S.B.&#39;s PhD grant is funded by Region Bretagne (ARED grant Number COH20020) and Sorbonne Universit&#233;. I.T.is PhD grant is funded by Region Bretagne (ARED grant Number COH18020) and the Norvegian NMBU University. This project has received financial support from the CNRS through the MITI interdisciplinary programs.&#160;MRic is member of the national infrastructure France-BioImaging supported by the French National Research Agency (ANR-10-INBS-04).

1. Production of Saccharina latissima gametophytes
<ol>
	<li>Collect mature sporophytes of S. latissima from the wild as previously described20,21. Ensure that the selected sporophytes are devoid of epiphytes (small organisms visible on the blade&#39;s surface) or internal parasites (found in the bleached areas or spots on the blade). </li>
	<li>Using a scalpel, cut the darkest part in the center of the blade (fertile spore-producing tissue22) into 1-5 square pieces (1 cm&#178;), avoiding any bleached spots, if present.</li>
	<li>Remove any remaining epiphytes by gently cleaning the cut pieces with the back of a scalpel and some absorbent paper.</li>
	<li>Place the cleaned pieces in a glass dish filled with sterile natural seawater (see Table of Materials) for 45 min to release spores following previously published report22.</li>
	<li>Remove the blade pieces and filter the seawater through a 40 &#181;m cell strainer to remove debris or unwanted organisms.</li>
	<li>Dilute the spores in the filtrate to a 20-40 spores/mL concentration in plastic Petri dishes22.</li>
	<li>Place the spore solution in a culture cabinet (see Table of Materials) configured with the optimal culture conditions (13 &#176;C, 24 &#181;E.m-2.s-1, photoperiod 16:8 L:D).</li>
	<li>Allow the spores to germinate and develop into gametophytes. 
	NOTE: Spore germination is visible after 2 days in the cabinet, and the first cell division of the gametophytic cells usually occurs within the following 48 h.</li>
	<li>Replace the growth medium after 5 days with micro-filtered natural seawater enriched with a 0.5x Provasoli solution (NSW1/2) (see Table of Materials). 
	NOTE: To avoid repeating these steps, specific male and female gametophytes can be selected and vegetatively propagated for several months. The gametophytes remain vegetative when grown under red light (4 &#181;E.m-2.s-1 with a wavelength of at least 580 nm)23 in the same culture conditions described above (step 1.7).</li>
</ol>
2. Fragmentation and induction of oogenesis
<ol>
	<li>Harvest gametophytes with a cell scraper.</li>
	<li>Using a small plastic pestle, crush the collected gametophytes in a 1.5 mL tube into 4-5-celled pieces.</li>
	<li>Fill the tube with 1 mL NSW1/2 (step 1.9).</li>
	<li>Add 2.5 &#181;L of the crushed gametophyte solution into 3 mL of natural seawater enriched with 1x Provasoli solution (NSW) and place them in a Petri dish. 
	NOTE: A 25 mm, glass-bottomed Petri dish is recommended for easier handling.</li>
	<li>Place the prepared dishes in a culture cabinet and induce gametogenesis at 13 &#176;C under white light with an intensity of 24 &#181;E.m-2.s-1&#160;(dim light, photoperiod 16:8 L:D). 
	NOTE: The first gametangia (female oogonia and male archegonia) can be observed after 5 days. The male is hyper-branched with small cells, and the female is composed of larger cells forming long filaments15,22. The first eggs are observed ~10 days later, and the first division of zygotes usually occurs within the following 2 days.</li>
	<li>Six days after observing the first eggs, transfer the dishes to brighter white light conditions: 50 &#181;E.m-2.s-1, photoperiod 16:8 L:D, still at 13 &#176;C.</li>
</ol>
3. Image acquisition for selecting embryos for ablation and monitoring subsequent growth
<ol>
