>
		<li>To visualize the timing of the current applications and their influence on eye movements, click on the &#34;PenPlots&#34; menu. Select &#34;X Gaze Position&#34;, &#34;Y Gaze Position&#34;, &#34;Torsion&#34;, and &#34;Pupil Width&#34; to show real time raw data plots for X and Y eye positions, torsion, and pupil width. Also select the &#34;Seconds &#38; Markers&#34; from the &#34;PenPlots&#34; menu to show a timing plot with tick marks, which appear at 1 s intervals. 
		NOTE: A capital letter &#34;T&#34; will appear marking the onset of the TTL pulse, occurring simultaneously with the current application.</li>
		<li>To store the data of eye movements evoked by currents, click on the &#34;File&#34; menu and select &#34;New Data File&#34; under &#34;Data&#34;. Input a file name and press &#34;Enter&#34;. Saving data can be paused and then restarted using the combination of key commands, &#34;Ctrl&#34; + &#34;p&#34;. When an experimental session is completed, select &#34;Close Data File&#34; off &#34;File&#34; menu under &#34;Data&#34;.</li>
		<li>To help keep track of the type of currents applied, click on the &#34;Windows&#34; menu and select &#34;Data Pad&#34;. The &#34;KeyPad/DataMarker&#34; window will appear. Click on a letter or a number to identify parameters of the current stimulations being delivered to the nerve. 
		NOTE: For example, &#34;X&#34; could stand for 10 &#181;A. Clicking on &#34;X&#34; stores its entry into the data file in real time for post hoc analysis. It also appears on the &#34;PenPlot&#34; for &#34;Seconds &#38; Markers&#34; for ongoing observation.</li>
	</ol>
	</li>
	<li>Analyze data from the eye tracker system.
	<ol>
		<li>Open the saved data file, which is in a text delimited format, into a spread sheet program of choice to organize the data and carry out statistical analysis.
		<ol>
			<li>Convert the pupil width values stored in the file to real sizes in mm.</li>
			<li>Convert the values of X and Y eye positions and torsions to units of degree and use conventions for describing eye movements; therefore, positive directions of rotations: intorsion, elevation, and adduction; and negative directions of rotations: extorsion, depression, and abduction.</li>
		</ol>
		</li>
		<li>Copy the header information onto a new worksheet. This will include values for the screen size (width and height) and the viewing distance. Eight columns of data collected at a rate of 30 Hz will follow below the header information.</li>
		<li>Go back to the worksheet containing the raw data. Do a &#34;Find&#34; for &#34;X&#34; in the last column titled &#34;Marker&#34; to locate where stimulation was applied with 10 &#181;A. Find incidence of &#34;T&#34;, marking the beginning of the current stimulation. Copy 15 rows (0.5 s) of data occurring before &#34;T&#34; and 90 frames after &#34;T&#34; (3.0 s); i.e., 3.5 s total. Paste the data in the new worksheet below the header information.</li>
		<li>Insert a blank column after &#34;PupilWidth&#34;. In the blank column, convert to the calibrated dimension in mm: 
		Horizontal pupil diameter = &#34;PupilWidth&#34; &#215; the dimension of camera view width</li>
		<li>Insert 2 blank columns after both &#34;X_Gaze&#34; and &#34;Y_Gaze&#34;. Normalize positions to the dimensions of the viewing screen: coordinates (0, 0) at the top left of the screen extending to (1, 1) at the bottom right. In the first blank column, translate positions to a coordinate system having (0, 0) at the center of the screen. Include the conversion to the dimensions of the screen in mm: 
		X = (0.5 &#215; width)&#160; &#8211; (&#34;X_Gaze&#34; &#215; width); Y = (0.5 &#215; height) &#8211; (&#34;Y_Gaze&#34; &#215; height) 
		NOTE: The sequence of operation is for the left eye. The sequence will need to be reversed for the right eye in order to follow convention of negativity for abduction and positivity for adduction.</li>
		<li>In the second columns, use trigonometry functions to convert to angles (&#176;) of rotation: 
		horizontal rotation = arctan (X/viewing distance);vertical rotation = arctan (Y/viewing distance)</li>
		<li>Torsion is already shown in units of degrees, but to conform to convention of positivity for intorsion, if measuring is done on the left eye, multiply by -1. For the right eye, no multiplication is necessary. The program codes clockwise rotation as positive.</li>
		<li>Plot the pupil diameter and rotations as a function of time.</li>
	</ol>
	</li>
</ol>

