We thank RIKEN for funding. P10-Gal4 was a kind gift of Alain Vincent (Universit&eacute; Paul Sabatier, Toulouse, France).

1. Preparation of reagents
Notes before beginning: Dissection and immunohistochemical staining are carried out in a magnetic chamber and the larva is pinned down using specially shaped insect pins. Detailed instructions on the construction of a magnetic chamber, and preparation of these pins can be found in related references 14,15. In brief, a 1x1cm square hole is cut into a magnetic sheet and a coverslip affixed to the rear of the sheet to make a small chamber. The sides of the chamber are sealed with epoxy glue; after this glue has set the chamber is washed several times with 70% ethanol before use. Dissection insect pins are prepared by bending to the required shape and then glued onto a metal tab14,15. Alternatively to a metal tab, we have used an inverted flat-head steel drawing pin with a handle made from a cut-off yellow tip. Use of this magnetic chamber arrangement allows close control over pin positioning and tissue stretching during dissection.
To drive reporter gene expression in different subsets of DA neurons investigators may use several different Gal4 lines (summarized by Shimono and colleagues16). Many of these lines are available from public stock centers. In this representative protocol, we carry out immunostaining of a line in which two contrasting classes of DA neuron are co-labeled: the most simple- class I and most complex- class IV (P10-Gal417,18, UAS-mCD8::Kusabira-Orange (KO)).
<ol>
 <li>Prepare Ca++-free HL3.1 saline
 <ol>
 <li>In mM: 70 NaCl, 5 KCl, 20 MgCl2 , 10 NaHCO3 , 5 HEPES, 115 sucrose, and 5 trehalose; pH 7.219. Filter-sterilize and store at 4&deg;C. Note: Ca++-free solution prevents muscle contraction during dissection.</li>
 </ol>
 </li>
 <li>Prepare 2x PHEM buffer
 <ol>
 <li>In mM: 130 PIPES, 60 HEPES, 20 EGTA, 4 MgSO4; pH 7.0. Filter-sterilize and store at 4&deg;C.
 Note: These materials will not dissolve until the pH approaches 7.0.</li>
 </ol>
 </li>
 <li>Prepare the fixative freshly immediately before the fixation.
 <ol>
 <li>In order to prepare a 25ml solution, first mix 2g paraformaldehyde, 100&mu;l 1M NaOH and 10ml water in a 50ml Falcon tube.</li>
 <li>Shake fixative in a 55&deg;C water bath with shaker until the solution is clear. (Total volume will be about 11.5 ml.)</li>
 <li>Cool fixative on ice.</li>
 <li>Add 12.5ml 2x PHEM buffer.</li>
 <li>Adjust pH to 7.0 with 1M HCl.</li>
 <li>Fill the solution with water until the volume is 25ml.</li>
 <li>Filter the solution using Whatman paper.</li>
 </ol>
 </li>
</ol>
2. Larval dissection
Note before beginning: Microtubule networks, and particularly those in sensory dendrites, will breakdown rapidly after the initiation of dissection. Achieving fast dissection in less than five minutes followed by immediate fixation are key factors in the success of this protocol.
<ol>
 <li>Wash the larva in water and quickly move them into this drop using a loop of hair.</li>
 <li>Orient the larva dorsal side up and the ventral side on the glass. Note: This orientation is to examine the ventral cluster neurons. To examine dorsal cluster neurons, invert the orientation.</li>
 <li>Use the center insect-pin to pin the anterior end near the mouth hooks. Place the pin close to the end for best results.</li>
 <li>Place a drop of HL3.1 saline in the dissection chamber.</li>
 <li>Cut the very posterior tip of the larva off with microscissors. Note: This step opens an aperture at the posterior end of the larva that will allow access for the microscissors (step 2.7).</li>
 <li>Grab with the forceps the region of gut that is now poking out of the aperture at the posterior end of the larva. Gently pull out the whole gut.</li>
 <li>Place the tip of one blade of the microscissors through the aperture and cut along the ventral midline towards the anterior.</li>
 <li>Using the corner pins, pin the now free corners, posterior first, then anterior. Simultaneously gently stretch open the larval fillet.</li>
</ol>
3. Fixation, blocking, staining, and mounting of larval fillets
Notes before beginning: All fixation and staining steps are carried out in the dissection chamber. During this process, be careful not to knock the insect pins holding the larva as this may lead to tissue damage. To prevent the experiment from drying out, do all staining steps in a small Tupperware container surrounded by moistened tissues.
<ol>
 <li>Aspirate the Ca++-free HL3.1 buffer using a yellow tip. Immediately add the fixative using another pipetteman.</li>
 <li>Gently pipette up and down to mix the fixative with the remaining traces of Ca++-free HL3.1 buffer in the chamber. Immediately aspirate it and then add fresh fixative into the chamber.</li>
 <li>Incubate at room temperature for 20 minutes.</li>
 <li>Wash 6x 10mins in PBST (0.1% Triton X-100 in PBS).</li>
 <li>Block with 5% goat serum in PBST for 20 mins at RT.</li>
