This work was supported in part by Grants from the National Key Research and Development Program of China (2022YFA1105503), the State Key Laboratory of Neuroscience (SKLN-202103), Zhejiang Natural Science Foundation of China (Y21H120019).

The care and handling of animals were approved by the Regulation of the Ethics Committee of Wenzhou Medical University in accordance with the ARVO guidelines. Adult mice (C57BL/6J, male and female, 8 to 12 weeks of age) were utilized in this research. The equipment and reagents needed for the study are listed in the Table of Materials.
1. Preparation for retina fixation
<ol>
	<li>Assemble the following materials and tools: a dissecting microscope, two forceps with very fine tips, scissors, a 1 mL syringe needle (needle size: G26), and filter paper.</li>
	<li>Deeply anesthetize mice by intraperitoneal injection of 2,2,2-tribromoethanol at 0.25 mg/g of body weight. Subsequently, decapitate the mice and cut their heads16.</li>
	<li>Enucleate their eyes with elbow scissors and place them in a glass dish containing 4% paraformaldehyde-0.2% picric acid in 0.1 M phosphate buffer (PB; pH 7.4).</li>
	<li>Under the dissecting microscope, punch a hole at the corneal limbus using a 1 mL syringe needle and cut off the anterior segment with scissors, following the hole. Then, remove the lens from the inner retinal surface with forceps.</li>
	<li>Use two forceps to carefully peel the sclera until the retina is completely isolated from the eyecup (the cuppy retina tissue separated from the choroid completely). Subsequently, cut the retina into four pieces.</li>
	<li>Fix these sections in 4% paraformaldehyde-0.2% picric acid in 0.1 M PB (pH 7.4) for 2 h at room temperature (RT), and then transfer the tissue to 4% paraformaldehyde in 0.1 M PB (pH 10.4) overnight at 4 &#176;C.</li>
</ol>
2. Pre-embedding immunostaining
<ol>
	<li>After washing in 0.01 M PBS (pH 7.4) six times for 10 min each, incubate the retinal tissues in 1% sodium borohydride (NaBH4) in 0.01 M PBS (pH 7.4) for 30 min.</li>
	<li>Wash the retinal sections in 0.01 M PBS (pH 7.4) at least six times. Meanwhile, remove the vitreous with filter paper and then cut the retina into small slices between 100-300 &#181;m using a double-edged razor blade.</li>
	<li>After blocking with 5% normal goat serum (NGS) in 0.1 M PB (pH 7.4) for 1 h at RT, incubate the retinal slices with primary antibodies (Anti-rabbit PKC&#945;, 1:80; Anti-Rabbit SP, 1:100) along with 2% NGS in 0.01 M PBS (pH 7.4) for 2 h at RT on a shaker. Subsequently, incubate for 96 h (5 days) at 4 &#176;C on a shaker.</li>
	<li>After washing six times for 10 min each in 0.01 M PBS (pH 7.4), incubate the retinal slices with the secondary antibody at 1:200, such as goat anti-rabbit IgG, along with 2% NGS in 0.01 M PBS (pH 7.4) for 2 h at RT on a shaker. Next, incubate for 48 h (2 days) at 4 &#176;C on a shaker. 
	NOTE: Secondary antibodies were applied alone in control experiments.</li>
	<li>Wash the retinal stripes in 0.01 M PBS (pH 7.4) six times for 10 min each before incubating with reagent A and reagent B from the ABC kit at 1:100 in 0.01M PBS (pH 7.4) for 2 days at 4&#176;C. (Both reagent A and reagent B were diluted with 0.01 M PBS. For example: A solution: 6 &#956;l; B solution: 6 &#956;l; 0.01 M PBS: 588 &#956;l, total of 600 &#956;l).</li>
	<li>Wash the retinal stripes in 0.05 M Tris buffer (pH 7.2) three times for 10 min each. Then, pre-incubate the retinal stripes with 5% reagent 1 and 5% reagent 3 from the DAB kit in distilled water for 1 hour at room temperature. (The ratio for example: reagent 1: 50 &#956;l; reagent 3: 50 &#956;l; distilled water: 900 &#956;l, total of 1000 &#956;l).</li>
	<li>Stain the retinal stripes with DAB. Add the same volume of solution from three tubes in the DAB kit in sequence (reagent 1-reagent 2-reagent 3). Observe the staining condition using the dissection microscope. Stop the staining when cells become brown; this process takes 10-20 min.</li>
	<li>Wash the retinal stripes with 0.05 M Tris buffer (pH 7.2) three times for 10 min each and then wash in 0.01 M PBS six times for 10 min each.</li>
</ol>
3. Post-fixation and embedding
<ol>
	<li>Fix the retinal stripes in 2% glutaraldehyde for 1-2 h at room temperature (RT), and then wash in 0.01 M PBS (pH 7.4) six times for 10 min each.</li>
	<li>Incubate the retinal stripes with 1% osmium tetroxide (OsO4) in 0.1 M PB for 1 h at RT and keep them in a dark place.</li>
	<li>After washing in distilled water (ddH2O) six times for 10 min each, incubate the retinal tissues with uranyl acetate for 1 h at RT and keep them in a dark place to stain these tissues.</li>
	<li>Put the retinal tissues in acetone solutions (50%, 70%, 80%, and 90%) for 10 min each, then in 100% twice for 10 min each.</li>
	<li>Dip the retinal tissues in a mixture containing the same volume of uranyl acetate and an epoxy resin for 1 h at 37 &#176;C drying oven, followed by a mixture containing uranyl acetate and the resin (1:4) overnight at 37 &#176;C drying oven.</li>
	<li>Transfer the retinal tissues gently using a toothpick into the new resin for 1 h at 45 &#176;C in a drying oven, and then the orientated retinal stripe is embedded in the embedding plate with the resin.</li>
	<li>Put the sample in a 45 &#176;C drying oven for 3 h and a 65 &#176;C drying oven for 48 h.</li>
	<li>Shape the embedding blocks into trapezoids and cut the block into 1 &#181;m thick sections with an ultramicrotome, stain these sections with toluidine blue, and prescreen regions of interest under a light microscope.</li>
	<li>Collect ultrathin sections (70-90 nm) on copper grids and view them under an electron microscope.</li>
</ol>

Synaptic Circuits Analysis and Protein Localization in the Mouse Retina

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The authors have no disclosures.

This article has described three critical steps for the successful observation of synaptic circuits and protein localization: (1) quick and weak fixation, (2) pre-embedding immunostaining, and (3) post-fixation and embedding.
We propose that fixation is the key step for a successful pre-embedding immuno-EM approach. Thus, the importance of fresh fixative and fast fixation is emphasized here, naming this principle the &#34;4F principle,&#34; which is crucial in tissue preparation. However, achi...

The complexity of neuronal circuits in the retina presents challenges for exploration, considering the diverse cell types within its structure1,2. The initial step involves identifying synaptic connections between different cells and determining the cellular localization of specific neurotransmitters or neuropeptides. As molecular biology advances introduce new proteins, precise localization in the retina becomes crucial for understanding their functions and analyzing retinal circuits and synaptic connections3,4,5.
Due to the limited resolution of light microscopy, electron microscopy (EM) is commonly used to detect the subcellular structures of nerve cells. EM has various classifications, with conventional transmission electron microscopy (TEM) utilized for observing cell ultrastructures6,7,8,9. Immunoelectron microscopy (immuno-EM), which combines the spatial resolution of EM with the chemical identification ability of antibodies binding specifically to proteins10, stands out as the optimal and exclusive method for investigating synaptic connections and subcellular protein localization in the retina11,12.
Immuno-EM techniques can be divided into pre-embedding and post-embedding methods based on the order of embedding and antibody incubation. Compared with the post-embedding method, the pre-embedding approach is capable of large-scale and long-distance identification13,14,15, offering an optimal approach for studying cell processes like axons and dendrites. Additionally, this technique provides a strong signal and broad field of view, making it advantageous for comprehensive investigations of protein expression and molecular localization in the cytoplasm. This method proves particularly valuable in ensuring chemically identified structures that are visible throughout the entire cytoplasm, cells, or retina.
However, the post-embedding method, while having lower penetration or diffusion compared to the pre-embedding method, is not as sensitive16,17. In simple terms, if the goal is to explore the localization of specific neurotransmitters in the cytoplasm or synaptic terminals, the pre-embedding immuno-EM is the preferred method. Conversely, for identifying the localization of membrane receptors, it is more recommended to utilize post-embedding immunogold EM.
Given these considerations, we opt for the pre-embedding immuno-EM method to delve into retinal circuits, including the interaction between cone and rod pathways, and molecular localization, such as the distribution and synaptic organization of substance P-like immunoreactivity (SP-IR) in the mouse retina.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 mL syringe needle</td>
			<td>kangdelai</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1% OsO4</td>
			<td>Electron Microscopy Science</td>
			<td>19100</td>
			<td></td>
		</tr>
		<tr>
			<td>2,2,2-Tribromoethanol</td>
			<td>Sigma-Aldrich</td>
			<td>T48402</td>
			<td></td>
		</tr>
		<tr>
			<td>8% Glutaraldehyde</td>
			<td>Electron Microscopy Science</td>
			<td>16020</td>
			<td></td>
		</tr>
		<tr>
			<td>8% Paraformaldehyde</td>
			<td>Electron Microscopy Science</td>
			<td>157-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetone</td>
