Name: cochlear surface preparation in the adult mouse
Uploaded: 2019-11-06T16:00:00
Description: Watch this Scientific Journal Video about Cochlear Surface Preparation in the Adult Mouse at JoVE.com

The research project described was supported by grant R01 DC009222 from the National Institute on Deafness and Other Communication Disorders, National Institutes of Health. This work was conducted in the WR Building at MUSC in renovated space supported by grant C06 RR014516. Animals were housed in MUSC CRI animal facilities supported by grant C06 RR015455 from the Extramural Research Facilities Program of the National Center for Research Resources. The authors thank Dr. Jochen Schacht for his valuable comments and Andra Talaska for proofreading of the manuscript.

All research protocols involving male adult CBA/J mice at the ages of 10&#8211;12 weeks and C57BL/6J mice at the ages of 6&#8211;8 weeks were approved by the Institutional Animal Care and Use Committee (IACUC) at the Medical University of South Carolina (MUSC). Animal care was under the supervision of the Division of Laboratory Animal Resources at MUSC.
NOTE: For the procedures presented below, mice are anesthetized with ketamine (100 mg/kg) and xylazine (10 mg/kg) via intrape...

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Cochlear microdissection of whole-mount surface preparations in combination with immunolabeling provides a basic tool for investigation of inner ear pathologies and molecular mechanisms. This modified adult mouse cochlear dissection method using the cell and tissue adhesive simplifies this difficult procedure.
Although this modified cochlear surface preparation method is relatively easy and accessible, it still requires practice in order to achieve proficiency. To make the correct cuts, the di...

The cochlea is dedicated to the detection of sound and responsible for hearing. The cochlear duct is coiled in a spiral shape in the bony labyrinth and holds the auditory sensory end organ, the organ of Corti (OC). The OC rests on the basilar membrane, making up the cochlear epithelium, with a length of about 5.7 mm when uncoiled in adult CBA/CaJ mice1,2. Because the OC is tonotopically organized with high frequencies detected in the base and low frequencies in the apex, the cochlear epithelium is often divided into three parts for analytical comparisons: the apical, middle, and basal turns corresponding to low, middle, and high frequency detection, respectively. In addition to an array of supporting cells, the OC is composed of one row of inner hair cells (IHCs) located medially and three rows of outer hair cells (OHCs) located laterally with respect to the cochlear spiral.
Correct auditory processing depends on the integrity of the sensory hair cells in the cochlea. Damage to or loss of sensory hair cells is a common pathological feature of acquired hearing loss, caused by numerous etiologies such as exposure to excessive noise, the use of ototoxic medications, bacterial or viral ear infections, head injuries, and the aging process3. Additionally, the integrity and function of the inner hair cell/auditory nerve synapses can be impaired by mild insults4. With the availability of molecular and genetic information and manipulation of genes by knockout and knock-in techniques, mice have been widely used in hearing science. Although the adult mouse cochlea is minuscule and the cochlear epithelium is surrounded by a bony capsule resulting in technically difficult microdissections, surface preparations of the epithelium in combination with immunolabeling or immunohistochemistry and confocal imagery have been broadly used for investigation of cochlear pathologies, including losses of ribbon synapses and hair cells, changes in levels of proteins in sensory hair cells and supporting cells, and hair cell regeneration. Cochlear surface preparations have also been used to determine the pattern of expression of reporter genes (i.e., GFP) and confirm successful transduction and identify transduced cell types. These techniques have been previously used for the study of molecular mechanisms underlying noise-induced hearing loss using adult CBA/J mice5,6,7,8,9.
Unlike immunohistochemistry using paraffin sections or cryosections to obtain small cross-sectional portions of the cochlea that contain three outer hair cells (OHCs) and one inner hair cell (IHC) on each section, cochlear surface preparations allow visualization of the entire length of the OC for counting sensory hair cells and ribbon synapses and immunolabeling of sensory hair cells corresponding to specific functional frequencies. Table 1 shows the mapping of hearing frequencies as a function of distance along the length of the cochlear spiral in adult CBA/J mouse according to studies from Muller1 and Viberg and Canlon1,2. Cochlear surface preparations have been widely used for investigation of cochlear pathologies4,5,6,7,8,9,10,11,12,13,14,15. The whole-mount cochlear dissection method was originally described in a book edited by Hans Engstrom in 196616. This technique was subsequently refined and adapted to a variety of species as described in the literature by a number of scientists in hearing science10,11,12,13,15,17 and by the Eaton-Peabody Laboratories at Massachusetts Eye and Ear18. Recently, another cochlear dissection method was reported by Montgomery et al.19. Microdissection of the cochlea is an essential and critical step for cochlear surface preparations. However, dissecting mouse cochleae is a technical challenge and requires considerable practice. Here, a modified cochlear surface preparation method is presented for use in adult mouse cochleae. This method requires decalcification and use of cell and tissue adhesive (i.e., Cell-Tak) to adhere the pieces of cochlear epithelium to 10 mm round coverslips for immunolabeling. Cell and tissue adhesive has been widely used for immunohistochemistry20. This modified cochlear microdissection method is relatively simple compared to those previously reported18,19.

