We are thankful to Yingtian He for sharing data of direction tuning. This work was supported by grants from the Canadian Foundation of Innovation and Ontario Research Fund (CFI/ORF project no. 37597), NSERC (RGPIN-2019-06479), CIHR (Project Grant 437007), and Connaught New Researcher Awards.

All experimental procedures performed in this study were approved by the Biological Sciences Local Animal Care Committee, in accordance with guidelines established by the University of Toronto Animal Care Committee and the Canadian Council on Animal Care.
1. Implantation of a head bar on top of the skull
NOTE: To avoid the contribution of VOR behavior to the eye movements, the head of the mouse is immobilized during the OKR test. Therefore, a head bar is surgically implanted on top of the skull.
<ol>
	<li>Anesthetize a mouse (2-5-month-old female and male C57BL/6) by a mixture of 4% isoflurane (v/v) and O2 in a gas chamber. Transfer the mouse to a customized surgery platform and reduce the concentration of isoflurane to 1.5%-2%. Monitor the depth of anesthesia by checking the toe-pinch response and the respiration rate throughout the surgery.</li>
	<li>Place a heating pad underneath the animal&#39;s body to maintain its body temperature. Apply a layer of lubricant eye ointment to both eyes to protect them from drying. Cover the eyes with aluminum foil to protect them from light illumination.</li>
	<li>Subcutaneously inject carprofen at a dose of 20&#160;mg/kg to reduce pain. After wetting the fur with chlorhexidine gluconate skin cleaner, shave the fur on top of the skull. Disinfect the exposed scalp with 70% isopropyl alcohol and chlorhexidine alcohol twice.</li>
	<li>Inject bupivacaine (8 mg/kg) subcutaneously at the site of incision, then remove the scalp (~1 cm2) with scissors to expose the dorsal surface of the skull, including the posterior frontal bone, parietal bone, and interparietal bone.</li>
	<li>Apply several drops of 1% lidocaine and 1:100,000 epinephrine on the exposed skull to reduce local pain and bleeding. Scrape the skull with a Meyhoefer curette to remove the fascia and clean it with phosphate-buffered saline (PBS). 
	NOTE: The temporalis muscle is separated from the skull to increase the surface area for a head bar to attach.</li>
	<li>Dry the skull by gently blowing compressed air toward the skull surface until the moisture is gone and the bone turns whitish. Apply a thin layer of superglue on the exposed surface of the skull, including the edge of the cut scalp, followed by a layer of acrylic resin. 
	NOTE: The surface of the skull needs to be free of blood or water before the application of superglue.</li>
	<li>Place a stainless-steel head bar (see Figure 1A) along the midline on top of the skull. Apply more acrylic resin, starting from the edge of the head bar until the base of the head bar is completely embedded in the acrylic resin. Apply acrylic resin two or three times to build up the thickness.</li>
	<li>Wait for about 15 min until the acrylic resin hardens. Subcutaneously inject 1 mL of lactated ringer&#39;s solution. Then, return the mouse to a cage placed on a heating pad until the animal is fully mobile.</li>
	<li>Allow the mouse to recover in the home cage for at least 5 days after surgery. Once the animal is in good shape, fix its head with the head bar in the OKR setup for 15-30 min to familiarize it with the head fixation and the experimental environment. Repeat the familiarization once a day for at least 3 days.</li>
</ol>
2. Setup of the virtual drum and video-oculography
<ol>
	<li>Mount three monitors orthogonally to each other to form a square enclosure that covers ~270&#176; of the azimuth and 63&#176; of the elevation in the visual space (Figure 1B left).</li>
	<li>With a discrete graphic card, merge the three monitors into a simple display to ensure synchronization across all the monitors.</li>
	<li>Calibrate the luminance of monitors as described below.
	<ol>
		<li>Turn on the computer to which the monitors are connected and wait for 15 min. The warm-up is essential to have stable luminance.</li>
		<li>Systematically change the brightness setting on the monitor from 0 to 100 by steps of 25.</li>
		<li>For each brightness value, measure the luminance of the monitors under various pixel values (0-255, steps of 15) with a luminance meter.</li>
		<li>Fit the relationship between luminance and brightness for pixel value 255 with linear regression and estimate the brightness value that gives rise to 160 cd/m2.</li>
		<li>For each pixel value used in the luminance measurement (step 2.3.3), estimate the luminance for the brightness value derived in step 2.3.4 based on linear regression. Use the power function lum = A * pixel&#947; to fit the relationship between the new set of luminance values (under the brightness value derived in 2.3.4) and their corresponding pixel values to derive the gamma factor &#947; and the coefficient A. These will be used to generate sinusoidal gratings of desired luminance values.</li>
		<li>Set the brightness of all three monitors to the values derived in step 2.3.4 to ensure their luminance values are the same for the same pixel value.</li>
	</ol>
	</li>
	<li>Generate a virtual drum, which is used to stimulate OKR behavior, with the visual stimulation toolkit, as described below.
	<ol>
		<li>Present a vertical sinusoidal grating on the monitors and adjust the period (spacing between stripes) along the azimuth to ensure the projection of the grating on the eye has constant spatial frequency (drum grating; Figure 1B middle and right).</li>
		<li>Ensure the animal head-fixed at the center of the enclosure so that it sees the grating has a constant spatial frequency across the surface of the virtual drum.</li>
