This work was supported by the German Federal Ministry of Education and Research (Bernstein Focus for Neurotechnology grant 01GQ0810, to T. Moser, and MED-EL Germany); the German Research Foundation through the Center for Nanoscale Microscopy and Molecular Physiology of the Brain (FZT 103, T. Moser) and through the SFB889, to N. Strenzke and T. Moser).

All experiments presented in this work were conducted with the ethical standards defined by the German law for the protection of experimental animals. The University of Goettingen board for animal welfare and the animal welfare office of the state of Lower Saxony approved the experiments.
1. Preparation of &#181;LED-stimulator
<ol>
	<li>For &#181;LEDs, first prepare the &#181;LED-stimulator. Use blue LEDs with 200 by 200 &#956;m active surface (&#181;LED, see Materials Table).</li>
	<li>Solder wires to the &#181;LED. Then, encapsulate the &#956;LED and the connections using epoxy glue. Leave the device overnight to let the epoxy cure.</li>
	<li>For mechanical stability and electrical insulation funnel the wires through a thin glass capillary, use 1 mm outer diameter borosilicate capillaries and seal the junction with epoxy (Figure 1D) such that the capillary can be mounted on a mechanical manipulator. 
	NOTE: There is a trade-off of making the capsule thick enough to get a good electrical isolation (avoiding stimulation artifacts), but thin enough to let the LED move freely inside the bulla and to locate it onto the cochleostomy. It is a good practice to prepare more than one LED.</li>
	<li>To confirm good electrical isolation, first prepare a Petri dish with 0.9% NaCl and dip 3&#160;bare ended copper wires into the liquid. This serves to simulate an animal with ABR electrodes. Connect the wires to the ABR-amplifier and from there to an oscilloscope.</li>
	<li>Then, connect the LED to the stimulator and immerse it into the solution until it is totally covered. Drive the LED with pulses (see step 3.1.1) at the maximum allowed current for longer than 20 min and check for stimulation artifacts. If they appear, discard the LED and try a new one.</li>
</ol>
2. Surgical Procedure
<ol>
	<li>Use 4-10 week-old transgenic mice expressing ChR2 under the Thy1.2 promoter (line 9 in Wang&#160;et al.13). To manipulate the animal and perform&#160;the surgery, always use clean and well identified surgical tools.</li>
	<li>Anesthetize animals with an i.p. injection of a mixture of urethane (1.32 mg/kg), xylazine (5 mg/kg), and buprenorphine 0.1 &#181;g/g diluted in 0.9% sterile NaCl solution and place the mouse onto a heating plate for maintaining body temperature at 37 &#176;C. With this dose, anesthesia typically lasts for the whole experimental protocol.
	<ol>
		<li>Beginning 10 min after initial anesthesia induction check for signs of arousal by the paw withdrawal reflex.&#160;In case of movements, apply maintenance doses of urethane (.66 mg/kg) and buprenorphine (0.05 &#181;g/g) diluted in 0.9% sterile NaCl solution (without Xylazine).&#160;Again check the paw withdrawal reflex and proceed only if the reflex is absent.</li>
	</ol>
	</li>
	<li>Carefully shave&#160;the retroauricular area in preparation of the incision. Use a retroauricular approach to reach the bulla (air-containing middle ear cavity).
	<ol>
		<li>With a micro forceps carefully dissect and/or remove part of the neck muscles, e.g., platysma, sternocleidomastoid, and scalenes.</li>
		<li>Find and expose the bulla (Figure 1A). Identify the bulla as a bony structure with a characteristic spherical shape with a ring on the bulla surface that indicates the insertion of the tympanic membrane (Figure 1A). Make a hole with the tip of an insulin needle just below this ring. Start there, remove the bone that covers the bulla with a fine Rongeur (gnawer) or microforceps in such a way that the promontorium of the cochlea is exposed (Figure 1B).</li>
	</ol>
	</li>
	<li>For the cochleostomy, gently shave off the bony capsule to open a small window (cochleostomy: ~500-800 &#956;m), taking care to leave intact the membranous labyrinth (Figure 1B). Perform the cochleostomy on the second cochlear turn as it is far enough from the stapedial artery and from the apex. Opening the apex could produce rupture of the cochlea including a fracture of the modiolus.</li>
	<li>For a round window insertion of an optical fiber or &#181;LED-implant to scala tympani, carefully enlarge the round window niche by scratching the bony niche with the tip of a sharp scalpel (a blade number 11 is a good option). Always use a new blade or scalpel. Be very careful as the stapedial artery runs very close to the round window (Figure 1B).</li>
</ol>
3. Optical Stimulation Using Two Approaches
<ol>
	<li>For transcochlear stimulation, use &#181;LEDs or a fiber coupled laser.
	<ol>