			<td>Electron Microscopy Science</td>
			<td>10000</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-rabbit PKC</td>
			<td>Sigma-Aldrich</td>
			<td>P4334</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-Rabbit SP</td>
			<td>Abcam</td>
			<td>ab67006</td>
			<td></td>
		</tr>
		<tr>
			<td>DAB Substrate kit</td>
			<td>MXB Biotechnologies</td>
			<td>KIT-9701/9702/9703</td>
			<td></td>
		</tr>
		<tr>
			<td>Elbow scissors</td>
			<td>Suzhou66 vision company</td>
			<td>54010</td>
			<td></td>
		</tr>
		<tr>
			<td>Electron microscope</td>
			<td>Phillips</td>
			<td>CM120</td>
			<td></td>
		</tr>
		<tr>
			<td>Epon resin</td>
			<td>Electron Microscopy Science</td>
			<td>14910</td>
			<td></td>
		</tr>
		<tr>
			<td>forcep</td>
			<td>Suzhou66 vision company</td>
			<td>S101A</td>
			<td></td>
		</tr>
		<tr>
			<td>Millipore filter paper</td>
			<td>Merck Millipore&#160;</td>
			<td>PR05538</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2HPO4&#183; 12H2O</td>
			<td>Sigma</td>
			<td>71650</td>
			<td>A component of phosphate buffer</td>
		</tr>
		<tr>
			<td>NaH2PO4&#183; H2O</td>
			<td>Sigma</td>
			<td>71507</td>
			<td>A component of phosphate buffer</td>
		</tr>
		<tr>
			<td>Picric acid</td>
			<td>Electron Microscopy Science</td>
			<td>19550</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium borohydride (NaBH4)&#160;</td>
			<td>Sigma</td>
			<td>215511</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris</td>
			<td>Solarbio</td>
			<td>917R071</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultramicrotome</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Uranyl acetate</td>
			<td>Electron Microscopy Science</td>
			<td>22400</td>
			<td></td>
		</tr>
		<tr>
			<td>VACTASTAIN ABC kit, Peroxidase (Rabbit IgG)</td>
			<td>Vector Laboratories</td>
			<td>PK-4001</td>
			<td></td>
		</tr>
	</tbody>
</table>

investigating retinal circuits and molecular localization by pre&#45;embedding immunoelectron microscopy

The retina comprises numerous cells forming diverse neuronal circuits, which constitute the first stage of the visual pathway. Each circuit is characterized by unique features and distinct neurotransmitters, determining its role and functional significance. Given the intricate cell types within its structure, the complexity of neuronal circuits in the retina poses challenges for exploration. To better investigate retinal circuits and cross-talk, such as the link between cone and rod pathways, and precise molecular localization (neurotransmitters or neuropeptides), such as the presence of substance P-like immunoreactivity in the mouse retina, we employed a pre-embedding immunoelectron microscopy (immuno-EM) method to explore synaptic connections and organization. This approach enables us to pinpoint specific intercellular synaptic connections and precise molecular localization and could play a guiding role in exploring its function. This article describes the protocol, reagents used, and detailed steps, including (1) retina fixation preparation, (2) pre-embedding immunostaining, and (3) post-fixation and embedding.

Figure 1 shows examples of control experiments without the incubation of primary antibodies against either protein kinase C alpha (PKC&#945;) or SP, in which no immunoreactivity (IR) was found.
Figure 2 depicts the PKC&#945;-IR in the mouse retina. PKC&#945; serves as a marker for all rod bipolar cells (RBC) in the retina18. At the electron microscopy (EM) level, RBC can be identified through PKC&#945;-IR, visu...

Author Spotlight: Understanding the Ultrastructural Basis of Retinal Synaptic Connectivity and Neurotransmitter Localization in Mice 

This protocol outlines the detailed steps of pre-embedding immunoelectron microscopy, with a focus on exploring synaptic circuits and protein localization in the retina.

Investigating Retinal Circuits and Molecular Localization by Pre&#45;Embedding Immunoelectron Microscopy

The retina comprises numerous cells forming diverse neuronal circuits, which constitute the first stage of the visual pathway. Each circuit is ...

investigating-retinal-circuits-molecular-localization-pre-embedding

Research

JoVE Journal

Neuroscience

253 Views.  Wenzhou Medical University. This protocol outlines the detailed steps of pre-embedding immunoelectron microscopy, with a focus on exploring synaptic circuits and protein localization in the retina.

Video: Author Spotlight: Understanding the Ultrastructural Basis of Retinal Synaptic Connectivity and Neurotransmitter Localization in Mice 

Mapping Inhibitory Neuronal Circuits by Laser Scanning Photostimulation

Inhibitory neurons are crucial to cortical function. They comprise about 20% of the entire cortical neuronal population and can be further subdivided into diverse subtypes based on their immunochemical, morphological, and physiological properties1-4. Although previous research has revealed much about intrinsic properties of individual types of inhibitory neurons, knowledge about their local circuit connections is still relatively limited3,5,6. Given that each individual neuron's function is shaped by its excitatory and inhibitory synaptic input within cortical circuits, we have been using laser scanning photostimulation (LSPS) to map local circuit connections to specific inhibitory cell types. Compared to conventional electrical stimulation or glutamate puff stimulation, LSPS has unique advantages allowing for extensive mapping and quantitative analysis of local functional inputs to individually recorded neurons3,7-9. Laser photostimulation via glutamate uncaging selectively activates neurons perisomatically, without activating axons of passage or distal dendrites, which ensures a sub-laminar mapping resolution. The sensitivity and efficiency of LSPS for mapping inputs from many stimulation sites over a large region are well suited for cortical circuit analysis.
Here we introduce the technique of LSPS combined with whole-cell patch clamping for local inhibitory circuit mapping. Targeted recordings of specific inhibitory cell types are facilitated by use of transgenic mice expressing green fluorescent proteins (GFP) in limited inhibitory neuron populations in the cortex3,10, which enables consistent sampling of the targeted cell types and unambiguous identification of the cell types recorded. As for LSPS mapping, we outline the system instrumentation, describe the experimental procedure and data acquisition, and present examples of circuit mapping in mouse primary somatosensory cortex. As illustrated in our experiments, caged glutamate is activated in a spatially restricted region of the brain slice by UV laser photolysis; simultaneous voltage-clamp recordings allow detection of photostimulation-evoked synaptic responses. Maps of either excitatory or inhibitory synaptic input to the targeted neuron are generated by scanning the laser beam to stimulate hundreds of potential presynaptic sites. Thus, LSPS enables the construction of detailed maps of synaptic inputs impinging onto specific types of inhibitory neurons through repeated experiments. Taken together, the photostimulation-based technique offers neuroscientists a powerful tool for determining the functional organization of local cortical circuits.

This paper introduces an approach of combining laser scanning photostimulation with whole cell recordings in transgenic mice expressing GFP in limited inhibitory neuron populations. The technique allows for extensive mapping and quantitative analysis of local synaptic circuits of specific inhibitory cortical neurons.

Inhibitory neurons are crucial to cortical function. They comprise about 20% of the entire cortical neuronal population and can be further subdivided into ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

SNARE-mediated Fusion of Single Proteoliposomes with Tethered Supported Bilayers in a Microfluidic Flow Cell Monitored by Polarized TIRF Microscopy

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. During neurotransmitter or hormone release via exocytosis, the fusion pore can transiently open and close repeatedly, regulating cargo release kinetics. Pore dynamics also determine the mode of vesicle recycling; irreversible resealing results in transient, &#34;kiss-and-run&#34; fusion, whereas dilation leads to full fusion. To better understand what factors govern pore dynamics, we developed an assay to monitor membrane fusion using polarized total internal reflection fluorescence (TIRF) microscopy with single molecule sensitivity and ~15 msec time resolution in a biochemically well-defined in vitro system. Fusion of fluorescently labeled small unilamellar vesicles containing v-SNARE proteins (v-SUVs) with a planar bilayer bearing t-SNAREs, supported on a soft polymer cushion (t-SBL, t-supported bilayer), is monitored. The assay uses microfluidic flow channels that ensure minimal sample consumption while supplying a constant density of SUVs. Exploiting the rapid signal enhancement upon transfer of lipid labels from the SUV to the SBL during fusion, kinetics of lipid dye transfer is monitored. The sensitivity of TIRF microscopy allows tracking single fluorescent lipid labels, from which lipid diffusivity and SUV size can be deduced for every fusion event. Lipid dye release times can be much longer than expected for unimpeded passage through permanently open pores. Using a model that assumes retardation of lipid release is due to pore flickering, a pore &#34;openness&#34;, the fraction of time the pore remains open during fusion, can be estimated. A soluble marker can be encapsulated in the SUVs for simultaneous monitoring of lipid and soluble cargo release. Such measurements indicate some pores may reseal after losing a fraction of the soluble cargo.

Here, we present a protocol to detect single, SNARE-mediated fusion events between liposomes and supported bilayers in microfluidic channels using polarized TIRFM, with single molecule sensitivity and ~15 msec time resolution. Lipid and soluble cargo release can be detected simultaneously. Liposome size, lipid diffusivity, and fusion pore properties are measured.

In the ubiquitous process of membrane fusion the opening of a fusion pore establishes the first connection between two formerly separate compartments. ...

Localization of Odorant Receptor Genes in Locust Antennae by RNA In Situ Hybridization

Insects have evolved sophisticated olfactory reception systems to sense exogenous chemical signals. These chemical signals are transduced by Olfactory Receptor Neurons (ORNs) housed in hair-like structures, called chemosensilla, of the antennae. On the ORNs&#39; membranes, Odorant Receptors (ORs) are believed to be involved in odor coding. Thus, being able to identify genes localized to the ORNs is necessary to recognize OR genes, and provides a fundamental basis for further functional in situ studies. The RNA expression levels of specific ORs in insect antennae are very low, and preserving insect tissue for histology is challenging. Thus, it is difficult to localize an OR to a specific type of sensilla using RNA in situ hybridization. In this paper, a detailed and highly effective RNA in situ hybridization protocol particularly for lowly expressed OR genes of insects, is introduced. In addition, a specific OR gene was identified by conducting double-color fluorescent in situ hybridization experiments using a co-expressing receptor gene, Orco, as a marker.

Localization of Odorant Receptor Genes in Locust Antennae by RNA In Situ Hybridization

This paper describes a detailed and highly effective RNA in situ hybridization protocol particularly for low-level expressed Odorant Receptor (OR) genes, as well as other genes, in insect antennae using digoxigenin (DIG)-labeled or biotin-labeled probes.

Insects have evolved sophisticated olfactory reception systems to sense exogenous chemical signals. These chemical signals are transduced by Olfactory ...

Optogenetic Manipulation of Neural Circuits During Monitoring Sleep/wakefulness States in Mice

In recent years, optogenetics has been widely used in many fields of neuroscientific research. In many cases, an opsin, such as channel rhodopsin 2 (ChR2), is expressed by a virus vector in a particular type of neuronal cells in various Cre-driver mice. Activation of these opsins is triggered by application of light pulses which are delivered by laser or LED through optic cables, and the effect of activation is observed with very high time resolution. Experimenters are able to acutely stimulate neurons while monitoring behavior or another physiological outcome in mice. Optogenetics can enable useful strategies to evaluate function of neuronal circuits in the regulation of sleep/wakefulness states in mice. Here we describe a technique for examining the effect of optogenetic manipulation of neurons with a specific chemical identity during electroencephalogram (EEG) and electromyogram (EMG) monitoring to evaluate the sleep stage of mice. As an example, we describe manipulation of GABAergic neurons in the bed nucleus of the stria terminalis (BNST). Acute optogenetic excitation of these neurons triggers a rapid transition to wakefulness when applied during NREM sleep. Optogenetic manipulation along with EEG/EMG recording can be applied to decipher the neuronal circuits that regulate sleep/wakefulness states.

Here, we describe methods of optogenetic manipulation of particular types of neurons during monitoring of sleep/wakefulness states in mice, presenting our recent work on the bed nucleus of the stria terminalis as an example.

In recent years, optogenetics has been widely used in many fields of neuroscientific research. In many cases, an opsin, such as channel rhodopsin 2 ...

Localization of the Locus Coeruleus in the Mouse Brain

The locus coeruleus (LC) is a major hub of norepinephrine producing neurons that modulate a number of physiological functions. Structural or functional abnormalities of LC impact several brain regions including cortex, hippocampus, and cerebellum and may contribute to depression, bipolar disorder, anxiety, as well as Parkinson disease and Alzheimer disease. These disorders are often associated with metal misbalance, but the role of metals in LC is only partially understood. Morphologic and functional studies of LC are needed to better understand the human pathologies and contribution of metals. Mice are a widely used experimental model, but the mouse LC is small (~0.3 mm diameter) and hard to identify for a non-expert. Here, we describe a step-by-step immunohistochemistry-based protocol to localize the LC in the mouse brain. Dopamine-&#946;-hydroxylase (DBH), and alternatively, tyrosine hydroxylase (TH), both enzymes highly expressed in the LC, are used as immunohistochemical markers in brain slices. Sections adjacent to LC-containing sections can be used for further analysis, including histology for morphological studies, metabolic testing, as well as metal imaging by X-ray fluorescence microscopy (XFM).

The locus coeruleus is a small cluster of neurons involved in a variety of physiological processes. Here, we describe a protocol to prepare mouse brain sections for studies of proteins and metals in this nucleus.

The locus coeruleus (LC) is a major hub of norepinephrine producing neurons that modulate a number of physiological functions. Structural or functional ...

Analysis of Spliceosomal snRNA Localization in Human Hela Cells Using Microinjection

Biogenesis of spliceosomal snRNAs is a complex process involving both nuclear and cytoplasmic phases and the last step occurs in a nuclear compartment called the Cajal body. However, sequences that direct snRNA localization into this subnuclear structure have not been known until recently. To determine sequences important for accumulation of snRNAs in Cajal bodies, we employed microinjection of fluorescently labelled snRNAs followed by their localization inside cells. First, we prepared snRNA deletion mutants, synthesized DNA templates for in vitro transcription and transcribed snRNAs in the presence of UTP coupled with Alexa488. Labelled snRNAs were mixed with 70 kDa-Dextran conjugated with TRITC, and microinjected to the nucleus or the cytoplasm of human HeLa cells. Cells were incubated for 1 h and fixed and the Cajal body marker coilin was visualized by indirect immunofluorescence, while snRNAs and dextran, which serves as a marker of nuclear or cytoplasmic injection, were observed directly using a fluorescence microscope. This method allows for efficient and rapid testing of how various sequences influence RNA localization inside cells. Here, we show the importance of the Sm-binding sequence for efficient localization of snRNAs into the Cajal body.

Biogenesis of spliceosomal snRNAs is a complex process involving various cellular compartments. Here, we employed microinjection of fluorescently labelled snRNAs in order to monitor their transport inside the cell.

Biogenesis of spliceosomal snRNAs is a complex process involving both nuclear and cytoplasmic phases and the last step occurs in a nuclear compartment ...

Investigating Long-term Synaptic Plasticity in Interlamellar Hippocampus CA1 by Electrophysiological Field Recording

The study of synaptic plasticity in the hippocampus has focused on the use of the CA3-CA1 lamellar network. Less attention has been given to the longitudinal interlamellar CA1-CA1 network. Recently however, an associational connection between CA1-CA1 pyramidal neurons has been shown. Therefore, there is the need to investigate whether the longitudinal interlamellar CA1-CA1 network of the hippocampus supports synaptic plasticity.
We designed a protocol to investigate the presence or absence of long-term synaptic plasticity in the interlamellar hippocampal CA1 network using electrophysiological field recordings both in vivo and in vitro. For in vivo extracellular field recordings, the recording and stimulation electrodes were placed in a septal-temporal axis of the dorsal hippocampus at a longitudinal angle, to evoke field excitatory postsynaptic potentials. For in vitro extracellular field recordings, hippocampal longitudinal slices were cut parallel to the septal-temporal plane. Recording and stimulation electrodes were placed in the stratum oriens (S.O) and the stratum radiatum (S.R) of the hippocampus along the longitudinal axis. This enabled us to investigate the directional and layer specificity of evoked excitatory postsynaptic potentials. Already established protocols were used to induce long-term potentiation (LTP) and long-term depression (LTD) both in vivo and in vitro. Our results demonstrated that the longitudinal interlamellar CA1 network supports N-methyl-D-aspartate (NMDA) receptor-dependent long-term potentiation (LTP) with no directional or layer specificity. The interlamellar network, however, in contrast to the transverse lamellar network, did not present with any significant long-term depression (LTD).

We used recording and stimulation electrodes in longitudinal hippocampal brain slices and longitudinally positioned recording and stimulation electrodes in the dorsal hippocampus in vivo to evoke extracellular postsynaptic potentials and demonstrate long-term synaptic plasticity along the longitudinal interlamellar CA1.