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D., Ariel, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+"beat-to-beat"+interval+generator+for+optokinetic+nystagmus.">A "beat-to-beat" interval generator for optokinetic nystagmus.</a> Biol Cybern. 66, 203-216 (1992).</li><li>Keifer, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+eye-blink+reflex+model:+Role+of+excitatory+amino+acid+receptors+and+labeling+of+network+activity+with+sulforhodamine.">In vitro eye-blink reflex model: Role of excitatory amino acid receptors and labeling of network activity with sulforhodamine.</a> Exp Brain Res. 97, 239-253 (1993).</li><li>Keifer, J., Armstrong, K. E., Houk, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+classical+conditioning+of+abducens+nerve+discharge+in+turtles.">In vitro classical conditioning of abducens nerve discharge in turtles.</a> J Neurosci. 15, 5036-5048 (1995).</li><li>Rosenberg, A. F., Ariel, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+model+for+optokinetic+eye+movements+in+turtles+that+incorporates+properties+of+retinal+slip+neurons.">A model for optokinetic eye movements in turtles that incorporates properties of retinal slip neurons.</a> Vis Neurosci. 13, 375-383 (1996).</li><li>Ariel, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Open-loop+optokinetic+responses+of+the+turtle.">Open-loop optokinetic responses of the turtle.</a> Vis Res. 37, 925-933 (1997).</li><li>Anderson, C. W., Keifer, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Properties+of+conditioned+abducens+nerve+responses+in+a+highly+reduced+in+vitro+brainstem+preparation+from+the+turtle.">Properties of conditioned abducens nerve responses in a highly reduced in vitro brainstem preparation from the turtle.</a> J Neurophysiol. 81, 1242-1250 (1999).</li><li>Keifer, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+classical+conditioning+of+the+turtle+eyeblink+reflex:+Approaching+cellular+mechanisms+of+acquisition.">In vitro classical conditioning of the turtle eyeblink reflex: Approaching cellular mechanisms of acquisition.</a> Cerebell. 2, 55-61 (2003).</li><li>Zhu, D., Keifer, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pathways+controlling+trigeminal+and+auditory+nerve-evoked+abducens+eyeblink+reflexes+in+pond+turtles.">Pathways controlling trigeminal and auditory nerve-evoked abducens eyeblink reflexes in pond turtles.</a> Brain Behav Evol. 64, 207-222 (2004).</li><li>Jones, M. S., Ariel, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effects+of+unilateral+eighth+nerve+block+on+fictive+VOR+in+the+turtle.">The effects of unilateral eighth nerve block on fictive VOR in the turtle.</a> Br Res. 1094, 149-162 (2006).</li><li>Jones, M. S., Ariel, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphology,+intrinsic+membrane+properties,+and+rotation-evoked+responses+of+trochlear+motoneurons+in+the+turtle.">Morphology, intrinsic membrane properties, and rotation-evoked responses of trochlear motoneurons in the turtle.</a> J Neurophysiol. 99 (3), 1187-1200 (2008).</li><li>Krenz, J. G., Naylor, G. J. P., Shaffer, H. B., Janzen, F. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+phylogenetics+and+evolution+of+turtles.">Molecular phylogenetics and evolution of turtles.</a> Mol Phylogenet Evol. 37 (1), 178-191 (2005).</li><li>Dearworth, J. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+the+trochlear+nerve+in+eye+abduction+and+frontal+vision+of+the+red-eared+slider+turtle+(Trachemys+scripta+elegans).">Role of the trochlear nerve in eye abduction and frontal vision of the red-eared slider turtle (Trachemys scripta elegans).</a> J Comp Neur. 52, 3464-3477 (2013).</li><li>Dearworth, J. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pupil+constriction+evoked+in+vitro+by+stimulation+of+the+oculomotor+nerve+in+the+turtle+(Trachemys+scripta+elegans).">Pupil constriction evoked in vitro by stimulation of the oculomotor nerve in the turtle (Trachemys scripta elegans).</a> Vis Neurosci. 26, 309-318 (2009).</li><li>Mead, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=IFEL+TOUR:+a+description+of+the+introduction+to+FUN+electrophysiology+labs+workshop+at+Bowdoin+College,+July+27-30,+and+the+resultant+faculty+learning+community.">IFEL TOUR: a description of the introduction to FUN electrophysiology labs workshop at Bowdoin College, July 27-30, and the resultant faculty learning community.</a> J Undergrad Neurosci Educ. 5, A42-A48 (2007).</li><li>Jackson, D. C., Ultsch, G. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physiology+of+hibernation+under+the+ice+by+turtles+and+frogs.">Physiology of hibernation under the ice by turtles and frogs.</a> J Exp Zool A Ecol Genet Physiol. 313 (6), 311-327 (2010).</li><li>Romano, J. M., Dearworth, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pupil+constriction+evoked+by+stimulation+of+the+ciliary+nerve+in+the+red-eared+slider+turtle+(Trachemys+scripta+elegans).">Pupil constriction evoked by stimulation of the ciliary nerve in the red-eared slider turtle (Trachemys scripta elegans).</a> J Penns Acad Sci. 85, 4-8 (2011).</li><li>Miller, J. M., Robins, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extraocular-muscle+forces+in+alert+monkey.">Extraocular-muscle forces in alert monkey.</a> Vis Res. 32, 1099-1113 (1992).</li><li>Gamlin, P. D., Miller, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extraocular+muscle+motor+units+characterized+by+spike-triggered+averaging+in+alert+monkey.">Extraocular muscle motor units characterized by spike-triggered averaging in alert monkey.</a> J Neurosci Meth. 204, 159-167 (2011).</li><li>Quaia, C., Ying, H. S., Optican, L. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Viscoelastic+properties+of+passive+eye+muscle+in+primates.+III:+Force+elicited+by+natural+elongations.">The Viscoelastic properties of passive eye muscle in primates. III: Force elicited by natural elongations.</a> PLOS ONE. 5, A236-A254 (2010).</li><li>Anderson, S. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamics+of+primate+oculomotor+plant+revealed+by+effects+of+abducens+microstimulation.">Dynamics of primate oculomotor plant revealed by effects of abducens microstimulation.</a> J Neurophys. 101, 2907-2923 (2009).</li><li>Maxwell, J. H., Harless, M., Morlock, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anesthesia+and+surgery.">Anesthesia and surgery.</a> Turtles: Perspective and Research. , 127-152 (1979).</li><li>AVMA Panel on Euthanasia. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=American+Veterinary+Medical+Association.">American Veterinary Medical Association.</a> J Am Vet Med Assoc. 218 (5), 669-696 (2001).</li><li>Clarke, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shaping+the+pupil's+response+to+light+in+the+hooded+rat.">Shaping the pupil's response to light in the hooded rat.</a> Exp Br Res. 176, 641-651 (2007).</li><li>Bennett, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+review+of+anesthesia+and+chemical+restraint+in+reptiles.">A review of anesthesia and chemical restraint in reptiles.</a> J Zoo Wild Med. 22 (3), 282-303 (1991).</li><li>Bickler, P. E., Buck, L. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hypoxia+Tolerance+in+Reptiles,+Amphibians,+and+Fishes:+Life+with+Variable+Oxygen+Availability.">Hypoxia Tolerance in Reptiles, Amphibians, and Fishes: Life with Variable Oxygen Availability.</a> Ann Rev Physiol. 69, 145-170 (2007).</li></ol>