		<li>Mount the capillary carrying the &#181;LED on a mechanical manipulator and carefully position the &#181;LED onto the cochleostomy. Use oABR, elicited by 3-10 msec long stimulation from a current source (typically 5-40 mA at 2.7 V) at 1-5 Hz, as a means to optimize the position and orientation of the &#181;LED.</li>
		<li>For laser stimulation, use a 473 nm continuous wave laser and a 250 &#181;m optical fiber (see Materials). Couple the optical fiber to a blue laser source that ideally offers fast analogue control of power (else use an acousto-optical device or a fast shutter to define the optical stimulus). Using a micromanipulator, locate the tip of the fiber directly onto the cochleostomy and fix it there using dental cement (transcochlear stimulation).</li>
	</ol>
	</li>
	<li>For intracochlear stimulation, insert the fiber through the round window into the scala tympani. Then use 3-10 msec laser pulses of 10-30 mW (power of fiber output) at 1-5 Hz to elicit oABR. The large amplitude of oABR enables the experimenter to observe oABR evoked by single stimuli on the oscilloscope for optimizing the position and orientation of the emitter (&#181;LED or fiber) using oABR amplitude as a read-out.</li>
	<li>Once good oABR have been achieved, fix the fiber with cyanoacrylate glue or dental cement and wait until the glue is solid. 
	NOTE: It is normal to observe some liquid coming out from the cochlea. However to avoid problems with the polymerization of the cement, it is advisable to dry up the region using fine cotton wicks.</li>
</ol>
4. Recordings of oABR
<ol>
	<li>Insert needle electrodes underneath the pinna, on the vertex and on the back near the legs and connect them to the amplifier.</li>
	<li>Amplify the difference potential between vertex and mastoid subdermal needles. Sample at a rate of 50 kHz and filter (1-10,000 Hz). Visualize the difference potential on an oscilloscope ideally near the recording setup for optimizing the position of the optical stimulator. Use signal averaging (50-1,000x) and store the data on a computer for offline analysis.</li>
</ol>
5. Recordings of Optically Evoked Local Field Potentials
<ol>
	<li>Place the animal into a stereotactic frame. Using forceps centrally pull the skin overlying the cranium and perform a median incision with scissors roughly extending from between the eyes to the point where the neck muscles attach to the skull.
	<ol>
		<li>Reflect the skin laterally. Cut the periosteum lengthwise along the sagittal suture and remove it sideward by gently rubbing the exposed cranial bone with a cotton swab.</li>
	</ol>
	</li>
	<li>Position a custom made metal plate onto the exposed skull leaving approximately 1 mm space between the caudal end of the plate and lambda. With a 0.7 mm diameter drill carefully drill a hole into the parietal bone.</li>
	<li>Gently screw in a 0.85 mm diameter screw &#8211; to serve as reference for potentials recorded from the inferior colliculus (IC) &#8211; holding the screw with forceps as long as the screw is not securely fastened. Ensure good contact between screw and dura without penetrating the brain. Glue the screw and the metal plate onto the skull with cyanoacrylate glue.</li>
	<li>Loosen the neck muscles gently from the skull. Pull the neck muscles caudally for circa 1-2 mm. Identify the intersection of the superior sagittal sinus (SSS) and the transverse sinus (TS). If necessary, increase the transparency of the skull by moistening the bone with physiological NaCl solution.
	<ol>
		<li>Identify the IC in mice as lying directly caudal to the SSS and TS intersection. After identification of the location of the IC slowly and carefully perform a trepanation extending ca. 0.5 mm rostrally to 1.5 mm caudally to the TS and roughly 3 mm laterally.</li>
		<li>Avoid any injury to the SSS and TS as these would preclude a successful experiment. Here, test whether the trepanned bone has become loose by softly pushing it. Using forceps slowly lift off the bone.</li>
	</ol>
	</li>
	<li>With an appropriate adaptor attach a multichannel array and headstage onto a micromanipulator. To prevent destroying the multichannel recording array during insertion into the IC, carefully remove the dura with a bent tip of a small hypodermic needle starting close to a bone edge.
	<ol>
		<li>Remove the animal from the stereotaxic frame. Fix the metal plate onto a bar to remain stereotaxic fixation. Position the recording probe over the IC and insert it under microscopic control such that the top most channel is just visible at the surface.</li>
	</ol>
	</li>
	<li>Amplify the difference potential between the individual channels of the recording array and the reference screw in the parietal bone, sample at a rate of 32 kHz, low-pass filter (300 Hz) and store on a computer for offline analysis.</li>
	<li>All animals should be humanely euthanized at the end of the procedure prior to anesthetic recovery in accordance with relevant euthanasia guidelines. Suitable methods of euthanasia may include cervical dislocation or CO2 inhalation.</li>
</ol>

