Name: confocal and super-resolution imaging of polarized intracellular trafficking and secretion of basement membrane proteins during drosophila oogenesis
Uploaded: 2022-05-19T14:00:00
Description: Watch this Scientific Journal Video about Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis at JoVE.c

The authors are grateful to Julie Merkle for her helpful comments on the manuscript. This work was supported by NIH grant R15GM137236 to O.D. The confocal and super-resolution images were acquired using a Zeiss LSM 900 with Airyscan 2, purchased with NSF MRI grant 2018748.

1. Fly preparation for ovary dissections
<ol>
	<li>Put 10-15 Drosophila melanogaster&#160;female flies (1-2 days old) of the desired genotype in a narrow vial containing ~8 mL of Drosophila fly medium sprinkled with a small amount of granulated baker&#39;s yeast 2-3 days prior to dissection at 25 &#176;C. Adding a few males to the vial can boost egg chamber yield. However, ensure that the total number of flies does not exceed 20 as this can negatively affect ovary development. 
	...

The BM is critical for embryonic and organ morphogenesis, and adult physiological functions. Moreover, the BM acts as a signaling platform for the establishment and maintenance of epithelial polarity and provides tissues with support2. Yet, the mechanisms that regulate the proper placement of BM proteins are poorly understood. A better understanding of the biological pathways dedicated to the intracellular trafficking and polarized secretion of BM proteins requires a careful analysis of the compon...