		<li>Modify the parameters of moving grating, such as the oscillatory amplitude, spatial frequency, temporal/oscillation frequency, direction, contrast, etc., in the visual stimulation codes. Use two types of visual motion: (1) the grating drifts clockwise or counterclockwise in an oscillatory manner following a sinusoidal function: 
		<img alt="Equation 1" src="/files/ftp_upload/65281/65281eq01v2.jpg" style="margin: auto;" /> 
		Here, Amp is the amplitude of drum trajectory, f is the oscillation frequency, and t is the time (oscillation amplitude: 5&#176;; grating spatial frequency: 0.04-0.45 cpd; oscillation frequency: 0.1-0.8 Hz, corresponding to a peak velocity of the stimulus of 3.14-25.12 &#176;/s [drum velocity = Amp x 2&#960; x f x cos (2&#960; x f x t); contrast: 80%-100%; mean luminance: 35-45 cd/m2; (2) the grating drifts unidirectionally at a constant velocity: 
		<img alt="Equation 2" src="/files/ftp_upload/65281/65281eq02v2.jpg" style="margin: auto;" /> 
		&#8203;(Spatial frequency: 0.04-0.64 cpd; temporal frequency: 0.25-1 Hz; drum velocity = temporal frequency/spatial frequency.)</li>
	</ol>
	</li>
	<li>Set up the video-oculography as described below.
	<ol>
		<li>To avoid blockage of animal&#39;s visual field, place an infrared (IR) mirror 60&#176; from the midline to form an image of the right eye.</li>
		<li>Place an IR&#160;camera on the right side behind the mouse (Figure 1C left) to capture an image of the right eye.</li>
		<li>Mount the high-speed IR camera on a camera arm that allows the camera to rotate by &#177; 10&#176; around the image of the right eye (Figure 1C right).</li>
		<li>Use a photodiode attached to one of the monitors to provide an electrical signal to synchronize the timing of video-oculography and visual stimulation.</li>
		<li>Place four IR light emitting diodes (LEDs) supported by gooseneck arms around the right eye to provide IR illumination of the eye.</li>
		<li>Place two IR&#160;LEDs on the camera to provide corneal reflection (CR) references: one is fixed above the camera (X-CR), while the other is on the left side of the camera (Y-CR; Figure 1D).</li>
		<li>Measure the optical magnification of the video-oculography system with a calibration slide. 
		NOTE: The reference CRs are used to cancel out the translational eye movements when the eye angle is calculated based on the rotational eye movements.</li>
	</ol>
	</li>
	<li>Fix the animal&#39;s head at the center of the enclosure formed by the monitors, as described below.
	<ol>
		<li>Fix the animal&#39;s head with the head plate to the center of the rig and make it face forward. Adjust the tilt of the head so that the left and right eyes are leveled, and the nasal and temporal corners of the eyes are aligned horizontally (Figure 1E).</li>
		<li>Move the animal&#39;s head horizontally by coarse adjustment provided by the head-fixation apparatus and fine adjustment provided by a 2D translational stage, and vertically through the head-fixation apparatus and a post/post holder pair, until the animal&#39;s right eye appears in the live video of the camera. Before the calibration and measurement of eye movements, overlay the image of the animal&#39;s right eye reflected by the hot mirror with the pivot point of the camera arm (see details in step 3.4 below).</li>
	</ol>
	</li>
	<li>Build a customized enclosure around the OKR rig to block room light (Figure 1F).</li>
</ol>
3. Calibration of eye movements
NOTE: Rotational eye movements are calculated based on movements of the pupil and the radius of the orbit of the pupillary movements (Rp, the distance from the center of the pupil to the center of the eyeball). For each individual mouse, this radius is measured experimentally21.
<ol>
	<li>Fix the animal&#39;s head at the center of the enclosure formed by the three monitors, as described in step 2.6.1.</li>
	<li>Turn the camera on and adjust the four LEDs surrounding the right eye to achieve uniform IR illumination.</li>
	<li>Under visual guidance, adjust the position of the right eye until it appears at the center of the video, as described in step 2.6.2.</li>
	<li>Align the virtual image of the right eye with the pivot point of the camera arm, as described below.
	<ol>
		<li>Manually rotate the camera arm to the left extreme end (-10&#176;). Manually move the animal&#39;s right eye position on the horizontal plane perpendicular to the optical axis with fine adjustment of the 2D translational stage (Figure 1C, green arrow), until the X-CR is at the horizontal center of the image.</li>
		<li>Manually rotate the camera arm to the other end (+10&#176;). If the X-CR runs away from the center of the image, move the right eye along the optical axis with fine adjustment until the X-CR comes to the center (Figure 1C, blue arrow).</li>
		<li>Repeat steps 3.4.1-3.4.2 a few times until the X-CR stays at the center when the camera arm swings leftward and rightward. If the right eye moves in the middle of one repetition, restart the adjustment process.</li>
	</ol>
	</li>
	<li>Measure the vertical distance between the Y-CR and X-CR after locking the camera arm at the central position. Turn the Y-CR LED on and record its position on the video, and then switch to the X-CR LED and record its position. 
	NOTE: The vertical distance between the Y-CR and X-CR will be used to derive the position of the Y-CR during the measurement of eye movements in which only the X-CR LED is turned on.</li>
	<li>Measure the radius of pupil rotation Rp, as described&#160;below.
	<ol>
		<li>Rotate the camera arm to the left end (-10&#176;) and record the positions of the pupil (Pp1) and X-CR (PCR1) on the video.</li>
		<li>Then, rotate the camera arm to the right end (+10&#176;) and record the positions of the pupil (Pp2) and X-CR (PCR2) on the video. Repeat this step multiple times. 
		NOTE: The animal&#39;s right eye needs to remain stationary during each repetition so that the amount of pupil movements in the movie accurately reflects the degree of swinging the camera arm.</li>
