Name: Immunostaining and Dye Penetration Experiments to Define Core Pleated Septate Junction Proteins in Drosophila Embryonic Epithelia (Video) | JoVE Journal
Uploaded: 2026-02-27T14:25:00.000+00:00
Duration: 9 min 37 s
Description: Learn immunostaining and dye penetration methods to evaluate pleated septate junction organization and barrier function in Drosophila embryonic epithelia. These protocols enable identification of core pSJ components.

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5 minutes

We thank the Bloomington Drosophila Stock Center for the fly stocks used in this study. We thank the Developmental Studies Hybridoma Bank for antibodies used in the study. We also thank the Department of Biology at Case Western Reserve University for the use of their Leica MZ10F fluorescence stereomicroscope for genotyping and staging embryos for dissection and their DMi8 inverted fluorescence microscope for the dye injection experiments. We thank Helen Salz, Professor of Genetics and Genome Science, CWRU School of Medicine for providing the micromanipulator and for pulling several of the needles used in the dye injection experiment presented in this study. This project was supported by a grant from the National Science Foundation (IOS 2111069) to REW.

1. Staging of embryos
<ol>
	<li>Cross healthy male and female flies of the appropriate genotypes (see Table of Materials for examples) and place them in embryo collection cages with fresh apple juice plates smeared with yeast paste. Let the flies acclimate to egg collection cages for a day or two before starting embryo collections.</li>
	<li>Collect embryos on apple juice plates and allow them to develop to stage 16 (~13 h after egg laying (AEL) to 16 h AEL at 25 &#176;C). Allow the adults to lay fertilized eggs for 1-2 h on apple juice plates, remove the plate from the egg collection cage, and then age them to 13-16 AEL at 25 &#176;C. Alternatively, collect embryos overnight at 25 &#176;C and then select stage 16 embryos prior to fixation or during the imaging after fixation and immunostaining. 
	NOTE: Embryonic staging can be assessed by gut morphology, which is most easily visualized by autofluorescence in the UV spectrum (i.e., DAPI channel). In stage 16 embryos, the gut appears as three parallel, long ovals perpendicular to the body&#39;s long axis (Figure 1H).</li>
</ol>
2. Fixation and immunostaining of embryos
<ol>
	<li>Assemble an embryo collection basket with a nylon mesh (120 mm mesh). Cover the opening and the basket&#39;s threads with the nylon mesh, then screw on the cap.</li>
	<li>Gently dislodge the embryos from the apple juice plate with a paintbrush and embryo wash solution. Pour the embryo mix into the embryo collection basket and rinse the embryos with distilled water. 
	NOTE: Be careful to direct the water down the inside of the embryo collection basket and avoid splashing the embryos onto the basket sides, which can result in embryo loss due to adhesion.</li>
	<li>Dechorionate the embryos by immersing the embryo collection basket and embryos into a tray containing 50% commercial bleach (final concentration of 3.75% sodium hypochlorite). Swirl the embryos in the bleach and keep them immersed for 3-5 min at room temperature. Remove the basket from the bleach and rinse the embryos with distilled water, again directing the flow of water down the inside of the basket.</li>
	<li>Prepare a fixation vial by adding equal volumes of fresh 4% paraformaldehyde solution and heptane to a 20 mL scintillation vial. Make paraformaldehyde solution by dissolving the appropriate amount of paraformaldehyde in hot (~90 &#176;C) 1x phosphate-buffered saline (PBS) in a chemical fume hood. To speed the dissolving process, add 10 &#181;L of 14 N Sodium Hydroxide (per 10 mL of PBS) to the fix until the formaldehyde goes into solution, then add 10 &#181;L of 14 N Hydrochloric acid to adjust the pH back to neutral. 
	CAUTION: Always wear a lab coat, goggles, and gloves when handling paraformaldehyde and heptane, and use only under a chemical fume hood. Paraformaldehyde is a carcinogen and is corrosive, whereas heptane is highly flammable and a skin and eye irritant.</li>
	<li>Disassemble the embryo collection basket and immerse the nylon mesh with dechorionated embryos in the heptane. The embryos will dislodge from the nylon and settle to the interface between the heptane (upper layer) and fix (lower layer). Fix the embryos by shaking the fixation vial on a platform shaker at 240 revolutions per minute (RPM) for 20 min.</li>
	<li>Remove the fix from the fixation vial with a Pasteur pipette in a chemical fume hood. Dispose of the fix in an approved hazardous chemical waste container. Add an equal volume of methanol, close the fixation chamber, and vigorously shake the chamber for 5-10 s. The fixed embryos will tear free from the vitelline envelope and sink to the bottom of the methanol (lower layer). Vitelline envelopes and any embryos that failed to tear free from the vitelline envelope will remain at the interface and will be removed in step 2.7. 
