This work is supported by an R21 grant from the National Institutes of Health and by a grant from Hearing Research, Incorporated.

NOTE: All procedures and animal handling complied with NIH ethics guidelines and approved protocol requirements of the Institutional Animal Care and Use Committee of the University of California, San Francisco.
1. Preparing the Animal for Surgery
<ol>
	<li>Carry out surgical procedures in a clean, dedicated space. Autoclave all surgical instruments, sterilize with a glass-bead sterilizer&#160;prior to surgery. 
	NOTE: In this protocol, use postnatal day 10-12 (P10-12) FVB mice. Different ages and strains of mice can be used to meet the needs of a specific project. Mice older than P40 can be challenging because the bulla bone gets harder.</li>
	<li>Anesthetize the mice with intraperitoneal injection of a mixture of ketamine hydrochloride (100 mg/kg), xylazine hydrochloride (10 mg/kg), and acepromazine (2 mg/kg). Begin surgical preparation only after the animal no longer responds to painful stimuli, such as toe pinch. If necessary, administer a booster dose (one-fifth the original dose) of the anesthetic cocktail to restore the original anesthetic plane. 
	NOTE: Take care at this step because most of the mortality observed at this age is caused by anesthetic overdose.</li>
	<li>Cover the animal&#39;s eyes with a protective ophthalmic ointment to keep the eyes moist during anesthesia, suppressing the animal&#39;s blink reflex.</li>
	<li>Position the mouse with neck extended on a heating pad throughout the procedure until the mouse is totally awake, to prevent post-anesthesia hypothermia.</li>
	<li>Shave the left post-auricular region with a clipper and disinfect with 70% ethanol and povidone-iodine before surgical manipulation.</li>
</ol>
2. The Surgical Procedure and Vector Injection
<ol>
	<li>Use a post-auricular approach to expose the tympanic bulla. Incise the subcutaneous tissue with small scissors to expose the post-auricular muscle. After retracting the adipose tissue to the posterior side of the incision, separate the muscles to the right and left side perpendicular to the incision to expose the temporal bone. Ensure that this incision is long enough to work comfortably with adequate visualization.</li>
	<li>Perforate the tympanic bulla with a 25 G needle and expand the hole as necessary with forceps by peeling the bone back to allow access to the basal turn of the cochlea, and then widen sufficiently to visualize the stapedial artery and the round window membrane (RWM).</li>
	<li>Puncture the RWM gently in the center with a borosilicate capillary pipette. Observe fluid efflux through the RWM at this point; this is normal. Wait until the efflux has stabilized (the effluxed fluid from the cochlea is dried with a sterile filter paper). 
	NOTE: Take care when holding and advancing the glass needle so that the RWM perforation is as small as possible. Depending on the microscope used, hold the pipettes using a micromanipulator or by hand using a pipette holder.</li>
	<li>Prepare the pipettes for injection using a pipette puller the same way patch clamp pipettes are prepared. Ensure that the tip diameter is large (about 15 &#181;m in diameter) so that pipetting the virus in and out is done easily.</li>
	<li>Draw up 1 to 2 &#181;l of AAV1-VGLUT3, AAV1-GFP or AAV2-GFP into an injection pipette. After the efflux is stabilized (5 to 10 min), microinject this fixed volume into the scala tympani through the same hole previously made in RWM.</li>
	<li>Seal the RW niche quickly after pulling out the pipette with a small plug of muscle and secure it with a small drop of tissue adhesive placed on the muscle to avoid leakage from the round window. 
	NOTE : Seal this hole very well after the needle is removed with a small plug of muscle&#160;to prevent any perilymph leakage from the cochlea-this step is critical to preserve hearing. Failure to seal completely the RWM will result in hearing loss over time.</li>