		<li>Based on the values recorded above, calculate the radius of pupil rotation Rp (Figure 2A) with the following formula: 
		<img alt="Equation 3" src="/files/ftp_upload/65281/65281eq03.jpg" style="margin: auto;" /> 
		NOTE: The distance between the corneal reflection and pupil center in the physical space is calculated based on their distance in the movie: 
		PCR - Pp = number of pixels in the movie x pixel size of camera chip x magnification</li>
	</ol>
	</li>
	<li>Develop the relationship between Rp and pupil diameter, as described below. Rp changes when the pupil dilates or constricts; proximately, its value is inversely proportional to the pupil size (Figure 2B top).
	<ol>
		<li>Change the luminance of the monitors systematically from 0 to 160 cd/m2 to regulate the pupil size.</li>
		<li>For each luminance value, repeat step 3.6 8-10 times and record the diameter of the pupil.</li>
		<li>Apply linear regression to the relationship between Rp and pupil diameter based on the values measured above to derive the slope and intercept (Figure 2B bottom). 
		NOTE: The outliers caused by occasional eye movements are removed before the linear fitting. For repetitive measurements in multiple sessions, the calibration needs to be done only once for one animal, unless its eye grows bigger during the experiment.</li>
	</ol>
	</li>
</ol>
4. Record eye movements of the OKR
<ol>
	<li>Head-fix a mouse in the rig following steps 3.1-3.4. Skip this step if the recording occurs right after the calibration is done. Lock the camera arm at the central position.</li>
	<li>Set up the monitors and the animal for scotopic OKR as described below. Skip this step for photopic OKR.
	<ol>
		<li>Cover the screen of each monitor with a customized filter, which is made of five layers of 1.2 neutral density (ND) film. Make sure no light leaks out through the gap between the filter and the monitor.</li>
		<li>Turn the room light off. The following steps are done with the aid of an IR goggle.</li>
		<li>Apply one drop of pilocarpine solution (2% in saline) to the right eye and wait 15 min. Ensure the drop stays on the eye and is not wiped away by the mouse. If the solution is wiped away by the animal, apply another drop of pilocarpine solution. This shrinks the pupil to a proper size for eye tracking under the scotopic condition. 
		NOTE: Under the scotopic condition, the pupil dilates substantially so that its edge is partially hidden behind the eyelid. This affects the precision of estimating the pupil center by video-oculography. Pharmacologically shrinking the pupil of the right eye diminishes its visual input, and thus the visual stimuli are presented to the left eye.</li>
		<li>Rinse the right eye with saline to thoroughly wash away the pilocarpine solution. Pull down the curtain to completely seal the enclosure, which prevents stray light from interfering with the scotopic vision.</li>
		<li>Give the animal 5 min to fully accommodate to the scotopic environment before starting the OKR test.</li>
	</ol>
	</li>
	<li>Run the visual stimulation software and the eye tracking software. For photopic OKR measurement, ensure the drum grating oscillates horizontally with a sinusoidal trajectory; for scotopic OKR measurement, ensure the drum grating drifts at a constant velocity from left to right, which is the temporo-nasal direction in reference to the left eye. 
	NOTE: When the pupil of the right eye, but not of the left eye, is shrunk by pilocarpine under the scotopic condition, the OKR elicited by oscillatory drum stimulation is highly asymmetric. Thus, for scotopic OKR measurement, the left eye is stimulated while the movement of the right eye is monitored.</li>
	<li>The eye tracking software automatically measures the pupil size, CR position, and pupil position for each frame, and computes the angle of eye position based on the following formula (Figure 2C): 
	<img alt="Equation 4" src="/files/ftp_upload/65281/65281eq04.jpg" style="margin: auto;" /> 
	Here, PCR is the CR position, Pp is the pupil position, and Rp is the radius of pupil rotation. The distance between corneal reflection and pupil center in the physical space is calculated based on their distance in the movie: 
	PCR - Pp = number of pixels in the movie x pixel size of camera chip x magnification 
	Rp of the corresponding pupil size is derived based on the linear regression model in step 3.7.3 (Figure 2B bottom).</li>
</ol>
5. Analysis of eye movements of the OKR with the eye analysis software
<ol>
	<li>Process the eye traces using a median filter (filter window = 0.05 s) to remove high frequency noise (Figure 3A middle).</li>
	<li>Remove the saccades or nystagmus as described below.
	<ol>
		<li>Estimate the eye velocity by calculating the first order derivative of eye movements (Figure 3A bottom). Identify the saccades or nystagmus by applying a velocity threshold of 50 &#176;/s (Figure 3A bottom).</li>
		<li>Replace the saccades or nystagmus by extrapolating the eye positions during these fast eye movements from the segment before the saccades or nystagmus based on linear regression (Figure 3B).</li>
	</ol>
	</li>
	<li>Compute the amplitude of OKR eye movements by fast Fourier transform (Goertzel algorithm) if the drum grating oscillates (Figure 3C), or compute the average velocity of eye movements during the visual stimulation if the drum grating moves at a constant velocity in one direction (Figure 3B bottom). 
	NOTE: The amplitude of oscillatory eye movements derived from Fourier transform is similar to the amplitude derived from the fitting of the eye trajectory with a sinusoidal function (Figure 3D).</li>
	<li>Calculate the OKR gain. For oscillatory drum motion, OKR gain is defined as the ratio of the amplitude of eye movements to the amplitude of drum movements (Figure 3C right). For unidirectional drum motion, OKR gain is defined as the ratio of eye velocity to drum grating velocity (Figure 3B bottom).</li>
</ol>

