Name: dopamine release at individual presynaptic terminals visualized with ffns
Uploaded: 2009-08-31T11:00:00
Duration: 9 min 37 s
Description: Watch this Scientific Journal Video about Dopamine Release at Individual Presynaptic Terminals Visualized with FFNs at JoVE.com

D. Sames thanks The G. Harold &amp; Leila Y. Mathers Charitable Foundation and Columbia University's Initiatives in Science and Engineering. D. Sames and D. Sulzer thank the McKnight Foundation for The McKnight Technological Innovations in Neuroscience Award D. Sulzer thanks NIDA, NIMH, and the Picower and Parkinson's Disease Foundations. H. Zhang thanks NARSAD. R.H. Edwards thanks the Michael J. Fox Foundation, the National Parkinson
Foundation, NIDA and NIMH. We thank Robert Burke for 6-OHDA injections and advice, Mark Sonders for useful discussion, Merek Siu for imaging analysis programming, and Jan Schmoranzer for technical support with the TIRF microscopy setup.

This method was used in the research reported in <a href="http://www.sciencemag.org/cgi/content/full/324/5933/1441">Gubernator et al., Science 324 (5933). 1441-1444 (2009)</a>.
1. Preparing acute striatal slices
<ol>
<li>Before preparing acute striatal slices, one must oxygenate the artificial cerebrospinal fluid (ASCF) and chill it with ice for at least 15 minutes prior to the brain extraction. Keep the ACSF ice cold and oxygenated during the dissection. ACSF (in mM): NaCl 125, KCl 2.5, NaHCO3 26, CaCl2 2.4, MgSO4 1.3, KH2PO4 0.3, glucose 10, HEPES 5; pH 7.3-7.4, 290-295 mOsm.</li>
<li>Decapitate male mouse without anesthesia.</li>
<li>Extract the whole brain from the mouse, and mount it on the vibratome tray placed on the vibratome using Krazy&reg;Glue. Brain extraction and mounting must be done within 2 minutes. During this time, make sure to keep the ACSF ice cold and oxygenated to keep the tissue healthy.</li>
<li>Cut coronal striatal brain slices at 250&mu;m thickness between bregma +1.54 to +0.621. Three whole slices will be obtained which can then be cut into 6 half slices with a needle.</li>
<li>Incubate the brain slices in oxygenated ACSF at room temperature for at least 1 hour before probe loading. Slices display good FFN511 loading and remain active for imaging up to 4 hours after slice preparation.</li>
</ol>
2. Loading FFN511
<ol>
<li>Prepare FFN511 loading solution (10&mu;M FFN511 in ACSF); also prepare 100&mu;M ADVASEP-7 in ACSF. All solutions should be prepared freshly and oxygenated at least 15 minutes prior to loading into the slices.</li>
<li>Individually incubate slice with FFN511 loading solution for 30 minutes at room temperature.</li>
<li>Then remove the dye bound to extracellular tissue by incubating the loaded slice in oxygenated 100&mu;M ADVASEP-7 ACSF for 30 minutes at room temperature2. Slices are now ready for imaging.</li>
</ol> 
3. Imaging FFN511 in the brain slices
Imaging of FFN511 in the slices is performed using multiphoton laser scanning microscope. We are using a Prairie Ultima multiphoton laser scanning microscope (We were using a Zeiss LSM 510 NLO multiphoton laser scanning microscope previously and some results were obtained with the Zeiss microscope).
<ol>
<li>Place the FFN511-loaded slice in the recording chamber (RC-27L, Warner) and superfuse with oxygenated ACSF at a flow rate of 1-2 ml per minute. Then place a platinum harp with nylon strings on top of the slice to minimize movement during the experiment. To further minimize movement, allow the slice to stabilize for at least 10 minutes in the chamber prior to imaging.</li>
<li>Visualize and locate the dorsal striatum with bright field luminescence under a 10x water immersion objective.</li>
<li>Place and position the bipolar twisted tungsten electrodes in this region if performing a stimulation experiment. The ideal imaging area is located within 300 &mu;m in between the two tips of the stimulating bipolar electrode. Place this region in the center of the visual field.</li>
<li>Image FFN511-labeled striatal terminals under a 63x (0.9 NA) water immersion ultraviolet objective. FFN511 is excited at 760 nm with a Mai Tai laser from Spectral Physics and optimal fluorescence of striatal terminals is seen through a band pass filter (480-520 nm). Images are captured in 12-bit format with 75 x 75 &mu;m regions of interest at 512 x 512 pixel resolution. For stimulated destaining experiments, to compensate for z-axis shift, a z-series of 5-7 images, separated by 1 &mu;m in the z-plane, is obtained for each time period.</li>
</ol>
4. FFN511 Destaining
The FFN511 label can be destained either by high concentration of KCl (we use 70 mM KCl), (+)-amphetamine sulphate (AMPH, 20 &mu;M), or electrical stimulation. In the KCl destaining experiment, when a high potassium ACSF solution is applied to the brain slice, the FFN511 label destains within 2 minutes. Record xyz-t images to track FFN511 destaining over time. The following describes electrical stimulation-dependent dopamine terminal destaining:
<ol>
<li>To track FFN511 destaining over time, xyz-t images are recorded. To ensure minimal movement of the slice (less than 4 &mu;m in z-axis) and no photobleaching by the laser, control time series images of the z-stack should be obtained for at least 5 minutes prior to stimulation. Otherwise, adjust the laser power.</li>
<li>Following these control images, continue the xyz-t imaging while start the stimulation at frequencies of 1, 4 or 20 Hz. Stimuli at 1, 4 or 20 Hz (300 &mu;s x 1 mA) are applied to the striatum locally by an Iso-Flex stimulus isolator triggered by a Master-8 pulse generator (AMPI, Jerusalem, Israel) using bipolar electrodes. To minimize the variation in the depolarization of release sites, we have used a stimulation protocol applying 150% of the maximal stimulation intensity determined by cyclic voltammetry.</li>
<li>Use proper software for imaging analysis. In our lab, we use Image J (Wayne Rosband, National Institutes of Health, Rockville, MD) and custom-written software in IDL (Research Systems, Boulder, CO) to quantify the puncta fluorescence and the changes with time3.</li>
</ol>
Represenative Results
Figure 1 shows FFN511 loading into striatal slice is now complete. Representative images using 10x and 60x objective are shown in Fig.1. The selective labeling of dopamine terminals can be destained by 20 &mu;M amphetamine. Figure 2 shows destaining of FFN511 labeling by high KCl. Figure 3 shows
destaining of FFN511 labeling by local electrical stimulation. 
<img src="/files/ftp_upload/1562/1562fig1.jpg" alt="Figure 1" /> Figure 1. FFN511 labels dopamine terminals in live cortical-striatal acute slices. (A) Labeling by FFN511 in acute live cortical-striatal slice: abundant labeling in the striatum (STR), sparser labeling in cortex (CTX), and no label in corpus callosum (CC). Scale bar: 100 &mu;m. (B) Destaining of FFN511 from the striatum by amphetamine. Left panels: before amphetamine; right panels: after 20 minutes of 20 &mu;M amphetamine. Scale bar: 10 &mu;m.
<img src="/files/ftp_upload/1562/1562fig2.jpg" alt="Figure 2" /> Figure 2. Destaining of FFN511 labeling by high KCl. FFN511 labeling is destained within 2 min application of 70 mM KCl in ACSF. Scale bar: 10&mu;m.
<img src="/files/ftp_upload/1562/1562fig3.jpg" alt="Figure 3" /> Figure 3. 
Frequency-dependent destaining of FFN511 labeling in striatum. (A) Local stimulation at 4 Hz resulted in destaining from the terminals. Stimulation began at t = 0. Scale bar: 5 &mu;m. (B) Destaining of FFN511 at 4 Hz is Ca2+- and frequency-dependent. Controls received no stimulation (153 puncta from 3 slices). Destaining with cadmium chloride (200 &mu;M) was identical to unstimulated controls (475 puncta from 5 slices). The destaining curves for each stimulation frequency were fit by a single exponential decay function and half-life (t1/2) values calculated as &tau; x 0.693 (1 Hz: 765 puncta from 9 slices, 4 Hz: 410 puncta from 7 slices, 20 Hz: 416 puncta from 6 slices). Please refer to the video micrograph from 2-photon microscopy of FFN511 destaining during exocytosis in a striatal brain slice preparation.

<ol>
	<li>Franklin, K. B. J., Paxinos, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The mouse brain in stereotaxi coordinators. , (1997).</li><li>Kay, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+synaptic+activity+in+intact+brain+and+slices+with+FM1-43+in+C.+elegans,+lamprey,+and+rat.">Imaging synaptic activity in intact brain and slices with FM1-43 in C. elegans, lamprey, and rat.</a> Neuron. 24, 8098-8117 (1999).</li><li>Bamford, N. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heterosynaptic+dopamine+neurotransmission+selects+sets+of+corticostriatal+terminals.">Heterosynaptic dopamine neurotransmission selects sets of corticostriatal terminals.</a> Neuron. 42, 653-653 (2004).</li></ol>

The authors have nothing to disclose.

In this video, we demonstrate a method to visualize neurotransmission optically using fluorescent dopamine analogs. FFN511 is the first generation of FFNs we have developed. Although it was designed by targeting the neuronal vesicular monoamine transporter (VMAT2) that carries monoamine neurotransmitters from the cytoplasm into synaptic vesicles, and specifically labels dopamine terminals in the striatum and presumed catecholamine and/or serotonin terminals in the cortex (as shown in the Science paper), an appropriate loading period is critical for the specificity since FFN511 is relatively hydrophobic. We found that incubation for longer than 40 minutes will result in extensive nonspecific staining in striatal slices. The specificity can be straightforwardly determined by destaining using a high concentration of KCl, which should cause the release of FFN within functional synaptic vesicles. Exposure of the slice to FFN511 for less than 15 minutes will result in weak fluorescent signal due to insufficient loading of the dye in the terminals. In this protocol, we use 100&mu;M ADVASEP-7 to remove the dye bound to extracellular tissue. This step is not necessary, but if it is omitted, the washout time in ACSF needs to be prolonged. In summary, depending on the preparation you chose, you will need to modify the concentration and loading period to determine optimal labeling.

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>FFN511 (8-(2-Amino-ethyl)-2,3,5,6-tetrahydro-1H,4H-11-oxa-3a-aza-benzo[de]anthracen-10-one)</td>
<td>&nbsp;</td>
<td>Dalibor Sames&#x2019;s laboratory at Columbia University</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>ADVASEP-7 </td>
<td>&nbsp;</td>
<td>CyDex, Overland Park, KS</td>
<td>AR-OA7-005</td>
<td>&nbsp;</td>
<tr>
<td>RC-27L Recording chamber</td>
<td>&nbsp;</td>
<td>Warner Instrument</td>
<td>64-0375</td>
<td>&nbsp;</td>
<tr>
<td>PELCO PrepEze 6-Well Holder</td>
<td>&nbsp;</td>
<td>Ted Pella, Inc.</td>
<td>36157-1</td>
<td>For slice incubation</td>
</tr>
<tr>
<td>Master-8 pulse stimulator and Iso- Flex stimulus isolator</td>
<td>&nbsp;</td>
<td>AMPI, Jerusalem, Israel</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
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dopamine release at individual presynaptic terminals visualized with ffns

The nervous system transmits signals between neurons via neurotransmitter release during synaptic vesicle fusion. To observe neurotransmitter uptake and release from individual presynaptic terminals directly, we designed fluorescent false neurotransmitters as substrates for the synaptic vesicle monoamine transporter. Using these probes to image dopamine release in the striatum, we made several observations pertinent to synaptic plasticity. We found that the fraction of synaptic vesicles releasing neurotransmitter per stimulus was dependent on the stimulus frequency. A kinetically distinct "reserve" synaptic vesicle population was not observed under these experimental conditions. A frequency-dependent heterogeneity of presynaptic terminals was revealed that was dependent in part on D2 dopamine receptors, indicating a mechanism for frequency-dependent coding of presynaptic selection.

Hui Zhang and Niko G. Gubernator contributed equally to this work.

Watch this Scientific Journal Video about Dopamine Release at Individual Presynaptic Terminals Visualized with FFNs at JoVE.com

Dopamine Release at Individual Presynaptic Terminals Visualized with FFNs

A new means to measure neurotransmission optically using fluorescent dopamine analogs.

The nervous system transmits signals between neurons via neurotransmitter release during synaptic vesicle fusion. To observe neurotransmitter uptake and ...

dopamine-release-at-individual-presynaptic-terminals-visualized-with

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Biology

Loading Drosophila Nerve Terminals with Calcium Indicators

Calcium plays many roles in the nervous system but none more impressive than as the trigger for neurotransmitter release, and none more profound than as the messenger essential for the synaptic plasticity that supports learning and memory. To further elucidate the molecular underpinnings of Ca2+-dependent synaptic mechanisms, a model system is required that is both genetically malleable and physiologically accessible. Drosophila melanogaster provides such a model. In this system, genetically-encoded fluorescent indicators are available to detect Ca2+ changes in nerve terminals. However, these indicators have limited sensitivity to Ca2+ and often show a non-linear response. Synthetic fluorescent indicators are better suited for measuring the rapid Ca2+ changes associated with nerve activity. Here we demonstrate a technique for loading dextran-conjugated synthetic Ca2+ indicators into live nerve terminals in Drosophila larvae. Particular emphasis is placed on those aspects of the protocol most critical to the technique's success, such as how to avoid static electricity discharges along the isolated nerves, maintaining the health of the preparation during extended loading periods, and ensuring axon survival by providing Ca2+ to promote sealing of severed axon endings. Low affinity dextran-conjugated Ca2+-indicators, such as fluo-4 and rhod, are available which show a high signal-to-noise ratio while minimally disrupting presynaptic Ca2+ dynamics. Dextran-conjugation helps prevent Ca2+ indicators being sequestered into organelles such as mitochondria. The loading technique can be applied equally to larvae, embryos and adults.

Calcium is a ubiquitous messenger in the nervous system, essential for triggering neurotransmitter release and changes in synaptic strength. Here we demonstrate a technique for loading Ca2+-indicators into Drosophila nerve terminals. We also demonstrate fabrication of the required apparatus and emphasize points critical for the technique's success.

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Labeling hESCs and hMSCs with Iron Oxide Nanoparticles for Non-Invasive in vivo Tracking with MR Imaging

In recent years, stem cell research has led to a better understanding of developmental biology, various diseases and its potential impact on regenerative medicine. A non-invasive method to monitor the transplanted stem cells repeatedly in vivo would greatly enhance our ability to understand the mechanisms that control stem cell death and identify trophic factors and signaling pathways that improve stem cell engraftment. MR imaging has been proven to be an effective tool for the in vivo depiction of stem cells with near microscopic anatomical resolution. In order to detect stem cells with MR, the cells have to be labeled with cell specific MR contrast agents. For this purpose, iron oxide nanoparticles, such as superparamagnetic iron oxide particles (SPIO), are applied, because of their high sensitivity for cell detection and their excellent biocompatibility. SPIO particles are composed of an iron oxide core and a dextran, carboxydextran or starch coat, and function by creating local field inhomogeneities, that cause a decreased signal on T2-weighted MR images. This presentation will demonstrate techniques for labeling of stem cells with clinically applicable MR contrast agents for subsequent non-invasive in vivo tracking of the labeled cells with MR imaging.

For the evaluation of new stem cell therapies it is important to non-invasively track the injected cells in vivo. This video will show you how to label human mesenchymal and embryonic stem cells with iron oxide based contrast agents in vivo for subsequent MR imaging in vivo.

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Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

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Primary Dissociated Midbrain Dopamine Cell Cultures from Rodent Neonates

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The process involves several steps, including the plating of cortical glial astrocytes, the conditioning of neuronal cell culture media by the glial substrate, the dissection of the midbrain in neonates, the digestion, extraction and plating of dopamine neurons and the addition of neurotrophic factors to ensure cell survival.
The applications suitable for such a preparation include electrophysiology, immunocytochemistry, quantitative PCR, video microscopy (i.e., of real-time vesicular fusion with the plasma membrane), cell viability assays and other toxicological screens.

Primary dissociated midbrain dopamine cell cultures allow for the study of presynaptic characteristics of dopamine neurons. They can be used to monitor real-time dopamine release kinetics and protein/mRNA levels of regulators of dopamine exocytosis. Here, we show you how to generate these cultures from rodent neonates.

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Murine Model for Parkinson's Disease: from 6-OH Dopamine Lesion to Behavioral Test

Parkinson's disease (PD) affects at least 6.5 million people worldwide, irrespective of gender, social, ethnic, economic, or geographic boundaries. Key symptoms, such as tremor, rigidity and bradikinesia, develop when about 3/4 of dopaminergic cells are lost in the substantia nigra, and fail to provide for the smooth, coordinated regulation of striatal motor circuits. Depression and hallucinations are common, and dementia eventually occurs in 20% of patients. At this time, there is no treatment to delay or stop the progression of PD. Rather, the medications currently available aim more towards the alleviation of these symptoms. New surgical strategies may reversibly switch on the functionally damaged circuits through the electrical stimulation of deep brain structures, but although deep brain stimulation is a major advance, it is not suitable for all patients. It remains therefore necessary to test new cell therapy approaches in preclinical models.
Selective neurotoxic disruption of dopaminergic pathways can be reproduced by injection of 6-hydroxydopamine (6-OHDA) or MPTP (1-methyl-4-phenyl-1,2,3,6-tertahydropyridine) whereas depleting drugs and oxidative-damaging chemicals may also reproduce specific features of PD in rodents. Unlike MPTP, 6-OHDA lesions cause massive irreversible neuronal loss, and can be uni- or bilateral. The 6-OHDA lesion model is reliable, leads to robust motor deficits, and is the most widely used after 40 years of research in rats1. As interactions between grafted cells and host can now be studied more thoroughly in mice rather than in rats, the model has been transposed to mice2,3, where it has been recently characterized4.
In this video, we demonstrate how to lesion the left nigro-striatal pathway of anesthetized mice by slowly delivering 2.0 &mu;L of 6-OHDA through a stereotaxically inserted micro-syringe needle. The loss of dopaminergic input occurs within days, and the functional impairments can be monitored over post-operative weeks and months by rating animal rotations induced by dopaminergic agents5. Here, we show full-body contralateral rotations occurring 10 minutes after a single subcutaneous administration of apomorphine, measured one month after the lesion. Outcomes and drawbacks are discussed below.

Murine Model for Parkinson&#039;s Disease: from 6-OH Dopamine Lesion to Behavioral Test

Parkinson disease is caused by loss of dopaminergic innervation to the striatum, which can be experimentally induced by 6-OH-dopamine. We describe how to perform a stereotaxic lesion and to monitor apomorphine-induced rotational behavior in mice. This model is useful and reliable for testing new therapies for Parkinson disease.

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Cell Electrofusion Visualized with Fluorescence Microscopy

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves application of short high-voltage electric pulses to cells that are in close contact. Application of short, high-voltage electric pulses causes destabilization of cell plasma membranes. Destabilized membranes are more permeable for different molecules and also prone to fusion with any neighboring destabilized membranes. Electrofusion is thus a convenient method to achieve a non-specific fusion of very different cells in vitro. In order to obtain fusion, cell membranes, destabilized by electric field, must be in a close contact to allow merging of their lipid bilayers and consequently their cytoplasm. In this video, we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method. In this method, cells are allowed to attach only slightly to the surface of the well, so that medium can be exchanged and cells still preserve their spherical shape. Fusion visualization is assessed by pre-labeling of the cytoplasm of cells with different fluorescent cell tracker dyes; half of the cells are labeled with orange CMRA and the other half with green CMFDA. Fusion yield is determined as the number of dually fluorescent cells divided with the number of all cells multiplied by two.

In this video we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method using electroporation and the subsequent detection of fused cells visualization with fluorescence microscopy.

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves ...

iCLIP - Transcriptome-wide Mapping of Protein-RNA Interactions with Individual Nucleotide Resolution

The unique composition and spatial arrangement of RNA-binding proteins (RBPs) on a transcript guide the diverse aspects of post-transcriptional regulation1. Therefore, an essential step towards understanding transcript regulation at the molecular level is to gain positional information on the binding sites of RBPs2.
Protein-RNA interactions can be studied using biochemical methods, but these approaches do not address RNA binding in its native cellular context. Initial attempts to study protein-RNA complexes in their cellular environment employed affinity purification or immunoprecipitation combined with differential display or microarray analysis (RIP-CHIP)3-5. These approaches were prone to identifying indirect or non-physiological interactions6. In order to increase the specificity and positional resolution, a strategy referred to as CLIP (UV cross-linking and immunoprecipitation) was introduced7,8. CLIP combines UV cross-linking of proteins and RNA molecules with rigorous purification schemes including denaturing polyacrylamide gel electrophoresis. In combination with high-throughput sequencing technologies, CLIP has proven as a powerful tool to study protein-RNA interactions on a genome-wide scale (referred to as HITS-CLIP or CLIP-seq)9,10. Recently, PAR-CLIP was introduced that uses photoreactive ribonucleoside analogs for cross-linking11,12.
Despite the high specificity of the obtained data, CLIP experiments often generate cDNA libraries of limited sequence complexity. This is partly due to the restricted amount of co-purified RNA and the two inefficient RNA ligation reactions required for library preparation. In addition, primer extension assays indicated that many cDNAs truncate prematurely at the crosslinked nucleotide13. Such truncated cDNAs are lost during the standard CLIP library preparation protocol. We recently developed iCLIP (individual-nucleotide resolution CLIP), which captures the truncated cDNAs by replacing one of the inefficient intermolecular RNA ligation steps with a more efficient intramolecular cDNA circularization (Figure 1)14. Importantly, sequencing the truncated cDNAs provides insights into the position of the cross-link site at nucleotide resolution. We successfully applied iCLIP to study hnRNP C particle organization on a genome-wide scale and assess its role in splicing regulation14.

The spatial arrangement of RNA-binding proteins on a transcript is a key determinant of post-transcriptional regulation. Therefore, we developed individual-nucleotide resolution UV crosslinking and immunoprecipitation (iCLIP) that allows precise genome-wide mapping of the binding sites of an RNA-binding protein.

The unique composition and spatial arrangement of RNA-binding proteins (RBPs) on a transcript guide the diverse aspects of post-transcriptional ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

Protein-protein Interactions Visualized by Bimolecular Fluorescence Complementation in Tobacco Protoplasts and Leaves

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular fluorescence complementation (BiFC) is an in vivo method to monitor such interactions in plant cells. In the presented protocol the investigated candidate proteins are fused to complementary halves of fluorescent proteins and the respective constructs are introduced into plant cells via agrobacterium-mediated transformation. Subsequently, the proteins are transiently expressed in tobacco leaves and the restored fluorescent signals can be detected with a confocal laser scanning microscope in the intact cells. This allows not only visualization of the interaction itself, but also the subcellular localization of the protein complexes can be determined. For this purpose, marker genes containing a fluorescent tag can be coexpressed along with the BiFC constructs, thus visualizing cellular structures such as the endoplasmic reticulum, mitochondria, the Golgi apparatus or the plasma membrane. The fluorescent signal can be monitored either directly in epidermal leaf cells or in single protoplasts, which can be easily isolated from the transformed tobacco leaves. BiFC is ideally suited to study protein-protein interactions in their natural surroundings within the living cell. However, it has to be considered that the expression has to be driven by strong promoters and that the interaction partners are modified due to fusion of the relatively large fluorescence tags, which might interfere with the interaction mechanism. Nevertheless, BiFC is an excellent complementary approach to other commonly applied methods investigating protein-protein interactions, such as coimmunoprecipitation, in vitro pull-down assays or yeast-two-hybrid experiments.

Formation of protein complexes in vivo can be visualized by bimolecular fluorescence complementation. Interaction partners are fused to complementary parts of fluorescent tags and transiently expressed in tobacco leaves, resulting in a reconstituted fluorescent signal upon close proximity of the two proteins.

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular ...

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane. Here a summary of the FIONA technique is reported and examples of research that have been performed using FIONA are briefly described. First, how to set up the required equipment for FIONA experiments, i.e., a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, followed by the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot, is illustrated. Lastly, recent effort to extend the application of FIONA to thick samples is reported. It is shown that, using a water immersion objective and quantum dots soaked deep in sol-gels and rabbit eye corneas (&#62;200 &#181;m), localization precision of 2-3 nm can be achieved.

Fluorescence Imaging with One-nanometer Accuracy FIONA

Single fluorophores can be localized with nanometer precision using FIONA. Here a summary of the FIONA technique is reported, and how to carry out FIONA experiments is described.

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in ...