	<li>After injection, cover the hole in the auditory bulla with adipose tissue, return the post-auricular muscles and adipose tissue to their normal position and suture the wound in layers with a 6-0 or smaller absorbable chromic suture.</li>
	<li>Disinfect the wound with povidone-iodine.</li>
</ol>
3. Postoperative Care
<ol>
	<li>Place mice in a warm cage and do not leave them unattended until they are fully recovered then&#160;move them back with the mother. It is recommended to remove the male from the cage before putting the pups back with the mother.</li>
	<li>Administer subcutaneous Carprofen (2 mg/kg) for analgesia postoperatively and every 24 hr thereafter for 3 days, to manage inflammation and pain. Monitor the animals daily for signs of distress, abnormal weight loss, pain, or infection. Generally by the 3rd day after the surgery all mice should be acting normally. If any signs of distress or disease appear in a mouse after the 3rd day, consider euthanizing the mouse as per institutional guidelines.</li>
</ol>
4. Assessment of Cochlear Function Following Viral Delivery Using Auditory Brainstem Response (ABR) Recordings
NOTE: The auditory brainstem response (ABR)&#160;is an&#160;auditory evoked potential&#160;extracted from ongoing electrical activity in the brain and recorded via electrodes placed under the scalp. The animal is stimulated with sound. The resulting recording consists of five waves that reflect the electrical activity of successive points in the auditory pathway in the first 10 msec after onset of an auditory stimulus.
<ol>
	<li>Anesthetize mice by intraperitoneal injection of a mixture of ketamine hydrochloride and xylazine hydrochloride as described in section 1.2 of this protocol, except without acepromazine.</li>
	<li>Place the mouse on a heating pad throughout the hearing test until the mouse is totally awake, to prevent post-anesthesia hypothermia. 
	NOTE: Use the following sound stimuli in this experiment: click (5 msec duration; 31 Hz presentation rate).</li>
	<li>Place subdermal needle electrodes at the vertex, below the pinna of the left ear (reference), and below the contralateral ear (ground) and start recording ABRs as previously described in a sound-proof chamber27,28. Record ABR waveforms in 5 dB sound pressure level intervals down from maximum amplitude.</li>
	<li>Determine ABR thresholds postoperatively as early as 4 days after viral delivery 
	NOTE: Threshold is defined as the lowest stimulus level at which response peaks for waves I&#8211;V were clearly and repeatedly present on visual inspection.</li>
	<li>Measure Wave I to analyze the activity from the cochlear nerve. The lowest stimulus level that yields a detectable ABR waveform is defined as the threshold.</li>
</ol>
5. Cochlear Transgene Protein Expression Using Immunofluorescence
<ol>
	<li>Anesthetize mice by an overdose of intraperitoneal injection of a mixture of ketamine hydrochloride and xylazine hydrochloride as described in section 1.2 of this protocol, except without acepromazine.</li>
	<li>Decapitate the head only after the animal no longer responds to painful stimuli, such as toe pinch.</li>
	<li>Dissect the cochleae out and process for whole-mount immunofluorescence as described26. A detailed protocol&#160;of cochlear whole-mount immunofluorescence using anti-VGLUT3 and anti-Myosin 7a antibodies is described in Akil et al., 201329.</li>
	<li>For GFP labeling, incubate the cochlear whole-mounts overnight at 4 &#176;C with a rabbit anti-GFP antibody at 1:250 dilution.</li>
	<li>Rinse the cochleae twice for 10 min with PBS and then incubate for 2 hours in goat anti-rabbit IgG conjugated to Cy2 diluted 1:4,000 in PBS.</li>
	<li>Rinse the cochleae with PBS twice for 10 minutes and incubate them with DAPI for 15 min.</li>
	<li>Mount the cochleae on glass slides and observe under a microscope with confocal immunofluorescence.</li>
</ol>

