This work was supported by grants to D.K. from the Medical Research Foundation of Oregon and from NIH/NINDS (NS047663). E.S. was supported by a training grant from the NIH (T32AG023477).

1. Fixing the Head on Collars and Embedding in Paraffin
Note: All of the steps in the fixation process should be done in a fume hood. Methylbenzoate, while not posing a health risk, has a highly distinct odor, which can be overwhelming if not handled in a fume hood.
<ol>
	<li>Before anesthetizing the flies, make up 50 mL of Carnoy solution by adding 15 mL of chloroform and 5 mL of glacial acetic acid to 30 mL of 99% ethanol (do not mix the chloroform and acetic acid). Pour it in a glass container with a flat bottom, such as a crystalizing dish, to ensure that the collars can lay flat and are completely covered by the solution.</li>
	<li>Anesthetize the flies with CO2 or ether.</li>
	<li>Thread flies (up to 20 with most collars) by their necks into the collars using forceps. Remember to align all of the heads in the same orientation, as seen in Figure 1A, and be gentle to ensure that no damage occurs to the head or eyes.</li>
	<li>Include sine oculis flies (arrows, Figure 1A) at random positions so that the order of the flies can be easily identified in the sections. In addition, if the experimental flies have a light or white eye color, thread some red-eyed flies, such as wild type, between them to ensure that sufficient pigment is present to stain the slide. Record the order of the flies on a protocol sheet together with the collar number if using more than one collar.</li>
	<li>Once a collar has been finished, place it in the prepared Carnoy solution for 3.5 - 4 h.</li>
	<li>Dump out the Carnoy solution into the appropriate disposal canister and begin the ethanol washes. Make sure to pour slowly so as not to disturb the placement of the collars in the container.</li>
	<li>Wash the collars for 30 min in 99% ethanol twice.</li>
	<li>Wash the collars in 100% ethanol for 1 h. Be sure to change the washes on time to prevent overdehydration.</li>
	<li>Put the collars in methylbenzoate O/N at RT. Seal the container with parafilm to prevent the evaporation of the methylbenzoate.</li>
	<li>Pour the methylbenozate into the proper disposable container in the fume hood. Add a previously-prepared mixture of 1:1 low melting point (56 - 57 &#176;C) paraffin wax and methylbenzoate for 1h. From this point on, the collars need to be kept in an incubator at 65 &#176;C to make sure that the paraffin does not harden.</li>
	<li>Pour out the methylbenozate and paraffin mixture into the proper disposable container, and pour molten pure paraffin wax, kept at 65 &#176;C, onto the collars.</li>
	<li>Change the paraffin after 30 min and repeat this at least 5 times. At least 6 - 8 washes should be performed.</li>
	<li>Once the washes are complete, place the collars into a rubber ice cube tray with slots approximately the size of the collars. Pour molten paraffin over them until completely covered and allow it to harden O/N (try to avoid air bubbles).</li>
	<li>Remove the paraffin blocks containing the collars from the ice cube tray. Separate the paraffin block from the collar using a razor blade, gently breaking off the collar. The heads will be in the paraffin block while the bodies will stay in the collar. The blocks can be kept at room temperature.</li>
	<li>To clean the collars, soak them in a deparafinization agent at 65 &#176;C to remove the paraffin, clean with light scrubbing, and wash in ethanol before reusing.</li>
</ol>
2. Sectioning and Mounting
<ol>
	<li>Warm a heating plate to 50 &#176;C. Place the object holders (or metal mounting blocks) and razor blades on the plate and let them warm up.</li>