	CAUTION: Always wear a lab coat, goggles, and gloves when handling methanol, as it is toxic and flammable.</li>
	<li>Remove the heptane with a Pasteur pipette and discard it in an approved hazardous chemical waste container. Then, remove most of the methanol, leaving the embryos in a small pool of methanol, and discard the methanol in hazardous chemical waste. Add 5 mL of methanol, swirl the embryos, and let them settle. Remove most of the methanol again and repeat this process 2 more times.</li>
	<li>Transfer the embryos in methanol into a 0.8 mL glass culture tube. Remove the methanol and add 750 mL of block solution (PBS plus 1% normal donkey serum and 0.1% Triton X-100). Wash the embryos 3x in PBS, then incubate them in block solution for 30 min at room temperature with gentle mixing on a platform rocker.</li>
	<li>Rinse the embryos one more time in PBS and then add block plus primary antibodies. Use antibodies against a core pSJ protein (e.g., Coracle) and an adherens junction protein (e.g., E cadherin). Incubate the embryos in primary antibody overnight at 4 &#176;C or for 3-4 h at room temperature with gentle mixing on a platform rocker.</li>
	<li>Remove the block with primary antibodies and rinse the embryos three times in PBS. Add Block and wash the embryos for 30 min at room temperature on a platform rocker.</li>
	<li>Rinse the embryos one more time in PBS, then add the blocking solution and fluorescently labeled secondary antibodies specific to the primary antibodies used in step 2.9. Incubate embryos at room temperature for 3-4 h or overnight at 4 &#176;C.</li>
	<li>Remove the block with secondary antibodies and rinse the embryos three times in PBS. Add Block and wash the embryos for 30 min at room temperature on a platform rocker.</li>
	<li>Rinse the embryos once more in PBS, then transfer them to a microscope slide using a Pasteur pipette. Wick away excess PBS using small strips of Whatman paper, taking care not to touch the embryos. Cover the embryos with mounting media (commercially available or made from 90% glycerol, 10% 1 M Tris, pH 8.0, with 0.5% n-propyl gallate as an antifading agent). Cover with an appropriately sized number 1.5 cover glass, and seal the edges with nail polish.</li>
</ol>
3. Imaging and data analysis of immunostained embryos
<ol>
	<li>For best results, image embryos on a confocal microscope with a 40x oil immersion objective. Use control embryos to set the laser intensity and gain on the confocal to achieve a strong fluorescent signal without oversaturating any pixels. Identify stage 16 embryos of choice (i.e., those not expressing balancer-encoded GFP to identify mutant embryos) using midgut morphology. Focus the scan on the salivary gland or hindgut as these organs are polarized and have long lateral membranes that are ideal for distinguishing the adherens junction, pSJ, and remaining lateral surface. Image in a z position that will include the largest possible diameter of the lumen of the salivary gland or hindgut to observe the full lateral membrane of the epithelial cell. 
	NOTE: The example shown in Figure 1C was taken on a confocal microscope with an HC PL APO 40x/1.3 CS2 oil immersion lens with a laser intensity of 1.59 and a gain of 3.3 for the 561 nm laser and a laser intensity of 2.0 and a gain of 21.7 for the 638 nm laser (see Table of Materials).</li>
	<li>Image a polarized tissue in at least 20 control and mutant stage 16 embryos. Only record data from embryos in which the adherens junction marker is tightly localized to the apical lateral region of the membrane to ensure the embryo was fixed and stained appropriately. Next, record the distribution of the pSJ marker along the lateral membrane. In wildtype stage 16 embryos, look for strong enrichment of pSJ proteins along the ~10-15% of the lateral membrane just basal to the adherens junction. 
	NOTE: Mislocalization of even 10-20% of pSJ protein along the length of the lateral membrane is diagnostic for a pSJ organization defect. pSJ organization defects are strongly penetrant in all core and accessory pSJ mutant embryos, so you should expect to see 75-100% of mutant embryos displaying clearly mislocalized pSJ protein.</li>
</ol>
4. Preparation for dye injection
<ol>
	<li>Make a fresh dye solution by dissolving rhodamine-labeled 10 kDa Dextran to 1 mg/mL in 0.5 M Sodium Phosphate, pH 7.5, and 5 mM Potassium Chloride.</li>
	<li>Pull a capillary tube (1.0 mm OD x 0.5 mm ID with Fiber) into a glass needle using a needle puller. 
	NOTE: Needle pullers are idiosyncratic and settings for ideal needles often differ from day to day. Good needles have a gradual taper to a sharp point and should fall between the shapes shown in Figure 2A. The needle on the left is longer and may bend as it is pushed into the embryo. That can be corrected by rebreaking the needle further from the point. The needle on the right is good but will have a larger bore when initially broken and may push too much liquid into the embryo. If that happens, change to a new needle with a finer point. It is best to pull several needles at once when the conditions on the needle puller create needles of the proper shape. Excess needles can be stored in a closed container with a piece of foam or tape to secure the needles independently.</li>
	<li>Load the needle into the needle holder on a micromanipulator. Arrange the micromanipulator next to an inverted microscope (Figure 2F). Orient the needle as close to parallel to the microscope slide as possible and lower it so that it is in the same z plane as the side of the slide.</li>
	<li>Break the tip of the needle by striking the side of the microscope slide. Raise the needle above the level of the slide and remove it for loading.</li>
	<li>Fill the capillary tube with dye using the smallest pipette tip available (P2 or P10 tip) or by capillary action by placing the back end of the needle in a 1.5 mL microcentrifuge tube with the dye.</li>
	<li>Reload the filled needle into the needle holder and test the flow of the dextran by injecting a bit into a drop of halocarbon oil on the surface of a microscope slide. 
	NOTE: A well broken needle will not leak without adding pressure, and the rate of flow can be adjusted by how much pressure is applied to the plunger on the syringe.</li>
</ol>
5. Mounting embryos and dye injection
<ol>
	<li>Select stage 16 or 17 embryos (control and mutant) using midgut morphology as a staging marker.</li>
	<li>Create an indented line on an apple juice plate using a 22 x 22 # 2 thickness coverslip. Line up approximately 25 staged embryos on the apple juice plate with their posterior ends touching the line and the ventral surface facing up.</li>
	<li>Apply a piece of double-sided tape to a 22 x 22 #2 coverslip. Touch the tape to the embryo line and lightly rub the coverslip to move the embryos from the plate to the double-sided tape on the coverslip (Figure 2B).</li>
	<li>Attach the coverslip to a microscope slide by adding a drop of halocarbon oil to the slide and using capillary action to create a bond. Orient the coverslip closer to the side of the slide opposite to any frosted portion. Desiccate the embryos for 3-5 min in a container of dessicant or in the air on a lab bench for 10-15 min (Figure 2C-E). Cover the embryos completely with Halocarbon oil. 
	NOTE: Embryos will be slightly less turgid after desiccation but will not be strongly flaccid. The embryos shown in Figure 2E were left to air dry for 15 min in a room with 42% relative humidity at 21.2 &#176;C.</li>
	<li>Mount the slide with embryos onto the inverted microscope with the embryos facing up (away from the objective). Align the micromanipulator so that the needle is facing the posterior end of the embryos at an angle as close to parallel to the embryos as possible (Figure 2G). Adjust the z axis of the micromanipulator such that the tip of the needle is aligned to the midpoint of the embryo (along dorsal-ventral axis).</li>
	<li>Inject the embryos by quickly moving the stage into the needle so that the needle tip enters the hemocoel of the embryo. Inject a small amount of dye (~0.2-0.5 nL) using a 10, 25 or 50 mL syringe. Quickly withdraw the needle, move the stage to the next embryo, and repeat the process. Inject as many embryos as possible in 10 min. 
	NOTE: It is important that the needle is inserted about 25% of the way into the embryo so that the dye is not directly injected into the perivitelline space that is continuous with the lumen of the fore- and hindgut. The injected dye should appear to &#34;inflate&#34; the embryo, making it appear more turgid. Overinjection will result in a bubble of dye plus hemolymph exiting the injection site and should be avoided.</li>
	<li>Add additional halocarbon oil and cover the embryos with a 22 x 22 #2 coverslip. The additional halocarbon oil forms a more uniform layer between the two coverslips, preventing the embryos from being crushed.</li>
</ol>
6. Imaging and data analysis
<ol>
	<li>Perform imaging on either a fluorescent compound microscope or a confocal microscope (10x or 20x air objectives are sufficient to yield unambiguous results). On the confocal, z stack imaging through about a quarter of the depth (along dorsal ventral axis) of the embryo will reveal if the labeled dextran has entered the lumen of trachea and hindguts (deeper sections or reorienting the embryos with ventral surface at the top is necessary to see the salivary glands). 
	NOTE: The examples shown in Figure 3 were taken on a confocal microscope with an HC PL APO 10x/0.4 CS2 dry lens with a laser intensity of 2.0 and a gain of 13.6 for the 561 nm laser. 24 z slices were obtained with a z step size of 2.4 nm (see the Table of Materials).</li>
	<li>Allow the injected embryo to age for 10 min and then examine the lumen of the trachea, salivary gland, or hindgut for accumulation of the labeled dextran. Record the genotype and stage of each embryo and whether there is accumulation of labeled dextran into any tubular organ. Note any embryos in which the perivitelline space is also strongly stained with dye as these may represent embryos that were inadvertently injected into that space.</li>
</ol>

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<li>Laprise, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Epithelial+polarity+proteins+regulate+Drosophila+tracheal+tube+size+in+parallel+to+the+luminal+matrix+pathway%5BTitle%5D)+AND+%22Curr.+Biol%22%5BJournal%5D)">Epithelial polarity proteins regulate Drosophila tracheal tube size in parallel to the luminal matrix pathway.</a> Curr. Biol. 20 (1), 55-61 (2010).</li>
<li>Batz, T., Förster, D., Luschnig, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+transmembrane+protein+macroglobulin+complement-related+is+essential+for+septate+junction+formation+and+epithelial+barrier+function+in+Drosophila%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">The transmembrane protein macroglobulin complement-related is essential for septate junction formation and epithelial barrier function in Drosophila.</a> Development. 141 (4), 899-908 (2014).</li>
<li>Behr, M., Riedel, D., Schuh, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+claudin-like+Megatrachea+is+essential+in+septate+junctions+for+epithelial+barrier+function+in%C3%82%C2%A0Drosophila%5BTitle%5D)+AND+%22Dev.+Cell%22%5BJournal%5D)">The claudin-like Megatrachea is essential in septate junctions for epithelial barrier function inÂ Drosophila.</a> Dev. Cell. 5 (4), 611-620 (2003).</li>
<li>Wu, V. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Sinuous+is+a+Drosophila+claudin+required+for+septate+junction+organization+and+epithelial+tube+size+control%5BTitle%5D)+AND+%22J.+Cell+Biol%22%5BJournal%5D)">Sinuous is a Drosophila claudin required for septate junction organization and epithelial tube size control.</a> J. Cell Biol. 164 (2), 313-323 (2004).</li>
<li>Wu, V. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Drosophila+Varicose%2C+a+member+of+a+new+subgroup+of+basolateral+MAGUKs%2C+is+required+for+septate+junctions+and+tracheal+morphogenesis%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">Drosophila Varicose, a member of a new subgroup of basolateral MAGUKs, is required for septate junctions and tracheal morphogenesis.</a> Development. 134 (5), 999-1009 (2007).</li>
<li>Jaspers, M. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+claudin+Megatrachea+protein+complex%5BTitle%5D)+AND+%22J.+Biol.+Chem%22%5BJournal%5D)">The claudin Megatrachea protein complex.</a> J. Biol. Chem. 287 (44), 36756-36765 (2012).</li>
<li>Dietzl, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+genome-wide+transgenic+RNAi+library+for+conditional+gene+inactivation+in+Drosophila%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">A genome-wide transgenic RNAi library for conditional gene inactivation in Drosophila.</a> Nature. 448 (7150), 151-156 (2007).</li>
<li>Zirin, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Large-scale+transgenic+Drosophila+resource+collections+for+loss-+and+gain-of-function+studies%5BTitle%5D)+AND+%22Genetics%22%5BJournal%5D)">Large-scale transgenic Drosophila resource collections for loss- and gain-of-function studies.</a> Genetics. 214 (4), 755-767 (2020).</li>
</ol>

The authors have no conflicts of interest to disclose.

Here we present a protocol to identify core and accessory pSJ proteins using two relatively simple assays that can be achieved in modestly appointed labs with fluorescence microscopy capabilities. The first assay examines the molecular organization of the pSJ in mature polarized epithelia using confocal imaging of fixed mutant embryos. The second assay tests the integrity of the pSJ&#39;s occluding function by injecting dye-labeled 10 kDa dextran molecules into stage 16 living embryos. The simplicity of these protocols a...

A critical function of epithelial tissues is to compartmentalize distinct regions within an organism. Creation of these distinct environments is essential for organ function and for protecting the organism from external threats. The occluding junction formed between the cells of an epithelium is responsible for establishing and maintaining these environments by preventing the paracellular flow of solutes between the apical and basal sides of that epithelium1,2. In vertebrate organisms, the occluding junction is referred to as the tight junction, whereas it is referred to as the septate junction (SJ) in invertebrates such as Drosophila melanogaster1,3. There are two types of SJ in arthropods distinguished by their tissue distribution and ultrastructure4. Pleated septate junctions (pSJs) are found in ectodermally derived tissues such as the epidermis, salivary glands, trachea, and hindgut. They are characterized ultrastructurally by a ladder-like array of electron-dense material in the extracellular space between cells that lies just basal to the adherens junction3,5. In contrast, smooth septate junctions (sSJ) are found in endodermally derived tissues such as midgut and Malpighian tubules and do not show electron-dense septae between cells6.
Genetic studies in Drosophila have identified 30 proteins that are required for pSJ structure and function and have identified additional proteins that reside in the junction but do not appear to have any function in their establishment or maintenance (reviewed in7). From these analyses, scientists have categorized SJ proteins into three classes. Core SJ proteins reside in the junction and are required for the establishment and maintenance of the junction. Seventeen proteins have been identified as Core pSJ proteins, including Coracle (Cora), Neurexin IV, Macroglobulin complement-related (Mcr), Kune-kune, and ATPase alpha8,9,10,11,12. Nine proteins have been identified as accessory pSJ proteins that are required for the establishment and/or maintenance of the junction, but do not necessarily reside in the junction. This class includes proteins such as Rab5, Rab11, and Coiled13,14. Finally, pSJ resident proteins localize to the pSJ but are dispensable for the establishment and maintenance of the junction. Examples of the four members of this group include Discs Large and Fasciclin III15,16.
The establishment of a functionally intact pSJ is a multistep process that begins midway through embryogenesis when all the core pSJ proteins are fully expressed. Some core pSJ genes are expressed maternally, but all show strong zygotic expression by stage 1217. At stage 12, the core pSJ proteins localize all along the length of the lateral membrane. During stages 13-16, core pSJ proteins are endocytosed and then recycled back to the lateral membrane in the region of the pSJ. This process requires pSJ accessory proteins, including Rab5 and Rab1114. By stage 16, the core pSJ proteins are tightly localized to the region of the pSJ along the lateral membrane. Consistent with these observations, the full occluding function of the junction occurs in late stage 15 of embryogenesis12. This is best demonstrated using a dye permeability assay. Prior to the formation of the junction, a 10 kDa dextran bead labeled with a fluorescent marker such as rhodamine can pass through the paracellular space between cells. If the labeled dextran is injected into the hemocoel, it will rapidly accumulate into the lumen of tubular epithelia such as the trachea, salivary gland, and hindgut. After the pSJ is physiologically tight at the end of stage 15, the injected dextran is restricted to the basal side of the epithelium and cannot accumulate in the lumen (apical domain) of these organs.
Core pSJ proteins share a characteristic feature in which each core pSJ protein requires the presence of all other core pSJ proteins for correct subcellular localization18,19. This interdependence in pSJ formation is a defining feature of core pSJ proteins and underlies the protocols presented in this article. Mutations in core or accessory pSJ genes lead to embryonic lethality and are characterized by the mislocalization of other core pSJ proteins along the lateral membrane in stage 16 embryos, rather than being tightly localized to their normal apical lateral region. In contrast, mutations in pSJ-resident genes do not lead to mislocalization of core pSJ proteins15. The functional disruption of the pSJ in core and accessory pSJ gene mutations is revealed by the accumulation of labeled 10 kDa dextran in the lumen of trachea, salivary glands or hindguts in stage 16 or 17 mutant embryos. Finally, the distinguishing feature between a core and accessory pSJ protein is their subcellular localization. Core pSJ proteins are tightly associated with the pSJ once they form, whereas accessory pSJ proteins are critical for assembly of the junction but do not necessarily reside in the junction. The best examples of accessory pSJ proteins are Rab5 and Rab11. Both are cytoplasmic proteins that aid in shuttling cargo from the plasma membrane to early and recycling endosomes and then back to the plasma membrane14. To distinguish whether a new potential pSJ protein is a core component or an accessory protein it is generally helpful to create a specific antibody or express a fluorescently tagged recombinant protein to determine where these proteins localize at various points in the pSJ maturation process.
The purpose of these protocols is to unambiguously distinguish potential core and accessory pSJ proteins from pSJ resident proteins. Several studies have revealed additional, non-occluding roles for pSJ proteins during development, including roles in morphogenesis and apical-basal polarity17,20,21,22. As interest in pSJ biology increases and more potential pSJ proteins are identified, the simple, easily interpretable protocols presented here will aid researchers in correctly characterizing the pSJ organization and occluding functions of these proteins. In the literature, many additional experimental approaches for characterizing pSJ proteins have been described, including Fluorescence Recovery After Photobleaching (FRAP) and transmission electron microscopy6,23. Although these approaches are often very useful, the immunostaining assay presented here provides rapid visualization of pSJ protein localization without expensive and time-consuming ultrastructural methods, whereas the dye permeability assay offers a functional readout of pSJ integrity not achievable by imaging alone. Additionally, these assays can be performed using standard lab equipment such as a good-quality fluorescence microscope and a rudimentary microinjection setup. It should be noted that these assays are optimized for analysis of Drosophila stage 16 embryos and are not appropriate for evaluating the organization or function of sSJs.

<table><tbody><tr><td>Capillary tube for needles (Boro 1 X 0.5/Fiber )</td><td>FHC</td><td>30-30-1</td><td></td></tr><tr><td>Confocal microscope</td><td>Leica</td><td>Stellaris 5&amp;nbsp;</td><td></td></tr><tr><td>Cover glass 22 x 22 2 thickness for injections</td><td>Fisher Scientific</td><td>12-540-B</td><td></td></tr><tr><td>Cover glass 22 x 30 1.5 thickness for imaging</td><td>Corning</td><td>2980-233</td><td></td></tr><tr><td>Culture tube for immunostaining (6 x 50 mm)&amp;nbsp;</td><td>Fisher Scientific</td><td>14-958-A</td><td></td></tr><tr><td>Embryo collection cage</td><td>Genesee Scientific</td><td>59-100</td><td></td></tr><tr><td>Fluorescence stereomicroscope</td><td>Leica</td><td>MZ10 F</td><td></td></tr><tr><td>Halocarbon oil (700)</td><td>Genesee Scientific</td><td>59-131</td><td></td></tr><tr><td>Heptane</td><td>Fisher Scientific</td><td>H350-4</td><td></td></tr><tr><td>Inverted fluorescence microscope</td><td>Leica</td><td>DMi8</td><td></td></tr><tr><td>Mesh baskets for dechorionating embryos</td><td>Genesee Scientific</td><td>46-101</td><td></td></tr><tr><td>Methanol</td><td>Fisher Scientific</td><td>A412-4</td><td></td></tr><tr><td>Micromanipulator</td><td>Narishige</td><td>MN151</td><td></td></tr><tr><td>Microscope slides</td><td>Fisher Scientific</td><td>12-544-2</td><td></td></tr><tr><td>Needle puller</td><td>David Kopf instruments</td><td>model 720</td><td></td></tr><tr><td>Normal Donkey Serum</td><td>Jackson ImmunoReseaerch</td><td>017-000-121</td><td></td></tr><tr><td>Nylon mesh (120 um)</td><td>Genesee Scientific</td><td>57-102</td><td></td></tr><tr><td>Objective lens HC PL FLUOTR 10X/0.32 PH 1</td><td>Leica</td><td>11506537</td><td></td></tr><tr><td>Objective lens HC PL APO 10x/0.40 CS2</td><td>Leica</td><td>15506407</td><td></td></tr><tr><td>Objective lens HC PL APO 40x/1.30 CS2</td><td>Leica</td><td>15506358</td><td></td></tr><tr><td>Petri plates 60 x 15 mm</td><td>Corning</td><td>351007</td><td></td></tr><tr><td>Platform rocker</td><td>Globe Scientific</td><td>GTR-FS</td><td></td></tr><tr><td>Primary antibody (Coracle)</td><td>DSHB</td><td>C566.9</td><td></td></tr><tr><td>Primary antibody (Coracle)</td><td>DSHB</td><td>C615.16</td><td></td></tr><tr><td>Primary antibody (E cadherin)</td><td>DSHB</td><td>DCAD2</td><td></td></tr><tr><td>Primary antibody (Mcr)</td><td>Ward lab, CWRU</td><td>guinea pig polyclonal</td><td></td></tr><tr><td>Rhodamine labeled 10 kDa dextran</td><td>Molecular Probes</td><td>D1824</td><td></td></tr><tr><td>Secondary antibody (donkey anti-guinea pig Cy3)</td><td>Jackson ImmunoReseaerch</td><td>706-165-148</td><td></td></tr><tr><td>Secondary antibody (donkey anti-mouse Cy2)</td><td>Jackson ImmunoReseaerch</td><td>715-225-151</td><td></td></tr><tr><td>Secondary antibody (donkey anti-rat Cy5)</td><td>Jackson ImmunoReseaerch</td><td>712-175-150</td><td></td></tr><tr><td>Table top shaker</td><td>New Brunswick Scientific</td><td>C2 platform shaker</td><td></td></tr><tr><td>Tegosept (methyl 4-hydroxybenzoate)</td><td>TCI</td><td>H0216</td><td></td></tr><tr><td>Triton X-100</td><td>Fisher Scientific</td><td>BP151500</td><td></td></tr><tr><td>Vials for fixation (20 ml WHEATON liquid scintillation vials)</td><td>Fisher Scientific</td><td>03-341-73C</td><td></td></tr><tr><td>Whatman 3MM&amp;nbsp; Chromatography Paper</td><td>Fisher Scientific</td><td>05-716-6H</td><td></td></tr><tr><td>&lt;strong&gt;Fly stocks&lt;/strong&gt;</td><td>&lt;strong&gt;Full genotype&lt;/strong&gt;</td><td>&lt;strong&gt;stock number&lt;/strong&gt;</td><td>&lt;strong&gt;Source&lt;/strong&gt;</td></tr><tr><td>cor4</td><td>w*; P{neoFRT}43D cora4/CyO</td><td>52232</td><td>BDSC</td></tr><tr><td>w1118</td><td>w[1118]</td><td>5905</td><td>BDSC</td></tr><tr><td>&lt;strong&gt;Solution&lt;/strong&gt;</td><td>&lt;strong&gt;Recipe&lt;/strong&gt;</td><td></td><td></td></tr><tr><td>10x PBS</td><td>Mix 90 g of NaCl, 20 g of Na2HPO4, and 8.3 g of NaH2PO4.H2O and dH20 to 1L. Filter sterilize. Dilute to 1X with dH2O for washing steps in immunostaining.</td><td></td><td></td></tr><tr><td>10% Triton X-100 solution</td><td>Mix 1 ml of Triton X-100 to 9 ml of dH2O on a platform rocker until dissolved. Store at room temperature.</td><td></td><td></td></tr><tr><td>50% bleach</td><td>Mix 10 ml of commercial bleach (7.5% sodium hypochlorite) to 10 ml of water prior to dechorionating embryos.</td><td></td><td></td></tr><tr><td>Apple juice plates</td><td>Boil 15 g Drosophila agar in 725 ml dH2O. Add 25 g sugar and 250 ml apple juice and mix until blended. Let the solution cool to 55 C and then add 1.3 g tegosept and gently mix. Pour into 60 x 15 petri plates and let cool. Store at 4 C.</td><td></td><td></td></tr><tr><td>Blocking solution</td><td>Mix 9.8 ml of 1X PBS with 100 ul of 10% Triton X-100 and 100 ul of Normal Donkey Serum in a 15 conical tube.</td><td></td><td></td></tr><tr><td>Embryo wash solution</td><td>Make 10X embryo wash solution by mixing 70 g NaCl and 2 ml Triton X-100 to 1L of dH2O. Dilute to 1X with dH2O for working solution.</td><td></td><td></td></tr><tr><td>Fix</td><td>Preheat 10 ml of 1X PBS in the microwave. Add 0.4 g&amp;nbsp; of paraformaldehyde and pipet to mix. Add 10 ul of 14N NaOH to speed dissolving. Place on a 90 C heatblock until dissolved. Once no solid paraformaldehyde remains add 10 ul of 14N HCL to adjust pH to neutral. Always wear a lab coat, gloves and googles when using paraformaldehyde.</td><td></td><td></td></tr><tr><td>Mounting media</td><td>Mix 9 ml of glycerol and 1 ml of 1M Tris base pH 8.0. Add 0.05 g n-propyl-gallate,&amp;nbsp; mix and heat to dissolve. Aliquot 0.5 ml into eppendorf tubes.&amp;nbsp;</td><td></td><td></td></tr><tr><td>Rhodamine dextran solution</td><td>Make fresh dye solution by dissolving Rhodamine-labeled 10 kDa Dextran to 1 mg/ml in 0.5M Sodium Phosphate, pH 7.5 and 5 mM Potassium Chloride.</td><td></td><td></td></tr><tr><td>Yeast paste</td><td>Mix 50 g of commercial yeast (Red Star) with a pinch of sucrose and enough water to make a thick paste. Store at 4 C.</td><td></td><td></td></tr></tbody></table>

