We thank the members of Matsunami laboratory for critical reading of this manuscript.

1. Procedure:
The procedure comprises of three parts completed over a total time of 3 days: transferring cells, transfection and immunocytochemistry, one step carried out per day. Transfer and transfection of cells over the first two days must be carried out in sterile conditions in a laminar flow chamber.
Day 1: Transfer of HEK293T cells for surface expression assay.
<ol>
<li>Transfer medium (M10): Mix minimum essential medium supplemented with 10% (by volume) fetal bovine serum in sterile condition.</li>
<li>Grow cells to a desired confluency in a 100mm culture plate, depending on the number of plates required to set up. It is important to determine the fraction of cells to be transferred to avoid overgrown or sparse cells: for instance, from a 100% confluent 100 mm dish, one may transfer about 3% to a 35 mm dish to obtain approximately 30% confluency in the latter.</li>
<li>Prior to transferring cells, in the sterile laminar flow chamber, coat 22 X 22 mm glass cover slips with sterile poly D lysine (1mg/ mL) and place one in each 35 mm cell culture dish, coated side up for plating cells (Figure 1). Allow the solution to dry for 5- 10 minutes. During this interval, the UV source in the laminar chamber may be turned on to sterilize the cover slips, if desired. Poly D lysine helps heterologous cells stick to the cover slips.</li>
<li>To transfer cells, aspirate out the existing media in the 100mm dish, wash by carefully adding 8 mL sterile PBS, aspirate PBS and trypsinize the cells using 3 mL 0.05% Trypsin-EDTA; when cells detach from the dish, block further enzymatic reaction by adding 5 mL M10. Triturate the cells and transfer requisite volume to a sterile conical tube; centrifuge at 200g for 5 minutes to pellet the cells. Aspirate out supernatant containing Trypsin-EDTA, leaving the pellet of cells intact. To transfer the cells to 35 mm dishes, add 1 mL M10 for each dish and triturate to dissociate the cells. Add 1 mL of the resuspended cells to each dish. Incubate cells overnight at 37&deg;C, 5% CO2. <img src="/files/ftp_upload/2405/2405fig1.jpg" alt="Figure 1" /> Figure 1. Coat 22 X 22 mm glass coverslips with poly D lysine and place one coated side up in each 35 mm dish.</li>
</ol>
Day 2: Transfection of HEK293T cells.
<ol>
<li>18-24 hours after transferring, cells are ready for transfection of the receptor to be assayed. Use receptor and RTP1S cloned in mammalian expression vector pCI, prepared from bacterial cultures using Qiagen Miniprep Kit following protocol described before4. Use receptors that are N terminally tagged with 20 amino acid long rhodopsin epitope to facilitate immunostaining.</li>
<li>Use 1000ng of receptor construct, 250 ng of RTP1S construct and 50 ng of green fluorescence protein each transfection. Perform transfection using Lipofectamine 2000, as per manufacturers manual.</li>
</ol>
Day 3: Immunocytochemistry to visualize cell surface stained receptors.
<ol>
<li>Prior to starting, prepare staining and washing solutions and cool them to 4&deg;C. Staining medium: Minimum Essential Medium, 45 mL; Fetal Bovine Serum, 5 mL; HEPES (1M), 500 &mu;L (final concentration 10mM); Sodium Azide (1.5M), 500 &mu;L (final concentration 15mM). Wash solution: HBSS, 500 mL; HEPES (1M), 5 mL (final concentration 10mM); Sodium Azide (1.5M), 5 mL (final concentration 15mM). Both solutions may be stored at 4&deg;C for reuse. Prepare antibody solutions by diluting required antibodies in the staining medium 100 fold.</li>
<li>Start immunochemistry for surface staining for odorant receptors about 24 hours post transfection. To perform live cell staining, the experimental set up must be cooled to 4&deg;C: fill a large contained with ice and pat the surface of ice to make it as even as possible, place a tray on the ice and spray it with 70% ethanol. Cut a piece of parafilm (smaller than the length of the tray) and affix it on the ethanol sprayed tray, to obtain a smooth surface. Transfer the glass cover slips containing transfected cells on the parafilm, one at a time, cell side up, using tweezers. Layer each cover slip with 100 &mu;L cold primary staining solution, extruding carefully from a corner of the slip (Figure 2). After layering all the cover slips with antibody solution, cover the tray to prevent drying up of the solution in air. Carry out primary incubation for 45-60 minutes.</li>
<li>After primary incubation, quickly transfer cover slips to the original 35mm culture dishes, containing the media. Aspirate the media and add 2 mL chilled wash solution gently from the edge of the dish to prevent losing cells from the cover slip. Aspirate the wash solution and repeat twice to complete three washes. At the end of the third wash, do not aspirate the wash solution out.</li>
<li>Prepare secondary antibody solution in the staining media and transfer the cover slips from the wash solution to a newly affixed parafilm layer in the same tray. Again gently extrude 100 &mu;L of antibody solution to layer each cover slip, from the corner like the primary incubation. Carry out secondary incubation for 30- 45 minutes and transfer back to the wash solution in the original 35 mm dishes. Once more, wash the cover slips three times as described.</li>
<li>Aspirate wash solution from last wash and add 2 mL 1% chilled paraformaldehyde. Fix cells in chilled 1% paraformaldehyde solution, on ice for 15 minutes.</li>
<li>Mount the cover slips on clean microscope glass slides using mounting reagent Mowiol, with the cell side down and cautiously exclude all air bubbles (Figure 3). Once Mowiol air dries, wash cover slips gently by adding distilled water on the surface and aspirating immediately. <img src="/files/ftp_upload/2405/2405fig2.jpg" alt="Figure 2" /> Figure 2. Layer each coverslip with 100 &mu;L cold primary staining solution. <img src="/files/ftp_upload/2405/2405fig3.jpg" alt="Figure 3" /> Figure 3. Mount the coverslips with the cell side down on clean glass microscope slides using Mowiol mounting reagent.</li>
</ol>
2. Representative Results:
Live cell staining enables one to visualize proteins on the surface of cells, on transfection of receptors with chaperones (in this case odorant receptor Olfr62, with Receptor Transporting Protein RTP1S) (Figure 4). Cell surface staining of receptors is characterized by punctate pattern of staining (Figure 4B, inset). Staining carried out inappropriately may lead to higher noise, making it difficult for the observer to distinguish the real surface staining from noise. <img src="/files/ftp_upload/2405/2405fig4.jpg" alt="Figure 4" /> Figure 4. Transfection of receptors with chaperones allows for the visualization of proteins on the surface of cells using live cell staining. Transfection of the odorant receptor Olfr62 alone (A) and transfection of Olfr62 with RTP1S chaperone (B). GFP expression levels serve as a transfection control (A' and B').

<ol>
	<li>Buck, L., Axel, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+multigene+family+may+encode+odorant+receptors:+a+molecular+basis+for+odor+recognition.">A novel multigene family may encode odorant receptors: a molecular basis for odor recognition.</a> Cell. 65, 175-187 (1991).</li><li>Malnic, B., Hirono, J., Sato, T., Buck, L. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Combinatorial+receptor+codes+for+odors.">Combinatorial receptor codes for odors.</a> Cell. 96, 713-723 (1999).</li><li>Saito, H., Kubota, M., Roberts, R. W., Chi, Q., Matsunami, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RTP+family+members+induce+functional+expression+of+mammalian+odorant+receptors.">RTP family members induce functional expression of mammalian odorant receptors.</a> Cell. 119, 679-691 (2004).</li><li>Zhuang, H., Matsunami, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluating+cell-surface+expression+and+measuring+activation+of+mammalian+odorant+receptors+in+heterologous+cells.">Evaluating cell-surface expression and measuring activation of mammalian odorant receptors in heterologous cells.</a> Nat Protoc. 3, 1402-1413 (2008).</li></ol>

No conflicts of interest declared.

Each step must be carried out carefully to ensure prominent and distinctive surface staining. The entire staining process must be carried out in cold (on ice), and the tray on which cover slips are laid must be cooled prior to use to ensure cells stay alive. Moreover fixation is done at the end of the surface staining process. This is in sharp contrast with internal staining where fixation precedes staining to permeabilize the cell membrane to primary and secondary antibodies. Time of exposure of cover slips to air must ...