<table>
	<tbody>
		<tr>
			<td>10-mm Rund Coverslips</td>
			<td>Microscopy products for science and industry</td>
			<td>260367</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 Goat Anti-mouse IgG2</td>
			<td>Thermo Fisher Scientific</td>
			<td>A-21131</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 Phalloidin</td>
			<td>Thermo Fisher Scientific</td>
			<td>A12379</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 594 Goat Anti-mouse IgG1</td>
			<td>Thermo Fisher Scientific</td>
			<td>A-21125</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 594 Goat Anti-rabbit IgG (H+L)</td>
			<td>Thermo Fisher Scientific</td>
			<td>A11012</td>
			<td></td>
		</tr>
		<tr>
			<td>Carboard Micro Slide Trays</td>
			<td>Fisher Scientific</td>
			<td>12-587-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell-Tak</td>
			<td>BD Biosciences</td>
			<td>354240</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning Petri Dishes</td>
			<td>Fisher Scientific</td>
			<td>353004</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Thermo Fisher Scientific</td>
			<td>62247</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #5 Forceps</td>
			<td>FST fine science tools</td>
			<td>11251-20</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA Disodium Salt</td>
			<td>Sigma-Aldrich</td>
			<td>E5134</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoro-gel with Tris Buffer</td>
			<td>Electron Microscopy Sciences</td>
			<td>17985-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Four-well Cell Culture Dishes</td>
			<td>Greiner Bio-One</td>
			<td>627170</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat Anti-myosin VIIa Antibody</td>
			<td>Proteus Biosciences</td>
			<td>25-6790</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Slides</td>
			<td>Fisher Scientific</td>
			<td>12-544-7</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse Anti-CtBP2 Antibody</td>
			<td>BD Biosciences</td>
			<td>#612044</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse Anti-Glu2R Antibody</td>
			<td>Millipore</td>
			<td>MAB397</td>
			<td></td>
		</tr>
		<tr>
			<td>Normal Goat Serum</td>
			<td>Thermo Fisher Scientific</td>
			<td>31872</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>441244</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline</td>
			<td>Fisher Scientific</td>
			<td>BP665-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>VWR</td>
			<td>100491-038</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>X100-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Vannas Spring Scissors</td>
			<td>Fine Science Tools</td>
			<td>15001-08</td>
			<td></td>
		</tr>
	</tbody>
</table>

cochlear surface preparation in the adult mouse

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous etiologies such as exposure to excessive noise, the use of ototoxic medications, bacterial or viral ear infections, head injuries, and the aging process. Loss of sensory hair cells is a common pathological feature of the varieties of acquired hearing loss. Additionally, the inner hair cell synapse can be damaged by mild insults. Therefore, surface preparations of cochlear epithelia, in combination with immunolabeling techniques and confocal imagery, are a very useful tool for the investigation of cochlear pathologies, including losses of ribbon synapses and sensory hair cells, changes in protein levels in hair cells and supporting cells, hair cell regeneration, and determination of report gene expression (i.e., GFP) for verification of successful transduction and identification of transduced cell types. The cochlea, a bony spiral-shaped structure in the inner ear, holds the auditory sensory end organ, the organ of Corti (OC). Sensory hair cells and surrounding supporting cells in the OC are contained in the cochlear duct and rest on the basilar membrane, organized in a tonotopic fashion with high-frequency detection occurring in the base and low-frequency in the apex. With the availability of molecular and genetic information and the ability to manipulate genes by knockout and knock-in techniques, mice have been widely used in biological research, including in hearing science. However, the adult mouse cochlea is miniscule, and the cochlear epithelium is encapsulated in a bony labyrinth, making microdissection difficult. Although dissection techniques have been developed and used in many laboratories, this modified microdissection method using cell and tissue adhesive is easier and more convenient. It can be used in all types of adult mouse cochleae following decalcification.