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J., Mogri, M., Deisseroth, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optogenetics+in+neural+systems.">Optogenetics in neural systems.</a> Neuron. 71 (1), 9-34 (2011).</li><li>Hernandez, V. H., 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=Optogenetic+stimulation+of+the+auditory+pathway.">Optogenetic stimulation of the auditory pathway.</a> The Journal of clinical investigation. 124 (3), 1114-1129 (2014).</li><li>Gunaydin, L. A., Yizhar, O., Berndt, A., Sohal, V. S., Deisseroth, K., Hegemann, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ultrafast+optogenetic+control.">Ultrafast optogenetic control.</a> Nature neuroscience. 13 (3), 387-392 (2010).</li><li>Klapoetke, N. C., Murata, Y., 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=Independent+optical+excitation+of+distinct+neural+populations.">Independent optical excitation of distinct neural populations.</a> Nature methods. 11 (3), 338-346 (2014).</li></ol>

The authors declare that they have no competing financial interests.

The described experiments demonstrate the optogenetic stimulation of the SGNs, and can, in principle, also be used to stimulate inner and/or outer hair cells, provided the expression of opsins. These experiments require much patience and care. As mentioned before, the most critical steps are a good cochleostomy/round window insertion as well as an appropriate position and orientation of the light source.
There are limitations with optogenetic stimulation when using ChR2. In our case oABR ampli...

