Name: time-lapse imaging of neuronal arborization using sparse adeno-associated virus labeling of genetically targeted retinal cell populations
Uploaded: 2021-03-19T14:00:00
Description: Watch this Scientific Journal Video about Time-Lapse Imaging of Neuronal Arborization using Sparse Adeno-Associated Virus Labeling of Genetically Targeted Retinal Cell Populations at JoVE.com

We thank Madison Gray for giving me a hand when I didn&#39;t have any. This research was supported by an NSERC Discovery Grant (RGPIN-2016-06128), a Sloan Fellowship in Neuroscience and a Canada Research Chair Tier 2 (to J.L.L). S. Ing-Esteves was supported by the Vision Science Research Program and NSERC Postgraduate Scholarships-Doctoral.

NOTE: This protocol spans 2 days with a minimum period of 4-5 days for viral transduction between experimental days (Figure 1A). Animal experiments are performed in accordance with the Canadian Council on Animal Care Guidelines for Use of Animals in Research and Laboratory Animal Care under protocols approved by the Laboratory of Animal Services Animal Use and Care Committee at the Hospital for Sick Children (Toronto, Canada).
1. Preparations for the neonatal AAV in...

This video demonstrates an experimental pipeline that utilizes existing genetic tools to image dendrite dynamics of developing retinal neurons with confocal live-imaging. Also demonstrated are intraocular injections of Cre-dependent AAVs encoding membrane-targeted fluorescent proteins into neonatal mice. Single cells of genetically targeted populations are brightly labelled as early as 4-5 dpi. Retinal flat-mounts were prepared for standard imaging chambers to perform live-cell confocal imaging. This method produces high...

Dendrites of retinal neurons form intricate, yet specific, patterns that influence their function within neural circuits. In the vertebrate retina, diverse types of retinal ganglion cells (RGCs) and amacrine cell interneurons bear unique dendritic morphologies that differ in arbor size, location, branch length, and density1. During postnatal development, RGCs and amacrine cells extend exuberant dendritic processes into a neuropil called the inner plexiform layer (IPL), where they receive bipolar cell inputs transmitting photoreceptor signals2. As captured by time-lapse imaging of fluorescently labelled retinal populations in chick or zebrafish larvae, dendrite morphogenesis is highly dynamic3,4,5. Within days, dendritic arbors expand, remodel, and ramify to narrow sublayers of the IPL, where they synapse with select partners. The arbors exhibit different structural dynamics over development, with changes in relative rates of branch addition, retraction, and stabilization. Amacrine and RGC dendrites also exhibit different outgrowth and remodeling behaviors that might reflect type-specific arborization. However, these studies tracked broad amacrine or RGC populations and focused on laminar targeting, which is just one aspect of morphology.
The mechanisms that produce the vast morphological diversity observed across retinal subtypes are poorly understood. The objective of this group was to develop a method to capture dendrite dynamics and arbor remodeling of defined retinal subtypes in mice. Identifying cell type-specific mechanisms of dendrite patterning requires methods to visualize and measure dendrite behaviors of cells of interest. Organotypic cultures of mouse retinas are well suited for live-cell imaging studies using confocal or multiphoton microscopy. Developing retinas are dissected and mounted into a flat explant that can be imaged for several hours in a recording chamber or cultured over a few days with limited effects on the circuitry6,7. Live retinal neurons can be labeled by a variety of techniques, including dye-filling by electrodes, electroporation, biolistic delivery of particles coated with lipophilic dyes or plasmids encoding fluorescent proteins (e.g., Gene Gun), as well as genetically encoded cell labels7,8,9,10. However, these approaches are inefficient for imaging dendrite dynamics of specific retinal subtypes. For instance, dye-filling methods are low-throughput and require electrophysiology apparatus and additional genetic labels to reliably target cells of interest. Moreover, the strong fluorescence signals in the soma can obscure nearby dendrites.
Biolistic gene delivery methods can simultaneously label dozens of cells, but steps involving high-pressure particle delivery and overnight incubation of isolated retina can compromise cell physiology and dendritic outgrowth. This paper proposes that recent genetic tools can be employed to capture early dendrite dynamics with cell type and structural resolution, given the following experimental criteria. First, to resolve the fine branches and filopodia that dominate developing arbors, the method should label neurons with bright, fluorescent proteins that fill processes in the entire arbor. The fluorescence labeling should not fade due to photobleaching during the imaging period. A variety of fluorescent protein variants have been generated and compared for suitability for in vivo/ex vivo&#160;imaging11 based on brightness and photostability. Second, the fluorescent proteins (XFPs) must be expressed at sufficiently high levels by the earliest stage of dendrite morphogenesis, so that the narrow developmental window is not missed. In analyses of static timepoints in the mouse retina, dendrite development occurs during the first postnatal week and includes phases of outgrowth, remodeling, and stabilization10,12,13,14,15. Third, the method should lead to selective labelling or to an increased probability of labelling of the neuronal subpopulation of interest.&#160;Fourth, labelling of the target subpopulation must be sufficiently sparse so that the entire neuronal arbor can be identified and traced.&#160;Although RGC and amacrine subtypes can be distinguished by their mature morphological characteristics and IPL stratification patterns16,17,18,19,20, the challenge is to identify subtypes during development based on immature structures. This task is facilitated by the expansion of transgenic tools to label specific retinal cell types during development.
Transgenic and knock-in mouse lines in which cellular and temporal expression of fluorescent proteins or Cre is determined by gene regulatory elements are widely used to study retinal cell types13,21,22,23. Key observations on subtype-specific patterns of dendrite development have come from studies of transgenic mouse retinas at static timepoints10,14,24,25. The Cre-Lox system, in particular, enables exquisite gene manipulation and monitoring of subtypes using a variety of recombinase-dependent reporters, sensors, and optogenetic activators. These tools have led to discoveries of subtype-specific molecular programs and functional properties that underlie retinal circuit assembly26,27,28,29,30. However, they have yet to be leveraged to study subtype-specific dendrite dynamics in the mouse retina. Low-density labeling can be achieved by combining Cre mouse lines with transgenes introduced by electroporation or by recombinant AAVs. If available, tamoxifen-inducible Cre lines or intersectional genetic strategies can also be used. Finally, the cell should be labelled in a minimally invasive manner and imaged using acquisition parameters so as not to compromise the tissue or interfere with cellular function required for dendrite morphogenesis.
Presented here is a method to apply transgenic tools and confocal microscopy to investigate dendrite dynamics in live mouse retinal explants. Cre transgenic mouse lines have been combined with AAV vectors that express fluorescent proteins upon Cre recombination, which allows for sparse labeling of retinal cells of interest. Commercially available AAVs are delivered to neonatal retina by intravitreal injections. This paper demonstrates that AAVs produce significantly high and cell type-specific fluorescent expression by 4 dpi, allowing access to postnatal time points. To illustrate this approach, the cholinergic &#34;starburst&#34; amacrine interneuron was labelled by delivering Brainbow AAV in neonatal mice expressing the choline acetyltransferase (ChAT)-internal ribosome entry site (IRES)-Cre transgene, which is active in the early postnatal retina31,32. Starburst amacrine cells develop a stereotyped and radial arbor morphology that is shaped by dendrite self-avoidance mediated by the clustered protocadherins33,34. This paper shows that the resolution of starburst dendrites and filopodia is significantly improved by XFPs to the plasma membrane with the addition of the CAAX motif which undergoes farnesylation, as used for the Brainbow AAVs31. Finally, time-lapse imaging and post-processing protocols have been determined&#160;that produce high-quality images amenable for dendrite reconstruction and morphometric quantification. This protocol can be used to identify factors controlling dendrite morphogenesis and to capture several cellular behaviors in the intact retina.

