The authors thank Dr. F. Bryan Pickett and Dr. Rodney Dale for use of equipment for injection. This work was funded by research funding from Loyola University Chicago to M.D., D.T., and J.J.

1. Collection of Samples
<ol>
	<li>Embryo collection
	<ol>
		<li>In each embryo collection cage, use a minimum of 50 virgin females with 20&#8722;25 males for collections. Incubate these flies in a bottle with cornmeal-agar food (Table of Materials) for 1&#8722;2 days before beginning collections23. 
		NOTE: More flies can be used, but the female-to-male ratio should be kept at 2:1.</li>
		<li>Pre-warm apple juice agar plates (Table 1) at 25 &#176;C overnight. 
		NOTE: This is required for proper staging of embryos. If plates are drying out quickly, add a bowl with water to the incubator to increase the humidity of the chamber.</li>
		<li>Anesthetize flies from step 1.1.1 with CO2 and transfer flies to a collection cage. Place a pre-warmed apple juice agar plate with a small smear of yeast paste on the open end and secure to the cage with the red sleeve (Table of Materials). In order to clear older embryos, allow flies to lay embryos/eggs on an apple juice agar plate for 1 h at 25 &#176;C.</li>
		<li>Remove the collection cage from the incubator. Invert the cage mesh side down and tap flies down to the bottom of the cage. Replace the apple juice agar with a new pre-warmed apple juice agar plate with a small smear of yeast paste. Discard the first plate.</li>
		<li>Allow flies to lay embryos/eggs on the new apple juice agar plate for 1 h at 25 &#176;C. Discard this plate following the 1 h collection and proceed to the next step to collect embryos for injection.</li>
		<li>To collect late stage 17 embryos (20&#8722;21 h AEL), allow flies of desired genotype from steps 1.1.1&#8722;1.1.5 to lay on a new pre-warmed apple juice agar plates with a small smear of yeast paste at 25 &#176;C for 1 h. Age plate for 19 h in a 25 &#176;C incubator, so embryos will be 20&#8722;21 h of age at the time of imaging. 
		NOTE: This step can be repeated as desired for multiple rounds of sample collection, injection, and imaging.</li>
		<li>Collect embryos from plates into a cell strainer with 70 &#956;m nylon mesh by adding phosphate-buffered saline (PBS) with 0.1% nonionic surfactant (PBTx; Table 1) to cover the surface of the plate and loosen embryos from the surface using a paintbrush.</li>
		<li>Dechorionate embryos collected in the cell strainer in a 50% bleach solution (Table 1) in a 100 mm Petri dish for 5 min with occasional agitation at room temperature. Rinse embryos 3x by swirling the cell strainer in PBTx in a Petri dish, using a fresh dish of PBTx each time.</li>
		<li>If all embryos are of the correct genotype, proceed directly to step 1.1.10. If generation of embryos of the correct genotype requires a cross with heterozygous flies, select embryos of the correct genotype using the presence or absence of fluorescently marked balancer chromosomes. Use a stereomicroscope with fluorescent capabilities for genotype selection. 
		NOTE: Balancer chromosomes marked with Deformed-yellow fluorescent protein; Kruppel-Gal4, UAS-green fluorescent protein (GFP); and twist-Gal4, UAS-GFP work well for genotype selection in late embryogenesis (Table 2)24,25,26.</li>
		<li>Using a glass pipette, transfer embryos in PBTx to a 2% agarose gel slab (Table 1). Remove excess liquid with filter paper. Align ~6&#8722;8 embryos on the 2% agarose gel slab with posterior to the right and dorsal side facing up (Figure 1B). 
		NOTE: The micropyle, the small hole through which spermatozoa enter the egg, is located at the anterior end of the embryo. The posterior end is more rounded. The trachea appears white and is located dorsally in the embryo, allowing for the distinction of the dorsal and ventral sides of the embryo (Figure 1B).</li>
		<li>Prepare a slide with one piece of double-sided tape. Firmly press the slide on top of the embryos to transfer them to the double-sided tape.</li>
		<li>Desiccate embryos by incubation at room temperature for ~25 min (no desiccant is used). Following desiccation, cover embryos with halocarbon oil. 
		NOTE: Desiccation periods may vary depending on the temperature, humidity, and ventilation in the room. The incubation period should be used to set up the apparatus for injection and the confocal microscope for imaging, as described in sections 2 and 3 of this protocol.</li>
	</ol>
	</li>
	<li>Larval collection
	<ol>
		<li>Set up a cross with 5&#8722;10 virgin female flies of the desired genotype and half as many males of the desired genotype in a vial with cornmeal-agar food and incubate at 25 &#176;C23.</li>
		<li>After 5&#8722;7 days, depending on the genotype, collect wandering third instar larvae from the vial gently with forceps. Rinse larvae in 1x PBS to remove food stuck to the larvae. Transfer larvae to an apple juice agar plate for genotyping as described in step 1.1.9 if necessary.</li>