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No conflicts of interest declared.

In this work, we describe in detail a technique that can be used for cochlear gene therapy, with the goal of restoring or rescuing normal auditory function that is compromised by a genetic defect. As it is typically atraumatic, this approach is safe for cochlear gene transfer or other potential molecular therapies30. Other approaches for cochlear therapy have been described, including a ventral approach24, cochleostomy31,32 and endolymphatic sac delivery33 in mouse and guinea p...

Gene therapy has long been suggested as a potential treatment for genetic hearing loss, but success in this area has remained elusive1. To date, virally mediated methodologies have predominated due the theoretic ability to target specific cell types within the relatively inaccessible cochlea. Both adenovirus (AV) and adeno-associated virus (AAV) &#160;have been used for cochlear gene delivery. AAVs are advantageous in the cochlea for a number of reasons. They are replication-deficient viruses and can efficiently transfer transgenic molecules to different cell types including neurons, an important target for a number of causes of hearing loss. AAV entry into the cell is mediated by specific receptors2; thus, the choice of a particular serotype must be compatible with the cell types to be transduced. AAVs can effectively transfect hair cells3 and incorporate into the host genome, resulting in stable, long-term expression of the transgenic protein and phenotypic change in the cell4. While not necessarily advantageous for short-term applications such as hair-cell regeneration, long-term expression is very important for stable rescue of genetic defects. Because AAVs are not associated with any human disease or infection and demonstrate no ototoxicity5,6,7, they are an ideal candidate for use in gene therapy for inherited forms of hearing loss8.
Transfer of exogenous genetic material into the mammalian inner ear using viral vectors has been studied over the last decade and is emerging as a promising technique for treating both genetic and acquired forms of hearing loss9. The cochlea is potentially an ideal target for gene therapy for several reasons: 1) its small volume necessitates&#160;a limited amount of the virus needed; 2) its relative isolation from other organ systems limits side effects; and 3) its fluid-filled chambers facilitate viral delivery throughout the labyrinth10,11,12,13,14,15.
Mouse models of congenital deafness allow for use of many methods of study to monitor development of the inner ear in a systematic, replicable way. While the small size of mouse cochleae does present some surgical difficulty, the mouse serves as an extremely important model in the study of genetic hearing loss, with several experimental advantages over other species16. Mouse models allow assessment of a range of characteristics through genetic linkage analysis, collection of detailed morphological observations, and simulating pathogenic scenarios; as such, they are good candidates for virally mediated gene therapy. Extensive genetic studies in mice combined with technological advances have made it possible to generate genetically modified mice in a reproducible way across laboratories17,18,19,20,21. Furthermore, there exist numerous models for both acquired and inherited hearing loss phenotypes in mice, allowing rigorous testing in this animal model22,23,24. Thus, correcting hearing using virally mediated gene therapy in a mouse model is an appropriate first step in the search for a cure for human disease.
We have previously shown that transgenic mice lacking vesicular glutamate transporter 3 (VGLUT3) are born deaf due to lack of glutamate release at the IHC ribbon synapse25. Because this mutation does not lead to a primary degeneration of the sensory hair cells, these mutant mice are &#160;potentially an excellent model in which to test cochlear gene therapy for congenital hearing loss.
To date, a number of viral delivery techniques for cochlear gene therapy have been described, including round window membrane diffusion, round window membrane injection, and delivery via a cochleostomy. There are potential advantages and disadvantages of each of these approaches9.
Here we report a surgical method for virally mediated gene delivery to the VGLUT3 KO mouse inner ear through the round window membrane (RWM). The post-auricular RWM injection method is minimally invasive with excellent hearing preservation, and is relatively fast. As we have previously published, in an effort to restore hearing in this mouse model, an AAV1 vector carrying the VGLUT3 gene (AAV1-VGLUT3) was introduced into the cochlea of these deaf mice at postnatal day 12(P!@), resulting in the restoration of hearing26. Hearing in the VGLUT3 KO mice was verified by auditory brainstem response (ABR), while transgene protein expression was verified using immunofluorescence (IF). This methodology thus demonstrates that virally-mediated gene therapy can correct a genetic defect that would otherwise results in deafness.

