This work was supported by NINDS 5P30NS069250. We thank Diala Chehayeb, Jeffrey Simpson, Taylor Rosenbaum, and Christopher Hoover for technical assistance.

Implementing the headphones system consists of four major steps. Section 1 below details the assembly of the headphones frame, which houses the electronics (speakers and a miniaturized microphone). Section 2 describes how the frame is attached to the bird. Section 3 describes the assembly of the electronics. Section 4 explains how the electronics are connected to sound-processing and data-collection hardware and details a procedure for testing that the system is functioning correctly.
1. Fabricate Headphone Frame
<ol>
	<li>Cut carbon fiber components for crossbar and earbuds. Use the cutting wheel on a Dremel tool to prepare the following pieces: 
	1 crossbar (3 cm of 1 x 3 mm carbon fiber strip) 
	2 vertical struts (1.5 cm of 1 x 3 mm carbon fiber strip) 
	2 earbuds (4 mm of 4 mm ID/ 6 mm OD carbon fiber cylinder)</li>
	<li>Assemble crossbar. Drill two small holes 4.5 mm apart in the center of the crossbar (Figure 1a, top) using a 1.4 mm diameter bit. Holes should be just large enough to accommodate the 0-80 screw. Insert two 0-80 screws though the holes and attach hex nuts. Cut two 6 mm strips of syringe needle (any small-diameter needle will work) and flatten with pliers (needle strips shown in red in Figure 1a). Turn the crossbar over so the hex nuts lie on top. To prevent epoxy from gluing the tip of the screws to the hex nuts put a drop of mineral oil (shown in green in Figure 1a) on the tip of each screw. Place the two needle segments on top of the hex nuts. Epoxy hex nuts to each other and the needle segments, covering both (Figure 1a, bottom). After epoxy has fully cured (24 hr), remove both screws. The crossbar unit should separate into four components: two screws, the 3 cm length of carbon fiber, and a single piece consisting of 2 hex nuts and 2 needle segments glued together with epoxy. Reassemble crossbar unit.</li>
	<li>Fabricate foam rings. Use a 1/4 in (6.35 mm) hole punch to cut a foam cylinder from a foam earplug. Use scissors to cut perpendicular to the long axis of the foam cylinder yielding a 2 mm (length) by 6.35 mm (diameter) cylinder. Then use a 1/8 in (3.175 mm) hole punch to cut out the center of the 2 mm foam cylinder, forming a foam ring (shown in orange in Figure 1b).</li>
	<li>Assemble earbuds. Use epoxy to glue vertical struts (1.5 cm length of carbon fiber strip) to the 4mm length of carbon fiber cylinder as shown in Figure 1b, bottom. When epoxy has hardened, use a small amount of epoxy to glue the foam ring to the end of the cylinder (Figure 1b, bottom). The foam ring will rest against the bird's head.</li>
</ol>
2. Attach Headphones Frame to Bird
<ol>
	<li>Prepare stereotax. Prior to surgery, insert stereotax earbars through earbuds with the foam pads facing inwards (Figure 1c, top).</li>
	<li>Attach crossbar. Anesthetize the bird using isoflurane and/or ketamine plus midazolam. The anesthetized bird should have closed eyes and be unresponsive to toe pinch. Place bird in stereotax and make a 5-10 mm incision in scalp. The incision should be along the midline and extend anterior from a point 2 mm anterior of the attachment of the neck muscles at the back of the skull. Rub the exposed skull with 70% ethanol to dry the bone surface. Use an alligator clip holder to place crossbar such that the epoxy-covered hex nuts rest on the top of the skull (Figure 1c, d). Lift the crossbar slightly and glue the hex nuts (with epoxy or dental acrylic) to the skull, being careful that the adhesive touches ONLY the hex nuts and not the crossbar itself. This will allow you to remove the headphones (by removing the screws), leaving only the hex nuts permanently attached to the head (Figure 1d).</li>
	<li>Attach earbuds. Slide earbuds along ear bars until foam pads rest against the bird's head (Figure 1c, top). Use alligator clips on the ear bars to press foam pads firmly against the bird's head (Figure 1c, bottom), this is important to getting a good acoustic seal. Rotate earbuds so that the vertical posts touch the back of the crossbar, leaving space for the connector strip socket (Figure 2) in front of the post. Glue posts to crossbar using epoxy or acrylic.</li>
	<li>Recovery. Allow 24 hr for epoxy/acrylic to fully cure and bird to recover. After recovery, examine frames to make sure foam pads form a tight seal around ear canals. If seal is loose, foam pads can be removed and replaced with larger ones to improve acoustic seal. Gently remove screws using a 0.05 in (1.27 mm) hexagonal wrench and remove assembled headphones frame from the bird.</li>
</ol>
3. Assemble Electronics
<ol>
	<li>Fabricate speaker adapters. Make speaker adapters by inserting a pipet tip (a 5.2 cm long tip with a diameter that tapers from 5 mm to 1 mm) into a scrap piece of carbon fiber cylinder (use the same 4 mm ID/ 6 mm OD cyinder stock as for the earbuds; length of the cylinder scrap is not important), then cut pipet tip 1 mm past the end of the cylinder (Figure 2a). Remove from cylinder and cut so that the total length of the adapter is 3 mm.</li>
	<li>Connect speakers. Solder 5 cm lengths of insulated 36 ga. wire to the positive and negative terminals of both speakers (green and black wires, respectively, in Figure 2d, e). Use an ohmmeter to verify that neither terminal is in electrical contact with the speaker's metal casing.</li>
