Name: visualization of the axonal projection pattern of embryonic motor neurons in drosophila
Uploaded: 2017-06-16T14:00:00
Description: Watch this Scientific Journal Video about Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila at JoVE.com

I thank Alex L. Kolodkin, as I learned this filleted preparation protocol in his laboratory. I also thank Young Gi Hong for technical assistance. &#160;This study was supported by NRF-2013R1A1A4A01011329 (S.J.).

1. Preparation
<ol>
	<li>Prepare 500 mL of phosphate-buffered saline (PBS) with t-Octylphenoxypolyethoxyethanol (PBT) solution by adding 0.5 g of bovine serum albumin (BSA) and 0.5 mL of t-Octylphenoxypolyethoxyethanol (see the Table of Materials) to 500 mL of 1X PBS and stirring for at least 30 min. Store at 4 &#176;C. Use when relatively fresh and store the solution in a clean bottle.</li>
	<li>Make 10 mL of 4% paraformaldehyde by adding 2.5 mL of 16% stock paraformaldehyde solution and 1 mL of 10x PBS to 6.5 mL of d...

<ol>
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The details of motor axon guidance defects are scored faster and with better accuracy by the filleted preparation of DAB-stained embryos than by laser scanning confocal microscopy of fluorescently labeled ones. Therefore, the filleted preparation of fixed and 1D4-stained embryos is best suited for the functional characterization of guidance cue molecules. Four major classes of guidance cues, including netrins, Slits, semaphorins (Semas), and ephrins, and their cognate receptors have evolutionarily conserved roles across ...

Precise and selective connections between motor axons and target muscles during embryonic development are essential for normal locomotion in Drosophila larvae. The embryonic patterning of 30 muscle fibers in each of the abdominal hemisegments A2-A7 is established by stage 161. The 36 motor neurons that are generated in the ventral nerve cord extend their axons into the periphery to innervate specific target muscles2. Motor axon pathfinding and target recognition can be visualized by immunohistochemistry with an antibody (mouse monoclonal antibody 1D4)3,4. Multiple images of the motor axon projection patterns in wildtype embryos are available on the web5. The 1D4 antibody labels all motor axons and three longitudinal axon fascicles on each side of the midline of the embryonic central nervous system (CNS)4,6 (Figure 1C and Figure 2A). Therefore, immunohistochemistry with FasII antibody provides a powerful tool for identifying genes required for neuromuscular connectivity for demonstrating the molecular mechanisms underlying motor axon guidance and target recognition.
In each of the abdominal hemisegments A2-A7, motor axons project and selectively fasciculate into two principal nerve branches, the segmental nerve (SN) and the intersegmental nerve (ISN)2,4, and a minor nerve branch, the transverse nerve (TN)7. The SN selectively defasciculates to give rise to two nerve branches called the SNa and SNc, whereas the ISN splits into three nerve branches called the ISN, ISNb, and ISNd2,4. Among them, ISN, ISNb, and SNa motor axon projection patterns are most precisely visualized when late stage-16 embryos are stained with FasII antibody and are filleted (Figure 1C and Figure 2A). The ISN motor neurons extend their axons to innervate dorsal muscles 1, 2, 3, 4, 9, 10, 11, 18, 19, and 202,4 (Figure 2A). The ISNb motor neurons innervate ventrolateral muscles 6, 7, 12, 13, 14, 28, and 302,4 (Figure 2A and 2B). The SNa nerve branch projects to innervate lateral muscles 5, 8, 21, 22, 23, and 242,4 (Figure 2A). The TN, which consists of two motor axons, projects ipsilaterally along the segmental border to innervate muscle 25 and makes synapses with the lateral bipolar dendritic neuron (LBD) in the periphery7 (Figure 2A). These target muscle innervations require not only selective defasciculation of motor axons at specific choice points, but also target muscle recognition. In addition, some putative mesodermal guidepost cells that act as intermediate targets were found in both the ISN and SNa pathways, but not along the ISNb pathway4. This might suggest that ISNb motor axon pathfinding can be regulated in a distinct manner compared to ISN and SNa motor axon guidance, and it also indicates that peripheral motor axon guidance provides an attractive experimental model to study the differential or conserved roles of a single guidance cue molecule8.
This work presents a standard method to visualize the axonal projection patterns of embryonic motor neurons in Drosophila. The described protocols include how to dissect fixed embryos stained with 1D4 antibody and processed in 3,3&#8242;-diaminobenzidine (DAB) for filleted preparations. One critical advantage of the flat preparations of fixed embryos is the better visualization of the axonal projections and muscle patterns in the periphery. Furthermore, this work also shows how to genotype fixed embryos to sort the desired mutant embryos using the LacZ staining method.