	<li>Image the entire Petri dish to (re)locate the embryos selected during the ablation step (no need to return the dish to the ocular microscope) and monitor the subsequent development of the selected embryos. 
	NOTE: Use an inverted laser scanning confocal microscope (see Table of Materials) for imaging (Figure 1), and the laser ablation is described in step 5.</li>
	<li>Place the Petri dish on the stage and orientate it with a visual mark (e.g., draw a line with a permanent marker).</li>
	<li>Use the 10x/0.45 objective to focus on an embryo. Record the position of the four cardinal points of the Petri dish.</li>
	<li>Start the tile scan. Acquire transmitted/fluorescent images of the whole Petri dish at low resolution: 256 x 256 pixels, a pixel dwell time of 1.54 &#181;s with bidirectional scanning, and a digital zoom of 0.6x using a 561 nm laser at 1.2% transmission. 
	NOTE: The scan time for a whole 2.5 cm Petri dish is ~6 min for 225 tiles (Figure 2). Here, the 561 nm laser was used for transmission and fluorescence imaging.&#160;The fluorescence signal was collected between 580&#8211;720 nm on the confocal photomultipliers (PMT) and the transmitted light was collected on the transmitted PMT. The 561 nm laser can also monitor chlorophyll at this step, but it is unnecessary because it only helps distinguish the signal noise and the organisms correctly.</li>
	<li>Save the tile scan image and keep it open in the image acquisition software (see Table of Materials) window.</li>
	<li>Change the objective, do not remove the Petri dish. 
	NOTE: The 40x/1.2 water objective was moved to the side of the stage so that the immersion medium (water) could be added to the objective without moving the Petri dish from its initial position.</li>
	<li>Navigate through the previously acquired tile scan image to select the appropriate embryo. Once an embryo has been identified on this image, move the stage to the exact position of the embryo and acquire transmitted/fluorescent images of that embryo at high resolution. 
	NOTE: High-resolution settings: 512 x 512 pixels, 0.130 &#181;m/pixel, 0.208 &#181;s pixel dwell time with mono-directional scanning and 2x digital zoom using a 561 nm laser at 0.9% transmission.</li>
	<li>Annotate the tile scan image for each embryo candidate for laser ablation (Figure 2B) and proceed to the laser ablation step.</li>
</ol>
4. Laser calibration
<ol>
	<li>Calibrate the laser and synchronize the image acquisition software with the laser-driver software in the &#34;Click &#38; Fire mode&#34; and a pulsed 355 nm laser. 
	NOTE: This step is crucial to ensure perfect synchronization between the mouse cursor&#39;s position in the laser-driver software (see Table of Materials) with the position in the live image of the acquisition software.</li>
	<li>Open the laser-driver software package and click on Live in the image acquisition software package.</li>
	<li>Synchronize both software packages by clicking on Start acquisition in the laser-driver software package. The live image is now also recorded in the UV laser-driver software.</li>
	<li>Define an area of interest (AOI) by clicking on Choose AOI button and clicking on the edges of the image (right, left, top and bottom) in the UV laser-driver software package. 
	NOTE: After this calibration step, the settings for pixel size, image format, and zoom in the acquisition software package must remain constant.</li>
	<li>Select an empty area on the dish and lower the level of the stage to 20 &#181;m below the sample focal plane to focus on the glass bottom.</li>
	<li>Set the ablation laser and imaging laser trajectories by clicking on Start calibration and choose Manual calibration.</li>