The authors have nothing to disclose.

Critical Steps:
The critical steps within this protocol are the following: 1) the dissection and the care taken to maintain the viability of the transected nerves; 2) the matching of sizes by the suction electrodes to cranial nerves to provide consistent responses; and 3) the placement of the head in the gimbal to provide adequate calibration of the rotations of the eye.
Troubleshooting:
The dissection can be challenging, but after completing i...

Rationale for Using Red-eared Slider Turtles in Electrophysiological Experiments:
Red-eared slider turtles (Trachemys scripta elegans), are considered one of world&#39;s worst invasive species1 and can indicate that an ecosystem is in trouble. The reason why red-eared slider turtles are so successful is poorly understood but it may in part be due to their tolerant physiology and possession of nervous tissues that can survive under hypoxic conditions2,3,4. Using them for experimentation does not threaten their numbers and with minimal efforts, electrophysiological preparations can remain viable over extended durations, as long as 18 hours5,6. The benefit is similar to the advantage of using invertebrate animals such as crayfish7, which also have the ability to withstand low levels of oxygen8.
Techniques for Measuring Eye Movements:
Approaches to measure eye movements in frontal-eyed animals using non-human primates have been well developed9. The eye rotates in the orbit around three axes: horizontal, vertical, and torsional. The magnetic search coil method is generally considered the most reliable for measuring rotations, but is invasive, requiring small coils to be inserted into the scleras of animals10,11. Video-based systems also can measure rotations and have the advantage of being non-invasive. The development of better cameras along with innovative image processing have enhanced their functionality making video-based systems an attractive alternative to consider12,13,14.
The techniques developed for measuring eye movements in nonmammals have been much less significant. Measures are either low resolution or describe only some of the rotations15,16,17,18. The lack of development can be partially blamed on the difficulty in training nonmammals to follow visual targets. Although eye movements have been well studied in red-eared slider turtles19,20,21,22,23,24,25,26,27,28,29,30, because of the challenge in training animals to track targets, the precise kinematics of their eye movements is poorly understood.
Red-eared slider turtles are generally considered lateral-eyed vertebrates, but because they can fully retract their heads into their shell31, significant occlusion of the lateral visual fields by the carapace occurs32. The result is that their visual line of sight is forced toward the front, making them behave more like frontal-eyed mammals. Therefore, their use as a model for developing approaches for measuring eye movements also offers a unique evolutionary perspective.
The protocol described in this work uses an in vitro isolated head preparation to identify the kinematics of the eye movements in red-eared slider turtles. Brains are dissected from the skulls leaving the cranial nerves intact. Heads are placed into a gimbal to calibrate eye movements and evoke responses by electrical stimulation of the cranial nerves innervating the eye muscles. Measures of rotations by the eyes are done by a video-based system, using software algorithms, which track the dark pupil and the markings of the iris. The preparation provides the opportunity to measure kinematics of both extraocular (i.e., horizontal, vertical, and torsional rotations)32 and intraocular (i.e., pupil changes)33 movements.
Model System for Analysis of Efferent Neural Pathways:
More generally, the approach provides investigators the chance to study how efferent neural signals generate eye movements when muscles start from their relaxed states and in the absence of integrated sensory information processed by the brain32,33. Therefore, the eye kinematics can be examined in a model system in which they are solely processed by the efferent neural pathway leaving the brain and synapsing onto the muscles.