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visualization of the axonal projection pattern of embryonic motor neurons in drosophila

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons extend growth cones to navigate along specific pathways by responding to a large number of axon guidance cues that emanate from the surrounding extracellular environment. Growth cone target recognition also plays a critical role in neuromuscular specificity. This work presents a standard immunohistochemistry protocol to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. This protocol includes a few key steps, including a genotyping procedure, to sort the desired mutant embryos; an immunostaining procedure, to tag embryos with fasciclin II (FasII) antibody; and a dissection procedure, to generate filleted preparations from fixed embryos. Motor axon projections and muscle patterns in the periphery are much better visualized in flat preparations of filleted embryos than in whole-mount embryos. Therefore, the filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition, and it can also be applied to both loss-of-function and gain-of-function genetic screens.

Precise connections between motor axons and target muscles during neural development depend on selective axon-axon repulsion and target recognition at specific choice points4. In Drosophila, selective repulsion between motor axons is in part regulated by the combined action of class 1 and 2 semaphorins (Semas), including Sema-1a, Sema-2a, and Sema-2b8,14,15,<...

Watch this Scientific Journal Video about Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila at JoVE.com

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

This work details a standard immunohistochemistry method to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. The filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition during neural development.

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons ...

The overall goal of this experiment is to analyze the motor neuron projection patterns in late stage embryos of Drosophila melanogaster. This method can help answer key questions in the field of neural development, such as motor axon guidance and neural subcu formation. The main advantage of this technique is that it provides precise visualization of motor axon in developing embryos, which allows for reliable analysis of axonal pathfinding and target the condition defect.One day after setting-up egg collection plates replace the plates with new ones containing a dab of freshly-made yeast paste. After three hours on the fresh plates, transfer the flies to new food bottles. And incubate the egg plates overnight at 25 degrees Celsius.In the morning, add 1.8 milliliters of PBT solution to the plates. And use a cotton swab to dislodge the embryos. Then, using a 1 milliliter pipette, transfer the embryos and solution to a micro centrifuge tube.Once the embryos settle to the bottom, aspirate as much solution as possible. Then rinse the embryos twice using one milliliter of PBT per rinse. To decoreonate the embryos, replace the last rinse with one milliliter of 50%bleach and rock the tube for three minutes at room temperature.To remove the bleach, rinse the embryos three times with PBT. Then, replace the last rinse with a half milliliter of 100%heptane and a half milliliter of 4%paraformaldehyde. Now, incubate the embryos for 15 minutes with gentle rocking.Following the incubation, remove the underlying solution layer and add back 500 microliters of 100%methanol. Now, for 30 seconds, shake the tube vigorously to devitellinize the embryos. Then, remove the overlying solution, add back 500 microliters of 100%methanol and shake the tube for another 10 seconds.After the shaking the tube, tap it to help the embryos settle and remove as much solution as possible. Next, briefly rinse the embryos with 100%methanol. Then, immediately rinse the embryos with PBT three times.Now, resuspend the embryos in one milliliter of fresh PBT. And incubate the embryos at 37 degrees Celsius for 15 minutes to prepare them for LacZ staining. While the embryos incubate, prepare a one milliliter aliquot of X-gal staining solution.Warm the aliquot in a 65 degree Celsius water bath until it gets cloudy. Then transfer it to a 37 degree Celsius heat block. After the embryos have been warmed for 15 minutes remove the PBT solution, and replace it with the one milliliter aliquot of staining solution.Then add 20 microliters of X-gal substrate. For about two to four hours, incubate the tube at 37 degrees Celsius with gentle rocking until a blue precipitate forms. Then, remove the staining solution and rinse the embryos three times with room temperature PBT.After the rinses, use a needle probe or forceps to hand-sort the embryos under a dissection microscope. Collect the stained embryos of interest into a half millimeter micro centrifuge tube using a one milliliter pipette. Next, wash the selected embryos with 0.4 milliliters of PBT.Then, replace the PBT with 0.3 milliliters of blocking solution and allow the embryos to incubate with gentle agitation for 15 minutes. After 15 minutes, add 75 microliters of the primary antibody and continue the incubation overnight. The following morning, wash the embryos with PBT for about two hours, changing the wash solution approximately every 30 minutes.After the wash, add 0.3 milliliters of blocking solution containing the secondary antibody and incubate the tube with gentle rocking at room temperature overnight. The next morning, wash the embryos with PBT for about three hours. Change the PBT at least five times during the wash period.After the wash, resuspend the embryos in 0.3 milliliters of dab solution and add two microliters of 3%hydrogen peroxide. Then, incubate them in the dark with gentle rocking at room temperature until enough precipitate is produced. This generally takes 30 to 60 minutes.Then, rinse the embryos with PBT four times and resuspend them in 0.2 milliliters of mounting solution. Now, allow the embryos to equilibrate in the dark at 4 degrees Celsius. To filet the embryos, stage them on a glass slide.First move them into a drop of glycerol ventral side up. Next, under a dissection microscope, cut off the anterior part and 1/4 of the body length and remove the most posterior region, which is free of motor axons. Now, using a needle probe, move the preparation out of the glycerol, oriented dorsal side up and position it horizontally in the field of view.The next step requires a fine tungsten needle that has been inserted into the hollow region of a needle probe, and then bent at the end. Using the tungsten needle, make a small cut at the posterior end of the embryo and continue to cut down the dorsal midline. Make sure the tip of the tungsten needle does not touch the surface of the glass slide during the dissection, because the needle will easily bend against the glass.Then, with the probe return preparation back to the glycerol drop and position it dorsal side up. There, detach the gut from the body wall by unfolding each body wall in a ventral-lateral direction. The two body walls should come apart.Now using the probe, carefully move the body wall flaps onto a glass slide. On the slide, use a tungsten needle to remove the internal organs by pushing them laterally. To stabilize the tungsten needle and keep it from being damaged, keep the hollow needle probed and hold the tungsten needle braced against the surface of the glass slide while manipulating the tungsten needle.Now, mount the preparations in eight microliters of mounting solution on a new glass slide. Attach a cover slip and seal the edges down with regular nail polish. Then, capture images at high resolution using DIC optics.Using the described method, stage 16 embryos were stained with Fas2 antibody and prepared for visualization of the motor neurons and abdominal hemi-segments A3 and A4.Normally, at least seven motor neurons extend and fasciculate their axons to form an ISNb nerve branch. Many surrounding nerve bundles were also identifiable in this preparation. When the intersegmental nerve bundle reaches the choice point at muscle 6 and 7, two axons selectively de-fasciculate and enervate muscles 6 and 7.The same occurs at other choice points as the bundle extends dorsally. Then, at the last choice point, two axons enervate muscle 12. By contrast, in Sema-1a mutant embryos the same axons fail to defasciculate at their choice points.This is not unexpected as the Sema-1 aegean product is known to help form connections between motor axons and target muscles during neural development. Thus, the described method can be used to analyze mutant phenotypes. After watching this video, you should have a good understanding of how to sort the desired mutant embryos, how to immunostain the axonal projection patterns of motor neurons, and how to dissect fixed embryos for flat preparation.

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Research

JoVE Journal

Neuroscience

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the procedure for the visualization of Drosophila embryonic nervous system using antibodies to neuronal proteins. Since the entire embryonic peripheral nervous and central nervous systems are well characterized at the level of individual cells (Dambly-Chaudi&egrave;re et al., 1986; Bodmer et al., 1987; Bodmer et al., 1989), any aberrations to these systems can be easily identified using antibodies to different neuronal proteins. The developing embryos are collected at certain times to ensure that the embryos are in the proper developmental stages for visualization. After collection, the outer layers of the embryo, the chorion membrane and the vitelline envelope that surrounds the embryo, are removed before fixation. Embryos are then incubated with neuronal antibodies and visualized using fluorescently labeled secondary antibodies. Embryos at stages 12-17 are visualized to access the embryonic nervous system. At stage 12 the CNS germ band starts shortening and by stage 15 the definitive pattern of the commissure has been achieved. By stage 17 the CNS contracts and the PNS is fully developed (Campos-Ortega et al. 1985). Thus changes in the pattern of the PNS and CNS can be easily observed during these developmental stages.

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

We describe the procedure to prepare staged Drosophila embryos for the visualization of the embryonic nervous system during embryogenesis.

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the ...

Mechanical Manipulation of Neurons to Control Axonal Development

Cell manipulations and extension of neuronal axons can be accomplished with calibrated glass micro-fibers capable of measuring and applying forces in the 10-1000 &mu;dyne range1,2. Force measurements are obtained through observation of the Hookean bending of the glass needles, which are calibrated by a direct and empirical method3. Equipment requirements and procedures for fabricating, calibrating, treating, and using the needles on cells are fully described. The force regimes previously used and different cell types to which these techniques have been applied demonstrate the flexibility of the methodology and are given as examples for future investigation4-6. The technical advantages are the continuous 'visualization' of the forces produced by the manipulations and the ability to directly intervene in a variety of cellular events. These include direct stimulation and regulation of axonal growth and retraction7; as well as detachment and mechanical measurements on any type of cultured cell8.

Application and direct measurements of forces on neurons in the 2-1000 microdyne range are achieved with high precision using calibrated glass needles. This methodology can be used to control and measure several aspects of axonal development, including axonal initiation, axonal tension, velocity of axonal elongation, and force vectors.

Cell manipulations and extension of neuronal axons can be accomplished with calibrated glass micro-fibers capable of measuring and applying forces in the ...

DiI-Labeling of DRG Neurons to Study Axonal Branching in a Whole Mount Preparation of Mouse Embryonic Spinal Cord

Here we present a technique to label the trajectories of small groups of DRG neurons into the embryonic spinal cord by diffusive staining using the lipophilic tracer 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI)1. The comparison of axonal pathways of wild-type with those of mouse lines in which genes are mutated allows testing for a functional role of candidate proteins in the control of axonal branching which is an essential mechanism in the wiring of the nervous system. Axonal branching enables an individual neuron to connect with multiple targets, thereby providing the physical basis for the parallel processing of information. Ramifications at intermediate target regions of axonal growth may be distinguished from terminal arborization. Furthermore, different modes of axonal branch formation may be classified depending on whether branching results from the activities of the growth cone (splitting or delayed branching) or from the budding of collaterals from the axon shaft in a process called interstitial branching2 (Fig. 1).
The central projections of neurons from the DRG offer a useful experimental system to study both types of axonal branching: when their afferent axons reach the dorsal root entry zone (DREZ) of the spinal cord between embryonic days 10 to 13 (E10 - E13) they display a stereotyped pattern of T- or Y-shaped bifurcation. The two resulting daughter axons then proceed in rostral or caudal directions, respectively, at the dorsolateral margin of the cord and only after a waiting period collaterals sprout from these stem axons to penetrate the gray matter (interstitial branching) and project to relay neurons in specific laminae of the spinal cord where they further arborize (terminal branching)3. DiI tracings have revealed growth cones at the dorsal root entry zone of the spinal cord that appeared to be in the process of splitting suggesting that bifurcation is caused by splitting of the growth cone itself4 (Fig. 2), however, other options have been discussed as well5.
This video demonstrates first how to dissect the spinal cord of E12.5 mice leaving the DRG attached. Following fixation of the specimen tiny amounts of DiI are applied to DRG using glass needles pulled from capillary tubes. After an incubation step, the labeled spinal cord is mounted as an inverted open-book preparation to analyze individual axons using fluorescence microscopy.

The stereotyped projections of sensory afferents into the rodent spinal cord offer an easily accessible experimental system to study axonal branching through the tracing of single axons.

Here we  present a technique to label the trajectories of small groups of DRG neurons  into the embryonic spinal cord by diffusive staining using the ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.
This extracellular technique has advantages. First, extracellular electrodes can stimulate and record neurons through the sheath5, so it does not need to be removed. Thus, neurons will be healthier in extracellular experiments than in intracellular ones. Second, if ganglia are rotated by appropriate pinning of the sheath, extracellular electrodes can access neurons on both sides of the ganglion, which makes it easier and more efficient to identify multiple neurons in the same preparation. Third, extracellular electrodes do not need to penetrate cells, and thus can be easily moved back and forth among neurons, causing less damage to them. This is especially useful when one tries to record multiple neurons during repeating motor patterns that may only persist for minutes. Fourth, extracellular electrodes are more flexible than intracellular ones during muscle movements. Intracellular electrodes may pull out and damage neurons during muscle contractions. In contrast, since extracellular electrodes are gently pressed onto the sheath above neurons, they usually stay above the same neuron during muscle contractions, and thus can be used in more intact preparations.
To uniquely identify motor neurons for a motor pool (in particular, the I1/I3 muscle in Aplysia) using extracellular electrodes, one can use features that do not require intracellular measurements as criteria: soma size and location, axonal projection, and muscle innervation4,6,7. For the particular motor pool used to illustrate the technique, we recorded from buccal nerves 2 and 3 to measure axonal projections, and measured the contraction forces of the I1/I3 muscle to determine the pattern of muscle innervation for the individual motor neurons.
We demonstrate the complete process of first identifying motor neurons using muscle innervation, then characterizing their timing during motor patterns, creating a simplified diagnostic method for rapid identification. The simplified and more rapid diagnostic method is superior for more intact preparations, e.g. in the suspended buccal mass preparation8 or in vivo9. This process can also be applied in other motor pools10,11,12 in Aplysia or in other animal systems2,3,13,14.

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.

In animals with large  identified neurons (e.g. mollusks), analysis of motor pools is done using  intracellular techniques1,2,3,4. Recently,  we developed ...

Real-time Imaging of Axonal Transport of Quantum Dot-labeled BDNF in Primary Neurons

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are increasingly viewed as an early feature of neurodegenerative diseases, including Alzheimer&#8217;s disease (AD) and Huntington&#8217;s disease (HD). As yet unclear is the mechanism(s) by which axonal injury is induced. We reported the development of a novel technique to produce biologically active, monobiotinylated BDNF (mBtBDNF) that can be used to trace axonal transport of BDNF. Quantum dot-labeled BDNF (QD-BDNF) was produced by conjugating quantum dot 655 to mBtBDNF. A microfluidic device was used to isolate axons from neuron cell bodies. Addition of QD-BDNF to the axonal compartment allowed live imaging of BDNF transport in axons. We demonstrated that QD-BDNF moved essentially exclusively retrogradely, with very few pauses, at a moving velocity of around 1.06 &#956;m/sec. This system can be used to investigate mechanisms of disrupted axonal function in AD or HD, as well as other degenerative disorders.

Axonal transport of BDNF, a neurotrophic factor, is critical for the survival and function of several neuronal populations. Some degenerative disorders are marked by disruption of axonal structure and function. We demonstrated the techniques used to examine live trafficking of QD-BDNF in microfluidic chambers using primary neurons.

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are ...

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic pathways. Quantitative descriptions are rarely performed, although genetic manipulations produce a range of phenotypic effects and variations are observed even among individuals within control groups.&#160;Emerging evidence shows that morphology, size and location of organelles play a previously underappreciated, yet fundamental role in cell function and survival. Here we provide step-by-step instructions for performing quantitative analyses of phenotypes at the Drosophila larval neuromuscular junction (NMJ). We use several reliable immuno-histochemical markers combined with bio-imaging techniques and morphometric analyses to examine the effects of genetic mutations on specific cellular processes. In particular, we focus on the quantitative analysis of phenotypes affecting morphology, size and position of nuclei within the striated muscles of Drosophila larvae. The Drosophila larval NMJ is a valuable experimental model to investigate the molecular mechanisms underlying the structure and the function of the neuromuscular system, both in health and disease. However, the methodologies we describe here can be extended to other systems as well.

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Morphology, size and location of intracellular organelles are evolutionarily conserved and appear to directly affect their function. Understanding the molecular mechanisms underlying these processes has become an important goal of modern biology. Here we show how these studies can be facilitated by the application of quantitative techniques.

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic ...

Retrograde Neuroanatomical Tracing of Phrenic Motor Neurons in Mice

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic nerves innervating the respiratory diaphragm. In spinal cord slices, phrenic motor neurons cannot be identified from other motor neurons on morphological or biochemical criteria. We provide the description of procedures for visualizing phrenic motor neuron cell bodies in mice, following intrapleural injections of cholera toxin subunit beta (CTB) conjugated to a fluorophore. This fluorescent neuroanatomical tracer has the ability to be caught up at the diaphragm neuromuscular junction, be carried retrogradely along the phrenic axons and reach the phrenic cell bodies. Two methodological approaches of intrapleural CTB delivery are compared: transdiaphragmatic versus transthoracic injections. Both approaches are successful and result in similar number of CTB-labeled phrenic motor neurons. In conclusion, these techniques can be applied to visualize or quantify the phrenic motor neurons in various experimental studies such as those focused on the diaphragm-phrenic circuitry.

Here, we describe a protocol for identifying phrenic motor neurons in mice after intrapleural delivery of fluorophore conjugated cholera toxin subunit beta. Two techniques are compared to inject the pleural cavity: transdiaphragmatic versus transthoracic approaches.

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.