The basement membrane (BM) is a thin sheet of layered cell-adherent extracellular matrix (ECM) critical for epithelial structure and morphogenesis1. It comprises ~50 proteins and is found ubiquitously underlying the epithelial and endothelial cells, and ensheathing skeletal, smooth, and heart muscle cells and adipocytes1,2,3. The three main components of the BM at the basal side of the epithelial cells are Collagen IV, Perlecan, and Laminins. The BM underlies the epithelial cells and is responsible for many functions, including tissue separation and barrier, growth and support, and cell polarization2,3,4,5,6,7,8,9,10,11,12. Its role as a signaling platform regulates the morphology and differentiation of epithelial cells and tissues during development3,13,14. Moreover, the mis-regulation of the BM and/or a breach in its integrity are the primary causes of many pathological conditions, including tumor metastasis2,15,16. Despite the essential functions performed by the BM during tissue and organ morphogenesis, the components of the biological pathway(s) dedicated to the polarized intracellular trafficking and secretion of BM proteins are vaguely known.
To study the intracellular trafficking of BM-containing vesicles and the secretion of BM proteins by epithelial cells, the follicular epithelium (FE) of the Drosophila ovary is a powerful model system (Figure 1). A Drosophila ovary comprises 16-20 long, tube-like structures, called ovarioles (Figure 1A,B)17,18,19. Each ovariole can be thought of as an egg assembly line, with the age progression of egg chambers (which each gives rise to an egg) that begins at the anterior end and moves posteriorly, until the mature egg exits through the oviduct. Each egg chamber is encapsulated by the FE, a monolayer of somatic follicle cells (FCs), that surrounds the central germline cells (GCs). The FE is highly polarized with a distinct apical-basal polarity where the apical domain faces the germline, and the BM proteins are secreted basally18,19. The FCs actively secrete all of the major components of the BM, including Collagen IV, Perlecan, and Laminins20,21. In epithelial cells such as FCs, the BM components are produced and require a specialized polarized secretion pathway for their deposition extracellularly. For example, in the case of the most abundant component of the BM, Collagen IV (Coll IV), the details surrounding its polarized intracellular trafficking and secretion are vague despite its production and deposition being the focus of many studies. Coll IV is translated in the endoplasmic reticulum (ER), which is also where each fibril - composed of three polypeptides (two &#945;1 chains and one &#945;2 chain) - is assembled into a triple helix22. Proper Coll IV folding and function require ER chaperones and enzymes, including lysyl and prolyl-hydroxylases such as Plod and PH4&#945;EFB20,22,23,24,25,26. These posttranslational enzymes regulate the ER sorting of Coll IV, as the loss of each causes Coll IV to be trapped in the basal ER20,23,24,25,26. Then, newly synthesized Coll IV exits the ER for the Golgi in COPII-coated vesicles. The cargo receptor Tango1 aids in packaging collagens into sizable Golgi-bound vesicles that can accommodate large multimeric proteins20,27. Once Coll IV is packaged into intracellular exocytic vesicles, it is specifically secreted basally from epithelial cells. To direct BM deposition to the basal side, epithelial cells require another set of factors specifically dedicated to polarized BM secretion. Using the FE of the Drosophila ovary, a few components of this novel cellular process have been characterized, including the nucleotide exchange factors (GEFs) Crag and Stratum, the GTPases Rab8 and Rab10, as well as the levels of the phosphoinositide PI(4,5)P2, and Kinesin 1 and 3 motor proteins20,28,29,30,31. These components are critical in ensuring the polarized distribution of BM proteins.
To monitor the intracellular localization of BM proteins in the FE, endogenously tagged basement membrane proteins (protein traps), such as Viking-GFP (Vkg-GFP or &#945;2 Coll IV-GFP) and Perlecan-GFP (Pcan-GFP) can be used32,33. These protein trap lines have been shown to accurately reflect the endogenous distribution of BM proteins and allow for more sensitive detection of vesicular trafficking28,30. The components involved in the polarized deposition of BM in the FE were first characterized using protein trap lines for Vkg-GFP and Pcan-GFP20,28,29,30. Protein traps can be used in different genetic backgrounds, including mutants and Gal4 lines34. Moreover, protein traps can be used in combination with fluorescent dyes and/or fluorescence immunostaining, allowing for precise characterization of the localization of BM proteins when comparing wild-type and mutant conditions35.
To accurately and efficiently assess the distribution and localization of BM protein-containing vesicles, confocal laser scanning microscopy (CLSM) and super-resolution imaging techniques present a significant advantage to other imaging approaches. These approaches couple high-resolution imaging with relative ease of use. CLSM is a microscopy technique that allows for an improved optical resolution by scanning the specimen with a laser in a raster scan manner using galvanometers. The pinhole aperture is a core component of a confocal microscope. By blocking the out-of-focus signals coming from above or below the focal plane, the pinhole aperture results in a highly superior resolution in the z-axis36. This also makes it possible to obtain a series of images in the z-plane, called a z-stack, corresponding to a series of optical sections. z-stacks subsequently create a 3D image of the specimen, via 3D reconstruction, with the aid of imaging software. Conventional epifluorescence (widefield) microscopes, unlike confocal microscopes, allow out-of-focus light to contribute to image quality, decreasing image resolution and contrast36,37. This makes epifluorescence microscopy a less attractive candidate when studying protein localization or colocalization.
Although CLSM is a suitable approach for various applications, including imaging and characterization of the intracellular trafficking of BM proteins, it still presents an issue when imaging samples below Abbe&#39;s diffraction limit of light (200-250 nm). When imaging such samples, confocal microscopy, especially when using an oil objective, can result in high resolution. However, super-resolution techniques surpass the limit of confocal microscopy. There are various approaches to achieve super-resolution microscopy, each with specific resolution limits, and each appropriate for different analyses. These approaches include photoactivated localization microscopy (PALM) or stochastic optical reconstruction microscopy (STORM), stimulated emission depletion microscopy (STED), structured illumination microscopy (SIM), and Airyscan (super-resolution) microscopy38,39,40,41,42,43,44,45,46. Although Airyscan has a coarser resolution than PALM/STORM, STED, and SIM, it can still achieve a resolution of up to ~120 nm (about twice the resolution of CLSM). Furthermore, this super-resolution microscopy approach has been shown to have an advantage over SIM and other super-resolution techniques when imaging thick samples and samples with a low signal-to-noise ratio47,48.