		<li>Roll larvae on a tissue using a paintbrush to dry them off. Transfer 6&#8722;8 larvae to a slide prepared with double-sided tape using forceps.</li>
	</ol>
	</li>
</ol>
2. Preparation of Needles and Specimen Injection
<ol>
	<li>Pull needles on a micropipette puller prior to the initiation of this protocol. Secure capillary tubes into the needle puller and pull according to the standard needle shape and parameters for D. melanogaster injections (Table 3)27. Store needles in a Petri dish by anchoring in clay until use for injection.</li>
	<li>Load a needle with 5 &#956;L of 10 kDa dextran conjugated to sulforhodamine 101 acid chloride (Table of Materials) using a 20 &#956;L gel-loading pipette tip during the 25-min desiccation period for embryos (step 1.1.12), or immediately following the transfer of larvae to the slide (step 1.2.3).</li>
	<li>Load the needle into a needle holder and position in a micromanipulator secured to a steel base (Table of Materials). 
	NOTE: The needle should be nearly parallel to the microscope stage for embryo injection and angled slightly downward for larval injections.</li>
	<li>Set the injection apparatus (Table of Materials) to 50 psi, 5&#8722;10 ms with a range of 10. 
	NOTE: It may be necessary to alter these settings for the particular injection apparatus being used.</li>
	<li>Place the slide on stage and brush edge of the needle against the edge of the double-sided tape at a 45&#176; angle to create an angled, broken tip. 
	NOTE: For embryos it is only necessary to break the tip enough to allow for flow of the 10 kDa dextran. A perfect needle has a slightly angled tip and only a small drop of dye will come out with each injection. For larvae, it is necessary to break the tip more, but with an angled tip to penetrate the larval body wall. A larger drop of dye will come out.</li>
	<li>Pump foot pedal until the dye is at the tip of the needle.</li>
	<li>Align the needle so that it is parallel with the embryo or angled slightly downward toward the larva.</li>
	<li>Move the needle to puncture the posterior end of the specimen and inject the specimen by pumping the foot pedal. Inject ~2 nL of dye into embryos, and ~220 nL of dye into larvae. 
	NOTE: The embryo or larva should flood with dye if the injection is successful.</li>
	<li>Note the time of injection for incubation purposes. Incubate embryos for 10 min at room temperature. Incubate larvae for 30 min at room temperature.</li>
	<li>Continue down the slide to inject additional specimens, noting the time of injection for each specimen. 
	NOTE: Depending on the speed with which subsequent dissection and imaging steps can be performed, 4&#8722;8 specimens can be injected at a time.</li>
</ol>
3. Preparation of Samples for Imaging
<ol>
	<li>Imaging of embryos
	<ol>
		<li>Following injection, prepare embryos for imaging. Apply petroleum jelly with a cotton-tipped applicator on the right and left sides of the samples on the slide as a spacer to prevent damage to the embryos upon placement of the coverslip.</li>
		<li>Image samples using a confocal microscope throughout the depth of the embryo with a 20x objective. Calculate the percentage of total samples with dye observed in the ventral nerve cord (VNC) using the following equation: % of samples with compromised BBB = Number of samples with dye accumulation in the VNC/total number of samples assayed.</li>
	</ol>
	</li>
	<li>Dissection and imaging of larvae
	<ol>
		<li>Prepare slides for larval samples ahead of time. Mount two coverslips spaced approximately 0.5 cm apart to the slide with nail polish. 
		NOTE: The coverslips function as spacers for the brain, so it is not damaged during the mounting process.</li>
		<li>Following the 30 min incubation, dissect the larvae in 1x PBS directly on the slide that will be used in imaging. First, use one pair of forceps to grab the larva halfway down the larval body, and use a second pair of forceps to separate the anterior and posterior halves of the larva.</li>
		<li>Next, use one pair of forceps to grip the anterior region at the mouth hooks, and use a second pair of forceps to invert the body wall over the tip of the forceps gripping the mouth hooks. The brain and VNC will be exposed.</li>
		<li>Separate the brain and VNC from the body wall by severing the nerves, and remove the body wall from the slide (Figure 1C,D). Remove imaginal discs if desired.</li>
		<li>Cover the sample with 10 &#956;L of 80% glycerol and place a coverslip on top of the sample for imaging.</li>
		<li>Image through the depth of the nervous system tissue using a 20x objective. Calculate the percentage of total samples with dye observed in the VNC.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

