Name: stimulating and analyzing adult neurogenesis in the drosophila central brain
Uploaded: 2021-10-08T14:30:00.000+00:00
Duration: 6 min 31 s
Description: Watch this Scientific Journal Video about Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain at JoVE.com

We are grateful to Stacey Rimkus and Becky Katzenberger for technical assistance and to Eduardo Moreno for sharing the perma-twin stocks. We would like to thank Barry Ganetzky and David Wassarman for lively discussions that undoubtedly improved science and Kent Mok, Cayla Guerra, and Bailey Spiegelberg for their contributions to the laboratory. The FasII antibodies were developed by Corey Goodman and obtained from the Developmental Studies Hybridoma Bank, created by the NICHD of the NIH and maintained at The University of Iowa, Department of Biology, Iowa City, IA 52242. Most of the Drosophila strains used in this study were obtained from the Bloomington Drosophila Stock Center (BDSC; NIH P40OD018537). This work was supported by NIH T32 GM007133 (KLC); NIH NS090190 (GBF); NIH NS102698 (GBF); the University of Wisconsin Graduate School (GBF); and the UW-Madison Women in Science and Engineering Leadership Institute (WISELI) (GBF).

This protocol follows the animal care guidelines of UW-Madison.
1. Generating adult Drosophila for PTBI
<ol>
	<li>For the standard cross, place 20 virgin y[1] w[1];UAS-mCD8-GFP;; OK107-GAL417&#160;adult females and 10 y[1] w[1]17 adult male flies together in vials (see Table of Materials) containing food. To have large numbers of synchronous offspring, set up 10-20 crosses at the same time. The standard cross gives rise to F1 progeny of the genotype: y[1] w[1];UAS-mCD8-GFP/+;; OK107-GAL4/+.
	<ol>
		<li>Place the vials at 25 &#176;C for mating and egg-laying. To maximize progeny, transfer parents into new vials every 2-4 days, maintaining the earlier vials at 25 &#176;C. Discard each vial 18 days after the parents were first placed in it to ensure that there is no F2 progeny. 
		NOTE: 3 sets of offspring (&#34;broods&#34;) can be generated from each set of parents.</li>
		<li>Check after ~10 days, when the F1 progeny will begin to eclose.</li>
	</ol>
	</li>
	<li>For lineage studies, use F1 perma-twin males15 of genotype: w; FRT40A, UAS-CD2-RFP, UAS-GFP-Mir/FRT40A, UAS-CD8-GFP, UAS-CD2-Mir; tub-GAL80ts/act-GAL4 UAS-flp. For consistency, do this cross in the same direction each time.
	<ol>
		<li>To generate perma-twin males, place 20 virgin w; FRT40A, UAS-CD2-RFP, UAS-GFP-Mir; tub-GAL80ts/TM6B females and 10 w; FRT40A, UAS-CD8-GFP, UAS-CD2-Mir; act-GAL4 UAS-flp/TM6B15 adult males together in vials containing food.</li>
		<li>Place the vials at 17 &#176;C for mating and egg-laying. Transfer parents into vials of new food every 7 days, maintaining all vials at 17 &#176;C. Discard each vial 35 days after the parents were first placed in it to ensure that there are no F2 progeny.</li>
		<li>Check after ~21 days, when the F1 progeny will begin to eclose.</li>
	</ol>
	</li>
</ol>
2. Penetrating traumatic brain injury (PTBI;&#160;&#160;Figure 1)
<ol>
	<li>Sort newly eclosed F1 flies. Select young males within 6 h post-eclosion. Place these males in clean vials containing food, with 40 or fewer flies per vial. 
	NOTE: This is most easily accomplished in mid-morning by anesthetizing flies on the CO2 pad and identifying adult males that still have meconium (visible as a dark greenish spot through the abdominal wall) in their guts.</li>
	<li>Pre-feed with 5-ethynyl-2&#39;-deoxyuridine (EdU) (see Table of Materials) for 6 h before the injury if planning to label dividing cells with EdU. See step 3 for details.</li>
	<li>Sanitize Minutien pins (see Table of Materials) for at least 5 min by placing ~100 pins in a 1.5 mL microcentrifuge tube filled with 70% ethanol.&#160;</li>
	<li>Sanitize the CO2 pad and a paintbrush by spraying 70% ethanol and wipe dry with a clean lint-free tissue. Once the tools are clean and dry, transfer 40 or fewer sorted F1 males back onto the clean pad.</li>
	<li>Separate the F1 males into 2 groups on the fly pad. One group will serve as a control, uninjured flies. The second experimental group will be subjected to PTBI.</li>
	<li>Using forceps, pull 4-5 new Minutien pins out of the microcentrifuge tube and place them near the edge of the CO2 pad. Under the dissecting scope, choose a straight Minutien pin with a sharp point. 
	NOTE: Using a single Minutien pin for an experiment will reduce variability. Reuse sharp pins. Place damaged or blunt pins in a separate 1.5 mL microcentrifuge tube containing 70% ethanol for safe disposal.</li>
	<li>If working with flies from the standard cross, switch on the stereo microscope light-emitting diode (LED) lamp equipped with the appropriate excitation and emission filters for green fluorescence protein (GFP). This permits excitation at 440-460 nM and allows visualization of 500-560 nM. 
	NOTE: With this lamp and filter set, the cell bodies of the mushroom body will fluoresce green through the head cuticle (Figure 1C). For perma-twin flies or flies of other genotypes, a standard white light illuminator for the stereo microscope can be used to visualize landmarks on the head cuticle for targeting the injury to the mushroom body (Figure 1C).</li>
	<li>Use the forceps to pick up and hold the selected Minutien pin in one hand (for right-handed people, this is usually the right hand) and the paintbrush in the other (usually left) hand. Choose a fly from the experimental group and position the fly such that you have a dorsal view of the head capsule with the fly&#39;s head to the right. Place the brush on the anterior of the dorsal thorax and push down gently to stabilize the fly.</li>
	<li>Aim the Minutien pin&#39;s tip at the mushroom body&#39;s cell bodies on the right side of the head and penetrate the head capsule. If using landmarks, target the dorsal head cuticle between the ocelli and the dorsal rim of the eye (Figure 1).</li>
	<li>After completing the injury, use the paintbrush to push the head gently off the Minutien pin.</li>
	<li>If using the brain for RNA-Seq or qRT-PCR, make a second injury on the left side of the head.</li>
	<li>Repeat steps 2.8-2.10 to injure all of the flies in the experimental group.</li>
	<li>Once all flies have been injured, place the control and injured flies into labeled, separate vials containing food. Lay the vials horizontally (i.e., on their sides) while the flies recover from the anesthesia and during subsequent aging to prevent flies from getting caught in the food.</li>
	<li>Place standard cross flies at 25 &#176;C, and perma-twin flies at 30 &#176;C to age.</li>
	<li>For flies aged longer than 24 h, place them on clean food every 1-2 days.</li>
</ol>
3. EdU labeling
<ol>
	<li>Prepare a stock of 10 mM 5-ethynyl-2&#39;-deoxyuridine (EdU) in dimethyl sulfoxide (DMSO). This can be stored at -20 &#176;C for up to 12 months.</li>
	<li>Prepare 200 &#181;L of 50&#160;&#181;M EdU in 10% sucrose. Pre-feed flies with EdU for 6 h before PTBI.</li>
	<li>Place 200 &#181;L of 50 &#181;M EdU in 10% sucrose on a 23 mm round grade 3 filter paper (see Table of Materials) in an otherwise empty vial.</li>
	<li>Place the flies in the vial. Then seal the vial with a cotton plug.</li>