	<li>Depending on the desired orientation for sectioning, attach the paraffin block either with the heads towards the side (for horizontal sections) or facing upwards (for frontal sections) to a heated mounting block (briefly melting the block at the contact side). Remove the block from the heating plate and allow it to cool for at least 10 min to ensure that the paraffin is hardened enough for a proper seal onto the mounting block. Keep the row of heads aligned in parallel with the surface of the block as much as possible to prevent uneven sections.</li>
	<li>Take a razor blade and trim the excess paraffin away from the fly heads so that only a small row with the embedded heads remains (the razor blade can be warmed up for easier trimming). Make sure not to trim too much so that the paraffin does not break during sectioning (more trimming can be done during sectioning).</li>
	<li>Place the mounting block into the object holder of the microtome and ensure that the alignment of the row of heads is as parallel as possible to the edge of the blade.</li>
	<li>Prepare microscope slides by covering them with a thin layer of poly-L-Lysine (PLL) solution and let them dry for 5 min. Cover them with water shortly before use.</li>
	<li>Cut 7&#160;&#181;m sections and transfer the ribbon of sections to the slide floating on the water. 
	NOTE: To obtain the entire brain for horizontal sections, we collect the ribbon from when starting to cut into the eye until the head has been completely cut (cutting from the proboscis into the brain). More than one slide may be needed for the entire head.</li>
	<li>Place the slide on a heat plate at 37 &#176;C and allow the ribbon to expand for about 1 min.</li>
	<li>Remove excess water (by pouring it off or using a tissue) and dry the slides O/N.</li>
	<li>Remove the paraffin wax from the slide by placing the slides in a tall, vertical slide-staining jar filled with a deparafinization agent (completely covering the sections). Perform 3 washes of 30 min - 60 min each.</li>
	<li>Remove the slide from the final wash. Place 2 drops of embedding media onto the slide and cover it with a large coverslip.</li>
</ol>
3. Photographing and Analyzing the Sections
<ol>
	<li>Allow the prepared slides to dry for 1 - 2 d. Then, examine them on a fluorescence microscope under blue light.</li>
	<li>Use a lower magnification to determine the orientation of the flies and to find the region of interest if focusing on a specific region. 
	NOTE: For the sws flies (see Figure 2), we find the section that contains the great commissure and take an image (usually at 40X magnification). When analyzing the entire brain (as in Figure 3), we scroll through all the sections from a head and either photograph the section with the most severe phenotype or all sections that show vacuoles.</li>
	<li>For a double-blind analysis, take and number images without knowing the genotype and record the collar number and position of the head in the row to identify them later.</li>
	<li>Once the images have been taken, analyze them using an imaging software.</li>
	<li>Count the number of vacuoles per section or per head. To measure the vacuole size, open the images in a software program and select the vacuoles with a selection tool. Determine the amount of pixels in the selected vacuoles.</li>
	<li>For conversion to &#181;m2, take a photo of a stage micrometer calibration slide at the magnification used for acquiring photos. Determine the amount of pixels in 100 &#181;m2 to calculate a conversion factor.</li>
	<li>Convert the total pixel number into &#181;m2 by dividing the number of pixels by the conversion factor calculated in the above step.</li>
</ol>

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The authors have nothing to disclose.

The described method provides a means to quantify neurodegeneration in the brain of Drosophila. While other methods, like counting a specific cell type, can be used to identify neurodegeneration, the advantage of this method is that it can be applied more generally. Counting cells requires that these cells can be reliably identified using either a specific antibody or the expression of a cell-specific marker, which is not always available. Furthermore, it has been shown that dramatically different results can be...

With the increase in life expectancy, neurodegenerative diseases like Alzheimer&#39;s or Parkinson&#39;s have become an increasing health threat for the general population. According to the National Institutes of Health, 115 million people worldwide are predicted to be affected by dementia in 2050. Although significant progress has been made in identifying genes and risk factors involved in at least some of these diseases, for many of them, the underlying molecular mechanisms are still unknown or not well understood.
Simple invertebrate model organisms like Caenorhabditis elegans and Drosophila melanogaster offer a variety of experimental advantages to study the mechanisms of neurodegenerative diseases, including a short life cycle, large number of progeny, and the availability of well-established and sometimes unique genetic and molecular methods1-12. Furthermore, these organisms are amenable to unbiased interaction screens that can identify factors contributing to these diseases by their aggravating or ameliorating effects on neurodegenerative phenotypes.
Analyzing such genetic interactions and assessing aging effects requires quantitative protocols to detect neurodegeneration and to measure its severity. This assessment can be done relatively easily when measuring behavioral aspects in Drosophila, such as olfactory learning, negative geotaxis, or fast phototaxis, which provide a numeric performance value13-21. It is also possible to determine the effects on neuronal survival by counting neurons. However, this is only possible when focusing on a specific population that is clearly identifiable, like the dopaminergic neurons that are affected in PD, and even then, the results have been controversial22-24.
The protocol described here uses the collar method to perform paraffin serial sections, a method that was originally developed by Heisenberg and B&#246;hl, who used it to isolate anatomical brain mutants in Drosophila25. The use of the collar method has subsequently been adapted, including in cryosections, vibratome sections, and plastic sections26-28. Here, this method is employed to obtain serial sections of the entire fly head, which can then be used to measure the vacuoles that develop in flies with neurodegenerative phenotypes16,21,29-32. These measurements can be done in specific brain areas or can cover the entire brain; the latter approach allows one to identify even weak degenerative phenotypes, as observed during aging. Finally, when using the collars, up to 20 flies can be processed as one preparation, which is not only less time-consuming, but also allows for the analysis of control and experimental flies in the same preparation, minimizing artifacts due to slight changes in the preparation.

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			<td>Name of the Reagent/Equipment</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td></td>
		</tr>
		<tr>
			<td>Collar</td>
			<td>Genesee Scientific</td>
			<td>TS 48-100</td>
			<td>We are using custom made collars that are made from one piece of metal instead of layers as the ones by Genesee. A discription to make collars can be found at http://flybrain.neurobio.arizona.edu/Flybrain/html/atlas/fluorescent/index.html&#160;</td>
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			<td>Rubber ice cube tray for embedding</td>
			<td>Household store</td>
			<td></td>
			<td>The size can be made to fit by glueing in additional walls&#160;</td>
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			<td>Crystallizing dish</td>
			<td>Fisher Scientific company</td>
			<td>08-762-3</td>
			<td></td>
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			<td>Ether</td>
			<td>Fisher Scientific Company</td>
			<td>E138-1</td>
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			<td>Decon Laboratories Inc.</td>
			<td>2701</td>
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			<td>Fisher Scientific Company</td>
			<td>C298-500</td>
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			<td>Glacial Acetic Acid</td>
			<td>Fisher Scientific Company</td>
			<td>A38-212</td>
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			<td>Fisher Scientific Company</td>
			<td>M205-500</td>
			<td>Distinct Odor</td>
		</tr>
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			<td>Use in fume hood</td>
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			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Low Melting Point Paraffin Wax</td>
			<td>Fisher Scientific Company</td>
			<td>T565</td>
			<td>Make sure to keep extra melted in a 65&#176;C waterbath</td>
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			<td>Microtome</td>
			<td>Leica Biosystems</td>
			<td>Reichert Jung 2040 Autocut</td>
			<td></td>
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			<td>Microscope Slide</td>
			<td>Fisher Scientific Company</td>
			<td>12-550D</td>
			<td></td>
		</tr>
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			<td>Microscope Cover Glass</td>
			<td>Fisher Scientific Company</td>
			<td>12-545-M</td>
			<td></td>
		</tr>
		<tr>
			<td>SafeClear</td>
			<td>Fisher Scientific Company</td>
			<td>314-629</td>
			<td>Three different containers for washes</td>
		</tr>
		<tr>
			<td>Vertical Staining Jar with Cover</td>
			<td>Ted Pella Inc.&#160;</td>
			<td>432-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Permount</td>
			<td>Fisher Scientific Company</td>
			<td>SP15-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-L-lysine Solution</td>
			<td>Sigma Life Science</td>
			<td>P8290-500</td>
			<td></td>
		</tr>
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mass histology to quantify neurodegeneration in drosophila

Progressive neurodegenerative diseases like Alzheimer&#39;s disease (AD) or Parkinson&#39;s disease (PD) are an increasing threat to human health worldwide. Although mammalian models have provided important insights into the underlying mechanisms of pathogenicity, the complexity of mammalian systems together with their high costs are limiting their use. Therefore, the simple but well-established Drosophila model-system&#160;provides an alternative for investigating the molecular pathways that are affected in these diseases. Besides behavioral deficits, neurodegenerative diseases are characterized by histological phenotypes such as neuronal death and axonopathy. To quantify neuronal degeneration and to determine how it is affected by genetic and environmental factors, we use a histological approach that is based on measuring the vacuoles in adult fly brains. To minimize the effects of systematic error and to directly compare sections from control and experimental flies in one preparation, we use the &#39;collar&#39; method for paraffin sections. Neurodegeneration is then assessed by measuring the size and/or number of vacuoles that have developed in the fly brain. This can either be done by focusing on a specific region of interest or by analyzing the entire brain by obtaining serial sections that span the complete head. Therefore, this method allows one to measure not only severe degeneration but also relatively mild phenotypes that are only detectable in a few sections, as occurs during normal aging.

Using the described method results in serial sections stained by the eye pigment33 that encompass the entire fly head. A part of this is shown in Figure 1B, where the sections from an individual head are shown from top to bottom. The sections from different flies are seen left to right in this example. To facilitate orientation and identification of the flies, an eyeless fly (sine oculis) is inserted as a marker at position 3 (arrow, Figure 1B<...

Watch this Scientific Journal Video about Mass Histology to Quantify Neurodegeneration in Drosophila at JoVE.com

Mass Histology to Quantify Neurodegeneration in Drosophila

Drosophila is widely used as a model system to study neurodegeneration. This protocol describes a method by which degeneration, as determined by vacuole formation in the brain, can be quantified. It also minimizes effects due to the experimental procedure by processing and sectioning control and experimental flies as one sample.

Mass Histology to Quantify Neurodegeneration in Drosophila

Progressive neurodegenerative diseases like Alzheimer&#39;s disease (AD) or Parkinson&#39;s disease (PD) are an increasing threat to human health ...

mass-histology-to-quantify-neurodegeneration-in-drosophila

Research

JoVE Journal

Neuroscience

10.1K Views.  Oregon Health & Sciences University. Drosophila is widely used as a model system to study neurodegeneration. This protocol describes a method by which degeneration, as determined by vacuole formation in the brain, can be quantified. It also minimizes effects due to the experimental procedure by processing and sectioning control and experimental flies as one sample.

Video: Mass Histology to Quantify Neurodegeneration in Drosophila

Quantifying Synapses: an Immunocytochemistry-based Assay to Quantify Synapse Number

One of the most important goals in neuroscience is to understand the molecular cues that instruct early stages of synapse formation. As such it has become imperative to develop objective approaches to quantify changes in synaptic connectivity. Starting from sample fixation, this protocol details how to quantify synapse number both in dissociated neuronal culture and in brain sections using immunocytochemistry. Using compartment-specific antibodies, we label presynaptic terminals as well as sites of postsynaptic specialization. We define synapses as points of colocalization between the signals generated by these markers. The number of these colocalizations is quantified using a plug in Puncta Analyzer (written by Bary Wark, available upon request, c.eroglu@cellbio.duke.edu) under the ImageJ analysis software platform. The synapse assay described in this protocol can be applied to any neural tissue or culture preparation for which you have selective pre- and postsynaptic markers. This synapse assay is a valuable tool that can be widely utilized in the study of synaptic development.

This protocol details how to quantify synapse number both in dissociated neuronal culture and in brain sections using immunocytochemistry. Using compartment-specific antibodies, we label presynaptic terminals as well as sites of postsynaptic specialization. We define synapses as points of colocalization between the signals generated by these markers.