immunostaining and dye penetration experiments to define core pleated septate junction proteins in drosophila embryonic epithelia

Immunostaining and Dye Penetration Experiments to Define Core Pleated Septate Junction Proteins in Drosophila Embryonic Epithelia

The purpose of these protocols is to evaluate the organization and barrier function of pleated Septate Junctions (pSJs) in the ectodermally-derived epithelial tissues of Drosophila embryos. pSJ form an occluding barrier on the lateral membrane of ectodermal tissues including the epidermis, salivary glands, trachea, and hindgut that is functionally similar to the tight junction in vertebrate tissues. In Drosophila, 30 proteins have been identified that are required for the formation of pSJs. Core pSJ proteins are initially localized along the length of the lateral membrane but become strongly enriched at the apical lateral region of the junction, just basal to the adherens junction, by stage 16 of embryogenesis. By late stage 15 of embryogenesis, these tissues possess a fully functional occluding junction. This paper will describe immunostaining and dye permeability protocols that can be used to interrogate the organization and function of pSJs in control and mutant embryos. Successful employment of these protocols can be used to demonstrate the occluding function of pSJs, and to test whether an unknown gene encodes a core component of the junction.

The protocols presented here are used to investigate the organization and function of pSJs in ectodermal tissues in stage 16 Drosophila embryos. These protocols are only applicable to pSJs and cannot be used to interrogate sSJs. A mutation in a gene that results in failure to localize other pSJ proteins to the pSJ region in stage 16 embryos and to create an occluding junction at stage 16 is either in a core pSJ gene or an accessory pSJ gene. The main distinction between the two is whether the encoded protein is subcellul...

These protocols outline how to characterize the organization and occluding function of pleated septate junctions in control and mutant Drosophila embryonic epithelia.

The purpose of these protocols is to evaluate the organization and barrier function of pleated Septate Junctions (pSJs) in the ectodermally-derived ...

Our research investigates the developmental nonoccluding functions of septate junction proteins, focusing on identifying and characterizing new components involved in the structure and biological roles. This protocol is meant to characterize the organization and occluding function of pleated septate junctions in the ectodermal epithelia of Drosophila embryos. To begin, cross healthy male and female Drosophila flies of the appropriate genotypes, and place them in embryo collection cages with fresh apple juice plates smeared with yeast paste.Allow the adults to lay fertilized eggs on the apple juice plates. Remove the plate from the egg collection cage and age the embryos for 13 to 16 hours after egg-laying at 25 degrees Celsius to reach stage 16. Then fix the stage 16 embryos in an appropriate fixative solution and perform immunostaining with the desired antibodies.For best results, image the embryos on a confocal microscope with a 40x oil immersion objective. After focusing on the embryo of interest, set the laser intensity and gain to achieve a strong fluorescent signal without oversaturating any pixels. Identify stage 16 embryos that do not express balancer-encoded green fluorescent protein, using midgut morphology.Focus the scan on the salivary gland or hindgut, and position the Z-scan to include the largest possible diameter of the lumen to visualize the full lateral membrane of the epithelial cell. Set a suitable line average and image a polarized tissue in at least 20 control and 20 Cora 4 mutant stage 16 embryos. Record data only from embryos in which the adherence junction marker is tightly localized to the apical-lateral region of the membrane.In wild-type stage 16 embryos, assess enrichment of pleated septate junction proteins along the apical region of the lateral membrane near the adherens junction. Record the persistent distribution of the pleated septate junction marker all along the lateral membrane in the mutant. Make a fresh dye solution by dissolving rhodamine-labeled 10 kilodalton dextran to 1 milligram per milliliter in 0.5 molar sodium phosphate at pH 7.5, with 5 millimolar potassium chloride.Using a needle puller, pull a 1-millimeter outer diameter by 0.5-millimeter inner diameter capillary tube with fiber, into a glass needle. Next, load the glass needle into the needle holder on a micromanipulator and arrange the micromanipulator next to an inverted microscope. Orient the needle as close to parallel to the microscope slide as possible, and lower it to the same Z-plane as the side of the slide.Break the tip of the needle by striking it against the side of the microscope slide. Raise the needle above the level of the slide and remove it for loading. Next, fill the capillary tube with dye using the smallest available pipette tip.Reload the filled needle into the needle holder. Test the flow of the 10 kilodalton dextran by injecting a small amount into a drop of halocarbon oil on the surface of a microscope slide. Using a 22 x 22 millimeter number 2 thickness cover slip, create an indented line on an apple juice plate.Line up approximately 25 embryos each of control and Cora 4 mutant at stage 16 or 17 on the apple juice plate. Ensure that their posterior ends align along the line, and that the ventral surfaces face upward. Then apply a piece of double-sided tape to a 22 x 22 millimeter number 2 cover slip.Touch the tape to the embryo line and lightly rub the cover slip to transfer the embryos from the plate onto the double-sided tape. Next, add a drop of halocarbon oil to a microscope slide and attach the cover slip to the slide by allowing capillary action to create a bond. Orient the cover slip closer to the side of the slide opposite any frosted portion.Desiccate the embryos for three to five minutes in a container of desiccant, and cover the embryos completely with halocarbon oil. Mount the slide with embryos onto the inverted microscope with the embryos facing up and away from the objective. Align the micromanipulator so that the needle faces the posterior end of the embryos at an angle as close to parallel to the embryos as possible, and adjust the Z-axis of the micromanipulator so that the needle tip aligns with the midpoint of the embryo along the dorsal-ventral axis.Next, inject the embryos by quickly moving the stage into the needle so that the needle tip enters the hemocoel. Inject approximately 0.2 to 0.5 nanoliters of dye using a 10-25-or 50-milliliter syringe. Quickly withdraw the needle and move the stage to the next embryo for dye injection.Then add additional halocarbon oil over the embryos. Cover the embryos with a 22 x 22 millimeter number 2 cover slip to create a uniform oil layer between the cover slips and prevent crushing. Perform imaging on a fluorescent compound microscope or a confocal microscope after dextran dye injection.On the confocal microscope, acquire a Z-stack through approximately one quarter of the embryo depth along the dorsal-ventral axis to assess the lumen of the trachea. Examine the lumen of the trachea, a salivary gland, or hindgut for accumulation of labeled dextran. Record the genotype and developmental stage of each embryo and whether labeled dextran accumulates in any tubular organ.Note embryos in which the perivitelline space is strongly stained with dye, as these may represent inadvertent injection into that space. In wild-type embryos, macroglobulin complement-related, or Mcr, was uniformly localized along the lateral membrane at stage 13, became enriched in the apical region of the lateral membrane at stage 14, and was strongly localized to the apical-lateral region, just basal to the adherens junction by stage 16. Cortical antibody staining defined the mature pleated septate junctions in stage 16 wild-type embryos similarly to Mcr.In Cora 4 mutant embryos, Mcr failed to achieve strong apical localization by stage 16 and remained distributed along the lateral membrane. E-cadherin localization was similar in wild-type and Cora 4 mutant stage 16 embryos. In wild-type stage 17 embryos injected with rhodamine-labeled 10 kilodalton dextran, the dye surrounded the trachea but did not penetrate the lumen.In Cora 4 mutant stage 17 embryos, the dye accumulated in the lumen of the trachea within 10 minutes of injection. This protocol allows researchers to confidently determine whether a gene encodes the core or the accessory component of the pleated septate junctions in the Drosophila embryonic tissues. Embryo staging is absolutely crucial while performing this technique, as this requires stage 16 embryos.The septate junctions matures over time until it is fully established in stage 16. We believe this protocol can be effectively implemented in open-ended genetic screening using RNA interference lines to identify new pleated septate junction genes.

immunostaining-dye-penetration-experiments-to-define-core-pleated

Research

JoVE Journal

Developmental Biology

This protocol demonstrates how to perform immunohistochemistry on dissected Drosophila larva.

Drosophila Larval NMJ Immunohistochemistry

The neuromuscular junction (NMJ) of Drosophila melanogaster is an important model system for studying normal synaptic function as well as perturbations to synaptic function found in certain neurological diseases. We present a protocol for dissection of the Drosophila larval motor system and immunostaining for active zone proteins within the NMJ.

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

Immunostaining is an effective technique for visualizing specific cell types and proteins within tissues. By utilizing sonication, the protocol described here alleviates the need to dissect Drosophila melanogaster tissues from late-stage embryos and larvae before immunostaining. We provide an efficient methodology for the immunostaining of formaldehyde-fixed whole mount larvae.

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

The adult structures of Drosophila are derived from sac-like structures called imaginal discs. Analysis of these discs provides insight into many developmental processes including tissue determination, compartment boundary establishment, cell proliferation, cell fate specification, and planar cell polarity. This protocol is used to prepare imaginal discs for light/fluorescent microscopy.

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

This protocol demonstrates how Proximity Ligation Assay can be used to detect in situ protein-protein interactions at the Drosophila larval neuromuscular junction. With this technique, Discs large and Hu-li tai shao are shown to form a complex at the postsynaptic region, an association previously identified through co-immunoprecipitation.

Detection of In Situ Protein-protein Complexes at the Drosophila Larval Neuromuscular Junction Using Proximity Ligation Assay

A protocol of whole-mount immunostaining on aphid embryos is presented, in which critical conditions for decreasing background staining and increasing tissue permeability are addressed. These conditions are specifically developed for effective detection of protein expression in the embryonic tissues of aphids.

Identification of Critical Conditions for Immunostaining in the Pea Aphid Embryos: Increasing Tissue Permeability and Decreasing Background Staining

We present here an improved protocol for the whole-mount preparation and immunostaining of Drosophila testes, suitable for confocal microscopy and also allowing reproducible and reliable labeling. We illustrate this protocol by 3D representation and quantification of colocalization experiments on the large S4-S5 spermatocytes.

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

Here, we describe a protocol for detection and localization of Drosophila embryo protein and RNA from collection to pre-embedding and embedding, immunostaining, and mRNA in situ hybridization.

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

The present protocol details the preparation and visualization of fixed tissue of the Drosophila pupal notum. It can be used for either intact or wounded tissue, and the original architecture of the tissue is preserved. The procedures for dissecting, fixing, and staining are all described in this article.

Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum

Here, we present a detailed protocol to visualize the microtubule networks in neuromuscular junctions and muscle cells. Combined with the powerful genetic tools of Drosophila melanogaster, this protocol greatly facilitates genetic screening and microtubule-dynamics analysis for the role of microtubule network regulatory proteins in the nervous system.

Using Drosophila Larval Neuromuscular Junction and Muscle Cells to Visualize Microtubule Network

This protocol describes the customized antibody-based fluorescence labeling and injection into early Drosophila embryos to enable live imaging of low-abundance proteins or post-translational modifications that are challenging to detect using traditional GFP/mCherry-tag approaches.

Visualizing Low-Abundance Proteins and Post-Translational Modifications in Living Drosophila Embryos via Fluorescent Antibody Injection

Learn immunostaining and dye penetration methods to evaluate pleated septate junction organization and barrier function in Drosophila embryonic epithelia. These protocols enable identification of core pSJ components.

Immunostaining and Dye Penetration Experiments to Define Core Pleated Septate Junction Proteins in Drosophila Embryonic Epithelia (Video) | JoVE Journal

Immunostaining and Dye Penetration Experiments to Define Core Pleated Septate Junction Proteins in Drosophila Embryonic Epithelia

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