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			<td>&#160;Tucker-Davis Technologies</td>
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surgical method for virally mediated gene delivery to the mouse inner ear through the round window membrane

Gene therapy, used to achieve functional recovery from sensorineural deafness, promises to grant better understanding of the underlying molecular and genetic mechanisms that contribute to hearing loss. Introduction of vectors into the inner ear must be done in a way that widely distributes the agent throughout the cochlea while minimizing injury to the existing structures. This manuscript describes a post-auricular surgical approach that can be used for mouse cochlear therapy using molecular, pharmacologic, and viral delivery to mice postnatal day 10 and older via the round window membrane (RWM). This surgical approach enables rapid and direct delivery into the scala tympani while minimizing blood loss and avoiding animal mortality. This technique involves negligible or no damage to essential structures of the inner and middle ear as well as neck muscles while wholly preserving hearing. To demonstrate the efficacy of this surgical technique, the vesicular glutamate transporter 3 knockout (VGLUT3 KO) mice will be used as an example of a mouse model of congenital deafness that recovers hearing after delivery of VGLUT3 to the inner ear using an adeno-associated virus (AAV-1).

To verify the technical features and utility of the post-auricular approach for cochlear molecular therapy, AAV1-VGLUT3, AAV1-GFP and AAV2-GFP were delivered into P10-12 mice inner ear via the RWM. This approach demonstrates successful transgene expression within inner hair cells (IHC) (VGLUT3 Figure 1 and GFP Figure 2 and GFP Figure 3A), outer hair cells (OHC) (GFP Figure 2) and supporting cells (GFP Figure 2 and Figure 3A</stro...

Watch this Scientific Journal Video about Surgical Method for Virally Mediated Gene Delivery to the Mouse Inner Ear through the Round Window Membrane at JoVE.com

Surgical Method for Virally Mediated Gene Delivery to the Mouse Inner Ear through the Round Window Membrane

The described post-auricular surgical approach allows rapid and direct delivery into the mouse cochlear scala tympani while minimizing blood loss and animal mortality. This method can be used for cochlear therapy using molecular, pharmacologic and viral delivery to postnatal mice through the round window membrane.

Gene therapy, used to achieve functional recovery from sensorineural deafness, promises to grant better understanding of the underlying molecular and ...

surgical-method-for-virally-mediated-gene-delivery-to-mouse-inner-ear

Research

JoVE Journal

Neuroscience

15.2K vues.  University of California, San Francisco. The described post-auricular surgical approach allows rapid and direct delivery into the mouse cochlear scala tympani while minimizing blood loss and animal mortality. This method can be used for cochlear therapy using molecular, pharmacologic and viral delivery to postnatal mice through the round window membrane. 

Vidéo: Surgical Method for Virally Mediated Gene Delivery to the Mouse Inner Ear through the Round Window Membrane

Intravital Microscopy of the Mouse Brain Microcirculation using a Closed Cranial Window

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, inflammatory and metabolic conditions, using in vivo fluorescence microscopy. A closed cranial window is placed over the left parieto-occipital cortex of the mice. Local microcirculation is recorded in real time through the window using epi and fluorescence illumination, and measurements of vessels diameters and red blood cell (RBC) velocities are performed. RBC velocity is measured using real-time cross-correlation and/or fluorescent-labeled erythrocytes. Leukocyte and platelet adherence to pial vessels and assessment of perfusion and vascular leakage are made with the help of fluorescence-labeled markers such as Albumin-FITC and anti-CD45-TxR antibodies. Microcirculation can be repeatedly video-recorded over several days. We used for the first time the close window brain intravital microscopy to study the pial microcirculation to follow dynamic changes during the course of Plasmodium berghei ANKA infection in mice and show that expression of CM is associated with microcirculatory dysfunctions characterized by vasoconstriction, profound decrease in blood flow and eventually vascular collapse.

