Funding: This work was supported by a Foundation Bertarelli grant (D.J.L), a MED-EL grant (D.J.L), and a National Institutes of Health Grants DC01089 (M.C.B.).

NOTE: All experimental procedures are performed in accordance with the Animal Care and Use Committee of the Massachusetts Eye and Ear Infirmary and Harvard Medical School, which follow national animal care guidelines, including Public Health Service Policy on Humane Care and Use of laboratory Animals, the ILAR Guide, and the Animal Welfare Act.&#160;Experimental procedures listed below detail exposure of the left DCN. Use sterile&#160;instruments while performing survival surgery.
1. Primary Craniotomy and Dorsal Cochlear Nucleus Exposure
<ol>
	<li>Anesthesia
	<ol>
		<li>Anesthetize a mouse, aged 8&#8211;12 weeks, weighing 18&#8211;24 g, with xylazine 20 mg/kg and ketamine 100 mg/kg via an intraperitoneal administration. Confirm proper anesthesia by monitoring heart rate, respiratory rate, as well as toe pinch withdrawal reflex. Place vet ointment on eyes to prevent dryness while under anesthesia. Once adequately under anesthesia, shave hair overlying the scalp to provide unobstructed access to the surgical site.</li>
	</ol>
	</li>
	<li>Surgical positioning
	<ol>
		<li>Place the mouse securely in a small animal stereotaxic holder, held in place by a snout clamp.</li>
		<li>Ensure that the snout clamp is loose enough to allow for adequate respiration but tight enough to completely immobilize the head of the mouse. If the mouse head is loose, the animal is likely to fall out of the head holder during the craniotomy portion of procedure.</li>
		<li>Place auditory brainstem implant electrodes in standard fashion, which&#160; allows for heart rate monitoring. Respiratory rate should be monitored by visualization. Note that normal heart rate and respiratory rate will vary depending on age of the mouse.</li>
		<li>A thermometer is placed and euthermia is ensured via placement on a homeothermic heating blanket.</li>
	</ol>
	</li>
	<li>Skin incision and exposure of the interparietal and occipital bones of the skull
	<ol>
		<li>Under a microscope, make a vertical incision through the skin starting at the midline, directly between the pinna, and extending to the caudal portion of the occiput. Following midline skin incision, displace the skin laterally. Visualize the muscles covering the left the posterior aspect of the skull.</li>
		<li>Remove muscle overlying the left parietal, interparietal, and occipital bones of the skull by disarticulation of muscle attachments to their respective bones with a scalpel or iris scissor. Observe a small degree of bleeding along cut muscle edges. Minimize this by gentle pressure with a cotton-tipped applicator for 10-15 sec.</li>
	</ol>
	</li>
	<li>Craniotomy overlying Cochlear Nucleus
	<ol>
		<li>Identify relevant suture lines, including sagittal and lambda sutures lines (Figure 1, Left Panel).</li>
		<li>Using rongeurs, make a craniotomy over the interparietal bone, left of midline, ~2 mm caudal to the lambda suture line. This region overlies the DCN.</li>
		<li>Following craniotomy, observe a thin layer of dura that overlies the cerebellum. Using a scalpel blade, remove the dura. (Figure 1, Middle Panel). 
		NOTE: Removal of dura may cause a small degree of bleeding. This process of using the scalpel blade to remove the dura is intended to reduce the amount of blood loss during the cerebellum aspiration, which will pool on the brainstem surface and obscure identification of the DCN.&#160;Total blood volume of mouse is ~1.5 ml should not exceed &#62; than 15%. If bleeding exceeds this amount, the mouse should be provided with&#160; with fluid supplementation.&#160;</li>
		<li>Remove coagulated blood overlaying the cerebellum with the cotton tip applicator. Optionally, gently drip saline using a dental point onto the cerebellum to clear blood away blood.</li>
	</ol>
	</li>
	<li>Cerebellar Aspiration
	<ol>
		<li>Using a 5 French suction, aspirate the lateral-most portion of the left cerebellum overlying the DCN until the DCN is visual (Figure 1, Right Panel). Remove approximately 1/4 to 1/3 of the left cerebellum. The main landmark adjacent to the DCN is the ampulla of the superior semicircular canal. 