	<li>Select a laser power high enough to see a black dot in the center of the live image corresponding to the hole in the glass coverslip (all the shutters must be open).</li>
	<li>Click on this central black dot with the mouse cursor and click on 18 additional dots proposed by the software to complete the alignment procedure.</li>
	<li>Check the calibration in the &#34;Click &#38; Fire mode&#34; on the same coverslip. 
	NOTE: Laser calibration depends on the imaging parameters. Once the laser has been calibrated, ensure that the imaging parameters (i.e., 512 x 512 pixels, 0.130 &#181;m/pixel, 0.208 &#181;s pixel dwell time with mono-directional scanning and 2x digital zoom) have not changed.</li>
</ol>
5. Laser ablation
<ol>
	<li>Select an embryo of interest. Start a time-lapse recording in the image-acquisition software.</li>
	<li>Acquire transmitted/fluorescence images with a 40x/1.2 W objective at high resolution (i.e., 512 x 512 pixels, 0.130 &#181;m/pixel, 1.54 &#181;s pixel dwell time with mono-directional scanning and 2x digital zoom using a 561 nm laser at 0.9% transmission). Acquire the time-lapse recording at maximum speed.</li>
	<li>Zoom out of the area at the beginning of the time-lapse recording. Zoom in on the AOI.</li>
	<li>Use the &#34;Click &#38; Fire&#34; function of the laser-driver software to apply the damaging irradiation on the cell of interest in the embryo. Use the following parameters: 45% laser transmission (corresponding to a maximum of 40 &#181;W) and 1 ms pulse time duration (step 4). 
	NOTE: Video recording during the laser shooting is recommended.</li>
	<li>Under 688 nm, monitor the ejection of autofluorescent chloroplasts from the cytoplasm.</li>
	<li>If cell contents remain in the cell, use the &#34;Click &#38; Fire&#34; function once more to increase the size of the breach in the cell. Repeat, keeping the number of shots to a minimum until most of the cell contents have been released.</li>
	<li>Stop the time-lapse recording after the embryo has stabilized (i.e., no further intracellular movement can be detected (~1-5 min).</li>
	<li>Update the annotation on the tile scan image, if necessary.</li>
</ol>
6. Monitoring the growth of irradiated embryos
NOTE: Monitoring is carried out over several days.
<ol>
	<li>Determine the survival rate by monitoring the number of embryos that develop after laser ablation and compare them to those that die. 
	NOTE: Some embryos die immediately after ablation for various reasons. A high and rapid mortality rate is usually a sign of inappropriate laser parameters or higher/longer exposure to stress during the experiment or subsequent transport.</li>
	<li>Determine the growth delay by measuring the length of the laser-shot embryos every day and comparing it to intact embryos. 
	NOTE: The growth rate of laser-shot organisms is generally slower than that of untreated organisms. However, some (inappropriate) laser settings can inhibit growth for more than a week, with growth resuming after that.</li>
	<li>Find out the adjacent damage by monitoring the reaction of cells adjacent to the ablated cells. In some cases, post-burst depressurization may cause neighboring cells to burst.</li>
	<li>Check for microbe contaminations. Monitor the growth of microalgae and bacteria in the medium. If an unusual level of microbes are present in the dish, then discard it and repeat the protocol from step 2 or step 3. 
	NOTE: After laser ablation, damaged S. latissima embryos are already highly stressed, and additional external stress can cause increased mortality. Bacterial or viral outbreaks are possible because the treated embryos cannot grow in axenic conditions.</li>
	<li>Check the global development of the shot organisms by studying the phenotype and understanding the role in the development of the targeted region.</li>
</ol>