 <li>Replace the blocking solution with primary antibodies diluted in 5% goat serum in PBST. The primary antibodies used are mouse anti-&alpha;-Tubulin (DM1A) and rat anti-CD8 (5H10) both diluted 1/1000. 
 Note: The investigator may wish to substitute mouse anti-&alpha;-Tubulin (DM1A) with mouse anti-Futsch (22C10)20,21 diluted 1/1000 in some circumstances (see discussion).</li>
 <li>Incubate overnight (for at least 16h) at 4&deg;C.</li>
 <li>Wash 6x 10mins in PBST.</li>
 <li>Add secondary antibody solution diluted in 5% goat serum in PBST. The secondary antibodies used are Alexa Fluor 488 goat anti-mouse IgG and Cy3 donkey anti-rat IgG. Keep the sample covered to prevent fluorophore photo-bleaching22.</li>
 <li>Incubate either at RT for two hours, or overnight at 4&deg;C followed by one hour at room temperature.</li>
 <li>Wash 6x 10mins in PBST.</li>
 <li>Place the larval fillet on the slide cuticle-side down, mount in 80% glycerol, and seal the sides of the coverslip with nail polish for a 'quick' mount. 
 Note: For improved tissue clearing, and a permanent stable sample, mount in DPX as previously described23.</li>
</ol>
4. Representative Results:
Fluorescent staining was examined under a confocal microscope. In Figures 1-2, different branches within a dendrite arbor have different cytoskeletal organization. Figure 1 shows a region of the arbor of a class IV DA neuron at the 1st instar larval stage. The whole arbor is marked with mCD8::KO and detected using an anti-CD8 antibody and fluorescent secondary (Cy3). Tubulin is detected using anti &alpha;-Tubulin antibody and fluorescent secondary (Alexa Fluor 488). The main branches are positive for Tubulin, some thin side branches are Tubulin-negative. Movie 1 is a set of serial sections through a similar staining of a class I DA neuron. Figure 2 shows a region of the arbor of a class IV DA neuron stained with antibodies against Futsch and CD8 at the 1st instar larval stage. The main branches are Futsch-positive, some thin side branches are Futsch-negative. Figure 3. Tracheae in the larval body wall show a complex microtubule organization. Movies 2 and 3 show serial sections of staining through body wall muscles and trachea.
<img alt="Figure 1" src="/files/ftp_upload/3662/3662fig1.jpg" /> 
Figure 1. Fig. 1 shows a region of the arbor of a class IV DA neuron at the 1st instar larval stage. The whole arbor is marked with mCD8::KO and detected using anti-CD8 antibody and fluorescent secondary (Cy3). Tubulin is detected using anti-&amp;-Tubulin antibody and fluorescent secondary (Alexa 488). Panels A-C sequential confocal z-sections (0.5&mu;m), C'-C", single antibody staining from panel C. Example of branches with (yellow arrowhead) or without (purple arrowhead) microtubules are highlighted. Red arrowheads highlight microtubules in the underlying epithelial cells.
<img alt="Figure 2" src="/files/ftp_upload/3662/3662fig2.jpg" /> 
Figure 2. Fig. 2 shows a similar region of the arbor of a class IV da neuron stained with antibodies against Futsch and CD8 at the 1st instar larval stage. The whole arbor is marked with mCD8::KO and detected using anti-CD8 antibody and fluorescent secondary (Cy3). Futsch is detected using anti-Futsch antibody and fluorescent secondary (Alexa 488). The main branches are Futsch-positive, some thin side branches are Futsch-negative. Example of branches with (yellow arrowhead) or without (purple arrowhead) Futsch are highlighted.
<img alt="Figure 3" src="/files/ftp_upload/3662/3662fig3.jpg" /> 
Figure 3. Trachea stained using the anti-Tubulin protocol described above. This larva is third instar and dissected as previously described23,24. Movie 3 shows enlarged serial sections from the field marked by a square.
Movie 1. Serial sections tracing Tubulin staining throughout the dendritic arbor of a class I neuron. The whole arbor is marked with mCD8::KO (Magenta) and detected using anti-CD8 antibody and fluorescent secondary (Cy3). Tubulin (Green) is detected using an anti-&alpha;-Tubulin antibody and fluorescent secondary (Alexa Fluor 488). Scale: one side of the video image corresponds to 46.88&mu;m in the section. <a href="https://www.jove.com/files/ftp_upload/3662/YalginMooreMovie1.avi">Click here to view the Movie.</a>
Movie 2. Serial sections tracing Tubulin staining in body wall muscles of a third instar larva. Tubulin is detected using anti-&alpha;-Tubulin antibody and fluorescent secondary (Alexa Fluor 488). Scale: one side of the video image corresponds to 46.88&mu;m in the section. <a href="https://www.jove.com/files/ftp_upload/3662/YalginMooreMovie2.avi">Click here to view the Movie.</a>
Movie 3. Serial sections tracing Tubulin staining in a body wall trachea of a third instar larva, marked with a square in Figure.3. Tubulin is detected using anti-&alpha;-Tubulin antibody and fluorescent secondary (Alexa Fluor 488). Scale: one side of the video image corresponds to 46.88&mu;m in the section. <a href="https://www.jove.com/files/ftp_upload/3662/YalginMooreMovie3.avi">Click here to view the Movie.</a>

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We have nothing to disclose.

To understand how complex cell shapes are achieved it is important to be able to accurately assay microtubule organization. Here we describe a robust immunohistological labeling method to assay microtubule organization of dendritic arborization neuron sensory dendrites. In addition to staining sensory neurons, this method achieves robust immunohistological staining of trachea, muscles and other body wall tissues.
We use this protocol to examine microtubule organization in the developing sensory dendrites of DA neurons. These dendrites are very sensitive to injury or stress25, and break down rapidly after the beginning of dissection23. Dissection should be carried out in five minutes or less. Microtubules in the trachea and other body wall tissues are more stable. This protocol is adapted from those used prepare larval fillets for analysis of the neuromuscular junction14,15, and is optimized for quick fixation after a rapid dissection.
When analyzing the dendrites of DA neurons, robust anti-Tubulin staining in overlying muscle, and underlying epithelial cells26 can lead to a complicated staining pattern (Figure 1, Movie1). Hence careful tracing of dendritic staining through several serial sections (Movie1) may be required for a full analysis of microtubule organization in the DA neuron dendrite arbor. Monoclonal antibodies to the specific Drosophila homologue of MAP1B, Futsch20,21, can also be used to label the microtubule cytoskeleton in the Drosophila peripheral nervous system5,9,20. Futsch is expressed only in neurons and hence it produces a less complicated staining pattern than &alpha;-Tubulin (Figure 2). GFP-tagged-endogenous-Tau has also been used as a marker of the DA neuron dendritic microtubules27. Use of these markers is effective, but it should be noted that in the axon-termini of the neuromuscular junction Futsch specifically labels bundled microtubules28. The exact spatial relationships between Futsch, Tau, and microtubules in DA dendrites are yet to be fully characterized. In this protocol we have addressed endogenous microtubule organization. It is also possible to label microtubules using overexpression of GFP tagged Tau or Tubulin. Care should be taken against inducing unanticipated phenotypes when overexpression approaches are used29.
In our hands, immunohistochemical staining of microtubules in DA neurons and other body wall tissues is best achieved when the secondary antibodies used to detect the anti-Tubulin primary antibody is coupled to an easily-visualized fluorophore such as Alexa Fluor 488. Hence when using transgenic tools to label DA neurons in concert with anti-tubulin labeling e.g. via the Gal4/UAS system, LexA system, or in MARCM23 and MARCM derivatives, we prefer marker proteins without spectral overlap with this secondary, such as mCD8::Cherry and mCD8::KO.

<table border="1">
	<tbody>
		<tr>
			<th>
				Name of the reagent</th>
			<th>
				Company</th>
			<th>
				Catalogue 
				number</th>
			<th>
				Comments 
				(optional)</th>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Dumont</td>
			<td>11251-20</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Microscissors</td>
			<td>FST</td>
			<td>15000-08</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Mouse anti-&alpha;-tubulin (Clone: DM1A)</td>
			<td>Sigma</td>
			<td>T9026</td>
			<td>Dilution 1/1000</td>
		</tr>
		<tr>
			<td>Mouse anti-Futsch (Clone: 22C10), 
			supernatant</td>
			<td>Developmental 
			Studies 
			Hybridoma Bank</td>
			<td>22C10</td>
			<td>Dilution 1/1000</td>
		</tr>
		<tr>
			<td>Rat anti-CD8 (Clone: 5H10)</td>
			<td>Caltag</td>
			<td>MCD0800</td>
			<td>Dilution 1/1000</td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 anti-mouse IgG</td>
			<td>Invitrogen</td>
			<td>A-11001</td>
			<td>Dilution 1/500</td>
		</tr>
		<tr>
			<td>Cy3 anti-Rat IgG</td>
			<td>Jackson Immunoresearch</td>
			<td>712-166-150</td>
			<td>Dilution 1/200</td>
		</tr>
	</tbody>
</table>

immunohistological labeling of microtubules in sensory neuron dendrites, tracheae, and muscles in the drosophila larva body wall

To understand how differences in complex cell shapes are achieved, it is important to accurately follow microtubule organization. The Drosophila larval body wall contains several cell types that are models to study cell and tissue morphogenesis. For example tracheae are used to examine tube morphogenesis1, and the dendritic arborization (DA) sensory neurons of the Drosophila larva have become a primary system for the elucidation of general and neuron-class-specific mechanisms of dendritic differentiation2-5 and degeneration6.
The shape of dendrite branches can vary significantly between neuron classes, and even among different branches of a single neuron7,8. Genetic studies in DA neurons suggest that differential cytoskeletal organization can underlie morphological differences in dendritic branch shape4,9-11. We provide a robust immunological labeling method to assay in vivo microtubule organization in DA sensory neuron dendrite arbor (Figures 1, 2, Movie 1). This protocol illustrates the dissection and immunostaining of first instar larva, a stage when active sensory neuron dendrite outgrowth and branching organization is occurring 12,13.
In addition to staining sensory neurons, this method achieves robust labeling of microtubule organization in muscles (Movies 2, 3), trachea (Figure 3, Movie 3), and other body wall tissues. It is valuable for investigators wishing to analyze microtubule organization in situ in the body wall when investigating mechanisms that control tissue and cell shape.

Watch this Scientific Journal Video about Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall at JoVE.com

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

To understand how complex cell shapes, such as neuronal dendrites, are achieved during development, it is important to be able to accurately assay microtubule organization. Here we describe a robust immunohistological labeling method to examine microtubule organization of dendritic arborization neuron sensory dendrites, trachea, muscle, and other Drosophila larva body wall tissues.

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

To understand how differences in complex cell shapes are achieved, it is important to accurately follow microtubule organization. The Drosophila larval ...

immunohistological-labeling-microtubules-sensory-neuron-dendrites

Nanyang Technological University

Research

JoVE Journal

Neuroscience

12.3K Views.  RIKEN Brain Science Institute. To understand how complex cell shapes, such as neuronal dendrites, are achieved during development, it is important to be able to accurately assay microtubule organization. Here we describe a robust immunohistological labeling method to examine microtubule organization of dendritic arborization neuron sensory dendrites, trachea, muscle, and other Drosophila larva body wall tissues.

Video: Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic development and axonal wiring. Recently the dendritic arborization (DA) sensory neurons of the Drosophila larval peripheral nervous system (PNS) have become powerful genetic models in which to elucidate both general and class-specific mechanisms of neuron differentiation.
There are four main DA neuron classes (I-IV)1. They are named in order of increasing dendrite arbor complexity, and have class-specific differences in the genetic control of their differentiation2-10. The DA sensory system is a practical model to investigate the molecular mechanisms behind the control of dendritic morphology11-13 because: 1) it can take advantage of the powerful genetic tools available in the fruit fly, 2) the DA neuron dendrite arbor spreads out in only 2 dimensions beneath an optically clear larval cuticle making it easy to visualize with high resolution in vivo, 3) the class-specific diversity in dendritic morphology facilitates a comparative analysis to find key elements controlling the formation of simple vs. highly branched dendritic trees, and 4) dendritic arbor stereotypical shapes of different DA neurons facilitate morphometric statistical analyses.
DA neuron activity modifies the output of a larval locomotion central pattern generator14-16. The different DA neuron classes have distinct sensory modalities, and their activation elicits different behavioral responses14,16-20. Furthermore different classes send axonal projections stereotypically into the Drosophila larval central nervous system in the ventral nerve cord (VNC)21. These projections terminate with topographic representations of both DA neuron sensory modality and the position in the body wall of the dendritic field7,22,23. Hence examination of DA axonal projections can be used to elucidate mechanisms underlying topographic mapping7,22,23, as well as the wiring of a simple circuit modulating larval locomotion14-17.
We present here a practical guide to generate and analyze genetic mosaics24 marking DA neurons via MARCM (Mosaic Analysis with a Repressible Cell Marker)1,10,25 and Flp-out22,26,27 techniques (summarized in Fig. 1).

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

The dendritic arborization sensory neurons of the Drosophila larval peripheral nervous system are useful models to elucidate both general and neuron class-specific mechanisms of neuron differentiation. We present a practical guide to generate and analyze dendritic arborization neuron genetic mosaics.

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

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

Visualization of Proprioceptors in Drosophila Larvae and Pupae

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

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

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this information to the brain. As such, determining how olfactory information is transferred to the brain, modifying both perception and behavior, merits investigation. Interestingly, there is emerging evidence that cellular transduction and transcriptional factors are involved in the diversification of olfactory receptor neuron. Here we provide a robust whole mount immunological labeling method to assay in vivo olfactory receptor neuron organization. Using this method, we identified all olfactory receptor neurons with anti-ELAV antibody, a known pan-neural marker and Or49a-mCD8::GFP, an olfactory receptor neuron specifically expressed in Nba neuron using anti-GFP antibody.

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Herein we describe the process of whole mount immunostaining of Drosophila antennae, which enables us to better understand the molecular mechanisms involved in the diversification of olfactory receptor neurons (ORN)s.

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this ...

Olfactory Behaviors Assayed by Computer Tracking Of Drosophila in a Four-quadrant Olfactometer

A key challenge in neurobiology is to understand how neural circuits function to guide appropriate animal behaviors. Drosophila melanogaster is an excellent model system for such investigations due to its complex behaviors, powerful genetic techniques, and compact nervous system. Laboratory behavioral assays have long been used with Drosophila to simulate properties of the natural environment and study the neural mechanisms underlying the corresponding behaviors (e.g. phototaxis, chemotaxis, sensory learning and memory)1-3. With the recent availability of large collections of transgenic Drosophila lines that label specific neural subsets, behavioral assays have taken on a prominent role to link neurons with behaviors4-11. Versatile and reproducible paradigms, together with the underlying computational routines for data analysis, are indispensable for rapid tests of candidate fly lines with various genotypes. Particularly useful are setups that are flexible in the number of animals tested, duration of experiments and nature of presented stimuli. The assay of choice should also generate reproducible data that is easy to acquire and analyze. Here, we present a detailed description of a system and protocol for assaying behavioral responses of Drosophila flies in a large four-field arena. The setup is used here to assay responses of flies to a single olfactory stimulus; however, the same setup may be modified to test multiple olfactory, visual or optogenetic stimuli, or a combination of these. The olfactometer setup records the activity of fly populations responding to odors, and computational analytical methods are applied to quantify fly behaviors. The collected data are analyzed to get a quick read-out of an experimental run, which is essential for efficient data collection and the optimization of experimental conditions.

Olfactory Behaviors Assayed by Computer Tracking Of Drosophila in a Four-quadrant Olfactometer

We describe here a behavioral setup and data analysis method for assaying olfactory responses of up to 100 vinegar flies (Drosophila melanogaster). This system may be used with single or multiple olfactory stimuli, and adaptable for optogenetic activation or silencing of neuronal subsets.

A key challenge in neurobiology is to understand how neural circuits function to guide appropriate animal behaviors. Drosophila melanogaster is an ...

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

Labeling and Imaging of Amyloid Plaques in Brain Tissue Using the Natural Polyphenol Curcumin

Deposition of amyloid beta protein (A&#946;) in extra- and intracellular spaces is one of the hallmark pathologies of Alzheimer&#39;s disease (AD). Therefore, detection of the presence of A&#946; in AD brain tissue is a valuable tool for developing new treatments to prevent the progression of AD. Several classical amyloid binding dyes, fluorochrome, imaging probes, and A&#946;-specific antibodies have been used to detect A&#946; histochemically in AD brain tissue. Use of these compounds for A&#946; detection is costly and time consuming. However, because of its intense fluorescent activity, high-affinity, and specificity for A&#946;, as well as structural similarities with traditional amyloid binding dyes, curcumin (Cur) is a promising candidate for labeling and imaging of A&#946; plaques in postmortem brain tissue. It is a natural polyphenol from the herb Curcuma longa. In the present study, Cur was used to histochemically label A&#946; plaques from both a genetic mouse model of 5x familial Alzheimer&#39;s disease (5xFAD) and from human AD tissue within a minute. The labeling capability of Cur was compared to conventional amyloid binding dyes, such as thioflavin-S (Thio-S), Congo red (CR), and Fluoro-jade C (FJC), as well as A&#946;-specific antibodies (6E10 and A11). We observed that Cur is the most inexpensive and quickest way to label and image A&#946; plaques when compared to these conventional dyes and is comparable to A&#946;-specific antibodies. In addition, Cur binds with most A&#946; species, such as oligomers and fibrils. Therefore, Cur could be used as the most cost-effective, simple, and quick fluorochrome detection agent for A&#946; plaques.

Curcumin is an ideal fluorophore for labeling and imaging of amyloid beta protein plaques in brain tissue due to its preferential binding to amyloid beta protein as well as its structural similarities with other traditional amyloid binding dyes. It can be used to label and image amyloid beta protein plaques more efficiently and inexpensively than traditional methods.

Deposition of amyloid beta protein (A&#946;) in extra- and intracellular spaces is one of the hallmark pathologies of Alzheimer&#39;s disease (AD). ...

Ballistic Labeling of Pyramidal Neurons in Brain Slices and in Primary Cell Culture

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of pyramidal neurons and dendritic spines, a ballistic labeling technique can be utilized. In the present protocol, pyramidal neurons are labeled with DilC18(3) dye and analyzed using neuronal reconstruction software to assess neuronal morphology and dendritic spines. To investigate neuronal structure, dendritic branching analysis and Sholl analysis are performed, allowing researchers to draw inferences about dendritic branching complexity and neuronal arbor complexity, respectively. The evaluation of dendritic spines is conducted using an automatic assisted classification algorithm integral to the reconstruction software, which classifies spines into four categories (i.e., thin, mushroom, stubby, filopodia). Furthermore, an additional three parameters (i.e., length, head diameter, and volume) are also chosen to assess alterations in dendritic spine morphology. To validate the potential of wide application of the ballistic labeling technique, pyramidal neurons from in vitro cell culture were successfully labeled. Overall, the ballistic labeling method is unique and useful for visualizing neurons in different brain regions in rats, which in combination with sophisticated reconstruction software, allows researchers to elucidate the possible mechanisms underlying neurocognitive dysfunction.

We present a protocol to label and analyze pyramidal neurons, which is critical for evaluating potential morphological alterations in neurons and dendritic spines that may underlie neurochemical and behavioral abnormalities.

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of ...