<table>
	<tbody>
		<tr>
			<td>Red-eared slider turtles</td>
			<td>Kons Scientific</td>
			<td>Trachemys scripta elegans</td>
			<td>Large size (carapace length 15-20 cm)</td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sigma-Aldrich Co. LLC.</td>
			<td>S5886</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Sigma-Aldrich Co. LLC.</td>
			<td>P5405</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium choride</td>
			<td>Sigma-Aldrich Co. LLC.</td>
			<td>M7304</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Sigma-Aldrich Co. LLC.</td>
			<td>S5761</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextrose</td>
			<td>Sigma-Aldrich Co. LLC.</td>
			<td>C5767</td>
			<td></td>
		</tr>
		<tr>
			<td>Concentrated hydrochloric acid</td>
			<td>Sigma-Aldrich Co. LLC.</td>
			<td>H7020</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride</td>
			<td>Sigma-Aldrich Co. LLC.</td>
			<td>C7902</td>
			<td></td>
		</tr>
		<tr>
			<td>pH meter</td>
			<td>Oakton</td>
			<td>pH 6+</td>
			<td></td>
		</tr>
		<tr>
			<td>Suction stimulation electrode</td>
			<td>A-M Systems</td>
			<td>573000</td>
			<td>Bipolar suction electrode. Note that 573000 has been replaced with 573050.</td>
		</tr>
		<tr>
			<td>Capillary glass</td>
			<td>A-M systems</td>
			<td>626000</td>
			<td>Single-barrel borosilicate capillary glass without microfilament, length 10 cm, outside diameter 1.0 mm, inner diameter 0.50 mm</td>
		</tr>
		<tr>
			<td>Alternative suction stimulation electrode</td>
			<td>A-M Systems</td>
			<td>573050</td>
			<td>Bipolar suction electrode. Requires larger diameter capillary glass: 627000, outside diameter 1.2 mm, inner diameter 0.68 mm</td>
		</tr>
		<tr>
			<td>Stereoscope</td>
			<td>Lieca</td>
			<td>GZ7</td>
			<td>Magnification range, 10x &#8211; 70x</td>
		</tr>
		<tr>
			<td>Fiber optic light source</td>
			<td>Amscope</td>
			<td>HL250-A</td>
			<td>150W Fiber optical microscope illuminator light box</td>
		</tr>
		<tr>
			<td>Rongeurs</td>
			<td>Carolina Biological Supply Company</td>
			<td>625654</td>
			<td>stainless steel, straight spring, 5.25&#34;</td>
		</tr>
		<tr>
			<td>Blunt dissection probe</td>
			<td>Carolina Biological Supply Company</td>
			<td>627405</td>
			<td>Huber mall probe, double-ended probe and seeker, 6&#34;</td>
		</tr>
		<tr>
			<td>Microscissors</td>
			<td>Carolina Biological Supply Company</td>
			<td>623555</td>
			<td>Iris microdissecting scissors, stainless steel, 0.5&#34; blades, 4.75&#34; long</td>
		</tr>
		<tr>
			<td>Fine forceps</td>
			<td>Sigma-Aldrich Co. LLC.</td>
			<td>F6521</td>
			<td>Jewelers forceps, dumont No. 5, inox alloy, 4.25&#34;</td>
		</tr>
		<tr>
			<td>Curved forceps</td>
			<td>Sigma-Aldrich Co. LLC.</td>
			<td>Z168696</td>
			<td>Medium tip, curved forceps, stainless steel, 4&#34;</td>
		</tr>
		<tr>
			<td>Scalpel handle</td>
			<td>Sigma-Aldrich Co. LLC.</td>
			<td>S2896</td>
			<td>Scalpel handles, No. 3, stainless steel</td>
		</tr>
		<tr>
			<td>Scalpel blade</td>
			<td>Sigma-Aldrich Co. LLC.</td>
			<td>S2771</td>
			<td>Scalpel blades, No. 11, steel</td>
		</tr>
		<tr>
			<td>Guillotine</td>
			<td>Harvard Apparatus</td>
			<td>73-1918</td>
			<td>Kleine guillotine type 7575</td>
		</tr>
		<tr>
			<td>Spatula</td>
			<td>Sigma</td>
			<td>Z648299</td>
			<td>Micro spoon and spatula weighing set. Use small spatula: 5.9&#8221; long x 0.07&#8221; diameter handle with square end: 0.17&#8221; x 1.3&#8221; long, other end round: 0.17&#8221; x 1.27&#8221; long</td>
		</tr>
		<tr>
			<td>Hook</td>
			<td>Autozone</td>
			<td>98069</td>
			<td>SureBilt hook and pick set. Use grinder to dull sharp points of hook to prevent injury to animals mouth.</td>
		</tr>
		<tr>
			<td>95/5% O2/CO2</td>
			<td>Airgas, Inc.</td>
			<td>X02OX95C2003102</td>
			<td>5% Carbon dioxide balance oxygen certified standard gas mixture, size 200 Cylinder, CGA-296</td>
		</tr>
		<tr>
			<td>Regulator</td>
			<td>Airgas, Inc.</td>
			<td>Y11244D296-AG</td>
			<td>Single stage brass 0-100 psi analytical cylinder regulator CGA-296 with needle outlet. Use brass adjustable airline pipe valve to go from 3/8&#34;, inner diameter, vinyl airline tubing connected to regulator to a 3/16&#34;, inner diameter, airline connection going to airstone or glass pasteur pipette.</td>
		</tr>
		<tr>