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			<td>Addgene viral prep #45185-AAV9</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Addgene viral prep #45186-AAV9</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
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			<td>Dissection tools</td>
			<td></td>
			<td></td>
			<td></td>
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		<tr>
			<td>Cellulose filter paper</td>
			<td>Whatman</td>
			<td>1001-070</td>
			<td></td>
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		<tr>
			<td>Dumont #5 fine forceps</td>
			<td>FST</td>
			<td>11252-20</td>
			<td>Two Dumont #5 forceps are required for retinal micro-dissection</td>
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			<td>Dumont forceps</td>
			<td>VWR</td>
			<td>82027-426</td>
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			<td>Fine Scissors</td>
			<td>FST</td>
			<td>14058-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Mixed cellulose ester membrane (MCE) filter papers, hydrophilic, 0.45 &#181;m pore size</td>
			<td>Millipore</td>
			<td>HABG01 300</td>
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			<td>Petri Dish, 50 &#215; 15 mm</td>
			<td>VWR</td>
			<td>470313-352</td>
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			<td>Polyethylene disposable transfer pipette</td>
			<td>VWR</td>
			<td>470225-034</td>
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			<td>Round tip paint brush, size 3/0</td>
			<td>Conventional art supply store</td>
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			<td>Two size 3/0 paint brushes (or smaller) are required for retinal flat-mounting</td>
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			<td>Surgical Scissors</td>
			<td>FST</td>
			<td>14007-14</td>
			<td></td>
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			<td>Vannas Spring Scissors - 2.5 mm Cutting Edge</td>
			<td>FST</td>
			<td>15000-08</td>
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			<td>Live-imaging incubation system</td>
			<td></td>
			<td></td>
			<td></td>
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			<td>Chamber polyethylene tubing, PE-160 10&#39;</td>
			<td>Warner Instruments</td>
			<td>64-0755</td>
			<td></td>
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			<td>Dual channel heater controller, Model TC-344C</td>
			<td>Warner Instruments</td>
			<td>64-2401</td>
			<td></td>
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			<td>HC FLUOTAR L 25x/0.95 W VISIR dipping objective</td>
			<td>Leica</td>
			<td>15506374</td>
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			<td>Heater controller cable</td>
			<td>Warner Instruments</td>
			<td>CC-28</td>
			<td></td>
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			<td>Large bath incubation chamber with slice support</td>
			<td>Warner Instruments</td>
			<td>RC-27L</td>
			<td></td>
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			<td>MPII Mini-Peristaltic Pump</td>
			<td>Harvard Apparatus</td>
			<td>70-2027</td>
			<td></td>
		</tr>
		<tr>
			<td>PM-6D Magnetic Heated Platform (incubation chamber heater)</td>
			<td>Warner Instruments</td>
			<td>PM-6D</td>
			<td></td>
		</tr>
		<tr>
			<td>Pump Head Tubing Pieces For MPII Mini-Peristaltic Pump</td>
			<td>Harvard Apparatus</td>
			<td>55-4148</td>
			<td></td>
		</tr>
		<tr>
			<td>Sample anchor (Harps)</td>
			<td>Warner Instruments</td>
			<td>64-0260</td>
			<td>Sample anchor must be compatible with incubation chamber</td>
		</tr>
		<tr>
			<td>Sloflo In-line Solution Heater</td>
			<td>Warner Instruments</td>
			<td>SF-28</td>
			<td></td>
		</tr>
		<tr>
			<td>Neonatal Injections</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10 &#181;L Microliter Syringe Series 700, Removable Needle</td>
			<td>Hamilton Company</td>
			<td>80314</td>
			<td></td>
		</tr>
		<tr>
			<td>30 G Hypodermic Needles (0.5 inch)</td>
			<td>BD PrecisionGlide</td>
			<td>305106</td>
			<td></td>
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		<tr>
			<td>4 inch thinwall glass capillary, no filament (1.0 mm outer diameter/0.75 mm)&#160;</td>
			<td>WPI World Precision Instruments</td>
			<td>TW100-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol 99.8% (to dilute to 70% with double-distilled water [ddH2O])</td>
			<td>Sigma-Aldrich</td>
			<td>V001229&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>AAV9.hEF1a.lox.TagBFP. lox.eYFP.lox.WPRE.hGH-InvBYF</td>
			<td>Penn Vector Core</td>
			<td>AV-9-PV2453</td>
			<td>Addgene Plasmid #45185&#160;</td>
		</tr>
		<tr>
			<td>AAV9.hEF1a.lox.mCherry.lox.mTFP 
			1.lox.WPRE.hGH-InvCheTF</td>
			<td>Penn Vector Core</td>
			<td>AV-9-PV2454</td>
			<td>Addgene Plasmid #45186</td>
		</tr>
		<tr>
			<td>ChAT-IRES-Cre knock-in transgenic mouse line</td>
			<td>The Jackson Laboratory</td>
			<td>6410</td>
			<td></td>
		</tr>
		<tr>
			<td>Fast Green FCF Dye content &#8805;85 %</td>
			<td>Sigma-Aldrich</td>
			<td>F7252-25G</td>
			<td></td>
		</tr>
		<tr>
			<td>Flaming/Brown Micropipette Puller, model P-97</td>
			<td>Sutter Instrument Co.</td>
			<td>P-97</td>
			<td></td>
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		<tr>
			<td>Green tattoo paste</td>
			<td>Ketchum MFG Co</td>
			<td>329A</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate-Buffered Saline, pH 7.4, liquid, sterile-filtered, suitable for cell culture</td>
			<td>Sigma-Aldrich</td>
			<td>806552</td>
			<td></td>
		</tr>
		<tr>
			<td>Pneumatic PicoPump</td>
			<td>WPI World Precision Instruments</td>
			<td>PV-820</td>
			<td></td>
		</tr>
		<tr>
			<td>Oxygenated artifiial cerebrospinal fluid (aCSF) Reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate (CaCl2&#183;2H2O)</td>
			<td>Sigma-Aldrich</td>
			<td>C7902</td>
			<td></td>
		</tr>
		<tr>
			<td>Carbogen (5% CO2, 95% O2)</td>
			<td>AirGas</td>
			<td>X02OX95C2003102</td>
			<td>Supplier may vary depending on region</td>
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		<tr>
			<td>D-(+)-Glucose</td>
			<td>Sigma-Aldrich</td>
			<td>G7021</td>
			<td></td>
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			<td>HEPES, Free Acid</td>
			<td>Bio Basic</td>
			<td>HB0264</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrochloric acid solution, 1 N</td>
			<td>Sigma-Aldrich</td>
			<td>H9892</td>
			<td></td>
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		<tr>
			<td>Magnesium chloride hexahydrate (MgCl2&#183;6H2O)</td>
			<td>Sigma-Aldrich</td>
			<td>M2670</td>
			<td></td>
		</tr>
		<tr>
			<td>pH-Test strips (6.0-7.7)</td>
			<td>VWR</td>
			<td>BDH35317.604</td>
			<td></td>
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		<tr>
			<td>Potassium chloride (KCl)</td>
			<td>Sigma-Aldrich</td>
			<td>P9541</td>
			<td></td>
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		<tr>
			<td>Sodium chloride (NaCl)</td>
			<td>Bio Basic</td>
			<td>DB0483</td>
			<td></td>
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			<td>Sodium phosphate monobasic (NaH2PO4)</td>
			<td>Sigma-Aldrich</td>
			<td>RDD007</td>
			<td></td>
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			<td>Software</td>
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			<td>ImageJ</td>
			<td>National Institutes of Health (NIH)</td>
			<td>Open source</td>
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time-lapse imaging of neuronal arborization using sparse adeno-associated virus labeling of genetically targeted retinal cell populations

Discovering mechanisms that pattern dendritic arbors requires methods to visualize, image, and analyze dendrites during development. The mouse retina is a powerful model system for the investigation of cell type-specific mechanisms of neuronal morphogenesis and connectivity. The organization and composition of retinal subtypes are well-defined, and genetic tools are available to access specific types during development. Many retinal cell types also constrain their dendrites and/or axons to narrow layers, which facilitates time-lapse imaging. Mouse retina explant cultures are well suited for live-cell imaging using confocal or multiphoton microscopy, but methods optimized for imaging dendrite dynamics with temporal and structural resolution are lacking. Presented here is a method to sparsely label and image the development of specific retinal populations marked by the Cre-Lox system. Commercially available adeno-associated viruses (AAVs) used here expressed membrane-targeted fluorescent proteins in a Cre-dependent manner. Intraocular delivery of AAVs in neonatal mice produces fluorescent labeling of targeted cell types by 4-5 days post-injection (dpi). The membrane fluorescent signals are detectable by confocal imaging and resolve fine branch structures and dynamics. High-quality videos spanning 2-4 h are acquired from imaging retinal flat-mounts perfused with oxygenated artificial cerebrospinal fluid (aCSF). Also provided is an image postprocessing pipeline for deconvolution and three-dimensional (3D) drift correction. This protocol can be used to capture several cellular behaviors in the intact retina and to identify novel factors controlling neurite morphogenesis. Many developmental strategies learned in the retina will be relevant for understanding the formation of neural circuits elsewhere in the central nervous system.

Using the above protocol, a high-resolution 3D video of developing starburst cell dendrites was acquired, deconvolved, and corrected for 3D drift. Z-plane maximum projections were produced to make 2D videos for analysis (Supplementary Video 1, Figure 5A). 3D deconvolution of each time point increased the resolution of fine filopodia projections (Figure 5B,C). Fine filopodia protrusions are a feature of developing retinal dendrit...

Watch this Scientific Journal Video about Time-Lapse Imaging of Neuronal Arborization using Sparse Adeno-Associated Virus Labeling of Genetically Targeted Retinal Cell Populations at JoVE.com

Time-Lapse Imaging of Neuronal Arborization using Sparse Adeno-Associated Virus Labeling of Genetically Targeted Retinal Cell Populations

Here, we present a method for investigating neurite morphogenesis in postnatal mouse retinal explants by time-lapse confocal microscopy. We describe an approach for sparse labeling and acquisition of retinal cell types and their fine processes using recombinant adeno-associated virus vectors that express membrane-targeted fluorescent proteins in a Cre-dependent manner.

Discovering mechanisms that pattern dendritic arbors requires methods to visualize, image, and analyze dendrites during development. The mouse retina is a ...

Single-cell labeling and visualization of dendritic or axonal arbors is required to study neuronal morphogenesis. By labeling developing arbors in a minimally-invasive manner, retinal tissue remains healthy and suitable for prolonged confocal live imaging. The main advantage of this method is the ability to visualize individual arbors with multi-color fluorescent proteins within four days post-injection.This makes studying neonatal arbor morphologies possible. This injection imaging and analysis protocol can be used to study neuronal morphologies across the central nervous system. Appropriate Cre selection and injection location must be determined to target the cell population of interest.Begin by selecting a Cre mouse line to label the retinal cell populations of interest. Obtain recombinase-dependent AAV virus encoding fluorescent proteins. For optimal labeling of fine processes, select vectors that express modified fluorescent proteins targeting the plasma membrane.For the AAV aliquot on ice, and prepare a one to four AAV dilution using sterile saline or PBS. To visualize the injections, color the solution blue by adding approximately one microliter of 0.02%Fast Green FCF dye solution for every 15 microliters of AAV dilution. Sterilize the injection area with 70%ethanol.Backfill the micropipette with the AAV dilution using a microsyringe. Under a stereo microscope, break the micropipette tip with a 30-gauge needle to unseal the tip. After anesthetizing the neonatal mice, tattoo their paw pads with tattoo ink using a 30-gauge needle to identify the animals.Collect tail clippings for DNA isolation and genotyping. Swab the skin overlying the eyes with 70%ethanol. Use a 30-gauge needle to open the fused eyelid while applying light pressure with the fingers to open the eye.Poke a small hole through the cornea at the cornea sclera junction. Insert the glass micropipette into the hole and press the micro injector foot pedal two to four times to inject the AAV into the intravitreal space. Slowly remove the micropipette and confirm AAV injection by visualizing the blue dye in the pupil.After preparing the retinal aCSF as described in the text manuscript, oxygenate the retinal aCSF by bubbling with carbogen for a minimum of 15 minutes, then adjust the pH to 7.4. Embed a 60 millimeter diameter Petri dish into an ice tray and fill it with oxygenated retinal aCSF. Place the filled Petri dish under a stereo microscope.Next, cut the eyelid flap of the euthanized mouse to expose the eye. Use blunt forceps to enucleate the eyes and transfer them into the cold retinal aCSF placed under the stereo microscope. Under the stereo microscope, stabilize the eye by clasping the optic nerve using Dumont number five forceps and poke a hole in the center of the cornea with a 30 gauge needle, then insert one tip of the micro scissors into the hole to make an incision from the hole to the end of the cornea.Repeat to make four slices in the cardinal directions, creating four flaps. Grasp and pull the two adjacent flaps apart, gently peeling the sclera from the retina for all the cornea flaps. Then remove the lens from the retinal cup using the forceps.Finally, use micro scissors to make four radial incisions from the edge of the retina towards the optic nerve, creating four equal petals. If using large diameter gray MCE membrane filters for mounting, cut the disc into quadrants roughly one centimeter across, then place the MCE disc onto the center of a larger white filter paper. Using two size three by zero paint brushes, flip one retinal cup onto a paint brush with the retinal ganglion cell side down.Gently lift the retina out of the aCSF, making sure the water tension does not tear the retina. While still holding the paint brush with the retina, move the Petri dish containing aCSF out of the way and place the white filter paper with the MCE membrane under the stereoscope. Using a transfer pipette, place a droplet of aCSF in the center of the MCE filter paper.Float the retina into the aCSF droplet created by the surface tension and charged MCE membrane. Use paint brushes to position the retina within the droplet with the retinal ganglion cell side up, then unfold the four petals. Once positioned, create a water bridge between the paintbrush and white filter paper to break the surface tension of the droplet.Assemble the live imaging incubation chamber and fill it with oxygenated aCSF. Turn on the pump and temperature controller, making sure that the temperature does not rise above 34 degrees Celsius. It can take up to an hour for the chamber temperature to stabilize.It is recommended to set up the incubation chamber before beginning retinal dissection. Stable chamber temperature helps reduce sample drift. To transfer the retinal flat mound into the perfusion chamber, stop the pump and remove the aCSF that is in the chamber.Then place the MCE disc with the retinal flat mount into the incubation chamber. Pre-wet a sample weight to break the surface tension, then place it onto the flat mount ensuring that the sample weight is placed on the MCE paper to reduce sample drift. Refill the chamber with the warmed aCSF and circulate aCSF at approximately one milliliter per minute.Position the nosepiece with the 25 times water dipping objective into the imaging chamber. Screen for labeled cells of interest using epifluorescent light, and adjust the imaging volume to capture dendritic features of interest. Using a lookup table that identifies both oversaturated and undersaturated pixels, adjust the laser power such that no pixels are oversaturated.Acquire the 3D imaging volume and repeat acquisition at the desired frame rate. Imaging volume can be adjusted between each time point to compensate for sample drift. Import the image series, splitting the images by time.Save all time points manually or by running a macro plugin. Create a theoretical point spread function using the ImageJ plugin by clicking Defraction PSF 3D, lens numerical aperture, fluorophore emission wavelength, pixel size, and Z-spacing are required to create an accurate PSF. Select plugins, macros, and run to perform batch parallel iterative deconvolution for all time points using the provided macro.Import all deconvolved time points as a stack by clicking file, import, then image sequence. Convert the stack into a hyperstack by clicking image, hyperstacks, then stacks to hyperstack. Add the number of Z-slices and timeframes, then correct the 3D drift using the correct 3D drift plugin.Save the 3D drift corrected hyperstack and convert the image back to a regular stack by clicking image, hyperstacks, hyperstack to stack, then split the time points by clicking image, stacks, tools, and stacks splitter. For the number of substacks to split, enter the number of time points. Use batch processing to create a maximum projection for all time points.Import the time lapse image sequence by clicking file, import, then image sequence. And use conventional ImageJ tools for the desired analysis of deconvolved and post processed two dimensional video, A high resolution 3D video of developing starburst cell dendrites was acquired, deconvolved, and corrected for 3D drift. Z-plane maximum projections were produced to make 2D videos for analysis, showing an area of dendritic refinement and an area of dendritic outgrowth.3D deconvolution of each time point increased the resolution of fine filopodia projections of single P6 starburst amacrine cell before and after deconvolution with ImageJ. Prolonged AAV infection periods do not necessarily lead to increased fluorescent signal as shown by similar levels of protein expression at five and 11 days post-injection. Reducing sample drift during imaging is important.Drift is minimized by maintaining a consistent imaging temperature and by appropriately positioning the weight on the sample. If drift occurs, manual acquisition of the time series allows for adjustment of the imaging volume between frames. This ensures the features of interest remain in frame.This protocol produces high resolution, deconvolved, and drift-corrected 3D videos of developing neurites. Such videos lead to a better understanding of neuron wiring and are required to advance computational analysis methods of 3D morphogenesis. Better automatic tracing and tracking softwares are required to achieve high throughput analysis of neurite development.

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Research

JoVE Journal

Neuroscience

Intracranial Injection of Adeno-associated Viral Vectors

Intracranial injection of viral vectors engineered to express a fluorescent protein is a versatile labeling technique for visualization of specific subsets of cells in different brain regions both in vivo and in brain sections. Unlike the injection of fluorescent dyes, viral labeling offers targeting of individual cell types and is less expensive and time consuming than establishing transgenic mouse lines. In this technique, an adeno-associated viral (AAV) vector is injected intracranially using stereotaxic coordinates, a micropipette and an automated pump for precise delivery of AAV to the desired area with minimal damage to the surrounding tissue. Injection parameters can be tailored to individual experiments by adjusting the animal age at injection, injection location, volume of injection, rate of injection, AAV serotype and the promoter driving gene expression. Depending on the conditions chosen, virally-induced transgene expression can allow visualization of groups of cells, individual cells or fine cellular processes, down to the level of dendritic spines. The experiment shown here depicts an injection of double-stranded AAV expressing green fluorescent protein for the labeling of neurons and glia in the mouse primary visual cortex.

Here we present the intracranial injection of AAV vectors for fluorescent labeling of neurons and glia in the visual cortex.

Intracranial injection of viral vectors engineered to express a fluorescent protein is a versatile labeling technique for visualization of specific ...

An Organotypic Slice Assay for High-Resolution Time-Lapse Imaging of Neuronal Migration in the Postnatal Brain

Neurogenesis in the postnatal brain depends on maintenance of three biological events: proliferation of progenitor cells, migration of neuroblasts, as well as differentiation and integration of new neurons into existing neural circuits. For postnatal neurogenesis in the olfactory bulbs, these events are segregated within three anatomically distinct domains: proliferation largely occurs in the subependymal zone (SEZ) of the lateral ventricles, migrating neuroblasts traverse through the rostral migratory stream (RMS), and new neurons differentiate and integrate within the olfactory bulbs (OB). The three domains serve as ideal platforms to study the cellular, molecular, and physiological mechanisms that regulate each of the biological events distinctly. This paper describes an organotypic slice assay optimized for postnatal brain tissue, in which the extracellular conditions closely mimic the in vivo environment for migrating neuroblasts. We show that our assay provides for uniform, oriented, and speedy movement of neuroblasts within the RMS. This assay will be highly suitable for the study of cell autonomous and non-autonomous regulation of neuronal migration by utilizing cross-transplantation approaches from mice on different genetic backgrounds.

This protocol describes an organotypic slice assay optimized for the postnatal brain and high-resolution time-lapse imaging of neuroblast migration in the rostral migratory stream.

Neurogenesis in the postnatal brain depends on maintenance of three biological events: proliferation of progenitor cells, migration of neuroblasts, as ...

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

Motor Nerve Transection and Time-lapse Imaging of Glial Cell Behaviors in Live Zebrafish

The nervous system is often described as a hard-wired component of the body even though it is a considerably fluid organ system that reacts to external stimuli in a consistent, stereotyped manner, while maintaining incredible flexibility and plasticity. Unlike the central nervous system (CNS), the peripheral nervous system (PNS) is capable of significant repair, but we have only just begun to understand the cellular and molecular mechanisms that govern this phenomenon. Using zebrafish as a model system, we have the unprecedented opportunity to couple regenerative studies with in vivo imaging and genetic manipulation. Peripheral nerves are composed of axons surrounded by layers of glia and connective tissue. Axons are ensheathed by myelinating or non-myelinating Schwann cells, which are in turn wrapped into a fascicle by a cellular sheath called the perineurium. Following an injury, adult peripheral nerves have the remarkable capacity to remove damaged axonal debris and re-innervate targets. To investigate the roles of all peripheral glia in PNS regeneration, we describe here an axon transection assay that uses a commercially available nitrogen-pumped dye laser to axotomize motor nerves in live transgenic zebrafish. We further describe the methods to couple these experiments to time-lapse imaging of injured and control nerves. This experimental paradigm can be used to not only assess the role that glia play in nerve regeneration, but can also be the platform for elucidating the molecular mechanisms that govern nervous system repair.

Although the peripheral nervous system (PNS) is capable of significant repair after injury, little is known about the cellular and molecular mechanisms that govern this phenomenon. Using live, transgenic zebrafish and a reproducible nerve transection assay, we can study dynamic glial cell behaviors during nerve degeneration and regeneration.

The  nervous system is often described as a hard-wired component of the body even  though it is a considerably fluid organ system that reacts to external ...

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 ...

Subpial Adeno-associated Virus 9 (AAV9) Vector Delivery in Adult Mice

The successful development of a subpial adeno-associated virus 9 (AAV9) vector delivery technique in adult rats and pigs has been reported on previously. Using subpially-placed polyethylene catheters (PE-10 or PE-5) for AAV9 delivery, potent transgene expression through the spinal parenchyma (white and gray matter) in subpially-injected spinal segments has been demonstrated. Because of the wide range of transgenic mouse models of neurodegenerative diseases, there is a strong desire for the development of a potent central nervous system (CNS)-targeted vector delivery technique in adult mice. Accordingly, the present study describes the development of a spinal subpial vector delivery device and technique to permit safe and effective spinal AAV9 delivery in adult C57BL/6J mice. In spinally immobilized and anesthetized mice, the pia mater (cervical 1 and lumbar 1-2 spinal segmental level) was incised with a sharp 34 G needle using an XYZ manipulator. A second XYZ manipulator was then used to advance a blunt 36G needle into the lumbar and/or cervical subpial space. The AAV9 vector (3-5 &#181;L; 1.2 x 1013 genome copies (gc)) encoding green fluorescent protein (GFP) was then injected subpially. After injections, neurological function (motor and sensory) was assessed periodically, and animals were perfusion-fixed 14 days after AAV9 delivery with 4% paraformaldehyde. Analysis of horizontal or transverse spinal cord sections showed transgene expression throughout the entire spinal cord, in both gray and white matter. In addition, intense retrogradely-mediated GFP expression was seen in the descending motor axons and neurons in the motor cortex, nucleus ruber, and formatio reticularis. No neurological dysfunction was noted in any animals. These data show that the subpial vector delivery technique can successfully be used in adult mice, without causing procedure-related spinal cord injury, and is associated with highly potent transgene expression throughout the spinal neuraxis.

Subpial Adeno-associated Virus 9 AAV9 Vector Delivery in Adult Mice

The goal of the present study was to develop and validate the potency and safety of spinal adeno-associated virus 9 (AAV9)-mediated gene delivery by using a novel subpial gene delivery technique in adult mice.

The successful development of a subpial adeno-associated virus 9 (AAV9) vector delivery technique in adult rats and pigs has been reported on previously. ...

Visualization and Live Imaging of Oligodendrocyte Organelles in Organotypic Brain Slices Using Adeno-associated Virus and Confocal Microscopy

Neurons rely on the electric insulation and trophic support of myelinating oligodendrocytes. Despite the importance of oligodendrocytes, the advanced tools currently used to study neurons, have only partly been taken on by oligodendrocyte researchers. Cell type-specific staining by viral transduction is a useful approach to study live organelle dynamics. This paper describes a protocol for visualizing oligodendrocyte mitochondria in organotypic brain slices by transduction with adeno-associated virus (AAV) carrying genes for mitochondrial targeted fluorescent proteins under the transcriptional control of the myelin basic protein promoter. It includes the protocol for making organotypic coronal mouse brain slices. A procedure for time-lapse imaging of mitochondria then follows. These methods can be transferred to other organelles and may be particularly useful for studying organelles in the myelin sheath. Finally, we describe a readily available technique for visualization of unstained myelin in living slices by Confocal Reflectance microscopy (CoRe). CoRe requires no extra equipment and can be useful to identify the myelin sheath during live imaging.

Myelinating oligodendrocytes promote rapid action potential propagation and neuronal survival. Described here is a protocol for oligodendrocyte-specific expression of fluorescent proteins in organotypic brain slices with subsequent time-lapse imaging. Further, a simple procedure for visualizing unstained myelin is presented.

Neurons rely on the electric insulation and trophic support of myelinating oligodendrocytes. Despite the importance of oligodendrocytes, the advanced ...

Quantitative Analysis of Neuronal Dendritic Arborization Complexity in Drosophila

Dendrites are the branched projections of a neuron, and dendritic morphology reflects synaptic organization during the development of the nervous system. Drosophila larval neuronal dendritic arborization (da) is an ideal model for studying morphogenesis of neural dendrites and gene function in the development of nervous system. There are four classes of da neurons. Class IV is the most complex with a branching pattern that covers almost the entire area of the larval body wall. We have previously characterized the effect of silencing the Drosophila ortholog of SOX5 on class IV neuronal dendritic arborization complexity (NDAC) using four parameters: the length of dendrites, the surface area of dendrite coverage, the total number of branches, and the branching structure. This protocol presents the workflow of NDAC quantitative analysis, consisting of larval dissection, confocal microscopy, and image analysis procedures using ImageJ software. Further insight into da neuronal development and its underlying mechanisms will improve the understanding of neuronal function and provide clues about the fundamental causes of neurological and neurodevelopmental disorders.

Quantitative Analysis of Neuronal Dendritic Arborization Complexity in Drosophila

This protocol focuses on quantitative analysis of neuronal dendritic arborization complexity (NDAC) in Drosophila, which can be used for studies of dendritic morphogenesis.

Dendrites are the branched projections of a neuron, and dendritic morphology reflects synaptic organization during the development of the nervous system. ...

Adeno-associated Virus-mediated Transgene Expression in Genetically Defined Neurons of the Spinal Cord

Selective manipulation of spinal neuronal subpopulations has mainly been achieved by two different methods: 1) Intersectional genetics, whereby double or triple transgenic mice are generated in order to achieve selective expression of a reporter or effector gene (e.g., from the Rosa26 locus) in the desired spinal population. 2) Intraspinal injection of Cre-dependent recombinant adeno-associated virus (rAAV); here Cre-dependent AAV vectors coding for the reporter or effector gene of choice are injected into the spinal cord of mice expressing Cre recombinase in the desired neuronal subpopulation. This protocol describes how to generate Cre-dependent rAAV vectors and how to transduce neurons in the dorsal horn of the lumbar spinal cord segments L3-L5 with rAAVs. As the lumbar spinal segments L3-L5 are innervated by those peripheral sensory neurons that transmit sensory information from the hindlimbs, spontaneous behavior and responses to sensory tests applied to the hindlimb ipsilateral to the injection side can be analyzed in order to interrogate the function of the manipulated neurons in sensory processing. We provide examples of how this technique can be used to analyze genetically defined subsets of spinal neurons. The main advantages of virus-mediated transgene expression in Cre transgenic mice compared to classical reporter mouse-induced transgene expression are the following: 1) Different Cre-dependent rAAVs encoding various reporter or effector proteins can be injected into a single Cre transgenic line, thus overcoming the need to create several multiple transgenic mouse lines. 2) Intraspinal injection limits manipulation of Cre-expressing cells to the injection site and to the time after injection. The main disadvantages are: 1) Reporter gene expression from rAAVs is more variable. 2) Surgery is required to transduce the spinal neurons of interest. Which of the two methods is more appropriate depends on the neuron population and research question to be addressed.

Intraspinal injection of recombinase dependent recombinant adeno-associated virus (rAAV) can be used to manipulate any genetically labelled cell type in the spinal cord. Here we describe how to transduce neurons in the dorsal horn of the lumbar spinal cord. This technique enables functional interrogation of the manipulated neuron subtype.

Selective manipulation of spinal neuronal subpopulations has mainly been achieved by two different methods: 1) Intersectional genetics, whereby double or ...

Production, Purification, and Quality Control for Adeno-associated Virus-based Vectors

Gene delivery tools based on adeno-associated viruses (AAVs) are a popular choice for the delivery of transgenes to the central nervous system (CNS), including gene therapy applications. AAV vectors are non-replicating, able to infect both dividing and non-dividing cells&#160;and provide long-term transgene expression. Importantly, some serotypes, such as the newly described PHP.B, can cross the blood-brain-barrier (BBB) in animal models, following systemic delivery. AAV vectors can be efficiently produced in the laboratory. However, robust and reproducible protocols are required to obtain AAV vectors with sufficient purity levels and titer values high enough for in vivo applications. This protocol describes an efficient and reproducible strategy for AAV vector production, based on an iodixanol gradient purification strategy. The iodixanol purification method is suitable for obtaining batches of high-titer AAV vectors of high purity, when compared to other purification methods. Furthermore, the protocol is generally faster than other methods currently described. In addition, a quantitative polymerase chain reaction (qPCR)-based strategy is described for a fast and accurate determination of the vector titer, as well as a silver staining method to determine the purity of the vector batch. Finally, representative results of gene delivery to the CNS, following systemic administration of AAV-PHP.B, are presented. Such results should be possible in all labs using the protocols described in this article.

Here, we describe an efficient and reproducible strategy to produce, titer, and quality-control batches of adeno-associated virus vectors. It allows the user to obtain a vector preparation with high-titer&#160;(&#8805;1 x 1013 vector genomes/mL) and a high purity, ready for in vitro or in vivo use.

Gene delivery tools based on adeno-associated viruses (AAVs) are a popular choice for the delivery of transgenes to the central nervous system (CNS), ...