Surface preparations of the cochlear epithelium, in combination with immunolabeling and confocal imaging, have been used broadly in hearing science for the investigation of cochlear pathologies, such as quantification of ribbon synapses, sensory hair cells, and protein expression in sensory hair cells5,6,7,8. Although the dissection of adult mouse cochleae for surface preparations is not simple...

Watch this Scientific Journal Video about Cochlear Surface Preparation in the Adult Mouse at JoVE.com

Cochlear Surface Preparation in the Adult Mouse

This article presents a modified cochlear surface preparation method that requires decalcification and use of a cell and tissue adhesive to adhere the pieces of cochlear epithelia to 10 mm round cover slips for immunohistochemistry in adult mouse cochleae.

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous ...

Cochlear surface preparations allow visualization of the entire lens of the organ of Corti using immunohistochemistry and confocal imaging. It has been broadly used for investigation of specific cochlear pathologies of interest. We will modify the cochlear microdissection method.The main advantage of this technique is the adherence of a piece of cochlear epithelium to 10 millimeter round coverslips for the immunolabeling procedure while avoiding tissue loss during the multiple washing steps. In addition cochlear pathologies, cochlear surface preparation can also be utilized for the assessment expression and hair cell regeneration. Basic microscope skills are required for this technique and visual demonstration of the method can help to ensure that each step is performed correctly.Demonstrating the procedure will be Qiaojun Fang, a graduate student from my laboratory. For temporal bone extraction, immediately post-mortem pull the skin anteriorly to expose the skull bone and use scissors to cut the bone from the posterior aspect forward along the center line of the skull. Use forceps to remove the brain tissue before using the thumb and index finger to manually remove the temporal bones from mice three months of age or older.Place the bones into a 30 millimeter diameter Petri dish containing fresh ice cold 4%PFA and PBS under a dissection microscope and remove the stapes from the oval window. Puncture the round window membrane with number five forceps and use a 27 gauge needle attached to a one milliliter syringe containing fresh 4%PFA to make a small hole into the apex of the cochlea. Gently and slowly perfuse the cochlea with a fixative solution via the round and oval windows until the solution washes out of the small hole at the apex.Then transfer up to two perfused cochleae into individual 20 milliliter scintillation vials containing 10 milliliters of 4%PFA per vial and gently agitate the samples for two hours at room temperature, followed by overnight storage at four degrees Celsius on a rotator. The next morning, wash the cochleae with three five-minute washes with fresh PBS per wash. After the last wash, add 20 milliliters of 4%EDTA to each vial and rotate the samples for 48 hours at four degrees Celsius with gentle agitation.At the end of the incubation, touch the bony vestibular portion of each cochlea with forceps to assess the elasticity of the samples. If the bones are elastic rather than firm, the cochlea have been decalcified. Replace the EDTA for each sample with fresh PBS.For microdissection of the cochlear epithelium, grasp the vestibular portion of the temporal bone with forceps and use a scalpel to cut the apical turn at a 45 degree angle. Cut vertically along the faint line between the round window and the oval window to separate the cochlea from the vestibular portion and orient the cochlear portion with the basal turn toward the bottom and middle turns toward the top of the Petri dish. From this position, cut the bony capsule and lateral wall of the middle turn toward the end from which the apical section was removed and continue cutting to completely separate the middle portion from the basal and hook regions.Placing the basal and hook portion with the basilar membrane site oriented toward the bottom of the dish, vertically cut the medialis off of the hook region to remove the medialis and cut the basal and hook regions to separate the hook portion. To avoid distortion of the tissue, cut the medialis off of the hook region before separation of the basal turn and hook region. Cut away the relatively large portions of bony capsule and lateral wall tissue of the middle turn and holding the lateral wall with forceps to align the bony capsule and lateral wall with the bottom of the Petri dish, cut these tissues from the basilar membrane side.Next, flatten the specimen to orient the sensory hair cell surface side up and trim away the rest of the bony capsule and lateral wall. Then use forceps to remove the tectorial membrane to completely separate the middle region and remove the bony capsule, lateral wall tissue, and tectorial regions of the remaining turns as just demonstrated. When all of the cochlear turns have been dissected, spread 0.5 microliters of cell and tissue adhesive onto the center of one round 10 millimeter coverslip and allow the adhesive to dry for three to five minutes at room temperature.Place the dried coverslips into the Petri dish and stick all four pieces of each sensory epithelium sample onto one coverslip. Then use forceps to grasp one edge of each coverslip to transfer the specimens into individual wells of a four-well cell culture dish for immunolabeling. For immunolabeling of surface cochlear synapse preparations, wash each sensory epithelium three times for five minutes in fresh PBS per wash, followed by the treatment of each sample with two milliliters of 2%non-ionic surfactant for 30 minutes at room temperature on a rotator.At the end of the incubation, aspirate the surfactant solution from each well and block any nonspecific binding with 100 microliters of blocking solution per well for one hour on the rotator with gentle agitation at room temperature. At the end of the blocking incubation, wash the samples three times with PBS as demonstrated under gentle agitation before labeling each epithelium with 100 microliters of the primary antibodies of interest for 24 hours at 37 degrees Celsius protected from light. The next day, wash the samples three times with fresh PBS and gentle agitation per wash and label the specimens with 100 microliters of the appropriate secondary antibodies of interest for two hours at 37 degrees Celsius protected from light.At the end of the incubation, wash the samples with three five-minute washes in fresh PBS per wash before placing each coverslip onto individual glass microscope slides with the samples facing up. Next, carefully add eight microliters of an appropriate mounting medium into the center of each coverslip and use forceps to mount another 10 millimeter coverslip onto each sample. Then seal the sides of each coverslip sandwich with clear nail polish, place the samples into a cardboard slide folder, and store the slides at four degrees Celsius.Although the dissection of adult mouse cochleae for surface preparations is not simple, researchers new to the technique should be able to learn the method after practicing with 10 to 15 ears. Immunolabeling of surface preparations of untreated 10 to 12-week-old CBAJ mice with CTBP2 and GluA2 demonstrates that both presynaptic ribbons and postsynaptic terminals respectively are located below the inner hair cell nuclei and are juxtaposed indicating functional synapses. Immunolabeling for myosin 7A and counterstaining with phalloidin and DAPI reveals the presence of sensory hair cells including the outer hair cells and inner hair cells and their nuclei.Immunolabeling for myosin 7A and counterstaining with phalloidin shows no differences in immunoreactivity or uniformity with or without the use of cell and tissue adhesive. In addition, scanning electron microscopy of surface preparations from untreated six to eight-week-old C57 black 6 mice demonstrate a well-organized V-shaped stereocilia of outer hair cells in three rows of the sample. Remember to fill the cochlear with fixative solution gently and slowly through the round and oval windows until the solution washes out of the small hole at the apex.Before separating the hook region from the basal turn, take care to cut the medialis off of the hook region to facilitate the removal of the medialis.