		NOTE: Aspiration of the cerebellum is the key step of the protocol. Directed aspiration of the cerebellum is most successfully performed if it occurs in a single attempt and not with multiple passes of the suction as this will cause bleeding. To assist in aspiration, set the focal plane of the microscope at the expected depth of the CN, which will be slightly distal to the surface of the cerebellum. Setting the focal plane to the CN will improve visualization and ensure a sharp image.</li>
		<li>Following aspiration, further bleeding, cerebrospinal fluid (CSF) build-up, and cerebellum displacement on the DCN are expected. Instill 0.5 cc of sterile saline quickly into the craniotomy to prevent blood coagulation.&#160;A combination of gentle dabbing with a dental point and suction may then be used to clear away a path for direct visualization of the DCN. Do not directly contact the surface of the DCN.</li>
	</ol>
	</li>
</ol>
2. Pressure Microinjection for Virus-mediated Gene Transfer and Surgical Recovery
<ol>
	<li>After the DCN is free of overlying blood and CSF and is clearly visible, make pressure microinjections into the DCN using a 10 &#181;l Hamilton syringe over a 2 min period.</li>
	<li>Introduce the needle with a micromanipulator until the tip is no longer visible under the surface of the DCN.</li>
	<li>For optimal performance, use a 33 or 34 gauge needle (use a gastight syringe with the 34 gauge needle) with a shallow bevel, such as 45&#176;, to minimize blunt trauma and localize the injection volume within the DCN, which is a thin and shallow brainstem structure (&#60;300 &#181;m thick). 
	NOTE: Other microinjection instruments may be utilized based. Ideally, the injection instrument should be flexible to allow for some slight bend, if needed to directly target the DCN. Further, as the mouse moves slightly throughout the procedure due to respiration, the instrument should be robust enough to withstand minor amounts of movement.</li>
</ol>
3. Surgical Recovery
<ol>
	<li>Immediately following injection, re-approximate the skin and allow the mouse to recover per standard recovery procedure. Remaining cerebellum and scar tissue will fill in the cavity caused by aspiration of the cerebellum. Constantly monitor animals until sufficient consciousness is regained to maintain sternal recumbency. Monitor heart rate, respiratory rate, as well as ability to feed. 
	NOTE: No animal that has undergone surgery is returned to a cage with other animals until fully recovered.</li>
	<li>If a mouse begins to walk in circles post-operatively (&#60;5% of cases), typically 2-4 hr&#160;after closure of the skin incision, immediately sacrifice the mouse, as it would likely have a difficult time feeding. This side effect is likely due to aspiration of the cerebellum.&#160;The animal should be monitored for pain postoperatively, and appropriate narcotics should be given to ensure animal comfort based on institutional standards. Antibiotics may be necessary, if evidence of infection.</li>
</ol>
4. Secondary Craniotomy and Cochlear Nucleus Exposure
<ol>
	<li>After 2-4 weeks of healing and virus-mediated gene transfer incubation, re-anesthetize a previously injected mouse and repeat step 1.1-1.5 Optimal incubation time varies with the type of gene transferred.</li>
	<li>Remove scar tissue over the craniotomy site by a combination of tweezers, scalpel, and rongeurs.</li>
	<li>After visualizing the craniotomy, identify a combination of remaining cerebellum and scar tissue overlying the DCN. Use a 5 French suction to aspirate overlying cerebellum/scar tissue (similar to the previous cerebellar aspiration).</li>
	<li>Following aspiration, expect further bleeding, CSF build-up, and remaining bits of cerebellum. Quickly introduce saline into the craniotomy to prevent blood coagulation. Use a combination of gentle dental point dabbing and suction to clear the path for direct visualization of the DCN.</li>
	<li>Observe the surface of the DCN. Perform optogenetics-based physiology experiments as DCN is now accessible. Introduce a light stimulus, for example with an optical fiber (Figure 2), to drive an optogenetics-based experiment.</li>
</ol>
5. Histology
<ol>
	<li>Following conclusion of experiments, euthanize the mouse with an overdose of ketamine. Perfuse the mouse with normal saline followed by 4% paraformaldehyde.</li>
	<li>Extract the brainstem from the skull and post-fix for 2 hr. Cryoprotect the brainstem in 30% sucrose for 24-48 hr. Section the brainstem using a standard cryostat using 60 &#181;m sections.</li>
</ol>