Evaluating Optokinetic Reflex Responses in Mice: A New High-Precision Protocol

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L., Kaja, S., Koulen, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantification+of+deficits+in+spatial+visual+function+of+mouse+models+for+glaucoma.">Quantification of deficits in spatial visual function of mouse models for glaucoma.</a> Investigative Ophthalmology &amp; Visual Science. 52 (6), 3654-3659 (2011).</li><li>Wakita, 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=Differential+regulations+of+vestibulo-ocular+reflex+and+optokinetic+response+by+&#946;-+and+&#945;2-adrenergic+receptors+in+the+cerebellar+flocculus.">Differential regulations of vestibulo-ocular reflex and optokinetic response by &#946;- and &#945;2-adrenergic receptors in the cerebellar flocculus.</a> Scientific Reports. 7 (1), 3944 (2017).</li><li>Dehmelt, F. A., 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=Spherical+arena+reveals+optokinetic+response+tuning+to+stimulus+location,+size,+and+frequency+across+entire+visual+field+of+larval+zebrafish.">Spherical arena reveals optokinetic response tuning to stimulus location, size, and frequency across entire visual field of larval zebrafish.</a> eLife. 10, e63355 (2021).</li><li>Magnusson, M., Pyykko, I., Jantti, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+alertness+and+visual+attention+on+optokinetic+nystagmus+in+humans.">Effect of alertness and visual attention on optokinetic nystagmus in humans.</a> American Journal of Otolaryngology. 6 (6), 419-425 (1985).</li><li>Collins, W. E., Schroeder, D. J., Elam, G. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+D-amphetamine+and+of+secobarbital+on+optokinetic+and+rotation-induced+nystagmus.">Effects of D-amphetamine and of secobarbital on optokinetic and rotation-induced nystagmus.</a> Aviation, Space, and Environmental Medicine. 46 (4), 357-364 (1975).</li><li>Reimer, J., 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+fluctuations+track+fast+switching+of+cortical+states+during+quiet+wakefulness.">Pupil fluctuations track fast switching of cortical states during quiet wakefulness.</a> Neuron. 84 (2), 355-362 (2014).</li><li>Sakatani, T., Isa, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PC-based+high-speed+video-oculography+for+measuring+rapid+eye+movements+in+mice.">PC-based high-speed video-oculography for measuring rapid eye movements in mice.</a> Neuroscience Research. 49 (1), 123-131 (2004).</li><li>Sakatani, T., Isa, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+analysis+of+spontaneous+saccade-like+rapid+eye+movements+in+C57BL/6+mice.">Quantitative analysis of spontaneous saccade-like rapid eye movements in C57BL/6 mice.</a> Neuroscience Research. 58 (3), 324-331 (2007).</li><li>Vinck, M., Batista-Brito, R., Knoblich, U., Cardin, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Arousal+and+locomotion+make+distinct+contributions+to+cortical+activity+patterns+and+visual+encoding.">Arousal and locomotion make distinct contributions to cortical activity patterns and visual encoding.</a> Neuron. 86 (3), 740-754 (2015).</li><li>Bradley, M. M., Miccoli, L., Escrig, M. A., Lang, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+pupil+as+a+measure+of+emotional+arousal+and+autonomic+activation.">The pupil as a measure of emotional arousal and autonomic activation.</a> Psychophysiology. 45 (4), 602-607 (2008).</li><li>Hess, E. H., Polt, 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=Pupil+size+as+related+to+interest+value+of+visual+stimuli.">Pupil size as related to interest value of visual stimuli.</a> Science. 132 (3423), 349-350 (1960).</li><li>Di Stasi, L. L., Catena, A., Canas, J. J., Macknik, S. L., Martinez-Conde, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Saccadic+velocity+as+an+arousal+index+in+naturalistic+tasks.">Saccadic velocity as an arousal index in naturalistic tasks.</a> Neuroscience and Biobehavioral Reviews. 37 (5), 968-975 (2013).</li></ol>

The authors declare no competing interests.