Intravital microscopy to follow temporal and spatial hemodynamic and inflammatory events in the pial microcirculation.

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, ...

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

Isolating LacZ-expressing Cells from Mouse Inner Ear Tissues using Flow Cytometry

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues presents a unique challenge because of the paucity of cells of interest and few transgenic reporter mouse models. Here, we describe a protocol using fluorescence-conjugated probes to selectively label LacZ-positive cells from the neonatal cochleae.
The most common underlying pathology of sensorineural hearing loss is the irreversible damage and loss of cochlear sensory hair cells, which are required to transduce sound waves to neural impulses. Recent evidence suggests that the murine auditory and vestibular organs harbor stem/progenitor cells that may have regenerative potential1,2. These findings warrant further investigation, including identifying specific cell types with stem/progenitor cell characteristics. The Wnt signaling pathway has been demonstrated to play a critical role in maintaining stem/progenitor cell populations in several organ systems3-7. We have recently identified Wnt-responsive Axin2-expressing cells in the neonatal cochlea, but their function is largely unknown8.
To better understand the behavior of these Wnt-responsive cells in vitro, we have developed a method of isolating Axin2-expressing cells from cochleae of Axin2-LacZ reporter mice9. Using flow cytometry to isolate Axin2-LacZ positive cells from the neonatal cochleae, we could in turn execute a variety of experiments on live cells to interrogate their behavior as stem/progenitor cells. Here, we describe in detail the steps for the microdissection of neonatal cochlea, dissociation of these tissues, labeling of the LacZ-positive cells using a fluorogenic substrate, and cell sorting. Techniques for dissociating cochleae into single cells and isolating cochlear cells via flow cytometry have been described2,10-12. We have made modifications to these techniques to establish a novel protocol to isolate LacZ-expressing cells from the neonatal cochlea.

Flow cytometry is a powerful tool allowing for the isolation and study of specific cell populations. This protocol describes steps for isolating LacZ-expressing cells from cochlear tissues from neonatal transgenic mice. Dissociated cochlear cells were labeled using fluorescent-conjugated substrates of &#946;-galactosidase prior to separation via flow cytometry.

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues ...

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and the organ of Corti in the coiled cochlea. The cristae and maculae contain vestibular hair cells that transduce mechanical stimuli to subserve the special sense of balance, while auditory hair cells in the organ of Corti are the primary transducers for hearing 1. Cell fate specification in these sensory epithelia and morphogenesis of the semicircular canals and cochlea take place during the second week of gestation in the mouse and are largely completed before birth 2,3. Developmental studies of the mouse inner ear are routinely conducted by harvesting transgenic embryos at different embryonic or postnatal stages to gain insight into the molecular basis of cellular and/or morphological phenotypes 4,5. We hypothesize that gene transfer to the developing mouse inner ear in utero in the context of gain- and loss-of-function studies represents a complimentary approach to traditional mouse transgenesis for the interrogation of the genetic mechanisms underlying mammalian inner ear development6.
The experimental paradigm to conduct gene misexpression studies in the developing mouse inner ear demonstrated here resolves into three general steps: 1) ventral laparotomy; 2) transuterine microinjection; and 3) in vivo electroporation. Ventral laparotomy is a mouse survival surgical technique that permits externalization of the uterus to gain experimental access to the implanted embryos7. Transuterine microinjection is the use of beveled, glass capillary micropipettes to introduce expression plasmid into the lumen of the otic vesicle or otocyst. In vivo electroporation is the application of square wave, direct current pulses to drive expression plasmid into progenitor cells8-10. 
We previously described this electroporation-based gene transfer technique and included detailed notes on each step of the protocol11. Mouse experimental embryological techniques can be difficult to learn from prose and still images alone. In the present work, we demonstrate the 3 steps in the gene transfer procedure. Most critically, we deploy digital video microscopy to show precisely how to: 1) identify embryo orientation in utero; 2) reorient embryos for targeting injections to the otocyst; 3) microinject DNA mixed with tracer dye solution into the otocyst at embryonic days 11.5 and 12.5; 4) electroporate the injected otocyst; and 5) label electroporated embryos for postnatal selection at birth. We provide representative examples of successfully transfected inner ears; a pictorial guide to the most common causes of otocyst mistargeting; discuss how to avoid common methodological errors; and present guidelines for writing an in utero gene transfer animal care protocol.

