In invertebrate epithelial tissues, the septate junction (SJ) forms an occluding barrier that is functionally similar to the tight junction in vertebrate tissues. In mature epithelia, the SJ localizes along the apical lateral membrane just basal to the adherens junction. In Drosophila, more than 30 proteins have been identified as either core components of the junction, accessory proteins that reside in the junction, or non-resident proteins that are required for the formation of the junction. In wild type conditions, SJ proteins are initially localized along the length of the lateral membrane but become enriched in the presumptive SJ by stage 15 of embryogenesis. By late stage 15, the epithelium is physiologically tight as assessed by the inability of a 10 kDa rhodamine-labeled dextran to traverse the paracellular route between the basal side of tubular organs and their lumen. During stages 16 and 17 of embryogenesis, the SJ appears to mature ultrastructurally, as an electron dense ladder-like structure forms between epithelial cells. As new potential SJ proteins are discovered, it is important to identify which ones are core components of the junction. Core SJ proteins show interdependence for subcellular localization, meaning that loss of one component leads to the mislocalization of other core components. Here, we will demonstrate the immunostaining procedures we use to interrogate whether a mutation in a novel gene disrupt the structure and function of the SJ. We will subsequently describe additional experiments including Fluorescence Recovery After Photobleaching (FRAP) and transmission electron microscopy that are often used to confirm these observations.

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>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 and the fact that a defect in pSJ organization and function is completely diagnostic for core and accessory pSJs mutations make them advantageous over more labor-intensive, technically challenging approaches, such as FRAP and electron microscopy. These protocols are specific to the organization and function of pSJ during embryogenesis in Drosophila and do not apply to sSJs or to later developmental stages.
The mislocalization of all core pSJ proteins in core and accessory pSJ mutant late-stage embryos is completely diagnostic. Therefore, it is possible to test any potential pSJ gene by simply immunostaining mutant embryos at stage 16 with an antibody against a core pSJ protein. The protocol we present here includes a very standard paraformaldehyde fixation step. Slight variations from this protocol should be fine so long as control stage 16 embryos show the expected localization of adherens junction and pSJ markers. Several useful pSJ antibodies are available from the Developmental Studies Hybridoma Bank (DSHB) including Cora, ATPase alpha, and Nervana. There are two antibodies against Cora: one against the amino terminus of the protein and the other against the carboxy terminus. We typically mix both Cora antibodies equally for use in these studies. Cora is a cytoplasmic protein but localizes tightly to the membrane through its interaction with the cytoplasmic domain of Neurexin IV in ectodermally-derived epithelial cell19. We also often use an antibody against Mcr since it was made in guinea pigs10 and allows for co-staining with antibodies made in mice, rabbits, and rats. E-cadherin is a very useful counterstain as a marker for the adherens junction. Rat antibodies against E-cadherin are available from the DSHB, as are rat antibodies made against alpha catenin, another adherens junction protein. Although we used confocal imaging for the analysis shown here, the spreading of pSJ proteins along the lateral membrane is obvious in the polarized epithelia of salivary glands and hindguts and can be easily identified with any fluorescent imaging. An important consideration for these studies is embryonic timing. Because the pSJ matures over time, it is critical to analyze core pSJ localization in stage 16 embryos. The deposition of cuticle prevents immunostaining at later stages of development, and so there is only a 2-3-h window (at 25 &#176;C) when mutant embryos can be examined. Since the mislocalization of pSJ protein is so highly penetrant in pSJ mutant stage 16 embryos, finding an intermediate penetrance of the phenotype in an unknown mutation would be suspicious and would necessitate confirming the developmental staging of the embryos. Many pSJ mutant embryos show defects in head involution and dorsal closure and develop at slightly slower rates than wild type embryos17 and so comparable timing to control embryos or staging based upon head morphology should be avoided. Instead, use developmental timing based upon midgut morphology24. Most pSJ proteins are not expressed in the midgut or mesoderm and mutations in pSJ genes do not appear to alter midgut morphology.
The main function of the pSJ is to act as a physical barrier to prevent the paracellular flow of solutes between the basal and apical sides of an ecdoermally-derived epithelium. Previous work has shown that the size of this paracellular blockade will exclude molecules as small as 10 kDa25. We can therefore use a fluorescently tagged 10 kDa dextran to interrogate the integrity of the pSJ in early and late-stage embryos. The assay is simple, but there are a few important considerations. Prior to injecting the labeled dextran, the embryos must be slightly desiccated so that they will not burst upon injection. The time and conditions (in the air versus in a chamber filled with desiccant) should be empirically determined and will depend on the room&#39;s relative humidity and temperature. The injection must also be made about a quarter of the way into the body cavity from the posterior end, and the dye should not be delivered until the needle has penetrated that far into the embryo. The primary reason for this caution is that the perivitelline space is continuous with the fore- and hindgut and the external spiracles, and thus dye that gets into the perivitelline space can enter the lumen of the salivary glands, trachea, and hindguts. It is nearly impossible to prevent any dye from entering the perivitelline space, so some accumulation in these lumens will occur over time. That is why it is also important to inject only as many embryos as can be accomplished in 10 min and then assess the luminal accumulation of dye at that point. In stage 13-14 wild-type embryos or stage 16-17 SJ mutant embryos, the dye quickly penetrates into the lumen of the trachea, salivary gland, and hindgut. This is in stark contrast to the lack of accumulation in stage 16 or 17 wildtype embryos, after the SJ has formed. Finding an intermediate penetrance of leaky pSJs in a mutant line would be suspicious and would necessitate verifying the genotype to eliminate heterozygous embryos and ensure there was no strong leak into the perivitelline space. In the examples presented here, we have visualized the embryo to show the tracheal lumen, but the salivary gland and hindgut are tubular organs that can also be easily assessed. Since the salivary glands lie closer to the ventral surface of the embryo, mounting the embryos with the ventral surface facing up prior to injection would make imaging them easier. We also used a confocal microscope to image the trachea, but the phenotype is completely diagnostic and easy to score on any fluorescent microscope. For these experiments, we mounted the micromanipulator adjacent to an inverted fluorescent compound microscope, but it is possible to do these injections on simpler microscopes, including dissecting stereomicroscopes26.
Finally, we want to discuss the utility of these protocols for gene discovery. Interest in pSJ proteins has increased over the years as more pSJ genes have been identified and additional roles for pSJ proteins have been identified17,22,27. In the past, many pSJ genes have been identified through screens of tracheal development since pSJ mutant embryos often show long and convoluted trachea28,29,30,31. A small systematic screen for pSJ components has been undertaken using the dye injection functional assay described here18. pSJ genes have even been identified through a mass spectrometric biochemical analysis32. Since most of those approaches are time and labor-intensive, we suggest that a simple RNA interference (RNAi)-based screen using the imaging protocol we presented might be highly productive and cost-effective. The mislocalization of core pSJ proteins in the polarized epithelia of stage 16 embryos is diagnostic for a defect in the formation or maintenance of the pSJ. We and others have shown that RNAi transgenes for known core or accessory pSJ proteins recapitulate the zygotic loss-of-function pSJ localization phenotypes shown here11,13,17,28. With the expansive collection of readily available RNAi stocks33,34, ubiquitous and tissue-specific GAL4 lines, and inexpensive pSJ-specific antibodies, it would be possible to conduct a large-scale screen of candidate genes for new pSJ components. We have tested both the ubiquitous GAL4 driver, daughterless, and the salivary-specific GAL4 driver, forkhead, and found comparable mislocalization of pSJ proteins in the salivary glands of pSJ-RNAi embryos to those with zygotic loss of function alleles of the same gene17. We have also found that both Bloomington TRiP RNAi lines and VDRC RNAi lines can be effective in these assays (not shown), but as with any RNAi screen, we recommend using multiple RNAi stocks for each gene tested. For any promising candidates, the dye-injection protocol can be employed to assess the function of the pSJ in candidate RNAi-expressing embryos. Finally, it would be useful to create a specific antibody or fluorescently tagged transgene to determine if the candidate gene encodes a core component of the junction.