	<li>Lay the vial horizontally in a humidified incubator at 25 &#176;C for 6 h.</li>
	<li>Carry out PTBI as described in steps 2.3-2.10.</li>
	<li>Place the flies back in the EdU-containing vial. Seal the vial with a cotton plug.</li>
	<li>Lay the vial horizontally in a humidified incubator at 25 &#176;C for up to 24 h. Follow one of the steps described below (step 3.8.1-3.8.3).
	<ol>
		<li>Dissect and fix brains as described in step 4 using fixative, wash buffer, and blocking buffer without azide and with the EdU detection reaction carried out before antibody staining.</li>
		<li>Transfer the flies to a clean vial containing a new filter paper and 200&#160;&#181;L of 50&#160;&#181;M EdU in 10% sucrose. Lay the vial horizontally in a 25 &#176;C incubator. Repeat every 24 h during the duration of the labeling. Then, dissect and fix brains as described in step 4 using buffers without azide with the EdU detection reaction carried out before antibody staining.</li>
		<li>To pulse-chase label with EdU, feed EdU for the pulse period, transfer the flies every 24 h to a clean vial containing a new filter paper and 200&#160;&#181;L of 50&#160;&#181;M in 10% sucrose. After the pulse period (e.g., 4 days), transfer the flies to a vial containing standard Drosophila food. Place the vial on its side in a 25 &#176;C incubator for an additional 3 days. Then, dissect and fix brains as described in step 4 using buffers without azide with the EdU detection reaction carried out before antibody staining.</li>
	</ol>
	</li>
</ol>
4. Dissection, immunohistochemistry, and mounting
<ol>
	<li>Prepare 1.5 mL microcentrifuge tubes with 100 &#181;L of fixative: 4% formaldehyde in PEM (100 mM piperazine-N,N&#39;-bis(2-ethanesulfonic acid) [PIPES], 1 mM EGTA, 1 mM MgSO4, pH 7.0) (see Table of Materials) and place on ice. 
	NOTE: As many as 20 brains of a single genotype and condition can be processed in a single tube.</li>
	<li>Prepare a dissection plate with one small pool (~100 &#181;L) of 70% ethanol and three small pools of phosphate-buffered saline (PBS; 100 mM of K2HPO4, 140 mM of NaCl, pH 7.0).</li>
	<li>Anesthetize ~10 control or experimental flies on a CO2 pad that has been sanitized with 70% ethanol.</li>
	<li>Separate the head from the trunk of each fly using a scalpel.</li>
	<li>Collect the heads with a paintbrush wetted in 70% ethanol and place heads for 2-5 min in the ethanol pool on the dissection plate. 
	NOTE: This slightly dehydrates the brains and makes them easier to dissect away from the head cuticle.</li>
	<li>Transfer the heads to a ~100 &#181;L pool of PBS and dissect out the brains, moving each brain to a clean ~100 &#181;L pool of PBS. Perform this using two pairs of Watchmakers forceps to open the back of the head cuticle and holding the cuticle with one pair of forceps while gently prying the brain out of the cuticle using the closed tip of the second pair of forceps.</li>
	<li>Transfer the dissected brains to a microcentrifuge tube containing 100 &#181;L of the fixing solution using a P200 pipettor equipped with a plastic tip that has been cut and beveled to permit entry of the brains.</li>
	<li>Fix for 20-25 min at room temperature. Carefully remove the fix with either a P200 or a glass pipette.</li>
	<li>Wash the fixed brains four times with 1 mL of &#39;PT&#39; (PBS plus 0.1% Triton X-100), allowing the brains to settle for several minutes between each wash. 
	NOTE: If brains do not settle rapidly between the washes, they are likely to have still fat body and/or trachea attached.</li>
	<li>Block samples in 1 mL of PBS plus 0.1% Triton X-100 and 2% bovine serum albumin (PBT) for ~1 h at room temperature.</li>
	<li>Remove the blocking solution and incubate samples with 100 &#181;L of primary antibody solution overnight at 4 &#176;C in PBT. The primary antibodies used in this study are rabbit anti-PH3 (1:500) and mouse anti-Fasiclin II (1:20) (see Table of Materials).</li>
	<li>Wash samples five times with 1 mL of PT, allowing the brains to settle for several minutes between each wash.</li>
	<li>Remove the final wash and incubate in 100 &#181;L of secondary antibody solution overnight at 4 &#176;C. The secondary antibodies used here are anti-rabbit 568 (1:400) and anti-mouse Cy5 (1:100) (see Table of Materials).</li>
	<li>Wash samples five times with 1 mL of PT, allowing the brains to settle for several minutes between each wash. During the final wash, add 4&#39;,6-Diamidino-2-Phenylindole, Dihydrochloride (DAPI; 1:100 of a 10 &#181;M solution) for 10 min to stain the nuclei. Remove the wash, leaving 50-100 &#181;L in each tube.</li>
	<li>Prepare microscope slides by placing a single, self-adhesive reinforcement label on the middle of each slide and a 50 &#181;L droplet of anti-fade mounting media in the center of each label (see Table of Materials). The reinforcement label holds the coverslip slightly above the slides and prevents the mounted brains from becoming overly flattened.</li>
	<li>Carefully transfer the brains from each microcentrifuge tube to a prepared slide with as little wash buffer as possible using a P200 equipped with a cut and beveled plastic tip. As many as 10 brains can be mounted on a single slide.</li>
	<li>Use forceps to gently reposition the brains before placing a coverslip on each slide and sealing the slide with nail polish. Orient the brains with either posterior side up or anterior side up but should not be touching one another. 
	NOTE: Because it is difficult to obtain high-quality confocal images through an entire brain, some brains should be mounted in each orientation.</li>
	<li>Store prepared slides flat and in the dark at 4 &#176;C until imaging. For longer-term storage (up to 1 year), slides may be stored at -20 &#176;C.</li>
</ol>
5. Confocal imaging
NOTE: Image brains using a laser-scanning confocal microscope with excitation lasers and emission filter cubes appropriate to DAPI and the fluorescent secondary antibodies (i.e., 405 nm, 488 nm, and 568 nm, 633 nm, respectively).
<ol>
	<li>Turn on the power of the microscope, lasers, controller, and computer.</li>
	<li>Open the Acquisition software.</li>
	<li>In the Acquisition mode, select up to four channels and set the parameter for sequential scanning of the desired channels. Increase the laser power for each channel to 5-10%.</li>
	<li>Place a slide on the microscope.</li>
	<li>Choose a brain to be imaged using the epifluorescence attachment (see Table of Materials).</li>
	<li>In the Acquisition mode, select 1024 x 1024 pixels as the frame dimension.</li>
	<li>In the Acquisition mode, select the Z-stack option, indicate Z-stack should be taken at 2 &#181;m sections, and focus through the sample, selecting the top and bottom focal planes to be imaged.</li>
	<li>Collect Z-stack images of entire brains using a 20x objective and specific brain regions such as the mushroom body using a 60x objective.</li>
</ol>
6. Data analysis
<ol>
	<li>Quantify the proliferating cells and the numbers of perma-twin clones manually and/or using the image analysis software (see Table of Materials). When using software, select regions of interest (ROIs) with areas of at least 10 &#181;m.</li>
</ol>

