This work was funded by National Institutes of Health Grant AR059397 (KSE).

Note: All animal work must be carried out according to institutional and animal care guidelines.
1. Preparation of Animal and Labeling
<ol>
	<li>Anesthetize rats: Anesthetize a rat (typically 150-400 g) using an intraperitoneal injection of a mixture of ketamine (50 mg/kg body weight), xylazine (5 mg/kg) and acepromazine (1 mg/kg). Check for reflexes by pinching the foot, wait till there is no response. Since the anesthesia response varies between animals, it is best to determine the effect of the anesthetics by testing pain reflexes.</li>
	<li>DiI Solution: Weigh DiI powder and dissolve in sterile 100% DMSO to achieve a 1.5% solution (15 mg/ml). Store the solution at room temperature in a dry atmosphere until use.</li>
	<li>DiI Injections: shave the hair over the calf area on both legs of the anesthetized rat using an electric razor.
	<ol>
		<li>Fill a 1 ml sterile syringe (1&#188; in&#160;27 G needle) with 200 &#181;l of 1.5% DiI solution. Slowly insert the needle parallel to the long-axis of each gastrocnemius muscle, and inject 100 &#181;l DiI into the muscle while slowly backing the needle out of the muscle (~10 sec) without withdrawing from the muscle.</li>
		<li>Prior to withdrawing the needle from the muscle, pull back on the syringe plunger to ensure the dye no longer flows from the tip, wait 10 sec, and then withdraw the needle from the muscle. This step prevents the leakage of DiI from the needle onto the skin, which could label cutaneous afferents.</li>
		<li>Repeat for other leg.</li>
		<li>Allow 4-5 days for the DiI to be transported from muscle to the DRG.</li>
	</ol>
	</li>
</ol>
2. Dissection and Neuronal Isolation
<ol>
	<li>Solutions:
	<ol>
		<li>Prepare Hank&#8217;s Balanced Salt Solution (HBSS) and Earle&#8217;s Balanced Salt Solution (EBSS) according to manufacturer&#8217;s instructions, filter sterilize and store at 4 &#176;C.</li>
		<li>Culture media (MEM Plus): Mix 89 ml minimal essential media (MEM), 10 ml fetal bovine serum (FBS), 1 ml penicillin-streptomycin solution, filter sterilize and store at 4 &#176;C.</li>
		<li>Polylysine solution: dissolve poly-L-lysine hydrobromide powder (10 mg/100 ml) in 0.1 M sodium borate solution, pH 8.4, filter sterilize and store aliquots at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Coating coverslips
	<ol>
		<li>Clean the glass coverslips (12 mm round) in 70% ethanol for 3-4 hr and air dry in sterile culture hood.</li>
		<li>Place the dried coverslips in the polylysine solution O/N.</li>
		<li>Remove from polylysine solution and wash in sterile distilled water 2x for 5 min each to remove any excess polylysine.</li>
		<li>Separate the coverslips, dry in culture hood, and store at 4 oC.</li>
	</ol>
	</li>
	<li>Setting up for dissection
	<ol>
		<li>Place on ice a beaker with 10 ml HBSS, a 35 mm Petri dish and a 100 mm Petri dish.</li>
		<li>Weigh out enzymes: 7 mg collagenase, 1 mg of DNAse and 10 mg of trypsin.</li>
		<li>Preheat a shaking water bath to 37 oC and place into the bath a 15 ml centrifuge tube containing 10 ml of EBSS to warm up.</li>
	</ol>
	</li>
	<li>Dissection:
	<ol>
		<li>Sacrifice rat according to an IACUC approved protocol.&#160;Carbon dioxide was used to euthanize the rat.</li>
		<li>Quickly remove the back by cutting open the skin on the back, remove the connective and muscle tissue along the spinal column for better exposure and remove the lumbar section of the vertebral column (end of the thoracic region to the iliac bone; Figure 1).</li>
		<li>Place the isolated vertebral column into the 100 mm Petri dish, and using a sterile Pasteur pipette apply chilled HBSS solution to keep the preparation moist and cold.</li>
		<li>Cut the muscles from the sides of the vertebral column. Perform a dorsal laminectomy, using rongeurs or scissors, and gently remove the bone enclosing the recesses containing the DRG.</li>
		<li>Repeatedly apply ice cold HBSS (~1 ml) during this procedure.</li>
		<li>Isolate the DRG by cutting the peripheral and central nerves (leaving 1-2 mm of nerve to facilitate handling of the ganglia)</li>
		<li>Place the isolated DRG into the 35 mm Petri dish containing ice-cold HBSS buffer.</li>
		<li>Repeat until all L-4 and L-5 DRG have been isolated. Thoracic DRG can be isolated and processed separately as a negative control for DiI labeling.</li>
		<li>Place the dish with isolated DRG under a dissection microscope and remove the excess nerves and blood vessels using micro scissors and forceps.</li>
		<li>Cut open the DRG for better exposure to the enzymes, but do not desheath the DRG.</li>
	</ol>
	</li>
	<li>DRG neuron dissociation:
	<ol>
		<li>Add the enzymes into the warm EBSS media (kept in the water bath). Shake briefly to dissolve, and filter sterilize.</li>
		<li>Using a sterile Pasteur pipette, place all DRG into the enzyme solution. Minimize the volume of HBSS that is added with DRG since the HBSS will dilute the enzyme mixture.&#160; Gas the flask with 95% O2 and 5% CO2 for 60 sec before tightly capping the flask.</li>
		<li>Incubate the flask in the shaking water bath set at 37 &#176;C for 60 min at 240 rpm.</li>
		<li>At the end of incubation, vigorously shake the flask 50x to dissociate the DRG5.</li>
		<li>Under sterile conditions, add 5 ml of MEM Plus to the flask to stop the enzymes.</li>
		<li>Transfer flask contents to a sterile 15 ml centrifuge tube and centrifuge at 16 x g for 6 min.</li>
		<li>Remove and discard the supernatant, add 5 ml of MEM Plus to the sample. Triturate gently with pipette to break up the pellet.</li>
		<li>Repeat centrifugation step, remove and discard the supernatant and add 500 &#181;l of MEM Plus. Gently triturate with a 1 ml pipette tip on a pipette to make a cell suspension.</li>
		<li>Place one poly-lysine coated glass coverslip into an empty sterile 35 mm dish. Add a drop (~50 &#181;l) of the cell suspension to the center of the coverslip.</li>
		<li>Repeat until cell solution is exhausted.</li>
		<li>Place the 35 mm dishes into a 37 &#176;C and 5% CO2 incubator for 1 hr to allow the neurons to adhere to the coverslips.</li>
		<li>Add an additional 2 ml of media into each dish and return to the incubator.</li>
	</ol>
	</li>
</ol>
3. Patch Clamp Recording
<ol>
	<li>Solutions for recording: these solutions are for recording NaV currents. Examples of solutions with differing composition can be found in these references5-7.
	<ol>
		<li>External solution: (in mM) 45 NaCl, 100 N-methyl-d-glucamine (NMG)&#8226;Cl, 4 MnCl2, 10 Na&#8226;HEPES and 10 glucose, with pH = 7.4 and osmolarity = 320 mOsm.</li>
		<li>Internal solution: (in mM) 104 NMG&#8226;Cl, 14 Creatine&#8226;PO4, 6 MgCl2, 10 NMG&#8226;HEPES, 5 Tris&#8226;ATP, 10 NMG2&#8226;EGTA, and 0.3 Tris2&#8226;GTP with pH 7.4 and osmolarity = 300 mOsm.</li>
	</ol>
	</li>
	<li>Measurement of ionic currents:
	<ol>
		<li>Pull recording electrodes from glass capillaries on an electrode puller and heat-polish the tips using a microforge. The resistance of the electrodes is typically 2-3 M&#937;</li>
		<li>Identify the DiI labeled DRG neurons using a microscope with epifluorescence and a TRITC cube. DiI has an excitation wavelength of 550 nm and emission wavelength of 570 nm.</li>
		<li>Fill the glass electrode with internal solution (only filing ~ 1 cm of the electrode tip) and gently tap to remove any trapped bubbles. Place in electrode holder and move to cell using a micromanipulator.</li>
		<li>Once the electrode touches the cell, apply gentle suction via the suction tube, until a tight seal is formed ( determined by the increase in electrode resistance).</li>
		<li>Clamp the membrane voltage at the desired holding potential (typically -60 to -120 mV).</li>
		<li>Obtain the whole-cell patch clamp configuration by applying stronger suction to remove the membrane patch at the tip of the electrode. The transition to whole-cell configuration is observed electrically as an increase in capacitative current8-10.</li>
		<li>Compensate the series resistance using the compensation circuitry of the patch clamp amplifier (use at least 80% compensation)8,10.</li>
	</ol>
	</li>
</ol>
4. Immunocytochemistry
<ol>
	<li>Fix the neurons on the coverslips with 4% paraformaldehyde for 1 hr and then permeabilize with 2% Tween 20 in phosphate buffered saline (PBS) for 10 min.</li>
	<li>Incubate the fixed cells O/N in a blocking solution (normal serum or Bovine Serum Albumin) plus the primary antibody, which is an antibody developed to bind to a specific antigen of interest.</li>
	<li>Incubate a control set of neurons O/N in blocking solution that lacks the primary antibodies.</li>
	<li>Wash the coverslips with PBS buffer 3x for 5 min followed by 2 washes of 5 min each with 1% Tween 20 to remove unbound primary antibody.</li>
	<li>Incubate test and control coverslips with the appropriate fluorescent tagged secondary antibody, which binds to the primary antibody to permit visualization.</li>
	<li>Wash the coverslips with PBS 3x for 5 min to remove excess secondary antibody.</li>
	<li>Mount the coverslips on a clean glass slide using mounting media.</li>
	<li>Apply clear nail polish to edge of the coverslip to fix it place. Once dry, visualize the neurons and capture images using the software accompanying the fluorescence microscope. 
	Note: In this work, a Nikon Microscope was used with NIS-elements BR 3.2 software. The exposure time was optimized and kept the same for control and test slides.</li>
	<li>Measure fluorescence intensity and cell size from individual neurons using imaging software such as ImageJ (rsbweb.nih.gov/ij/index.html).</li>
</ol>

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The authors declare no conflicts of interest.

Muscle afferents originating from triceps surae muscles (e.g. gastrocnemius) have been used in multiple studies to investigate the exercise pressor reflex (EPR)13. Given the interest in studying sensory neurons mediating the EPR, the gastrocnemius muscles were selected for dye injection. DiI was chosen due to its lipophilic nature and since it is not lost from the neuron by diffusion through the intact plasma membrane3,14. Using this method as a guide, one can inject the dye into any target...

Dorsal root ganglia (DRG) are comprised of soma from afferent neurons innervating various parts of the body, and important insights can be gained from studying sensory neurons with different innervation targets1. Various indirect or invasive methods are used to isolate and identify afferent neurons for e.g.&#160;injecting the nerve with markers like fluorogold1, injecting dye into the muscles by surgical method2. Of the different dyes that have been successfully used to label neurons and neuronal tracts, fluorescent dyes like DiI are widely used in neuroanatomical tracings owing to their rapid uptake by neuron terminals, long term retention in neurons, and the lack of transfer between neurons3. The afferent neurons labeled with DiI can be easily visualized using a fluorescent microscope for detailed study. However, one limitation is that markers such as DiI can be lost from cells during cell permeabilization needed for immunocytochemistry with intracellular epitopes4. Here we describe a method that uses DiI injected into skeletal muscle to identify muscle afferent neurons in the DRG. We also describe a cell permeabilization method that retains DiI in the neuron while permitting antibody access to intracellular epitopes. We demonstrate use of both patch clamp electrophysiology and immunocytochemistry to study these identified neurons, as well as describe the distinct advantages of each technique in studying ion channels that control neuronal excitability. Finally, we describe potential problems that can hinder interpretation of electrophysiological data. We have used these methods to identify muscle and cutaneous afferent neurons and determine the voltage-gated sodium (NaV) channels expressed in these identified neurons8. However, these methods can be adapted to a variety of preparations for studying identified subsets of neurons innervating a particular target tissue or brain nucleus.

<table border="0" cellpadding="0" cellspacing="0" width="480">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="22">
			<td height="22" style="height:22px;width:195px;">NaV 1.8</td>
			<td style="width:71px;">Abcam</td>
			<td style="width:136px;">Ab93616</td>
			<td style="width:79px;"></td>
		</tr>
		<tr height="81">
			<td height="81" style="height:81px;width:195px;">NaV 1.1 / 1.6/ 1.7 and 1.9</td>
			<td style="width:71px;">Alamone Labs, Jerusalem, Israel</td>
			<td style="width:136px;">ASC- 001/ 009 /008 /017</td>
			<td style="width:79px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:195px;">Alexa Fluor secondary antibodies</td>
			<td style="width:71px;">Invitrogen</td>
			<td style="width:136px;"></td>
			<td style="width:79px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:195px;">Normal goat serum</td>
			<td style="width:71px;"></td>
			<td align="right" style="width:136px;"></td>
			<td style="width:79px;"></td>
		</tr>
		<tr height="101">
			<td height="101" style="height:101px;width:195px;">Fluoro-Gel</td>
			<td style="width:71px;">Electron Microscopy Sciences</td>
			<td align="right" style="width:136px;">17985</td>
			<td style="width:79px;"></td>
		</tr>
	</tbody>
</table>

