The authors gratefully thank Maria Herrero and Eliseo Rivas for their helpful discussion and advice. This work was supported by the Ministerio de Econom&#237;a y Competitividad &#8212; European Regional Development Fund, European Union [grant number BES- 2010-037992 to E. F.]; the Instituto Nacional de Investigaci&#243;n y Tecnolog&#237;a Agraria y Alimentaria [grant numbers RFP2015-00015-00, RTA2014-00085-00, RTA2017-00003-00]; and the Gobierno de Arag&#243;n &#8212; European Social Fund, European Union [Grupo Consolidado A12-17R].

1. Dormancy Determination and Plant Material Collection
<ol>
	<li>Sample the flower buds in the field. Dormancy studies are long-term experiments and require adult trees big enough to collect buds and shoots all winter without compromising the trees&#8217; development during the next spring. Special orchard management could be required depending on the training system; thus, pruning may be less severe than for fruit production purposes.
	<ol>
		<li>Each week, from the start of autumn until the onset of bud break, collect and weigh 10 flower buds.</li>
		<li>Fix the flower buds by placing them into a 10-mL glass tube with a cap and soak the samples in a fixative solution of ethanol/acetic acid (3:1) for at least 24 h at 4 &#176;C. Then, discard the fixative and add 75% ethanol, generously ensuring that it covers the samples. The samples can be conserved in this solution at 4 &#176;C until use. 
		Note: Vacuum infiltration can be used to remove the air bubbles inside the bud and prevent them from floating. This facilitates the penetration of the fixative into the tissues but could damage the structure of the tissues. Try to avoid it if not necessary.</li>
	</ol>
	</li>
	<li>Sample the shoots in the field.
	<ol>
		<li>Each week, from the start of autumn until the onset of bud break, take three shoots of 15 - 30 cm in length and 5 mm in diameter, containing at least 10 flower buds each. Place them on water-soaked florist foam in a growth chamber at 22 &#177; 1 &#176;C with a 12 h light photoperiod.</li>
		<li>After 10 days in the growth chamber, pick and weigh 10 flower buds from the shoots.</li>
	</ol>
	</li>
	<li>Evaluate the bud growth and determine the dormancy status. The sampling period has to be adjusted to the conditions of the location. In the orchard conditions (Zaragoza, Spain, 41&#176;44&#8217;30&#8221; N, 0&#176;47&#8217;00&#8221; W, and 220 m above sea level), the sampling of the shoots was carried out from November 30 until the end of February or the beginning of March.
	<ol>
		<li>Weekly evaluate the response of the flower buds to the suitable growth conditions of the chamber, from the beginning of autumn until budburst in the spring, by comparing the weight of the 10 buds picked from the field.</li>
		<li>If there are no differences or those differences are less than 30%, consider that the buds have not fulfilled their chilling requirements and are endodormant22. If the differences are more than 30%, consider that the buds have fulfilled their chilling requirements and are ecodormant22.</li>
	</ol>
	</li>
</ol>
2. Plant Material Preparation for Starch Quantification
<ol>
	<li>Select about six fixed buds from each sampling date (see step 1.1). Depending on cultivars, each sweet cherry flower bud contains up to five flower primordia.
	<ol>
		<li>Remove the external bud scales and place the bud on a watchmaker&#8217;s glass with 75% ethanol to prevent it from drying out.</li>
		<li>Dissect the bud and extract at least one flower primordium per bud with the help of precision forceps and an ophthalmologic scalpel under a stereoscopic microscope. The flower primordium can be conserved in a 10-mL glass tube with 75% ethanol at 4 &#176;C, or proceed immediately to the next step.</li>
	</ol>
	</li>
	<li>Dehydrate the samples in a tertiary butyl alcohol series.
	<ol>
		<li>Replace the 75% ethanol solution with 10 mL of 75% tertiary butyl alcohol (TBA) to generously cover the samples and incubate them for 1.5 h. A Pasteur pipette can be useful to discard the solution.</li>
		<li>Repeat step 2.2.1, adding TBA with rising concentrations (85%, 95%, and 100%, v/v; and 3x with pure TBA) in a drying stove at 30 &#176;C with air extraction, since pure TBA crystalizes at room temperature (&#60; 20 &#176;C) and is very volatile and toxic. If TBA crystalizes with the sample, it would damage the tissues.</li>
	</ol>
	</li>
	<li>Embed the samples in paraffin18.
	<ol>
		<li>Melt the paraffin by introducing the paraffin pearls in the drying stove at 60 &#176;C with air extraction the previous day. The paraffin pearls can be also melted on a hotplate, but ensure that the liquid paraffin is at 60 &#176;C and not at a higher temperature, to avoid tissue damage.</li>
		<li>Replace the TBA with a mix of TBA and paraffin oil (1:1) and incubate for 24 h inside a drying stove at 60 &#176;C. Then, replace the mix of TBA and paraffin oil with pure melt paraffin and incubate for at least 6 h inside a drying stove at 60 &#176;C. Repeat it 2x and incubate the last change for at least 4 - 6 d.</li>
		<li>Place each sample on a small metal base mold over a heat surface, embedded in paraffin wax, and pace the embedding cassette. Place it over a cold surface and remove the block once the wax is solidified.</li>
	</ol>
	</li>
	<li>Section and rehydrate the preparations.
	<ol>
		<li>Prepare Haupt&#8217;s adhesive: dissolve 1 g of plain Knox gelatin in 100 mL of distilled water at 30 &#176;C; then, add 2 g of phenol loose crystals (C6H5OH) and 15 mL of glycerol. Spread the Haupt&#8217;s adhesive over a glass slide with a brush and add a drop of a 1% formaldehyde solution.</li>
		<li>Section each paraffin block at 10 &#181;m in a rotatory microtome and place the sections onto the glass slide covered with Haupt&#8217;s adhesive.</li>
		<li>Place the glass slides with the sections over a heat surface at 35 - 40 &#176;C until dry; then, move the glass slides to a slide rack.</li>
		<li>Prepare the dewax and rehydration solutions. Place 200 mL of Histoclear II in three glass-staining dishes, another one with Histoclear II:ethanol (1:1, v/v), an ethanol series (100%, 70%, 40%, v/v), and a final distilled-water wash.</li>