	<li>Assemble speaker/microphone component. Epoxy a strip of tape (any electrically insulating tape will work) to one side of one speaker and epoxy the headphones mic on top of the tape (Figure 2b). Solder the ground wire from the microphone to the negative terminal on the speaker (connection shown in black in Figure 2e).</li>
	<li>Epoxy both the speaker and the speaker/microphone component into the wider end of each adapter (Figure 2c, right). Both the speaker and microphone port will (just) fit inside the pipet. Be sure that epoxy does not cover either port.</li>
	<li>Gently insert adapters into the earbuds (Figure 2d, right). Glue each adapter to its earbud with a small drop of epoxy.</li>
	<li>Fabricate connector strip. Cut a segment of connector strip socket consisting of four terminals and place a reference mark (white dot in Figure 2d) on one corner of the socket. Position the connector strip near one of the vertical posts. Solder the speaker and microphone wires to the pins in the configuration shown in Figure 2e. Use an ohmmeter to check for short circuits on connector strip and repair if necessary. Epoxy the connector strip to the headphones frame. Wrap strips of electrical or scotch tape around exposed wires to protect wires and add thin layer of epoxy over tape ends to prevent unraveling. The finished weight of the headphones (including frame and electronics) should be 1.3-1.5 g.</li>
	<li>Build a flexible lead by soldering four 15 cm lengths of insulated 36 ga. wire to a segment of connector strip header consisting of four terminals (Figure 3a). Place a reference mark (white dot in Figure 3) on one corner of the header to ensure that the lead is plugged into the socket in the correct orientation. Braid wires and finish lead by soldering wires to an adapter that plugs into commutator (Figure 3b).</li>
</ol>
4. Connect Headphones Electronics to Power and Signal Processing Equipment
<ol>
	<li>Collect, filter, pitch shift, and amplify acoustic input. Hang an omnidirectional condenser microphone ("Cage mic," Figure 4) directly above the bird's cage. Preamplify and low-pass filter this signal (cutoff at 10 kHz for finch song) and input it into the Harmonizer (or other realtime sound-processing hardware). Use the pitch shifter module on the Harmonizer to generate a shifted acoustic signal. Amplify this signal and route it (via the commutator and flexible lead) to the positive channel on the speaker headphones (green in Figure 2e).</li>
	<li>Provide power to phones mic. Attach a 9V battery to the wall of the bird's soundbox. Connect the positive terminal of the battery to the V+ channel (red in Figure 2e) via the commutator and flexible lead, connecting the battery's negative terminal to a common ground on the signal processing equipment. Connect this common ground to the speaker and phones mic ground (black in Figure 2e).</li>
	<li>Calibrate feedback gain. Attach finished headphones to the hex nuts on bird's head using 0-80 screws. When bird begins singing, record three channels of data (cage mic/unshifted signal, shifted signal, and phones mic) as shown in Figure 4. Examine the sound recorded from the phones mic. This channel will record the superposition of the virtual feedback (played through the headphones speaker) and the bird's actual song "leaking" directly into the headphones frame. Adjust amplitude setting of speaker amplifier (Figure 4) so that the virtual auditory feedback is 15-20 dB louder than the direct/leaking signal. Note that increasing the amplitude beyond this threshold can result in the virtual signal bleeding through the headphones and being picked up by the cage mic, resulting in feedback distortion. An example of a well-calibrated system (in which the signal recorded by the headphones mic is dominated by the pitch-shifted signal rather than the direct/leaking signal) is shown in Figure 5. Here, the signal recorded through the headphones microphone (Figure 5a, right) is dominated by the pitch-shifted signal (rather than the bird's natural vocalization), as shown in the example power spectrum in Figure 5b.</li>
	<li>Monitor system performance. Check headphones twice daily to insure that both speakers and the microphone are functioning properly. Note that the headphones mic only monitors one of the speakers. To test the other speaker, tap the cage mic while holding the non-monitored speaker next to your ear - you will hear a clicking sound if the non-monitored speaker is working. Another recommended check is to disconnect the power source and signal output from the phone mic and use an ohmmeter to measure the resistance across the speakers (black and green in Figure 2e). When both speakers are connected, this will be half of the resistance across a single speaker since speakers are connected in parallel.</li>
</ol>
5. A Note on the Cost of Materials
With two notable exceptions, all items listed in the Table of Materials are relatively inexpensive (less than a few hundred US dollars). The most costly components are the commutator and the Harmonizer listed in the Table, which each cost 2,000 US dollars or more. We note that less expensive versions of both items may be available from different manufacturers than the ones listed (although we have not tested them) and might allow researchers to implement this protocol at a lower cost.