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The authors have no conflicts of interest.

Although penetrating injuries to the adult Drosophila brain have been described previously15,17,18, these injuries focused on the optic lobes and not the central brain. Further, detailed instructions for how to carry out the injuries are thus far lacking. This protocol describes a model for penetrating injury to the adult Drosophila central brain that reproduces statistically significant evidence for adult neurogenesis after PTBI.
The reproducibility of this PTBI protocol is due, in part, to the mushroom body as the injury target region. The mushroom body is large, consisting of ~2200 neurons with complex dendrite and axon arbors in large and highly stereotyped arrays18. The cell bodies of mushroom body neurons lie near the brain&#39;s surface and can be visualized through the head cuticle using the expression of green fluorescent protein (GFP) (Figure 1C). Mushroom body precursors are the last neural stem cells to undergo apoptosis during development13,12,19. Thus, many mushroom body neurons are pretty young at the time of eclosion. This led to the hypothesis that the mushroom body might have more mitotic potential than other brain regions16. In addition, the mushroom body is critical for learning and memory18. This allows one to ask whether PTBI-triggered neurogenesis leads to functional recovery.
Other factors that contribute to the reproducibility of the results include using outcrossed flies of consistent genotypes, performing crosses in the same direction each time, precisely controlling the rearing and aging temperatures, and analyzing males and females separately. Using F1 flies from an outcross reduces the probability of analyzing brains homozygous for spontaneous mutations. The standard cross of y[1] w[1];UAS-mCD8-GFP;; OK107-GAL4 adult&#160;females to y[1] w[1] adult male flies&#160;results in F1 progeny of the genotype y[1] w[1];UAS-mCD8-GFP/+;; OK107-GAL4/+. OK107-GAL4 is expressed in all intrinsic neurons of the mushroom body and drives expression of the membrane-bound reporter UAS-mCD8-GFP permitting visualization of mushroom bodies and their projections. For the perma-twin crosses, crosses must remain at 17 &#176;C at all times to keep the lineage tracing system switched off. This ensures that no dividing cells are labeled during development and that only adult-born neurons and glia are labeled. To this end, the fly room can also be maintained at 17 &#176;C. Although the initial description of the perma-twin system15 recommended rearing flies at 18 &#176;C, this can lead to significant background labeling.
For consistency, it also is recommended to keep the control uninjured flies on the CO2 pad as one carries out the PTBI. This ensures that both sets of flies have identical anesthetic exposure. In addition, it is desirable for reproducibility to completely penetrate the head. However, care must be taken not to bend the tip of the pin against the pad, making it unusable for future injuries. For skilled practitioners, there is little unintended harm to PTBI flies. Nonetheless, pressing too hard on the thorax to stabilize the fly during injury can be lethal. One way to assess the extent of the unintended injury is to quantify the mortality of PTBI flies 24 h post-injury. For unilaterally injured flies, this can be 50% or higher for beginners. Therefore, to ensure that observed outcomes are due to PTBI and not to unintended injury, it is advised that beginners practice administering PTBI on ~20 flies daily over several weeks and do not analyze the resulting brains until the 24 h mortality is consistently &#60;10%.
To quantify the amount of proliferation stimulated by central brain PTBI, both anti-phosphohistone H3 (PH3) immunostaining and 5-ethynyl-2&#180;-deoxyuridine (EdU) incorporation can be employed. Anti-PH3 labels cells before and throughout metaphase, limiting detection to only a fraction of the actively dividing cells. Thus, anti-PH3 staining provides only a partial glimpse of proliferation. EdU is a thymidine analog that can be incorporated into newly synthesized DNA. By feeding flies EdU before and after injury, it is possible to gain a more complete picture of the cells that are either dividing or have divided following the injury. The fact that any cells that divide are permanently marked is helpful both for the identification of slowly cycling cells and assay the survival of cells after initial proliferation. For unclear reasons, but maybe due to limited permeability of the blood-brain barrier, EdU labeling is inefficient and under-reports cell proliferation in the adult brain. This is evidenced by the similar numbers of PH3+ and EdU+ cells in both control and experimental brains at 24 h post-PTBI and by observing that only a subset of new cells in perma-twin clones incorporate EdU16. For maximal labeling, it is essential to pre-feed the flies with EdU because injured flies do not resume feeding for several hours post-PTBI. Feeding was assessed by adding food coloring to the EdU solution and monitoring the amount of dye in the gut through the abdominal cuticle16.
It is to be noted that while we have provided a brain dissection protocol in step 4, alternative techniques may be used. Several of these are available in previously published protocols20,21,22. Drosophila melanogaster offers a low-cost model with powerful genetic and molecular tools that can be used to study the mechanisms underlying regeneration of multiple tissues, including the gut and components of the nervous system. A novel and reproducible injury model that can be used to study the response to brain injury is outlined here. Data obtained using these protocols support the idea that the adult Drosophila central brain retains the proliferative ability, generating new neurons in response to injury. These observations warrant further investigation of both the extent of adult neurogenesis and its underlying molecular mechanisms. Once the components involved in neural regeneration are identified in this system, we can convert our knowledge of adult Drosophila neurogenesis to humans.