cochlear-surface-preparation-in-the-adult-mouse

Research

JoVE Journal

Neuroscience

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls gastrointestinal motility. A procedure for the isolation and culture of a mixed population of neurons and glia from the myenteric plexus is described. The primary cultures can be maintained for over 7 days, with connections developing among the neurons and glia. The longitudinal muscle strip with the attached myenteric plexus is stripped from the underlying circular muscle of the mouse ileum or colon and subjected to enzymatic digestion. In sterile conditions, the isolated neuronal and glia population are preserved within the pellet following centrifugation and plated on coverslips. Within 24-48 hr, neurite outgrowth occurs and neurons can be identified by pan-neuronal markers. After two days in culture, isolated neurons fire action potentials as observed by patch clamp studies. Furthermore, enteric glia can also be identified by GFAP staining. A network of neurons and glia in close apposition forms within 5 - 7 days. Enteric neurons can be individually and directly studied using methods such as immunohistochemistry, electrophysiology, calcium imaging, and single-cell PCR. Furthermore, this procedure can be performed in genetically modified animals. This methodology is simple to perform and inexpensive. Overall, this protocol exposes the components of the enteric nervous system in an easily manipulated manner so that we may better discover the functionality of the ENS in normal and disease states.

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

We demonstrate a cell culture protocol for the direct study of neuronal and glial components of the enteric nervous system. A neuron/glia mixed culture on coverslips is prepared from the myenteric plexus of adult mouse providing the ability to examine individual neuron and glia function by electrophysiology, immunohistochemical, etc.

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls ...