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F., Ghez, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Independent+learning+of+internal+models+for+kinematic+and+dynamic+control+of+reaching.">Independent learning of internal models for kinematic and dynamic control of reaching.</a> Nature Neuroscience. 2, 1026-1031 (1999).</li><li>Chen, S. H., Liu, H., Xu, Y., Larson, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Voice+F0+responses+to+pitch-shifted+voice+feedback+during+English+speech.">Voice F0 responses to pitch-shifted voice feedback during English speech.</a> J. Acoust. Soc. Am. 121, 1157-1163 (2007).</li><li>Houde, J. F., Jordan, M. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sensorimotor+adaptation+in+speech+production.">Sensorimotor adaptation in speech production.</a> Science. 279, 1213-1216 (1998).</li><li>Jones, J. A., Munhall, K. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perceptual+calibration+of+F0+production:+evidence+from+feedback+perturbation.">Perceptual calibration of F0 production: evidence from feedback perturbation.</a> J. Acoust. Soc. Am. 108, 1246-1251 (2000).</li><li>Liu, H., Larson, C. R. <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+perturbation+magnitude+and+voice+F0+level+on+the+pitch-shift+reflex.">Effects of perturbation magnitude and voice F0 level on the pitch-shift reflex.</a> J. Acoust. Soc. Am. 122, 3671-3677 (2007).</li><li>Doupe, A. J., Kuhl, P. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Birdsong+and+human+speech:+common+themes+and+mechanisms.">Birdsong and human speech: common themes and mechanisms.</a> Annu. Rev. Neurosci. 22, 567-631 (1999).</li><li>Mooney, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+mechanisms+for+learned+birdsong.">Neural mechanisms for learned birdsong.</a> Learn Mem. 16, 655-669 (2009).</li><li>Brainard, M. S., Doupe, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interruption+of+a+basal+ganglia-forebrain+circuit+prevents+plasticity+of+learned+vocalizations.">Interruption of a basal ganglia-forebrain circuit prevents plasticity of learned vocalizations.</a> Nature. 404, 762-766 (2000).</li><li>Chi, Z., Margoliash, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporal+Precision+and+Temporal+Drift+in+Brain+and+Behavior+of+Zebra+Finch+Song.">Temporal Precision and Temporal Drift in Brain and Behavior of Zebra Finch Song.</a> Neuron. 32, 899-910 (2001).</li><li>Sober, S. J., Wohlgemuth, M. J., Brainard, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Central+contributions+to+acoustic+variation+in+birdsong.">Central contributions to acoustic variation in birdsong.</a> J. Neurosci. 28, 10370-10379 (2008).</li><li>Kao, M. H., Doupe, A. J., Brainard, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contributions+of+an+avian+basal+ganglia-forebrain+circuit+to+real-time+modulation+of+song.">Contributions of an avian basal ganglia-forebrain circuit to real-time modulation of song.</a> Nature. 433, 638-643 (2005).</li><li>Sober, S. J., Brainard, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adult+birdsong+is+actively+maintained+by+error+correction.">Adult birdsong is actively maintained by error correction.</a> Nat. Neurosci. 12, 927-931 (2009).</li><li>Choe, C. S., Welch, R. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Variables+affecting+the+intermanual+transfer+and+decay+of+prism+adaptation.">Variables affecting the intermanual transfer and decay of prism adaptation.</a> J. Exp. Psychol. 102, 1076-1084 (1974).</li><li>Tumer, E. C., Brainard, M. 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No conflicts of interest declared.