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>

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 subcellularly localized to the pSJ once a functional junction forms in wild type embryos. In that case, the protein would be classified as a core pSJ protein. To demonstrate this principle, we first show how the localization of a core pSJ protein matures from stage 13 to stage 16 of embryogenesis in wild-type embryos (Figure 1A-D). Here, we focus on the salivary gland, which is an epithelial tube with the apical surface facing the lumen of the gland. During stages 13-16 of embryogenesis, the salivary gland is comprised of tall columnar cells, allowing for direct observation of the lateral membrane in confocal optical sections of the embryo. Using an antibody against the core pSJ protein Macroglobulin complement-related (Mcr; in A-C), we see that Mcr is uniformly localized all along the lateral membrane in stage 13 embryos. By stage 14, Mcr is more strongly enriched in an apical region of the lateral membrane, just below the adherens junction (visualized by the expression of the adherens junction protein E-cadherin), although substantial Mcr immunostaining can still be observed along the remainder of the lateral membrane. At stage 16, Mcr strongly localizes to an apical lateral region just basal to the adherens junction (~10-15% of the length of the lateral membrane), with little or no basal spreading. This region of the lateral membrane is the mature pSJ. Mcr is a transmembrane core pSJ protein10, whereas Coracle (Cora) is a cytoplasmic core pSJ protein9. Note that antibodies against both proteins work equally well to define the mature pSJ (Figure 1C,D). Embryonic staging is best determined by midgut morphology. The midgut can be visualized by autofluorescence of the yolk proteins using UV illumination as shown for a stage 16 embryo in Figure 1H. The stage 16 midgut appears as 3 distinct parallel chambers.
In embryos with mutations in core pSJ genes, the organization of the pSJ is disrupted, and all other core pSJ proteins are mislocalized. This lack of pSJ organization is most clearly demonstrated by observing the localization of core pSJ proteins in stage 16 mutant embryos. As shown in Figure 1E-G, the maturation of the pSJ as determined by Mcr localization fails to occur in cora4 mutant embryos. cora4 is a zygotic loss-of-function mutation resulting from an in-frame deletion in the conserved amino-terminal region of Cora19. Mcr is expressed normally in stage 13 cora4 mutant embryos but fails to achieve a strong apical localization by stage 16 (Figure 1G). The expression of Mcr protein along the entire length of the lateral membrane in the stage 16 salivary gland, even with some enrichment in the region just basal to the adherens junction, is diagnostic of a core pSJ mutation. This phenotype is highly penetrant in cora4 mutant embryos and would be observed in all or nearly all cora4 mutant embryos. Note that E-cadherin is similarly localized in wild-type and cora4 mutant stage 16 embryos, demonstrating that gross apical-basal polarity is normal in these mutant animals.
To test the functional integrity of the pSJ, we employed a dye permeability assay in which we injected a rhodamine-labeled 10 kDa dextran solution into the hemocoel of wild-type and pSJ mutant embryos (cora4). Previous work has demonstrated that the occluding junction is functionally tight by the end of stage 15 of embryogenesis12. Once the pSJ establishes a functional occluding junction, any labeled dextran injected into the hemocoel is trapped on the basal side of the trachea, salivary glands, and hindgut and cannot enter the lumen through a paracellular route. This can be easily observed by fluorescence or confocal microscopy. Here, we used a confocal microscope to take a z-stack of fluorescent images from the dorsal surface. In wildtype embryos, we observe labeled dextran spreading throughout the body cavity, but no accumulation in the lumen of the dorsal trunk of the trachea (Figure 3, top panels). Instead, the dye pools along the basal side of tracheal epithelium, showing a clear outline of the tube (red arrows in the top panel of Figure 3). In contrast, injection of rhodamine-labeled 10 kDa dextran solution into the hemocoel of cora4 mutant stage 17 embryo results in strong accumulation of the dye in the lumen of the trachea within 10 minutes (red arrows in the bottom panels of Figure 3), indicating that the occluding junction has failed to form. The rapid accumulation of dye into the lumen of the trachea (or salivary gland or hindgut) in stage 16 or 17 embryos is diagnostic for a pSJ functional defect. The penetrance of this phenotype is very high in a core pSJ mutant and so all or nearly all of the mutant embryos would be expected to show this defect.
<img alt="Confocal microscopy images of Drosophila embryonic stages showing Mcr and Ecad expression levels." class="xfigimg" src="/files/ftp_upload/69986/69986fig1.jpg" data-altupdatedat="1782792046000" data-alt="Figure 1"> 