Damage to the brain and nervous system is a major cause of death and disability worldwide. Approximately 1.5 million Americans suffer traumatic brain injuries (TBI) every year1, while an estimated 6 million individuals in the United States alone suffer from neurodegenerative diseases, such as Parkinson&#39;s and Alzheimer&#39;s Disease2. Both disease and injury to the brain can cause neural degeneration, leading to sensory, cognitive, and motor defects3. Developing therapeutic strategies for human brain repair has been difficult due to the complex physiology of the brain. Model organisms such as Drosophila melanogaster provide a simple system for identifying the fundamental mechanisms underlying neurodegeneration and potential therapeutic targets4.
The fruit fly Drosophila melanogaster has been a powerful model organism for more than a century, advancing the fields of genetics, developmental biology, and neuroscience5,6. The Drosophila brain comprises only ~90,000 neurons7, a million-fold fewer than the average human brain8, yet they have many similarities. Both human and fly brains utilize the neurotransmitters GABA, glutamate, acetylcholine, and the biogenic amines dopamine and serotonin9. Drosophila and human neurons also function similarly, with a shared synaptic architecture and analogous neural cell types10. The smaller brain size of Drosophila and the availability of advanced genetics techniques, in combination with the conservation of molecular, cellular, and physiological mechanisms between Drosophila and mammals, permits Drosophila researchers to ask questions that are impractical or difficult to answer in mammalian models.
Our current understanding of adult neurogenesis in Drosophila, both during homeostasis and following injury, remains limited. More is known about neurogenesis during normal development. For example, neurons and glia are created during development from precursor cells, called neuroblasts10,11. At least three different types of neuroblasts have been distinguished in the central brain. Both Type I and Type II lineage neuroblasts exit the cell cycle ~20-30 h after puparium formation12. In contrast, the mushroom body neuroblasts are the last to terminate cell division and do so via Reaper-dependent apoptosis ~85-90 h after puparium formation13. Following eclosion, the adult Drosophila brain has few dividing cells (~1 cell/brain), predominantly glia14. The adult optic lobes possess slowly cycling neuroblasts capable of neurogenesis15, while the adult central brain has no known neuroblasts. The scarcity of neural progenitors and limited cell proliferation strongly resembles the situation in the adult mammalian brain, underscoring the potential relevance of the mechanisms of adult neurogenesis in Drosophila to humans.
The discovery of low levels of adult neurogenesis in the adult Drosophila optic lobes after injury15 led to the hypothesis that the adult Drosophila central brain also might be capable of adult neurogenesis16. This protocol describes creating a rigorous, reproducible model of central brain injury in adult Drosophila that can be used to investigate neurogenesis in the adult central brain. Given the similarities between human and Drosophila brain architecture and function, these discoveries could lead to the identification of critical targets for therapeutic neurogenesis in injured and diseased human brains.