Airyscan is a relatively new super-resolution confocal microscopy technology46. Unlike traditional CLSMs, which use the pinhole and single point detectors to reject out-of-focus light, this super-resolution approach uses a 32-channel gallium arsenide phosphide (GaAsP) photomultiplier tube area detector that collects all of the light at every scan position45. Each of the 32 detectors work as a small pinhole, reducing the pinhole size from the traditional 1.0 Airy Unit (A.U.) to an enhanced 0.2 A.U., enabling an even higher resolution and signal-to-noise ratio, while maintaining the efficiency of a 1.25 A.U. diameter45. Furthermore, the linear deconvolution used by Airyscan results in up to a 2x increase in resolution45. Taking this into consideration, CLSM, and specifically super-resolution microscopy, are&#160;well-suited to study BM proteins and proteins that regulate the basal deposition of BM proteins, as they can produce very high-resolution images for localization and colocalization studies, thereby providing new insights in the spatial, temporal, and molecular events that control these processes.
An alternative approach to confocal microscopy that can be used to perform localization experiments is image deconvolution. Since widefield microscopy allows out-of-focus light to reach the detectors, mathematical and computational deconvolution algorithms can be applied to remove or reassign out-of-focus light from images obtained by widefield microscopy, thereby improving the resolution and contrast of the image49. Deconvolution algorithms can also be applied to confocal images to further increase resolution and contrast, producing final images almost comparable to that of super-resolution microscopy50. Airyscan makes use of Weiner filter-based deconvolution along with Sheppard&#39;s pixel reassignment, resulting in a highly improved spatial resolution and signal-to-noise ratio. Compared to confocal microscopy, an increase of 2x in resolution in all three spatial dimensions (120 nm in x and y, and 350 nm in z) is observed when using this super-resolution microscopy technique45,51.
This manuscript provides detailed and optimized protocols to stain, acquire, and visualize the intracellular trafficking and deposition of BM proteins using the FE of the Drosophila ovary as a model system coupled with confocal and super-resolution microscopy. Drosophila lines expressing endogenously tagged basement membrane proteins, e.g., Vkg-GFP and Pcan-GFP, are efficient and accurate tools to visualize BM protein trafficking and secretion. In addition, they can be easily used in different genetic backgrounds, including mutant and Gal4/UAS lines34. Although endogenously tagged basement membrane proteins are recommended, the use of antibodies against specific BM proteins is also compatible with the described protocols. These protocols are particularly useful for scientists who are interested in studying intracellular trafficking and the secretion of BM proteins in intact epithelial tissue using confocal and super-resolution imaging. Moreover, the ability to combine epithelial tissue analysis with the expansive tools of Drosophila genetics makes this approach especially powerful. Finally, these protocols could be easily adapted to study vesicular trafficking and sorting of other proteins of interest.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Alexa Fluor 546 phalloidin</td>
			<td>Invitrogen</td>
			<td>A22283</td>
			<td>F-Actin Stain (1/500 of 66&#181;M)</td>
		</tr>
		<tr>
			<td>Alexa Fluor 647 phalloidin</td>
			<td>Invitrogen</td>
			<td>A22287</td>
			<td>F-Actin Stain (1/100 of 66&#181;M))</td>
		</tr>
		<tr>
			<td>Anti-GM130 Antibody</td>
			<td>abcam</td>
			<td>ab30637</td>
			<td>For Golgi Stain (colocalization); use as concentration of 7&#181;g/uL</td>
		</tr>
		<tr>
			<td>Aqua-Polymount</td>
			<td>Polysciences, Inc.</td>
			<td>1860620</td>
			<td>Mounting Medium</td>
		</tr>
		<tr>
			<td>Bakers Yeast (Active Dry Yeast)</td>
			<td>Genesee Scientific</td>
			<td>62-103</td>
			<td>To fatten the overies for dissection</td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (30% solution)</td>
			<td>Sigma-Aldrich</td>
			<td>A7284</td>
			<td>For blocking solution</td>
		</tr>
		<tr>
			<td>Depression wells</td>
			<td>Electron Microscopy Sciences</td>
			<td>7156101</td>
			<td>For dissection (glass concavity slide can be used instead)</td>
		</tr>
		<tr>
			<td>Dissecting needle</td>
			<td>Fisher scientifc</td>
			<td>13-820-024</td>
			<td></td>
		</tr>
		<tr>
			<td>Drosophila Incubator</td>
			<td>Genesee Scientific/Invictus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fly Stock: Perlecan-GFP Drosophila line (ZCL1700)</td>
			<td></td>
			<td></td>
			<td>Morin et al., 2001</td>
		</tr>
		<tr>
			<td>Fly Stock: UAS-Crag RNAi line (TRIP line HMS00241)</td>
			<td>Bloomington Drosophila Stock Center</td>
			<td>33594</td>
			<td>RNAi against Crag</td>
		</tr>
		<tr>
			<td>Fly Stock: Viking-GFP Drosophila line (CC00791)</td>
			<td></td>
			<td></td>
			<td>Buszczak et al., 2007</td>
		</tr>
		<tr>
			<td>Fly Stock: Vkg-GFP, tj-Gal4</td>
			<td></td>
			<td></td>
			<td>Devergne et al., 2017. Drive the expression of Crag RNAi in the FE</td>
		</tr>
		<tr>
			<td>Forceps (Dumont 5)</td>
			<td>Fine Science Tools</td>
			<td>11251-30</td>
			<td>For dissection</td>
		</tr>
		<tr>
			<td>Glass Concavity Slide</td>
			<td>Electron Microscopy Sciences</td>
			<td>7187804</td>
			<td>For dissection (depression wells can be used instead)</td>
		</tr>
		<tr>
			<td>Goat anti-Rabbit IgG, Alexa Fluor 568</td>
			<td>Invitrogen</td>
			<td>A11036</td>
			<td>Secondary antibody (GM130 antibody) (5 &#181;g/mL)</td>
		</tr>
		<tr>
			<td>Hoechst (Hoechst 33342)</td>
			<td>Invitrogen</td>
			<td>H3570</td>
			<td>DNA Stain (1 ug/mL)</td>
		</tr>
		<tr>
			<td>Kimwipes</td>
			<td>Kimtech</td>
			<td>Fisher Scientific: 06-666</td>
			<td>Delicate task wipers</td>
		</tr>
		<tr>
			<td>Leica Fluorescent Stereo Microscope&#160; M165 FC</td>
			<td>Leica</td>
			<td></td>
			<td>For ovary imaging</td>
		</tr>
		<tr>
			<td>Microscope Slides</td>
			<td>Corning</td>
			<td>294875X25</td>
			<td>Microscope Slides</td>
		</tr>
		<tr>
			<td>Nutating platform rocker</td>
			<td>Corning Life Sciences</td>
			<td>6720</td>
			<td>For ovary fixation and staining</td>
		</tr>
		<tr>
			<td>Nutri-Fly BF</td>
			<td>Genesee Scientific</td>
			<td>66-121</td>
			<td>Fly Food</td>
		</tr>
		<tr>
			<td>Paraformaldehyde 20% Solution</td>
			<td>Electron Microscopy Sciences</td>
			<td>Fisher Scientific: 15713</td>
			<td>For PFA 4%</td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline Tablets</td>
			<td>Fisher scientific</td>
			<td>BP2944100</td>
			<td>For PBS solution</td>
		</tr>
		<tr>
			<td>ProLong Glass Antifade Mountant</td>
			<td>Invitrogen</td>
			<td>P36980</td>
			<td>Mounting Medium</td>
		</tr>
		<tr>
			<td>Square Cover Glass</td>
			<td>Corning</td>
			<td>285022</td>
			<td>Cover glass for microscope slides</td>
		</tr>
		<tr>
			<td>Triton x-100</td>
			<td>Sigma-Aldrich</td>
			<td>9036-19-5</td>
			<td>For PBST</td>
		</tr>
		<tr>
			<td>Zeiss LSM 900 with Airyscan 2</td>
			<td>Zeiss</td>
			<td></td>
			<td>Confocal and super-resolution Microscope</td>
		</tr>
		<tr>
			<td>Zeiss Stemi 305 Stereo Microscope</td>
			<td>Zeiss</td>
			<td></td>
			<td>Dissecting microscope</td>
		</tr>
		<tr>
			<td>Zeiss Zen Software version 3.3 (Blue Edition)</td>
			<td>Zeiss</td>
			<td></td>
			<td>Image acquisition and processing</td>
		</tr>
	</tbody>
</table>