identification of specific sensory neuron populations for study of expressed ion channels

Sensory neurons transmit signals from various parts of the body to the central nervous system. The soma for these neurons are located in the dorsal root ganglia that line the spinal column. Understanding the receptors and channels expressed by these sensory afferent neurons could lead to novel therapies for disease. The initial step is to identify the specific subset of sensory neurons of interest. Here we describe a method to identify afferent neurons innervating the muscles by retrograde labeling using a fluorescent dye DiI (1,1&#39;-dioctadecyl-3,3,3&#39;,3&#39;-tetramethylindocarbocyanine perchlorate). Understanding the contribution of ion channels to excitation of muscle afferents could help to better control excessive excitability induced by certain disease states such as peripheral vascular disease or heart failure. We used two approaches to identify the voltage dependent ion channels expressed by these neurons, patch clamp electrophysiology and immunocytochemistry. While electrophysiology plus pharmacological blockers can identify functional ion channel types, we used immunocytochemistry to identify channels for which specific blockers were unavailable and to better understand the ion channel distribution pattern in the cell population. These techniques can be applied to other areas of the nervous system to study specific neuronal groups.

Using the method described here, we identified muscle afferent neurons in DRG and went on to determine the voltage-gated sodium channels (NaV) expressed by these neurons. DiI labeled muscle afferent neurons (Figure 1C) comprised a small fraction of the total population of neurons in the DRG with an average of 16% of neurons (167/1047) labeled. The quantification was done by first measuring the fluorescence using ImageJ and plotting histograms of the intensity data to determine the threshold for positivel...

Watch this Scientific Journal Video about Identification of Specific Sensory Neuron Populations for Study of Expressed Ion Channels at JoVE.com

Identification of Specific Sensory Neuron Populations for Study of Expressed Ion Channels

Afferent sensory neurons signal sensory information from the periphery to the central nervous system. Identifying specific afferent neurons will help in understanding their physiology. We describe a method of retrograde labeling to identify afferent neurons, and study the voltage-gated ion channels in these neurons using patch clamp electrophysiology and immunocytochemistry.

Sensory neurons transmit signals from various parts of the body to the central nervous system. The soma for these neurons are located in the dorsal root ...

identification-specific-sensory-neuron-populations-for-study

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Neuroscience

6.7K Görüntüleme.  AT Still University of Health Sciences. Afferent sensory neurons signal sensory information from the periphery to the central nervous system. Identifying specific afferent neurons will help in understanding their physiology. We describe a method of retrograde labeling to identify afferent neurons, and study the voltage-gated ion channels in these neurons using patch clamp electrophysiology and immunocytochemistry.

Video: Identification of Specific Sensory Neuron Populations for Study of Expressed Ion Channels

Laser Capture Microdissection of Drosophila Peripheral Neurons

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the molecular mechanisms underlying class-specific dendrite morphogenesis1,2. To facilitate molecular analyses of class-specific da neuron development, it is vital to obtain these cells in a pure population. Although a range of different cell, and tissue-specific RNA isolation techniques exist for Drosophila cells, including magnetic bead based cell purification3,4, Fluorescent Activated Cell Sorting (FACS)5-8, and RNA binding protein based strategies9, none of these methods can be readily utilized for isolating single or multiple class-specific Drosophila da neurons with a high degree of spatial precision. Laser Capture Microdissection (LCM) has emerged as an extremely powerful tool that can be used to isolate specific cell types from tissue sections with a high degree of spatial resolution and accuracy. RNA obtained from isolated cells can then be used for analyses including qRT-PCR and microarray expression profiling within a given cell type10-16. To date, LCM has not been widely applied in the analysis of Drosophila tissues and cells17,18, including da neurons at the third instar larval stage of development. 
Here we present our optimized protocol for isolation of Drosophila da neurons using the infrared (IR) class of LCM. This method allows for the capture of single, class-specific or multiple da neurons with high specificity and spatial resolution. Age-matched third instar larvae expressing a UAS-mCD8::GFP19 transgene under the control of either the class IV da neuron specific ppk-GAL420 driver or the pan-da neuron specific 21-7-GAL421 driver were used for these experiments. RNA obtained from the isolated da neurons is of very high quality and can be directly used for downstream applications, including qRT-PCR or microarray analyses. Furthermore, this LCM protocol can be readily adapted to capture other Drosophila cell types a various stages of development dependent upon the cell type specific, GAL4-driven expression pattern of GFP.

Laser Capture Microdissection of Drosophila Peripheral Neurons

In this video-article we present a method for isolating single or multiple Drosophila da neurons from third instar larvae using the infrared capture (IR) class of Laser Capture Microdissection (LCM). RNA obtained from the isolated neurons can be readily used for downstream applications including qRT-PCR or microarray analyses.

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the ...

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 ...

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.

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.

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

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 ...

Mouse Hindbrain Ex Vivo Culture to Study Facial Branchiomotor Neuron Migration

Embryonic neurons are born in the ventricular zone of the brain, but subsequently migrate to new destinations to reach appropriate targets. Deciphering the molecular signals that cooperatively guide neuronal migration in the embryonic brain is therefore important to understand how the complex neural networks form which later support postnatal life. Facial branchiomotor (FBM) neurons in the mouse embryo hindbrain migrate from rhombomere (r) 4 caudally to form the paired facial nuclei in the r6-derived region of the hindbrain. Here we provide a detailed protocol for wholemount ex vivo culture of mouse embryo hindbrains suitable to investigate the signaling pathways that regulate FBM migration. In this method, hindbrains of E11.5 mouse embryos are dissected and cultured in an open book preparation on cell culture inserts for 24 hr. During this time, FBM neurons migrate caudally towards r6 and can be exposed to function-blocking antibodies and small molecules in the culture media or heparin beads loaded with recombinant proteins to examine roles for signaling pathways implicated in guiding neuronal migration.

Mouse Hindbrain Ex Vivo Culture to Study Facial Branchiomotor Neuron Migration

Embryonic neurons are born in the ventricular zone of the neural tube, but migrate to reach appropriate targets. Facial branchiomotor (FBM) neurons are a useful model to study neuronal migration. This protocol describes the wholemount ex vivo culture of mouse embryo hindbrains to investigate mechanisms that regulate FBM migration.

Embryonic neurons are born in the ventricular zone of the brain, but subsequently migrate to new destinations to reach appropriate targets. Deciphering ...

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 ...

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat open surfaces is complex,&#160;including: pausing, turning, and meandering. This complexity in the repertoire of movement hinders detailed analysis of the events occurring during a single crawl stride cycle. To overcome this obstacle, linear agarose channels were made that constrain larval behavior to straight, sustained, rhythmic crawling. In principle, because agarose channels and the Drosophila larval body are both optically clear, the movement of larval structures labeled by genetically-encoded fluorescent probes can be monitored in intact, freely-moving larvae. In the past, larvae were placed in linear channels and crawling at the level of whole organism, segment, and muscle were analyzed1. In the future, larvae crawling in channels can be used for calcium imaging to monitor neuronal activity. Moreover, these methods can be used with larvae of any genotype and with any researcher-designed channel. Thus the protocol presented below is widely applicable for studies using the Drosophila larva as a model to understand motor control.

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat ...

Physiological Preparation of Hair Cells from the Sacculus of the American Bullfrog (Rana catesbeiana)

The study of hearing and balance rests upon insights drawn from biophysical studies of model systems. One such model, the sacculus of the American bullfrog, has become a mainstay of auditory and vestibular research. Studies of this organ have revealed how sensory cells hair can actively detect signals from the environment. Because of these studies, we now better understand the mechanical gating and localization of a hair cell&#39;s transduction channels, calcium&#39;s role in mechanical adaptation, and the identity of hair cell currents. This highly accessible organ continues to provide insight into the workings of hair cells. Here we describe the preparation of the bullfrog&#39;s sacculus for biophysical studies on its hair cells. We include the complete dissection procedure and provide specific protocols for the preparation of the sacculus in specific contexts. We additionally include representative results using this preparation, including the calculation of a hair bundle&#39;s instantaneous force-displacement relation and measurement of a bundle&#39;s spontaneous oscillation.

Physiological Preparation of Hair Cells from the Sacculus of the American Bullfrog Rana catesbeiana

The American bullfrog&#39;s (Rana catesbeiana) sacculus permits direct examination of hair-cell physiology. Here the dissection and preparation of the bullfrog&#39;s sacculus for biophysical studies is described. We show representative experiments from these hair cells, including the calculation of a bundle&#39;s force-displacement relation and measurement of its unforced motion.

The study of hearing and balance rests upon insights drawn from biophysical studies of model systems. One such model, the sacculus of the American ...

Stimulus-specific Cortical Visual Evoked Potential Morphological Patterns

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using 128-channel high-density electroencephalography (EEG). The specific aim of the described stimuli and analyses is to examine whether it is feasible to replicate previously reported CVEP morphological patterns elicited by an apparent motion stimulus, designed to simultaneously stimulate both ventral and dorsal central visual networks, using object and motion stimuli designed to separately stimulate ventral and dorsal visual cortical networks.&#160; Four visual paradigms are presented: 1. Randomized visual objects with consistent temporal presentation. 2. Randomized visual objects with inconsistent temporal presentation (or jitter).&#160; 3. Visual motion via a radial field of coherent central dot motion without jitter.&#160; 4. Visual motion via a radial field of coherent central dot motion with jitter.&#160; These four paradigms are presented in a pseudo-randomized order for each participant.&#160; Jitter is introduced in order to view how possible anticipatory-related effects may affect the morphology of the object-onset and motion-onset CVEP response.&#160; EEG data analyses are described in detail, including steps of data exportation from and importation to signal processing platforms, bad channel identification&#160;and removal, artifact rejection, averaging, and categorization of average CVEP morphological pattern type based upon latency ranges of component peaks. Representative data show that the methodological approach is indeed sensitive in eliciting differential object-onset and motion-onset CVEP morphological patterns and may, therefore, be useful in addressing the larger research aim. Given the high temporal resolution of EEG and the possible application of high-density EEG in source localization analyses, this protocol is ideal for the investigation of distinct CVEP morphological patterns and the underlying neural mechanisms which generate these differential responses.&#160; &#160; &#160;&#160;

In this paper, we present a protocol to investigate differential cortical visual evoked potential morphological patterns through stimulation of ventral and dorsal networks using high-density EEG. Visual object and motion stimulus paradigms, with and without temporal jitter, are described. Visual evoked potential morphological analyses are also outlined.&#160;&#160;

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using ...

Assessment of Sensory Thresholds in Dogs Using Mechanical and Hot Thermal Quantitative Sensory Testing

Quantitative sensory testing (QST) is used to evaluate the function of the somatosensory system in dogs by assessing the response to applied mechanical and thermal stimuli. QST is used to determine normal dogs&#39; sensory thresholds and evaluate alterations in peripheral and central sensory pathways caused by various disease states, including osteoarthritis, spinal cord injury, and cranial cruciate ligament rupture. Mechanical sensory thresholds are measured by electronic von Frey anesthesiometers and pressure algometers. They are determined as the force at which the dog exhibits a response indicating conscious stimulus perception. Hot thermal sensory thresholds are the latency to respond to a fixed or ramped temperature stimulus applied by a contact thermode.
Following a consistent protocol for performing QST and paying attention to details of the testing environment, procedure, and individual study subjects are critical for obtaining accurate QST results for dogs. Protocols for the standardized collection of QST data in dogs have not been described in detail. QST should be performed in a quiet, distraction-free environment that is comfortable for the dog, the QST operator, and the handler. Ensuring that the dog is calm, relaxed, and properly positioned for each measurement helps produce reliable, consistent responses to the stimuli and makes the testing process more manageable. The QST operator and handler should be familiar and comfortable with handling dogs and interpreting dogs&#39; behavioral responses to potentially painful stimuli to determine the endpoint of testing, reduce stress, and maintain safety during the testing process.

This work describes a standard protocol for mechanical and hot thermal quantitative sensory testing to evaluate the somatosensory system in dogs. Sensory thresholds are measured using an electronic von Frey anesthesiometer, pressure algometer, and hot contact thermode.

Quantitative sensory testing (QST) is used to evaluate the function of the somatosensory system in dogs by assessing the response to applied mechanical ...