According to the World Health Organization, 360 million people worldwide suffer from hearing loss. In deaf subjects, direct electrical stimulation of SGNs by CIs&#160;enable open speech comprehension in the majority of them1,2,4,5. Even though CIs have been implanted in more than 200,000 people, therefore being the most successful neuroprosthesis, sound encoding driven by the current cochlear implants is limited. CIs are based on electrical stimulation by a certain number of electrodes where each one activates a tonotopic region of the auditory nerve thus bypassing the dysfunctional sensory organ of Corti in the cochlea. Normal hearing listeners can discriminate more than 2,000 frequencies, however today&#39;s CIs use only up to 12-22 frequency channels4. This is due to widespread current flow from each stimulating electrode7,9, activating a large number of SGNs that represent many different sound frequencies8,15. This limitation can be improved using multipolar stimulation but at the expense of higher power consumption16,17. Their output dynamic range for sound intensity is also limited, typically below 6-20 dB4,18. For these reasons, improving frequency and intensity resolution are important objectives for increasing CI performance to ameliorate speech recognition in noisy environments, prosody comprehension and music perception.
A different option to stimulate the auditory nerve is optical stimulation. Light can be conveniently focused to target a small SGN population, promising better spatial confinement, increasing frequency resolution and also widening dynamic range, resulting in a better intensity resolution. Indeed, cochlear stimulation with infrared light has shown excellent frequency resolution in animal models10,11,19. One disadvantage of this kind of stimulation is that it requires higher energies than electrical stimulation10,11. Moreover, concerns about the ability of the method to directly stimulate auditory neurons have been raised12,20.
As an alternative to infrared stimulation, we employ optogenetics to render SGNs light sensitive. Optogenetics is a novel approach that combines genetic and optical techniques to non-invasively and specifically control cells with high temporal precision (reviews21-23). The currently most frequently used modality employs the expression of the microbial channelrhodopsin 2 (ChR2) gene of Chlamydomonas reinhardtii and variants thereof, encoding a light-gated cation channel24. ChR2 is a 7-transmembrane-helix protein that, when transduced into neurons and activated by blue light, acts as non-selective cation channel, thus depolarizing the cells24-27. ChR2 has been well characterized24,28-31 and many variants have been developed to modify action spectrum, gating and permeability properties32,33. The aim of our work is to establish cochlear optogenetics for the activation of the auditory pathway. We note that the optogenetic approach to stimulate the auditory nerve requires genetic manipulation of the spiral ganglion for the expression of channelrhodopsin. Working with mice and rats allows the use of available transgenic animals13,34,35, which provide expression of channelrhodopsin with little variability along the tonotopic axis and across animals36. Combining conditional alleles37 with appropriate Cre-lines provides for cell-specific expression. Gene transfer into the spiral ganglion of other animals requires the use of virus such as adeno-associated virus that is a standard approach in optogenetics38 and that we showed to work well in mice36. Genetic manipulation and expression of transgenes encoding alien proteins bear risks for adverse effects such as immune responses and/or proliferation, compromised condition or even death of genetically manipulated cells. For the purpose of this demonstration we use transgenic mice expressing ChR2 in spiral ganglion neurons under the Thy-1 promotor13 to optically stimulate the auditory pathway. We note that other channelrhodopsin variants can be used for the same purpose as we demonstrated using virus-mediated transfer of the variant CatCh14 into SGNs39.
While cochlear optogenetics requires genetic manipulation, it offers molecular tuning for optimized SGN stimulation and promises improved frequency and intensity resolution when compared to electrical stimulation. Optogenetic stimulation of the auditory pathway is highly relevant for hearing research. For example, it promises advances in studies of the activity-dependent refinement of tonotopy during development, in the analysis of the requirement for spectral integration in sound localization and of the extent of interaction between frequency-specific afferent projections in the central auditory system.

<table border="0" cellpadding="0" cellspacing="0" width="769">
	<tbody>
		<tr>
			<td>Urethane</td>
			<td>Sigma Aldrich</td>
			<td>U2500-100G</td>
			<td>Anesthetic</td>
		</tr>
		<tr>
			<td>Xylazine HCl</td>
			<td>RXV</td>
			<td></td>
			<td>Sedative and analgesic</td>
		</tr>
		<tr>
			<td>Buprenorphine</td>
			<td>Reckitt Benckiser</td>
			<td></td>
			<td>Analgesic</td>
		</tr>
		<tr>
			<td>Dumont #5 Forceps</td>
			<td>Fine Science Tools</td>
			<td>11251-10</td>
			<td>It is used to hold hard tissue, e.g. bone or materials. Never use them to hold soft delicated tissue&#160;</td>
		</tr>
		<tr>
			<td>Dumont #5 - Fine Forceps</td>
			<td>Fine Science Tools</td>
			<td>11254-20</td>
			<td>Only to be used to hold soft tissue</td>
		</tr>
		<tr>
			<td>Fine Scissors - Sharp</td>
			<td>Fine Science Tools</td>
			<td>14060-09</td>
			<td>To open the skin and help with the muscle dissection</td>
		</tr>
		<tr>
			<td>Lempert Rongeurs&#160;</td>
			<td>Fine Science Tools</td>
			<td>16004-16</td>
			<td>They are very useful to easily remove the bone from the bulla</td>
		</tr>
		<tr>
			<td>473-nm laser&#160;</td>
			<td>Changchun New Industries</td>
			<td>MLL-III473</td>
			<td>100 mW solid state 473 nm laser</td>
		</tr>
		<tr>
			<td>Laser driver&#160;</td>
			<td>Changchun New Industries</td>
			<td>DPSSL MLL 100 mW</td>
			<td>TTL operated laser driver</td>
		</tr>
		<tr>
			<td>250 &#181;m optical fiber</td>
			<td>Any comercial ; e.g. Thorlabs</td>
			<td>M42L05</td>
			<td></td>
		</tr>
		<tr>
			<td>Acousto-optical modulator</td>
			<td>Crystal Technology, Inc.</td>
			<td>PCAOM VIS</td>
			<td>Control the amount of light coupled into the fiber from the laser</td>
		</tr>
		<tr>
			<td>Controller for Acousto-optical modulator</td>
			<td>Crystal Technology, Inc.</td>
			<td>160T1-8SAR-24-0.8</td>
			<td>Control the acousto-optic modulator</td>
		</tr>
		<tr>
			<td>Solo2 laser power &#38; energy meter</td>
			<td>Gentec-EO</td>
			<td></td>
			<td>Used to measure light intensity of the LED and the fiber coupled laser</td>
		</tr>
		<tr>
			<td>Blue &#181;LED</td>
			<td>Cree</td>
			<td>C470UT200</td>
			<td>It is necessary to build several &#956;LED devices because easily get damaged or the isolation is not good enough</td>
		</tr>
		<tr>
			<td>TDT System&#160;</td>
			<td>Tucker-Davis Technologies</td>
			<td>RZ6-A-P1</td>
			<td>It can be used any system for stimulus generation&#160; presentation and data acquisition</td>
		</tr>
		<tr>
			<td>Single-shank, 16-channel silicon probe</td>
			<td>Neuronexus</td>
			<td>a1x16-5mm-100-177-CM16LP&#160;</td>
			<td>These are fragile devises, must be handled carefully and cleaned after use</td>
		</tr>
		<tr>
			<td>Omnidrill</td>
			<td>World Precision Instruments</td>
			<td>503598</td>
			<td>Perform craniotomy for IC recordings and reference screw implantation</td>
		</tr>
		<tr>
			<td>Micro Drill Steel Burrs</td>
			<td>any commercial; e.g. Fine Science Tools</td>
			<td>19007-07</td>
			<td></td>
		</tr>
		<tr>
			<td>Self tapping bone screw</td>
			<td>any commercial; e.g. Fine Science Tools</td>
			<td>19010-10</td>
			<td>Reference screw</td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>any commercial; e.g. Luigs+NeumannInVivo Unit Junior 4 axis</td>
			<td></td>
			<td>Positioning of recording probe</td>
		</tr>
	</tbody>
</table>

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.

An optimal cochleostomy is critical and increases the probability of a successful experiment. This means the window is regular, small, and there is no injury of the internal cochlear structures. For example, bleeding indicates damage of the stria vascularis. A good example is presented in Figure 1B.
Using ChR2-transgenic mice, ChR2 is expressed in the SGNs within the cochlea (Figure 1C). Blue light illumination, either by &#181;LED or laser, e...

Watch this Scientific Journal Video about Optogenetic Stimulation of the Auditory Nerve at JoVE.com

Optogenetic Stimulation of the Auditory Nerve

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 ...

optogenetic-stimulation-of-the-auditory-nerve

Research

JoVE Journal

Neuroscience

14.5K Görüntüleme.  University Medical Center Goettingen.  Koklear implantlar (CI) işitme siniri doğrudan elektriksel stimülasyon ile işitme sağlarlar. Bununla birlikte, kötü sıklığı ve yoğunluğu çözünürlüğü CI ile işitme kalitesini sınırlar. Burada işitsel araştırma ve gelecekteki CI'ler geliştirmek için alternatif bir strateji olarak farelerde işitme sinirinin Optogenetic uyarılmasını açıklamaktadır. 