confocal and super-resolution imaging of polarized intracellular trafficking and secretion of basement membrane proteins during drosophila oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment and maintenance of epithelial tissue morphology and organ morphogenesis. Moreover, the BM is essential for tissue modeling, serving as a signaling platform, and providing external forces to shape tissues and organs. Despite the many important roles that the BM plays during normal development and pathological conditions, the biological pathways controlling the intracellular trafficking of BM-containing vesicles and how basal secretion leads to the polarized deposition of BM proteins are poorly understood. The follicular epithelium of the Drosophila ovary is an excellent model system to study the basal deposition of BM membrane proteins, as it produces and secretes all major components of the BM. Confocal and super-resolution imaging combined with image processing in fixed tissues allows for the identification and characterization of cellular factors specifically involved in the intracellular trafficking and deposition of BM proteins. This article presents a detailed protocol for staining and imaging BM-containing vesicles and deposited BM using endogenously tagged proteins in the follicular epithelium of the Drosophila ovary. This protocol can be applied to address both qualitative and quantitative questions and it was developed to accommodate high-throughput screening, allowing for the rapid and efficient identification of factors involved in the polarized intracellular trafficking and secretion of vesicles during epithelial tissue development.

The methods described herein can be used to efficiently and accurately image and characterize the intracellular trafficking and secretion of BM proteins in polarized epithelial cells, such as the FE of the Drosophila ovary. Next, we provide anticipated results obtained using the described methods, as well as helpful advice and potential pitfalls. To do so, Vkg-GFP, an endogenously tagged Vkg (Drosophila Col IV) is used. However, the same results can be achieved with other endogenously tagged BM proteins...

Watch this Scientific Journal Video about Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis at JoVE.c

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane is essential for tissue and organ morphogenesis during development. To better understand the mechanisms leading to proper placement of this structure, the protocol presented describes methods to visualize and characterize the intracellular trafficking and secretion of basement membrane proteins in epithelial cells using confocal and super-resolution microscopy.

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment ...