One of the most important goals in neuroscience is to understand the molecular cues that instruct early stages of synapse formation. As such it has become ...

Combining Lipophilic dye, in situ Hybridization, Immunohistochemistry, and Histology

Going beyond single gene function to cut deeper into gene regulatory networks requires multiple mutations combined in a single animal. Such analysis of two or more genes needs to be complemented with in situ hybridization of other genes, or immunohistochemistry of their proteins, both in whole mounted developing organs or sections for detailed resolution of the cellular and tissue expression alterations. Combining multiple gene alterations requires the use of cre or flipase to conditionally delete genes and avoid embryonic lethality. Required breeding schemes dramatically enhance effort and cost proportional to the number of genes mutated, with an outcome of very few animals with the full repertoire of genetic modifications desired. Amortizing the vast amount of effort and time to obtain these few precious specimens that are carrying multiple mutations necessitates tissue optimization. Moreover, investigating a single animal with multiple techniques makes it easier to correlate gene deletion defects with expression profiles. We have developed a technique to obtain a more thorough analysis of a given animal; with the ability to analyze several different histologically recognizable structures as well as gene and protein expression all from the same specimen in both whole mounted organs and sections. Although mice have been utilized to demonstrate the effectiveness of this technique it can be applied to a wide array of animals. To do this we combine lipophilic dye tracing, whole mount in situ hybridization, immunohistochemistry, and histology to extract the maximal possible amount of data.

Combining Lipophilic dye, in situ Hybridization, Immunohistochemistry, and Histology

A combination of different techniques to maximize data collection from mouse tissue is presented.

Going beyond single gene function to cut deeper into gene regulatory networks requires multiple mutations combined in a single animal.  Such analysis of ...

Assaying Locomotor, Learning, and Memory Deficits in Drosophila Models of Neurodegeneration

Advances in genetic methods have enabled the study of genes involved in human neurodegenerative diseases using Drosophila as a model system1. Most of these diseases, including Alzheimer's, Parkinson's and Huntington's disease are characterized by age-dependent deterioration in learning and memory functions and movement coordination2. Here we use behavioral assays, including the negative geotaxis assay3 and the aversive phototaxic suppression assay (APS assay)4,5, to show that some of the behavior characteristics associated with human neurodegeneration can be recapitulated in flies. In the negative geotaxis assay, the natural tendency of flies to move against gravity when agitated is utilized to study genes or conditions that may hinder locomotor capacities. In the APS assay, the learning and memory functions are tested in positively-phototactic flies trained to associate light with aversive bitter taste and hence avoid this otherwise natural tendency to move toward light. Testing these trained flies 6 hours post-training is used to assess memory functions. Using these assays, the contribution of any genetic or environmental factors toward developing neurodegeneration can be easily studied in flies.

Assaying Locomotor, Learning, and Memory Deficits in Drosophila Models of Neurodegeneration

Behavior assays for measuring locomotor functions, learning, and memory abilities in Drosophila.

Advances in genetic methods have enabled the study of genes involved in human neurodegenerative diseases using Drosophila as a model system1. Most of ...

In vivo Laser Axotomy in C. elegans

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron is damaged by injury or disease, it may regenerate. Cell-intrinsic and extrinsic factors influence the ability of a neuron to regenerate and restore function. Recently, the nematode C. elegans has emerged as an excellent model organism to identify genes and signaling pathways that influence the regeneration of neurons1-6. The main way to initiate neuronal regeneration in C. elegans is laser-mediated cutting, or axotomy. During axotomy, a fluorescently-labeled neuronal process is severed using high-energy pulses. Initially, neuronal regeneration in C. elegans was examined using an amplified femtosecond laser5. However, subsequent regeneration studies have shown that a conventional pulsed laser can be used to accurately sever neurons in vivo and elicit a similar regenerative response1,3,7.
We present a protocol for performing in vivo laser axotomy in the worm using a MicroPoint pulsed laser, a turnkey system that is readily available and that has been widely used for targeted cell ablation. We describe aligning the laser, mounting the worms, cutting specific neurons, and assessing subsequent regeneration. The system provides the ability to cut large numbers of neurons in multiple worms during one experiment. Thus, laser axotomy as described herein is an efficient system for initiating and analyzing the process of regeneration.