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The authors have nothing to disclose.

Local cellular laser ablation allows for temporal and spatial ablation with a high level of precision. However, its efficiency can be hampered by the non-accessibility of target cells; for example, all the cells are of a three-dimensional embryo. This protocol was developed on the embryo of the alga Saccharina latissima, which develops a monolayered lamina in which all cells can be easily distinguished and destroyed individually with a laser beam.
Laser power and wavelength</s...

Laser ablation has been used for decades to study embryo development. Irradiating embryo cells with a laser beam makes it possible to monitor the regenerative potential and the modification of the cell lineage during embryogenesis and investigate the impact of targeted ablation on cell division and cell fate. The model organisms used in laser ablation methods are typically animals, such as insects1,2, nematodes3,4, vertebrates5,6, and occasionally plants7,8. In addition, a laser micro-ablation approach was used on the brown alga Fucus in 1994 and 1998 to demonstrate the role of the cell wall in the photopolarization of the early embryo9,10.
Brown algae belong to the group Stramenopiles, diverged at the root of the eukaryotic tree 1.6 billion years ago. As a result, they are phylogenetically independent of other multicellular organisms, such as animals and plants11. Saccharina latissima belongs to the order Laminariales, more commonly known as kelps, and they are among the largest organisms on earth, reaching sizes of over 30 m. Saccharina sp.&#160;is a large seaweed used for many applications such as food and feed, and its polysaccharides are extracted for use in the agricultural, pharmacological and cosmetic industries worldwide12,13. Its cultivation, mainly in Asia and more recently in Europe, requires the preparation of embryos in hatcheries before releasing juveniles in the open sea. Like all kelps, it&#160;has a biphasic life cycle composed of a microscopic gametophytic phase, during which a haploid gametophyte grows and produces gametes for fertilization, and a diploid macroscopic sporophytic phase, where a large planar blade develops from its holdfast attached to the seafloor or rocks. The sporophyte releases haploid spores at maturity, thereby completing the life cycle14,15,16.
S. latissima presents some interesting&#160;morphological features17. Its embryo develops as a monolayered planar sheet15,18,19&#160;before acquiring a multilayered structure coinciding with the emergence of different tissue types. In addition, Laminariales is one of the only taxa of brown algae whose embryos remain attached to their maternal gametophytic tissue (Desmarestiales and Sporochnales do too15). This feature offers the opportunity to study the role of maternal tissue in this developmental process and compare maternal control mechanisms in brown algae with those in animals and plants.
This article presents the first complete protocol for laser ablation in an early kelp embryo. This protocol involving UV ns-pulsed technique results in the specific destruction of individual embryo cells to study their respective roles during embryogenesis. The procedure offers a reliable approach for investigating cell interactions and cell fate during embryogenesis in Laminariales.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>25 mm glass bottom petri dish</td>
			<td>NEST</td>
			<td>801001</td>
			<td></td>
		</tr>
		<tr>
			<td>Autoclaved sea water</td>
			<td>-</td>
			<td></td>
			<td>Collected offshore near the Astan buoy (48&#176;44.934 N 003&#176;57.702 W) close to Roscoff, France, at a depth of 20 m.</td>
		</tr>
		<tr>
			<td>Cell scraper</td>
			<td>MED 2</td>
			<td>83.3951</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell strainer 40 &#181;m</td>
			<td>Corning / Falcon</td>
			<td>352340</td>
			<td></td>
		</tr>
		<tr>
			<td>Culture cabinets</td>
			<td>Snijders Scientific Plant Growth Cabinet ECD01</td>
			<td></td>
			<td>Any other brand is suitable provided that the light intensity, the photoperiod and the temperature can be controlled.</td>
		</tr>
		<tr>
			<td>LSM 880 Zeiss confocal microscope</td>
			<td>Carl Zeiss microscopy, Jena, Germany</td>
			<td></td>
			<td>Ablation and imaging were performed using a 40x/1.2 water objective</td>
		</tr>
		<tr>
			<td>Pellet pestles</td>
			<td>Sigma Aldrich</td>
			<td>Z359947</td>
			<td>Blue polypropylene (autoclavable)</td>
		</tr>
		<tr>
			<td>Provasoli supplement</td>
			<td>-</td>
			<td></td>
			<td>Recipe is available here: http://www.sb-roscoff.fr/sites/www.sb-roscoff.fr/files/documents/station-biologique-roscoff-preparation-du-provasoli-2040.pdf</td>
		</tr>
		<tr>
			<td>Pulsed 355 laser (UGA-42 Caliburn 355/25)</td>
			<td>Rapp OptoElectronic, Wedel, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Paramount</td>
			<td>PDSS 11</td>
			<td></td>
		</tr>
		<tr>
			<td>SysCon software</td>
			<td>Rapp OptoElectronic, Wedel, Germany</td>
			<td></td>
			<td>Laser-driver software</td>
		</tr>
		<tr>
			<td>ZEN software</td>
			<td>Carl Zeiss microscopy, Jena, Germany</td>
			<td></td>
			<td>Imaging software, used together with the SysCon software; Black 2.3 version</td>
		</tr>
	</tbody>
</table>

targeted laser ablation in the embryo of saccharina latissima

In Saccharina latissima, the embryo&#160;develops as a monolayered cell sheet called the lamina or the blade. Each embryo cell is easy to observe, readily distinguishable from its neighbors, and can be individually targeted. For decades, laser ablation has been used to study embryo development. Here, a protocol for cell-specific laser ablation was developed for early embryos of the brown alga S. latissima. The presented work includes: (1) the preparation of Saccharina embryos, with a description of the critical parameters, including culture conditions, (2) the laser ablation settings, and (3) the monitoring of the subsequent growth of the irradiated embryo using time-lapse microscopy. In addition, details are provided on the optimal conditions for transporting the embryos from the imaging platform back to the lab, which can profoundly affect subsequent embryo development. Algae belonging to the order Laminariales display embryogenesis patterns similar to Saccharina; this protocol can thus be easily transferred to other species in this taxon.

Gametophytes of S. latissima were grown, and gametogenesis was induced to produce zygotes and embryos. Twelve days after the induction of gametogenesis, the embryos underwent laser ablation. Here, the experiment aimed to assess the role of specific cells in the overall development of S. latissima embryos. The most apical cell, the most basal cell, and the median cells were targeted. After tile scanning, the entire Petri dish (Figure 2A), an embryo of interest, was identifie...

Watch this Scientific Journal Video about Targeted Laser Ablation in the Embryo of Saccharina latissima at JoVE.com

Targeted Laser Ablation in the Embryo of Saccharina latissima

The destruction of specific cells in the embryo is a powerful tool for studying cellular interactions involved in cell fate. The present protocol describes techniques for the laser ablation of targeted cells in the early embryo of the brown alga Saccharina latissima.

Targeted Laser Ablation in the Embryo of Saccharina latissima

In Saccharina latissima, the embryo&#160;develops as a monolayered cell sheet called the lamina or the blade. Each embryo cell is easy to observe, readily ...

targeted-laser-ablation-in-the-embryo-of-saccharina-latissima

Research

JoVE Journal

Developmental Biology

1.6K Views.  UMR8227, CNRS / Sorbonne University. The destruction of specific cells in the embryo is a powerful tool for studying cellular interactions involved in cell fate. The present protocol describes techniques for the laser ablation of targeted cells in the early embryo of the brown alga Saccharina latissima.

Video: Targeted Laser Ablation in the Embryo of Saccharina latissima

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance to man of conventional invertebrate model systems and the complexity of genomes in vertebrates are critical challenges to modeling human normal and pathological conditions. The chordate Ciona intestinalis is an invertebrate chordate that emerged from a common ancestor with the vertebrates and may represent features at the interface between invertebrates and vertebrates. A common body plan with much simpler cellular and genomic composition should unveil gene regulatory network (GRN) links and functional genomics readouts explaining phenomena in the vertebrate condition. The compact genome of Ciona, a fixed embryonic lineage with few divisions and large cells, combined with versatile community tools foster efficient gene functional analyses in this organism. Here, we present several crucial methods for this promising model organism, which belongs to the closest sister group to vertebrates. We present protocols for transient transgenesis by electroporation, along with microinjection-mediated gene knockdown, which together provide the means to study gene function and genomic regulatory elements. We extend our protocols to provide information on how community databases are utilized for in silico design of gene regulatory or gene functional experiments. An example study demonstrates how novel information can be gained on the interplay, and its quantification, of selected neural factors conserved between Ciona and man. Furthermore, we show examples of differential subcellular localization in embryonic cells, following DNA electroporation in Ciona zygotes. Finally, we discuss the potential of these protocols to be adapted for tissue specific gene interference with emerging gene editing methods. The in vivo approaches in Ciona overcome major shortcomings of classical model organisms in the quest of unraveling conserved mechanisms in the chordate developmental program, relevant to stem cell research, drug discovery, and subsequent clinical application.

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

We present transient transgenesis and gene knockdown in Ciona intestinalis, a chordate sister group to vertebrates, using microinjection and electroporation techniques. Such methods facilitate functional genomics in this simple invertebrate that features rudimentary characteristics of vertebrates, including notochord and head sensory epithelia, and many orthologs of human disease associated genes.

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance ...

Surgical Ablation Assay for Studying Eye Regeneration in Planarians

In the study of adult stem cells and regenerative mechanisms, planarian flatworms are a staple in vivo model system. This is due in large part to their abundant pluripotent stem cell population and ability to regenerate all cell and tissue types after injuries that would be catastrophic for most animals. Recently, planarians have gained popularity as a model for eye regeneration. Their ability to regenerate the entire eye (comprised of two tissue types: pigment cells and photoreceptors) allows for the dissection of the mechanisms regulating visual system regeneration. Eye ablation has several advantages over other techniques (such as decapitation or hole punch) for examining eye-specific pathways and mechanisms, the most important of which is that regeneration is largely restricted to eye tissues alone. The purpose of this video article is to demonstrate how to reliably remove the planarian optic cup without disturbing the brain or surrounding tissues. The handling of worms and maintenance of an established colony is also described. This technique uses a 31 G, 5/16-inch insulin needle to surgically scoop out the optic cup of planarians immobilized on a cold plate. This method encompasses both single and double eye ablation, with eyes regenerating within 1-2 weeks, allowing for a wide range of applications. In particular, this ablation technique can be easily combined with pharmacological and genetic (RNA interference) screens for a better understanding of regenerative mechanisms and their evolution, eye stem cells and their maintenance, and phototaxic behavioral responses and their neurological basis.

This protocol shows how to consistently excise planarian eyes (optic cups) without disturbing surrounding tissues. Using an insulin needle and syringe, either one or both eyes can be ablated to facilitate investigations into the mechanisms regulating eye regeneration, the evolution of visual regeneration, and the neural basis of light-induced behavior.

In the study of adult stem cells and regenerative mechanisms, planarian flatworms are a staple in vivo model system. This is due in large part to their ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. Various propositions claim to have been important contributions to the understanding of the mechanical properties of cells and tissues in their spatiotemporal organization in different developmental and morphogenetic processes. However, due to the lack of reliable and accessible tools to measure material properties and tensional parameters in vivo, validating these hypotheses has been difficult. Here we present methods employing atomic force microscopy (AFM) and particle tracking with the aim of quantifying the mechanical properties of the intact zebrafish embryo yolk cell during epiboly. Epiboly is an early conserved developmental process whose study is facilitated by the transparency of the embryo. These methods are simple to implement, reliable, and widely applicable since they overcome intrusive interventions that could affect tissue mechanics. A simple strategy was applied for the mounting of specimens, AFM recording, and nanoparticle injections and tracking. This approach makes these methods easily adaptable to other developmental times or organisms.

The lack of tools to measure material properties and tensional parameters in vivo prevents validating their roles during development. We employed atomic force microscopy (AFM) and nanoparticle-tracking to quantify mechanical features on the intact zebrafish embryo yolk cell during epiboly. These methods are reliable and widely applicable avoiding intrusive interventions.

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. ...

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout (Oncorhynchus Mykiss) and Brown Trout (Salmo Trutta Morpha fario)

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on fish abdomens to expel the eggs. An alternative method uses the pressure of gas injected into the body cavity, which causes the subsequent release of the eggs. This method is believed to have less negative effects on both the welfare and egg quality of the broodstocks. Herein, we compare the results of air and hand stripping methods with respect to one-year survival and egg quantity and quality in two salmonid fish, rainbow trout (Oncorhynchus mykiss) and brown trout (Salmo trutta morpha fario). Our results indicate that air stripping yielded a better quality of eggs and higher one-year survival rate in rainbow trout. In addition, air stripping resulted in lower mortality rate than the group subjected to hand stripping (25% vs. 35%). The pH and hatching rate of the hand stripped group was lower than those of the air stripped group. In the case of brown trout, the quality of eggs obtained by both hand and air-stripping methods was similar; however, the one-year losses in fish were higher in air stripped group (15% compared to 0% in hand stripped fish). Although the advantages of air stripping method over hand stripping in terms of egg quality might not be observed in all salmonid species, the air-stripping procedure might be a promising option to be adopted in hatcheries as it ensures a high level of reproducibility and efficiency.

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout Oncorhynchus Mykiss and Brown Trout Salmo Trutta Morpha fario

The principal aim of this study is to standardize and test the pneumatic method (air stripping) of collecting eggs in rainbow trout and brown trout. This method allows effective and simple collection of the eggs without the necessity of fish abdomen massage.

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on ...

Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal and postnatal consequences. To ensure the innocuousness of ART applications, studies in animal models are necessary. In addition, as a last step, embryo development studies require evaluation of their capacity to develop full-term healthy offspring. Here, embryo transfer to the uterus is indispensable to perform any ARTs-related experiment.
The rabbit has been used as a model organism to study mammalian reproduction for over a century. In addition to its phylogenetic proximity to the human species and its small size and low maintenance cost, it has important reproductive characteristics such as induced ovulation, a chronology of early embryonic development similar to humans and a short gestation that allow us to study the consequences of ART application easily. Moreover, ARTs (such as intracytoplasmic sperm injection, embryo culture, or cryopreservation) are applied with suitable efficiency in this species.
Using the laparoscopic embryo transfer technique and the cryopreservation protocol presented in this article, we describe 1) how to transfer embryos through an easy, minimally invasive technique and 2) an effective protocol for long-term storage of rabbit embryos to provide time-flexible logistical capacities and the ability to transport the sample. The outcomes obtained after transferring rabbit embryos at different developmental stages indicate that morula is the ideal stage for rabbit embryo recovery and transfer. Thus, an oviductal embryo transfer is required, justifying the surgical procedure. Furthermore, rabbit morulae are successfully vitrified and laparoscopically transferred, proving the effectiveness of the described techniques.

Assisted reproductive techniques (ARTs) are in continuous evaluation to improve outcomes and reduce the associated risks. This manuscript describes a minimally invasive embryo transfer procedure with an efficient cryopreservation protocol that allows the use of rabbits as an ideal animal model of human reproduction.

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal ...

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a general challenge in developmental biology research. In the embryonic heart, this can be particularly difficult as regional differences in the expression of transcriptional regulators, paracrine/juxtacrine signaling cues, and hemodynamic force are all known to influence cardiomyocyte maturation. A simplified method to alter a developing cardiomyocyte's molecular and biomechanical microenvironment would, therefore, serve as a powerful technique to examine how local conditions influence cell fate and function. To address this, we have optimized a method to physically transplant juvenile cardiomyocytes into ectopic locations in the heart or the surrounding embryonic tissue. This allows us to examine how microenvironmental conditions influence cardiomyocyte fate transitions at single cell resolution within the intact embryo. Here, we describe a protocol in which embryonic myocytes can be isolated from a variety of cardiac sub-domains, dissociated, fluorescently labeled, and microinjected into host embryos with high precision. Cells can then be directly analyzed in situ using a variety of imaging and histological techniques. This protocol is a powerful alternative to traditional grafting experiments that can be prohibitively difficult in a moving tissue such as the heart. The general outline of this method can also be adapted to a variety of donor tissues and host environments, and its ease of use, low cost, and speed make it a potentially useful application for a variety of developmental studies.

In this method, embryonic cardiac tissues are surgically microdissected, dissociated, fluorescently labeled, and implanted into host embryonic tissues. This provides a platform for studying the individual or tissue level developmental organization under ectopic hemodynamic conditions, and/or altered paracrine/juxtacrine environments.

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a ...

In Vitro Reconstitution of Spatial Cell Contact Patterns with Isolated Caenorhabditis elegans Embryo Blastomeres and Adhesive Polystyrene Beads

In multicellular systems, individual cells are surrounded by the various physical and chemical cues coming from neighboring cells and the environment. This tissue complexity confounds the identification of causal link between extrinsic cues and cellular dynamics. A synthetically reconstituted multicellular system overcomes this problem by enabling researchers to test for a specific cue while eliminating others. Here, we present a method to reconstitute cell contact patterns with isolated Caenorhabditis elegans blastomere and adhesive polystyrene beads. The procedures involve eggshell removal, blastomere isolation by disrupting cell-cell adhesion, preparation of adhesive polystyrene beads, and reconstitution of cell-cell or cell-bead contact. Finally, we present the application of this method to investigate the orientation of cellular division axes that contributes to the regulation of spatial cellular patterning and cell fate specification in developing embryos. This robust, reproducible, and versatile in vitro method enables the study of direct relationships between spatial cell contact patterns and cellular responses.

In Vitro Reconstitution of Spatial Cell Contact Patterns with Isolated Caenorhabditis elegans Embryo Blastomeres and Adhesive Polystyrene Beads

Tissue complexities of multicellular systems confound the identification of causal relationship between extracellular cues and individual cellular behaviors. Here, we present a method to study the direct link between contact-dependent cues and division axes using C. elegans embryo blastomeres and adhesive polystyrene beads.

In multicellular systems, individual cells are surrounded by the various physical and chemical cues coming from neighboring cells and the environment. ...