An In Vitro Adult Mouse Muscle-nerve Preparation for Studying the Firing Properties of Muscle Afferents

Muscle sensory neurons innervating muscle spindles and Golgi tendon organs encode length and force changes essential to proprioception. Additional afferent fibers monitor other characteristics of the muscle environment, including metabolite buildup, temperature, and nociceptive stimuli. Overall, abnormal activation of sensory neurons can lead to movement disorders or chronic pain syndromes. We describe the isolation of the extensor digitorum longus (EDL) muscle and nerve for in vitro study of stretch-evoked afferent responses in the adult mouse. Sensory activity is recorded from the nerve with a suction electrode and individual afferents can be analyzed using spike sorting software. In vitro preparations allow for well controlled studies on sensory afferents without the potential confounds of anesthesia or altered muscle perfusion. Here we describe a protocol to identify and test the response of muscle spindle afferents to stretch. Importantly, this preparation also supports the study of other subtypes of muscle afferents, response properties following drug application and the incorporation of powerful genetic approaches and disease models in mice.

An In Vitro Adult Mouse Muscle-nerve Preparation for Studying the Firing Properties of Muscle Afferents

Muscle sensory neurons are involved in proprioceptor signaling and also report on metabolic state and injury related events. We describe an adult mouse in vitro muscle-nerve preparation for studies on stretch-activated muscle afferents.

Muscle sensory neurons innervating muscle spindles and Golgi tendon organs encode length and force changes essential to proprioception. Additional ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...

Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a spiral shape and have a tonotopic gradient for sound detection. The mouse cochlea is approximately 6 mm long and often divided into three turns (apex, middle, and base) for analysis. To investigate cell loss, cell division, or mosaic gene expression, the whole mount or surface preparation of the cochlea is useful. This dissection method allows visualization of all cells within the organ of Corti when combined with immunostaining and confocal microscopy to image cells at different planes in the z-axis. Multiple optical cross-sections can also be obtained from these z-stack images. In addition, the whole mount dissection method can be used for scanning electron microscopy, although a different fixation method is needed. Here, we present a method to isolate the organ of Corti as three intact cochlear turns (apex, middle, and base). This method can be used for mice ranging from one week of age through adulthood and differs from the technique used for neonatal samples where calcification of the cochlea is incomplete. A slightly modified version can be used for dissection of the rat cochlea. We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies.

We present a method to isolate the adult organ of Corti as three intact cochlear turns (apex, middle, and base). We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies. Together these techniques allow the study of hair cells, supporting cells, and other cell types found in the cochlea.

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a ...

Preparation of Primary Mixed Glial Cultures from Adult Mouse Spinal Cord Tissue

It has been well-accepted that spinal cord glial responses contribute significantly to the development of neuropathic pain. Tremendous information regarding glial activities at the cellular and molecular levels has been obtained through in vitro cell culture systems. The in vitro systems utilized, mainly include primary glia derived from neonatal brain cortical tissue and immortalized cell lines. However, these systems may not reflect the characteristics of spinal cord glial cells in vivo. In order to further investigate the roles of spinal cord glial cells in the development of peripheral nerve injury-induced neuropathic pain using a culture system that better reflects the in vivo condition, our laboratory has developed a method to establish primary spinal cord mixed glial cultures from adult mice. Briefly, spinal cords are collected from adult mice and processed through papain digestion followed by myelin removal with a density-gradient medium. Single cell suspensions are cultured in complete Dulbecco&#39;s modified Eagle media (cDMEM) supplemented with 2-mercaptoethanol (2-ME) at 35.9 oC. These culture conditions were optimized specifically for the growth of mixed glial cells. Under these conditions, cells are ready to be used for experimentation between 12 - 14 d&#160;(cells are usually in log phase during this time) after the establishment of the culture (D 0) and can be kept in culture conditions up to D 21. This culture system can be used to investigate the responses of spinal cord glial cells upon stimulation with various substances and agents. Besides neuropathic pain, this system can be used to study glial responses in other diseases that involve pathological changes of spinal cord glial cells.

The development of neuropathic pain involves pathological changes of spinal cord glial cells. A reliable glial culture system derived from adult spinal cord tissue and designed for studying these cells in vitro is lacking. Therefore, we show here how&#160;to establish primary mixed glial cultures from adult mouse spinal cord tissue.

It has been well-accepted that spinal cord glial responses contribute significantly to the development of neuropathic pain. Tremendous information ...