This protocol provides a comprehensive description of the steps needed to assay for BBB integrity during the late embryonic and third instar larval stages of D. melanogaster development. Similar approaches have been described elsewhere to assay the integrity of the BBB during development, as well as in adult stages5,7,29,30. However, descriptions of procedures in materials and methods ...

During development, cell-cell communication and interactions are critical for the establishment of tissue and organ structure and function. In some cases, these cell-cell interactions seal off organs from the surrounding environment to ensure proper organ function. This is the case for the nervous system, which is insulated by the blood-brain barrier (BBB). Dysfunction of the BBB in humans has been linked to neurological disorders including epilepsy, and breakdown of the barrier has been observed in neurodegenerative diseases including multiple sclerosis and amyotrophic lateral sclerosis1. In mammals, the BBB is formed by tight junctions between endothelial cells2,3. Other animals, including the fruit fly, Drosophila melanogaster, have a BBB composed of glial cells. These glial cells form a selectively permeable barrier to control movement of nutrients, waste products, toxins, and large molecules into and out of the nervous system4. This allows for the maintenance of the electrochemical gradient necessary to fire action potentials, allowing for mobility and coordination4. In D. melanogaster, the glia protect the nervous system from the potassium-rich, blood-like hemolymph5.
In the central nervous system (CNS) and peripheral nervous system (PNS) of D. melanogaster, two outer glial layers, the subperineurial glia and the perineurial glia, as well as an outer network of extracellular matrix, the neural lamella, form the hemolymph-brain and hemolymph-nerve barrier6, referred to as the BBB throughout this article. During development subperineurial glia become polyploid and enlarge to surround the nervous system5,6,7,8,9,10,11. The subperineurial glia form septate junctions, which provide the main diffusion barrier between the hemolymph and the nervous system5,6,12. These junctions are molecularly similar to the septate-like junctions found at the paranodes of myelinating glia in vertebrates, and they perform the same function as tight junctions in the BBB of mammals13,14,15,16,17. The perineurial glia divide, grow, and wrap around the subperineurial glia to regulate the diffusion of metabolites and large molecules6,10,18,19. BBB formation is complete by 18.5 h after egg laying (AEL) at 25 &#176;C5,8. Previous studies have identified genes that are critical regulators of BBB formation20,21,22. To better understand the exact roles of these genes, it is important to examine the effect of mutation of these potential regulators on BBB integrity. While previous studies have outlined approaches for assaying BBB integrity in embryos and larvae, a comprehensive protocol for this assay has yet to be described5,7. This step-by-step protocol describes methods for assaying BBB integrity during D. melanogaster embryonic and third instar larval stages.