		<li>Dewax the sections with three washes, each 5 min, in Histoclear II and in Histoclear II:ethanol (1:1, v/v). Place the slide rack inside the glass-staining dishes, ensuring that the solution completely covers the slides, and, then, move the slide rack from one solution to the next.</li>
		<li>Rehydrate the sections by the following series of 2-min washes: an ethanol series (100%, 70%, 40%, v/v) and a final distilled-water wash. Finally, dry the glass slides at room temperature.</li>
	</ol>
	</li>
	<li>Stain the sections18.
	<ol>
		<li>Prepare I2KI stain: dissolve 2 g of potassium iodide (KI) and 0.2 g of iodine (I2) in 100 mL of distilled water. Apply a drop of fresh I2KI over the section(s) for 5 min and, then, discard the excess of stain by absorbing it with a blotting paper. Quickly proceed to the next step.</li>
		<li>Apply a small drop of a synthetic mounting media, place a small cover glass on top, and press hard. Once the mounting media dries, observe under a bright-field microscope for a preliminary evaluation of the ovary starch. 
		Note: This step is not mandatory, although a better contrast between the starch and the background is obtained. If the section has to be reused with other stains after starch quantification, place the cover glass over the drop of stain and discard the excess by absorbing it with a blotting paper (see 2.5.1). Then, after starch quantification, wash out the I2KI stain with distilled water and place the preparations over a heat surface at 35 - 40 &#176;C until dry.</li>
	</ol>
	</li>
</ol>
3. Quantification of the Starch Content
<ol>
	<li>Calibrate the optical conditions. 
	Note: The detection levels used by the image analyzer to detect the stained starch are directly dependent on light conditions and magnification of the microscope; thus, fix these conditions for all the preparations evaluated. Adapt the adjustment proposed here to the available microscope and light conditions.
	<ol>
		<li>Adjust the aperture diaphragm at the 20X magnification and the brightness control or light intensity.</li>
		<li>Make sure that there are no filters on the filter holder, and select a bright-field condition in the microscope. Select a suitable magnification (e.g., 40X for sweet cherry ovary primordium).</li>
	</ol>
	</li>
	<li>Control the image acquisition conditions. Adjust the camera settings with a stained preparation without tissue via Image | Acquisition | Pre-view.
	<ol>
		<li>Fix the brightness at 50%, the gain at 1.0x, and the histogram indicators the gamma value at 1.00 and the contrast at 0 - 100, lining up with the limits of the brightness distribution histogram.</li>
		<li>Activate the overexposure/underexposure function and adjust the exposure time at the limit of overexposure.</li>
		<li>Apply the white balance function to the complete image to display all neutral-colored components of the image without any color tone and the shading correction to the complete image to create a corrected and homogeneous image.</li>
	</ol>
	</li>
	<li>Calibrate the Image Analysis System to obtain the control values of the grey level (0, black; 255, white) of different optical densities (OD) that are obtained by the transmittance (T) values.
	<ol>
		<li>Acquire an image of a stained preparation without tissue, considered control white, and measure the grey level of the black and white image via Measure | Grey measure | Calibrate grey | Reference value = 0 | Measure | Calibrate | OK. This corresponds with a 100% transmittance; thus, an optical density of 0, according to OD = 2 - log T.</li>
		<li>Acquire an image of the same preparation with a 4N filter, which reduces the amount of light 4x, and measure the grey level of the black and white image via Measure | Grey measure | Calibrate grey | Reference value = 0.6 | Measure | Calibrate | OK. This corresponds with a 25% transmittance; thus, an optical density of 0.6, according to OD = 2 - log T.</li>
		<li>Acquire an image of the same preparation without light, considered black, and measure the grey level of the black and white image via Measure | Grey measure | Calibrate grey | Reference value = 1 | Measure | Calibrate | OK. This corresponds with a 0% transmittance; thus, an optical density of 1, according to OD = 2 - log T.</li>
	</ol>
	</li>
	<li>Detect starch.
	<ol>
		<li>Acquire a color image of the field to measure in TIFF format with a resolution of at least 300 dots per inch (dpi).</li>
		<li>Create a binary image corresponding to the area stained. Set the three color thresholds (values between 0 - 255 for each) until the binary image exactly reflects the stained starch granules observed, via Image | Detect | Select the thresholds of red, blue and green | OK. Make repeatedly visual comparisons in various preparations and tissues to tune up the final detection levels. Store and use these levels for all preparations.</li>
		<li>Quantify the starch. Convert the original color image into a black and white image with the image analysis system. Use the binary image as a superimposed mask on the black and white image via Image | Binary Edit. Measure the sum of the optical density of every pixel under the mask via Measure | Grey level | OK and consider this value as the starch content in the measured field.</li>
		<li>Repeat steps 3.4.1 - 3.4.3 in four places of the ovary primordia to obtain a representative value of the starch content in the ovary of flower primordia.</li>
		<li>Repeat steps 3.4.1 - 3.4.3 and step 3.4.5 in different flowers of each collecting date.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Dormancy in woody perennials presents clear implications in fruit production and forestry in a changing climate, although the biological process behind dormancy remains unclear. Dormancy studies can be approached from different points of view, but the research looking for a biological marker for winter dormancy has intensified over the last years. However, most attempts to find an unequivocal indicator showing when a bud has broken dormancy have been unsuccessful3. The methodology described herein...

Temperate woody perennials adapt to the seasons by modulating their growth and development. While they develop during spring and summer, they stop growing during the autumn to go dormant in winter1. Although dormancy allows them to survive at low winter temperatures, chilling is a prerequisite for a proper budburst in spring2. The important implications of dormancy in temperate fruit production and forestry have led to diverse efforts to determine and predict the dormancy period3. In fruit tree species, empirical experiments transferring shoots to forcing conditions and statistical predictions based on data of flowering are current approaches to determine the date of the breaking of dormancy, which allows researchers to estimate the chilling requirements for each cultivar. However, how to determine the dormancy status based on biological processes remains unclear3. 
Flowering in temperate fruit trees, such as sweet cherry (Prunus avium L.), occurs once a year and lasts about a fortnight. However, flowers start to differentiate and develop about 10 months earlier, during the previous summer4. Flower primordia stop growing during the autumn to remain dormant inside the buds during winter. In this period, each cultivar needs to accumulate a particular chilling requirement for proper flowering4. Despite the lack of phenological changes in the buds during winter, flower primordia are physiologically active during dormancy, and the accumulation of chilling temperatures has been recently associated with the dynamic of starch accumulation or decrease within the cells of the ovary primordium, offering a new approach for dormancy determination5. However, the small size and the location of the ovary primordium require a special methodology.
Starch is the major storage carbohydrate in woody plant species6. Thus, changes in starch have been related to the physiological activity of the flower tissues, which need carbohydrates to support their development7,8. Different key events during the reproductive process are also related to variations in starch content in different floral structures, such as anther meiosis9, the growth of the pollen tubes through the style or ovule fertilization10. Histochemical techniques allow the detection of starch in each particular tissue of the flower primordia during dormancy. However, the difficulty remains in quantifying that starch to allow following its pattern of accumulation/decrease over time or comparing the starch content among tissues, cultivars, or years. This is due to the little amount of tissue available for the analytical techniques11. As an alternative, image analysis linked to microscopy12 allows the quantification of the starch in very small samples of plant tissue13. 
Approaches combining microscopy and image analysis have been used to quantify the content of different components in plant tissues, such as callose14, microtubes15, or starch16, by measuring the size of the area dyed by specific stains. For starch, it can be easily detected using the potassium iodide-iodine (I2KI) reaction17. This method is highly specific; I2KI intercalates within the laminar structure of starch grains and forms a dark blue or reddish-brown color, depending on the amylose content of the starch18. Sections stained with I2KI stain show adequate contrast between starch and the background tissue, allowing an unequivocal starch detection and the subsequent quantification by the image analysis system19. Although this dye is not stoichiometric, the accumulation of iodine is proportional to the length of the starch molecule, which can highly vary17. Thus, the size of the stained area expressed as the number of pixels may not reflect accurately the content of starch, since high differences in starch content could be found between fields with stained areas of similar size. As an alternative, the starch content may be evaluated by measuring the optical density of the stained granules on black and white images obtained from the microscope, as it has been reported in different tissues in apricot8,13,19, avocado10,20, and olive21.
Here, we describe a methodology that combines the experimental determination of dormancy status with the quantification of starch content in the ovary primordium tissue from autumn to spring in sweet cherry, offering a new tool for the understanding and prediction of dormancy based on the study of the biological mechanisms linked with dormancy.

<table border="0" cellpadding="0" cellspacing="0" width="767">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Precision scale</td>
			<td style="width:251px;">Sartorius</td>
			<td style="width:133px;">CP225D</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Stereoscopic microscope</td>
			<td style="width:251px;">Leica Microsystems</td>
			<td style="width:133px;">MZ-16</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Drying-stove</td>
			<td style="width:251px;">Memmert</td>
			<td style="width:133px;">U15</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Paraffin Embedding station</td>
			<td style="width:251px;">Leica Microsystems</td>
			<td style="width:133px;">EG1140H</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Rotatory microtome</td>
			<td style="width:251px;">Reichert-Jung</td>
			<td style="width:133px;">1130/Biocut</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Microtome blade</td>
			<td style="width:251px;">Feather</td>
			<td style="width:133px;">S35</td>
			<td>Stainless steel</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Bright field microscope</td>
			<td style="width:251px;">Leica Microsystems</td>
			<td style="width:133px;">DM2500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Digital Camera</td>
			<td style="width:251px;">Leica Microsystems</td>
			<td style="width:133px;">DC-300</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Image Analysis System</td>
			<td style="width:251px;">Leica Microsystems</td>
			<td style="width:133px;">Quantiment Q550</td>
			<td></td>
		</tr>
	</tbody>
</table>

combining histochemical staining and image analysis to quantify starch in the ovary primordia of sweet cherry during winter dormancy

Changes in starch in small structures are associated with key events during several plant developmental processes, including the reproductive phase from pollination to fertilization and the onset of fruiting. However, variations in starch during flower differentiation are not completely known, mainly due to the difficulty of quantifying the starch content in the particularly small structures of the flower primordia. Here, we describe a method for the quantification of starch in the ovary primordia of sweet cherry (Prunus avium L.) by using an image analysis system attached to the microscope, which allows relating the changes in starch content with the different phases of dormancy from autumn to spring. For this purpose, the dormancy status of flower buds is determined by evaluating the bud growth of shoots transferred to controlled conditions at different moments in winter time. For the quantification of starch in the ovary primordia, flower buds are sequentially collected, fixed, embedded in paraffin wax, sectioned, and stained with I2Kl (potassium iodide-iodine). Preparations are observed under the microscope and analyzed by an image analyzer that clearly distinguishes starch from the background. Starch content values are obtained by measuring the optical density of the image that corresponds to the stained starch, considering the sum of the optical density of each pixel as an estimation of the starch content of the frame studied.

Dormancy studies require the determination of the moment when the chilling requirements are fulfilled. Despite the lack of phenological changes during winter under field conditions (Figure 1A), cherry trees do not recover the capacity of growth in suitable conditions until they pass a certain period under low temperatures. The regular transference of shoots to a controlled conditions chamber (Figure 1B) during winter time allowed...

Watch this Scientific Journal Video about Combining Histochemical Staining and Image Analysis to Quantify Starch in the Ovary Primordia of Sweet Cherry during Winter Dormancy at JoVE.com

Combining Histochemical Staining and Image Analysis to Quantify Starch in the Ovary Primordia of Sweet Cherry during Winter Dormancy

We present a methodology to quantify the starch content in the ovary primordia in sweet cherry (Prunus avium L.) during winter dormancy by using an image analysis system combined with histochemical techniques.

Changes in starch in small structures are associated with key events during several plant developmental processes, including the reproductive phase from ...

combining-histochemical-staining-image-analysis-to-quantify-starch

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

6.2K Views.  Centro de Investigación y Tecnología Agroalimentaria de Aragón. We present a methodology to quantify the starch content in the ovary primordia in sweet cherry (Prunus avium L.) during winter dormancy by using an image analysis system combined with histochemical techniques.

Video: Combining Histochemical Staining and Image Analysis to Quantify Starch in the Ovary Primordia of Sweet Cherry during Winter Dormancy

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

The evolutionarily conserved&#160;extracellular signal transducing RTK-RAS-ERK pathway is an important kinase-signaling cascade that controls multiple cellular and developmental processes principally via activation of ERK, the terminal kinase of the pathway. Tight regulation of ERK activity is essential for normal development and homeostasis; overly active ERK results in excessive cellular proliferation, while underactive ERK causes cell death. C. elegans is a powerful model system that has helped characterize the function and regulation of RTK-RAS-ERK signaling pathway during development. In particular, the RTK-RAS-ERK pathway is essential for C. elegans germline development, which is the focus of this method. Using antibodies specific to the active, diphosphorylated form of ERK (dpERK), the stereotypical localization pattern can be visualized within the germline. Because this pattern is both spatially and temporally controlled, the ability to reproducibly assay dpERK is useful to identify regulators of the pathway that affect dpERK signal duration and amplitude and thus germline development. Here we demonstrate how to successfully dissect, stain, and image dpERK within the C. elegans gonad. This method can be adapted for spatial localization of any signaling or structural protein in the C. elegans gonad, provided an antibody compatible with immunofluorescence is available.

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

We present an immunofluorescence&#160;imaging-based method for spatial and temporal localization of active ERK in the dissected C. elegans gonad. The protocol described here can be adapted for visualization of any signaling or structural protein in the C. elegans gonad, provided a suitable antibody reagent is available.

The evolutionarily conserved&#160;extracellular signal transducing RTK-RAS-ERK pathway is an important kinase-signaling cascade that controls multiple ...

Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and scanning electron microscopy. The sub-diffraction limit resolution capabilities of super-resolution light microscopes allow improving the accuracy of the correlated data. Briefly, 110 nanometer thick cryo-sections are transferred to a silicon wafer and, after immunofluorescence staining, are imaged by super-resolution light microscopy. Subsequently, the sections are preserved in methylcellulose and platinum shadowed prior to imaging in a scanning electron microscope (SEM). The images from these two microscopy modalities are easily merged using tissue landmarks with open source software. Here we describe the adapted method for the larval zebrafish retina. However, this method is also applicable to other types of tissues and organisms. We demonstrate that the complementary information obtained by this correlation is able to resolve the expression of mitochondrial proteins in relation with the membranes and cristae of mitochondria as well as to other compartments of the cell.

This protocol describes the necessary steps to obtain subcellular protein localization results on zebrafish retina by correlating super-resolution light microscopy and scanning electron microscopy images.

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

Studying the cell biological processes during converting the identities of specific cell types provides important insights into mechanism that maintain and protect cellular identities. The conversion of germ cells into specific neurons in the nematode Caenorhabditis elegans (C. elegans) is a powerful tool for performing genetic screens in order to dissect regulatory pathways that safeguard established cell identities. Reprogramming of germ cells to a specific type of neurons termed ASE requires transgenic animals that allow broad over-expression of the Zn-finger transcription factor (TF) CHE-1. Endogenous CHE-1 is expressed exclusively in two head neurons and is required to specify the glutamatergic ASE neurons fate, which can easily be visualized by the gcy-5prom::gfp reporter. A trans gene containing the heat-shock promoter-driven che-1 gene expression construct allows broad mis-expression of CHE-1 in the entire animal upon heat-shock treatment. The combination of RNAi against the chromatin-regulating factor LIN-53 and heat-shock-induced che-1 over-expression leads to reprogramming of germ cell into ASE neuron-like cells. We describe here the specific RNAi procedure and appropriate conditions for heat-shock treatment of transgenic animals in order to successfully induce germ cell to neuron conversion.

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

This protocol describes how to study cellular processes during cell fate conversion in Caenorhabditis elegans in vivo. Using transgenic animals, allowing heat-shock promoter-driven overexpression of the neuron fate-inducing transcription factor CHE-1 and RNAi-mediated depletion of the chromatin-regulating factor LIN-53 germ cell to neuron reprogramming can be observed in vivo.

Studying the cell biological processes during converting the identities of specific cell types provides important insights into mechanism that maintain ...

Ovarian Tissue Culture to Visualize Phenomena in Mouse Ovary

Mammalian females periodically ovulate an almost constant number of oocytes during each estrus cycle. To sustain such regularity and periodicity, regulation occurs at the hypothalamic-pituitary-gonadal axis level and on developing follicles in the ovary. Despite active studies, follicle development mechanisms are not clear because of the several steps involved from the dormant primordial follicle activation to ovulation, and because of the regulation complexity that differs at each follicular stage. To investigate the mechanisms of follicle development, and the dynamics of follicles throughout the estrus cycle, we developed a mouse ovarian tissue culture model that can be used to observe follicle development using a microscope. Systematic follicle development, periodical ovulation, and follicle atresia can all be reproduced in the cultured ovary model, and the culture conditions can be experimentally modulated. Here, we demonstrate the usefulness of this method in the study of the regulatory mechanisms of follicle development and other ovarian phenomena.

Ovarian tissue cultures can be used as models of follicle development, ovulation, and follicle atresia and indicate regulatory mechanisms of dynamic ovarian processes.

Mammalian females periodically ovulate an almost constant number of oocytes during each estrus cycle. To sustain such regularity and periodicity, ...

A Hyperandrogenic Mouse Model to Study Polycystic Ovary Syndrome

Hyperandrogenemia plays a critical role in reproductive and metabolic function in females and is the hallmark of polycystic ovary syndrome. Developing a lean PCOS-like mouse model that mimics women with PCOS is clinically meaningful. In this protocol, we describe such a model. By inserting a 4 mm length of DHT (dihydrotestosterone) crystal powder pellet (total length of pellet is 8 mm), and replacing it monthly, we are able to produce a PCOS-like mouse model with serum DHT levels 2 fold higher than mice not implanted with DHT (no-DHT). We observed reproductive and metabolic dysfunction without changing body weight and body composition. While exhibiting a high degree of infertility, a small subset of these PCOS-like female mice can get pregnant and their offspring show delayed puberty and increased testosterone as adults. This PCOS-like lean mouse model is a useful tool to study the pathophysiology of PCOS and the offspring from these PCOS-like dams.

We describe the development of a lean PCOS-like mouse model with&#160;dihydrotestosterone pellet to study the pathophysiology of PCOS and the offspring from these PCOS-like dams.

Hyperandrogenemia plays a critical role in reproductive and metabolic function in females and is the hallmark of polycystic ovary syndrome. Developing a ...

An In Vitro 3D Model and Computational Pipeline to Quantify the Vasculogenic Potential of iPSC-Derived Endothelial Progenitors

Induced pluripotent stem cells (iPSCs) are a patient-specific, proliferative cell source that can differentiate into any somatic cell type. Bipotent endothelial progenitors (EPs), which can differentiate into the cell types necessary to assemble mature, functional vasculature, have been derived from both embryonic and induced pluripotent stem cells. However, these cells have not been rigorously evaluated in three-dimensional environments, and a quantitative measure of their vasculogenic potential remains elusive. Here, the generation and isolation of iPSC-EPs via fluorescent-activated cell sorting are first outlined, followed by a description of the encapsulation and culture of iPSC-EPs in collagen hydrogels. This extracellular matrix (ECM)-mimicking microenvironment encourages a robust vasculogenic response; vascular networks form after a week of culture. The creation of a computational pipeline that utilizes open-source software to quantify this vasculogenic response is delineated. This pipeline is specifically designed to preserve the 3D architecture of the capillary plexus to robustly identify the number of branches, branching points, and the total network length with minimal user input.

Endothelial progenitors derived from induced pluripotent stem cells (iPSC-EPs) have the potential to revolutionize cardiovascular disease treatments and to enable the creation of more faithful cardiovascular disease models. Herein, the encapsulation of iPSC-EPs in three-dimensional (3D) collagen microenvironments and a quantitative analysis of these cells&#8217; vasculogenic potential are described.

Induced pluripotent stem cells (iPSCs) are a patient-specific, proliferative cell source that can differentiate into any somatic cell type. Bipotent ...

Combining Non-reducing SDS-PAGE Analysis and Chemical Crosslinking to Detect Multimeric Complexes Stabilized by Disulfide Linkages in Mammalian Cells in Culture

The structures of many proteins are stabilized through covalent disulfide linkages. In recent work, this bond has also been classified as a post-translational modification. Thus, it is important to be able to study this modification in living cells. A simple method to analyze these cysteine-stabilized multimeric complexes is through a two-step method of non-reducing SDS-PAGE analysis and formaldehyde cross-linking. This two-step method is advantageous as the first step to uncovering multimeric complexes stabilized by disulfide linkages due to its technical ease and low cost of operation. Here, the human bone osteosarcoma cell line U-2 OS is used to illustrate this method by specifically analyzing the nuclear isoform of dUTPase.

Disulfide linkages have long been known to stabilize the structure of many proteins. A simple method to analyze multimeric complexes stabilized by these linkages is through non-reducing SDS-PAGE analysis. Here, this method is illustrated by analyzing the nuclear isoform of dUTPase from the human bone osteosarcoma cell line U-2 OS.

Combining Histochemical Staining and Image Analysis to Quantify Starch in the Ovary Primordia of Sweet Cherry during Winter Dormancy

Summary

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Chapters in this video

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

Ovarian Tissue Culture to Visualize Phenomena in Mouse Ovary

A Hyperandrogenic Mouse Model to Study Polycystic Ovary Syndrome

An In Vitro 3D Model and Computational Pipeline to Quantify the Vasculogenic Potential of iPSC-Derived Endothelial Progenitors

Combining Non-reducing SDS-PAGE Analysis and Chemical Crosslinking to Detect Multimeric Complexes Stabilized by Disulfide Linkages in Mammalian Cells in Culture