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

Preparation of Horizontal Slices of Adult Mouse Retina for Electrophysiological Studies

Vertical slice preparations are well established to study circuitry and signal transmission in the adult mammalian retina. The plane of sectioning in these preparations is perpendicular to the retinal surface, making it ideal for the study of radially oriented neurons like photoreceptors and bipolar cells. However, the large dendritic arbors of horizontal cells, wide-field amacrine cells, and ganglion cells are mostly truncated, leaving markedly reduced synaptic activity in these cells. Whereas ganglion cells and displaced amacrine cells can be studied in a whole-mounted preparation of the retina, horizontal cells and amacrine cells located in the inner nuclear layer are only poorly accessible for electrodes in whole retina tissue.
To achieve maximum accessibility and synaptic integrity, we developed a horizontal slice preparation of the mouse retina, and studied signal transmission at the synapse between photoreceptors and horizontal cells. Horizontal sectioning allows&#160;(1) easy and unambiguous visual identification of horizontal cell bodies for electrode targeting, and (2) preservation of the extended horizontal cell dendritic fields, as a prerequisite for intact and functional cone synaptic input to horizontal cell dendrites.
Horizontal cells from horizontal slices exhibited tonic synaptic activity in the dark, and they responded to brief flashes of light with a reduction of inward current and diminished synaptic activity. Immunocytochemical evidence indicates that almost all cones within the dendritic field of a horizontal cell establish synapses with its peripheral dendrites. The horizontal slice preparation is therefore well suited to study the physiological properties of horizontally extended retinal neurons as well as sensory signal transmission and integration across selected synapses.

We developed a horizontal slice preparation of the adult mammalian retina with the plane of sectioning parallel to the retinal surface. Dendritic fields of nonradially oriented retinal neurons were not truncated, allowing the study of signal processing with patch-clamp and imaging techniques.

Vertical slice preparations are well established to study circuitry and signal transmission in the adult mammalian retina. The plane of sectioning in ...

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the dorsal root ganglion (DRGs) extends a single axon with two branches. One branch goes to the peripheral nerve (peripheral branch), and the other branch enters the spinal cord through the dorsal root (central branch). After the neural injury, the peripheral branch regenerates robustly whereas the central branch does not regenerate. Due to the high regenerative capacity, sensory axon regeneration has been widely used as a model system to study mammalian axon regeneration in both the peripheral nervous system (PNS) and the central nervous system (CNS). Here, we describe a previously established approach protocol to manipulate gene expression in mature sensory neurons in vivo via electroporation. Based on transfection with plasmids or small RNA oligos (siRNAs or microRNAs), the approach allows for both loss- and gain-of-function experiments to study the roles of genes-of-interests or microRNAs in regulation of axon regeneration in vivo. In addition, the manipulation of gene expression in vivo can be controlled both spatially and temporally within a relatively short time course. This model system provides a unique tool to investigate the molecular mechanisms by which mammalian axon regeneration is regulated in vivo.

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the ...

Isolation of Adult Spinal Cord Nuclei for Massively Parallel Single-nucleus RNA Sequencing

Probing an individual cell&#39;s gene expression enables the identification of cell type and cell state. Single-cell RNA sequencing has emerged as a powerful tool for studying transcriptional profiles of cells, particularly in heterogeneous tissues such as the central nervous system. However, dissociation methods required for single cell sequencing can lead to experimental changes in the gene expression and cell death. Furthermore, these methods are generally restricted to fresh tissue, thus limiting studies on archival and bio-bank material. Single nucleus RNA sequencing (snRNA-Seq) is an appealing alternative for transcriptional studies, given that it accurately identifies cell types, permits the study of tissue that is frozen or difficult to dissociate, and reduces dissociation-induced transcription. Here, we present a high-throughput protocol for rapid isolation of nuclei for downstream snRNA-Seq. This method enables isolation of nuclei from fresh or frozen spinal cord samples and can be combined with two massively parallel droplet encapsulation platforms.

Here, we present a protocol to rapidly isolate high-quality nuclei from the fresh or frozen tissue for downstream massively parallel RNA sequencing. We include detergent-mechanical and hypotonic-mechanical tissue disruption and cell lysis options, both of which can be used for isolation of nuclei.

Probing an individual cell&#39;s gene expression enables the identification of cell type and cell state. Single-cell RNA sequencing has emerged as a ...