Figure 1: Maturation of pSJ organization in wild-type embryos and disrupted pSJ organization in cora4 mutant embryos. (A-C) Confocal optical sections of salivary glands from staged w1118 control embryos stained with antibodies against Mcr (in cyan and in panels A'-C') and Ecad (in magenta and in panels A"-C". In each panel, anterior is to the left and the dorsal surface of the embryo is to the top of the image. The salivary gland is an epithelial tube with the apical surface facing the lumen of the gland. Note that Mcr is localized all along the lateral membrane in stage 13 embryos (red arrow) and becomes more strongly enriched along the apical region of the lateral membrane in stage 14 embryo, and is completely restricted to the region of the pSJ by stage 16. The pSJ region lies just basal to the adherens junction, as visualized with Ecad (A"-C"). (D) Confocal optical section of a salivary gland from a stage 16 w1118 embryos stained with antibodies against Cora (in cyan and in panel D') and Ecad (in magenta and in panel D") to demonstrate that Cora antibody staining is an equally good marker for pSJ organization. (E-G) Confocal optical sections of salivary glands from staged cora4 mutant embryos stained with antibodies against Mcr (in cyan and in panels E'-G') and Ecad (in magenta and in panels E"-G"). Note the persistent localization of Mcr along lateral membranes in stage 15 and 16 mutant embryos (red arrows in F' and G'). (H) Bright field and fluorescence image of a stage 16 wild-type embryo to show the midgut morphology used in embryo staging. The embryo shown was imaged with brightfield microscopy and then with fluorescence microscopy using UV illumination to highlight the autofluorescence of the yolk proteins in the midgut. In stage 16 embryos, the midgut appears as three chambers. Scale bars in panels A and E are 10 µm. <a href="https://www.jove.com/files/ftp_upload/69986/69986fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="A: Microfabrication setup with microneedles; B: Optical microscope image with enhanced contrast; C: Temperature-controlled slide preparation; D-E: Microscopic analysis of cell structures; F: Advanced microscope for high-resolution imaging; G: Microfluidic system for fluid analysis and experimentation." class="xfigimg" src="/files/ftp_upload/69986/69986fig2.jpg" data-altupdatedat="1782792046000" data-alt="Figure 2"> 
Figure 2: Experimental details for dye penetration assay. (A) Examples of acceptable capillary needles for dye injection assays. Both needles have a short gradual taper that transitions to a sharp taper with a long point. The needle on the left is on the longer side of acceptable and may bend as it enters the embryo. If it does so, the needle can be re-broken to make a shorter point. The needle on the right is on the shorter side but has a small enough bore to work effectively. (B) Dechorionated embryos lined up on a piece of double-sided tape on a 22 x 22 #2 coverslip. After stage 16 and 17 embryos are lined up on an apple juice plate with posterior ends aligned and dorsal sides facing down, they are picked up on a coverslip covered by a piece of double-sided tape. (C) Conditions for desiccation are seen just after the coverslip has been affixed to a microscope slide using halocarbon oil. The embryos are then aged in air for 15 min in a room with 42% relative humidity and 21.2 °C temperature. (D,E) Photomicrograph of embryos (D) before and after (E) 15 min of desiccation. Note that the desiccated embryos are slightly less turgid but not substantially flaccid. (F,G) Our microinjection setup on an inverted microscope. (G) A higher magnification image to show the angle of the needle compared to the embryos. This is the shallowest angle we can achieve with this micromanipulator/microscope configuration, but a shallower angle (closer to parallel with the embryos) would be better. Note that the embryos are covered with halocarbon oil, and the slide is arranged with the embryos facing away from the objective. <a href="https://www.jove.com/files/ftp_upload/69986/69986fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Drosophila embryo development, fluorescence microscopy, w1118 vs cora4, dorsal-ventral inspection." class="xfigimg" src="/files/ftp_upload/69986/69986fig3.jpg" data-altupdatedat="1782792046000" data-alt="Figure 3"> 
Figure 3: Examples of dye permeability assays in wild-type and pSJ mutant embryos to interrogate pSJ occluding function. Confocal optical sections from a z-stack of a w1118 control stage 17 embryo (top panels) and a cora4 mutant stage 17 embryo (bottom panels) that had been injected with rhodamine-labeled 10 kDa dextran within the previous 10 min. Embryos are displayed with the anterior side to the top. Sections are 2.4 µm apart and shown sequentially from the dorsal surface (left side of the figure) towards the interior of the embryo (right side). Red arrows indicate the location of the trachea. Note that in the w1118 embryo, the dye surrounds the trachea but does not penetrate into the lumen. In contrast, in the cora4 mutant embryo, the interior of the trachea is filled with dye. Scale bar on top panel is 100 µm. <a href="https://www.jove.com/files/ftp_upload/69986/69986fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

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

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 ...

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 and dye penetration experiments to define core septate junction proteins in Drosophila embryonic epithelia

Allison Hall

Bethany Lucas

The establishment of polarity is critical for the proper development of all multicellular organisms. The study of how cells polarize on individual and tissue levels is of broad interest and has been studied across many model organisms. This Methods Collection aims to encapsulate methods used to study polarity across organisms and fields.&nbsp;

JoVE Developmental Biology publishes research methodologies in the field of developmental biology. This includes methods that are used to study biological development in vitro and in vivo at the molecular, cellular, tissue, organ and whole organism levels.

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Methods for Studying Polarity

The establishment of polarity is critical for the proper development of all multicellular organisms. Curated by Allison Hall, Bethany Lucas. Explore Developmental Biology techniques on JoVE.

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