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mouse inner ear is a placode-derived sensory organ whose developmental program is elaborated during gestation. We define an in utero gene transfer technique consisting of three steps: mouse ventral laparotomy, transuterine microinjection, and in vivo electroporation. We use digital video microscopy to demonstrate the critical experimental embryological techniques.

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and ...

A Polished and Reinforced Thinned-skull Window for Long-term Imaging of the Mouse Brain

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form a polished and reinforced thinned skull (PoRTS) window in the mouse skull that spans several millimeters in diameter and is stable for months. The skull is thinned to 10 to 15 &mu;m in thickness with a hand held drill to achieve optical clarity, and is then overlaid with cyanoacrylate glue and a cover glass to: 1) provide rigidity, 2) inhibit bone regrowth and 3) reduce light scattering from irregularities on the bone surface. Since the skull is not breached, any inflammation that could affect the process being studied is greatly reduced. Imaging depths of up to 250 &mu;m below the cortical surface can be achieved using two-photon laser scanning microscopy. This window is well suited to study cerebral blood flow and cellular function in both anesthetized and awake preparations. It further offers the opportunity to manipulate cell activity using optogenetics or to disrupt blood flow in targeted vessels by irradiation of circulating photosensitizers.

We present a method to form an imaging window in the mouse skull that spans millimeters and is stable for months without inflammation of the brain. This method is well suited for longitudinal studies of blood flow, cellular dynamics, and cell/vascular structure using two-photon microscopy.

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form ...

Patch Clamp Recordings in Inner Ear Hair Cells Isolated from Zebrafish

Patch clamp analyses of the voltage-gated channels in sensory hair cells isolated from a variety of species have been described previously1-4 but this video represents the first application of those techniques to hair cells from zebrafish. Here we demonstrate a method to isolate healthy, intact hair cells from all of the inner ear end-organs: saccule, lagena, utricle and semicircular canals. Further, we demonstrate the diversity in hair cell size and morphology and give an example of the kinds of patch clamp recordings that can be obtained. The advantage of the use of this zebrafish model system over others stems from the availability of zebrafish mutants that affect both hearing and balance. In combination with the use of transgenic lines and other techniques that utilize genetic analysis and manipulation, the cell isolation and electrophysiological methods introduced here should facilitate greater insight into the roles hair cells play in mediating these sensory modalities.

The purpose of this video is to demonstrate procedures for obtaining healthy, intact hair cells from the inner ear organs of adult zebrafish and then using them for patch clamp studies aimed at characterizing the biophysical properties of their voltage-gated channels.

Patch  clamp analyses of the voltage-gated channels in sensory hair cells isolated  from a variety of species have been described previously1-4 but this  ...

Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an experimental approach with excellent spatial and temporal resolutions. Moreover, such an approach must also have the ability to distinguish receptors localized on the PM from those in intracellular compartments. Most importantly, detecting receptors in a single vesicle requires outstanding detection sensitivity, since each vesicle carries only a small number of receptors. Standard approaches for examining receptor trafficking include surface biotinylation followed by biochemical detection, which lacks both the necessary spatial and temporal resolutions; and fluorescence microscopy examination of immunolabeled surface receptors, which requires chemical fixation of cells and therefore lacks sufficient temporal resolution1-6 . To overcome these limitations, we and others have developed and employed a new strategy that enables visualization of the dynamic insertion of receptors into the PM with excellent spatial and temporal resolutions 7-17 . The approach includes tagging of a pH-sensitive GFP, the superecliptic pHluorin 18, to the N-terminal extracellular domain of the receptors. Superecliptic pHluorin has the unique property of being fluorescent at neutral pH and non-fluorescent at acidic pH (pH &lt; 6.0). Therefore, the tagged receptors are non-fluorescent when within the acidic lumen of intracellular trafficking vesicles or endosomal compartments, and they become readily visualized only when exposed to the extracellular neutral pH environment, on the outer surface of the PM. Our strategy consequently allows us to distinguish PM surface receptors from those within intracellular trafficking vesicles. To attain sufficient spatial and temporal resolutions, as well as the sensitivity required to study dynamic trafficking of receptors, we employed total internal reflection fluorescent microscopy (TIRFM), which enabled us to achieve the optimal spatial resolution of optical imaging (~170 nm), the temporal resolution of video-rate microscopy (30 frames/sec), and the sensitivity to detect fluorescence of a single GFP molecule. By imaging pHluorin-tagged receptors under TIRFM, we were able to directly visualize individual receptor insertion events into the PM in cultured neurons. This imaging approach can potentially be applied to any membrane protein with an extracellular domain that could be labeled with superecliptic pHluorin, and will allow dissection of the key detailed mechanisms governing insertion of different membrane proteins (receptors, ion channels, transporters, etc.) to the PM.

By tagging the extracellular domains of membrane receptors with superecliptic pHluorin, and by imaging these fusion receptors in cultured mouse neurons, we can directly visualize individual vesicular insertion events of the receptors to the plasma membrane. This technique will be instrumental in elucidating the molecular mechanisms governing receptor insertion to the plasma membrane.

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an ...

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

Mouse Microsurgery Infusion Technique for Targeted Substance Delivery into the CNS via the Internal Carotid Artery

Animal models of central nervous system (CNS) diseases and, consequently, blood-brain barrier disruption diseases, require the delivery of exogenous substances into the brain. These exogenous substances may induce injurious impact or constitute therapeutic strategy. The most common delivery methods of exogenous substances into the brain are based on systemic deliveries, such as subcutaneous or intravenous routes. Although commonly used, these approaches have several limitations, including low delivery efficacy into the brain. In contrast, surgical methods that locally deliver substances into the CNS are more specific and prevent the uptake of the exogenous substances by other organs. Several surgical methods for CNS delivery are available; however, they tend to be very traumatic. Here, we describe a mouse infusion microsurgery technique, which effectively delivers substances into the brain via the internal carotid artery, with minimal trauma and no interference with normal CNS functionality.

Mouse Microsurgery Infusion Technique for Targeted Substance Delivery into the CNS via the Internal Carotid Artery

The present protocol describes a mouse microsurgery infusion technique, which effectively delivers substances directly into the brain via the internal carotid artery.

Animal models of central nervous system (CNS) diseases and, consequently, blood-brain barrier disruption diseases, require the delivery of exogenous ...

Laminectomy and Spinal Cord Window Implantation in the Mouse

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated digital vaporizer is utilized to achieve a stable plane of anesthesia at a low-flow rate of isoflurane. A single vertebral spine is removed, and a commercially available cover-glass is overlaid on a thin agarose bed. A 3D-printed plastic backplate is then affixed to the adjacent vertebral spines using tissue adhesive and dental cement. A stabilization platform is used to reduce motion artifact from respiration and heartbeat. This rapid and clamp-free method is well-suited for acute multi-photon fluorescence microscopy. Representative data are included for an application of this technique to two-photon microscopy of the spinal cord vasculature in transgenic mice expressing eGFP:Claudin-5 &#8212; a tight junction protein.

This protocol describes implantation of a glass window onto the spinal cord of a mouse to facilitate visualization by intravital microscopy.

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated ...