This work is supported by the Intramural Research Program of National Institute of Neurological Disorders and Stroke, National Institutes of Health. Project number 1ZIANS003137.

CAUTION: This protocol involves the use of class IV lasers and will require proper training and safety guidelines to be followed. Avoid eye or skin exposure to direct or scattered laser light. 
NOTE: The protocol includes six steps. The workflow is shown in Figure 1A.
1. Labeling Individual Neurons Using the Flip-out Technique
NOTE: The single labeling of LNvs is achieved by expressing mCD8::GFP in single LNvs using flippase -mediated stochastic labeling. The genotype of the fly line is: hs-flp; Pdf-Gal4; UAS-FRT-CD2-stop-FRT-mCD8::GFP11,12,13. The frequency of obtaining a single labeled LNv is around 10%. Fly stocks are maintained in standard medium in circadian- and humidity-controlled 25 &#176;C incubators.
<ol>
	<li>Collect 100-200 eggs within a 2 h window post fertilization on a grape juice plate with yeast supplement. Incubate the embryos at 25 &#176;C for 24 h and collect newly hatched first instar larvae for the next step.</li>
	<li>Heat shock the newly hatched first instar larvae at 37.5 &#176;C for 40 min two times, with a 40 min recovery period in between.</li>
	<li>Culture the larvae at 25 &#176;C with circadian and humidity controls to the desired developmental stage(s).</li>
</ol>
2. Dissecting and Mounting Larval Brain Explants
<ol>
	<li>Dissect larval brains in the physiological external saline solution (120 mM NaCl, 4 mM MgCl2, 3 mM KCl, 10 mM NaHCO3, 10 mM Glucose, 10 mM Sucrose, 5 mM TES, 10 mM HEPES, 2 mM Ca2+, PH 7.2) under a dissection microscope (4.5x magnification power) with two pairs of #5 standard tip dissection forceps (11 cm). Use one pair of forceps to hold the larval body in place and the other to carefully dissect out the brain. Preserve the eye disks, brain lobes and the ventral nerve cord. Remove attached muscles to minimize sample movements during imaging.</li>
	<li>Prepare a glass slide (25 x 75 x 1.0 mm3) and use a syringe to draw a square chamber with vacuum grease.</li>
	<li>Add 20 &#181;L of external saline solution to the square chamber with grease barriers.</li>
	<li>Transfer dissected larval brains into the chamber on the glass slide using forceps. Adjust the position of the brains under the dissection scope to ensure the dorsal side faces up.</li>
	<li>Cover the chamber with a glass cover slip (22 x 22 x 0.15 mm3). The larval brain is now mounted on the slide within a chamber filled with the external saline solution (Figure 1B). 
	NOTE: Pressing gently on the coverslip confines the brains and reduces sample drifting in the subsequent imaging session.</li>
</ol>
3. Time-lapse Live Imaging
NOTE: We perform time-lapse imaging experiments using a confocal microscope equipped with a multiphoton laser. The acquisition parameters need to be adjusted for other imaging setups.
<ol>
	<li>Identify brain explants containing individually labeled neurons using a 40X water immersion objective (NA 1.3) and an epifluorescent light source. For image collection, use a two-photon laser tuned to 920 nm and a non-descanned (NDD) detector.</li>
	<li>Collect images at 512 x 512 pixels per frame and 1 min per Z-stack for 10 min. Adjust the optical and digital zoom to achieve a sufficient x-y-z resolution while ensuring the coverage of the whole dendritic arbor within 1 min (Figure 2, Supplementary Video 1). The settings generate images with a typical x-y-z resolution of 0.11 x 0.11 x 0.25 &#181;m3.</li>
	<li>Collect the time lapse image series within 30 min of the brain dissection. Data with excessive drift or obvious morphological deterioration should be excluded from post- processing and quantification.</li>
</ol>
4. Drift Correction and Deconvolution
NOTE: Depending on the image quality, both steps are optional but strongly recommended.
<ol>
	<li>Open an acquired image file with a drift correction software. Edit microscopic parameters to match those used for the experiment. For the software (see Table of Materials), click Edit | Edit microscopic parameters. Set microscope type as widefield for two-photon images if there is no two-photon option and follow the workflow defined by the software. For example, open the tab Deconvolution | Object Stabilizer, and choose Stabilize time frames.</li>
	<li>Open the drift-corrected image in the deconvolution software and follow its workflow. For the software (see Table of Materials), select the stabilized image and then click Deconvolution | Deconvolution Express. To get a better result, fine-tune the parameters using Deconvolution Wizard.</li>
	<li>Save the deconvolved image in a file type that is supported by the subsequent image annotation software capable of analyzing 4D data and reporting the spatial coordinates of defined spots in the image (see Table of Materials).</li>
</ol>
5. Image Annotation
<ol>
	<li>Open the deconvolved image in the image annotation software. Examine and mark branch tips at all time points in 3D. The image annotation software reports and stores the spatial and temporal coordinates of the marked branch tips. Export the coordinate information as a .csv file for subsequent calculations. 
	NOTE: The following two steps are specific to the annotation software we used (see Table of Materials). The workflow may be different for other software.</li>
	<li>Within the Spots module, click &#34;Skip automatic creation, edit manually&#34; and check the bottom &#34;Auto-connect to selected Spot&#34; checkbox (Figure 3A). Go over the frames in the time series and select a branch for annotation. Hold the Shift key and click the branch terminal tip to add a spot. Click through all time points. 
	NOTE: The image annotation software connects the spots between frames and generates a trajectory automatically. The spatial and temporal information of the branch tips is now associated with marked spots. Repeat these steps until all branch tips are annotated (Figure 3B).</li>
	<li>Within the Spots module, click the Statistics tab, choose Detailed and select Specific Values | Position. The recorded spatial and temporal coordinate information will be on display. Click the Save bottom to export that information as a .csv file (Figure 3C).</li>
</ol>
6. Calculating Dendritic Branch Dynamics
NOTE: Using the coordinate information, displacement of the branch tips in 3D can be easily calculated. In our study, all dendritic branch movements are categorized as Extension or Retraction. The steps below describe how to process the .csv files using custom-written R scripts (Supplementary File) and by adding the directional information through manual editing.
<ol>
	<li>Open the .csv file as a spreadsheet. Select the Track ID column, click the Sort Smallest to Largest option and accept Expand the selection. After sorting, Track ID identifies each unique dendritic branch, and spots from the same branch share the same Track ID. The Time column stores the information of different time frames.</li>
	<li>In the spreadsheet, add a Distance column. Calculate the distance of every two temporally adjacent spots using their coordinates and put the values in the Distance column (Figure 4A).</li>
	<li>Minor movements at the single voxel level. Based on the imaging settings, 0.3 &#181;m are usually artifacts from uncorrected drifting or imperfect image annotation. Filter these movements out by resetting all Distance values smaller than 0.3 &#181;m to 0. Process multiple .csv files manually or use our R script batch column filtering.R (Supplementary File).</li>
	<li>Manually generate a new column named Displacement in the spreadsheet. Copy the values from Distance column to Displacement column. Manually assign the extension and retraction events for each branch tip. If it is an extension, leave the Displacement value unchanged, which is a positive value. If it is a retraction, change the corresponding Displacement value to a negative value (e.g., 0.35 to -0.35) (Figure 4B).</li>
	<li>Generate a new column named Event. In this column, manually sum the Displacement values for individual extension and retraction events (Figure 4B).</li>
	<li>Process the modified spreadsheet and quantify the extension and retraction events based on their displacement values using R script batch column sum.R (Supplementary File). The output parameters include Number of extension events, Number of retraction events, Cumulative length extended, Cumulative length retracted, Net length changed, and Total length traveled.</li>
</ol>

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The authors have nothing to disclose.

Here, we describe a protocol we developed to record and quantify the dynamic behavior of dendritic branches in individually labeled neurons in Drosophila larval brains. Notably, our live imaging protocol contains specific parameters that enable us to capture the whole dendritic arbor of a larval LNv and provide a global view of the dynamic state of its dendrite branches. However, because our quantification methods heavily rely on the annotation of branch terminals, neurons with complex dendritic structures and c...

Dendrites are specialized neuronal compartments that receive and process sensory and synaptic input. The complex and stereotyped structure of dendritic arbors has been under intense investigation since their discovery. A number of model systems, including Xenopus optic tectal neurons, chick retinal ganglion cells, and dendritic arborization (da) neurons in the Drosophila system, have been established to study the development, remodeling and plasticity of neuronal dendrites1,2,3,4. Drosophila ventral lateral neurons (LNvs) are a group of visual projection neurons initially identified for their important functions in circadian regulation of fly behaviors5. Studies also revealed the role of larval LNvs as the direct postsynaptic target of the larval photoreceptors (PRs)6,7. Importantly, culturing developing larvae in different light regimes strongly affects the size of LNvs&#39; dendritic arbors, demonstrating the suitability of LNvs as a new model for studying dendritic plasticity7. Recent work from our group further indicates that both the size of the LNv dendrite and the dynamic behavior of the dendritic branches display experience-dependent plasticity8,9. As part of this work, we developed a new live imaging and quantification protocol to perform analysis on the dendrite dynamics of LNvs from 2nd or 3rd instar larvae.
The transparent nature of the Drosophila larval brain makes it ideal for live imaging. However, the dendritic arbors of LNvs are situated in the densely innervated larval optic neuropil (LON) in the center of the larval brain lobe6. To capture images of fine dendrite branches and filopodia in the intact brain tissue, we utilize two-photon microscopy, which increases the depth of light penetration and reduces the phototoxicity during live imaging experiments10. Using this setup, we successfully performed live imaging experiments on LNvs for over 30 min without observing obvious morphological deterioration of the neuron. In addition, genetic manipulations using the Flip-out technique enabled us to label the individual LNvs with a membrane tagged GFP, which is also critical for monitoring the movements of individual branches11,12,13.
To capture the dynamic behavior of all branches on the LNv dendritic arbor with optimal optic resolution, we performed time-lapse 3D imaging on freshly dissected larval brain explants with a high spatial resolution at 1 min per frame for 10 to 30 min. Developing LNv dendrites are highly dynamic, with a large percentage of the branches displaying observable changes within the 10 min window. This leads to one of the main technical challenges in studying dendrite dynamics, quantifying branch behavior based on the 4D image data sets. Previously established methods have various limitations, including lack of accuracy and excessive time requirement. Therefore, we developed a semi-automatic method that combines image post-processing, manual marking of the branch terminals, and automatic 4D spot tracing using an image annotation software. We calculate the movements of branch terminals at different time points based on the 3D coordinates of the spots. The data are then exported and analyzed to produce quantitative measurements of the branch dynamics. This method accurately assesses the duration and extent of extension and retraction events of existing branches, as well as the formation of new branches, allowing us to monitor dendrite dynamics in a large number of neurons.

<table><tbody><tr><td>Chameleon Vision II multiphoton laser</td><td>Coherent</td><td></td><td></td></tr><tr><td>High vacuum grease</td><td>Dow Corning</td><td>79751-30</td><td></td></tr><tr><td>LSM 780 two-photon laser scanning confocal microscope</td><td>Carl Zeiss</td><td></td><td>upright configuration</td></tr><tr><td>Microscope Cover Glass</td><td>Fisher Scientific</td><td>12-544-E</td><td></td></tr><tr><td>Superfrost Plus Microscope Slides</td><td>Fisher Scientific</td><td>12-550-15</td><td></td></tr><tr><td>&lt;strong&gt;Software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Excel</td><td>Microsoft</td><td></td><td>for processing .csv files</td></tr><tr><td>Huygens Professional</td><td>Scientific Volume Imaging</td><td>for drift correction and deconvolution</td><td></td></tr><tr><td>Imaris</td><td>Oxford Instruments</td><td></td><td>for 3D visualization and image annotation</td></tr><tr><td>&lt;strong&gt;Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Glucose</td><td></td><td></td><td></td></tr><tr><td>HEPES</td><td></td><td></td><td></td></tr><tr><td>KCl</td><td></td><td></td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td></td><td></td><td></td></tr><tr><td>NaCl</td><td></td><td></td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td></td><td></td><td></td></tr><tr><td>PBS</td><td></td><td></td><td></td></tr><tr><td>Sucrose</td><td></td><td></td><td></td></tr><tr><td>TES</td><td></td><td></td><td></td></tr></tbody></table>

time-lapse live imaging and quantification of fast dendritic branch dynamics in developing drosophila neurons

Highly motile dendritic filopodia are widely present in neurons at early developmental stages. These exploratory dynamic branches sample the surrounding environment and initiate contacts with potential synaptic partners. Although the connection between dendritic branch dynamics and synaptogenesis is well established, how developmental and activity-dependent processes regulate dendritic branch dynamics is not well understood. This is partly due to the technical difficulties associated with the live imaging and quantitative analyses of these fine structures using an in vivo system. We established a method to study dendrite dynamics using Drosophila larval ventral lateral neurons (LNvs), which can be individually labeled using genetic approaches and are accessible for live imaging. Taking advantage of this system, we developed protocols to capture branch dynamics of the whole dendritic arbor of a single labeled LNv through time-lapse live imaging. We then performed post-processing to improve image quality through drift correction and deconvolution, followed by analyzing branch dynamics at the single-branch level by annotating spatial positions of all branch terminals. Lastly, we developed R scripts (Supplementary File) and specific parameters to quantify branch dynamics using the coordinate information generated by the terminal tracing. Collectively, this protocol allows us to achieve a detailed quantitative description of branch dynamics of the neuronal dendritic arbor with high temporal and spatial resolution. The methods we developed are generally applicable to sparsely labeled neurons in both in vitro and in vivo conditions.

Using the live imaging protocol described above, we capture high resolution image stacks for the subsequent analyses and quantification. Supplementary Video 1 shows the maximum intensity projected (MIP) image series collected from a representative individually labeled LNv. Figure 2B shows the corresponding montage of eight frames of the image series. In both panels, arrowheads mark retraction events and arrows mark extension ...

Watch this Scientific Journal Video about Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons at JoVE.com

Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

Here, we describe the method we employed to image highly motile dendritic filopodia in a live preparation of the Drosophila larval brain, and the protocol we developed to quantify time-lapse 3D imaging datasets for quantitative assessments of dendrite dynamics in developing neurons.

Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

Highly motile dendritic filopodia are widely present in neurons at early developmental stages. These exploratory dynamic branches sample the surrounding ...

time-lapse-live-imaging-quantification-fast-dendritic-branch-dynamics

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Neuroscience

6.2K Views.  National Institutes of Health. Here, we describe the method we employed to image highly motile dendritic filopodia in a live preparation of the Drosophila larval brain, and the protocol we developed to quantify time-lapse 3D imaging datasets for quantitative assessments of dendrite dynamics in developing neurons.

Video: Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Drosophila larvae are an attractive model system for live imaging due to their translucent cuticle and powerful genetics. This protocol describes how to utilize a single-layer PDMS device, called the &#39;larva chip&#39; for live imaging of cellular processes within neurons of 3rd instar Drosophila larvae.

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of ...

Bioengineering

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

Biology

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 ...

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. The molecular clock within each pacemaker neuron generates circadian rhythms in gene expression, which underlie the rhythmic neuronal functions essential to the operation of the circuit. Investigation of the properties of the individual molecular oscillators in different subclasses of pacemaker neurons and their interaction with neuronal signaling yields a better understanding of the circadian pacemaker circuit. Here, we present a time-lapse fluorescent microscopy approach developed to monitor the molecular clockwork in clock neurons of cultured Drosophila larval brain. This method allows the multi-day recording of the rhythms of genetically encoded fluorescent circadian reporters at single-cell resolution. This setup can be combined with pharmacological manipulations to closely analyze real-time response of the molecular clock to various compounds. Beyond circadian rhythms, this multipurpose method in combination with powerful Drosophila genetic techniques offers the possibility to study diverse neuronal or molecular processes in live brain tissue.

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The goal of this protocol is to establish ex vivo Drosophila larval brain culture optimized to monitor circadian molecular rhythms with long-term fluorescence time-lapse imaging. The application of this method to pharmacological assays is also discussed.

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. ...

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. ...

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. Learning-induced changes in synaptic transmission are often distributed across many neurons and levels of processing in the brain. Therefore, methods to visualize learning-dependent synaptic plasticity across neurons are needed. The fruit fly Drosophila melanogaster represents a particularly favorable model organism to study neuronal circuits underlying learning. The protocol presented here demonstrates a way in which the processes underlying the formation of associative olfactory memories, i.e., synaptic activity and their changes, can be monitored in vivo. Using the broad array of genetic tools available in Drosophila, it is possible to specifically express genetically encoded calcium indicators in determined cell populations and even single cells. By fixing a fly in place, and opening the head capsule, it is possible to visualize calcium dynamics in these cells whilst delivering olfactory stimuli. Additionally, we demonstrate a set-up in which the fly can be subjected, simultaneously, to electric shocks to the body. This provides a system in which flies can undergo classical olfactory conditioning - whereby a previously na&#239;ve odor is learned to be associated with electric shock punishment - at the same time as the representation of this odor (and other untrained odors) is observed in the brain via two-photon microscopy. Our lab has previously reported the generation of synaptically localized calcium sensors, which enables one to confine the fluorescent calcium signals to pre- or postsynaptic compartments. Two-photon microscopy provides a way to spatially resolve fine structures. We exemplify this by focusing on neurons integrating information from the mushroom body, a higher-order center of the insect brain. Overall, this protocol provides a method to examine the synaptic connections between neurons whose activity is modulated as a result of olfactory learning.

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Here we present a protocol with which pre- and/or postsynaptic calcium can be visualized in the context of Drosophila learning and memory. In vivo calcium imaging using synaptically localized calcium sensors is combined with a classical olfactory conditioning paradigm such that the synaptic plasticity underlying this type of associative learning may be determined.

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. ...

3D Particle Tracking for Noninvasive In Vivo Analysis of Synaptic Microtubule Dynamics in Dendrites and Neuromuscular Junctions of Drosophila

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. Furthermore, despite progress in understanding postsynaptic MTs, much less is known about the contributions of presynaptic MTs to neuronal morphogenesis. In particular, studies of in vivo MT dynamics in Drosophila sensory dendrites yielded significant insights into polymer-level behavior. However, the technical and analytical challenges associated with live imaging of the fly neuromuscular junction (NMJ) have limited comparable studies of presynaptic MT dynamics. Moreover, while there are many highly effective software strategies for automated analysis of MT dynamics in vitro and ex vivo, in vivo data often necessitate significant operator input or entirely manual analysis due to inherently inferior signal-to-noise ratio in images and complex cellular morphology. &#160;To address this, this study optimized a new software platform for automated and unbiased in vivo particle detection. Multiparametric analysis of live time-lapse confocal images of EB1-GFP labeled MTs was performed in both dendrites and the NMJ of Drosophila larvae and found striking differences in MT behaviors. MT dynamics were furthermore analyzed following knockdown of the MT-associated protein (MAP) dTACC, a key regulator of Drosophila synapse development, and identified statistically significant changes in MT dynamics compared to wild type. These results demonstrate that this novel strategy for the automated multiparametric analysis of both pre- and postsynaptic MT dynamics at the polymer-level significantly reduces human-in-the-loop criteria. The study furthermore shows the utility of this method in detecting distinct MT behaviors upon dTACC-knockdown, indicating a possible future application for functional screens of factors that regulate MT dynamics in vivo. Future applications of this method may also focus on elucidating cell type and/or compartment-specific MT behaviors, and multicolor correlative imaging of EB1-GFP with other cellular and subcellular markers of interest.&#160;

This study presents a noninvasive intravital neuronal imaging strategy combined with a new software strategy to achieve automated, unbiased tracking and analysis of in vivo microtubule (MT) plus-end dynamics in the sensory dendrites and the neuromuscular junctions of Drosophila.

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. ...

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.

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 ...

Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

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Chapters in this video

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Live Imaging of Drosophila Larval Neuroblasts

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

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

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

Quantitative Analysis of Neuronal Dendritic Arborization Complexity in Drosophila

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

3D Particle Tracking for Noninvasive In Vivo Analysis of Synaptic Microtubule Dynamics in Dendrites and Neuromuscular Junctions of Drosophila

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