<table><tbody><tr><td>10 kDa sulforhodamine 101 acid chloride (Texas Red) Dextran</td><td>ThermoFisher Scientific</td><td>D1863</td><td>Dextran should be diluted in autoclaved ddH&lt;sub&gt;2&lt;/sub&gt;O to a concentration of 25 mg/mL.</td></tr><tr><td>20 &amp;mu;L Gel-Loading Pipette Tips</td><td>Eppendorf</td><td>22351656</td><td></td></tr><tr><td>100% Ethanol (200 proof)</td><td>Pharmco-Aaper</td><td>11000200</td><td></td></tr><tr><td>Active Dry Yeast</td><td>Red Star</td><td></td><td></td></tr><tr><td>Agar</td><td>Fisher Scientific</td><td>BP1423</td><td></td></tr><tr><td>Agarose</td><td>Fisher Scientific</td><td>BP160-500</td><td></td></tr><tr><td>Air Compressor</td><td>DeWalt</td><td>D55140</td><td></td></tr><tr><td>Apple Juice</td><td>Mott&#039;s Natural Apple Juice</td><td></td><td></td></tr><tr><td>Bleach</td><td>Household Bleach</td><td></td><td>1-5% Hypochlorite</td></tr><tr><td>Borosilicate Glass Capillaries</td><td>World Precision Instruments</td><td>1B100F-4</td><td></td></tr><tr><td>Bottle Plugs</td><td>Fisher Scientific</td><td>AS-277</td><td></td></tr><tr><td>Cell Strainers</td><td>BD Falcon</td><td>352350</td><td></td></tr><tr><td>Confocal Microscope</td><td>Olympus</td><td>FV1000</td><td>Samples imaged using 20x objective (UPlanSApo 20x/ 0.75)</td></tr><tr><td>Cotton-Tipped Applicator</td><td>Puritan</td><td>19-062614</td><td></td></tr><tr><td>Double-Sided Tape 1/2&quot;</td><td>Scotch</td><td></td><td></td></tr><tr><td>Dumont Tweezers; Pattern #5; .05 x .01 mm Tip</td><td>Roboz</td><td>RS-5015</td><td></td></tr><tr><td>Fly Food Bottles</td><td>Fisher Scientific</td><td>AS-355</td><td></td></tr><tr><td>Fly Food Vials</td><td>Fisher Scientific</td><td>AS-515</td><td></td></tr><tr><td>Foot Pedal</td><td>Treadlite II</td><td>T-91-S</td><td></td></tr><tr><td>Gel Caster</td><td>Bio-Rad</td><td>1704422</td><td></td></tr><tr><td>Gel Tray</td><td>Bio-Rad</td><td>1704436</td><td></td></tr><tr><td>Glass Pipette</td><td>VWR</td><td>14673-010</td><td></td></tr><tr><td>Glycerol</td><td>Fisher Scientific</td><td>BP229-1</td><td></td></tr><tr><td>Granulated sugar</td><td></td><td></td><td>Purchased from grocery store.</td></tr><tr><td>Halocarbon Oil</td><td>Lab Scientific, Inc.</td><td>FLY-7000</td><td></td></tr><tr><td>Light Source</td><td>Schott</td><td>Ace I</td><td></td></tr><tr><td>Manipulator Stand</td><td>World Precision Instruments</td><td>M10</td><td></td></tr><tr><td>Micromanipulator</td><td>World Precision Instruments</td><td>KITE-R</td><td></td></tr><tr><td>Micropipette Puller</td><td>Sutter Instrument Co.</td><td>P-97</td><td></td></tr><tr><td>Needle Holder</td><td>World Precision Instruments</td><td>MPH310</td><td></td></tr><tr><td>Nightsea Filter Sets</td><td>Electron Microscopy Science</td><td>SFA-LFS-CY</td><td>For visualization of YFP</td></tr><tr><td>Nightsea Full Adapter System w/ Royal Blue Color Light Head</td><td>Electron Microscopy Science</td><td>SFA-RB</td><td>For visualization of GFP</td></tr><tr><td>Paintbrush</td><td>Simply Simmons</td><td>Chisel Blender #6</td><td></td></tr><tr><td>Pipetter</td><td>Fisher Scientific</td><td>13-683C</td><td></td></tr><tr><td>Pneumatic Pump</td><td>World Precision Instruments</td><td>PV830</td><td>This is also referred to as a microinjector or pressure regulator. Since the model used in our study is no longer available this is one alternative.</td></tr><tr><td>Potassium Chloride</td><td>Fisher Scientific</td><td>BP366-500</td><td></td></tr><tr><td>Potassium Phosphate Dibasic</td><td>Fisher Scientific</td><td>BP363-500</td><td></td></tr><tr><td>Small Embryo Collection Cages</td><td>Genesee Scientific</td><td>59-100</td><td></td></tr><tr><td>Sodium Chloride</td><td>Fisher Scientific</td><td>BP358-212</td><td></td></tr><tr><td>Sodium Phosphate Dibasic Anhydrous</td><td>Fisher Scientific</td><td>BP332-500</td><td></td></tr><tr><td>Steel Base Plate</td><td>World Precision Instruments</td><td>5052</td><td></td></tr><tr><td>Stereomicroscope</td><td>Carl Zeiss</td><td>Stemi 2000</td><td>Used for tissue dissection.</td></tr><tr><td>Stereomicroscope with transmitted light source</td><td>Baytronix</td><td></td><td>Used for injection.</td></tr><tr><td>Tegosept (p-hydroxybenzoic acid, methyl ester)</td><td>Genesee Scientific</td><td>20-258</td><td></td></tr><tr><td>Triton X-100</td><td>Fisher Scientific</td><td>BP151-500</td><td>Nonionic surfactant</td></tr><tr><td>Vial Plugs</td><td>Fisher Scientific</td><td>AS-273</td><td></td></tr></tbody></table>