<table><tbody><tr><td>#11 disposable scalpels</td><td>Santa Cruz Biotechnology</td><td>sc-395923</td><td>used for separating Drosophila heads from trunks prior to brain dissection</td></tr><tr><td>150 mm diameter black Sylgard dishes</td><td>Dow</td><td>1696157</td><td>made in the laboratory with reagents from Dow; used for brain dissection</td></tr><tr><td>18 mm coverslips</td><td>any</td><td></td><td>for mounting brains on microscope slides</td></tr><tr><td>4&#039;,6-Diamidino-2-Phenylindole, Dihydrochloride (DAPI)</td><td>ThermoFisher</td><td>D1306</td><td>for immunohistochemistry</td></tr><tr><td>70% Ethanol</td><td>made from 95% ethanol sourced variously</td><td></td><td></td></tr><tr><td>anti-mouse Cy5</td><td>Jackson ImmunoResearch</td><td>715-175-151</td><td>for immunohistochemistry</td></tr><tr><td>anti-rabbit 568</td><td>ThermoFisher</td><td>A11036</td><td>for immunohistochemistry</td></tr><tr><td>bovine serum albumin (BSA)</td><td>SIgma Aldrich</td><td>A7030</td><td>for immunohistochemisty</td></tr><tr><td>Clear nail polish</td><td>any</td><td></td><td>for sealing coverslips</td></tr><tr><td>Click-It EdU labeling kit</td><td>InVitrogen</td><td>C10640</td><td>to detect newly synthesized DNA</td></tr><tr><td>CO&lt;sub&gt;2&lt;/sub&gt; bubbler</td><td>Genesee Scientific</td><td>59-181</td><td>for anesthesia</td></tr><tr><td>CO&lt;sub&gt;2&lt;/sub&gt; pad</td><td>Genesee Scientific</td><td>59-114</td><td>for anesthesia</td></tr><tr><td>CO&lt;sub&gt;2&lt;/sub&gt; regulator and supply</td><td>any</td><td></td><td>for anesthesia</td></tr><tr><td>Confocal microscope</td><td>any</td><td></td><td>for imaging fixed, stained and mounted brains</td></tr><tr><td>cotton plugs</td><td>Genesee Scientific</td><td>51-101</td><td>for EdU labeling</td></tr><tr><td>Drosophila vials</td><td>Genesee Scientific</td><td>32-109</td><td>for EdU labeling</td></tr><tr><td>Fix buffer (Pipes, EGTA, Magnesium; PEM)</td><td>components sourced from various companies</td><td></td><td>for fixing adult brains; 100 mM piperazine-N,N&amp;rsquo;-bis(2-ethanesulfonic acid) [PIPES], 1 mM EGTA, 1 mM MgSO4, pH 7.0</td></tr><tr><td>Formaldehyde</td><td>Sigma Aldrich</td><td>252549</td><td>for fixing adult brains, added to PEM</td></tr><tr><td>Grade 3 round Whatman filters, 23 mm round</td><td>Tisch Scientific</td><td>1003-323</td><td>for EdU labeling</td></tr><tr><td>Microfuge tubes</td><td>any</td><td></td><td>for fixing and staining reactions and for storing Minutien pins</td></tr><tr><td>Microscope slides</td><td>any</td><td></td><td>for mounting brains</td></tr><tr><td>Minutien pins</td><td>Fine Science Tools</td><td>26002-10</td><td>for brain injury; 12.5 &amp;mu;m diameter tip and 100 &amp;mu;m diameter rod</td></tr><tr><td>mouse anti-Fasiclin II</td><td>Developmental Studies Hybridoma Bank</td><td>1D4-s</td><td>for immunohistochemistry</td></tr><tr><td>NIGHTSEA stereo microscope fluorescence adaptor</td><td>Electron Microscopy Sciences</td><td>SFA-GR</td><td>fluorescence setup for dissecting microscope</td></tr><tr><td>P20, P200 and P1000 pipettors and tips</td><td>any</td><td></td><td>for measuring solutions</td></tr><tr><td>phosphate buffered saliine (PBS)</td><td>components sourced from various companies</td><td></td><td>for dissecting brains and making immunohistochemistry blocking and washing solutions; 100 mM of K2HPO4, 140 mM of NaCl, pH 7.0</td></tr><tr><td>phosphate buffered saline with 0.1% Triton X-100 (PT)</td><td>components sourced from various companies</td><td></td><td>for washing dissected brains</td></tr><tr><td>phosphate buffered saline with 0.1% Triton X-100 + 2% bovine serum albumin (PBT)</td><td>components sourced from various companies</td><td></td><td>blocking solution for immunohistochemistry and for diluting antibodies</td></tr><tr><td>rabbit anti-PH3</td><td>Santa Cruz Biotechnology, Inc</td><td>sc-8656-R</td><td>for immunohistochemistry</td></tr><tr><td>Reinforcement labels</td><td>Avery</td><td>5721</td><td>to maintain space between the microscope slide and the coverslip</td></tr><tr><td>Size 0 paintbrushes</td><td>any</td><td></td><td>to manipulate and stabilize adult Drosophila during injury</td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>93443</td><td></td></tr><tr><td>Two pair of #5 watchmakers forceps</td><td>Fine Science Tools</td><td>11255-20</td><td>used to hold the Minutien pins and for brain dissections</td></tr><tr><td>Vectashield</td><td>Vector Laboratories</td><td>H-1000</td><td>mounting medium for microscope slides</td></tr></tbody></table>

stimulating and analyzing adult neurogenesis in the drosophila central brain

The molecular and cellular mechanisms underlying neurogenesis in response to disease or injury are not well understood. However, understanding these mechanisms is crucial for developing neural regenerative therapies. Drosophila melanogaster is a leading model for studies of neural development but historically has not been exploited to investigate adult brain regeneration. This is primarily because the adult brain exhibits very low mitotic activity. Nonetheless, penetrating traumatic brain injury (PTBI) to the adult Drosophila central brain triggers the generation of new neurons and new glia. The powerful genetic tools available in Drosophila combined with the simple but rigorous injury protocol described here now make adult Drosophila brain a robust model for neural regeneration research. Provided here are detailed instructions for (1) penetrating injuries to the adult central brain and (2) dissection, immunohistochemistry, and imaging post-injury. These protocols yield highly reproducible results and will facilitate additional studies to dissect mechanisms underlying neural regeneration.

PTBI stimulates cell proliferation 
To determine the extent of neurogenesis after a central brain PTBI, the proliferative response was measured in young adult males collected and injured within 6 h of eclosion. A significant increase in proliferation was observed 24 h post-injury using anti-phosphohistone 3 (PH3), a marker for cells actively undergoing mitosis. Approximately 3 PH3+ cells in control central brains and 11 PH3+ cells in the injured central brains are observed 24 h post-PTBI (Figure 2A-D). The majority of the dividing cells are located near the injury site. A second assay for cell division was used to quantify the cumulative cell proliferation from a single injury and to assess the extent to which the newly created cells survived. 5-ethynyl-2&#39;-deoxyuridine (EdU) is a thymidine analog that can be incorporated into newly synthesized DNA and permanently label cells that have undergone DNA synthesis. Flies were given a 4-day pulse of EdU, followed by a 3-day chase. This revealed that the labeled cells were viable and survived at least 3 days after proliferation. By 7 days, there were an average of 2 EdU+ cells in control central brains and an average of 11 EdU+ cells in the injured central brains, respectively (Figure 2E). This is similar to the results obtained 24 h post-injury using the PH3 antibody. When cell proliferation is measured at 14 days, the uninjured controls averaged 1 EdU+ cell per central brain, while the injured brains averaged 29 EdU+ cells (Figure 2E), demonstrating that cell proliferation continues at least into the second week following a PTBI.
Cell proliferation is age-dependent 
The greatest proliferative response in the central brain was observed within the first 24 h after eclosion (Figure 3). By 7 days post-eclosion, a penetrating injury still causes a significant increase in proliferation, with an average of 6 PH3+cells per central brain. Still, by 14 days post-eclosion, the ability for cells to divide following PTBI decreases significantly to 1 dividing cell, similar to that of control brains (Figure 3). Thus, the potential for cell proliferation post-PTBI is age-dependent.
Newly created neurons can project to correct target areas 
To evaluate neural regeneration post-PTBI, the perma-twin labeling system15 was used. Perma-twin lineage tracing permanently labels dividing cells and their progeny with a green fluorescent protein (GFP) or red fluorescent protein (RFP)15. More perma-twin clones were detected in injured samples, at 2 days and 2 weeks, than in controls (Figure 4A-E). Notably, there were new mushroom body neurons in ~50% of the PTBI brains 2 weeks post-injury (Figure 4N). These new neurons projected their dendrites appropriately to the mushroom body calyx and their axons appropriately to the mushroom body lobes (Figure 4D,F,G). This indicates that the newly created cells may be functional neurons involved in the repair of the damaged mushroom bodies. Other areas of the brain that appeared to regenerate include the ellipsoid body (EB) (Figure 4H,I), the antennal lobes (AL) (Figure 4J,K), and the lateral horn (LH) (Figure 4L,M) which possessed large clones approximately 26%, 26%, and 20% of the time, respectively (Figure 4N). These results underscore the utility of this system for the investigation of adult neurogenesis. A proposed model for the sequence of events following PTBI and leading to the generation of new neurons is shown in Figure 5.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63182/63182fig01.jpg" /> 
Figure 1: Penetrating Traumatic Brain Injury (PTBI) to the adult Drosophila central brain. (A)&#160;Schematic of the exterior of an adult fly head. This is a frontal view. Thus, the right side of the animal is to the viewer&#39;s left. (B)&#160;Schematic of the interior of an adult Drosophila head with the injury trajectory indicated in grey. This is a posterior view. Thus, in this image and subsequent figures, the right side of the brain is to the right. Central brain PTBI impacts multiple brain structures, including the mushroom body (green) and tissues outside the brain, including the fat body (blue) and hemocytes (red). CB = central brain region. OL= optic lobe region. (C)&#160;Dorsal view of a live adult head in which mushroom bodies (arrowheads) are labeled with a green fluorescent protein (GFP). This is the &#39;standard genotype&#39; (see text for details). The PTBI protocol reproducibly results in injury to the mushroom bodies. This Figure has been adapted from Reference16. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63182/63182fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63182/63182fig02.jpg" /> 
Figure 2: PTBI stimulates cell proliferation. Uninjured control (A) and PTBI (B) schematics. The blue boxes in the upper right corners indicate the brain regions shown at higher magnification in panels (C) and (D). (C,D) PH3 antibody (red) was used to assay cell proliferation 24 h after injury. In control brains (C), there are few PH3+ cells and none near the MB. However, in PTBI brains (D), there are PH3+ cells near the MB.(E)&#160;Quantification of proliferating cells. The numbers reflect proliferating cells throughout entire brains, not only in the vicinity of the mushroom body. At 24 h, uninjured control brains had an average of 3 PH3+ cells/brain (n = 11 brains, 28 cells), while 24 h post-PTBI, brains had an average of 11 PH3+ cells/brain (n = 17 brains, 181 cells). At 7 days, uninjured controls have few EdU+ cells, with an average of 2 EdU+ cells/brain (n = 15 brains, 24 cells), while 7-day post-PTBI brains had an average of 11 EdU+ cells/brain (n = 22 brains, 238 cells). At 14 days, uninjured controls have an average of 1 EdU+ cell/brain (n = 8 brains,11 cells), while 14-day post-PTBI brains have an average of 29 EdU+ cells/brain (n = 14 brains, 400 cells). For this set of experiments, young adult males within 6 h of eclosion were used. Unpaired t-tests of control and PTBI samples at the 3-time points yield values of p&#60;0.0001, p&#60;0.0001, and p&#60;0.0002, respectively. Error bars reflect the standard deviation (SD). This Figure has been adapted from Reference16. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63182/63182fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63182/63182fig03.jpg" /> 
Figure 3: The proliferative response to PTBI decreases with age. To explore whether age impacts the amount of cell proliferation that occurs post-injury, newly eclosed adult males were compared to animals aged to 7 days, 14 days, and 28 days before PTBI, using anti-PH3 to assay cell proliferation 24 h after injury. Flies injured within 6 h of eclosion had an average of 11 PH3+ cells/brain (n = 17 brains, 182 cells) compared to an average of 3 PH3+ cells/brain in age-matched controls (n = 11 brains, 28 cells). Flies aged to 7 days, then subjected to PTBI, had an average of 6 PH3+ cells/brain (n = 11 brains, 65 cells) compared to age-matched controls with an average of 2 PH3+ cells/brain (n = 5 brains, 12 cells). When flies were aged to 14 days before PTBI and assayed 24 h later, there was an average of 1 PH3+ cell/brain (n = 8 brains, 11 cells) similar to age-matched controls, which also averaged 1 PH3+ cell/brain (n = 4 brains, 2 cells). 28-day uninjured control (n = 4, 1 cell) and PTBI (n = 3, 1 cell) flies both averaged 0 PH3+cells/brain. Unpaired t-tests for PTBI to control comparisons at these 4-time points are p&#60;0.0001, p&#60;0.04, p&#60;0.07, and p&#60;0.84, respectively. This Figure has been adapted from Reference16. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63182/63182fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63182/63182fig04.jpg" /> 
Figure 4: Perma-twin lineage tracing demonstrates brain regeneration and appropriate targeting of axons following PTBI. The perma-twin lineage-tracing system15 was utilized to analyze neurogenesis after PTBI. This system permanently labels dividing cells and progeny with a green fluorescent protein (GFP) or red fluorescent protein (RFP). Flies were reared at 17 &#176;C to keep the system off during development. F1 males carrying perma-twin transgenes were collected upon eclosion, then injured and placed at 30 &#176;C to recover for 2 or 14 days. (A) In 2-day uninjured controls, there are some GFP+ cells scattered throughout the brain.&#160;(B)&#160;At 14 days, there are relatively few GFP+ cells present in the control central brain. (C)&#160;In comparison, 2-day injured brains have more GFP+ cells that tend to cluster near the injury (arrowhead). (D) At 14 days post-injury, there are large clones near the site of injury. Some of these clones have axons that project along the mushroom body tracts (arrowhead). Only the GFP channel is shown here; there were similar RFP+ clones in the PTBI samples. (E)&#160;The number of clones increases over time post-PTBI.Control uninjured brains (n = 13) have an average of 10 clones at 2 days, while 2-day PTBI brains (n = 20) have an average of 23 clones (p&#60;0.00002). At 7 days, control brains had an average of 9 clones per brain (n = 18), while 7-day PTBI brains had an average of 39 clones per brain (n = 16) (p-value&#60;0.00000002). This is significantly more than the number of clones seen at 2 days post-injury (p-value&#60;0.0009). In 14-day control brains, there is an average of 10 clones per brain, which is not significantly different from the 2-day and 7-day controls. However, at 14 days post-PTBI, there is an average of 66 GFP+ clones, which is significantly more than either age-matched controls (p&#60;0.0000003) or 2-day post-PTBI brains (p-value&#60;0.0001). Error bars reflect SD. (F-M) PTBI stimulates clone formation in multiple regions in the brain.&#160;Panels on the left side are schematics of brain regions where large clones were found 14 days post-PTBI (A, H, J, L). Panels on the right show high magnifications of representative brains (G, I, K, M). Many 14-day brains had clones that projected to particular target areas. These included the mushroom body (MB) (F,G), the ellipsoid body (EB) (H,I), the antennal lobe (AL) (J,K), and the lateral horn (LH) (L,M).&#160;(N)&#160;Both clone number and clone size increase with time post-PTBI.The proportions of brain regions with large clones were calculated at 2, 7, and 14 days in controls and injured brains. At 2 days, ~8% of control brains (n = 13) showed AL clones, while there were no AL clones in 2-day injured brains (n = 20). In 7-day control brains (n = 18), 6% had AL and 6% had EB clones. At 7 days post-PTBI (n = 16), 6% of brains also had AL clones, 6% had EB clones, and 19% had large MB clones. At 14 days, control brains (n = 9) did not exhibit any specific areas with clones, while 47% of PTBI brains (n = 15) had MB clones, 20% of PTBI brains had AL clones, and 27% of PTBI brains had EB clones, and 27% had LH clones. This Figure has been adapted from Reference16. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63182/63182fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63182/63182fig05.jpg" /> 
Figure 5:&#160;Summary model for regeneration following penetrating traumatic brain injury (PTBI). In young adult Drosophila, there are quiescent neuroblast-like cells within the central brain that lack expression of canonical neuroblast genes. By 24 h post-PTBI, the quiescent neuroblast-like cells are activated, express neuroblast genes, and have begun to proliferate. At both 4 h and 24 h post-PTBI, there is a wave of cell death16. At 7 days, the proliferation rate is still high, and many of the new cells have adopted mature cell identities, becoming neurons or glia. At 14 days post-PTBI, large clones of new neurons with axons and dendrites correctly projecting to their respective target areas. Locomotor defects are also restored by 14 days, suggesting that adult Drosophila can regenerate functionally and structurally. This Figure has been adapted from Reference16. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63182/63182fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain at JoVE.com