This protocol allows the characterization of the intracellular trafficking and secretion of basement membrane proteins using the Drosophila array as a model system. The main advantage of our technique is that it allows high-resolution imaging of the intracellular trafficking of basement membrane proteins using endogenously-tagged proteins and Airyscan super-resolution microscopy. Although this protocol is developed to image the intracellular trafficking of basement membrane proteins, it can be extended to study the trafficking of other proteins of interest and other biological systems, including cell culture and organoids.To begin, after dissecting Drosophila ovaries in PBS under a dissecting scope, add one milliliter of fixation solution and fix the ovaries on a nutating platform for 15 minutes. Remove the fixation solution and perform two quick washes, each with one milliliter of PBST. By gently inverting the microcentrifuge tubes five to six times, then perform four long washes of 10 minutes each with one milliliter of PBST on a nutating platform rocker.After performing fixation and washing, remove PBST, then add one milliliter of blocking solution and block the ovaries on a nutating platform rocker for a minimum of one hour. Next, remove the blocking solution and add 300 microliters of primary antibody solution, containing primary antibodies diluted at their appropriate concentrations in blocking solution. Incubate overnight on a nutating platform rocker at 4 degrees Celsius.After removing the primary antibody solution, perform two quick washes and four long washes, as demonstrated previously, then remove PBST and add 500 microliters of secondary antibody solution containing fluorescent secondary antibodies that will detect the primary antibodies used. Incubate the ovaries in the secondary antibody solution on a nutating platform rocker for two hours at room temperature, then perform two quick washes and four long washes in PBST as demonstrated previously on a nutating platform rocker. After the last wash, use a P-1000 pipette to gently pipette the ovaries up and down in the tube to separate the egg chambers.Allow the egg chambers to sink to the bottom by keeping the tube in an upright position for five to 10 minutes. Next, remove PBST using a Pasteur pipette, leaving approximately 50 microliters. Afterward, remove as much of the remaining PBST as possible using a P-200 pipette and add two drops of mounting medium, enough to spread evenly on a cover slip.To allow easy transfer of the viscous mounting medium to the slide from the microcentrifuge tube, cut off the end of a P-200 pipette tip. Next, slowly transfer all the egg chambers in the mounting medium to the glass slide, ensuring not to create bubbles. Under a dissecting microscope, gently spread out the mounting media and separated egg chambers using a new P-200 pipette tip or forceps to cover an area approximately the size of a cover slip.Using forceps, carefully place the cover slip on the egg chambers at an angle to avoid bubbles and store the slide at room temperature on a flat surface in the dark for two days to polymerize. Once the mounting media has cured, the slide can be stored at 4 degrees Celsius in the dark for a few weeks for imaging. To visualize the intracellular localization of basement membrane proteins, set the objective to 63x and gently place a drop of immersion oil on its lens, then position the slide on the objective with the cover slip facing the objective to locate the specimen.Locate the region of interest using the eyepiece of the epifluorescent microscope, then select a configuration with appropriate settings for the fluorophores to image as described in the manuscript. Once the configuration is set, image the sample by selecting Live under Acquisition Mode and adjust the zoom between 2 and 4x to optimize the scan area of the sample. For each individual channel, select a track under Channel and click on Live.Adjust the master gain and laser power while using the range indicator tool and follow all the guidelines to avoid saturated pixels as described in the manuscript, while repeating the same for each channel. In the Acquisition Mode toggle window, under Image Size, click on SR to maximize the capabilities of the detector and adjust the frame size automatically. Keep the averaging to None in Airyscan mode as it is usually not necessary and will decrease the scan time.However, in some cases, an averaging of 2x may improve the signal-to-noise ratio, then click on Snap to acquire an image. To acquire a Z-stack, click on Z-Stack checkbox under the Acquisition tab. Next, select the desired channel to observe the specimen.For example, the DAPI channel and click on Live to start a live scan, then set a range for the Z-stack using the fine adjustment knob on the microscope. Afterward, set the endpoints of the Z-stack by clicking Set First and Set Last. For optimal 3D reconstruction, set the interval for the Z-stack lower than 0.5 micrometers to assign the step size and click on Start Experiment to begin Z-stack acquisition.Once the images or Z-stack is obtained, click on the Processing, then Method option and select Airyscan Processing. Perform Auto Filter to start and if required, perform further manual processing by changing the super resolution value to obtain the best results for the sample. After the optimal SR value has been determined, click on Apply to generate a processed image.In the case of Z-stack images, process either as one Z-slice or as the whole Z-stack by clicking on the 3D Processing box. To visualize protein trafficking in 3D after the acquisition of a Z-stack, generate a 3D image by clicking on the 3D icon in the preview window, that will appear in the display control section of the Airyscan processed image. From the different 3D view options, use Surface or Mixed views when viewing the structure of vesicles.For the highest quality image, select the Precise setting as the Fastest setting will be less accurate and lead to poor 3D rendering. Once the image has been generated, manipulate the 3D image by zooming and rotating to focus on a preferred location. After the view has been obtained, under the 3D tab, select Displayed Resolution, and then click on Create Image, which will create a snapshot of the image in the same orientation as it was viewed and can be saved and exported in various file formats.Confocal microscopy can be used to visualize the intracellular localization and deposition of basement membrane proteins such as Viking-GFP when the acquisition parameters are optimized. When acquisition and image processing are performed properly, super-resolution microscopy increases the image resolution compared to confocal microscopy. As illustrated by the better defined images in the intracellular trafficking of basement membrane proteins, such as Viking.Orthogonal projection allows the visualization of the overall distribution of vesicles and compartments containing basement membrane proteins in a single image using confocal or super-resolution imaging. A 3D reconstruction by assembling a stack of optical Z-sections taken by confocal or super-resolution imaging was employed to assess the localization and distribution of intracellular basement membrane proteins. Super-resolution imaging can be used to determine the precise localization of basement membrane proteins in mutant conditions.For example, in Crag knockdown epithelial cells, basement membrane proteins accumulate both apically and basally, indicating that it controls the polarized secretion of basement membrane proteins. Confocal and super-resolution microscopy can also be employed in co-localization experiments. For example, this figure shows the partial co-localization of Viking-GFP and a Golgi marker, GM-130, confirming that Viking is sorted to the Golgi before secretion.To efficiently image the trafficking of basement membrane proteins using super-resolution microscopy, we recommend to carefully mount the ovaries and take particular attention when setting the acquisition and the convolution parameters. This protocol is optimized to visualize the intracellular trafficking and secretion of basement membrane proteins. However, it can be easily modified to efficiently image the trafficking of other proteins of interest.

confocal-super-resolution-imaging-polarized-intracellular-trafficking

Research

JoVE Journal

Developmental Biology

Live Confocal Imaging of Developing Arabidopsis Flowers

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. Recent advances in confocal microscopy, combined with the development of numerous fluorophores, have overcome these issues and opened the possibility to study the expression of several genes simultaneously, with a good cellular resolution, in live samples. Live confocal imaging provides plant biologists with a powerful tool to study development, and has been extensively used to study root growth and the formation of lateral organs on the flanks of the shoot apical meristem. However, it has not been widely applied to the study of flower development, in part due to challenges that are specific to imaging flowers, such as the sepals that grow over the flower meristem, and filter out the fluorescence from underlying tissues. Here, we present a detailed protocol to perform live confocal imaging on live, developing Arabidopsis flower buds, using either an upright or an inverted microscope.

Live confocal imaging provides biologists with a powerful tool to study development. Here, we present a detailed protocol for the live confocal imaging of developing Arabidopsis flowers.

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. ...

The C. elegans Excretory Canal as a Model for Intracellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis in a Single Cell: labeling by GFP-fusions, RNAi Interaction Screen and Imaging

The four C. elegans excretory canals are narrow tubes extended through the length of the animal from a single cell, with almost equally far extended intracellular endotubes that build and stabilize the lumen with a membrane and submembraneous cytoskeleton of apical character. The excretory cell expands its length approximately 2,000 times to generate these canals, making this model unique for the in vivo assessment of de novo polarized membrane biogenesis, intracellular lumen morphogenesis and unicellular tubulogenesis. The protocol presented here shows how to combine standard labeling, gain- and loss-of-function genetic or RNA interference (RNAi)-, and microscopic approaches to use this model to visually dissect and functionally analyze these processes on a molecular level. As an example of a labeling approach, the protocol outlines the generation of transgenic animals with fluorescent fusion proteins for live analysis of tubulogenesis. As an example of a genetic approach, it highlights key points of a visual RNAi-based interaction screen designed to modify a gain-of-function cystic canal phenotype. The specific methods described are how to: label and visualize the canals by expressing fluorescent proteins; construct a targeted RNAi library and strategize RNAi screening for the molecular analysis of canal morphogenesis; visually assess modifications of canal phenotypes; score them by dissecting fluorescence microscopy; characterize subcellular canal components at higher resolution by confocal microscopy; and quantify visual parameters. The approach is useful for the investigator who is interested in taking advantage of the C. elegans excretory canal for identifying and characterizing genes involved in the phylogenetically conserved processes of intracellular lumen and unicellular tube morphogenesis.

The C. elegans Excretory Canal as a Model for Intracellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis in a Single Cell: labeling by GFP-fusions, RNAi Interaction Screen and Imaging

The C. elegans excretory canal is a unique single-cell model for the visual in vivo analysis of de novo polarized membrane biogenesis. This protocol describes a combination of standard genetic/RNAi and imaging approaches, adaptable for the identification and characterization of molecules directing unicellular tubulogenesis, and apical membrane and lumen biogenesis.

The four C. elegans excretory canals are narrow tubes extended through the length of the animal from a single cell, with almost equally far extended ...