assay for blood-brain barrier integrity in drosophila melanogaster

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous system and protects neural tissue from toxins and pathogens. Defects in the formation of this barrier have been correlated with neuropathies, and the breakdown of this barrier has been observed in many neurodegenerative diseases. Therefore, it is critical to identify the genes that regulate the formation and maintenance of the blood-brain barrier to identify potential therapeutic targets. In order to understand the exact roles these genes play in neural development, it is necessary to assay the effects of altered gene expression on the integrity of the blood-brain barrier. Many of the molecules that function in the establishment of the blood-brain barrier have been found to be conserved across eukaryotic species, including the fruit fly, Drosophila melanogaster. Fruit flies have proven to be an excellent model system for examining the molecular mechanisms regulating nervous system development and function. This protocol describes a step-by-step procedure to assay for blood-brain barrier integrity during the embryonic and larval stages of D. melanogaster development.

The methods described here allow for the visualization of the integrity of the BBB throughout the CNS in D. melanogaster embryos and larvae (Figure 1). Upon completion of BBB formation in late embryogenesis, the BBB functions to exclude large molecules from the brain and VNC5. This protocol takes advantage of this function to assay BBB formation. When wild-type (Oregon R) late stage 17 (20&#8722;21 h old) embryos were injected with 10 kDa dextran conjugated t...

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Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Blood-brain barrier integrity is critical for nervous system function. In Drosophila melanogaster, the blood-brain barrier is formed by glial cells during late embryogenesis. This protocol describes methods to assay for blood-brain barrier formation and maintenance in D. melanogaster embryos and third instar larvae.

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous ...

assay-for-blood-brain-barrier-integrity-in-drosophila-melanogaster

Research

JoVE Journal

Developmental Biology

8.2K Views.  Loyola University Chicago. Blood-brain barrier integrity is critical for nervous system function. In Drosophila melanogaster, the blood-brain barrier is formed by glial cells during late embryogenesis. This protocol describes methods to assay for blood-brain barrier formation and maintenance in D. melanogaster embryos and third instar larvae.

Video: Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

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 ...

Neuroscience

A Burrowing/Tunneling Assay for Detection of Hypoxia in Drosophila melanogaster Larvae

Oxygen deprivation in animals can result from exposure to low atmospheric oxygen levels or from internal tissue damage that interferes with oxygen distribution. It is also possible that aberrant behavior of oxygen-sensing neurons could induce hypoxia-like behavior in the presence of normal oxygen levels. In D. melanogaster, development at low oxygen levels results in inhibition of growth and sluggish behavior during the larval phases. However, these established manifestations of oxygen deficit overlap considerably with the phenotypes of many mutations that regulate growth, stress responses or locomotion. As result, there is currently no assay available to identify i) cellular hypoxia induced by a mutation or ii) hypoxia-like behavior when induced by abnormal neuronal behavior.
We have recently identified two distinctive behaviors in D. melanogaster larvae that occur at normal oxygen levels in response to internal detection of hypoxia. First, at all stages, such larvae avoid burrowing into food, often straying far away from a food source. Second, tunneling into a soft substratum, which normally occurs during the wandering third instar stage is completely abolished if larvae are hypoxic. The assay described here is designed to detect and quantitate these behaviors and thus to provide a way to detect hypoxia induced by internal damage rather than low external oxygen. Assay plates with an agar substratum and a central plug of yeast paste are used to support animals through larval life. The positions and state of the larvae are tracked daily as they proceed from first to third instar. The extent of tunneling into the agar substratum during wandering phase is quantitated after pupation using NIH ImageJ. The assay will be of value in determining when hypoxia is a component of a mutant phenotype and thus provide insight into possible sites of action of the gene in question.

A Burrowing/Tunneling Assay for Detection of Hypoxia in Drosophila melanogaster Larvae

The protocol describes a simple assay to identify Drosophila melanogaster larvae that are experiencing hypoxia under normal atmospheric oxygen levels. This protocol allows hypoxic larvae to be distinguished from other mutants that show overlapping phenotypes such as sluggishness or slow growth.

Oxygen deprivation in animals can result from exposure to low atmospheric oxygen levels or from internal tissue damage that interferes with oxygen ...

Behavior

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 ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Assessing Neurodegenerative Phenotypes in Drosophila Dopaminergic Neurons by Climbing Assays and Whole Brain Immunostaining

Drosophila melanogaster is a valuable model organism to study aging and pathological degenerative processes in the nervous system. The advantages of the fly as an experimental system include its genetic tractability, short life span and the possibility to observe and quantitatively analyze complex behaviors. The expression of disease-linked genes in specific neuronal populations of the Drosophila brain, can be used to model human neurodegenerative diseases such as Parkinson's and Alzheimer's 5.
Dopaminergic (DA) neurons are among the most vulnerable neuronal populations in the aging human brain. In Parkinson's disease (PD), the most common neurodegenerative movement disorder, the accelerated loss of DA neurons leads to a progressive and irreversible decline in locomotor function. In addition to age and exposure to environmental toxins, loss of DA neurons is exacerbated by specific mutations in the coding or promoter regions of several genes. The identification of such PD-associated alleles provides the experimental basis for the use of Drosophila as a model to study neurodegeneration of DA neurons in vivo. For example, the expression of the PD-linked human &alpha;-synuclein gene in Drosophila DA neurons recapitulates some features of the human disease, e.g. progressive loss of DA neurons and declining locomotor function 2. Accordingly, this model has been successfully used to identify potential therapeutic targets in PD 8.
Here we describe two assays that have commonly been used to study age-dependent neurodegeneration of DA neurons in Drosophila: a climbing assay based on the startle-induced negative geotaxis response and tyrosine hydroxylase immunostaining of whole adult brain mounts to monitor the number of DA neurons at different ages. In both cases, in vivo expression of UAS transgenes specifically in DA neurons can be achieved by using a tyrosine hydroxylase (TH) promoter-Gal4 driver line 3, 10.

Assessing Neurodegenerative Phenotypes in Drosophila Dopaminergic Neurons by Climbing Assays and Whole Brain Immunostaining

Here we describe two assays that have been established to study age-dependent neurodegeneration of dopaminergic (DA) neurons in Drosophila: the climbing/startle-induced negative geotaxis assay which allows to study the functional effects of DA neurons degeneration and the tyrosine hydroxylase immunostaining which is used to identify and count DA neurons in whole brain mounts.

Drosophila melanogaster is a valuable model organism to study aging and pathological degenerative processes in the nervous system. The advantages of the ...

An In Vivo Blood-brain Barrier Permeability Assay in Mice Using Fluorescently Labeled Tracers

Blood-brain barrier (BBB) is a specialized barrier that protects the brain microenvironment from toxins and pathogens in the circulation and maintains brain homeostasis. The principal sites of the barrier are endothelial cells of the brain capillaries whose barrier function results from tight intercellular junctions and efflux transporters expressed on the plasma membrane. This function is regulated by pericytes and astrocytes that together form the neurovascular unit (NVU). Several neurological diseases such as stroke, Alzheimer&#39;s disease (AD), brain tumors are associated with an impaired BBB function. Assessment of the BBB permeability is therefore crucial in evaluating the severity of the neurological disease and the success of the treatment strategies employed.
We present here a simple yet robust permeability assay that have been successfully applied to several mouse models both, genetic and experimental. The method is highly quantitative and objective in comparison to the tracer fluorescence analysis by microscopy that is commonly applied. In this method, mice are injected intraperitoneally with a mix of aqueous inert fluorescent tracers followed by anesthetizing the mice. Cardiac perfusion of the animals is performed prior to harvesting brain, kidneys or other organs. Organs are homogenized and centrifuged followed by fluorescence measurement from the supernatant. Blood drawn from the cardiac puncture just before perfusion serves for normalization purpose to the vascular compartment. The tissue fluorescence is normalized to the wet weight and serum fluorescence to obtain a quantitative tracer permeability index. For additional confirmation, the contralateral hemi-brain preserved for immunohistochemistry can be utilized for tracer fluorescence visualization purposes.

An In Vivo Blood-brain Barrier Permeability Assay in Mice Using Fluorescently Labeled Tracers

Here we present a mouse brain vascular permeability assay using intraperitoneal injection of fluorescent tracers followed by perfusion that is applicable to animal models of blood-brain barrier dysfunction. One hemi-brain is used for assessing permeability quantitatively and the other for tracer visualization/immunostaining. The procedure takes 5 - 6 h for 10 mice.

Blood-brain barrier (BBB) is a specialized barrier that protects the brain microenvironment from toxins and pathogens in the circulation and maintains ...

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs provides a tissue-level understanding of energy utilization that cannot be captured when analyzing primary cells and cell lines. While this analysis is ex vivo, it allows for the measurement from a number of specialized cells working together to perform a function in one tissue and more closely models the in vivo organ. Metabolic reprogramming has been observed in many neurological diseases, including neoplasia, and neurodegenerative diseases. This protocol was developed to assist the D. melanogaster community&#39;s investigation of metabolism in neurological disease models using a commercially available metabolic analyzer. Measuring metabolism of whole brains in the metabolic analyzer is challenging due to the geometry of the brain. This analyzer requires samples to remain at the bottom of a 96-well plate. Cell samples and tissue punches can adhere to the surface of the cell plate or utilize spheroid plates, respectively. However, the spherical, three-dimensional shape of D. melanogaster brains prevents the tissue from adhering to the plate. This protocol requires a specially designed and manufactured micro-tissue restraint that circumvents this problem by preventing any movement of the brain while still allowing metabolic measurements from the analyzer&#39;s two solid-state sensor probes. Oxygen consumption and extracellular acidification rates are reproducible and sensitive to a treatment with metabolic inhibitors. With a minor optimization, this protocol can be adapted for use with any whole tissue and/or model system, provided that the sample size does not exceed the chamber generated by the restraint. While basal metabolic measurements and an analysis after a treatment with mitochondrial inhibitors are described within this protocol, countless experimental conditions, such as energy source preference and rearing environment, could be interrogated.

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

We present a protocol for measuring oxygen consumption and extracellular acidification in Drosophila melanogaster larval and adult brains. A metabolic analyzer is utilized with an adapted and optimized protocol. Micro-tissue restraints are a critical component of this protocol and were designed and created specifically for their use in this analysis.

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs ...

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield deleterious consequences, yet the current methods for recording nervous system activity of an individual animal is costly and laborious. This suction electrode preparation of the third-instar larval central nervous system of Drosophila melanogaster, is a tractable system for testing the physiological effects of neuroactive agents, determining the physiological role of various neural pathways to CNS activity, as well as the influence of genetic mutations to neural function. This ex vivo preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of insect neuronal activity. A wide variety of chemical modulators, including peptides, can then be applied directly to the nervous system in solution with the physiological saline to measure the influence on the CNS activity. Further, genetic technologies, such as the GAL4/UAS system, can be applied independently or in tandem with pharmacological agents to determine the role of specific ion channels, transporters, or receptors to arthropod CNS function. In this context, the assays described herein are of significant interest to insecticide toxicologists, insect physiologists, and developmental biologists for which D. melanogaster is an established model organism. The goal of this protocol is to describe an electrophysiological method to enable the measurement of electrogenesis of the central nervous system in the model insect, Drosophila melanogaster, which is useful for testing a diversity of scientific hypotheses.