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		<th>Catalogue Number</th>
		<th>Comment</th>
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<td>Cover slip 22 X 22 mm (#1)</td>
<td>&nbsp;</td>
<td>Thermo Scientific</td>
<td>72200-11</td>
<td>&nbsp;</td>
<tr>
<td>Fetal bovine serum</td>
<td>&nbsp;</td>
<td>Gibco</td>
<td>16000</td>
<td>&nbsp;</td>
<tr>
<td>Lipofectamine 2000</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>11668-019</td>
<td>&nbsp;</td>
<tr>
<td>Minimal essential medium</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>M4655</td>
<td>With L glutamate, Earle&#x2019;s salt and bicarbonate</td>
</tr>
<tr>
<td>Mowiol</td>
<td>&nbsp;</td>
<td>Calbiochem</td>
<td>475904</td>
<td>&nbsp;</td>
<tr>
<td>Paraformaldehyde</td>
<td>&nbsp;</td>
<td>EMS</td>
<td>19208</td>
<td>&nbsp;</td>
<tr>
<td>Hanks Balanced Salt Solution (HBSS) 1X </td>
<td>&nbsp;</td>
<td>Gibco </td>
<td>14025</td>
<td>With calcium chloride and magnesium chloride</td>
</tr>
<tr>
<td>HEPES Buffer Solution</td>
<td>&nbsp;</td>
<td>Gibco</td>
<td>15630</td>
<td>&nbsp;</td>
<tr>
<td>Phosphate Buffer Saline (PBS)</td>
<td>&nbsp;</td>
<td>Cellgro</td>
<td>21-040-CV</td>
<td>Without calcium and magnesium</td>
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<td>Poly D Lysine Hydrobromide</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P7280</td>
<td>&nbsp;</td>
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<td>Sodium Azide</td>
<td>&nbsp;</td>
<td>EM Science</td>
<td>SX0299-1</td>
<td>&nbsp;</td>
<tr>
<td>Tissue culture dish 35 X 10 mm</td>
<td>&nbsp;</td>
<td>Falcon</td>
<td>353801</td>
<td>&nbsp;</td>
<tr>
<td>Trypsin-EDTA </td>
<td>&nbsp;</td>
<td>Gibco</td>
<td>25300</td>
<td>0.05%</td>
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assaying surface expression of chemosensory receptors in heterologous cells

The vivid world of odors is recognized by the sense of olfaction. Olfaction in mice is mediated by a repertoire of about 1200 G Protein Coupled Receptors (GPCRs) 1 that are postulated to bind volatile odorant molecules and converting the extracellular signal into an intracellular signal by coupling with G protein G&alpha;olf. Binding of the odorants to the receptors is thought to follow a combinatorial rule, that is, one odorant may bind several receptors and one receptor may bind several odorants to varying degrees 2. Biochemical, signaling and ligand binding studies have been conveniently carried out for most GPCRs using heterologous cells. However use of heterologous cells for study of odorant receptors, was precluded for a long time since on transfection they failed to export to the surface. Saito et al have demonstrated single membrane pass Receptor Transporting Protein (RTP) family chaperones show enhanced expression in the olfactory sensory neurons and act as chaperones to traffic odorant receptors to the surface in heterologous cells, when co transfected together 3. To carry out biochemical assays for receptors using heterologous cells, one must first determine if the receptor shows robust surface expression in the cell line. This can be assayed by overexpressing the receptors with the chaperone RTP1S followed by live cell staining to fluorescently label the extracellular domain or a tag in the extracellular domain exclusively. Here we demonstrate a protocol to carry out live cell staining that can be used to detect odorant receptors on the surface of HEK293T cells conveniently. In addition, it may also be used to assay for surface expression of other chemosensory receptors or GPCRs.

Watch this Scientific Journal Video about Assaying Surface Expression of Chemosensory Receptors in Heterologous Cells at JoVE.com

Assaying Surface Expression of Chemosensory Receptors in Heterologous Cells

Here we demonstrate a protocol to carry out live cell staining that can be used to detect odorant receptors on the surface of HEK293T cells conveniently. In addition, it may also be used to assay for surface expression of other chemosensory receptors or GPCRs.

The vivid world of odors is recognized by the sense of olfaction. Olfaction in mice is mediated by a repertoire of about 1200 G Protein Coupled Receptors ...

assaying-surface-expression-chemosensory-receptors-heterologous

Research

JoVE Journal

Neuroscience

10.8K 조회수.  Duke University. 우리가 편리하게 HEK293T 세포의 표면에 odorant 수용체를 감지하는 데 사용할 수있는 라이브 세포 얼룩을 수행하는 프로토콜을 보여줍니다. 또한, 그것은 또한 다른 chemosensory 수용체 또는 GPCRs의 표면 표현 분석하는 데 사용할 수 있습니다.

비디오: Assaying Surface Expression of Chemosensory Receptors in Heterologous Cells

Assaying Locomotor Activity to Study Circadian Rhythms and Sleep Parameters in Drosophila

Most life forms exhibit daily rhythms in cellular, physiological and behavioral phenomena that are driven by endogenous circadian (&equiv;24 hr) pacemakers or clocks. Malfunctions in the human circadian system are associated with numerous diseases or disorders. Much progress towards our understanding of the mechanisms underlying circadian rhythms has emerged from genetic screens whereby an easily measured behavioral rhythm is used as a read-out of clock function. Studies using Drosophila have made seminal contributions to our understanding of the cellular and biochemical bases underlying circadian rhythms. The standard circadian behavioral read-out measured in Drosophila is locomotor activity. In general, the monitoring system involves specially designed devices that can measure the locomotor movement of Drosophila. These devices are housed in environmentally controlled incubators located in a darkroom and are based on using the interruption of a beam of infrared light to record the locomotor activity of individual flies contained inside small tubes. When measured over many days, Drosophila exhibit daily cycles of activity and inactivity, a behavioral rhythm that is governed by the animal's endogenous circadian system. The overall procedure has been simplified with the advent of commercially available locomotor activity monitoring devices and the development of software programs for data analysis. We use the system from Trikinetics Inc., which is the procedure described here and is currently the most popular system used worldwide. More recently, the same monitoring devices have been used to study sleep behavior in Drosophila. Because the daily wake-sleep cycles of many flies can be measured simultaneously and only 1 to 2 weeks worth of continuous locomotor activity data is usually sufficient, this system is ideal for large-scale screens to identify Drosophila manifesting altered circadian or sleep properties.

Assaying Locomotor Activity to Study Circadian Rhythms and Sleep Parameters in Drosophila

We describe procedures for recording daily locomotor activity rhythms of Drosophila and subsequent data analysis. Locomotor activity rhythms are a reliable behavioral output of animal circadian clocks and are used as the standard readout of clock function when studying circadian mutants or examining how the environment regulates the circadian system.

Most life forms exhibit daily rhythms in cellular, physiological and behavioral phenomena that are driven by endogenous circadian (&equiv;24 hr) ...

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

Advances in genetic methods have enabled the study of genes involved in human neurodegenerative diseases using Drosophila as a model system1. Most of these diseases, including Alzheimer's, Parkinson's and Huntington's disease are characterized by age-dependent deterioration in learning and memory functions and movement coordination2. Here we use behavioral assays, including the negative geotaxis assay3 and the aversive phototaxic suppression assay (APS assay)4,5, to show that some of the behavior characteristics associated with human neurodegeneration can be recapitulated in flies. In the negative geotaxis assay, the natural tendency of flies to move against gravity when agitated is utilized to study genes or conditions that may hinder locomotor capacities. In the APS assay, the learning and memory functions are tested in positively-phototactic flies trained to associate light with aversive bitter taste and hence avoid this otherwise natural tendency to move toward light. Testing these trained flies 6 hours post-training is used to assess memory functions. Using these assays, the contribution of any genetic or environmental factors toward developing neurodegeneration can be easily studied in flies.

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

Behavior assays for measuring locomotor functions, learning, and memory abilities in Drosophila.

Advances in genetic methods have enabled the study of genes involved in human neurodegenerative diseases using Drosophila as a model system1. Most of ...

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

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 ...

Isolating Nasal Olfactory Stem Cells from Rodents or Humans

The olfactory mucosa, located in the nasal cavity, is in charge of detecting odours. It is also the only nervous tissue that is exposed to the external environment and easily accessible in every living individual. As a result, this tissue is unique for anyone aiming to identify molecular anomalies in the pathological brain or isolate adult stem cells for cell therapy. 
Molecular abnormalities in brain diseases are often studied using nervous tissue samples collected post-mortem. However, this material has numerous limitations. In contrast, the olfactory mucosa is readily accessible and can be biopsied safely without any loss of sense of smell1. Accordingly, the olfactory mucosa provides an "open window" in the adult human through which one can study developmental (e.g. autism, schizophrenia)2-4 or neurodegenerative (e.g. Parkinson, Alzheimer) diseases4,5. Olfactory mucosa can be used for either comparative molecular studies4,6 or in vitro experiments on neurogenesis3,7.
The olfactory epithelium is also a nervous tissue that produces new neurons every day to replace those that are damaged by pollution, bacterial of viral infections. This permanent neurogenesis is sustained by progenitors but also stem cells residing within both compartments of the mucosa, namely the neuroepithelium and the underlying lamina propria8-10. We recently developed a method to purify the adult stem cells located in the lamina propria and, after having demonstrated that they are closely related to bone marrow mesenchymal stem cells (BM-MSC), we named them olfactory ecto-mesenchymal stem cells (OE-MSC)11.
Interestingly, when compared to BM-MSCs, OE-MSCs display a high proliferation rate, an elevated clonogenicity and an inclination to differentiate into neural cells. We took advantage of these characteristics to perform studies dedicated to unveil new candidate genes in schizophrenia and Parkinson's disease4. We and others have also shown that OE-MSCs are promising candidates for cell therapy, after a spinal cord trauma12,13, a cochlear damage14 or in an animal models of Parkinson's disease15 or amnesia16.
In this study, we present methods to biopsy olfactory mucosa in rats and humans. After collection, the lamina propria is enzymatically separated from the epithelium and stem cells are purified using an enzymatic or a non-enzymatic method. Purified olfactory stem cells can then be either grown in large numbers and banked in liquid nitrogen or induced to form spheres or differentiated into neural cells. These stem cells can also be used for comparative omics (genomic, transcriptomic, epigenomic, proteomic) studies.

We describe here a method for biopsying olfactory mucosa from rat and human nasal cavities. These biopsies can be used for either identifying molecular anomalies in brain diseases or isolating multipotent adult stem cells that can be utilized for cell transplantation in animal models of brain trauma/disease.

The olfactory mucosa, located in the nasal cavity, is in charge of detecting odours. It is also the only nervous tissue that is exposed to the external ...

High-resolution Live Imaging of Cell Behavior in the Developing Neuroepithelium

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central nervous system (CNS). Although much is known about the molecular mechanisms that pattern the spinal cord and elicit neuronal differentiation1, 2, we lack a deep understanding of these early events at the level of cell behavior. It is thus critical to study the behavior of neural progenitors in real time as they undergo neurogenesis.
In the past, real-time imaging of early embryonic tissue has been limited by cell/tissue viability in culture as well as the phototoxic effects of fluorescent imaging. Here we present a novel assay for imaging such tissue for long periods of time, utilizing a novel ex vivo slice culture protocol and wide-field fluorescence microscopy (Fig. 1). This approach achieves long-term time-lapse monitoring of chick embryonic spinal cord progenitor cells with high spatial and temporal resolution.
This assay may be modified to image a range of embryonic tissues3, 4 In addition to the observation of cellular and sub-cellular behaviors, the development of novel and highly sensitive reporters for gene activity (for example, Notch signaling5) makes this assay a powerful tool with which to understand how signaling regulates cell behavior during embryonic development.

Imaging embryonic tissue in real-time is challenging over long periods of time. Here we present an assay for monitoring cellular and sub-cellular changes in chick spinal cord for long periods with high spatial and temporal resolution. This technique can be adapted for other regions of the nervous system and developing embryo.

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central ...

Assaying DNA Damage in Hippocampal Neurons Using the Comet Assay

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been extensively evaluated. Traditional methods are time consuming, expensive, resource intensive and require replicating cells. In contrast, the comet assay, a single cell gel electrophoresis assay, is a faster, non-invasive, inexpensive, direct and sensitive measure of DNA damage and repair. All forms of DNA damage as well as DNA repair can be visualized at the single cell level using this powerful technique.
	The principle underlying the comet assay is that intact DNA is highly ordered whereas DNA damage disrupts this organization. The damaged DNA seeps into the agarose matrix and when subjected to an electric field, the negatively charged DNA migrates towards the cathode which is positively charged. The large undamaged DNA strands are not able to migrate far from the nucleus. DNA damage creates smaller DNA fragments which travel farther than the intact DNA. Comet Assay, an image analysis software, measures and compares the overall fluorescent intensity of the DNA in the nucleus with DNA that has migrated out of the nucleus. Fluorescent signal from the migrated DNA is proportional to DNA damage. Longer brighter DNA tail signifies increased DNA damage. Some of the parameters that are measured are tail moment which is a measure of both the amount of DNA and distribution of DNA in the tail, tail length and percentage of DNA in the tail. This assay allows to measure DNA repair as well since resolution of DNA damage signifies repair has taken place. The limit of sensitivity is approximately 50 strand breaks per diploid mammalian cell 1,2. Cells treated with any DNA damaging agents, such as etoposide, may be used as a positive control. Thus the comet assay is a quick and effective procedure to measure DNA damage.

The comet assay is an efficient way of detecting single- and double-strand breaks, including alkali-labile sites and DNA-DNA/DNA-protein cross-links on the DNA in all cells including hippocampal neurons. The method takes advantage of the differential migration of DNA in an electric field due to differences in amount of DNA damage.

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been ...

Proprioception and Tension Receptors in Crab Limbs: Student Laboratory Exercises

The primary purpose of these procedures is to demonstrate for teaching and research purposes how to record the activity of living primary sensory neurons responsible for proprioception as they are detecting joint position and movement, and muscle tension. Electrical activity from crustacean proprioceptors and tension receptors is recorded by basic neurophysiological instrumentation, and a transducer is used to simultaneously measure force that is generated by stimulating a motor nerve. In addition, we demonstrate how to stain the neurons for a quick assessment of their anatomical arrangement or for permanent fixation. Staining reveals anatomical organization that is representative of chordotonal organs in most crustaceans. Comparing the tension nerve responses to the proprioceptive responses is an effective teaching tool in determining how these sensory neurons are defined functionally and how the anatomy is correlated to the function. Three staining techniques are presented allowing researchers and instructors to choose a method that is ideal for their laboratory.

Physiological and anatomical techniques are demonstrated to address function and structure for joint proprioceptors and muscle tension receptors in crustacean walking limbs.

The primary purpose of these procedures is to demonstrate for teaching and research purposes how to record the activity of living primary sensory neurons ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

Determining Cell-surface Expression and Endocytic Rate of Proteins in Primary Astrocyte Cultures Using Biotinylation

Cell-surface proteins mediate a wide array of functions. In many cases, their activity is regulated by endocytic processes that modulate their levels at the plasma membrane. Here, we present detailed protocols for 2 methods that facilitate the study of such processes, both of which are based on the principle of the biotinylation of cell-surface proteins. The first is designed to allow for the semi-quantitative determination of the relative levels of a particular protein at the cell-surface. In it, the lysine residues of the plasma membrane proteins of cells are first labeled with a biotin moiety. Once the cells are lysed, these proteins may then be specifically precipitated via the use of agarose-immobilized streptavidin by exploiting the natural affinity of the latter for biotin. The proteins isolated in such a manner may then be analyzed via a standard western blotting approach. The second method provides a means of determining the endocytic rate of a particular cell-surface target over a period of time. Cell-surface proteins are first modified with a biotin derivative containing a cleavable disulfide bond. The cells are then shifted back to normal culture conditions, which causes the endocytic uptake of a proportion of biotinylated proteins. Next, the disulfide bonds of non-internalized biotin groups are reduced using the membrane-impermeable reducing agent glutathione. Via this approach, endocytosed proteins may thus be isolated and quantified with a high degree of specificity.

Two biotinylation-based methods, designed for determining the cell-surface expression and endocytic rate of proteins expressed at the plasma membrane, are presented in this report.

Cell-surface proteins mediate a wide array of functions. In many cases, their activity is regulated by endocytic processes that modulate their levels at ...