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The authors have nothing to disclose.

This paper describes the technique of direct visualization of the DCN in the murine model for manipulation of the central auditory system. The outlined approach of direct visualization provides significant advantages over the main alternative, which are stereotaxic approaches. Primarily, direct visualization of the DCN allows for immediate confirmation of the site of the brainstem, whereas stereotaxic approaches do not afford direct visualization. In experiments that necessitate prolonged incubation periods, as is the ca...

Virus-mediated gene transfer to reverse hearing loss has largely been focused on the peripheral auditory system.1 Targeting the cochlea, investigators have examined a host of delivery routes, including osmotic minipump infusion2, vector-transgene complex-soaked Gelfoam&#174;2 or gelatin sponge3, direct microinjection4; numerous gene transfer vectors, including adeno-associated viral vectors5,6, lentiviral vectors7, and cationic liposomes2; and the dissemination of gene transfer vectors beyond the target tissue2. Most recently, adeno-associated virus (AAV)-1 has been introduced in the cochlea in order to treat deafness in mice due to loss of vesicular glutamate transporter-3.8 Further, the application of optogenetics in peripheral auditory system has recently been described.9
Few studies, however, have examined gene transfer to the central auditory system. The dorsal cochlear nucleus (DCN) of the brainstem contain second order neurons of the auditory pathway. While gene transfer techniques in the cochlear nucleus (CN) may be utilized for a host of investigations, gene transfer of opsins, light-sensitive proteins, to the DCN may also be utilized to enable optogenetics-based experimental techniques. Following virus-mediated gene transfer delivery of an opsin, such as channelrhodopsin-2 (ChR2), the neurons of the DCN becomes sensitive to light stimuli. Optogenetic gene transfer has been previously attempted in several brainstem regions, including the rat retrotrapezoid nucleus, mouse locus coeruleus, monkey superior colliculus, and mouse ventral tegmental area.10-14
Recently, investigators have examined the use of optogenetics in the DCN.15,16 The DCN is the location of placement of auditory brainstem implants in humans, making it an attractive part of the auditory system to study for translational studies on auditory neuroprostheses. However, given the location of the DCN, surgical exposure is challenging. The technique described herein provides a protocol for maximal exposure of the DCN via posterior fossa approach to enable viral vector gene transfer and optogenetics-based experiments in a murine model. Previous studies used stereotactic microinjection into the DCN with channelrhodopsin-2.16 Stereotaxic injections, however, are potentially less accurate than injections made by direct visualization, especially in a nucleus as small and deep along the brainstem as the DCN. Transgenic mice expressing tissue specific proteins in the CN are also an attractive option and would obviate the need for gene transfer. Our protocol for exposure of the DCN would also work in transgenic mice as the DCN would need to be directly exposed for optical stimulation. This technique for surgical exposure of the DCN is adapted from previous protocols involving recordings from the auditory nerve and cochlear nucleus in mice and rat models.15,17-20
The overall goal of the protocol is to provide direct exposure to the CN to allow for gene transfer techniques. More specifically, the approach is compatible with both acute and survival surgery and the preparation can be repeated in the same animal for subsequent neurophysiological testing. The direct exposure of the DCN protocol has implications for optogenetics- and virus-mediated gene transfer-based experimentation in other nuclei of the brainstem.

<table border="0" cellpadding="0" cellspacing="0" width="560">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:197px;">Name of the Material / Equipment</td>
			<td style="width:227px;">Company</td>
			<td style="width:136px;">Catalog Number</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:197px;">Stereotaxic holder</td>
			<td style="width:227px;">Stoelting</td>
			<td>51500</td>
		</tr>
		<tr height="21">
			<td height="42" rowspan="2" style="height:42px;width:197px;">Homeothermic blanket</td>
			<td rowspan="2" style="width:227px;">Harvard</td>
			<td rowspan="2">507214</td>
		</tr>
		<tr height="21">
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:197px;">Scalpel blade #11</td>
			<td style="width:227px;">Fine Surgical Tools</td>
			<td>10011-00</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:197px;">Iris scissor</td>
			<td style="width:227px;">Fine Surgical Tools</td>
			<td>14084-08</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:197px;">5 French suction</td>
			<td style="width:227px;">Symmetry Surgical</td>
			<td>2777914</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:197px;">Dental Points</td>
			<td style="width:227px;">Henry Schein</td>
			<td>100-8170</td>
		</tr>
		<tr height="25">
			<td height="46" rowspan="2" style="height:46px;width:197px;">Bone rongeur</td>
			<td rowspan="2" style="width:227px;">Fine Surgical Tools</td>
			<td rowspan="2">16020-14</td>
		</tr>
		<tr height="21">
		</tr>
		<tr height="30">
			<td height="51" rowspan="2" style="height:52px;width:197px;">10 &#181;l Hamilton syringe</td>
			<td rowspan="2" style="width:227px;">Hamilton&#160;</td>
			<td rowspan="2">7633-01</td>
		</tr>
		<tr height="21">
		</tr>
		<tr height="21">
			<td height="42" rowspan="2" style="height:42px;width:197px;">34 gauge, needle</td>
			<td rowspan="2" style="width:227px;">Hamilton&#160;</td>
			<td rowspan="2">207434</td>
		</tr>
		<tr height="21">
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rongeurs</td>
			<td>Fine Surgical Tools</td>
			<td>16021-14</td>
		</tr>
	</tbody>
</table>

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.