The method of the OKR behavioral assay presented here provides several advantages. First, computer-generated visual stimulation solves the intrinsic issues of physical drums. Dealing with the issue that physical drums do not support the systematic examination of spatial frequency, direction, or contrast tuning8, the virtual drum allows these visual parameters to be changed on a trial-by-trial basis, thus facilitating a systematic and quantitative analysis of the feature selectivity of the OKR beha...

In response to visual stimuli in the environment, our eyes move to shift our gaze, stabilize retinal images, track moving targets, or align the foveae of two eyes with targets located at different distances from the observer, which are vital to proper vision1,2. Oculomotor behaviors have been widely used as attractive models of sensorimotor integration to understand the neural circuits in health and disease, at least partly because of the simplicity of the oculomotor system3. Controlled by three pairs of extraocular muscles, the eye rotates in the socket primarily around three corresponding axes: elevation and depression along the transverse axis, adduction and abduction along the vertical axis, and intorsion and extorsion along the anteroposterior axis1,2. Such a simple system allows researchers to evaluate the oculomotor behaviors of mice easily and accurately in a lab environment.
One prime oculomotor behavior is the optokinetic reflex (OKR). This involuntary eye movement is triggered by slow drifts or slips of images on the retina and serves to stabilize retinal images as an animal&#39;s head or its surroundings move2,4. The OKR, as a behavioral paradigm, is interesting to researchers for several reasons. First, it can be stimulated reliably and quantified accurately5,6. Second, the procedures of quantifying this behavior are relatively simple and standardized and can be applied to evaluate the visual functions of a large cohort of animals7. Third, this innate behavior is highly plastic5,8,9. Its amplitude can be potentiated when repetitive retinal slips occur for a long time5,8,9, or when its working partner vestibular ocular reflex (VOR), another mechanism of stabilizing retinal images triggered by vestibular input2, is impaired5. These experimental paradigms of OKR potentiation empower researchers to unveil the circuit basis underlying oculomotor learning.
Two non-invasive methods have primarily been used to evaluate the OKR in previous studies: (1) video-oculography combined with a physical drum7,10,11,12,13 or (2) arbitrary determination of head turns combined with a virtual drum6,14,15,16. Although their applications have made fruitful discoveries in understanding the molecular and circuit mechanisms of oculomotor plasticity, these two methods each have some drawbacks which limit their powers in quantitatively examining the properties of the OKR. First, physical drums, with printed patterns of black and white stripes or dots, do not allow easy and quick changes of visual patterns, which largely restricts the measurement of the dependence of the OKR on certain visual features, such as spatial frequency, direction, and contrast of moving gratings8,17. Instead, tests of the selectivity of the OKR to these visual features can benefit from computerized visual stimulation, in which visual features can be conveniently modified from trial to trial. In this way, researchers can systematically examine the OKR behavior in the multi-dimensional visual parameter space. Moreover, the second method of the OKR assay reports only the thresholds of visual parameters that trigger discernible OKRs, but not the amplitudes of eye or head movements6, 14,15,16. The lack of quantitative power thus prevents analyzing the shape of tuning curves and the preferred visual features, or detecting subtle differences between individual mice in normal and pathological conditions. To overcome the above limitations, video-oculography and computerized virtual visual stimulation had been combined to assay the OKR behavior in recent studies5,17,18,19,20. However, these previously published studies did not provide enough technical details or step-by-step instructions, and consequently it is still challenging for researchers to establish such an OKR test for their own research.
Here, we present a protocol to precisely quantify the visual feature selectivity of OKR behavior under photopic or scotopic conditions with the combination of video-oculography and computerized virtual visual stimulation. Mice are head-fixed to avoid the eye movement evoked by vestibular stimulation. A high-speed camera is used to record the ocular movements from mice viewing moving gratings with changing visual parameters. The physical size of the eyeballs of individual mice is calibrated to ensure the accuracy of deriving the angle of eye movements21. This quantitative method allows comparing OKR behavior between animals of different ages or genetic backgrounds, or monitoring its change caused by pharmacological treatments or visual-motor learning.

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			<td>8507-14-1</td>
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			<td>Graphic card</td>
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			<td>Kent Scientific</td>
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			<td>High-speed infrared (IR) camera</td>
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			<td>IR LED</td>
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			<td>Isoflurane</td>
			<td>FRESENIUS KABI</td>
			<td>CP0406V2</td>
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			<td>Labview</td>
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			<td>XYLOCAINE. Local anesthesia</td>
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			<td>Luminance Meter</td>
			<td>Konica Minolta</td>
			<td>LS-150</td>
			<td>for calibration of monitors</td>
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		<tr>
			<td>Matlab</td>
			<td>MathWorks</td>
			<td>version xxx</td>
			<td>analysis of eye movements</td>
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			<td>Meyhoefer Curette</td>
			<td>World Precision Instruments</td>
			<td>501773</td>
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		<tr>
			<td>Microscope calibration slide</td>
			<td>Amscope</td>
			<td>MR095</td>
			<td>to measure the magnification of video-oculography</td>
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			<td>Monitors</td>
			<td>Acer&#160;</td>
			<td>B247W</td>
			<td>Visual stimulation</td>
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			<td>Neutral density filter</td>
			<td>Lee filters</td>
			<td>299</td>
			<td>to generate scotopic visual stimulation</td>
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			<td>Nigh vision goggle</td>
			<td>Alpha optics</td>
			<td>AO-3277</td>
			<td>for scotopic OKR</td>
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		<tr>
			<td>Photodiode</td>
			<td>Digikey</td>
			<td>TSL254-R-LF-ND</td>
			<td>to synchronize visual stimulation and video-oculography</td>
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			<td>Sigma-Aldrich</td>
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			<td>version xxx</td>
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			<td>Scissor</td>
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			<td>Superglue</td>
			<td>Krazy Glue</td>
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quantification of visual feature selectivity of the optokinetic reflex in mice

