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

Summary

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

Abstract

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.

Introduction

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 epithelium^1,2. 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 melanogaster^1,3. 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 junction^3,5. 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 alpha^8,9,10,11,12. 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 Coiled^13,14. 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 III^15,16.

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 localization^18,19. 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 polarity^17,20,21,22. 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 microscopy^6,23. 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.

Protocol

1. Staging of embryos

1. 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.
2. 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 °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 °C. Alternatively, collect embryos overnight at 25 °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's long axis (Figure 1H).

2. Fixation and immunostaining of embryos

1. Assemble an embryo collection basket with a nylon mesh (120 mm mesh). Cover the opening and the basket's threads with the nylon mesh, then screw on the cap.
2. 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.
3. 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.
4. 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 °C) 1x phosphate-buffered saline (PBS) in a chemical fume hood. To speed the dissolving process, add 10 µL of 14 N Sodium Hydroxide (per 10 mL of PBS) to the fix until the formaldehyde goes into solution, then add 10 µL 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.
5. 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.
6. 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.
7. 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.
8. 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.
9. 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 °C or for 3-4 h at room temperature with gentle mixing on a platform rocker.
10. 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.
11. 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 °C.
12. 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.
13. 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.

3. Imaging and data analysis of immunostained embryos

1. 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).
2. 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.

4. Preparation for dye injection

1. 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.
2. 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.
3. 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.
4. 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.
5. 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.
6. 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.

5. Mounting embryos and dye injection

1. Select stage 16 or 17 embryos (control and mutant) using midgut morphology as a staging marker.
2. 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.
3. 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).
4. 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 °C.
5. 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).
6. 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 "inflate" 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.
7. 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.

6. Imaging and data analysis

1. 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).
2. 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.

Results

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...

Discussion

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's occluding function by injecting dye-labeled 10 kDa dextran molecules into stage 16 living embryos. The simplicity of these protocols a...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Capillary tube for needles (Boro 1 X 0.5/Fiber )	FHC	30-30-1
Confocal microscope	Leica	Stellaris 5
Cover glass 22 x 22 2 thickness for injections	Fisher Scientific	12-540-B
Cover glass 22 x 30 1.5 thickness for imaging	Corning	2980-233
Culture tube for immunostaining (6 x 50 mm)	Fisher Scientific	14-958-A
Embryo collection cage	Genesee Scientific	59-100
Fluorescence stereomicroscope	Leica	MZ10 F
Halocarbon oil (700)	Genesee Scientific	59-131
Heptane	Fisher Scientific	H350-4
Inverted fluorescence microscope	Leica	DMi8
Mesh baskets for dechorionating embryos	Genesee Scientific	46-101
Methanol	Fisher Scientific	A412-4
Micromanipulator	Narishige	MN151
Microscope slides	Fisher Scientific	12-544-2
Needle puller	David Kopf instruments	model 720
Normal Donkey Serum	Jackson ImmunoReseaerch	017-000-121
Nylon mesh (120 um)	Genesee Scientific	57-102
Objective lens HC PL FLUOTR 10X/0.32 PH 1	Leica	11506537
Objective lens HC PL APO 10x/0.40 CS2	Leica	15506407
Objective lens HC PL APO 40x/1.30 CS2	Leica	15506358
Petri plates 60 x 15 mm	Corning	351007
Platform rocker	Globe Scientific	GTR-FS
Primary antibody (Coracle)	DSHB	C566.9
Primary antibody (Coracle)	DSHB	C615.16
Primary antibody (E cadherin)	DSHB	DCAD2
Primary antibody (Mcr)	Ward lab, CWRU	guinea pig polyclonal
Rhodamine labeled 10 kDa dextran	Molecular Probes	D1824
Secondary antibody (donkey anti-guinea pig Cy3)	Jackson ImmunoReseaerch	706-165-148
Secondary antibody (donkey anti-mouse Cy2)	Jackson ImmunoReseaerch	715-225-151
Secondary antibody (donkey anti-rat Cy5)	Jackson ImmunoReseaerch	712-175-150
Table top shaker	New Brunswick Scientific	C2 platform shaker
Tegosept (methyl 4-hydroxybenzoate)	TCI	H0216
Triton X-100	Fisher Scientific	BP151500
Vials for fixation (20 ml WHEATON liquid scintillation vials)	Fisher Scientific	03-341-73C
Whatman 3MM  Chromatography Paper	Fisher Scientific	05-716-6H
Fly stocks	Full genotype	stock number	Source
cor4	w*; P{neoFRT}43D cora4/CyO	52232	BDSC
w1118	w[1118]	5905	BDSC
Solution	Recipe
10x PBS	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.
10% Triton X-100 solution	Mix 1 ml of Triton X-100 to 9 ml of dH2O on a platform rocker until dissolved. Store at room temperature.
50% bleach	Mix 10 ml of commercial bleach (7.5% sodium hypochlorite) to 10 ml of water prior to dechorionating embryos.
Apple juice plates	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.
Blocking solution	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.
Embryo wash solution	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.
Fix	Preheat 10 ml of 1X PBS in the microwave. Add 0.4 g  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.
Mounting media	Mix 9 ml of glycerol and 1 ml of 1M Tris base pH 8.0. Add 0.05 g n-propyl-gallate,  mix and heat to dissolve. Aliquot 0.5 ml into eppendorf tubes.
Rhodamine dextran solution	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.
Yeast paste	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.