Video: Optogenetic Stimulation of the Auditory Nerve

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

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

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

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

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

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

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

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

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells in the walls of the lateral ventricle give rise to neuroblasts that migrate for several millimeters along the rostral migratory stream (RMS) to reach and incorporate into the olfactory bulb. To study the different steps and the impact of adult-born neuron integration into preexisting olfactory circuits, it is necessary to selectively label and manipulate the activity of this specific population of neurons. The recent development of optogenetic technologies offers the opportunity to use light to precisely activate this specific cohort of neurons without affecting surrounding neurons4,5. Here, we present a series of procedures to virally express Channelrhodopsin2(ChR2)-YFP in a temporally restricted cohort of neuroblasts in the RMS before they reach the olfactory bulb and become adult-born neurons. In addition, we show how to implant and calibrate a miniature LED for chronic in vivo stimulation of ChR2-expressing neurons.

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Adult-born neurons of the olfactory bulb can be optogenetically controlled using Channelrhodopsin2-expressing lentiviral injection in the rostral migratory stream and chronic photostimulation with an implanted miniature LED.

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells ...

Fiber-optic Implantation for Chronic Optogenetic Stimulation of Brain Tissue

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard for analyzing patterns of synaptic connectivity, but paired electrophysiological recordings can be both cumbersome and experimentally limiting. The development of optogenetics has introduced an elegant method to stimulate neurons and circuits, both in vitro1 and in vivo2,3. By exploiting cell-type specific promoter activity to drive opsin expression in discrete neuronal populations, one can precisely stimulate genetically defined neuronal subtypes in distinct circuits4-6. Well described methods to stimulate neurons, including electrical stimulation and/or pharmacological manipulations, are often cell-type indiscriminate, invasive, and can damage surrounding tissues. These limitations could alter normal synaptic function and/or circuit behavior. In addition, due to the nature of the manipulation, the current methods are often acute and terminal. Optogenetics affords the ability to stimulate neurons in a relatively innocuous manner, and in genetically targeted neurons. The majority of studies involving in vivo optogenetics currently use a optical fiber guided through an implanted cannula6,7; however, limitations of this method include damaged brain tissue with repeated insertion of an optical fiber, and potential breakage of the fiber inside the cannula. Given the burgeoning field of optogenetics, a more reliable method of chronic stimulation is necessary to facilitate long-term studies with minimal collateral tissue damage. Here we provide our modified protocol as a video article to complement the method effectively and elegantly described in Sparta et al.8 for the fabrication of a fiber optic implant and its permanent fixation onto the cranium of anesthetized mice, as well as the assembly of the fiber optic coupler connecting the implant to a light source. The implant, connected with optical fibers to a solid-state laser, allows for an efficient method to chronically photostimulate functional neuronal circuitry with less tissue damage9 using small, detachable, tethers. Permanent fixation of the fiber optic implants provides consistent, long-term in vivo optogenetic studies of neuronal circuits in awake, behaving mice10 with minimal tissue damage.

The development of optogenetics now provides the means to precisely stimulate genetically defined neurons and circuits, both in vitro and in vivo. Here we describe the assembly and implantation of a fiber optic for chronic photostimulation of brain tissue.

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2 avoidance, proboscis extension and giant-fiber mediated startle response2-4. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-&beta;, encoded by the norpA gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

Genetically encoded optogenetic tools enable noninvasive manipulation of specific neurons in the Drosophila brain. Such tools can identify neurons whose activation is sufficient to elicit or suppress particular behaviors. Here we present a method for activating Channelrhodopsin2 that is expressed in targeted neurons in freely walking flies.

A  growing number of genetically encoded tools are becoming available that allow non-invasive  manipulation of the neural activity of specific neurons in ...