The optokinetic reflex (OKR) is an essential innate eye movement that is triggered by the global motion of the visual environment and serves to stabilize retinal images. Due to its importance and robustness, the OKR has been used to study visual-motor learning and to evaluate the visual functions of mice with different genetic backgrounds, ages, and drug treatments. Here, we introduce a procedure for evaluating OKR responses of head-fixed mice with high accuracy. Head fixation can rule out the contribution of vestibular stimulation on eye movements, making it possible to measure eye movements triggered only by visual motion. The OKR is elicited by a virtual drum system, in which a vertical grating presented on three computer monitors drifts horizontally in an oscillatory manner or unidirectionally at a constant velocity. With this virtual reality system, we can systematically change visual parameters like spatial frequency, temporal/oscillation frequency, contrast, luminance, and the direction of gratings, and quantify tuning curves of visual feature selectivity. High-speed infrared video-oculography ensures accurate measurement of the trajectory of eye movements. The eyes of individual mice are calibrated to provide opportunities to compare the OKRs between animals of different ages, genders, and genetic backgrounds. The quantitative power of this technique allows it to detect changes in the OKR when this behavior plastically adapts due to aging, sensory experience, or motor learning; thus, it makes this technique a valuable addition to the repertoire of tools used to investigate the plasticity of ocular behaviors.

With the procedure detailed above, we evaluated the dependence of the OKR on several visual features. The example traces shown here were derived using the analysis codes provided in Supplementary Coding File 1, and the example traces raw file can be found in Supplementary Coding File 2. When the drum grating drifted in a sinusoidal trajectory (0.4 Hz), the animal&#39;s eye automatically followed the movement of the grating in a similar oscillatory manner (Figure 3B</...

Watch this Scientific Journal Video about An Accurate and Quantitative Approach to Study Visual Feature Selectivity of the Optokinetic Reflex in Mice at JoVE.com

Author Spotlight: An Accurate and Quantitative Approach to Study Visual Feature Selectivity of the Optokinetic Reflex in Mice 

Here, we describe a standard protocol for quantifying the optokinetic reflex. It combines virtual drum stimulation and video-oculography, and thus allows precise evaluation of the feature selectivity of the behavior and its adaptive plasticity.

Quantification of Visual Feature Selectivity of the Optokinetic Reflex in Mice

The optokinetic reflex (OKR) is an essential innate eye movement that is triggered by the global motion of the visual environment and serves to stabilize ...

quantification-visual-feature-selectivity-optokinetic-reflex

Research

JoVE Journal

Neuroscience

2.2K Views.  University of Toronto Mississauga. Here, we describe a standard protocol for quantifying the optokinetic reflex. It combines virtual drum stimulation and video-oculography, and thus allows precise evaluation of the feature selectivity of the behavior and its adaptive plasticity. 

Video: Author Spotlight: An Accurate and Quantitative Approach to Study Visual Feature Selectivity of the Optokinetic Reflex in Mice 

The Optokinetic Response as a Quantitative Measure of Visual Acuity in Zebrafish

Zebrafish are a proven model for vision research, however many of the earlier methods generally focused on larval fish or demonstrated a simple response. More recently adult visual behavior in zebrafish has become of interest, but methods to measure specific responses are new coming. To address this gap, we set out to develop a methodology to repeatedly and accurately utilize the optokinetic response (OKR) to measure visual acuity in adult zebrafish. Here we show that the adult zebrafish's visual acuity can be measured, including both binocular and monocular acuities. Because the fish is not harmed during the procedure, the visual acuity can be measured and compared over short or long periods of time. The visual acuity measurements described here can also be done quickly allowing for high throughput and for additional visual procedures if desired. This type of analysis is conducive to drug intervention studies or investigations of disease progression.

Zebrafish have been a valuable tool for many research areas. Here we demonstrate a method to elicit a visual response and calculate a functional visual acuity in the adult zebrafish.

Zebrafish are a proven model for vision research, however many of the  earlier methods generally focused on larval fish or demonstrated a simple  ...

Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the surface of the cornea, electrical activity generated in response to light can be measured and used to assess the activity of retinal cells in vivo. This manuscript describes the use of the ERG to measure visual function in zebrafish. Zebrafish have long been utilized as a model for vertebrate development due to the ease of gene suppression by morpholino oligonucleotides and pharmacological manipulation. At 5-10 dpf, only cones are functional in the larval retina. Therefore, the zebrafish, unlike other animals, is a powerful model system for the study of cone visual function in vivo. This protocol uses standard anesthesia, micromanipulation and stereomicroscopy protocols that are common in laboratories that perform zebrafish research. The outlined methods make use of standard electrophysiology equipment and a low light camera to guide the placement of the recording microelectrode onto the larval cornea. Finally, we demonstrate how a commercially available ERG stimulator/recorder originally designed for use with mice can easily be adapted for use with zebrafish. ERG of larval zebrafish provides an excellent method of assaying cone visual function in animals that have been modified by morpholino oligonucleotide injection as well as newer genome engineering techniques such as Zinc Finger Nucleases (ZFNs), Transcription Activator-Like Effector Nucleases (TALENs), and Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9, all of which have greatly increased the efficiency and efficacy of gene targeting in zebrafish. In addition, we take advantage of the ability of pharmacological agents to penetrate zebrafish larvae to evaluate the molecular components that contribute to the photoresponse. This protocol outlines a setup that can be modified and used by researchers with various experimental goals.

We present a method for the electroretinographic (ERG) analysis of zebrafish larvae utilizing micromanipulation and electroretinography techniques. This is a simple and straightforward method for assaying visual function of zebrafish larvae in vivo.

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Visual Evoked Potential Recording in a Rat Model of Experimental Optic Nerve Demyelination

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to monitor the disease course in the follow-up period. Changes in the VEP parameters closely correlate with pathological damage in the optic nerve. This protocol provides a detailed description about the rodent model of optic nerve microinjection, in which a partial demyelination lesion is produced in the optic nerve. VEP recording techniques are also discussed. Using skull implanted electrodes, we are able to acquire reproducible intra-session and between-session VEP traces. VEPs can be recorded on individual animals over a period of time to assess the functional changes in the optic nerve longitudinally. The optic nerve demyelination model, in conjunction with the VEP recording protocol, provides a tool to investigate the disease processes associated with demyelination and remyelination, and can potentially be employed to evaluate the effects of new remyelinating drugs or neuroprotective therapies.

Focal demyelination is induced in the optic nerve using lysolecithin microinjection. Visual evoked potentials are recorded via skull electrodes implanted over the visual cortex to examine the signal conduction along the visual pathway in vivo. This protocol details the surgical procedures underlying electrode implantation and optic nerve microinjection.

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to ...

Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich structures suggest alterations in synaptic integrity and connectivity. Here we provide a detailed protocol for culturing primary rat cortical neurons, Phalloidin staining for F-actin puncta, and subsequent quantification techniques. First, the frontal cortex of E18 rat embryos are dissociated into low-density cell culture, then the neurons grown in vitro for at least 12-14 days. Following experimental treatment, the cortical neurons are stained with AlexaFluor 488 Phalloidin (to label the dendritic F-actin puncta) and microtubule-associated protein 2 (MAP2; to validate the neuronal cells and dendritic integrity). Finally, specialized software is used to analyze and quantify randomly selected neuronal dendrites. F-actin rich structures are identified on second order dendritic branches (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence. The protocol presented here will be a useful method for investigating changes in dendritic synapse structures subsequent to experimental treatments.

Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich ...

Whole-cell Patch-clamp Recordings for Electrophysiological Determination of Ion Selectivity in Channelrhodopsins

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate electrical processes in target cells. In this context, ion selectivity of a channelrhodopsin is of particular importance. This article describes the investigation of chloride selectivity for a recently identified anion-conducting channelrhodopsin of Proteomonas sulcata via electrophysiological patch-clamp recordings on HEK293 cells. The experimental procedure for measuring light-gated photocurrents demands a fast switchable &#8211; ideally monochromatic &#8211; light source coupled into the microscope of an otherwise conventional patch-clamp setup. Preparative procedures prior to the experiment are outlined involving preparation of buffered solutions, considerations on liquid junction potentials, seeding and transfection of cells, and pulling of patch pipettes. The actual recording of current-voltage relations to determine the reversal potentials for different chloride concentrations takes place 24 h to 48 h after transfection. Finally, electrophysiological data are analyzed with respect to theoretical considerations of chloride conduction.

This article describes how the ion selectivity of channelrhodopsin is determined with electrophysiological whole-cell patch-clamp recordings using HEK293 cells. Here, the experimental procedure for investigating chloride selectivity of an anion-selective channelrhodopsin is demonstrated. However, the procedure is transferable to other channelrhodopsins of distinct selectivity.

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate ...

A Volumetric Method for Quantification of Cerebral Vasospasm in a Murine Model of Subarachnoid Hemorrhage

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor determining patient outcome and is therefore frequently taken as a study endpoint. However, in small animal studies on SAH, quantification of cerebral vasospasm is a major challenge. Here, an ex vivo method is presented that allows quantification of volumes of entire vessel segments, which can be used as an objective measure to quantify cerebral vasospasm. In a first step, endovascular casting of the cerebral vasculature is performed using a radiopaque casting agent. Then, cross-sectional imaging data are acquired by micro computed tomography. The final step involves 3-dimensional reconstruction of the virtual vascular tree, followed by an algorithm to calculate center lines and volumes of the selected vessel segments. The method resulted in a highly accurate virtual reconstruction of the cerebrovascular tree shown by a diameter-based comparison of anatomical samples with their virtual reconstructions. Compared with vessel diameters alone, the vessel volumes highlight the differences between vasospastic and non-vasospastic vessels shown in a series of SAH and sham-operated mice.