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

This article provides detailed protocols for inflicting Penetrating Traumatic Brain Injury (PTBI) to adult Drosophila and examining the resulting neurogenesis.

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

The molecular and cellular mechanisms underlying neurogenesis in response to disease or injury are not well understood. However, understanding these ...

stimulating-analyzing-adult-neurogenesis-drosophila-central

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

2.7K Views.  University of Wisconsin-Madison, School of Medicine and Public Health. This article provides detailed protocols for inflicting Penetrating Traumatic Brain Injury (PTBI) to adult Drosophila and examining the resulting neurogenesis.

Video: Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the removal of brains from animals at 70-78 hrs after puparium formation. The isolated brains are shown after brief incubation in papain followed by several washes in serum-free growth medium. The process of mechanical dissociation of each brain in a 5 ul drop of media on a coverslip is illustrated. The axons and dendrites of the post-mitotic neurons are sheered off near the soma during dissociation but the neurons begin to regenerate processes within a few hours of plating. Images show live cultures at 2 days. Neurons continue to elaborate processes during the first week in culture. Specific neuronal populations can be identified in culture using GAL4 lines to drive tissue specific expression of fluorescent markers such as  GFP or RFP. Whole cell recordings have demonstrated the cultured neurons form functional, spontaneously active cholinergic and GABAergic synapses. A short video segment illustrates calcium dynamics in the cultured neurons using Fura-2 as a calcium indicator dye to monitor spontaneous calcium transients and nicotine evoked calcium responses in a dish of cultured neurons. These pupal brain cultures are a useful model system in which genetic and pharmacological tools can be used to identify intrinsic and extrinsic factors that influence formation and function of central synapses.

This video demonstrates the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. Views of live cultures show neurite outgrowth and imaging of calcium levels using Fura-2.

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the ...

Biology

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 ...

Developmental Biology

Ex vivo Culturing of Whole, Developing Drosophila Brains

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for investigating the cell biology and development of central brain structures by allowing acute pharmacological interventions and live imaging of cellular processes. As an example of the technique, prior work from our lab1 has shown that a previously unrecognized subcellular compartment lies between the axonal and somatodendritic compartments of axons of the Drosophila central brain. The development of this compartment, referred to as the axon initial segment (AIS)2, was shown genetically to depend on the neuron-specific cyclin-dependent kinase, Cdk5. We show here that ex vivo treatment of wild-type Drosophila larval brains with the Cdk5-specific pharmacological inhibitors roscovitine and olomoucine3 causes acute changes in actin organization, and in localization of the cell-surface protein Fasciclin 2, that mimic the changes seen in mutants that lack Cdk5 activity genetically.
A second example of the ex vivo culture technique is provided for remodeling of the connections of embryonic mushroom body (MB) gamma neurons during metamorphosis from larva to adult. The mushroom body is the center of olfactory learning and memory in the fly4, and these gamma neurons prune their axonal and dendritic branches during pupal development and then re-extend branches at a later timepoint to establish the adult innervation pattern5. Pruning of these neurons of the MB has been shown to occur via local degeneration of neurite branches6, by a mechanism that is triggered by ecdysone, a steroid hormone, acting at the ecdysone receptor B17, and that is dependent on the activity of the ubiquitin-proteasome system6. Our method of ex vivo culturing can be used to interrogate further the mechanism of developmental remodeling. We found that in the ex vivo culture setting, gamma neurons of the MB recapitulated the process of developmental pruning with a time course similar to that in vivo. It was essential, however, to wait until 1.5 hours after puparium formation before explanting the tissue in order for the cells to commit irreversibly to metamorphosis; dissection of animals at the onset of pupariation led to little or no metamorphosis in culture. Thus, with appropriate modification, the ex vivo culture approach can be applied to study dynamic as well as steady state aspects of central brain biology.

Ex vivo Culturing of Whole, Developing Drosophila Brains

This article describes a method by which one can mimic in vivo development of the Drosophila mushroom body in an ex vivo culture system.

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for ...

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. Understanding repair and regeneration in animals is a key question in biology. The damaged human CNS does not regenerate, and understanding how to promote the regeneration is one of main goals of medical neuroscience. The powerful genetic toolkit of Drosophila can be used to tackle the problem of CNS regeneration.
A lesion to the CNS ventral nerve cord (VNC, equivalent to the vertebrate spinal cord) is applied manually with a tungsten needle. The VNC can subsequently be filmed in time-lapse using laser scanning confocal microscopy for up to 24 hr to follow the development of the lesion over time. Alternatively, it can be cultured, then fixed and stained using immunofluorescence to visualize neuron and glial cells with confocal microscopy. Using appropriate markers, changes in cell morphology and cell state as a result of injury can be visualized. With ImageJ and purposely developed plug-ins, quantitative and statistical analyses can be carried out to measure changes in wound size over time and the effects of injury in cell proliferation and cell death. These methods allow the analysis of large sample sizes. They can be combined with the powerful genetics of Drosophila to investigate the molecular mechanisms underlying CNS regeneration and repair.

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. ...

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.

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.

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

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 ...

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various intrinsic and extrinsic inhibitory factors involved in the restriction of axon regeneration have been identified. However, simply removing these inhibitory cues is insufficient for successful regeneration, indicating the existence of additional regulatory machinery. Drosophila melanogaster, the fruit fly, shares evolutionarily conserved genes and signaling pathways with vertebrates, including humans. Combining the powerful genetic toolbox of flies with two-photon laser axotomy/dendriotomy, we describe here the Drosophila sensory neuron &#8211; dendritic arborization (da) neuron injury model as a platform for systematically screening for novel regeneration regulators. Briefly, this paradigm includes a) the preparation of larvae, b) lesion induction to dendrite(s) or axon(s) using a two-photon laser, c) live confocal imaging post-injury and d) data analysis. Our model enables highly reproducible injury of single labeled neurons, axons, and dendrites of well-defined neuronal subtypes, in both the peripheral and central nervous system.

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

Here, we present a protocol using the Drosophila sensory neuron - dendritic arborization (da) neuron injury model, which combines in vivo live imaging, two-photon laser axotomy/dendriotomy, and the powerful fly genetic toolbox, as a platform for screening potential promoters and inhibitors of neuroregeneration.

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various ...

Medicine

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are optimized to minimize their impact on behavior. This is first achieved by using a holder that minimizes movement hindrances. The back of the fly&#8217;s head is glued to this holder at an angle that allows optical access to the whole brain while retaining the fly&#8217;s ability to walk, groom, smell, taste and see. The back of the head is dissected to remove tissues in the optical path and muscles responsible for head movement artefacts. The fly brain can subsequently be imaged to record brain activity, for instance using calcium or voltage indicators, during specific behaviors such as walking or grooming, and in response to different stimuli. Once the challenging dissection, which requires considerable practice, has been mastered, this technique allows to record rich data sets relating whole brain activity to behavior and stimulus responses.

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method specifically tailored to image the whole brain of adult Drosophila during behavior and in response to stimuli. The head is positioned to allow optical access to the whole brain, while the fly can move its legs and the antennae, the tip of the proboscis, and the eyes can receive sensory stimuli.

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are ...

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

Neural stem cells (NSCs) have the ability to proliferate, differentiate, undergo apoptosis, and even enter and exit quiescence. Many of these processes are controlled by the complex interplay between NSC intrinsic genetic programs with NSC extrinsic factors, local and systemic. In the genetic model organism, Drosophila melanogaster, NSCs, known as neuroblasts (NBs), switch from quiescence to proliferation during the embryonic to larval transition. During this time, larvae emerge from their eggshells and begin crawling, seeking out dietary nutrients. In response to animal feeding, the fat body, an endocrine organ with lipid storage capacity, produces a signal, which is released systemically into the circulating hemolymph. In response to the fat body-derived signal (FBDS), Drosophila insulin-like peptides (Dilps) are produced and released from brain neurosecretory neurons and glia, leading to downstream activation of PI3-kinase growth signaling in NBs and their glial and tracheal niche. Although this is the current model for how NBs switch from quiescence to proliferation, the nature of the FBDS extrinsic cue remains elusive. To better understand how NB extrinsic systemic cues regulate exit from quiescence, a method was developed to culture early larval brains in vitro before animal feeding. With this method, exogenous factors can be supplied to the culture media and NB exit from quiescence assayed. We found that exogenous insulin is sufficient to reactivate NBs from quiescence in whole-brain explants. Because this method is well-suited for large-scale screens, we aim to identify additional extrinsic cues that regulate NB quiescence versus proliferation decisions. Because the genes and pathways that regulate NSC proliferation decisions are evolutionarily conserved, results from this assay could provide insight into improving regenerative therapies in the clinic.

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

A method to reactivate quiescent neural stem cells in cultured Drosophila brain explants has been established. Using this method, the role of systemic signals can be uncoupled from tissue-intrinsic signals in the regulation of neural stem cell quiescence, entry and exit.

Neural stem cells (NSCs) have the ability to proliferate, differentiate, undergo apoptosis, and even enter and exit quiescence. Many of these processes ...

Analyzing Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

Source: Naomi Keliinui, C., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/63189">Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants.</a> J. Vis. Exp. (2022).
This video demonstrates a protocol to identify exogenous factors that initiate the reactivation of Drosophila brain neural stem cells from their quiescent state. Freshly hatched larvae are dissected to isolate their brains, which are then cultured in media with or without a test hormone to evaluate the hormone&#39;s potential to induce neural stem cell reactivation and proliferation.

1. Culture media and tool preparation

	Spray the bench and work area with 70% ethanol and let dry.
	Spray the dissection tools, forceps, and two glass ...

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

Summary

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Chapters in this video

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

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

Ex vivo Culturing of Whole, Developing Drosophila Brains

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

Mass Histology to Quantify Neurodegeneration in Drosophila

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

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

Analyzing Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants