The work was supported by the Howard Hughes Medical Institute (LK and ASA) and the Helen Hay Whitney Foundation (LHG). 
 
Original data underlying this manuscript can be accessed from the Stowers Original Data Repository at http://www.stowers.org/research/publications/libpb-1628

1. Preparation
<ol>
	<li>Prepare worms: feed sexual planarians twice a week with organic liver paste to achieve sexual maturity. 
	NOTE: Generally, such worms are bigger than 1 cm in length and have a gonopore posterior to the pharynx opening.</li>
	<li>Prepare solutions.
	<ol>
		<li>Prepare the following reagents: 16% paraformaldehyde (PFA); 50% glutaraldehyde (GA) aqueous solution; N-acetyl-L-cysteine (NAC); 1x phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, and 2 mM KH2PO4.</li>
		<li>Dilute 50% GA and 16% PFA with PBS to a final mixture of 2.5% GA and 2% PFA. Dilute 16% PFA with PBS to a final concentration of 4% PFA.</li>
		<li>Dissolve 5 g of NAC in 100 mL of PBS to make a 5% working solution.</li>
	</ol>
	</li>
</ol>
2. Collecting ovaries
<ol>
	<li>Treat sexually mature worms with 5% NAC in a dish at room temperature for 5 min. 
	NOTE: Large worms may curl up. Brushes or pipette tips can be used to flatten the worms. The amount of 5% NAC used is minimal, enough to cover the surface of the Petri dish.</li>
	<li>Replace NAC with 10 mL of freshly prepared 4% paraformaldehyde and fix the worms for 1 h at room temperature with occasional shaking. Start the dissection 10 min after the addition of 4% PFA, and perform the dissection in 4% PFA as the worms are being fixed.</li>
	<li>Observe the epithelial pigmentation to locate the ventral nerve cords, which appear as two lines with lighter pigmentation running from the anterior cephalic ganglia toward the tail.</li>
	<li>Locate the ovaries. 
	NOTE: The ovary is located right next to the nerve cord on the medial side (Figure 1A). Rely on epithelia pigmentation to locate the position of the ovary in the anterior-posterior direction. Ideally, the pigmentation of the ventral epithelia in the ovary region is lighter than the surrounding regions. In some worms, the pigmentation of the ventral epithelia does not provide enough confidence. These worms can be discarded as some may not have well-developed ovaries.</li>
	<li>Validate the ovaries&#39; positions once the putative ovaries are located by pigmentation. 
	NOTE: First, the ovaries are posterior to the cephalic ganglia. Second, dark-colored testes are lateral to the dorsal surface above the ovary. Ovaries locate in front of the most anterior testis (Figure 1B). Sometimes, testis lobes are positioned at the level of or slightly anterior to the ovaries.</li>
	<li>Use two surgical knives to cut posterior and anterior to the two ovaries (Figure 1C) to remove the anterior and posterior worm fragments completely. Use one knife to anchor the worm and clean the other knife used to make the cuts.</li>
	<li>Cut in the middle line of the ovary fragment to separate the two ovaries. Flip the fragment to have the ventral side down.</li>
	<li>Peel off the dorsal tissue to expose the gut (Figure 1D) with two pairs of pointed tweezers (type 5-SA). 
	NOTE: The dome-shaped ovary is located beneath the gut branches.</li>
	<li>Gently remove the gut branches sitting above the ventral tissues with the tip of the tweezers or a soft brush to expose the ovary (Figure 1E).</li>
	<li>Remove the surrounding tissues to take out the ovary (Figure 1F,G).</li>
	<li>Take the ovaries out and transfer them to a 1.5 mL tube to wash with 1 mL of PBS.</li>
</ol>
3. Fixation for TEM
<ol>
	<li>Wash the ovaries in PBS for 10 min with gentle shaking. Keep the tubes upright, and let the ovaries sink to the bottom by gravity. Repeat the wash once. 
	NOTE: If the ovaries do not sink, apply a gentle spin for 15 s with a mini-benchtop centrifuge (maximum speed 2000 &#215; g).</li>
	<li>Replace the PBS with 2% PFA/2.5% GA/PBS.</li>
	<li>Fix the samples at room temperature for 1 h on a shaker at 40 rpm.</li>
	<li>Keep the samples in fixative at 4 &#176;C overnight on a shaker at 40 rpm.</li>
</ol>
4. Sample processing for TEM
<ol>
	<li>Wash the ovaries in PBS 3 times for 10 min each.</li>
	<li>Wash the ovaries in double-distilled water (ddH2O) 3 times for 10 min each.</li>
	<li>Post-fix the ovaries in 2% aqueous OsO4 for 1-2 h at room temperature or overnight at 4 &#176;C. 
	NOTE: The sample containers need to be sealed with parafilm during this step. Prepare 2% OsO4 with reverse-osmosis-treated water (see the Table of Materials).</li>
	<li>Rinse the samples with ddH2O 3 times, 10 min each.</li>
	<li>Pre-stain the ovaries in 2% aqueous uranyl acetate (UA) overnight at 4 &#176;C. 
	NOTE: The UA solution needs to be filtered before use; avoid light during the en bloc staining. Prepare 2% OsO4 with reverse-osmosis-treated water (see the Table of Materials).</li>
	<li>Rinse the samples in ddH2O 4 times with gentle agitation, 10 min each.</li>
	<li>Dehydrate the ovaries in a graded ethanol series (30%, 50%, 70%, 95%, and two times 100%, 10 min each solution) and then equilibrate them in two incubations (10 min) in propylene oxide.</li>
	<li>For infiltration with resin, incubate the samples in 50% propylene oxide/50% liquid epoxy resin mixture overnight at 4 &#176;C. Infiltrate the samples with 100% epoxy resin with 3 changes (1 h each change) with gentle agitation.</li>
	<li>Embed the ovaries in 100% epoxy resin and polymerize the resin at 60 &#176;C for 48 h.</li>
</ol>
5. Ultramicrotomy
<ol>
	<li>Block trimming and sample check with a light microscope
	<ol>
		<li>Trim the sample blocks into a pyramid shape with a razor blade.</li>
		<li>Cut sections of 1-2 &#956;m thickness with a glass or diamond knife.</li>
		<li>Transfer the sections onto a slide with a drop of water, then heat the slide on a hot plate until the sections flatten and adhere to the slide surface during water evaporation.</li>
		<li>Cover the sections with a drop of 1% toluidine blue O and heat them on a hot plate for ~10 s. Rinse the slide with running water, then let it dry. Check the sections under a regular light microscope.</li>
	</ol>
	</li>
	<li>Ultrathin sections cutting, collection, and post-staining.
	<ol>
		<li>Cut ultrathin sections of 50-70 nm thickness with a diamond knife. Transfer the sections from the knife water boat onto mesh or single-slot copper grids.</li>
		<li>Stain the sections in 2% UA for 8 min. Wash the grids in running ddH2O for 30 s.</li>
		<li>Stain the sections with 1% lead solution for 6 min. Wash the sections in running ddH2O for 1 min and air-dry.</li>
	</ol>
	</li>
</ol>
6. Data collection and analysis
<ol>
	<li>Collect TEM data using appropriate software, e.g., Digital Micrograph (see the Table of Materials for more details).</li>
	<li>Operate the scope at 80 kV. Insert the sample grid into the scope when the vacuum is ready.</li>
	<li>Turn on the beam by clicking Light on the menu. Turn the Intensity knob on the left control pad until the Auto Exposure time is ~1 s. Click on Start Acquire to get a final image.</li>
	<li>Construct three-dimensional EM models using the open-source image processing, modeling, and display (IMOD) package.</li>
</ol>
7. Antibody staining
<ol>
	<li>Wash the dissected ovaries (step 2.16) in a 1.5 mL microfuge tube with 1 mL of 1x PBS supplemented with 0.5% Triton X-100 (PBST). Place the tubes on a shaker for agitation at 40 rpm for 10 min. Let the tubes stand in a rack and the ovaries sink by gravity. Replace the wash with fresh PBST, and repeat twice.</li>
	<li>Digest the tissues with 2 &#181;g/mL of Proteinase K and 0.1% sodium dodecylsulfate for 10 min at room temperature in PBST.</li>
	<li>Wash the digested ovaries in PBST three times for 10 min each wash.</li>
	<li>Incubate the ovaries with 10% horse serum in PBST for 1 h at room temperature.</li>
	<li>Dilute primary antibodies 1 to 100 in the blocking solution (10% horse serum in PBS with 0.5% Triton X-100). Incubate the ovaries with primary antibodies overnight at 4 &#176;C with gentle shaking.</li>
	<li>Wash the ovaries in PBST three times for 10 min each wash.</li>
	<li>Incubate the ovaries with secondary antibodies overnight at 4 &#176;C with gentle shaking. Dilute all secondary antibodies 1:300 in the blocking solution.</li>
	<li>Wash the ovaries in PBST three times, ensuring that the first and third washes last for 10 min. Stain the ovarian nuclei with Hoechst 33342 at 1:300 dilution in PBST for 30 min at room temperature in the second wash.</li>
	<li>Mount the ovaries onto slides. 
	NOTE: Anti-fading mountant can be used to prolong the fluorescence signals.</li>
</ol>