The protocol presented here allows the experimenter to manipulate auditory feedback in singing birds. The lightweight construction allows such manipulations to be sustained over long periods, and birds will sing prolifically while wearing headphones for a month or more. Although some songbirds will sing for as long as 10 weeks wearing headphones, in some cases the amount of singing begins to decline after ~5 weeks of use. For this reason, we typically limit experiments to 4 weeks. In our experience, every songbird fitted with headphones can be expected to sing 100+ song bouts per day (and sometimes much more). Therefore, if properly employed, the headphones system offers a nearly 100% success rate (if success is defined by the acquisition of data from singing birds). Furthermore, after completing one learning experiment the headphones can be removed and subsequently reattached for further data collection. Provided that the animal is in good general health reattachment can take place at any time.
One important determinant of success is minimizing the weight and optimizing the comfort of the headphones. During construction, care should be taken to minimize the amount of epoxy or dental acrylic used as excess adhesive will increase the overall weight of the apparatus and potentially reduce the bird's willingness to sing. Additionally, several days after attaching the headphones, the apparatus should be briefly removed to verify that the skin around the ear canals has not become irritated by the earbuds, which can occur if the earbuds are too tight. The ear canals should appear just as they did at the time of headphones attachment (open and with no signs of redness or swelling). If irritation occurs, pressure can be alleviated by reducing the thickness of the foam pads. Take care to ensure that foam hardened by dried epoxy does not contact the bird's skin, as this will also cause irritation.
It is important to note that in addition to the pitch shift selected by the experimenter, virtual auditory feedback is also delayed (by ~10 msec, reflecting the processing latency of the Harmonizer) and is introduced at a greater amplitude than the bird's natural auditory feedback (in order to drown out the sound of the bird's natural song "leaking" into the headphones). For this reason, experiments should begin with a baseline period of several days in which the bird sings with the headphones on but with zero pitch shift14, allowing the effect of the pitch shift to be isolated from vocal changes resulting from other factors related to the headphones paradigm. In practice, changes in song pitch or amplitude are seldom observed when birds first begin to sing with headphones in the absence of a pitch shift. Furthermore, we have shown that extended exposure to unshifted feedback delivered via headphones does not cause a change in song pitch14.
We have previously used this design to demonstrate that in adult songbirds, both upward and downward shifts in the pitch of auditory feedback generate adaptive changes in vocal pitch (i.e. changes opposite in sign to the feedback shift) 14. Including both upward and downward shifts in any experiment employing this paradigm is important because such a design can demonstrate that song pitch changes in response to changes in the pitch of auditory feedback (and not in response to the delay or amplitude artifacts introduced by the headphones). Additionally, a key strength of this paradigm is that it can be used to introduce arbitrary auditory manipulations. The Harmonizer system can generate a wide variety of online perturbations, for example by altering the amplitude or spectral envelope of the acoustic signal. Expanding the range of manipulations beyond pitch shifts could therefore be used to examine a variety of vocal learning phenomena. Additionally, the headphones could be used to deliver white noise or other conditional reinforcement signals to drive learning in individual syllables16. Finally, this paradigm could in principle be employed in any small animal system that relies on auditory feedback during vocal behavior.
We note that our technique, which mimics auditory feedback manipulations used to study human speech 4-7, allows vocal plasticity to be investigated in a physiologically accessible animal model. Combining behavioral studies of vocal error correction with brain lesions, pharmacological manipulations, or neural recordings could be used to reveal how particular neural circuits contribute to the correction of errors in vocal performance.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Hex nuts</td>
 <td>Amazon supply</td>
 <td>B000FMW43Y</td>
 <td></td>
 </tr>
 <tr>
 <td>0-80 Screws, 1/8"</td>
 <td>Amazon supply</td>
 <td>B000FN0JXK</td>
 <td></td>
 </tr>
 <tr>
 <td>0.05" Hex wrench</td>
 <td>Amazon supply</td>
 <td>B003GDISE8</td>
 <td></td>
 </tr>
 <tr>
 <td>Headphones speakers</td>
 <td>Digikey</td>
 <td>423-1113-ND</td>
 <td></td>
 </tr>
 <tr>
 <td>Headphones microphone</td>
 <td>Digikey</td>
 <td>423-1062-ND</td>
 <td></td>
 </tr>
 <tr>
 <td>Harmonizer</td>
 <td>Sweetwater Sound</td>
 <td>H7600</td>
 <td></td>
 </tr>
 <tr>
 <td>(Eventide H7600)</td>
		<td></td>
		<td></td>
		<td></td>
 </tr>
 <tr>
 <td>Carbon fiber strip, 1 x 3 mm</td>
 <td>Hobby Lobby International</td>
 <td>GXS1030</td>
 <td></td>
 </tr>
 <tr>
 <td>Carbon fiber cylinder, 6 mm (OD) x 4 mm (ID)</td>
 <td>Hobby Lobby International</td>
 <td>GXT6040</td>
 <td></td>
 </tr>
 <tr>
 <td>Wire</td>
 <td>Cooner Wire &amp; Cable</td>
 <td>NUF36-2550</td>
 <td></td>
 </tr>
 <tr>
 <td>Connector strip header</td>
 <td>Digikey</td>
 <td>ED83100-ND</td>
 <td></td>
 </tr>
 <tr>
 <td>Connector strip socket</td>
 <td>Digikey</td>
 <td>ED85100-ND</td>
 <td></td>
 </tr>
 <tr>
 <td>Foam earplugs</td>
 <td>AO SAFETY</td>
 <td>92050</td>
 <td></td>
 </tr>
 <tr>
 <td>1/8" hole punch</td>
 <td>Paperwishes</td>
 <td>7260197000</td>
 <td></td>
 </tr>
 <tr>
 <td>1/4" hole punch</td>
 <td>Paperwishes</td>
 <td>7260198000</td>
 <td></td>
 </tr>
 <tr>
 <td>Pipet tips</td>
 <td>VWR</td>
 <td>89003-056</td>
 <td></td>
 </tr>
 <tr>
 <td>Dental acrylic</td>
 <td>Maxcem</td>
 <td>33873</td>
 <td></td>
 </tr>
 <tr>
 <td>5-minute epoxy</td>
 <td>Devcon</td>
 <td>14210</td>
 <td></td>
 </tr>
 <tr>
 <td>Cage microphone</td>
 <td>Countryman</td>
 <td>B3P4FF05B</td>
 <td></td>
 </tr>
 <tr>
 <td>Microphone preamp</td>
 <td>M-audio</td>
 <td>DMP3</td>
 <td></td>
 </tr>
 <tr>
 <td>Speaker amplifier (Crown D-45)</td>
 <td>Sweetwater sound</td>
 <td>D-45</td>
 <td></td>
 </tr>
 <tr>
 <td>Low-pass filter</td>
 <td>Krohn-hite</td>
 <td>FMB3002AC, 3FS8SL-10kg-N1U1</td>
 <td></td>
 </tr>
 <tr>
 <td>Commutator</td>
 <td>Dragonfly</td>
 <td>SL-88-10</td>
 <td></td>
 </tr>
 <tr>
 <td>Alligator clip holder</td>
 <td>GC Electronics</td>
 <td>12-051</td>
 <td></td>
 </tr>
 <tr>
 <td>Mineral oil</td>
 <td>Sigma</td>
 <td>M3516</td>
 <td></td>
 </tr>
 <tr>
 <td>Dremel tool</td>
 <td>Dremel</td>
 <td>8200</td>
 <td></td>
 </tr>
 </tbody>
 </table>

a lightweight, headphones-based system for manipulating auditory feedback in songbirds

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and sensorimotor plasticity1. Consistent sensory perturbations result in compensatory changes in motor output, reflecting changes in feedforward motor control that reduce the experienced feedback error. By quantifying how different sensory feedback errors affect human behavior, prior studies have explored how visual signals are used to recalibrate arm movements2,3 and auditory feedback is used to modify speech production4-7. The strength of this approach rests on the ability to mimic naturalistic errors in behavior, allowing the experimenter to observe how experienced errors in production are used to recalibrate motor output.
Songbirds provide an excellent animal model for investigating the neural basis of sensorimotor control and plasticity8,9. The songbird brain provides a well-defined circuit in which the areas necessary for song learning are spatially separated from those required for song production, and neural recording and lesion studies have made significant advances in understanding how different brain areas contribute to vocal behavior9-12. However, the lack of a naturalistic error-correction paradigm - in which a known acoustic parameter is perturbed by the experimenter and then corrected by the songbird - has made it difficult to understand the computations underlying vocal learning or how different elements of the neural circuit contribute to the correction of vocal errors13.
The technique described here gives the experimenter precise control over auditory feedback errors in singing birds, allowing the introduction of arbitrary sensory errors that can be used to drive vocal learning. Online sound-processing equipment is used to introduce a known perturbation to the acoustics of song, and a miniaturized headphones apparatus is used to replace a songbird's natural auditory feedback with the perturbed signal in real time. We have used this paradigm to perturb the fundamental frequency (pitch) of auditory feedback in adult songbirds, providing the first demonstration that adult birds maintain vocal performance using error correction14. The present protocol can be used to implement a wide range of sensory feedback perturbations (including but not limited to pitch shifts) to investigate the computational and neurophysiological basis of vocal learning.

Figure 6 shows a representative experiment performed on an adult Bengalese finch. Here, the headphones system was used to increase the pitch of auditory feedback by one semitone (one twelfth of an octave, representing an approximately 6% change in absolute frequency) for 16 days. This manipulation resulted in a gradual reduction of the pitch of all song syllables (colored lines). This change in the vocal motor program resulted in a reduction in the auditory error experienced by the bird (dashed line), demonstrating the bird's reliance on auditory feedback to correct apparent vocal errors. When the pitch shift was removed after day 16, the pitch of song eventually returned to baseline.
The data shown in Figure 6 are typical in that they reflect incomplete adaptation. Here, although the pitch of auditory feedback was shifted by 1.0 semitones, the bird changed the pitch of his song by only about 0.4 semitones. Across species and systems, incomplete adaptation is the norm when virtual feedback is used to perturb a single sensory modality 5,15, and in the present paradigm likely reflects a partial reliance on nonauditory (e.g. proprioceptive) signals as songbirds evaluate their ongoing vocal performance.
<img alt="Figure 1" src="/files/ftp_upload/50027/50027fig1.jpg" /> 
Figure 1. Headphones frame assembly. a. Crossbar assembly. Attach screws, crossbar, hex nuts, and syringe needles (red) using epoxy adhesive (blue) as shown. Cover the tip of each screw with mineral oil (green) to prevent epoxy from bonding to screws. b. Earbud assembly. Attach post, cylinder, and foam pad (orange) with epoxy. c. Fitting headphones frame. Prior to surgery, thread stereotax earbars (black) through earbuds. Attach crossbar to skull using epoxy or dental acrylic (blue), then attach earbuds to crossbar using alligator clips to gently press the foam pads against the bird's head. d. Left, side view illustrating positioning of crossbar and earbuds on bird's head. Right, side view of the hex nut assembly attached to the skull after the headphones frame has been removed by detaching the screws. Scale bar in b pertains to all panels.
<img alt="Figure 2" src="/files/ftp_upload/50027/50027fig2.jpg" /> 
Figure 2. Electronics assembly. a. Make an adapter by inserting a pipet tip into a scrap piece of carbon fiber cylinder and cutting pipet tip to length. b. Using epoxy, glue one speaker and headphones mic together separated by a piece of tape. c. Glue adapter onto speaker (shown) and speaker/phones mic component. d. Solder wires from speakers and headphones mic to a connector strip socket and glue the socket to the top of the headphones frame. White dot, alignment mark on connector strip socket. e. Wiring diagram showing connections between speakers, headphones mic, and socket.
<img alt="Figure 3" src="/files/ftp_upload/50027/50027fig3.jpg" /> 
Figure 3. Lead assembly. a. Fabricate a flexible lead by soldering four 15 cm lengths of wire to the pins on one side of a connector strip header and braid wires together. Solder the other end of the lead wires to an adapter connecting to commutator. White dot, alignment mark on connector strip header. b. Plug lead into connector strip socket on headphones to carry altered auditory feedback to headphones speaker, power the headphones mic, and record the signals from the headphones mic. Align dots on connector strip header and socket to ensure correct connectivity.
<img alt="Figure 4" src="/files/ftp_upload/50027/50027fig4.jpg" /> 
Figure 4. Circuit summary. Flowchart summarizing system connectivity. The three data channels record (1) the unshifted signal from the cage microphone, (2) a copy of the pitch-shifted signal sent to the headphones speakers, and (3) the sound waveform recorded by the headphones mic. Preamp, microphone preamplifier; LPF, low-pass filter.
<img alt="Figure 5" src="/files/ftp_upload/50027/50027fig5.jpg" /> 
Figure 5. Testing the system. a. Spectrograms of sound recorded on the three data channels. Each data channel shows three song syllables. Color represents power at each time and acoustic frequency. b. Power spectrum at the time indicated by the vertical red lines in a. Note that the peaks in the headphones mic spectrum (green) match the peaks in the pitch-shifted (red) rather than the unshifted (black) signal, indicating that the sound reaching the bird's ear is dominated by the shifted feedback.
<img alt="Figure 6" src="/files/ftp_upload/50027/50027fig6.jpg" /> 
Figure 6. Using the headphones system to drive vocal learning. Vocal error correction in an adult Bengalese finch. Colored lines show changes in the pitch of seven different song syllables during a 16-day period (gray box) in which the headphones system was used to shift the pitch of auditory feedback upwards by one semitone. Solid black line, mean pitch change across all song syllables. Dashed black line shows the mean pitch error experienced by the subject during the shift epoch. Note that the change in song pitch serves to reduce the experienced pitch error. After the pitch shift was set back to zero on day 17 pitch approaches baseline. On day 24 the headphones were removed, and then replaced on day 46, at which time pitch had recovered back to its baseline value.

Watch this Scientific Journal Video about A Lightweight, Headphones-based System for Manipulating Auditory Feedback in Songbirds at JoVE.com

A Lightweight, Headphones-based System for Manipulating Auditory Feedback in Songbirds

We describe the design and assembly of miniaturized headphones suitable for replacing a songbird&#x2019;s natural auditory feedback with a manipulated acoustic signal. Online sound processing hardware is used to manipulate song output, introduce real-time errors in auditory feedback via the headphones, and drive vocal motor learning.

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and ...

a-lightweight-headphones-based-system-for-manipulating-auditory

Research

JoVE Journal

Neuroscience

14.2K ビュー.  Emory University. 我々は、操られた音響信号と小鳥の自然聴覚フィードバックを交換するのに適した小型化されたヘッドフォンの設計と組立を説明します。オンライン音声処理ハードウェアは、歌の出力を操作するヘッドフォンを介して聴覚フィードバックにおけるリアルタイムエラーを導入すると、ボーカルの運動学習を駆動するために使用される。

ビデオ: A Lightweight, Headphones-based System for Manipulating Auditory Feedback in Songbirds

Efficient Derivation of Human Neuronal Progenitors and Neurons from Pluripotent Human Embryonic Stem Cells with Small Molecule Induction

There is a large unfulfilled need for a clinically-suitable human neuronal cell source for repair or regeneration of the damaged central nervous system (CNS) structure and circuitry in today's healthcare industry. Cell-based therapies hold great promise to restore the lost nerve tissue and function for CNS disorders. However, cell therapies based on CNS-derived neural stem cells have encountered supply restriction and difficulty to use in the clinical setting due to their limited expansion ability in culture and failing plasticity after extensive passaging1-3. Despite some beneficial outcomes, the CNS-derived human neural stem cells (hNSCs) appear to exert their therapeutic effects primarily by their non-neuronal progenies through producing trophic and neuroprotective molecules to rescue the endogenous cells1-3. Alternatively, pluripotent human embryonic stem cells (hESCs) proffer cures for a wide range of neurological disorders by supplying the diversity of human neuronal cell types in the developing CNS for regeneration1,4-7. However, how to channel the wide differentiation potential of pluripotent hESCs efficiently and predictably to a desired phenotype has been a major challenge for both developmental study and clinical translation. Conventional approaches rely on multi-lineage inclination of pluripotent cells through spontaneous germ layer differentiation, resulting in inefficient and uncontrollable lineage-commitment that is often followed by phenotypic heterogeneity and instability, hence, a high risk of tumorigenicity7-10. In addition, undefined foreign/animal biological supplements and/or feeders that have typically been used for the isolation, expansion, and differentiation of hESCs may make direct use of such cell-specialized grafts in patients problematic11-13. To overcome these obstacles, we have resolved the elements of a defined culture system necessary and sufficient for sustaining the epiblast pluripotence of hESCs, serving as a platform for de novo derivation of clinically-suitable hESCs and effectively directing such hESCs uniformly towards clinically-relevant lineages by small molecules14 (please see a schematic in Fig. 1). Retinoic acid (RA) does not induce neuronal differentiation of undifferentiated hESCs maintained on feeders1, 14. And unlike mouse ESCs, treating hESC-differentiated embryoid bodies (EBs) only slightly increases the low yield of neurons1, 14, 15. However, after screening a variety of small molecules and growth factors, we found that such defined conditions rendered retinoic acid (RA) sufficient to induce the specification of neuroectoderm direct from pluripotent hESCs that further progressed to neuroblasts that generated human neuronal progenitors and neurons in the developing CNS with high efficiency (Fig. 2). We defined conditions for induction of neuroblasts direct from pluripotent hESCs without an intervening multi-lineage embryoid body stage, enabling well-controlled efficient derivation of a large supply of human neuronal cells across the spectrum of developmental stages for cell-based therapeutics.

We have established a protocol for induction of neuroblasts direct from pluripotent human embryonic stem cells maintained under defined conditions with small molecules, which enables derivation of a large supply of human neuronal progenitors and neuronal cell types in the developing CNS for neural repair.

There is a large unfulfilled need for a clinically-suitable human neuronal cell source for repair or regeneration of the damaged central nervous system ...

A Fully Automated and Highly Versatile System for Testing Multi-cognitive Functions and Recording Neuronal Activities in Rodents

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be investigated in a single task sequence. The unique feature of this system is a custom-made, acoustically transparent chamber that eliminates many of the issues associated with auditory cue control in most commercially available chambers. The ease with which operant devices can be added or replaced makes this system quite versatile, allowing for the implementation of a variety of auditory, visual, and olfactory behavioral tasks. Automation of the system allows fine temporal (10 ms) control and precise time-stamping of each event in a predesigned behavioral sequence. When combined with a multi-channel electrophysiology recording system, multiple cognitive brain functions, such as motivation, attention, decision-making, patience, and rewards, can be examined sequentially or independently.

In this report, we present a fully automated and highly versatile system capable of simultaneously testing multi-cognitive behaviors and recording neuronal activities for rodents.

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be ...

Functional Imaging of Auditory Cortex in Adult Cats using High-field fMRI

Current knowledge of sensory processing in the mammalian auditory system is mainly derived from electrophysiological studies in a variety of animal models, including monkeys, ferrets, bats, rodents, and cats. In order to draw suitable parallels between human and animal models of auditory function, it is important to establish a bridge between human functional imaging studies and animal electrophysiological studies. Functional magnetic resonance imaging (fMRI) is an established, minimally invasive method of measuring broad patterns of hemodynamic activity across different regions of the cerebral cortex. This technique is widely used to probe sensory function in the human brain, is a useful tool in linking studies of auditory processing in both humans and animals and has been successfully used to investigate auditory function in monkeys and rodents. The following protocol describes an experimental procedure for investigating auditory function in anesthetized adult cats by measuring stimulus-evoked hemodynamic changes in auditory cortex using fMRI. This method facilitates comparison of the hemodynamic responses across different models of auditory function thus leading to a better understanding of species-independent features of the mammalian auditory cortex.

Functional studies of the auditory system in mammals have traditionally been conducted using spatially-focused techniques such as electrophysiological recordings. The following protocol describes a method of visualizing large-scale patterns of evoked hemodynamic activity in the cat auditory cortex using functional magnetic resonance imaging.

Current knowledge of sensory processing in the mammalian auditory system is mainly derived from electrophysiological studies in a variety of animal ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Optogenetic Stimulation of the Auditory Nerve

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of&#160;the cochlea has been used to evoke responses in the auditory nerve10. Nevertheless it requires higher energies than electrical stimulation10,11 and uncertainty remains as to the underlying mechanism12. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13 or virus-mediated expression of the ChR2-variant CatCh14. We used micro-light emitting diodes (&#181;LEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.

Cochlear implants (CIs) enable hearing by direct electrical stimulation of the auditory nerve. However, poor frequency and intensity resolution limits the quality of hearing with CIs. Here we describe optogenetic stimulation of the auditory nerve in mice as an alternative strategy for auditory research and developing future CIs.

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted ...

Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and awake animals. Therefore, methods allowing the manipulation of synaptic systems in awake animals are required in order to determine the contribution of synaptic inputs to neuronal processing unaffected by anesthetics. Here, we present methodology for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. By combining these procedures, we performed microiontophoretic injections of gabazine to selectively block GABAA receptors in neurons of the inferior colliculus of head-restrained mice. Gabazine successfully modified neural response properties such as the frequency response area and stimulus-specific adaptation. Thus, we demonstrate that our methods are suitable for recording single-unit activity and for dissecting the role of specific neurotransmitter receptors in auditory processing.
The main limitation of the described procedure is the relatively short recording time (~3 hr), which is determined by the level of habituation of the animal to the recording sessions. On the other hand, multiple recording sessions can be performed in the same animal. The advantage of this technique over other experimental procedures used to manipulate the level of neurotransmission or neuromodulation (such as systemic injections or the use of optogenetic models), is that the drug effect is confined to the local synaptic inputs to the target neuron. In addition, the custom-manufacture of electrodes allows adjustment of specific parameters according to the neural structure and type of neuron of interest (such as the tip resistance for improving the signal-to-noise ratio of the recordings).

We present methods for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. This technique allows the detailed analysis of putative local synaptic inputs to the neuron of interest.

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and ...

Measuring and Manipulating Functionally Specific Neural Pathways in the Human Motor System with Transcranial Magnetic Stimulation

Understanding interactions between brain areas is important for the study of goal-directed behavior. Functional neuroimaging of brain connectivity has provided important insights into fundamental processes of the brain like cognition, learning, and motor control. However, this approach cannot provide causal evidence for the involvement of brain areas of interest. Transcranial magnetic stimulation (TMS) is a powerful, noninvasive tool for studying the human brain that can overcome this limitation by transiently modifying brain activity. Here, we highlight recent advances using a paired-pulse, dual-site TMS method with two coils that causally probes cortico-cortical interactions in the human motor system during different task contexts. Additionally, we describe a dual-site TMS protocol based on cortical paired associative stimulation (cPAS) that transiently enhances synaptic efficiency in two interconnected brain areas by applying repeated pairs of cortical stimuli with two coils. These methods can provide a better understanding of the mechanisms underlying cognitive-motor function as well as a new perspective on manipulating specific neural pathways in a targeted fashion to modulate brain circuits and improve behavior. This approach may prove to be an effective tool to develop more sophisticated models of brain-behavior relations and improve diagnosis and treatment of many neurological and psychiatric disorders.

This article describes new approaches to measure and strengthen functionally specific neural pathways with transcranial magnetic stimulation. These advanced noninvasive brain stimulation methodologies can provide new opportunities for the understanding of brain-behavior relations and development of new therapies to treat brain disorders.

Understanding interactions between brain areas is important for the study of goal-directed behavior. Functional neuroimaging of brain connectivity has ...

Applying the RatWalker System for Gait Analysis in a Genetic Rat Model of Parkinson's Disease

Parkinson's disease (PD) is a progressive neurodegenerative disorder caused by the loss of dopaminergic (DA) neurons in the substantia nigra pars compacta. Gait abnormalities, including decreased arm swing, slower walking speed, and shorter steps are common in PD patients and appear early in the course of disease. Thus, the quantification of motor patterns in animal models of PD will be important for phenotypic characterization during disease course and upon therapeutic treatment. Most cases of PD are idiopathic; however, the identification of hereditary forms of PD uncovered gene mutations and variants, such as loss-of-function mutations in Pink1 and Parkin, two proteins involved in mitochondrial quality control that could be harnessed to create animal models. While mice are resistant to neurodegeneration upon loss of Pink1 and Parkin (single and combined deletion), in rats, Pink1 but not Parkin deficiency leads to nigral DA neuron loss and motor impairment. Here, we report the utility of FTIR imaging to uncover gait changes in freely walking young (2 months of age) male rats with combined loss of Pink1 and Parkin prior to the development of gross visually apparent motor abnormality as these rats age (observed at 4-6 months), characterized by hindlimb dragging as previously reported in Pink1 knockout (KO) rats.

Applying the RatWalker System for Gait Analysis in a Genetic Rat Model of Parkinson&#039;s Disease

Here we describe the RatWalker system, built by redesigning the MouseWalker apparatus to accommodate for the increased size and weight of rats. This system uses frustrated total internal reflection (FTIR), high-speed video capture, and open-access analysis software to track and quantify gait parameters.

Parkinson's disease (PD) is a progressive neurodegenerative disorder caused by the loss of dopaminergic (DA) neurons in the substantia nigra pars ...

Stereocilia Bundle Imaging with Nanoscale Resolution in Live Mammalian Auditory Hair Cells

Inner ear hair cells detect sound-induced displacements and transduce these stimuli into electrical signals in a hair bundle that consists of stereocilia that are arranged in rows of increasing height. When stereocilia are deflected, they tug on tiny (~5 nm in diameter) extracellular tip links interconnecting stereocilia, which convey forces to the mechanosensitive transduction channels. Although mechanotransduction has been studied in live hair cells for decades, the functionally important ultrastructural details of the mechanotransduction machinery at the tips of stereocilia (such as tip link dynamics or transduction-dependent stereocilia remodeling) can still be studied only in dead cells with electron microscopy. Theoretically, scanning probe techniques, such as atomic force microscopy, have enough resolution to visualize the surface of stereocilia. However, independent of imaging mode, even the slightest contact of the atomic force microscopy probe with the stereocilia bundle usually damages the bundle. Here we present a detailed protocol for the hopping probe ion conductance microscopy (HPICM) imaging of live rodent auditory hair cells. This non-contact scanning probe technique allows time lapse imaging of the surface of live cells with a complex topography, like hair cells, with single nanometers resolution and without making physical contact with the sample. The HPICM uses an electrical current passing through the glass nanopipette to detect the cell surface in close vicinity to the pipette, while a 3D-positioning piezoelectric system scans the surface and generates its image. With HPICM, we were able to image stereocilia bundles and the links interconnecting stereocilia in live auditory hair cells for several hours without noticeable damage. We anticipate that the use of HPICM will allow direct exploration of ultrastructural changes in the stereocilia of live hair cells for better understanding of their function.

Here we present a protocol for the Hopping Probe Ion Conductance Microscopy (HPICM), a non-contact scanning probe technique that allows nanoscale imaging of stereocilia bundles in live auditory hair cells.

Inner ear hair cells detect sound-induced displacements and transduce these stimuli into electrical signals in a hair bundle that consists of stereocilia ...

Evaluation of Auditory Brainstem Response in Chicken Hatchlings

The auditory brainstem response (ABR) is an invaluable assay in clinical audiology, non-human animals, and human research. Despite the widespread use of ABRs in measuring auditory neural synchrony and estimating hearing sensitivity in other vertebrate model systems, methods for recording ABRs in the chicken have not been reported in nearly four decades. Chickens provide a robust animal research model because their auditory system is near functional maturation during late embryonic and early hatchling stages. We have demonstrated methods used to elicit one or two-channel ABR recordings using subdermal needle electrode arrays in chicken hatchlings. Regardless of electrode recording configuration (i.e., montage), ABR recordings included 3-4 positive-going peak waveforms within the first 6 ms of a suprathreshold click stimulus. Peak-to-trough waveform amplitudes ranged from 2-11 &#181;V at high-intensity levels, with positive peaks exhibiting expected latency-intensity functions (i.e., increase in latency as a function of decreased intensity). Standardized earphone position was critical for optimal recordings as loose skin can occlude the ear canal, and animal movement can dislodge the stimulus transducer. Peak amplitudes were smaller, and latencies were longer as animal body temperature lowered, supporting the need for maintaining physiological body temperature. For young hatchlings (&#60;3 h post-hatch day 1), thresholds were elevated by ~5 dB, peak latencies increased ~1-2 ms, and peak to trough amplitudes were decreased ~1 &#181;V compared to older hatchlings. This suggests a potential conductive-related issue (i.e., fluid in the middle ear cavity) and should be considered for young hatchlings. Overall, the ABR methods outlined here permit accurate and reproducible recording of in-vivo auditory function in chicken hatchlings that could be applied to different stages of development. Such findings are easily compared to human and mammalian models of hearing loss, aging, or other auditory-related manipulations.

We have used standard auditory brainstem response (ABR) techniques and applied them to hatchling chickens, a precocious avian model for auditory function. The protocol outlines animal preparation and ABR acquisition techniques in detail, with steps that could translate to other avian or rodent models.

The auditory brainstem response (ABR) is an invaluable assay in clinical audiology, non-human animals, and human research. Despite the widespread use of ...