Selective Tracing of Auditory Fibers in the Avian Embryonic Vestibulocochlear Nerve

The embryonic chick is a widely used model for the study of peripheral and central ganglion cell projections. In the auditory system, selective labeling of auditory axons within the VIIIth cranial nerve would enhance the study of central auditory circuit development. This approach is challenging because multiple sensory organs of the inner ear contribute to the VIIIth nerve 1. Moreover, markers that reliably distinguish auditory versus vestibular groups of axons within the avian VIIIth nerve have yet to be identified. Auditory and vestibular pathways cannot be distinguished functionally in early embryos, as sensory-evoked responses are not present before the circuits are formed. Centrally projecting VIIIth nerve axons have been traced in some studies, but auditory axon labeling was accompanied by labeling from other VIIIth nerve components 2,3. Here, we describe a method for anterograde tracing from the acoustic ganglion to selectively label auditory axons within the developing VIIIth nerve. First, after partial dissection of the anterior cephalic region of an 8-day chick embryo immersed in oxygenated artificial cerebrospinal fluid, the cochlear duct is identified by anatomical landmarks. Next, a fine pulled glass micropipette is positioned to inject a small amount of rhodamine dextran amine into the duct and adjacent deep region where the acoustic ganglion cells are located. Within thirty minutes following the injection, auditory axons are traced centrally into the hindbrain and can later be visualized following histologic preparation. This method provides a useful tool for developmental studies of peripheral to central auditory circuit formation.

Here we describe a microdissection technique followed by fluorescent dye injection into the acoustic ganglion of early chick embryos for selective tracing of auditory axon fibers in the nerve and hindbrain.

The embryonic chick is a widely used model for the study of peripheral and central ganglion cell projections. In the auditory system, selective labeling ...

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 ...

Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. Few studies have examined gene transfer to the central auditory system. The dorsal cochlear nucleus (DCN) of the brainstem, which contains second order neurons of the auditory pathway, is a potential site for gene transfer. In this protocol, a technique for direct and maximal exposure of the murine DCN via a posterior fossa approach is demonstrated. This approach allows for either acute or survival surgery. Following direct visualization of the DCN, a host of experiments are possible, including injection of opsins into the cochlear nucleus and subsequent stimulation by an optical fiber coupled to a blue light laser.&#160;Other neurophysiology experiments, such as electrical stimulation and neural injector tracings are also feasible.&#160;The level of visualization and the duration of stimulation achievable make this approach applicable to a wide range of experiments.

The goal of this protocol is to outline a surgical approach to provide direct access to the dorsal cochlear nucleus in a murine model.

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. ...

Assessment of Neuromuscular Function Using Percutaneous Electrical Nerve Stimulation

Percutaneous electrical nerve stimulation is a non-invasive method commonly used to evaluate neuromuscular function from brain to muscle (supra-spinal, spinal and peripheral levels). The present protocol describes how this method can be used to stimulate the posterior tibial nerve that activates plantar flexor muscles. Percutaneous electrical nerve stimulation consists of inducing an electrical stimulus to a motor nerve to evoke a muscular response. Direct (M-wave) and/or indirect (H-reflex) electrophysiological responses can be recorded at rest using surface electromyography. Mechanical (twitch torque) responses can be quantified with a force/torque ergometer. M-wave and twitch torque reflect neuromuscular transmission and excitation-contraction coupling, whereas H-reflex provides an index of spinal excitability. EMG activity and mechanical (superimposed twitch) responses can also be recorded during maximal voluntary contractions to evaluate voluntary activation level. Percutaneous nerve stimulation provides an assessment of neuromuscular function in humans, and is highly beneficial especially for studies evaluating neuromuscular plasticity following acute (fatigue) or chronic (training/detraining) exercise.

We present a protocol to assess changes in neuromuscular function. Percutaneous electrical nerve stimulation is a non-invasive method that evokes muscular responses. Electrophysiological and mechanical properties of these responses permit the evaluation of neuromuscular function from brain to muscle (supra-spinal, spinal and peripheral levels).

Percutaneous electrical nerve stimulation is a non-invasive method commonly used to evaluate neuromuscular function from brain to muscle (supra-spinal, ...

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.