The authors acknowledge Dr. Kenzo Ivanovitch for previous work on this method and the group of Dr. Shigenori Nonaka (National Institutes of Natural Sciences, Japan) for providing the initial expertise on embryo mounting. This study was supported by Grant PGC2018-096486-B-I00 from the Spanish Ministerio de Ciencia e Innovaci&#243;n and Grant H2020-MSCA-ITN-2016-722427 from the EU Horizon 2020 program to MT and Grant 1380918 from the FEDER Andaluc&#237;a 2014-2020 Operating Program to JND. MS was supported by a La Caixa Foundation PhD fellowship (LCF/BQ/DE18/11670014) and The Company of Biologists travelling fellowship (DEVTF181145). The CNIC is supported by the Spanish Ministry of Science and the ProCNIC Foundation.

All animal procedures were approved by the CNIC Animal Experimentation Ethics Committee, by the Community of Madrid (Reference PROEX 220/15) and conformed to EU Directive 2010/63EU and Recommendation 2007/526/EC regarding the protection of animals used for experimental and other scientific purposes, enforced in Spanish law under Real Decreto 1201/2005.
This protocol includes the use of two males from the fluorescent transgenic mouse line reporting NOTCH activity Tg(CBF:H2BVenus,+)18. See the Table of Materials for details about materials, animals, and equipment used in this protocol.
1. Tools and holder customization
<ol>
	<li>Cut 1 cm long tungsten wire pieces and sharpen them to 0.02-0.05 mm diameter using any of the simple methods available, for example, submerging the tip of the wire in a saturated sodium nitrite solution19.</li>
	<li>Prepare elbowed glass capillaries.
	<ol>
		<li>Place the midpoint of a standard 1.0 mm glass capillary over a Bunsen lighter for 3-6 s, pulling slowly from both ends until the capillary turns thin and flexible. At that point, break the capillary in half and heat it briefly to produce a 90&#176; bend 2 cm from the tip.</li>
	</ol>
	</li>
	<li>Saw a piece of polymethyl methacrylate measuring approximately 7 mm long, 4 mm wide, and 2 mm thick (Figure 3D). 
	NOTE: Polymethyl methacrylate sheets can be either obtained from recycled laboratory equipment or ordered new.</li>
	<li>Hold the methacrylate piece using a bench vise. Next, drill custom-sized holes transversally through the whole piece (Figure 3D). 
	NOTE: Typically, E6.5 to E7.5 mouse embryos require using 0.2-0.5 mm diameter drills.
	<ol>
		<li>Rotate the drill slowly while applying low but constant pressure. If a drill breaks inside the holder, discard the piece, as the metal cannot be taken out.</li>
	</ol>
	</li>
	<li>Use a fine file to smooth the holder&#39;s edges and adjust its size. Additionally, create a mild slope on the edges of the holder to facilitate embryo positioning (Figure 3D).</li>
	<li>Place the finished holder in a dish with distilled water to rinse it.</li>
	<li>Check the holder under the stereomicroscope to see if the holes contain drilling dust. If dust is present, use tungsten needles to remove it.</li>
	<li>To sterilize the holder, place it in a conical tube full of distilled water and sonicate it for 20 min with a minimum 20 W power output.</li>
</ol>
2. Media preparation
<ol>
	<li>To heat-inactivate the complement system, leave two 500 &#956;L aliquots of commercial or homemade20 rat serum at 56 &#176;C for 30 min. 
	NOTE: For a detailed protocol on how to prepare rat serum, see21.</li>
	<li>In a cell culture hood, prepare two aliquots of the medium for culturing the embryos in the microscope. For each, mix 490 &#956;L of DMEM for live-cell imaging, 500 &#956;L of inactivated rat serum, and 10 &#956;L&#160;of penicillin-streptomycin to obtain a final concentration of 50 &#181;g/mL penicillin and 50 &#181;g/mL streptomycin22. 
	NOTE: The volume of culture medium depends on the time and number of embryos to be cultured. As a rule of thumb, optimal growth requires a minimum of 200 &#956;L per embryo per day in E7.0 embryos.</li>
	<li>Using a 2 mL syringe, filter the medium through a 0.22 &#181;m pore size membrane filter into a new tube and place it lid-open in a cell culture incubator at 37 &#176;C and 7% CO2 for 1 h before embryo culture23.</li>
	<li>Prepare the dissection medium by adding to a 500 mL of DMEM supplemented with L-glutamine bottle the following: 50 mL of fetal bovine serum, 10 mL of 25 mM HEPES-NaOH (pH 7.2), and 10 mL of penicillin and streptomycin (50 mg/mL).</li>
	<li>Next, separate it into 50 mL aliquots, place three aliquots in a 37 &#176;C bath and store the rest at 4 &#176;C for up to 3 months.</li>
</ol>
3. Embryo dissection
<ol>
	<li>Euthanize an E6.75-E7.5 pregnant mouse.</li>
	<li>Extract the uterus and place it on a dry and clean paper wipe. Then, cut the uterus, inserting the tip of fine scissors from the mesometrial side and sliding the blade along to expose the deciduae and transfer them to the dissection medium. For a detailed protocol, see24.</li>
	<li>Dissect the embryos, keeping the ectoplacental cone intact (Figure 3A,B). For a detailed dissection method, refer to25.</li>
	<li>Then, peel off the Reichert&#39;s membrane, leaving part of it on the extraembryonic region (Figure 3B). 
	NOTE: Fine forceps are essential for this step. Inexperienced users can dissect embryos on 2% agarose gel-coated dishes to avoid bending the forceps&#39; tips against the dish surface (Figure 3C).</li>
	<li>To keep the dissection medium warm, replace it every 10 min. Moreover, dissect deciduae in groups of 3-4 while keeping the rest in dissection medium placed in a 37 &#176;C bath. Immediately place the dissected embryos in culture medium.</li>
	<li>Select the embryos with the desired fluorescence features using a fluorescence stereomicroscope and place them in the tube containing culture medium in the cell culture incubator.</li>
	<li>If the embryos are at E6.5 to E7.0 stages, close the lid and leave the embryos to recover for 2 h before imaging. 
	&#8203;NOTE: E6.5 to E7.0 embryos are sensitive. They can also be placed directly under the microscope but preculturing allows for selection of the ones that best recover from dissection, increasing the chance of success.</li>
</ol>
4. Microscope preparation
<ol>
	<li>Turn on all microscope components, including lasers, computer, and software. Turn on the temperature controllers and set to 37 &#176;C 3 h in advance to ensure equilibration.</li>
	<li>If using an immersion detection objective, clean it using a residue-free disposable wipe. Dip the objective&#39;s tip in double-distilled water using a 60 mm dish and leave it until the acquisition starts.</li>
	<li>Turn on the CO2 controller and set it to 7%.</li>
	<li>Place a drop of high-vacuum silicon grease on a 35 mm dish with glass coverslip bottom (14 mm diameter), place the holder on top of the drop, and press gently to immobilize it.</li>
	<li>Under a stereomicroscope, add culture medium to fill the glass-bottom portion of the dish. While doing so, aim the stream to the holder&#39;s holes to prevent formation of air bubbles in them.</li>
	<li>If the bubbles remain, use the glass capillary prepared in step 1.2, connected to a silicone tube with a mouthpiece, to suck the bubbles out gently. 
	&#8203;NOTE: The elbowed end of the capillary will facilitate access through the holes.</li>
</ol>
5. Embryo mounting
<ol>
	<li>Transfer 2-3 embryos that were left to recover in the incubator to the dish with the holder. Leave the rest in the incubator as a backup.</li>
	<li>Set the embryos in the holes by using a pair of forceps and a tapered tungsten needle. Use the needle to hook the embryo by the ectoplacental cone and insert it into a size-matching hole. To immobilize the embryo, rotate the cone by 90-120&#176; so that the Reichter&#39;s membrane adheres to the hole walls (Figure 3C). For an alternative description, see26.</li>
	<li>Carefully move the dish containing the embryos and the holder to the microscope plate (Figure 3F,G). 
	NOTE: If not moved steadily, the embryo can move out of the holder.</li>
	<li>Lower the objective until a liquid meniscus is formed at the medium-objective interface.</li>
	<li>Locate the embryo using transmitted light under the microscope binocular and place the embryo in focus.</li>
	<li>Mount the incubation chamber and seal it around the objective using tape (Figure 3F).</li>
	<li>Using a Live Capture at the microscope control software, adjust the Laser Output and Gain levels. 
	NOTE: For two-photon acquisition of green fluorescent protein (GFP) and Tdtomato reporters, 30% output laser power at 980 nm and 70% gain on non-descanned detectors were used in this protocol (Figure 3G).</li>
	<li>To avoid evaporation, cover the medium with paraffin oil by dripping it slowly on the objective. 
	&#8203;NOTE: This is the last point where additional embryos can be mounted in the holder. Once covered with paraffin oil, one would have to clean the holder and replace the medium to do so. For alternative mounting methods see27.</li>
</ol>
6. Image acquisition
<ol>
	<li>Set up time-lapse recording. Set a 5-10 min time interval with 3-5 &#181;m z-space between stacks. Leave a blank space on top of the z-stack to anticipate embryo growth, a minimum of 50 &#181;m, adding 30 &#181;m for every hour the acquisition will not be supervised.</li>
	<li>If available, enable the Auto-Save option to allow supervision of the acquisition from a personal computer to allow the adjustment of the z-stack size, if needed, to fit the area of interest within focus.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64273/64273fig03.jpg" /> 
Figure 3: Live imaging tools and setup. (A) Dissecting mouse embryos with an agarose-coated dish under the stereomicroscope. (B) Diagram of the steps to dissect mouse embryos for live imaging. (C) Embryo positioning in the holder. (D) Embryo holder design and features. (F) Incubator chamber around the immersion objective. (G) Diagram of final setup for time-lapse acquisition. Scale bar = 500 &#181;m (A). <a href="https://www.jove.com/files/ftp_upload/64273/64273fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors have no conflicts of interest to disclose.

Early heart progenitors organize in a primitive heart tube that starts beating while it is still forming. Understanding how this process takes place is key to pinpoint the wide spectrum of congenital heart defects to specific morphogenetic events. For that, live imaging offers an opportunity to study normal and defective embryonic development with increased temporal resolution. This is especially useful to study early cardiac progenitor cells as they transition quickly through multiple differentiation and migration behav...

The heart forms early during embryogenesis to start pumping nutrients to the whole embryo, while it continues developing1. In mouse embryos, one and a half days after the initiation of gastrulation, a rudimentary heart organ assembles at the anterior pole2,3. By Early Streak (ES) stage, cardiac progenitors in the epiblast ingress through the primitive streak to the nascent mesodermal layer4,5,6 and start migrating to the anterior pole, where they differentiate to form the primitive heart tube. Throughout this process, early heart progenitors undergo cell rearrangements, shape transformations, and differentiation, in addition to migration7 (Figure 1).
Early cardiac progenitors have attracted researchers for nearly a century due to their remarkable ability to differentiate and build a functional organ simultaneously. Over the last two decades, clonal analysis and conditional knockout models have shown that early heart development implicates distinct cell sources in a highly dynamic process8,9,10. However, the 3D structure of the primitive heart tube and the dynamic nature of its morphogenesis make it challenging to study (Figure 1), and we are far from understanding its full complexity11.
To study these dynamic cellular processes, live imaging methods now offer an unprecedented detail7,12,13,14. In the mouse model, live approaches have been key to interrogating developmental topics that are difficult to address by static analysis7,13,15. While long-term ex vivo culture and robust microscope setups are advancing fast16,17, few researchers have the expertise to successfully image live embryos. Although paper-based publications provide enough technical details to reproduce live imaging experiments, some skills and tricks are hard to grasp without visual examples or peer-to-peer assistance. To accelerate this learning process and spread the use of live imaging among laboratories, we assembled a video protocol (Figure 2) that gathers the necessary skills to perform live imaging on gastrulating mouse embryos.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64273/64273fig01.jpg" /> 
Figure 1: Early differentiation of cardiac progenitor cells in the mouse embryo from the onset of gastrulation to the stage preceding primitive heart tube formation. Cardiac progenitor cells ingress the mesoderm soon after the start of gastrulation, migrating to the opposite side of the embryo. Morphological and embryonic day (E) stage are written on top of the diagrams. Dashed arrows depict the migration trajectory of primitive heart tube progenitors during gastrulation. This figure was adapted from11. Abbreviations: ES = Early Streak; MS = Middle Streak; EHF = Early Head Fold. <a href="https://www.jove.com/files/ftp_upload/64273/64273fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64273/64273fig02.jpg" /> 
Figure 2: Workflow diagram for live imaging of early heart progenitors. <a href="https://www.jove.com/files/ftp_upload/64273/64273fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>#55 Forceps</td>