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The authors have nothing to disclose.

These fixation-based procedures for dissecting planarian ovaries will facilitate the understanding of oocyte meiosis as well as ovary development and regeneration. The sizes of the oocytes and their somatic supportive cells can range from 20 &#181;m to 50 &#181;m. Dissection-based methods will provide accessibility to intact single-ovary cells that sectioning or whole-mount-based methods cannot achieve. This protocol will facilitate the study of intact planarian ovary anatomy and oocyte cell biology at cellular and subce...

Planarian anatomy has been examined by using TEM in many tissues1,2,3,4,5,6. However, little attention has been given to ovaries or oocytes. The paucity of oocyte literature is partly due to the difficulty accessing these cells, leaving the biology of planarian oocytes largely unexplored. Molecular tools have uncovered many regulatory mechanisms of ovary development in the planarians&#160;using light or fluorescence microscopy7,8,9,10,11,12,13,14,15,16,17,18,19,20. All these experiments were performed on whole worms or histological sections of whole worms. The antibody staining and in situ hybridization protocols on whole worms involve extensive bleaching, washing, and tissue clearing steps, which are time-consuming and will take several days.
The overall goal of the method described here is to provide accessibility to intact, dissected planarian ovaries and oocytes, which will remove the necessity of bleaching or histological sectioning and shorten the time for washing and tissue clearing in antibody staining and in situ hybridization. The dissected ovaries will also improve probe or antibody penetration and increase imaging depth and quality for light and electron microscopes. Accessibility to the dissected ovaries and oocytes allows cell biology research at cellular and subcellular resolution with whole intact oocytes. A recent study on dissected planarian ovaries characterized planarian oocyte meiosis for the first time with TEM and confocal microscopy21. The work provided a comprehensive description of a new phenomenon during meiosis called nuclear envelope vesiculation.
Here, we present the detailed procedures in the dissection of planarian ovaries. A fixation step was sufficient to preserve the ovary cell structure for dissection and downstream manipulation (i.e., processing for TEM and light microscope analysis). Given their similarity in body plans and tissue architecture, this protocol should also be broadly informative for studying oocytes and their nuclei in several other Platyhelminthes species (e.g., the genus of Dugesia or Polycelis). This protocol is likely irrelevant for Macrostomum lignano for their small sizes and almost transparent body architecture, which will allow for direct observation of the ovary and oocytes22,23,24. The body area containing the ovaries is more optically distinguishable (e.g., darker pigmented or lighter pigmented) in some species (e.g., Dugesia ryukyuensis9,25) than S. mediterranea. Studies in these species can rely less on the guidelines for locating the ovaries in S. mediterranea presented here but take advantage of the fixation and dissection conditions.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>16% paraformaldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>15710</td>
			<td>EM grade</td>
		</tr>
		<tr>
			<td>2% aqueous OsO4</td>
			<td>Electron Microscopy Sciences</td>
			<td>19152</td>
			<td></td>
		</tr>
		<tr>
			<td>50% glutaraldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>16320</td>
			<td>EM grade</td>
		</tr>
		<tr>
			<td>Digital Micrograph</td>
			<td>Gatan Inc.</td>
			<td></td>
			<td>Version 2.33.97.1, TEM data collection</td>
		</tr>
		<tr>
			<td>Epon resin</td>
			<td>Electron Microscopy Sciences</td>
			<td>14120</td>
			<td>Embed 812 Kit, liquid, epoxy resin</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Ted Pella</td>
			<td>19207</td>
			<td>Denatured</td>
		</tr>
		<tr>
			<td>Hoechst 33342</td>
			<td>Thermo Fisher Scientific</td>
			<td>H3570</td>
			<td></td>
		</tr>
		<tr>
			<td>Horse serum</td>
			<td>Sigma</td>
			<td>H1138</td>
			<td></td>
		</tr>
		<tr>
			<td>Lead Acetate</td>
			<td>Electron Microscopy Sciences</td>
			<td>6080564</td>
			<td></td>
		</tr>
		<tr>
			<td>MilliQ water</td>
			<td></td>
			<td></td>
			<td>reverse-osmosis treated water</td>
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		<tr>
			<td>N-Acetyl-L-cysteine</td>
			<td>Sigma</td>
			<td>A7250</td>
			<td></td>
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		<tr>
			<td>Parafilm</td>
			<td>sigma</td>
			<td>P7793</td>
			<td></td>
		</tr>
		<tr>
			<td>Prolong Diamond Antifade Mountant</td>
			<td>Thermo Fisher Scientific</td>
			<td>P36965</td>
			<td></td>
		</tr>
		<tr>
			<td>Propylene oxide</td>
			<td>Electron Microscopy Sciences</td>
			<td>75569</td>
			<td>EM grade</td>
		</tr>
		<tr>
			<td>&#160;Proteinase K</td>
			<td>Thermo Fisher Scientific</td>
			<td>25530049</td>
			<td></td>
		</tr>
		<tr>
			<td>Toluidine blue O</td>
			<td>Electron Microscopy Sciences</td>
			<td>92319</td>
			<td></td>
		</tr>
		<tr>
			<td>Transmission Electron Microscope</td>
			<td>FEI</td>
			<td></td>
			<td>Tecnai G2 Spirit BioTWIN</td>
		</tr>
		<tr>
			<td>Uranyl acetate</td>
			<td>Electron Microscopy Sciences</td>
			<td>541093</td>
			<td></td>
		</tr>
	</tbody>
</table>

planarian ovary dissection for ultrastructural analysis and antibody staining

Accessibility to germ cells allows the study of germ cell development, meiosis, and recombination. The sexual biotype of the freshwater planarian, Schmidtea mediterranea, is a powerful invertebrate model to study the epigenetic specification of germ cells. Unlike the large number of testis and male germ cells, planarian oocytes are relatively difficult to locate and examine, as there are only two ovaries, each with 5-20 oocytes. Deeper localization within the planarian body and lack of protective epithelial tissues also make it challenging to dissect planarian ovaries directly. 
This protocol uses a brief fixation step to facilitate the localization and dissection of planarian ovaries for downstream analysis to overcome these difficulties. The dissected ovary is compatible for ultrastructural examination by transmission electron microscopy (TEM) and antibody immunostaining. The dissection technique outlined in this protocol also allows for gene perturbation experiments, in which the ovaries are examined under different RNA interference (RNAi) conditions. Direct access to the intact germ cells in the ovary achieved by this protocol will greatly improve the imaging depth and quality and allow cellular and subcellular interrogation of oocyte biology.

The method presented here has been described by Guo et al.21. The key to successful dissection is to identify the ovary pigmentation and position guides correctly. The strategy of the method is to move from broad positions to a specific location. First, to achieve this, rely on dorsal and ventral pigmentation patterns (Figure 1A,B). Ventral pigmentation, where the ovaries reside, will turn white after 5% NAC treatment and 4% PFA fixation. If the worm ...

Watch this Scientific Journal Video about Planarian Ovary Dissection for Ultrastructural Analysis and Antibody Staining at JoVE.com

Planarian Ovary Dissection for Ultrastructural Analysis and Antibody Staining

This protocol presents steps taken to dissect ovaries in the freshwater planarians, Schmidtea mediterranea. The dissected ovaries are compatible for antibody immunostaining and ultrastructural analysis with transmission electron microscopy to study the cell biology of the oocytes and somatic cells, providing an imaging depth and quality that were previously inaccessible.

Accessibility to germ cells allows the study of germ cell development, meiosis, and recombination. The sexual biotype of the freshwater planarian, ...

planarian-ovary-dissection-for-ultrastructural-analysis-antibody

Research

JoVE Journal

Biology

1.5K Views.  Stowers Institute for Medical Research. This protocol presents steps taken to dissect ovaries in the freshwater planarians, Schmidtea mediterranea. The dissected ovaries are compatible for antibody immunostaining and ultrastructural analysis with transmission electron microscopy to study the cell biology of the oocytes and somatic cells, providing an imaging depth and quality that were previously inaccessible.

Video: Planarian Ovary Dissection for Ultrastructural Analysis and Antibody Staining

Method for Whole Mount Antibody Staining in Chick

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying antibody expression in developing brain, neural tube and somite. This video demonstrates the different steps in whole-mount antibody staining using HRP conjugated secondary antibodies; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, endogenous peroxidase is inactivated; The embryo is then exposed to primary antibody. After several washes, the embryo is incubated with secondary antibody conjugated to HRP. Peroxidase activity is revealed using reaction with diaminobenzidine substrate. Finally, the embryo is fixed and processed for photography and sectioning. The advantage of this method over the use of fluorescent antibodies is that embryos can be processed for wax sectioning, thus enabling the study of antigen sites in cross section. This method was originally introduced by Jane Dodd and Tom Jessell 1.

This video demonstrates whole mount immunohistochemistry, a method by which the spatial and temporal expression pattern of an antigen can be visualized in young chick embryos. This method was originally introduced by Jane Dodd and Tom Jessell.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

A Neuronal and Astrocyte Co-Culture Assay for High Content Analysis of Neurotoxicity

High Content Analysis (HCA) assays combine cells and detection reagents with automated imaging and powerful image analysis algorithms, allowing measurement of multiple cellular phenotypes within a single assay. In this study, we utilized HCA to develop a novel assay for neurotoxicity. Neurotoxicity assessment represents an important part of drug safety evaluation, as well as being a significant focus of environmental protection efforts. Additionally, neurotoxicity is also a well-accepted in vitro marker of the development of neurodegenerative diseases such as Alzheimer's and Parkinson's diseases. Recently, the application of HCA to neuronal screening has been reported. By labeling neuronal cells with &beta;III-tubulin, HCA assays can provide high-throughput, non-subjective, quantitative measurements of parameters such as neuronal number, neurite count and neurite length, all of which can indicate neurotoxic effects. However, the role of astrocytes remains unexplored in these models. Astrocytes have an integral role in the maintenance of central nervous system (CNS) homeostasis, and are associated with both neuroprotection and neurodegradation when they are activated in response to toxic substances or disease states. GFAP is an intermediate filament protein expressed predominantly in the astrocytes of the CNS. Astrocytic activation (gliosis) leads to the upregulation of GFAP, commonly accompanied by astrocyte proliferation and hypertrophy. This process of reactive gliosis has been proposed as an early marker of damage to the nervous system. The traditional method for GFAP quantitation is by immunoassay. This approach is limited by an inability to provide information on cellular localization, morphology and cell number. We determined that HCA could be used to overcome these limitations and to simultaneously measure multiple features associated with gliosis - changes in GFAP expression, astrocyte hypertrophy, and astrocyte proliferation - within a single assay. In co-culture studies, astrocytes have been shown to protect neurons against several types of toxic insult and to critically influence neuronal survival. Recent studies have suggested that the use of astrocytes in an in vitro neurotoxicity test system may prove more relevant to human CNS structure and function than neuronal cells alone. Accordingly, we have developed an HCA assay for co-culture of neurons and astrocytes, comprised of protocols and validated, target-specific detection reagents for profiling &beta;III-tubulin and glial fibrillary acidic protein (GFAP). This assay enables simultaneous analysis of neurotoxicity, neurite outgrowth, gliosis, neuronal and astrocytic morphology and neuronal and astrocytic development in a wide variety of cellular models, representing a novel, non-subjective, high-throughput assay for neurotoxicity assessment. The assay holds great potential for enhanced detection of neurotoxicity and improved productivity in neuroscience research and drug discovery.

This article describes a novel protocol and reagent set designed for sensitive measurement of neurotoxic effects of compounds and treatments on co-cultures of neurons and astrocytes using high content analysis. Results demonstrate that high content analysis represents an exciting novel technology for neurotoxicity assessment.

High Content Analysis (HCA) assays combine cells and detection reagents with automated imaging and powerful image analysis algorithms, allowing ...

Cancer Borealis Stomatogastric Nervous System Dissection

The stomatogastric ganglion (STG) is an excellent model for studying cellular and network interactions because it contains a relatively small number of cells (approximately 25 in C. borealis) which are well characterized. The cells in the STG exhibit a broad range of outputs and are responsible for the motor actions of the stomach. The stomach contains the gastric mill which breaks down food with three internal teeth, and the pylorus which filters the food before it reaches the midgut. The STG produces two rhythmic outputs to control the gastric mill and pylorus known as central pattern generators (CPGs). Each cell in the STG can participate in one or both of these rhythms. These CPGs allow for the study of neuromodulation, homeostasis, cellular and network variability, network development, and network recovery.
The dissection of the stomatogastric nervous system (STNS) from the Jonah crab (Cancer borealis) is done in two parts; the gross and fine dissection. In the gross dissection the entire stomach is dissected from the crab. During the fine dissection the STNS is extracted from the stomach using a dissection microscope and micro-dissection tools (see figure 1). The STNS includes the STG, the oesophageal ganglion (OG), and the commissural ganglia (CoG) as well as the nerves that innervate the stomach muscles. Here, we show how to perform a complete dissection of the STNS in preparation for an electrophysiology experiment where the cells in the STG would be recorded from intracellularly and the peripheral nerves would be used for extracellular recordings. The proper technique for finding the desired nerves is shown as well as our technique of desheathing the ganglion to reveal the somata and neuropil.

Cancer Borealis Stomatogastric Nervous System Dissection

The stomatogastric nervous system (STNS) of the Jonah crab (C. borealis) can be used for electrophysiology, immunohistochemistry, and cell culture studies. The STNS extraction is done in two parts: the gross and fine dissection.

The stomatogastric ganglion (STG) is an excellent model for studying cellular and network interactions because it contains a relatively small number of ...

Dissection of Hippocampal Dentate Gyrus from Adult Mouse

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its remarkable neuronal cell plasticity, and its involvement in epilepsy, neurodegenerative diseases, and psychiatric disorders. The hippocampus is composed of distinct regions; the dentate gyrus, which comprises mainly granule neurons, and Ammon's horn, which comprises mainly pyramidal neurons, and the two regions are connected by both anatomic and functional circuits. Many different mRNAs and proteins are selectively expressed in the dentate gyrus, and the dentate gyrus is a site of adult neurogenesis; that is, new neurons are continually generated in the adult dentate gyrus. To investigate mRNA and protein expression specific to the dentate gyrus, laser capture microdissection is often used. This method has some limitations, however, such as the need for special apparatuses and complicated handling procedures. In this video-recorded protocol, we demonstrate a dissection technique for removing the dentate gyrus from adult mouse under a stereomicroscope. Dentate gyrus samples prepared using this technique are suitable for any assay, including transcriptomic, proteomic, and cell biology analyses. We confirmed that the dissected tissue is dentate gyrus by conducting real-time PCR of dentate gyrus-specific genes, tryptophan 2,3-dioxygenase (TDO2) and desmoplakin (Dsp), and Ammon's horn enriched genes, Meis-related gene 1b (Mrg1b) and TYRO3 protein tyrosine kinase 3 (Tyro3). The mRNA expressions of TDO2 and Dsp in the dentate gyrus samples were detected at obviously higher levels, whereas Mrg1b and Tyro3 were lower levels, than those in the Ammon's horn samples. To demonstrate the advantage of this method, we performed DNA microarray analysis using samples of whole hippocampus and dentate gyrus. The mRNA expression of TDO2 and Dsp, which are expressed selectively in the dentate gyrus, in the whole hippocampus of alpha-CaMKII+/- mice, exhibited 0.037 and 0.10-fold changes compared to that of wild-type mice, respectively. In the isolated dentate gyrus, however, these expressions exhibited 0.011 and 0.021-fold changes compared to that of wild-type mice, demonstrating that gene expression changes in dentate gyrus can be detected with greater sensitivity. Taken together, this convenient and accurate dissection technique can be reliably used for studies focused on the dentate gyrus.

A dissection technique for removal of the dentate gyrus from adult mouse under a stereomicroscope was demonstrated in this video-recorded protocol.

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its ...

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations. In these preparations, the central nervous system runs down the middle, and is flanked by the body walls. Many different phenotypes have been examined using such preparations. In most cases, the fillets were generated by dissection of antibody-stained fixed whole-mount embryos. These "fixed dissections" have some disadvantages, however. They are time-consuming to execute, and it is difficult to sort mutant (GFP-negative) embryos from stocks in which mutations are maintained over GFP balancer chromosomes. Since 2002, our group has been conducting deficiency and ectopic expression screens to identify ligands for orphan receptors. In order to do this, we developed streamlined protocols for live embryo dissection and antibody staining of collections containing hundreds of balanced lines. We have concluded that it is considerably more efficient to examine phenotypes in large collections of stocks by live dissection than by fixed dissection. Using the protocol described here, a single trained individual can screen up to 10 lines per day for phenotypes, examining 4-7 mutant embryos from each line under a compound microscope. This allows the identification of mutations conferring subtle, low-penetrance phenotypes, since up to 70 hemisegments per line are scored at high magnification with a 40X water-immersion lens.

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

We describe a streamlined protocol for generating "fillet" preparations of Drosophila embryos of specific genotypes. This protocol allows efficient execution of a variety of genetic screens. It also allows excellent visualization of structures in the late embryo.

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations.  In these preparations, ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

Planarian Immobilization, Partial Irradiation, and Tissue Transplantation

The planarian, a freshwater flatworm, has proven to be a powerful system for dissecting metazoan regeneration and stem cell biology1,2. Planarian regeneration of any missing or damaged tissues is made possible by adult stem cells termed neoblasts3. Although these stem cells have been definitively shown to be pluripotent and singularly capable of reconstituting an entire animal4, the heterogeneity within the stem cell population and the dynamics of their cellular behaviors remain largely unresolved. Due to the large number and wide distribution of stem cells throughout the planarian body plan, advanced methods for manipulating subpopulations of stem cells for molecular and functional study in vivo are needed.
Tissue transplantation and partial irradiation are two methods by which a subpopulation of planarian stem cells can be isolated for further study. Each technique has distinct advantages. Tissue transplantation allows for the introduction of stem cells, into a na&iuml;ve host, that are either inherently genetically distinct or have been previously treated pharmacologically. Alternatively, partial irradiation allows for the isolation of stem cells within a host, juxtaposed to tissue devoid of stem cells, without the introduction of a wound or any breech in tissue integrity. Using these two methods, one can investigate the cell autonomous and non-autonomous factors that control stem cell functions, such as proliferation, differentiation, and migration.
Both tissue transplantation5,6 and partial irradiation7 have been used historically in defining many of the questions about planarian regeneration that remain under study today. However, these techniques have remained underused due to the laborious and inconsistent nature of previous methods. The protocols presented here represent a large step forward in decreasing the time and effort necessary to reproducibly generate large numbers of grafted or partially irradiated animals with efficacies approaching 100 percent. We cover the culture of large animals, immobilization, preparation for partial irradiation, tissue transplantation, and the optimization of animal recovery. Furthermore, the work described here demonstrates the first application of the partial irradiation method for use with the most widely studied planarian, Schmidtea mediterranea. Additionally, efficient tissue grafting in planaria opens the door for the functional testing of subpopulations of na&iuml;ve or treated stem cells in repopulation assays, which has long been the gold-standard method of assaying adult stem cell potential in mammals8. Broad adoption of these techniques will no doubt lead to a better understanding of the cellular behaviors of adult stem cells during tissue homeostasis and regeneration.