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

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

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

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

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and differentiation. Typically, pHi dynamics are determined in cultured cells, which are amenable to measuring and experimentally manipulating pHi. However, the recent development of new tools and methodologies has made it possible to study pHi dynamics within intact, live tissue. For Drosophila research, one important development was the generation of a transgenic line carrying a pHi biosensor, mCherry::pHluorin. Here, we describe a protocol that we routinely use for imaging live Drosophila ovarioles to measure pHi in the epithelial follicle stem cell (FSC) lineage in mCherry::pHluorin transgenic wild type lines; however, the methods described here can be easily adapted for other tissues, including the wing discs and eye epithelium. We describe techniques for expressing mCherry::pHluorin in the FSC lineage, maintaining ovarian tissue during live imaging, and acquiring and analyzing images to obtain pHi values.

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

We provide a protocol for imaging intracellular pH of an epithelial stem cell lineage in live Drosophila ovarian tissue. We describe methods to generate transgenic flies expressing a pH biosensor, mCherry::pHluorin, image the biosensor using quantitative fluorescence imaging, generate standard curves, and convert fluorescence intensity values to pH values.

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and ...

Live Cell Imaging of Chromosome Segregation During Mitosis

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled by multiple mechanisms that include the Spindle Assembly Checkpoint (SAC). The SAC is part of a complex feedback system that is responsible for prevention of a cell progress through mitosis unless all chromosomal kinetochores have attached to spindle microtubules. Chromosomal lagging and abnormal chromosome segregation is an indicator of dysfunctional cell cycle control checkpoints and can be used to measure the genomic stability of dividing cells. Deregulation of the SAC can result in the transformation of a normal cell into a malignant cell through the accumulation of errors during chromosomal segregation. Implementation of the SAC and the formation of the kinetochore complex are tightly regulated by interactions between kinases and phosphatase such as Protein Phosphatase 2A (PP2A). This protocol describes live cell imaging of lagging chromosomes in mouse embryonic fibroblasts isolated from mice that had a knockout of the PP2A-B56&#947; regulatory subunit. This method overcomes the shortcomings of other cell cycle control imaging techniques such as flow cytometry or immunocytochemistry that only provide a snapshot of a cell cytokinesis status, instead of a dynamic spatiotemporal visualization of chromosomes during mitosis.

This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

Analysis of Epididymal Protein Synthesis and Secretion

The mammalian epididymis generates one of the most complex intraluminal fluids of any endocrine gland in order to support the post-testicular maturation and storage of spermatozoa. Such complexity arises due to the combined secretory and absorptive activity of the lining epithelial cells. Here, we describe the techniques for the analysis of epididymal protein synthesis and secretion by focusing on the model protein family of dynamin (DNM) mechanoenzymes; large GTPases that have the potential to regulate bi-directional membrane trafficking events. For the study of protein expression in epididymal tissue, we describe robust methodology for immunofluorescence labeling of target proteins in paraffin-embedded sections and the subsequent detection of the spatial distribution of these proteins via immunofluorescence microscopy. We also describe optimized methodology for the isolation and characterization of exosome like vesicles, known as epididymosomes, which are secreted into the epididymal lumen to participate in intercellular communication with maturing sperm cells. As a complementary approach, we also describe the immunofluorescence detection of target proteins in an SV40-immortalized mouse caput epididymal epithelial (mECap18) cell line. Moreover, we discuss the utility of the mECap18 cell line as a suitable in vitro model with which to explore the regulation of epididymal secretory activity. For this purpose, we describe the culturing requirements for the maintenance of the mECap18 cell line and the use of selective pharmacological inhibition regimens that are capable of influencing their secretory protein profile. The latter are readily assessed via harvesting of conditioned culture medium, concentration of secreted proteins via trichloroacetic acid/acetone precipitation and their subsequent analysis via SDS-PAGE and immunoblotting. We contend that these combined methods are suitable for the analysis of alternative epididymal protein targets as a prelude to determining their functional role in sperm maturation and/or storage.

Here, we report the immunofluorescence localization of dynamin to illustrate the protocols for the detection of proteins in paraffin-embedded mouse epididymal sections and those of an immortalized epididymal cell line (mECap18). We also describe the protocols for the isolation of secretory proteins from both epididymal fluid and conditioned cell media.

The mammalian epididymis generates one of the most complex intraluminal fluids of any endocrine gland in order to support the post-testicular maturation ...

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the number of novel cell lines is expected to increase. The DGRC aims to familiarize researchers with using Drosophila cell lines as an experimental tool to complement and drive their research agenda. Procedures for working with a variety of Drosophila cell lines with distinct characteristics are provided, including protocols for thawing, culturing, and cryopreserving cell lines. Importantly, this publication demonstrates the best practices required to work with Drosophila cell lines to minimize the risk of contaminations from adventitious microorganisms or from other cell lines. Researchers who become familiar with these procedures will be able to delve into the many applications that use Drosophila cultured cells including biochemistry, cell biology and functional genomics.