In vivo Laser Axotomy in C. elegans

A protocol to cut neurons in C. elegans with a MicroPoint pulsed laser is presented. We describe setting up the system, immobilizing worms, and severing labeled neurons. Advantages include a relatively low-cost system and the ability to sever neuronal processes or ablate cells in vivo.

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron ...

Methods to Assay Drosophila Behavior

Drosophila melanogaster, the fruit fly, has been used to study molecular mechanisms of a wide range of human diseases such as cancer, cardiovascular disease and various neurological diseases1. We have optimized simple and robust behavioral assays for determining larval locomotion, adult climbing ability (RING assay), and courtship behaviors of Drosophila. These behavioral assays are widely applicable for studying the role of genetic and environmental factors on fly behavior. Larval crawling ability can be reliably used for determining early stage changes in the crawling abilities of Drosophila larvae and also for examining effect of drugs or human disease genes (in transgenic flies) on their locomotion. The larval crawling assay becomes more applicable if expression or abolition of a gene causes lethality in pupal or adult stages, as these flies do not survive to adulthood where they otherwise could be assessed. This basic assay can also be used in conjunction with bright light or stress to examine additional behavioral responses in Drosophila larvae. Courtship behavior has been widely used to investigate genetic basis of sexual behavior, and can also be used to examine activity and coordination, as well as learning and memory. Drosophila courtship behavior involves the exchange of various sensory stimuli including visual, auditory, and chemosensory signals between males and females that lead to a complex series of well characterized motor behaviors culminating in successful copulation. Traditional adult climbing assays (negative geotaxis) are tedious, labor intensive, and time consuming, with significant variation between different trials2-4. The rapid iterative negative geotaxis (RING) assay5 has many advantages over more widely employed protocols, providing a reproducible, sensitive, and high throughput approach to quantify adult locomotor and negative geotaxis behaviors. In the RING assay, several genotypes or drug treatments can be tested simultaneously using large number of animals, with the high-throughput approach making it more amenable for screening experiments.

Methods to Assay Drosophila Behavior

Drosophila melanogaster is a genetically and behaviorally tractable model system that has been used to understand the molecular and cellular basis of many important biological processes for over a century 1. Drosophila has been well exploited to gain insights into the genetic basis of fly behavior.

Drosophila melanogaster, the fruit fly, has been used to study molecular mechanisms of a wide range of human diseases such as cancer, cardiovascular ...

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 ...

Utilizing 3D Printing Technology to Merge MRI with Histology: A Protocol for Brain Sectioning

Magnetic resonance imaging (MRI) allows for the delineation between normal and abnormal tissue on a macroscopic scale, sampling an entire tissue volume three-dimensionally. While MRI is an extremely sensitive tool for detecting tissue abnormalities, association of signal changes with an underlying pathological process is usually not straightforward. In the central nervous system, for example, inflammation, demyelination, axonal damage, gliosis, and neuronal death may all induce similar findings on MRI. As such, interpretation of MRI scans depends on the context, and radiological-histopathological correlation is therefore of the utmost importance. Unfortunately, traditional pathological sectioning of brain tissue is often imprecise and inconsistent, thus complicating the comparison between histology sections and MRI. This article presents novel methodology for accurately sectioning primate brain tissues and thus allowing precise matching between histology and MRI. The detailed protocol described in this article will assist investigators in applying this method, which relies on the creation of 3D printed brain slicers. Slightly modified, it can be easily implemented for brains of other species, including humans.