Partial Cerebellar Aspiration Demonstrates Access to the Cochlear Nucleus
After the skin and muscle overlying the skull are removed, skull surface landmarks, such as the coronal and lamda suture lines, demonstrate the approximate localization of the craniotomy. Following craniotomy with rongeurs, the cerebellum is visualized. Careful aspiration of the small portion of the cerebellum demonstrates visualization of the CN, which can then be injected (Figure 1).
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Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway

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. ...

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9.2K ビュー.  Harvard Medical School. The goal of this protocol is to outline a surgical approach to provide direct access to the dorsal cochlear nucleus in a murine model. 

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

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 ...

Electrode Positioning and Montage in Transcranial Direct Current Stimulation

Transcranial direct current stimulation (tDCS) is a technique that has been intensively investigated in the past decade as this method offers a non-invasive and safe alternative to change cortical excitability2. The effects of one session of tDCS can last for several minutes, and its effects depend on polarity of stimulation, such as that cathodal stimulation induces a decrease in cortical excitability, and anodal stimulation induces an increase in cortical excitability that may last beyond the duration of stimulation6. These effects have been explored in cognitive neuroscience and also clinically in a variety of neuropsychiatric disorders &#x2013; especially when applied over several consecutive sessions4. One area that has been attracting attention of neuroscientists and clinicians is the use of tDCS for modulation of pain-related neural networks3,5. Modulation of two main cortical areas in pain research has been explored: primary motor cortex and dorsolateral prefrontal cortex7. Due to the critical role of electrode montage, in this article, we show different alternatives for electrode placement for tDCS clinical trials on pain; discussing advantages and disadvantages of each method of stimulation.

Transcranial direct current stimulation (tDCS) is an established technique to modulate cortical excitability1,2. It has been used as an investigative tool in neuroscience due to its effects on cortical plasticity, easy operation, and safe profile. One area that tDCS has been showing encouraging results is pain alleviation 3-5.

Transcranial direct current stimulation (tDCS) is a technique that has been intensively investigated in the past decade as this method offers a ...

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 ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

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

Visualization of Proprioceptors in Drosophila Larvae and Pupae

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

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

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 ...

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 ...

Analysis of Gene Expression Changes in the Rat Hippocampus After Deep Brain Stimulation of the Anterior Thalamic Nucleus

Deep brain stimulation (DBS) surgery, targeting various regions of the brain such as the basal ganglia, thalamus, and subthalamic regions, is an effective treatment for several movement disorders that have failed to respond to medication. Recent progress in the field of DBS surgery has begun to extend the application of this surgical technique to other conditions as diverse as morbid obesity, depression and obsessive compulsive disorder. Despite these expanding indications, little is known about the underlying physiological mechanisms that facilitate the beneficial effects of DBS surgery. One approach to this question is to perform gene expression analysis in neurons that receive the electrical stimulation. Previous studies have shown that neurogenesis in the rat dentate gyrus is elicited in DBS targeting of the anterior nucleus of the thalamus1. DBS surgery targeting the ATN is used widely for treatment refractory epilepsy. It is thus of much interest for us to explore the transcriptional changes induced by electrically stimulating the ATN. In this manuscript, we describe our methodologies for stereotactically-guided DBS surgery targeting the ATN in adult male Wistar rats. We also discuss the subsequent steps for tissue dissection, RNA isolation, cDNA preparation and quantitative RT-PCR for measuring gene expression changes. This method could be applied and modified for stimulating the basal ganglia and other regions of the brain commonly clinically targeted. The gene expression study described here assumes a candidate target gene approach for discovering molecular players that could be directing the mechanism for DBS.

The mechanism underlying the therapeutic effects of Deep Brain Stimulation (DBS) surgery needs investigation. The methods presented in this manuscript describe an experimental approach to examine the cellular events triggered by DBS by analyzing the gene expression profile of candidate genes that can facilitate neurogenesis post DBS surgery.

Deep brain stimulation (DBS) surgery, targeting various regions of the brain such as the basal ganglia, thalamus, and subthalamic regions, is an effective ...