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

Drosophila cell lines are important reagents for both fundamental and biomedical research. This article provides protocols for thawing, subculturing, and the cryopreservation of commonly used Drosophila cell lines to assist researchers in incorporating the use of these reagents in their research.

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the ...

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A remarkable example is given by sensory epithelia, where cells of the same or distinct identities are brought together via cell-cell adhesion showing highly organized planar patterns. Cells align to one another in the same direction and display equivalent polarity over large distances. This organization of the mature epithelia is established over the course of morphogenesis. To understand how the planar arrangement of the mature epithelia is achieved, it is crucial to track cell orientation and growth dynamics with high spatiotemporal fidelity during development in vivo. Robust analytical tools are also essential to identify and characterize local-to-global transitions. The Drosophila pupa is an ideal system to evaluate oriented cell shape changes underlying epithelial morphogenesis. The pupal developing epithelium constitutes the external surface of the immobile body, allowing long-term imaging of intact animals. The protocol described here is designed to image and analyze cell behaviors at both global and local levels in the pupal abdominal epidermis as it grows. The methodology described can be easily adapted to the imaging of cell behaviors at other developmental stages, tissues, subcellular structures, or model organisms.

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

This protocol is designed for the imaging and analysis of the dynamics of cell orientation and tissue growth in the Drosophila abdominal epithelia as the fruit fly undergoes metamorphosis. The methodology described here can be applied to the study of different developmental stages, tissues, and subcellular structures in Drosophila or other model organisms.

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A ...

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of embryogenesis, these organelles undergo dramatic movements powered by actin-based cytoplasmic streaming and motor-driven trafficking along microtubules. The development of a multitude of small, organelle-specific fluorescent probes (FPs) makes it possible to visualize a wide range of different lipid-containing structures in any genotype, allowing live imaging without requiring a genetically encoded fluorophore. This protocol shows how to inject vital dyes and molecular probes into Drosophila embryos to monitor the trafficking of specific organelles by live imaging. This approach is demonstrated by labeling lipid droplets (LDs) and following their bulk movement by particle image velocimetry (PIV). This protocol provides a strategy amenable to the study of other organelles, including lysosomes, mitochondria, yolk vesicles, and the ER, and for tracking the motion of individual LDs along microtubules. Using commercially available dyes brings the benefits of separation into the violet/blue and far-red regions of the spectrum. By multiplex co-labeling of organelles and/or cytoskeletal elements via microinjection, all the genetic resources in Drosophila are available for trafficking studies without the need to introduce fluorescently tagged proteins. Unlike genetically encoded fluorophores, which have low quantum yields and bleach easily, many of the available dyes allow for rapid and simultaneous capture of several channels with high photon yields.

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

In the early Drosophila embryo, many organelles are motile. In principle, they can be imaged live via specific fluorescent probes, but the eggshell prevents direct application to the embryo. This protocol describes how to introduce such probes via microinjection, and then analyze bulk organelle motion via particle image velocimetry.

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of ...

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

The workhorse of developmental biology is the confocal microscope, which allows researchers to determine the three-dimensional localization of tagged molecules within complex biological samples. While traditional confocal microscopes allow one to resolve two adjacent fluorescent point sources located a few hundred nanometers apart, observing the finer details of subcellular biology requires the ability to resolve signals in the order of tens of nanometers. Numerous hardware-based methods for super-resolution microscopy have been developed to allow researchers to sidestep such resolution limits, although these methods require specialized microscopes that are not available to all researchers. An alternative method for increasing resolving power is to isotropically enlarge the sample itself through a process known as expansion microscopy (ExM), which was first described by the Boyden group in 2015. ExM is not a type of microscopy per se but is rather a method for swelling a sample while preserving the relative spatial organization of its constituent molecules. The expanded sample can then be observed at an effectively increased resolution using a traditional confocal microscope. Here, we describe a protocol for implementing ExM in whole-mount Drosophila embryos, which is used to examine the localization of Par-3, myosin II, and mitochondria within the surface epithelial cells. This protocol yields an approximately four-fold increase in sample size, allowing for the detection of subcellular details that are not visible with conventional confocal microscopy. As proof of principle, an anti-GFP antibody is used to distinguish distinct pools of myosin-GFP between adjacent cell cortices, and fluorescently labeled streptavidin is used to detect endogenous biotinylated molecules to reveal the fine details of the mitochondrial network architecture. This protocol utilizes common antibodies and reagents for fluorescence labeling, and it should be compatible with many existing immunofluorescence protocols.

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

Here, a protocol for the implementation of expansion microscopy in early Drosophila embryos to achieve super-resolution imaging using a conventional laser-scanning confocal microscope is presented.

The workhorse of developmental biology is the confocal microscope, which allows researchers to determine the three-dimensional localization of tagged ...