			<td>Adjustable airline pipe valve</td>
			<td>Doctors Foster and Smith</td>
			<td>CD-12061</td>
			<td>Brass valve</td>
		</tr>
		<tr>
			<td>Rigid table</td>
			<td>Unknown</td>
			<td>Unknown</td>
			<td>Auto-clave door laid on top of a sturdy table. Nine 5&#34; diameter tennis balls isolate vibrations from the top surface of the table.</td>
		</tr>
		<tr>
			<td>5&#34; tennis ball</td>
			<td>Petco Animal Supplies, Inc.</td>
			<td>712868</td>
			<td>Petco Jumbo Pet Tennis Ball: balls are unsliced and held within an integrated frame on the underside part of the autoclave door.</td>
		</tr>
		<tr>
			<td>Alternative vibration isolation table</td>
			<td>Newport Corporation</td>
			<td>INT1-36-6-N</td>
			<td>Rigid vibration control system, integrity 1: Surface dimensions, 3&#39; x 6&#39;</td>
		</tr>
		<tr>
			<td>Gimbal</td>
			<td>ISI, International Scientific Instruments, Inc.</td>
			<td>Stage from SUPER III-A Scanning EM</td>
			<td>5-axis eucentric stage: X, Y, and Z linear movements, &#177;20 mm, 0.1 mm precision; Rotations, vertical, &#177;10&#176;, and horizontal, &#177;12.5&#176;, with 1.25&#176; precision. Note: from decommission instrument.</td>
		</tr>
		<tr>
			<td>Chuck for gimbal</td>
			<td>Unknown</td>
			<td>Unknown</td>
			<td>Chuck from an old microtome of unknown manufacture was machined to fit the shaft of the specimen holder of the Scanning EM stage</td>
		</tr>
		<tr>
			<td>Alternative gimbal</td>
			<td>ThorLabs, Inc.</td>
			<td>GN2/M with MBT602/M</td>
			<td>Dual-axis goniometer (GN2/M) mounted on 3-axis microblock stage with thumbscrew adjusters (MBT602/M): design a chuck to hold turtle head with eye at 12.7 mm above top surface of goniometer (distance to point of rotation)</td>
		</tr>
		<tr>
			<td>Video-based eye tracking system</td>
			<td>Arrington Research, Inc.</td>
			<td>ViewPoint EyeTracker, PC-60</td>
			<td>Tracking method: Infrared video by dark pupil; Black and white camera (Item BC02): 30 Hz, 640 x 480; System requirements: Windows 2000, XP, 7, 8, 8.1, 10; Visual range: Horizontal +/- 44&#176;; vertical +/- 20&#176;; Accuracy ~0.5&#176;; Spatial resolution ~0.15&#176;; Pupil size resolution ~0.03 mm; Eye data: X, Y position of gaze, pupil height and width, torsion, delta time, total time, and regions of interest (ROI); Real-time communication (Item 0022): 4-Channel AnalogOut with eight TTL input channels to mark codes into the data file</td>
		</tr>
		<tr>
			<td>Multi-position magnetic base</td>
			<td>Harbor Freight Tools</td>
			<td>Pittsburg, item #5645</td>
			<td>Magnetic holder reaches up to 12&#34; and produces 45 lbs. of magnetic pull. Use to position camera. Machine thread holes onto the end of the rod to mount cameras.</td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>Kopf</td>
			<td>900</td>
			<td>5 axis manipulation for mount of suction electrode: X, Y, Z linear travel, 2 axis of rotation</td>
		</tr>
		<tr>
			<td>Dissection scope on boom</td>
			<td>Lieca</td>
			<td>GZ6</td>
			<td>Magnification range, 6.7x &#8211; 40x</td>
		</tr>
		<tr>
			<td>Nerve/muscle stimulator</td>
			<td>Astro-Med Grass Telefactor</td>
			<td>Grass S88</td>
			<td>Dual pulse voltage stimulator: two output channels that can be operated independently or synchronized to generate non-isolated constant voltage pulses (10 mv to 150 V). Pulses can be single (10 &#956;sec to 10 sec), repetitive (0.01 Hz to 1 KHz), and trains (1 ms to 10 s) and synchronized with TTL inputs and output. Send TTL outputs via the output channels of a DB25 connector to the TTL input channels of the ViewPoint EyeTracker. Note: Astro-Med Grass Telefactor is no longer in business.</td>
		</tr>
		<tr>
			<td>Current isolation device</td>
			<td>Astro-Med Grass Telefactor</td>
			<td>PSIU6</td>
			<td>Current stimulus isolation unit: enables safe delivery of constant currents by the S88 to the preparation. The PSIU6 connects by a BNC cable to one of the output channels of the S88. Multiplier switches on the PSIU6 allow the S88 to generate a wide array of current amplitudes ranging from 0.1 &#181;A to 15 mA.</td>
		</tr>
		<tr>
			<td>Alternative nerve/muscle stimulator with isolation</td>
			<td>A-M Systems</td>
			<td>2100</td>
			<td>Isolated Pulse Stimulator: Unit has built-in isolator to produce constant currents.</td>
		</tr>
	</tbody>
</table>