The goal of this article is to present a method that allows a 3-dimensional reconstruction of the cerebrovascular tree in mice after micro computed tomography and determination of volumes of entire vessel segments that can be used to quantify cerebral vasospasm in murine models of subarachnoid hemorrhage.

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor ...

Using Optical Coherence Tomography and Optokinetic Response As Structural and Functional Visual System Readouts in Mice and Rats

Optical coherence tomography (OCT) is a fast, non-invasive, interferometric technique allowing high-resolution retinal imaging. It is an ideal tool for the investigation of processes of neurodegeneration, neuroprotection and neuro-repair involving the visual system, as these often correlate well with retinal changes. As a functional readout, visually evoked compensatory eye and head movements are commonly used in experimental models involving the visual function. Combining both techniques allows a quantitative in vivo investigation of structure and function, which can be used to investigate the pathological conditions or to evaluate the potential of novel therapeutics. A great benefit of the presented techniques is the possibility to perform longitudinal analyses allowing the investigation of dynamic processes, reducing variability and cuts down the number of animals needed for the experiments. The protocol described aims to provide a manual for acquisition and analysis of high quality retinal scans of mice and rats using a low cost customized holder with an option to deliver inhalational anesthesia. Additionally, the proposed guide is intended as an instructional manual for researchers using optokinetic response (OKR) analysis in rodents, which can be adapted to their specific needs and interests.

A detailed protocol for the assessment of structural and visual readouts in rodents by optical coherence tomography and optokinetic response is presented. The results provide valuable insights for ophthalmologic as well as neurologic research.

Optical coherence tomography (OCT) is a fast, non-invasive, interferometric technique allowing high-resolution retinal imaging. It is an ideal tool for ...

Stimulus-specific Cortical Visual Evoked Potential Morphological Patterns

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using 128-channel high-density electroencephalography (EEG). The specific aim of the described stimuli and analyses is to examine whether it is feasible to replicate previously reported CVEP morphological patterns elicited by an apparent motion stimulus, designed to simultaneously stimulate both ventral and dorsal central visual networks, using object and motion stimuli designed to separately stimulate ventral and dorsal visual cortical networks.&#160; Four visual paradigms are presented: 1. Randomized visual objects with consistent temporal presentation. 2. Randomized visual objects with inconsistent temporal presentation (or jitter).&#160; 3. Visual motion via a radial field of coherent central dot motion without jitter.&#160; 4. Visual motion via a radial field of coherent central dot motion with jitter.&#160; These four paradigms are presented in a pseudo-randomized order for each participant.&#160; Jitter is introduced in order to view how possible anticipatory-related effects may affect the morphology of the object-onset and motion-onset CVEP response.&#160; EEG data analyses are described in detail, including steps of data exportation from and importation to signal processing platforms, bad channel identification&#160;and removal, artifact rejection, averaging, and categorization of average CVEP morphological pattern type based upon latency ranges of component peaks. Representative data show that the methodological approach is indeed sensitive in eliciting differential object-onset and motion-onset CVEP morphological patterns and may, therefore, be useful in addressing the larger research aim. Given the high temporal resolution of EEG and the possible application of high-density EEG in source localization analyses, this protocol is ideal for the investigation of distinct CVEP morphological patterns and the underlying neural mechanisms which generate these differential responses.&#160; &#160; &#160;&#160;

In this paper, we present a protocol to investigate differential cortical visual evoked potential morphological patterns through stimulation of ventral and dorsal networks using high-density EEG. Visual object and motion stimulus paradigms, with and without temporal jitter, are described. Visual evoked potential morphological analyses are also outlined.&#160;&#160;

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using ...

How to Obtain Reliable Visual Event-related Potentials in Newborns

The present study discusses the characteristics of visual event-related potentials (VEPs) and outlines methodological steps for obtaining reliable measurements in newborns. Obtaining high-quality, reliable VEPs is crucial for the early detection of abnormal development of the central nervous system in at-risk newborns, and for implementing successful early interventions. Recommendations are based on a previous study which showed that when post-conceptional age, polysomnography-identified sleep stages, and light-emitting diodes (LEDs) googles as the luminous source are controlled, no more than 4 repetitions of VEP averages are required to obtain replicable recordings, variability decreases, and reliable VEPs can be obtained. By controlling for these sources of variability and using statistical analyses, we were able to clearly and reliably identify the amplitude and latency of three main components (NII, PII and NIII) present in 100% of newborns (n = 20) during active sleep. Recording VEPs during awake states, quiet sleep and transitional sleep is not recommended because VEP morphology may differ significantly from one average to the next, leading to the risk of misleading clinical prognoses. Moreover, it is easier to obtain VEPs during active sleep because this state can be clearly and reliably identified at this stage of development, sleep cycles are short enough to allow measurements to be taken in a reasonable time, and the method does not require new o expensive equipment.

Several important points for obtaining high-quality reliable visual evoked potentials (VEPs) in newborns while minimizing variability and the risk of misleading prognoses are presented.