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 case in virus-mediated gene transfer, there is the potential for low infection efficiency if the injection &#8220;misses&#8221; the target location. Further, a direct visualization approach allows for injection in a wide range animal ages. An analogous surgical approach may be used in survival experiments with other animal models, such as rats and guinea pigs.
There are several critical steps in this protocol. First, the mouse must be positioned properly in the stereotactic holder. Any movement of the head will result in difficulty making the craniotomy. Second, location of the craniotomy is crucial. If the craniotomy is not made in the proper position, visualization of the DCN following aspiration of the cerebellum will be challenging, if not impossible. Finally, the most critical step of the protocol is the partial directed aspiration of the cerebellum. There are two technical approaches to aspiration of the cerebellum. In the first aspiration approach, the suction is constantly held on the cerebellum, lifting it upwards, off the brainstem. In this approach, the DCN may be partially visualized underneath the cerebellum and suction is held until the CN is fully visualized and cerebellum removed. In the second approach, the cerebellum is partially aspirated in several layered passes until the DCN is visualized. Dental points may be used with gentle dabbing to clear blood or remaining cerebellum fragments from the CN if it cannot be fully visualized.
Bleeding is a major drawback of the direct visualization approach and principally occurs during the initial craniotomy and after cerebellar aspiration. In many cases during the initial craniotomy, bleeding will stop with time within 2-5 min. Placing a tissue paper or dental point at the site of bleeding and allowing for hemostasis. In terms of bleeding after cerebellar aspiration, it is important to inject normal saline (0.5-1 cc) at the site of the craniotomy to dilute the blood and prevent blood clot formation at the level of the CN. A combination of suction and gentle dental point dabbing may be used to clear away blood and saline. Indeed, the major limitation of this approach is the invasive nature of the procedure.
The protocol has implications beyond manipulation of the auditory system. In vivo gene transfer into the central nervous system enables manipulation of diverse neural pathways and has offered insights into multiple neural functions, including memory, motor control, olfaction, and audition.16,21-26 The surgical approach, enabling direct visualization for the purposes of gene transfer and stimulation may potentially be applied to other regions of the brainstem, both those listed above and others. Following mastery of direct visualization of the DCN, a host of experiments are possible, including injection of opsins into the CN and subsequent stimulation with a blue light laser. Other neurophysiology experiments, such as electrical stimulation and neural injector tracings,18 are also feasible. The level of visualization and the duration of stimulation of this approach allows for multiple applications for a wide range of experiments.

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><tbody><tr><td>Stereotaxic holder</td><td>Stoelting</td><td>51500</td><td></td></tr><tr><td>Homeothermic blanket</td><td>Harvard</td><td>507214</td><td></td></tr><tr><td>Scalpel blade #11</td><td>Fine Surgical Tools</td><td>10011-00</td><td></td></tr><tr><td>Iris scissor</td><td>Fine Surgical Tools</td><td>14084-08</td><td></td></tr><tr><td>5 French suction</td><td>Symmetry Surgical</td><td>2777914</td><td></td></tr><tr><td>Dental Points</td><td>Henry Schein</td><td>100-8170</td><td></td></tr><tr><td>Bone rongeur</td><td>Fine Surgical Tools</td><td>16020-14</td><td></td></tr><tr><td>10 &amp;micro;l Hamilton syringe</td><td>Hamilton&amp;nbsp;</td><td>7633-01</td><td></td></tr><tr><td>34 gauge, needle</td><td>Hamilton&amp;nbsp;</td><td>207434</td><td></td></tr><tr><td>Rongeurs</td><td>Fine Surgical Tools</td><td>16021-14</td><td></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).
Secondary Cochlear Nucleus Exposure for Stimulation with Blue Light Laser
Following initial exposure of the DCN, gene transfer with an opsin, and the incubation period, the DCN can then be stimulated by a blue light laser. The secondary craniotomy exposing the surface of the DCN for optical stimulation is similar in technical approach to the primary craniotomy (Supp. Movie 1). The DCN, being a surface structure that is less than 300 &#181;m in thickness. The experimental murine preparation remains viable for up to 6 hr.
<img alt="Figure 1" fo:content-width="7in" src="/files/ftp_upload/52426/52426fig1highres.jpg" width="700" /> 
Figure 1: Partial cerebellar aspiration demonstrates access to the dorsal cochlear nucleus. Left Panel: After the mouse is placed in the holder, an incision is made vertically through the skin and soft tissue along the posterior aspect of the skull to the soft tissue of the neck. The skin is lateralized, and the muscle overlying the coronal suture line is removed.&#160;Middle Panel: A craniotomy is then completed over the region of the cerebellum and underlying dorsal cochlear nucleus with the bone rongeurs.&#160;Right Panel: For exposure of the dorsal cochlear nucleus, a 5 French suction is then used to aspirate the cerebellum overlying the cochlear nucleus until the dorsal cochlear nucleus is visualized.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/52426/52426fig2highres.jpg" width="500" /> 
Figure 2: Exposure of the surface of the cochlear nucleus for stimulation with blue light laser to produce an optically-evoked auditory brainstem response. Placement of optical fiber coupled to a blue light laser on the surface of the dorsal cochlear nucleus allows for optical stimulation of the opsin-infected dorsal cochlea nucleus. Blue bar = 400 &#181;m.

Watch this Scientific Journal Video about Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway at JoVE.com

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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Nanyang Technological University

Research

JoVE Journal

Neuroscience

9.2K Views.  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. 

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

Canalostomy As a Surgical Approach to Local Drug Delivery into the Inner Ears of Adult and Neonatal Mice

Local delivery of therapeutic drugs into the inner ear is a promising therapy for inner ear diseases. Injection through semicircular canals (canalostomy) has been shown to be a useful approach to local drug delivery into the inner ear. The goal of this article is to describe, in detail, the surgical techniques involved in canalostomy in both adult and neonatal mice. As indicated by fast-green dye and adeno-associated virus serotype 8 with the green fluorescent protein gene, the canalostomy facilitated broad distribution of injected reagents in the cochlea and vestibular end-organs with minimal damage to hearing and vestibular function. The surgery was successfully implemented in both adult and neonatal mice; indeed, multiple surgeries could be performed if required. In conclusion, canalostomy is an effective and safe approach to drug delivery into the inner ears of adult and neonatal mice and may be used to treat human inner ear diseases in the future.

Here we describe canalostomy procedure which allows local drug delivery into the inner ears of adult and neonatal mice through the semicircular canal with minimum damage to hearing and vestibular function. This method can be used to inoculate viral vectors, pharmaceuticals, and small molecules into the mouse inner ear.

Local delivery of therapeutic drugs into the inner ear is a promising therapy for inner ear diseases. Injection through semicircular canals (canalostomy) ...

Genetics

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 ...

Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the interactions of these forebrain circuits has been aided recently by the development of optogenetic methods for light-mediated modulation of neuronal activity. Here, we describe a protocol for examining the functional organization of forebrain circuits&#160;in vitro using laser-scanning photostimulation of channelrhodopsin, expressed optogenetically via viral-mediated transfection. This approach also exploits the utility of cre-lox recombination in transgenic mice to target expression in specific neuronal cell types. Following transfection, neurons are physiologically recorded in slice preparations using whole-cell patch clamp to measure their evoked responses to laser-scanning photostimulation of channelrhodopsin expressing fibers. This approach enables an assessment of functional topography and synaptic properties. Morphological correlates can be obtained by imaging the neuroanatomical expression of channelrhodopsin expressing fibers using confocal microscopy of the live slice or post-fixed tissue. These methods enable functional investigations of forebrain circuits that expand upon more conventional approaches.

We describe a method for characterizing the functional topography and synaptic properties of forebrain circuits using an optogenetic approach to photostimulate neuronal populations&#160;in vitro.

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the ...

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti. 

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the inner ear (fig 1). Hair cells in the developing cochlea, which are the mechanosensory cells of the auditory system, are aligned in one row of inner hair cells and three (in the base and mid-turns) to four (in the apical turn) rows of outer hair cells that span the length of the organ of Corti. Hair cells transduce sound-induced mechanical vibrations of the basilar membrane into neural impulses that the brain can interpret. Most cases of sensorineural hearing loss are caused by death or dysfunction of cochlear hair cells.
An increasingly essential tool in auditory research is the isolation and in vitro culture of the organ explant 1,2,9. Once isolated, the explants may be utilized in several ways to provide information regarding normative, anomalous, or therapeutic physiology. Gene expression, stereocilia motility, cell and molecular biology, as well as biological approaches for hair cell regeneration are examples of experimental applications of organ of Corti explants.
This protocol describes a method for the isolation and culture of the organ of Corti from neonatal mice. The accompanying video includes stepwise directions for the isolation of the temporal bone from mouse pups, and subsequent isolation of the cochlea, spiral ligament, and organ of Corti. Once isolated, the sensory epithelium can be plated and cultured in vitro in its entirety, or as a further dissected micro-isolate that lacks the spiral limbus and spiral ganglion neurons. Using this method, primary explants can be maintained for 7-10 days. As an example of the utility of this procedure, organ of Corti explants will be electroporated with an exogenous DsRed reporter gene. This method provides an improvement over other published methods because it provides reproducible, unambiguous, and stepwise directions for the isolation, microdissection, and primary culture of the organ of Corti.

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti.

This procedure describes a method for the isolation and culture of the murine organ of Corti with or without the spiral limbus and spiral ganglion neurons. We also demonstrate a method for the expression of an exogenous reporter gene in the organ of Corti explant by electroporation.

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the ...

Biology

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 ...

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral cortex. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex using a method developed by Lima and colleagues1. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. This technique &#8211; termed &#8220;PINP&#8221;, or Photostimulation-assisted Identification of Neuronal Populations &#8211; can be implemented with standard extracellular recording equipment. It can serve as an inexpensive and accessible alternative to calcium imaging or visually-guided patching, for the purpose of targeting extracellular recordings to genetically-identified cells. Here we provide a set of guidelines for optimizing the method in everyday practice. We refined our strategy specifically for targeting parvalbumin-positive (PV+) cells, but have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons.

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching.

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral ...

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 ...

Neonatal Murine Cochlear Explant Technique as an In Vitro Screening Tool in Hearing Research

While there have been remarkable advances in hearing research over the past few decades, there is still no cure for Sensorineural Hearing Loss (SNHL), a condition that typically involves damage to or loss of the delicate mechanosensory structures of the inner ear. Sophisticated in vitro and ex vivo assays have emerged in recent years, enabling the screening of an increasing number of potentially therapeutic compounds while minimizing resources and accelerating efforts to develop cures for SNHL. Though homogenous cultures of certain cell types continue to play an important role in current research, many scientists now rely on more complex organotypic cultures of murine inner ears, also known as cochlear explants. The preservation of organized cellular structures within the inner ear facilitates the in situ evaluation of various components of the cochlear infrastructure, including inner and outer hair cells, spiral ganglion neurons, neurites, and supporting cells. Here we present the preparation, culture, treatment, and immunostaining of neonatal murine cochlear explants. The careful preparation of these explants facilitates the identification of mechanisms that contribute to SNHL and constitutes a valuable tool for the hearing research community.

Neonatal Murine Cochlear Explant Technique as an In Vitro Screening Tool in Hearing Research

The goal of this protocol is to demonstrate the preparation, culture, treatment, and immunostaining of neonatal murine cochlear explants. The technique can be utilized as an in vitro screening tool in hearing research.

While there have been remarkable advances in hearing research over the past few decades, there is still no cure for Sensorineural Hearing Loss (SNHL), a ...

Medicine

Multiscale Investigations of Cortical Processing by Integrating Laminar Polytrodes and Optogenetics with Micro Electrocorticography in Rodents

Electrocorticography (ECoG) is a methodological bridge between basic neuroscience and understanding human brain function in health and disease. ECoG records neurophysiological signals directly from the cortical surface at millisecond temporal resolution and columnar spatial resolution over large regions of cortical tissue simultaneously, making it uniquely positioned to study both local and distributed cortical computations. Here, we describe the design of custom, high-density micro-ECoG (&#181;ECoG) devices and their use in two procedures. These grids have 128 low-impedance electrodes with 200 &#181;m spacing fabricated on a clear polymer substrate with perforations between electrodes; these features enable simultaneous &#181;ECoG recording with laminar polytrode recordings and optogenetic manipulations. First, we present a protocol for combined epidural &#181;ECoG recording over the whisker somatosensory cortex of mice with optogenetic manipulation of specific genetically defined cortical cell types. This allows causal dissection of the distinct contributions of different neuronal populations to sensory processing while also monitoring their specific signatures in &#181;ECoG signals. Second, we present a protocol for acute experiments to record neural activity from the rat auditory cortex using &#181;ECoG grids and laminar polytrodes. This allows detailed topographic mapping of sensory-evoked neural responses across the cortical surface simultaneously with recordings from multiple neural units distributed across the cortical depth. These protocols enable experiments that characterize distributed cortical activity and may contribute to understanding and eventual interventions for diverse neurological disorders.

Here, we present two protocols for high-density micro-electrocorticography (&#181;EcoG) recording in rats and mice, including surgical, implantation, and recording methods. &#181;ECoG recordings are performed in combination with either laminar polytrode recording in the rat auditory cortex or with optogenetic manipulation of neural activity in the mouse somatosensory cortex.

Electrocorticography (ECoG) is a methodological bridge between basic neuroscience and understanding human brain function in health and disease. ECoG ...

Preparing Wedge Brain Slices to Visualize Auditory Neurons

Source: Fischl, M. J. et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/61664">In Vitro Wedge Slice Preparation for Mimicking In Vivo Neuronal Circuit Connectivity.</a>&#160;J. Vis. Exp.&#160;(2020)
This video demonstrates the process of preparing wedge-shaped brain slices in a laboratory setting. Using a vibratome, a wedge-shaped slice from a mouse&#39;s brain is crafted with varying thickness. This slice contains the auditory nerve root positioned on the thicker end, while the hearing-related neurons are located on the thinner end.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Brain ...

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

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