			<td>Dumont&#160;</td>
			<td>11295-51</td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm Dish with glass coverslip bottom 14 mm Diameter</td>
			<td>Mattek</td>
			<td>P35G-1.5-14-C</td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm vise table&#160;</td>
			<td>Grandado</td>
			<td>SKU 8798771617573</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL tubes</td>
			<td>BD Falcon</td>
			<td>352070</td>
			<td></td>
		</tr>
		<tr>
			<td>Distilled water</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM - Dulbecco&#39;s Modified Eagle Medium</td>
			<td>Gibco</td>
			<td>11966025</td>
			<td>&#160;with L-Glutamine, without Glucose, without Na Pyruvate</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Invitrogen</td>
			<td>10438-026</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescent reporter transgenic mice (Tg(CBF:H2BVenus,+)&#160;</td>
			<td>JAX</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorobrite DMEM</td>
			<td>ThermoFisher</td>
			<td>A1896701&#160;</td>
			<td>DMEM for live-cell imaging</td>
		</tr>
		<tr>
			<td>High-vacuum silicone grease</td>
			<td>Dow Corning</td>
			<td>Z273554-1EA</td>
			<td></td>
		</tr>
		<tr>
			<td>Holder for wires</td>
			<td>Perlen Pressen</td>
			<td>pwb1</td>
			<td></td>
		</tr>
		<tr>
			<td>LSM 780 Upright microscope</td>
			<td>Zeiss&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MaiTai Deepsee far red pulsed-laser tuned at 980 nm</td>
			<td>Spectra-Physics</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Non Descanned Detectors equipped with the filter sets 
			cyan-yellow (BP450-500/BP520-560), green-red (BP500-520/BP570-610) and yellow-red (BP520-560/BP645-710)</td>
			<td>Zeiss&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Obj: 20x water dipping 1.0 NA, long working distance</td>
			<td>Zeiss&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>P1000 and P200 pipettes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Paraffin Oil</td>
			<td>Nidacon</td>
			<td>VNI0049</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-streptomycin</td>
			<td>Invitrogen</td>
			<td>15070-063</td>
			<td>(the final concentration should be 50 &#956;g/mL penicillin and 50 &#956;g/mL streptomycin)</td>
		</tr>
		<tr>
			<td>Petri dishes 35 mm x 10 mm</td>
			<td>BD Falcon</td>
			<td>351008</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette tips</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Polymethyl methacrylate&#160;</td>
			<td></td>
			<td></td>
			<td>Reused from old laboratory equipment</td>
		</tr>
		<tr>
			<td>Rat Serum culture embryo, male rats SPRAGUE DAWLEY RjHan SD</td>
			<td>Janvier Labs</td>
			<td>9979</td>
			<td></td>
		</tr>
		<tr>
			<td>Set of 160 mm fines</td>
			<td>RS PRO</td>
			<td>541-6933</td>
			<td></td>
		</tr>
		<tr>
			<td>Standard 1.0 mm glass capillaries</td>
			<td>Anima Lab</td>
			<td>&#160;1B100F-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile 0.22 &#956;m syringe filter</td>
			<td>Corning</td>
			<td>431218</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile 5 mL syringe</td>
			<td>Fisher Scientific</td>
			<td>15809152</td>
			<td></td>
		</tr>
		<tr>
			<td>Tungsten needles</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasonic homogeniser (sonicator)</td>
			<td>Bandelin</td>
			<td>BASO_17021</td>
			<td></td>
		</tr>
	</tbody>
</table>

live imaging of early cardiac progenitors in the mouse embryo

The first steps of heart development imply drastic changes in cell behavior and differentiation. While analysis of fixed embryos allows studying in detail specific developmental stages in a still snapshot, live imaging captures dynamic morphogenetic events, such as cell migration, shape changes, and differentiation, by imaging the embryo as it develops. This complements fixed analysis and expands the understanding of how organs develop during embryogenesis. Despite its advantages, live imaging is rarely used in mouse models because of its technical challenges. Early mouse embryos are sensitive when cultured ex vivo and require efficient handling. To facilitate a broader use of live imaging in mouse developmental research, this paper presents a detailed protocol for two-photon live microscopy that allows long-term acquisition in mouse embryos. In addition to the protocol, tips are provided on embryo handling and culture optimization. This will help understand key events in early mouse organogenesis, enhancing the understanding of cardiovascular progenitor biology.

We used the protocol to visualize NOTCH signaling activation in early cardiac progenitors about to differentiate to endothelial cells during primitive heart tube morphogenesis. For that, we crossed wild-type C57BL/6-N mice with Tg(CBF:H2BVenus,+) mice18 to obtain embryos reporting NOTCH activity through yellow fluorescent protein Venus. At E7.5, Venus fluorescence is present throughout the neural ectoderm, with a few positive nuclei at the splanchnic and extraembryonic mesoderm. After 4 h, endothe...

Watch this Scientific Journal Video about Live Imaging of Early Cardiac Progenitors in the Mouse Embryo at JoVE.com

Live Imaging of Early Cardiac Progenitors in the Mouse Embryo

We present a detailed protocol for mouse embryo culture and imaging that enables 3D + time imaging of cardiac progenitor cells. This video-toolkit addresses the key skills required for successful live imaging otherwise hard to acquire from text-only publications.

The first steps of heart development imply drastic changes in cell behavior and differentiation. While analysis of fixed embryos allows studying in detail ...

live-imaging-of-early-cardiac-progenitors-in-the-mouse-embryo

Research

JoVE Journal

Developmental Biology

1.3K Views.  Centro Nacional de Investigaciones Cardiovasculares (CNIC). We present a detailed protocol for mouse embryo culture and imaging that enables 3D + time imaging of cardiac progenitor cells. This video-toolkit addresses the key skills required for successful live imaging otherwise hard to acquire from text-only publications.

Video: Live Imaging of Early Cardiac Progenitors in the Mouse Embryo

Protocols for Obtaining Zygotic and Somatic Embryos for Studying the Regulation of Early Embryo Development in the Model Legume Medicago truncatula

Early embryogenesis starting from a single cell zygote goes through rapid cell division and morphogenesis, and is morphologically characterized by pre-globular, globular, heart, torpedo and cotyledon stages. This progressive development is under the tight regulation of a complex molecular network. Harvesting sufficient early embryos at a similar stage of development is essential for investigating the cellular and molecular regulation of early embryogenesis. This is not straightforward since early embryogenesis undergoes rapid morphogenesis in a short while e.g. 8 days for Medicago truncatula to reach the early cotyledon stage. Here, we address the issue by two approaches. The first one establishes a linkage between embryo development and pod morphology in helping indicate the stage of the zygotic embryo. This is particularly based on the number of pod spirals and development of the spines. An alternative way to complement the in vivo studies is via culturing leaf explants to produce somatic embryos. The medium includes an unusual hormone combination - an auxin (1-naphthaleneacetic acid), a cytokinin (6-benzylaminopurine), abscisic acid and gibberellic acid. The different stages can be discerned growing out of the callus without dissection.

Protocols for Obtaining Zygotic and Somatic Embryos for Studying the Regulation of Early Embryo Development in the Model Legume Medicago truncatula

The goal is to illustrate that the model legume Medicago truncatula can be readily utilized to investigate the regulation of early plant embryogenesis to complement the non-legume Arabidopsis model. Pod morphology is linked to zygotic embryogenesis stages and a protocol to collect embryos using tissue culture is also provided.

Early embryogenesis starting from a single cell zygote goes through rapid cell division and morphogenesis, and is morphologically characterized by ...

Characterization of Thymic Settling Progenitors in the Mouse Embryo Using In Vivo and In Vitro Assays

Characterizing thymic settling progenitors is important to understand the pre-thymic stages of T cell development, essential to devise strategies for T cell replacement in lymphopenic patients. We studied thymic settling progenitors from murine embryonic day 13 and 18 thymi by two complementary in vitro and in vivo techniques, both based on the &#8220;hanging drop&#8221; method. This method allowed colonizing irradiated fetal thymic lobes with E13 and/or E18 thymic progenitors distinguished by CD45 allotypic markers and thus following their progeny. Colonization with mixed populations allows analyzing cell autonomous differences in biologic properties of the progenitors while colonization with either population removes possible competitive selective pressures. The colonized thymic lobes can also be grafted in immunodeficient male recipient mice allowing the analysis of the mature T cell progeny in vivo, such as population dynamics of the peripheral immune system and colonization of different tissues and organs. Fetal thymic organ cultures revealed that E13 progenitors developed rapidly into all mature CD3+ cells and gave rise to the canonical &#947;&#948; T cell subset, known as dendritic epithelial T cells. In comparison, E18 progenitors have a delayed differentiation and were unable to generate dendritic epithelial T cells. The monitoring of peripheral blood of thymus-grafted CD3-/- mice further showed that E18 thymic settling progenitors generate, with time, larger numbers of mature T cells than their E13 counterparts, a feature that could not be appreciated in the short term fetal thymic organ cultures.

Characterization of Thymic Settling Progenitors in the Mouse Embryo Using In Vivo and In Vitro Assays

This article describes in vivo and in vitro methodology to characterize the thymic settling progenitors by the analysis of the kinetics of generation, phenotype and numbers of their T cell progeny.

Characterizing thymic settling progenitors is important to understand the pre-thymic stages of T cell development, essential to devise strategies for T ...

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 ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Identification Of Erythromyeloid Progenitors And Their Progeny In The Mouse Embryo By Flow Cytometry

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where they act as front line sentinels of infection and tissue damage. While other immune cells are continuously renewed from hematopoietic stem and progenitor cells (HSPC) located in the bone marrow, a lineage of macrophages, known as resident macrophages, have been shown to be self-maintained in tissues without input from bone marrow HSPCs. This lineage is exemplified by microglia in the brain, Kupffer cells in the liver and Langerhans cells in the epidermis among others. The intestinal and colon lamina propria are the only adult tissues devoid of HSPC-independent resident macrophages. Recent investigations have identified that resident macrophages originate from the extra-embryonic yolk sac hematopoiesis from progenitor(s) distinct from fetal hematopoietic stem cells (HSC). Among yolk sac definitive hematopoiesis, erythromyeloid progenitors (EMP) give rise both to erythroid and myeloid cells, in particular resident macrophages. EMP are only generated within the yolk sac between E8.5 and E10.5 days of development and they migrate to the fetal liver as early as circulation is connected, where they expand and differentiate until at least E16.5. Their progeny includes erythrocytes, macrophages, neutrophils and mast cells but only EMP-derived macrophages persist until adulthood in tissues. The transient nature of EMP emergence and the temporal overlap with HSC generation renders the analysis of these progenitors difficult. We have established a tamoxifen-inducible fate mapping protocol based on expression of the macrophage cytokine receptor Csf1r promoter to characterize EMP and EMP-derived cells in vivo by flow cytometry.

While infiltrating macrophages are continuously recruited to adult tissues from circulating precursors, resident macrophages seed their tissue during development, where they are maintained without further input from progenitors. The progenitors for resident macrophages were recently identified. Here, we present methods for the genetic fate mapping of the resident macrophage progenitors.

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where ...

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

Bioengineering of Humanized Bone Marrow Microenvironments in Mouse and Their Visualization by Live Imaging

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth factors, and extracellular matrix. The ability of HSCs to remain quiescent, self-renew or differentiate, and acquire mutations and become malignant depends upon the complex interactions they establish with different stromal components. To observe the crosstalk between human HSCs and the human BM niche in physiological and pathological conditions, we designed a protocol to ectopically model and image a humanized BM niche in immunodeficient mice. We show that the use of different cellular components allows for the formation of humanized structures and the opportunity to sustain long-term human hematopoietic engraftment. Using two-photon microscopy, we can live-image these structures in situ at the single-cell resolution, providing a powerful new tool for the functional characterization of the human BM microenvironment and its role in regulating normal and malignant hematopoiesis.

A method to create and live-image different humanized bone-marrow niches in mice is presented. Based on the supportive niche created by human mesenchymal cells, the addition of human endothelial cells induces the formation of human vessels, while the addition of rhBMP-2 induces the formation of human-mouse chimeric mature bone tissue.

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth ...

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; for example, how the zygote polarizes to establish the body axis and how the embryo specifies various cell fates during organ formation. This manuscript describes an in vitro ovule culture method to perform live-cell imaging of developing zygotes and embryos of Arabidopsis thaliana. The optimized cultivation medium allows zygotes or early embryos to grow into fertile plants. By combining it with a poly(dimethylsiloxane) (PDMS) micropillar array device, the ovule is held in the liquid medium in the same position. This fixation is crucial to observe the same ovule under a microscope for several days from the zygotic division to the late embryo stage. The resulting live-cell imaging can be used to monitor the real-time dynamics of zygote polarization, such as nuclear migration and cytoskeleton rearrangement, and also the cell division timing and cell fate specification during embryo patterning. Furthermore, this ovule cultivation system can be combined with inhibitor treatments to analyze the effects of various factors on embryo development, and with optical manipulations such as laser disruption to examine the role of cell-cell communication.

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

This manuscript describes an in vitro ovule cultivation method that enables live-cell imaging of Arabidopsis zygotes and embryos. This method is utilized to visualize the intracellular dynamics during zygote polarization and the cell fate specification in developing embryos.

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

A Novel Use of Three-dimensional High-frequency Ultrasonography for Early Pregnancy Characterization in the Mouse

High-frequency ultrasonography (HFUS) is a common method to non-invasively monitor the real-time development of the human fetus in utero. The mouse is routinely used as an in vivo model to study embryo implantation and pregnancy progression. Unfortunately, such murine studies require pregnancy interruption to enable follow-up phenotypic analysis. To address this issue, we used three-dimensional (3-D) reconstruction of HFUS imaging data for early detection and characterization of murine embryo implantation sites and their individual developmental progression in utero. Combining HFUS imaging with 3-D reconstruction and modeling, we were able to accurately quantify embryo implantation site number as well as monitor developmental progression in pregnant C57BL6J/129S mice from 5.5 days post coitus (d.p.c.) through to 9.5 d.p.c. with the use of a transducer. Measurements included: number, location, and volume of implantation sites as well as inter-implantation site spacing; embryo viability was assessed by cardiac activity monitoring. In the immediate post-implantation period (5.5 to 8.5 d.p.c.), 3-D reconstruction of the gravid uterus in both mesh and solid overlay format enabled visual representation of the developing pregnancies within each uterine horn. As genetically engineered mice continue to be used to characterize female reproductive phenotypes derived from uterine dysfunction, this method offers a new approach to detect, quantify, and characterize early implantation events in vivo. This novel use of 3-D HFUS imaging demonstrates the ability to successfully detect, visualize, and characterize embryo-implantation sites during early murine pregnancy in a non-invasive manner. The technology offers a significant improvement over current methods, which rely on the interruption of pregnancies for gross tissue and histopathologic characterization. Here we use a video and text format to describe how to successfully perform ultrasounds of early murine pregnancy to generate reliable and reproducible data with reconstruction of the uterine form in mesh and solid 3-D images.

Mice are widely used to study gestational biology. However, pregnancy termination is required for such studies which precludes longitudinal investigations and necessitates the use of large numbers of animals. Therefore, we describe a non-invasive technique of high-frequency ultrasonography for early detection and monitoring of post-implantation events in the pregnant mouse.

High-frequency ultrasonography (HFUS) is a common method to non-invasively monitor the real-time development of the human fetus in utero. The mouse is ...