An effective method for grafting tissue of defined and consistent size between planaria is described. Also included is a description of how the immobilization technique used for transplantation can be adapted, in conjunction with lead shields, for the partial irradiation of live animals.

The planarian, a freshwater flatworm, has proven to be a powerful system for dissecting metazoan regeneration and stem cell biology1,2. Planarian ...

Induction and Analysis of Epithelial to Mesenchymal Transition

Epithelial to mesenchymal transition (EMT) is essential for proper morphogenesis during development. Misregulation of this process has been implicated as a key event in fibrosis and the progression of carcinomas to a metastatic state. Understanding the processes that underlie EMT is imperative for the early diagnosis and clinical control of these disease states. Reliable induction of EMT in vitro is a useful tool for drug discovery as well as to identify common gene expression signatures for diagnostic purposes. Here we demonstrate a straightforward method for the induction of EMT in a variety of cell types. Methods for the analysis of cells pre- and post-EMT induction by immunocytochemistry are also included. Additionally, we demonstrate the effectiveness of this method through antibody-based array analysis and migration/invasion assays.

A straightforward method is described for the induction of epithelial to mesenchymal transition (EMT) in a variety of cell types. Methods for the analysis of cells in EMT states by immunocytochemistry are included.

Epithelial  to mesenchymal transition (EMT) is essential for proper morphogenesis during  development. Misregulation of this  process has been implicated ...

Antibody Staining in C. Elegans Using &quot;Freeze-Cracking&quot;

To stain C. elegans with antibodies, the relatively impermeable cuticle must be bypassed by chemical or mechanical methods. "Freeze-cracking" is one method used to physically pull the cuticle from nematodes by compressing nematodes between two adherent slides, freezing them, and pulling the slides apart. Freeze-cracking provides a simple and rapid way to gain access to the tissues without chemical treatment and can be used with a variety of fixatives. However, it leads to the loss of many of the specimens and the required compression mechanically distorts the sample. Practice is required to maximize recovery of samples with good morphology. Freeze-cracking can be optimized for specific fixation conditions, recovery of samples, or low non-specific staining, but not for all parameters at once. For antibodies that require very hard fixation conditions and tolerate the chemical treatments needed to chemically permeabilize the cuticle, treatment of intact nematodes in solution may be preferred. If the antibody requires a lighter fix or if the optimum fixation conditions are unknown, freeze-cracking provides a very useful way to rapidly assay the antibody and can yield specific subcellular and cellular localization information for the antigen of interest.

Antibody Staining in C. Elegans Using "Freeze-Cracking"

"Freeze-cracking," a method for exposing the inner tissues of the nematode C. elegans to antibodies for protein localization, is demonstrated.

To stain C.  elegans with antibodies, the relatively impermeable cuticle must be  bypassed by chemical or mechanical methods.  "Freeze-cracking" is one ...

Dissection and Downstream Analysis of Zebra Finch Embryos at Early Stages of Development

The zebra finch (Taeniopygiaguttata) has become an increasingly important model organism in many areas of research including toxicology1,2, behavior3, and memory and learning4,5,6. As the only songbird with a sequenced genome, the zebra finch has great potential for use in developmental studies; however, the early stages of zebra finch development have not been well studied. Lack of research in zebra finch development can be attributed to the difficulty of dissecting the small egg and embryo. The following dissection method minimizes embryonic tissue damage, which allows for investigation of morphology and gene expression at all stages of embryonic development. This permits both bright field and fluorescence quality imaging of embryos, use in molecular procedures such as in situ hybridization (ISH), cell proliferation assays, and RNA extraction for quantitative assays such as quantitative real-time PCR (qtRT-PCR).&#160;This technique allows investigators to study early stages of development that were previously difficult to access.

The zebra finch (Taeniopygiaguttata) is a valuable model organism; however, early stages of zebra finch development have not been extensively studied. The protocol describes how to dissect early embryos for developmental and molecular applications.

The zebra finch (Taeniopygiaguttata) has become an increasingly important model organism in many areas of research including toxicology1,2, behavior3, and ...