ocular kinematics measured by in vitro stimulation of the cranial nerves in the turtle

After animals are euthanized, their tissues begin to die. Turtles offer an advantage because of a longer survival time of their tissues, especially when compared to warm-blooded vertebrates. Because of this, in vitro experiments in turtles can be performed for extended periods of time to investigate the neural signals and control of their target actions. Using an isolated head preparation, we measured the kinematics of eye movements in turtles, and their modulation by electrical signals carried by cranial nerves. After the brain was removed from the skull, leaving the cranial nerves intact, the dissected head was placed in a gimbal to calibrate eye movements. Glass electrodes were attached to cranial nerves (oculomotor, trochlear, and abducens) and stimulated with currents to evoke eye movements. We monitored eye movements with an infrared video tracking system and quantified rotations of the eyes. Current pulses with a range of amplitudes, frequencies, and train durations were used to observe effects on responses. Because the preparation is separated from the brain, the efferent pathway going to muscle targets can be examined in isolation to investigate neural signaling in the absence of centrally processed sensory information.

Figure 1 shows stills of images taken from a video describing the dissection. Images provide typical locations of the nerves prior to cutting from the brain.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56864/56864fig1.jpg" /> 
Figure 1: Stills of images captured from the video of the dissection to show locations of the optic nerve (nII), oculomotor nerve (nIII), trochlear...

Watch this Scientific Journal Video about Ocular Kinematics Measured by In Vitro Stimulation of the Cranial Nerves in the Turtle at JoVE.com

Ocular Kinematics Measured by In Vitro Stimulation of the Cranial Nerves in the Turtle

This protocol describes how to use an in vitro isolated turtle head preparation to measure the kinematics of their eye movements. After removal of the brain from the cranium, cranial nerves can be stimulated with currents to quantify rotations of the eye and changes in pupil sizes.

Ocular Kinematics Measured by In Vitro Stimulation of the Cranial Nerves in the Turtle

After animals are euthanized, their tissues begin to die. Turtles offer an advantage because of a longer survival time of their tissues, especially when ...

ocular-kinematics-measured-vitro-stimulation-cranial-nerves

Research

JoVE Journal

Neuroscience

8.4K Views.  Lafayette College. This protocol describes how to use an in vitro isolated turtle head preparation to measure the kinematics of their eye movements. After removal of the brain from the cranium, cranial nerves can be stimulated with currents to quantify rotations of the eye and changes in pupil sizes.

Video: Ocular Kinematics Measured by In Vitro Stimulation of the Cranial Nerves in the Turtle

TMS: Using the Theta-Burst Protocol to Explore Mechanism of Plasticity in Individuals with Fragile X Syndrome and Autism

Fragile X Syndrome (FXS), also known as Martin-Bell Syndrome, is a genetic abnormality found on the X chromosome.1,2 Individuals suffering from FXS display abnormalities in the expression of FMR1 - a protein required for typical, healthy neural development.3 Recent data has suggested that the loss of this protein can cause the cortex to be hyperexcitable thereby affecting overall patterns of neural plasticity.4,5
In addition, Fragile X shows a strong comorbidity with autism: in fact, 30% of children with FXS are diagnosed with autism, and 2 - 5% of autistic children suffer from FXS.6
Transcranial Magnetic Stimulation (a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate cortical excitability via the application of localized magnetic field pulses 7,8) represents a unique method of exploring plasticity and the manifestations of FXS within affected individuals. More specifically, Theta-Burst Stimulation (TBS), a specific stimulatory protocol shown to modulate cortical plasticity for a duration up to 30 minutes after stimulation cessation in healthy populations, has already proven an efficacious tool in the exploration of abnormal plasticity.9,10 
Recent studies have shown the effects of TBS last considerably longer in individuals on the autistic spectrum - up to 90 minutes.11 This extended effect-duration suggests an underlying abnormality in the brain's natural plasticity state in autistic individuals - similar to the hyperexcitability induced by Fragile X Syndrome.
In this experiment, utilizing single-pulse motor-evoked potentials (MEPs) as our benchmark, we will explore the effects of both intermittent and continuous TBS on cortical plasticity in individuals suffering from FXS and individuals on the Autistic Spectrum.

In this article, we examine the effects of Theta-Burst TMS stimulation on cortical plasticity in individuals suffering from Fragile X syndrome and individuals on the autistic spectrum.

Fragile X Syndrome (FXS), also known as Martin-Bell Syndrome, is a genetic abnormality found on the X chromosome.1,2 Individuals suffering from FXS ...

The NeuroStar TMS Device: Conducting the FDA Approved Protocol for Treatment of Depression

The Neuronetics NeuroStar Transcranial Magnetic Stimulation (TMS) System is a class II medical device that produces brief duration, pulsed magnetic fields. These rapidly alternating fields induce electrical currents within localized, targeted regions of the cortex which are associated with various physiological and functional brain changes.1,2,3
In 2007, O'Reardon et al., utilizing the NeuroStar device, published the results of an industry-sponsored, multisite, randomized, sham-stimulation controlled clinical trial in which 301 patients with major depression, who had previously failed to respond to at least one adequate antidepressant treatment trial, underwent either active or sham TMS over the left dorsolateral prefrontal cortex (DLPFC). The patients, who were medication-free at the time of the study, received TMS five times per week over 4-6 weeks.4
The results demonstrated that a sub-population of patients (those who were relatively less resistant to medication, having failed not more than two good pharmacologic trials) showed a statistically significant improvement on the Montgomery-Asberg Depression Scale (MADRS), the Hamilton Depression Rating Scale (HAMD), and various other outcome measures. In October 2008, supported by these and other similar results5,6,7, Neuronetics obtained the first and only Food and Drug Administration (FDA) approval for the clinical treatment of a specific form of medication-refractory depression using a TMS Therapy device (FDA approval K061053).
In this paper, we will explore the specified FDA approved NeuroStar depression treatment protocol (to be administered only under prescription and by a licensed medical profession in either an in- or outpatient setting).

In this article, we examine the methodology and considerations relevant to the FDA approved depression treatment protocol using the Neuronetics NeuroStar TMS device.

The Neuronetics NeuroStar Transcranial Magnetic Stimulation (TMS) System is a class II medical device that produces brief duration, pulsed magnetic ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Generation of Local CA1 &#947; Oscillations by Tetanic Stimulation

Neuronal network oscillations are important features of brain activity in health and disease and can be modulated by a range of clinically used drugs. A protocol is provided to generate a model for studying CA1 &#947; oscillations (20 - 80 Hz). These &#947; oscillations are stable for at least 30 min&#160;and depend upon excitatory and inhibitory synaptic activity in addition to activation of pacemaker currents. Tetanically stimulated oscillations have a number of reproducible and easily quantifiable characteristics including spike count, oscillation duration, latency and frequency that report upon the network state. The advantages of the electrically stimulated oscillations include stability, reproducibility and episodic acquisition enabling robust characterization of network function. This model of CA1 &#947; oscillations can be used to study cellular mechanisms and to systematically investigate how neuronal network activity is altered in disease and by drugs. Disease state pharmacology can be readily incorporated by the use of brain slices from genetically modified or interventional animal models to enable selection of drugs that specifically target disease mechanisms.

Oscillations are fundamental network properties and are modulated by disease and drugs. Studying brain-slice oscillations allows characterization of isolated networks under controlled conditions. Protocols are provided for the preparation of acute brain slices for evoking CA1 &#947; oscillations.

Neuronal network oscillations are important features of brain activity in health and disease and can be modulated by a range of clinically used drugs. A ...

Caspase-3 Activity in the Rat Amygdala Measured by Spectrofluorometry After Myocardial Infarction

Myocardial infarction (MI) has dramatic mid- and long-term consequences at the physiological and behavioral levels, but the mechanisms involved are still unclear. Our laboratory has developed a rat model of post-MI syndrome that displays impaired cardiac functions, neuronal loss in the limbic system, cognitive deficits and behavioral signs of depression. At the neuronal level, caspase-3 activation mediates post-MI apoptosis in different limbic regions, such as the amygdala &#8211; peaking at 3 days post-MI. Cognitive and behavioral impairments appear 2-3 weeks post-MI and these correlate statistically with measures of caspase-3 activity. The protocol described here is used to induce MI, collect amygdala tissue and measure caspase-3 activity using spectrofluorometry. To induce MI, the descending coronary artery is occluded for 40 min. The protocol for evaluation of caspase-3 activation starts 3 days after MI: the rats are sacrificed and the amygdala isolated rapidly from the brain. Samples are quickly frozen in liquid nitrogen and kept at -80 &#176;C until actual analysis. The technique performed to assess caspase-3 activation is based on cleavage of a substrate (DEVD-AMC) by caspase-3, which releases a fluorogenic compound that can be measured by spectrofluorometry. The methodology is quantitative and reproducible but the equipment required is expensive and the procedure for quantifying the samples is time-consuming. This technique can be applied to other tissues, such as the heart and kidneys. DEVD-AMC can be replaced by other substrates to measure the activity of other caspases.

Myocardial infarction (MI) is not only followed by impaired cardiac function but also by apoptosis in the amygdala, a brain region involved in the behavioral consequences of MI. This protocol describes how to induce MI, collect amygdala tissue and measure caspase-3 activity, a marker of apoptosis, therein.

Myocardial infarction (MI) has dramatic mid- and long-term consequences at the physiological and behavioral levels, but the mechanisms involved are still ...

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

The Power of Interstimulus Interval for the Assessment of Temporal Processing in Rodents

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in neurocognitive disorders. Despite the popularization of prepulse inhibition (PPI) in recent years, many current protocols promote using a percent of control measure, thereby precluding the assessment of temporal processing. The present study used cross-modal PPI and gap prepulse inhibition (gap-PPI) to demonstrate the benefits of employing a range of interstimulus intervals (ISIs) to delineate effects of sensory modality, psychostimulant exposure, and age. Assessment of sensory modality, psychostimulant exposure, and age reveals the utility of an approach varying the interstimulus interval (ISI) to establish the shape of the ISI function, including increases (sharper curve inflections) or decreases (flattening of the response amplitude curve) in startle amplitude. Additionally, shifts in peak response inhibition, suggestive of a differential sensitivity to the manipulation of ISI, are often revealed. Thus, the systematic manipulation of ISI affords a critical opportunity to evaluate temporal processing, which may reveal the underlying neural mechanisms involved in neurocognitive disorders.

Temporal processing, a preattentive process, may underlie deficits in higher-level cognitive processes, including attention, commonly observed in neurocognitive disorders. Using prepulse inhibition as an exemplar paradigm, we present a protocol for manipulating interstimulus interval (ISI) to establish the shape of the ISI function to provide an assessment of temporal processing.

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in ...

Assessing Pupil-linked Changes in Locus Coeruleus-mediated Arousal Elicited by Trigeminal Stimulation

Current scientific literature provides evidence that trigeminal sensorimotor activity associated with chewing may affect arousal, attention, and cognitive performance. These effects may be due to widespread connections of the trigeminal system to the ascending reticular activating system (ARAS), to which noradrenergic neurons of the locus coeruleus (LC) belongs. LC neurons contain projections to the whole brain, and it is known that their discharge co-varies with pupil size. LC activation is necessary for eliciting task-related mydriasis. If chewing effects on cognitive performance are mediated by the LC, it is reasonable to expect that changes in cognitive performance are correlated to changes in task-related mydriasis. Two novel protocols are presented here to verify this hypothesis and document that chewing effects are not attributable to aspecific motor activation. In both protocols, performance and pupil size changes observed during specific tasks are recorded before, soon after, and half an hour following a 2 min period of either: a) no activity, b) rhythmic, bilateral handgrip, c) bilateral chewing of soft pellet, and d) bilateral chewing of hard pellet. The first protocol measures level of performance in spotting target numbers displayed within numeric matrices. Since pupil size recordings are recorded by an appropriate pupillometer that impedes vision to ensure constant illumination levels, task-related mydriasis is evaluated during a haptic task. Results from this protocol reveal that 1) chewing-induced changes in performance and task-related mydriasis are correlated and 2) neither performance nor mydriasis are enhanced by handgrip. In the second protocol, use of a wearable pupillometer allows measurement of pupil size changes and performance during the same task, thus allowing even stronger evidence to be obtained regarding LC involvement in the trigeminal effects on cognitive activity. Both protocols have been run in the historical office of Prof. Giuseppe Moruzzi, the discoverer of ARAS, at the University of Pisa.

To verify whether trigeminal effects on cognitive performance involve locus coeruleus activity, two protocols are presented that aim to evaluate possible correlations between the performance and task-related pupil size changes induced by chewing. These protocols may be applied to conditions in which locus coeruleus contribution is suspected.

Current scientific literature provides evidence that trigeminal sensorimotor activity associated with chewing may affect arousal, attention, and cognitive ...

Wireless Electrophysiological Recording of Neurons by Movable Tetrodes in Freely Swimming Fish

The neural mechanisms governing fish behavior remain mostly unknown, although fish constitute the majority of all vertebrates. The ability to record brain activity from freely moving fish would advance research on the neural basis of fish behavior considerably. Moreover, precise control of the recording location in the brain is critical to studying coordinated neural activity across regions in fish brain. Here, we present a technique that records wirelessly from the brain of freely swimming fish while controlling for the depth of the recording location. The system is based on a neural logger associated with a novel water-compatible implant that can adjust the recording location by microdrive-controlled tetrodes. The capabilities of the system are illustrated through recordings from the telencephalon of goldfish.

A novel wireless technique for recording extracellular neural signals from the brain of freely swimming goldfish is presented. The recording device is composed of two tetrodes, a microdrive, a neural data logger, and a waterproof case. All parts are custom-made except for the data logger and its connector.

The neural mechanisms governing fish behavior remain mostly unknown, although fish constitute the majority of all vertebrates. The ability to record brain ...

Assessing Early Stage Open-Angle Glaucoma in Patients by Isolated-Check Visual Evoked Potential

Recently, the isolated-check visual evoked potential (icVEP) technique was designed and has been reported to detect glaucomatous damage earlier and faster. It creates low spatial frequency/high temporal frequency bright stimuli and records cortical activity initiated primarily by afferents in the magnocellular ON pathway. This pathway contains neurons with larger volumes and axonal diameters, and it is preferentially damaged in early glaucoma, which can result in visual field loss. The study presented here uses standard operative procedures (SOP) of icVEP to obtain reliable results. It can detect visual function loss using a signal-to-noise ratio (SNR) corresponding to the defects of retinal nerve fiber layer (RNFL) in early stage open-angle glaucoma (OAG). A setting of 10 Hz and condition of 15% positive-contrast (bright) are selected to differentiate OAG patients and control subjects, with each check containing eight runs. Each run persists for 2 s (for 20 total cycles). A flowchart is constructed, which consists of pupil size and intraocular pressure over a 30 min rest period before each examination. Additionally, the testing order of eyes is performed to obtain reliable electroencephalographic signals. VEPs are recorded and analyzed automatically by software, and SNRs are derived based on a multivariate statistic. An SNR of &#8804; 1 is considered abnormal. A receiver-operating-characteristic (ROC) curve is applied to analyze the accuracy of group classification. Then, the SOP is applied in a cross-sectional study, showing that icVEP can detect glaucomatous visual function abnormality in the central visual field in the form of SNR. This value also correlates with the thickness thinning of RNFL and produces high classification accuracy for early stage OAG. Thus, it serves as a useful and objective diagnostic technology for the early detection of glaucoma.

The isolated-check visual evoked potential (icVEP) method is implemented here to assess the magnocellular ON pathway that is initially damaged in glaucoma. The study shows standard operative procedures using icVEP to obtain reliable results. It is proved to serve as a useful objective diagnosing technology for the early detection of glaucoma.

Recently, the isolated-check visual evoked potential (icVEP) technique was designed and has been reported to detect glaucomatous damage earlier and ...

Ocular Kinematics Measured by In Vitro Stimulation of the Cranial Nerves in the Turtle

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Chapters in this video

TMS: Using the Theta-Burst Protocol to Explore Mechanism of Plasticity in Individuals with Fragile X Syndrome and Autism

The NeuroStar TMS Device: Conducting the FDA Approved Protocol for Treatment of Depression

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Generation of Local CA1 γ Oscillations by Tetanic Stimulation

Caspase-3 Activity in the Rat Amygdala Measured by Spectrofluorometry After Myocardial Infarction

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

The Power of Interstimulus Interval for the Assessment of Temporal Processing in Rodents

Assessing Pupil-linked Changes in Locus Coeruleus-mediated Arousal Elicited by Trigeminal Stimulation

Wireless Electrophysiological Recording of Neurons by Movable Tetrodes in Freely Swimming Fish

Assessing Early Stage Open-Angle Glaucoma in Patients by Isolated-Check Visual Evoked Potential