We would like to thank Dr. Jennifer A. Zallen for providing the Sqh-GFP Drosophila line and support for the initial development of this technique, and Dr. Francois Schweisguth for providing the Notch-GFP Drosophila line. This work was supported by funding from the National Natural Science Foundation of China (32270809) to H.H.Yu, generous financial and staff support from the School of Life Sciences, SUSTech, and funding to Y. Yan from Shenzhen Science and Technology Innovation Commission/JCYJ20200109140201722.

The experiments were conducted in accordance with the guidelines and approval of the School of Life Sciences, SUSTech University. The organism used is Drosophila melanogaster, and the genotypes are Notch-Knockin-GFP (Chromosome X) and Sqh-sqh-GFP (Chromosome II), generously provided by the labs of Dr. Francois Schweisguth (Institute Pasteur) and Dr. Jennifer Zallen (Sloan Kettering Institute), respectively. While this protocol mainly focuses on aspects of antibody labeling and live imaging, please refer to published reports for more detailed descriptions of Drosophila embryo collection and injection15,16.
1. Fluorescent labeling of antibodies
<ol>
	<li>Preferably, use monoclonal antibodies or nanobodies for the protein of interest. Prepare the stock concentration of antibodies at 1 mg/mL or higher. 
	&#8203;NOTE: In this study, GFP nanobody (see Table of Materials) is used as an example. The commercially available GFP nanobody is packaged at a concentration of 1.0 mg/mL and a volume of 250 &#181;L. Additionally, a commercially available antibody labeling kit (see Table of Materials) is utilized, which conjugates Alexa Fluor 594 to primary amines of proteins through a succinimidyl ester reaction11. Importantly, this labeling process does not significantly alter the antibody concentration.</li>
	<li>Prepare a 1 M sodium bicarbonate solution by resuspending Component B (provided in the antibody labeling kit) in 1 mL of deionized water.</li>
	<li>Adjust the antibody concentration to 1.0 mg/mL, then add 1/10th volume (10 &#181;L for 100 &#181;L GFP nanobody) of the 1 M sodium bicarbonate solution.</li>
	<li>Add 110 &#181;L of the antibody mix (from step 1.3) directly to the tube containing Alexa Fluor 594 dye. Invert to mix (do not vortex) and incubate on a rotator for 1 h at room temperature.</li>
	<li>Assemble the purification column from the conjugation kit (see Table of Materials). Prepare a 1.5 mL resin bed (provided in the antibody labeling kit) and centrifuge the column at 1100 x g for 3 min at room temperature (RT) to remove excess liquid from the resin.</li>
	<li>Add the reaction mix (from step 1.4) dropwise to the top of the resin column and centrifuge at 1100 x g for 5 min at 4 &#176;C to collect the labeled antibody. The collected antibody should appear pink in color, and the volume should be slightly less than 100 &#181;L. Store in foil-wrapped or dark-colored tubes at 4 &#176;C.</li>
</ol>
2. Preparation of Drosophila embryos
<ol>
	<li>Place 200 newly hatched adult Drosophila in a plastic cylinder-shaped cage (diameter = 5 cm, height = 8 cm) with a male-to-female ratio of 1: 10. Seal one side with porous metal mesh for ventilation and the other side with a fruit-juice/yeast-paste agar plate.</li>
	<li>Change the plate every 12 h for three days before embryo collection. This allows synchronization of Drosophila egg laying and improves collection efficiency.</li>
	<li>On the day of injection, change the cage to a fruit-juice plate with minimal yeast paste and collect embryos at 25 &#176;C for 1 h. If the first round of collection does not yield sufficient embryos (&#60;50 embryos), discard the first plate and repeat this step for a second round of collection until more than 100 embryos are laid within 1 h.</li>
	<li>Remove the plate from the cage to stop the egg laying and incubate at 25 &#176;C for another 2 h to obtain embryos that are 2-3 h old. The correct stage of embryos is critical for the success of antibody injection.</li>
	<li>To remove the eggshell of embryos, add 2 mL of 50% bleach to the fruit-juice plate and gently dislodge embryos from the plate using a paintbrush (Figure 2A-4). Often, embryos will be laid directly on top of the yeast paste. In this case, it is acceptable to brush the yeast paste into the bleach solution to collect all the embryos.</li>
	<li>Incubate the floating embryos in 50% bleach for 2 min and swirl the fruit-juice plate every 30 s to improve the efficiency of eggshell removal.</li>
	<li>Transfer the embryo/bleach mix by pouring it into a 70 &#181;m nylon cell strainer (Figure 2A-7). Wash the embryos thoroughly using a water bottle to remove any residual bleach and yeast paste. Dry the cell strainer completely with paper towels, ensuring that any excess water around the embryos is removed.</li>
</ol>
3. Embryo alignment and desiccation
<ol>
	<li>Prepare a 5 cm x 5 cm x 0.5 cm (width, length, height) 4% agarose gel (Figure 2A-9) and allow the gel to cool down to room temperature. Cut a 1 cm x 3 cm x 0.5 cm (width, length, height) agarose block using a clean razor blade and place the block on a glass slide (Figure 2A-2). Examine the gel block under the dissecting scope to ensure that the edges are cut straight, and the surface is flat and dry.</li>
	<li>Transfer embryos from the strainer onto the gel block using a paintbrush. Spread the embryos evenly along the midline of the block by brushing gently. Ideally, clusters of embryos are separated into single ones or doublets without touching each other.</li>
	<li>Prepare a tweezer with two legs taped tightly together to create a single fine tip (Figure 2A-5). Under a dissecting scope, pick up individual embryos using the tweezer and place them along the long edge of the gel block. Align 20-30 embryos, ensuring their anterior-posterior axis is parallel to the edge (Figure 2C).</li>
	<li>Pipette 10 &#181;L heptane glue (see Table of Materials) at the center of a 24 mm x 50 mm coverslip (Figure 2A-1) and spread the glue into a 0.5 cm x 3 cm rectangular area using a pipette tip. Wait for the glue to completely dry before use.</li>
	<li>Place the glass slide with the gel block at the edge of the desk, with embryos facing outward. Lift the coverslip with glue using a flat-tip tweezer (Figure 2A-6) and hold it still on top of the embryos, with the glue side facing toward the embryos at a tilted angle of around 45 degrees. 
	NOTE: Gently press the coverslip against the gel block so that the embryos will be in contact with the glue, and then quickly release the pressure and lift the coverslip. The amount of tension applied is critical so that embryos can be stably attached to the glue without being pressed too hard and bursting.</li>
	<li>Pipette 10 &#181;L water at the center of a new glass slide and place the coverslip with the embryos on top of it, with the side of the embryo facing upwards (Figure 2B). Ensure to use water instead of nail polish or other adhesive glue for attaching the coverslip to the glass slide, as this side of the coverslip will be placed directly on top of the confocal lens for live imaging.</li>
	<li>Dry the embryos in a desiccation chamber (Figure 2A-3) (see Table of Materials) for 10-15 min until the vitelline membrane wrinkles (Figure 2E). The required time may vary with the humidity of the room and should be experimentally tested for each lab condition.</li>
	<li>Pipette 40 &#181;L of halocarbon oil (a mix of type 27 and type 700 at a 1: 1 ratio in volume, see Table of Materials) at one end of the embryo strip. Tilt the slide until the halocarbon oil covers the entire surface of the embryos.</li>
</ol>
4. Antibody Injection and imaging
<ol>
	<li>Prepare a few injection glass needles using a micropipette puller (see Table of Materials for parameter settings) and load each needle with 5 &#181;L of AlexaFluor-labeled antibody solution (from step 1.6).</li>
	<li>Place a 25 mm x 25 mm coverslip on top of a glass slide. Pipette 40 &#181;L of halocarbon oil and spread it along the edge of the coverslip.</li>
	<li>Under the injection scope, align the tip of the needle against the edge of the coverslip under oil. Adjust the injection air pressure of the picopump (see Table of Materials) so that one pump generates one bubble filled with antibodies. The bubble diameter should be limited to 20-50 &#181;m (Figure 2F).</li>
	<li>Replace the empty glass slide with one containing embryos under oil. Align the injection needle tip perpendicularly against the anterior-posterior axis of the embryos (Figure 2D,G).</li>
	<li>Check the stage of the embryos to ensure that most have initiated cellularization but have not yet started gastrulation (stage 4 and stage 5), remaining as a syncytium (Figure 2F,G). 
	NOTE: The hallmark of this stage is the absence of any grooves or folds and the presence of an oval-shaped, dark-colored yolk sac visible at the center of the embryo. Morphological characterization of the embryo stage under oil is based on the book chapter &#34;Stages of Drosophila Embryogenesis&#34; by Volker Hartenstein17.</li>
	<li>Use the X-Y stage manipulator to move embryos toward the needle until the tip arrives at the center of the yolk. Pump two to three times to inject antibodies into the yolk. Successful injection is indicated by the quick disappearance of the vitelline membrane wrinkle as embryos gain volume from the antibody solution (Figure 2G).</li>
	<li>Use the X-Y stage manipulator to move embryos away from the needle after injection and move downward to the next embryo. Repeat step 6 until all embryos on the slide are injected.</li>
	<li>Transfer the glass slide with injected embryos to a humidity chamber (Figure 2A-8). Incubate at 25 &#176;C until the desired stage of embryo development is achieved.</li>
	<li>Lift the 24&#160;mm x 50 mm coverslip off the glass slide and place it directly into the slide holder of the microscope. The side without embryos will be facing the objectives. Locate the embryos using a low-magnification lens under bright field illumination and switch to a 40x or 63x lens for high-resolution fluorescent live imaging.</li>
</ol>

Enhancing Protein Visualization with Antibody-Mediated Live Imaging

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The authors have no conflicts of interest to declare.

This presented procedure outlines the specialized method of fluorescence labeling with custom antibodies and subsequent injection into early-stage Drosophila embryos. This technique facilitates real-time visualization of proteins or post-translational modifications that exist in low quantities and are typically difficult to observe through conventional GFP/mCherry tagging methods.
Caution should be exercised when extending this method to make quantitative comparisons between wild-type...

Immunofluorescence is a cornerstone technique of modern cell biology originally developed by Albert Coons, which enables the detection of molecules at their native cellular compartments and characterization of the molecular compositions of subcellular organelles or machineries1. Coupled with genetic manipulations, immunofluorescence helps establish the now well-accepted concept that protein localization is essential for its function2. Aside from specific primary antibodies and bright fluorescent dyes, the success of this technique relies on a preliminary process named fixation and permeabilization, which preserves cellular morphologies, immobilizes antigens, and increases the accessibility of antibodies into intracellular compartments. Inevitably, the fixation and permeabilization process would kill cells and terminate all biological processes3. Therefore, immunofluorescence only provides snapshots of the life journey of proteins. However, many biological processes such as cell migration and divisions are dynamic in nature, requiring investigation of protein behaviors in a spatial-temporally resolved manner4,5.
To examine protein dynamics in living organisms, live imaging methods based on genetically encoded fluorescent proteins such as green fluorescent protein (GFP)6 and high-speed confocal microscopes have been developed. Briefly, the protein of interest can be genetically manipulated to be fused with GFP7, and then ectopically expressed from viral or yeast promoters such as cytomegalovirus (CMV)8 or upstream activation sequence (UAS)9. Because GFP is autofluorescent in nature, no fluorophore-coupled antibodies are required to reveal the localization of target proteins, which bypasses the necessity of preliminary processes of fixation or permeabilization. Over the last two decades, fluorescent tags spanning the whole spectrum of wavelength have been developed10, enabling multi-color live imaging of several target proteins at the same time. However, compared to chemically engineered fluorescent dyes such as AlexaFluor or ATTO, the autofluorescence of these genetically encoded fluorescent proteins is relatively weak and unstable when expressed from endogenous promoters, especially during live imaging over longer time scales10. While this shortfall can be mitigated by over-expressing fluorescently tagged target proteins, many with enzymatic activities such as kinases and phosphatases severely disrupt normal biological processes if not expressed at physiological levels.
This protocol presents a method that enables photostable antibody-based target illumination in a live image setup, essentially allowing immunofluorescence without the process of fixation or permeabilization (Figure 1). Through a simple NHS-based primary amine reaction11, one can conjugate fluorescent dyes such as AlexaFluor 488 or 594 with essentially any primary antibody or GFP/HA/Myc nanobody12. Taking advantage of a developmental feature that all Drosophila embryonic cells share a common cytoplasm during the syncytium stage13, one can achieve antigen binding and illumination across entire embryos after the injection of dye-conjugated antibodies. With expanding libraries of endogenously tagged proteins available in Drosophila and other model systems14, this method can potentially broaden applications of these libraries by revealing dynamics of low-abundance fluorescently tagged proteins and other non-fluorescently tagged (HA/Myc-tagged) proteins in living tissues.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Agarose</td>
			<td>Sangon Biotech</td>
			<td>&#160;A620014&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 594 Antibody Labeling Kit</td>
			<td>Invitrogen&#160;</td>
			<td>A20185</td>
			<td>Purification column from step 1.6 is included in this kit</td>
		</tr>
		<tr>
			<td>Biological Microscope</td>
			<td>SOPTOP</td>
			<td>EX20</td>
			<td>Eyepiece lens: PL 10X/20. Objective lens: 10x/0.25</td>
		</tr>
		<tr>
			<td>Bleach</td>
			<td>Clorox&#174;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate Glass Capillaries</td>
			<td>World Precision Instruments</td>
			<td>TW100F-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>Eppendorf</td>
			<td>5245</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Strainer</td>
			<td>FALCON</td>
			<td>352350</td>
			<td></td>
		</tr>
		<tr>
			<td>Desiccation chamber&#160;</td>
			<td>LOCK&#38;LOCK</td>
			<td>HSM8200</td>
			<td>320ml</td>
		</tr>
		<tr>
			<td>Dissecting Microscope</td>
			<td>Mshot</td>
			<td>MZ62</td>
			<td>Eyepiece lens: WF10X/22mm.</td>
		</tr>
		<tr>
			<td>Double-sided Tape</td>
			<td>Scotch</td>
			<td>665</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Super Tweezer</td>
			<td>VETUS</td>
			<td>ST-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand&#8482; Cover Glasses: Rectangles</td>
			<td>Fisherbrand</td>
			<td>12-545F</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand&#8482; Superfrost&#8482; Plus Microscope Slides</td>
			<td>Fisherbrand</td>
			<td>12-550-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Forcep</td>
			<td>VETUS</td>
			<td>33A-SA</td>
			<td></td>
		</tr>
		<tr>
			<td>Halocarbon oil 27</td>
			<td>Sigma-Aldrich</td>
			<td>H8773-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Halocarbon oil 700</td>
			<td>Sigma-Aldrich</td>
			<td>H8898-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Heptane</td>
			<td>Sigma-Aldrich</td>
			<td>H2198-1L</td>
			<td>Heptane glue is made of double-sided tape immersed in heptane</td>
		</tr>
		<tr>
			<td>Dehydration reagent&#160;</td>
			<td>TOKAI</td>
			<td>1-7315-01</td>
			<td>Fill to 90% volume of the dessication chamber</td>
		</tr>
		<tr>
			<td>Manual Micromanipulator</td>
			<td>World Precision Instruments</td>
			<td>M3301R</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>World Precision Instruments</td>
			<td>PUL-1000</td>
			<td>Procedure: step 1, Heat: 290, Force:300, Distance:1.00, Delay:50. 
			Step 2, Heat: 290, Force:300, Distance:2.21, Delay:50&#160;</td>
		</tr>
		<tr>
			<td>Pneumatic picopump</td>
			<td>World Precision Instruments</td>
			<td>PV 830</td>
			<td>Eject: 20 psi;&#160; Range: 100ms; Duration: timed</td>
		</tr>
		<tr>
			<td>PY20</td>
			<td>Santa Cruz</td>
			<td>SC-508</td>
			<td></td>
		</tr>
		<tr>
			<td>Square petri dishes</td>
			<td>Biosharp</td>
			<td>BS-100-SD</td>
			<td></td>
		</tr>
		<tr>
			<td>GFP nanobody</td>
			<td>Chromotek</td>
			<td>gt</td>
			<td></td>
		</tr>
	</tbody>
</table>

visualizing low-abundance proteins and post-translational modifications in living drosophila embryos via fluorescent antibody injection

Visualization of proteins in living cells using GFP (Green Fluorescent Protein) and other fluorescent tags has greatly improved understanding of protein localization, dynamics, and function. Compared to immunofluorescence, live imaging more accurately reflects protein localization without potential artifacts arising from tissue fixation. Importantly, live imaging enables quantitative and temporal characterization of protein levels and localization, crucial for understanding dynamic biological processes such as cell movement or division. However, a major limitation of fluorescent tagging approaches is the need for sufficiently high protein expression levels to achieve successful visualization. Consequently, many endogenously tagged fluorescent proteins with relatively low expression levels cannot be detected. On the other hand, ectopic expression using viral promoters can sometimes lead to protein mislocalization or functional alterations in physiological contexts. To address these limitations, an approach is presented that utilizes highly sensitive antibody-mediated protein detection in living embryos, essentially performing immunofluorescence without the need for tissue fixation. As proof of principle, endogenously GFP-tagged Notch receptor that is barely detectable&#160;in living embryos can be successfully visualized after antibody injection. Furthermore, this approach was adapted to visualize post-translational modifications (PTMs) in living embryos, allowing the detection of temporal changes in tyrosine phosphorylation patterns during early embryogenesis and revealing a novel subpopulation of phosphotyrosine (p-Tyr) underneath apical membranes. This approach can be modified to accommodate other protein-specific, tag-specific, or PTM-specific antibodies and should be compatible with other injection-amenable model organisms or cell lines. This protocol opens new possibilities for live imaging of low-abundance proteins or PTMs that were previously challenging to detect using traditional fluorescent tagging methods.

To demonstrate the advantages of the antibody injection method over fluorescent-tag-based live imaging or immunofluorescence, two case studies are provided that characterize the dynamic localization of a low-abundance transmembrane receptor, Notch, and a type of post-translational modification called tyrosine phosphorylation in living embryos.
Notch signaling activity plays a major role in cell fate determination during embryogenesis and adult organ homeostasis18,<...

Watch this Scientific Journal Video about Unveiling the Dynamics of Mechanical and Biochemical Signals in Animal Morphogenesis Research at JoVE.com

Author Spotlight: Unveiling the Dynamics of Mechanical and Biochemical Signals in Animal Morphogenesis Research 

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

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

Visualization of proteins in living cells using GFP (Green Fluorescent Protein) and other fluorescent tags has greatly improved understanding of protein ...

visualizing-low-abundance-proteins-post-translational-modifications

Research

JoVE Journal

Developmental Biology

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

Video: Author Spotlight: Unveiling the Dynamics of Mechanical and Biochemical Signals in Animal Morphogenesis Research 

Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes

We report here a robust and efficient protocol for the expression of fluorescent proteins after mRNA injection into unfertilized oocytes of the cephalochordate amphioxus, Branchiostoma lanceolatum. We use constructs for membrane and nuclear targeted mCherry and eGFP that have been modified to accommodate amphioxus codon usage and Kozak consensus sequences. We describe the type of injection needles to be used, the immobilization protocol for the unfertilized oocytes, and the overall injection set-up. This technique generates fluorescently labeled embryos, in which the dynamics of cell behaviors during early development can be analyzed using the latest in vivo imaging strategies. The development of a microinjection technique in this amphioxus species will allow live imaging analyses of cell behaviors in the embryo as well as gene-specific manipulations, including gene overexpression and knockdown. Altogether, this protocol will further consolidate the basal chordate amphioxus as an animal model for addressing questions related to the mechanisms of embryonic development and, more importantly, to their evolution.

Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes

We report here the robust and efficient expression of fluorescent proteins after mRNA injection into unfertilized oocytes of Branchiostoma lanceolatum. The development of the microinjection technique in this basal chordate will pave the way for far-reaching technical innovations in this emerging model system, including in vivo imaging and gene-specific manipulations.

We report here a robust and efficient protocol for the expression of fluorescent proteins after mRNA injection into unfertilized oocytes of the ...

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological systems. In FDAP experiments, the clearance of proteins within living organisms can be measured. A protein of interest is tagged with a photoconvertible fluorescent protein, expressed in vivo and photoconverted, and the decrease in the photoconverted signal over time is monitored. The data is then fitted with an appropriate clearance model to determine the protein half-life. Importantly, the clearance kinetics of protein populations in different compartments of the organism can be examined separately by applying compartmental masks. This approach has been used to determine the intra- and extracellular half-lives of secreted signaling proteins during zebrafish development. Here, we describe a protocol for FDAP experiments in zebrafish embryos. It should be possible to use FDAP to determine the clearance kinetics of any taggable protein in any optically accessible organism.

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP

Protein levels in cells and tissues are often tightly regulated by the balance of protein production and clearance. Using Fluorescence Decay After Photoconversion (FDAP), the clearance kinetics of proteins can be experimentally measured in vivo.

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological ...

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. The observation of Tribolium embryos with light sheet-based fluorescence microscopy has multiple advantages over conventional widefield and confocal fluorescence microscopy. Due to the unique properties of a light sheet-based microscope, three dimensional images of living specimens can be recorded with high signal-to-noise ratios and significantly reduced photo-bleaching as well as photo-toxicity along multiple directions over periods that last several days. With more than four years of methodological development and a continuous increase of data, the time seems appropriate to establish standard operating procedures for the usage of light sheet technology in the Tribolium community as well as in the insect community at large. This protocol describes three mounting techniques suitable for different purposes, presents two novel custom-made transgenic Tribolium lines appropriate for long-term live imaging, suggests five fluorescent dyes to label intracellular structures of fixed embryos and provides information on data post-processing for the timely evaluation of the recorded data. Representative results concentrate on long-term live imaging, optical sectioning and the observation of the same embryo along multiple directions. The respective datasets are provided as a downloadable resource. Finally, the protocol discusses quality controls for live imaging assays, current limitations and the applicability of the outlined procedures to other insect species.
This protocol is primarily intended for developmental biologists who seek imaging solutions that outperform standard laboratory equipment. It promotes the continuous attempt to close the gap between the technically orientated laboratories/communities, which develop and refine microscopy methodologically, and the life science laboratories/communities, which require 'plug-and-play' solutions to technical challenges. Furthermore, it supports an axiomatic approach that moves the biological questions into the center of attention.

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

Imaging the morphogenesis of insect embryos with light sheet-based fluorescence microscopy has become state of the art. This protocol outlines and compares three mounting techniques appropriate for Tribolium castaneum embryos, introduces two novel custom-made transgenic lines well suited for live imaging, discusses essential quality controls and indicates current experimental limitations.

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and optical transparency. In particular, the zebrafish embryo has become an important organism to study vertebrate kidney organogenesis as well as multiciliated cell (MCC) development. To visualize MCCs in the embryonic zebrafish kidney, we have developed a combined protocol of whole-mount fluorescent in situ hybridization (FISH) and whole mount immunofluorescence (IF) that enables high resolution imaging. This manuscript describes our technique for co-localizing RNA transcripts and protein as a tool to better understand the regulation of developmental programs through the expression of various lineage factors.

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

Cilia development is vital to proper organogenesis. This protocol describes an optimized method to label and visualize ciliated cells of the zebrafish.

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and ...

Single-cell Photoconversion in Living Intact Zebrafish

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause disease when disrupted. Scientists have developed clever techniques to investigate characteristics and natural dynamics of these cells within intact tissue by expressing fluorescent proteins in subsets of cells. However, at times, experiments require more selected visualization of cells within the tissue, sometimes at the single-cell or population-of-cells manner. To achieve this and visualize single cells within a population of cells, scientists have utilized single-cell photoconversion of fluorescent proteins. To demonstrate this technique, we show here how to direct UV light to an Eos-expressing cell of interest in an intact, living zebrafish. We then image those photoconverted Eos+ cells 24 h later to determine how they changed in the tissue. We describe two techniques: single cell photoconversion and photoconversions of populations of cell. These techniques can be used to visualize cell-cell interactions, cell-fate and differentiation, and cell migrations, making it a technique that is applicable in numerous biological questions.

Here, we present a protocol to show how cell photoconversion is achieved through UV exposure to specific areas expressing the fluorescent protein, Eos, in living animals.

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

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 ...

Fast and Efficient Expression of Multiple Proteins in Avian Embryos Using mRNA Electroporation

We report that mRNA electroporation permits fluorescent proteins to label cells in living quail embryos more quickly and broadly than DNA electroporation. The high transfection efficiency permits at least 4 distinct mRNAs to be co-transfected with ~87% efficiency. Most of the electroporated mRNAs are degraded during the first 2 h post-electroporation, permitting time-sensitive experiments to be carried out in the developing embryo. Finally, we describe how to dynamically image live embryos electroporated with mRNAs that encode various subcellular targeted fluorescent proteins.

We report messenger RNA (mRNA) electroporation as a method that permits fast and efficient expression of multiple proteins in the quail embryo model system. This method can be used to fluorescently label cells and record their in vivo movements by time-lapse microscopy shortly after electroporation.

We report that mRNA electroporation permits fluorescent proteins to label cells in living quail embryos more quickly and broadly than DNA electroporation. ...

Microinjection of DNA into Eyebuds in Xenopus laevis Embryos and Imaging of GFP Expressing Optic Axonal Arbors in Intact, Living Xenopus Tadpoles

The primary visual projection of tadpoles of the aquatic frog Xenopus laevis serves as an excellent model system for studying mechanisms that regulate the development of neuronal connectivity. During establishment of the retino-tectal projection, optic axons extend from the eye and navigate through distinct regions of the brain to reach their target tissue, the optic tectum. Once optic axons enter the tectum, they elaborate terminal arbors that function to increase the number of synaptic connections they can make with target interneurons in the tectum. Here, we describe a method to express DNA encoding green fluorescent protein (GFP), and gain- and loss-of-function gene constructs, in optic neurons (retinal ganglion cells) in Xenopus embryos. We explain how to microinject a combined DNA/lipofection reagent into eyebuds of one day old embryos such that exogenous genes are expressed in single or small numbers of optic neurons. By tagging genes with GFP or co-injecting with a GFP plasmid, terminal axonal arbors of individual optic neurons with altered molecular signaling can be imaged directly in brains of intact, living Xenopus tadpoles several days later, and their morphology can be quantified. This protocol allows for determination of cell-autonomous molecular mechanisms that underlie the development of optic axon arborization in vivo.

Microinjection of DNA into Eyebuds in Xenopus laevis Embryos and Imaging of GFP Expressing Optic Axonal Arbors in Intact, Living Xenopus Tadpoles

This protocol aims to demonstrate how to microinject a DNA/DOTAP mixture into eyebuds of one day old Xenopus laevis embryos, and how to image and reconstruct individual green fluorescent protein (GFP) expressing optic axonal arbors in tectal midbrains of intact, living Xenopus tadpoles.

The primary visual projection of tadpoles of the aquatic frog Xenopus laevis serves as an excellent model system for studying mechanisms that regulate the ...