The overall goal of this protocol is to accurately align magnetic resonance imaging (MRI) image volumes with histology sections via the creation of customized 3D-printed brain holders and slicer boxes.

Magnetic resonance imaging (MRI) allows for the delineation between normal and abnormal tissue on a macroscopic scale, sampling an entire tissue volume ...

A Novel Method to Model Chronic Traumatic Encephalopathy in Drosophila

Chronic Traumatic Encephalopathy (CTE) is an established neurodegenerative disease that is closely associated with exposure to repetitive mild Traumatic Brain Injury (mTBI). The mechanisms responsible for its complex pathological changes remain largely elusive, despite a recent consensus to define the neuropathological criteria. Here, we describe a novel method to develop a model of CTE in Drosophila melanogaster&#160;(Drosophila&#160;)&#160;in an attempt to identify the key genes and pathways that lead to the characteristic hyperphosphorylated tau accumulation and neuronal death in the brain. Adjustable-strength impacts to inflict mild closed injury are delivered directly to the fly head, subjecting the head to rapid acceleration and deceleration. Our method eliminates the potential problems inherent with other Drosophila mTBI models (e.g.,animal death might be induced by damage to other parts of the body or to internal organs). The less labor- and cost-intensive animal care, short life span, and extensive genetic tools make the fruit fly ideal to study CTE pathogenesis and make it possible to perform large-scale, genome-wide forward genetic and pharmacological screens. We anticipate that the ongoing characterization of the model will generate important mechanistic insights on disease prevention and therapeutic approaches.

A Novel Method to Model Chronic Traumatic Encephalopathy in Drosophila

Here, we describe a new approach to inflict closed-head traumatic brain injury in Drosophila melanogaster. Our method has the advantage of directly delivering repetitive impacts with adjustable strength to the head alone. Further exploration of the invertebrate system will help to illuminate the pathogenesis of chronic traumatic encephalopathy.

Chronic Traumatic Encephalopathy (CTE) is an established neurodegenerative disease that is closely associated with exposure to repetitive mild Traumatic ...

Modeling Amyloid-&#946;42 Toxicity and Neurodegeneration in Adult Zebrafish Brain

Alzheimer's disease (AD) is a debilitating neurodegenerative disease in which accumulation of toxic amyloid-&#946;42 (A&#946;42) peptides leads to synaptic degeneration, inflammation, neuronal death, and learning deficits. Humans cannot regenerate lost neurons in the case of AD in part due to impaired proliferative capacity of the neural stem/progenitor cells (NSPCs) and reduced neurogenesis. Therefore, efficient regenerative therapies should also enhance the proliferation and neurogenic capacity of NSPCs. Zebrafish (Danio rerio) is a regenerative organism, and we can learn the basic molecular programs with which we could design therapeutic approaches to tackle AD. For this reason, the generation of an AD-like model in zebrafish was necessary. Using our methodology, we can introduce synthetic derivatives of A&#946;42 peptide with tissue penetrating capability into the adult zebrafish brain, and analyze the disease pathology and the regenerative response. The advantage over the existing methods or animal models is that zebrafish can teach us how a vertebrate brain can naturally regenerate, and thus help us to treat human neurodegenerative diseases better by targeting endogenous NSPCs. Therefore, the amyloid-toxicity model established in the adult zebrafish brain may open new avenues for research in the field of neuroscience and clinical medicine. Additionally, the simple execution of this method allows for cost-effective and efficient experimental assessment. This manuscript describes the synthesis and injection of A&#946;42 peptides into zebrafish brain.

This protocol describes the synthesis, characterization, and injection of monomeric amyloid-&#946;42 peptides for generating amyloid toxicity in adult zebrafish to establish an Alzheimer&#39;s disease model, followed by histological analyses and detection of aggregations.

Alzheimer's disease (AD) is a debilitating neurodegenerative disease in which accumulation of toxic amyloid-&#946;42 (A&#946;42) peptides leads to ...

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. ...