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield ...

Morphological and Functional Evaluation of Axons and their Synapses during Axon Death in Drosophila melanogaster

Axon degeneration is a shared feature in neurodegenerative disease and when nervous systems are challenged by mechanical or chemical forces. However, our understanding of the molecular mechanisms underlying axon degeneration remains limited. Injury-induced axon degeneration serves as a simple model to study how severed axons execute their own disassembly (axon death). Over recent years, an evolutionarily conserved axon death signaling cascade has been identified from flies to mammals, which is required for the separated axon to degenerate after injury. Conversely, attenuated axon death signaling results in morphological and functional preservation of severed axons and their synapses. Here, we present three simple and recently developed protocols that allow for the observation of axonal morphology, or axonal and synaptic function of severed axons that have been cut-off from the neuronal cell body, in the fruit fly Drosophila. Morphology can be observed in the wing, where a partial injury results in axon death side-by-side of uninjured control axons within the same nerve bundle. Alternatively, axonal morphology can also be observed in the brain, where the whole nerve bundle undergoes axon death triggered by antennal ablation. Functional preservation of severed axons and their synapses can be assessed by a simple optogenetic approach coupled with a post-synaptic grooming behavior. We present examples using a highwire loss-of-function mutation and by over-expressing dnmnat, both capable of delaying axon death for weeks to months. Importantly, these protocols can be used beyond injury; they facilitate the characterization of neuronal maintenance factors, axonal transport, and axonal mitochondria.

Morphological and Functional Evaluation of Axons and their Synapses during Axon Death in Drosophila melanogaster

Here, we provide protocols to perform three simple injury-induced axon degeneration (axon death) assays in Drosophila melanogaster to evaluate the morphological and functional preservation of severed axons and their synapses.

Axon degeneration is a shared feature in neurodegenerative disease and when nervous systems are challenged by mechanical or chemical forces. However, our ...

A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity

The Drosophila embryo has long been a powerful laboratory model for elucidating molecular and genetic mechanisms that control development. The ease of genetic manipulations with this model has supplanted pharmacological approaches that are commonplace in other animal models and cell-based assays. Here we describe recent advances in a protocol that enables application of small molecules to the developing fruit fly embryo. The method details steps to overcome the impermeability of the eggshell while maintaining embryo viability. Eggshell permeabilization across a broad range of developmental stages is achieved by application of a previously described d-limonene embryo permeabilization solvent (EPS1) and by aging embryos at reduced temperature (18 &#176;C) prior to treatments. In addition, use of a far-red dye (CY5) as a permeabilization indicator is described, which is compatible with downstream applications involving standard red and green fluorescent dyes in live and fixed preparations. This protocol is applicable to studies using bioactive compounds to probe developmental mechanisms as well as for studies aimed at evaluating teratogenic or pharmacologic activity of uncharacterized small molecules.

A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity

Introduction of small molecules to the developing Drosophila embryo offers great potential for characterizing biological activity of novel compounds, drugs, and toxins as well as for probing fundamental developmental pathways. Methods described herein outline steps that overcome natural barriers to this approach, expanding the utility of the Drosophila embryo model.

The Drosophila embryo has long been a powerful laboratory model for elucidating molecular and genetic mechanisms that control development. The ease of ...

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

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

A Burrowing/Tunneling Assay for Detection of Hypoxia in Drosophila melanogaster Larvae

Methods to Assay Drosophila Behavior

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Assessing Neurodegenerative Phenotypes in Drosophila Dopaminergic Neurons by Climbing Assays and Whole Brain Immunostaining

An In Vivo Blood-brain Barrier Permeability Assay in Mice Using Fluorescently Labeled Tracers

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

Morphological and Functional Evaluation of Axons and their Synapses during Axon Death in Drosophila melanogaster

A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity