We thank James Priess and Bruce Bowerman for advice and providing C. elegans strains, Don Moerman, Kota Mizumoto, and Life Sciences Institute Imaging Core Facility for sharing equipment and reagents, Aoi Hiroyasu, Lisa Fernando, Min Jee Kim for the maintenance of C. elegans and critical reading of our manuscript. Our work is supported by the Natural Sciences and Engineering Research Council of Canada (NSERC), (RGPIN-2019-04442).

1. Preparation of adhesive polystyrene bead
NOTE: This protocol does not require aseptic technique.
<ol>
	<li>Weigh 10 mg of carboxylate modified polystyrene beads in a 1.5 mL microcentrifuge tube.</li>
	<li>To wash the beads, add 1 mL of 2-(N-morpholino)ethanesulfonic acid (MES) buffer into the tube. Since MES buffer does not contain phosphate and acetate, which can reduce the reactivity of carbodiimide, it is suitable to use in protein coupling reaction. Vortex the tube to mix the beads.</li>
	<li>Spin the tube for 60 s at 2,000 x g via a benchtop centrifuge.</li>
	<li>Discard the supernatant by carefully pipetting out the buffer.</li>
	<li>Wash the beads again with 1 mL of MES buffer by following steps 1.2-1.4.</li>
	<li>Add 1 mL of MES buffer containing 10 mg of EDAC into the tube to activate the surface carboxyl groups. Vortex the tube to mix the beads.</li>
	<li>Rotate and incubate the tube for 15 min at room temperature.</li>
	<li>Spin down the beads for 60 s at 2,000 x g.</li>
	<li>Discard the supernatant by carefully pipetting out the buffer.</li>
	<li>To wash the beads, add 1 mL of phosphate buffered saline (PBS) into the tube. Vortex the tube to mix the beads.</li>
	<li>Spin down the tube for 60 s at 2,000 x g.</li>
	<li>Discard the supernatant by carefully pipetting out the buffer.</li>
	<li>Wash the beads again with 1 mL of PBS by following steps 1.10-1.12.</li>
	<li>The final concentration of Rhodamine used will depend on the strain being imaged. Prepare 1 mL of 1-, 10-, 100-, and 1000-fold dilution series of Rhodamine Red-X from the 0.65 mM Rhodamine Red-X stock solution.</li>
	<li>Pipette 20 &#181;L of beads into each serial dilution tube.</li>
	<li>Rotate and incubate the tube for 5 min at room temperature.</li>
	<li>Wash the beads twice with 1 mL of PBS by repeating steps 1.10-1.12.</li>
	<li>Add 1 mL of PBS into the tube and store it at 4 &#176;C for up to 6 weeks. Check the fluorescence intensity of the beads under a microscope used for live-imaging. The appropriate concentration of the Rhodamine Red-X succinimidyl ester is dependent on the imaging conditions (Figure 1). 
	NOTE: Beads without Rhodamine Red-X treatment do not adhere to cell. Rhodamine Red-X serves as fluorescence marker as well as adhesive molecule. The electro static interaction between positively charged Rhodamine Red-X and negatively charged plasma membrane is a putative cause of adhesion.</li>
</ol>
2. Assembly of mouth pipette
<ol>
	<li>Cut wide (6.35 mm inner diameter) and narrow (3.175 mm inner diameter) Tygon tubes to about 25 cm and 40 cm in length, respectively (Figure 2).</li>
	<li>Connect the Tygon tubes with a polytetrafluoroethylene (PTFE) filter (0.2 &#181;m pore size) (Figure 2).</li>
	<li>Disassemble a commercially available aspirator tube and attach its capillary holder and mouthpiece to the end of the narrow and wide Tygon tubes, respectively (Figure 2). 
	NOTE: PTFE filter was used to prevent the inhalation of fumes of hypochlorite solution via mouth pipette.</li>
</ol>
3. Isolation of embryo blastomere
NOTE: Wear gloves and lab coat to avoid cut and contact with the bleaching solution.
<ol>
	<li>Hold each end of a microcapillary (capacity; 10 &#181;L) with right and left hand.</li>
	<li>Pull the microcapillary towards both ends to apply tension and bring the center of the capillary over a burner to make two hand-pulled capillaries (Figure 3A).</li>
	<li>Trim the tips of the hand-pulled capillaries with forceps under the dissecting microscope and attach the pulled capillary into a mouth pipetting apparatus (Figure 2). Prepare two types of pipettes. The tip opening sizes for the pipettes should be approximately 2x and 1x the short axis length of C. elegans embryos (30 &#181;m) for the embryo transfer and eggshell removal, respectively Figure 3B-D).</li>
	<li>Pipette 45 &#956;L of egg salt solution onto a well of a multiwell slide (Figure 4A; bottom).</li>
	<li>Place 5-10 adult C. elegans onto a well containing egg salt solution.</li>
	<li>To obtain early C. elegans embryos, cut adults into pieces by positioning two needles to the right and left of C. elegans body and sliding the needles past each other (Figure 4A; upper schematics).</li>
	<li>Pipette 45 &#956;L of hypochlorite solution onto a well next to the well containing egg salt solution (Figure 4B).</li>
	<li>Pipette 45 &#956;L of Shelton&#39;s growth medium onto the subsequent three wells next to the well containing hypochlorite solution (Figure 4B).</li>
	<li>Transfer 1-cell stage and early 2-cell stage embryos into the hypochlorite solution by mouth pipetting with the hand-drawn capillary for embryo transfer (Figure 4B).</li>
	<li>Wait for 40&#8211;55 s.</li>
	<li>Wash the embryos by transferring the embryos from hypochlorite solution into Shelton&#39;s growth medium by mouth pipetting with the hand-drawn capillary for embryo transfer (Figure 4B).</li>
	<li>Wash the embryos again by transferring the embryos into a new well of Shelton&#39;s growth medium by mouth pipetting with the hand-drawn capillary for embryo transfer (Figure 4B).</li>
	<li>Transfer the washed embryos into a new well of Shelton&#39;s growth medium by mouth pipetting with the hand-drawn capillary for embryo transfer. Using the hand-drawn capillary for eggshell removal, carefully repeat the pipetting (Figure 4C; middle schematics). If the eggshell is successfully removed, the embryonic cells will become more spherical (Figure 4C; right).</li>
	<li>Separate the 2-cell stage embryonic blastomeres by gently and continuously pipetting with the hand-drawn capillary for eggshell removal (Figure 4D).</li>
</ol>
4. Reconstitution of contact patterns with blastomere and bead
NOTE: Work under the imaging microscope to avoid dissociation of a cell from a bead and to facilitate the timely image acquisition.
<ol>
	<li>To observe using an inverted microscope, prepare an imaging chamber as in Figure 5.
	<ol>
		<li>Place a coverslip onto a coverslip holder (Figure 5A).</li>
		<li>Tape the edges of the coverslip to stabilize it. The side with tape is the &#39;back&#39; side (Figure 5A,B).</li>
		<li>Flip the coverslip holder over to the &#39;front&#39; side and draw a circle on the coverslip with a hydrophobic pen (Figure 5C). 
		NOTE: Any glass-bottom dish also works, provided the embryos are manipulatable by the mouth pipette.</li>
	</ol>
	</li>
	<li>Add Shelton&#39;s growth medium within the circle drawn by the hydrophobic marker (Figure 5D).</li>
	<li>Transfer the isolated blastomere to the imaging chamber.</li>
	<li>Dispense a small volume of the chemically functionalized beads using the hand-drawn capillary for embryo transfer.</li>
	<li>Control the position of the polystyrene beads by blowing into the hand-drawn capillary until the bead attaches to the isolated blastomere.</li>
	<li>Mount a coverslip to avoid evaporation of medium (Figure 5E). Perform live imaging.</li>
</ol>
5. Preparation of important reagents
<ol>
	<li>Egg Salt Solution (10 mL): Combine 235 &#181;L of 5 M NaCl and 240 &#181;L of 2 M KCl with 9525 &#181;L of dH2O.</li>
	<li>Hypochlorite Solution (10 mL): Combine 7.5 mL of Clorox (containing approximately 7.5% sodium hypochlorite) with 2.5 mL of 10 N NaOH (final concentration of sodium hypochlorite is approximately 5.625%). 
	NOTE: Although many published methods have used chitinase to digest chitinous eggshells, we have adopted a method using hypochlorite solution11. By avoiding the batch-to-batch variations of chitinase activities, we believe this method is more reproducible and cost-effective approach.</li>
	<li>0.81 mM Inulin Solution (40 mL)
	<ol>
		<li>Add 0.2 g of Inulin to 40 mL of dH2O.</li>
		<li>Autoclave to dissolve. 
		NOTE: Keeps for 1 month at 4 &#176;C.</li>
	</ol>
	</li>
	<li>0.5 M MES Buffer (500 mL)
	<ol>
		<li>Dissolve 48.81 g of MES in 400 mL of distilled water (dH2O).</li>
		<li>Adjust pH to 6.0.</li>
		<li>Bring the total volume to 500 mL with dH2O.</li>
	</ol>
	</li>
	<li>PBS Solution (1 L)
	<ol>
		<li>To make 10-fold PBS solution, dissolve 1 package of PBS premix powder in 1 L of dH2O.</li>
		<li>Combine 100 mL of 10-fold PBS solution with 900 mL of dH2O.</li>
	</ol>
	</li>
	<li>5% Polyvinylpyrrolidone (PVP) Solution (4 mL)
	<ol>
		<li>Under sterile conditions, dissolve 0.2 g of PVP in 4 mL of Drosophila Schneider&#39;s Medium. 
		NOTE: Always open Schneider&#39;s Medium stock in tissue culture (TC) hood. Keep for 1 month at 4 &#176;C.</li>
	</ol>
	</li>
	<li>0.65 mM Rhodamine Red-X stock solution (2 mL)
	<ol>
		<li>Weigh 1 mg of Rhodamine Red-X.</li>
		<li>Add 2 mL of dimethyl sulfoxide (DMSO) to dissolve.</li>
		<li>Aliquot and store at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Shelton&#39;s Growth Medium (SGM) (10.25 mL)
	<ol>
		<li>Under sterile conditions, combine 8 mL of Drosophila Schneider&#39;s Medium, 1 mL of PVP solution, 1 mL of Inulin solution, 1 mL of Basal Medium Eagle (BME) Vitamins, 50 &#181;L of Penicillin-Streptomycin, and 100 &#181;L of Lipid concentrate together.</li>
		<li>Aliquot 325 &#181;L of SGM into 1.5 mL microtubes. 
		NOTE: Keep for about 1 month at 4 &#176;C.</li>
		<li>On the day of blastomere isolation, add 175 &#181;L of Fetal Bovine Serum (FBS) to the SGM (for total volume of 500 &#181;L). 
		NOTE: Store FBS at -20 &#176;C and thaw it before usage. FBS does not need to be heat killed.</li>
	</ol>
	</li>
</ol>

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The authors declare no conflict of interests.

Reconstitution of simplified cell contact patterns will let researchers to test the roles of specific cell contact patterns in different aspects of morphogenesis. We have used this technique to show that cell division axis is controlled by the physical contact with adhesive beads10. As division axis specification is crucial for multicellular development by contributing to morphogenesis14, stem cell division15,16, an...

During multicellular development, the cellular behaviors (e.g., division axis) of individual cells are specified by various chemical and physical cues. To understand how individual cell interprets this information, and how they regulate multicellular assembly as an emergent property is one of the ultimate goals of morphogenesis studies. The model organism C. elegans has contributed significantly to the understanding of cellular-level regulation of morphogenesis such as cell polarity1, cell division patterning1, cell fate decision2, and tissue-scale regulations such as neuronal wiring3 and organogenesis4,5. Although there are various genetic tools available, tissue engineering methods are limited.
The most successful tissue engineering method in C. elegans study is the classical blastomere isolation6; as C. elegans embryo is surrounded by an eggshell and a permeability barrier7, their removal is one of the main procedures of this method. While this blastomere isolation method enables reconstitution of cell-cell contact in a simplified manner, it does not allow for the elimination of unwanted cues; cell contact still poses both mechanical (e.g., adhesion) and chemical cues, thereby limiting our ability to fully analyze the causal relationship between the cue and cellular behavior.
The method presented in this paper uses carboxylate modified polystyrene beads that can covalently bind to any amine-reactive molecules including proteins as ligands. Particularly, we used an amine-reactive form of Rhodamine Red-X as a ligand to make beads both visually trackable and adhesive to the cell. The carboxyl groups of bead surface and primary amine groups of ligand molecule are coupled by water-soluble carbodiimide 1-ethyl-3-(dimethylaminopropyl) carbodiimide (EDAC)8,9. Obtained adhesive beads allow for the effects of the mechanical cue on cellular dynamics10. We have used this technique to identify mechanical cues required for cell division orientation10.

<table>
	<tbody>
		<tr>
			<td>1-(3-Dimethylaminopropyl)-3-ethylcarbodiimide hydrochloride</td>
			<td>Alfa Aesar</td>
			<td>AAA1080703</td>
			<td>For the bead preparation</td>
		</tr>
		<tr>
			<td>Aspirator Tube Assembly</td>
			<td>Drummond</td>
			<td>21-180-13</td>
			<td>For the blastomere isolation.</td>
		</tr>
		<tr>
			<td>Caenorhabditis elegans strain: N2, wild-type</td>
			<td>Caenorhabditis Genetics Center</td>
			<td>N2</td>
			<td>Strain used in this study</td>
		</tr>
		<tr>
			<td>Caenorhabditis elegans strain: KSG5, genotype: zuIs45; itIs37</td>
			<td>in house</td>
			<td>KSG5</td>
			<td>Strain used in this study</td>
		</tr>
		<tr>
			<td>Calibrated Mircopipets, 10 &#181;L</td>
			<td>Drummond</td>
			<td>21-180-13</td>
			<td>For the blastomere isolation</td>
		</tr>
		<tr>
			<td>Carboxylate-modified polystyrene beads (30 &#181;m diameter)</td>
			<td>KISKER Biotech</td>
			<td>PPS-30.0COOHP</td>
			<td>For the bead preparation</td>
		</tr>
		<tr>
			<td>CD Lipid Concentrate</td>
			<td>Life Technologies</td>
			<td>11905031</td>
			<td>For the blastomere isolation. Work in the tissue culture hood.</td>
		</tr>
		<tr>
			<td>Clorox</td>
			<td>Clorox</td>
			<td>N. A.</td>
			<td>For the blastomere isolation. Open a new bottle when the hypochlorite treatment does not work well.</td>
		</tr>
		<tr>
			<td>Coverslip holder</td>
			<td>In house</td>
			<td>N.A.</td>
			<td>For the blastomere isolation.</td>
		</tr>
		<tr>
			<td>Dissecting microscope: Zeiss Stemi 508 with M stand. Source of light is built-in LED. Magnification of eye piece is 10X.</td>
			<td>Carl Zeiss</td>
			<td>Stemi 508</td>
			<td>For the blastomere isolation.</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum, Qualified One Shot, Canada origin</td>
			<td>Gibco</td>
			<td>A3160701</td>
			<td>For the blastomere isolation. Work in the tissue culture hood.</td>
		</tr>
		<tr>
			<td>General Use and Precision Glide Hypodermic Needles, 25 gauge</td>
			<td>BD</td>
			<td>14-826AA</td>
			<td>For the blastomere isolation</td>
		</tr>
		<tr>
			<td>Inulin</td>
			<td>Alfa Aesar</td>
			<td>AAA1842509</td>
			<td>For the blastomere isolation</td>
		</tr>
		<tr>
			<td>MEM Vitamin Solution (100x)</td>
			<td>Gibco</td>
			<td>11120052</td>
			<td>For the blastomere isolation.</td>
		</tr>
		<tr>
			<td>MES (Fine White Crystals)</td>
			<td>Fisher BioReagents</td>
			<td>BP300-100</td>
			<td>For the bead preparation</td>
		</tr>
		<tr>
			<td>Multitest Slide 10 Well</td>
			<td>MP Biomedicals</td>
			<td>ICN6041805</td>
			<td>For the blastomere isolation</td>
		</tr>
		<tr>
			<td>PBS, Phosphate Buffered Saline, 10 x Powder</td>
			<td>Fisher BioReagents</td>
			<td>BP665-1</td>
			<td>For the bead preparation</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin (10,000 U/mL)</td>
			<td>Gibco</td>
			<td>15140148</td>
			<td>For the blastomere isolation.</td>
		</tr>
		<tr>
			<td>Polyvinylpyrrolidone</td>
			<td>Fisher BioReagents</td>
			<td>BP431-100</td>
			<td>For the blastomere isolation</td>
		</tr>
		<tr>
			<td>Potassium Chloride</td>
			<td>Bioshop</td>
			<td>POC888</td>
			<td>For the blastomere isolation</td>
		</tr>
		<tr>
			<td>Rhodamine Red-X, Succinimidyl Ester, 5-isomer</td>
			<td>Molecular Probes</td>
			<td>R6160</td>
			<td>For the bead preparation</td>
		</tr>
		<tr>
			<td>Schneider's Drosophila Sterile Medium</td>
			<td>Gibco</td>
			<td>21720024</td>
			<td>For the blastomere isolation. Work in the tissue culture hood.</td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>Bioshop</td>
			<td>SOD001</td>
			<td>For the blastomere isolation</td>
		</tr>
		<tr>
			<td>Sodium Hydroxide Solution, 10 N</td>
			<td>Fisher Chemical</td>
			<td>SS255-1</td>
			<td>For the blastomere isolation</td>
		</tr>
		<tr>
			<td>Spinning disk confocal microscope: Yokogawa CSU-X1, Zeiss Axiovert inverted scope, Quant EM 512 camera, 63X NA 1.4 Plan apochromat objective lens. System was controlled by Slidebook 6.0.</td>
			<td>Intelligent Imaging Innovation</td>
			<td>N.A.</td>
			<td>For live-imaging</td>
		</tr>
		<tr>
			<td>Syringe Filters, PTFE, Non-Sterile</td>
			<td>Basix</td>
			<td>13100115</td>
			<td>For the blastomere isolation.</td>
		</tr>
		<tr>
			<td>Tygon S3 Laboratory Tubing,, Formulation E-3603, Inner diameter 3.175 mm</td>
			<td>Saint Gobain Performance Plastics</td>
			<td>89403-862</td>
			<td>For the blastomere isolation.</td>
		</tr>
		<tr>
			<td>Tygon S3 Laboratory Tubing,, Formulation E-3603, Inner diameter 6.35 mm</td>
			<td>Saint Gobain Performance Plastics</td>
			<td>89403-854</td>
			<td>For the blastomere isolation.</td>
		</tr>
	</tbody>
</table>

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.

For beads preparation, we determined the optimal amount of Rhodamine Red-X succinimidyl ester for the transgenic strain expressing GFP-myosin II and mCherry-histone (Figure 1A-D). We used mCherry tagged histone as a marker of cell cycle progression. Because both Rhodamine Red-X and mCherry will be illuminated by a 561 nm laser, the optimal intensity of Rhodamine Red-X signal is comparable to that of histone to allow simultaneous imaging of cell and bead. For example, the flu...

Watch this Scientific Journal Video about In Vitro Reconstitution of Spatial Cell Contact Patterns with Isolated Caenorhabditis elegans Embryo Blastomeres and Adhesive Polystyrene Beads at JoVE.com

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 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. ...

in-vitro-reconstitution-spatial-cell-contact-patterns-with-isolated

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

7.0K Views.  The University of British Columbia. 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.

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

Comprehensive Assessment of Germline Chemical Toxicity Using the Nematode Caenorhabditis elegans

Identifying the reproductive toxicity of the thousands of chemicals present in our environment has been one of the most tantalizing challenges in the field of environmental health. This is due in part to the paucity of model systems that can (1) accurately recapitulate keys features of reproductive processes and (2) do so in a medium- to high-throughput fashion, without the need for a high number of vertebrate animals.
We describe here an assay in the nematode C. elegans that allows the rapid identification of germline toxicants by monitoring the induction of aneuploid embryos. By making use of a GFP reporter line, errors in chromosome segregation resulting from germline disruption are easily visualized and quantified by automated fluorescence microscopy. Thus the screening of a particular set of compounds for its toxicity can be performed in a 96- to 384-well plate format in a matter of days. Secondary analysis of positive hits can be performed to determine whether the chromosome abnormalities originated from meiotic disruption or from early embryonic chromosome segregation errors. Altogether, this assay represents a fast first-pass strategy for the rapid assessment of germline dysfunction following chemical exposure.

Comprehensive Assessment of Germline Chemical Toxicity Using the Nematode Caenorhabditis elegans

We describe the detailed steps of a high-throughput chemical assay in the nematode Caenorhabditis elegans used to assess germline toxicity. In this assay, disruption of germline function following chemical exposure is monitored using a fluorescent reporter specific to aneuploid embryos.

Identifying the reproductive toxicity of the thousands of chemicals present in our environment has been one of the most tantalizing challenges in the ...

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 ...

Application of Impermeable Barriers Combined with Candidate Factor Soaked Beads to Study Inductive Signals in the Chick

The chick embryo provides a superb vertebrate model that can be used to dissect developmental questions in a direct way. Its accessibility and robustness following surgical intervention are key experimental strengths. Mica plates were the first barriers used to prevent chick limb bud initiation1. Protocols that use aluminum foil as an impermeable barrier to wing bud or leg bud induction and or initiation are described. We combine this technique with bead placement lateral to the barrier to exogenously supply candidate endogenous factors that have been blocked by the barrier. The results are analyzed using in situ hybridization of subsequent gene expression. Our main focus is on the role of retinoic acid signaling in the induction and later initiation of the chick embryo fore and hindlimb. We use BMS 493 (an inverse agonist of retinoic acid receptors (RAR)) soaked beads implanted in the lateral plate mesoderm (LPM) to mimic the effect of a barrier placed between the somites (a source of retinoic acid (RA)) and the LPM from which limb buds grow. Modified versions of these protocols could also be used to address other questions on the origin and timing of inductive cues. Provided the region of the chick embryo is accessible at the relevant developmental stage, a barrier could be placed between the two tissues and consequent changes in development studied. Examples may be found in the developing brain, axis extension and in organ development, such as liver or kidney induction.

Protocols using impermeable barriers to block induction events between tissues of the main body axis and flank in the chick embryo required for limb formation are described. Beads soaked in candidate inductive signals are used to overcome the effect of barrier placement and analysis of gene expression confirms this.

The chick embryo provides a superb vertebrate model that can be used to dissect developmental questions in a direct way. Its accessibility and robustness ...

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

The evolutionarily conserved&#160;extracellular signal transducing RTK-RAS-ERK pathway is an important kinase-signaling cascade that controls multiple cellular and developmental processes principally via activation of ERK, the terminal kinase of the pathway. Tight regulation of ERK activity is essential for normal development and homeostasis; overly active ERK results in excessive cellular proliferation, while underactive ERK causes cell death. C. elegans is a powerful model system that has helped characterize the function and regulation of RTK-RAS-ERK signaling pathway during development. In particular, the RTK-RAS-ERK pathway is essential for C. elegans germline development, which is the focus of this method. Using antibodies specific to the active, diphosphorylated form of ERK (dpERK), the stereotypical localization pattern can be visualized within the germline. Because this pattern is both spatially and temporally controlled, the ability to reproducibly assay dpERK is useful to identify regulators of the pathway that affect dpERK signal duration and amplitude and thus germline development. Here we demonstrate how to successfully dissect, stain, and image dpERK within the C. elegans gonad. This method can be adapted for spatial localization of any signaling or structural protein in the C. elegans gonad, provided an antibody compatible with immunofluorescence is available.

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

We present an immunofluorescence&#160;imaging-based method for spatial and temporal localization of active ERK in the dissected C. elegans gonad. The protocol described here can be adapted for visualization of any signaling or structural protein in the C. elegans gonad, provided a suitable antibody reagent is available.

The evolutionarily conserved&#160;extracellular signal transducing RTK-RAS-ERK pathway is an important kinase-signaling cascade that controls multiple ...

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; ...

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. ...

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

Axonal transport and intraflagellar transport (IFT) are essential for axon and cilia morphogenesis and function. Kinesin superfamily proteins and dynein are molecular motors that regulate anterograde and retrograde transport, respectively. These motors use microtubule networks as rails. Caenorhabditis elegans (C. elegans) is a powerful model organism to study axonal transport and IFT in vivo. Here, I describe a protocol to observe axonal transport and IFT in living C. elegans. Transported cargo can be visualized by tagging cargo proteins using fluorescent proteins such as green fluorescent protein (GFP). C. elegans is transparent and GFP-tagged cargo proteins can be expressed in specific cells under cell-specific promoters. Living worms can be fixed by microbeads on 10% agarose gel without killing or anesthetizing the worms. Under these conditions, cargo movement can be directly observed in the axons and cilia of living C. elegans without dissection. This method can be applied to the observation of any cargo molecule in any cells by modifying the target proteins and/or the cells they are expressed in. Most basic proteins such as molecular motors and adaptor proteins that are involved in axonal transport and IFT are conserved in C. elegans. Compared to other model organisms, mutants can be obtained and maintained more easily in C. elegans. Combining this method with various C. elegans mutants can clarify the molecular mechanisms of axonal transport and IFT.

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

Caenorhabditis elegans (C. elegans) is a good model to study axonal and intracellular transport. Here, I describe a protocol for in vivo recording and analysis of axonal and intraflagellar transport in C. elegans.

Axonal transport and intraflagellar transport (IFT) are essential for axon and cilia morphogenesis and function. Kinesin superfamily proteins and dynein ...

Assessing Lysosomal Alkalinization in the Intestine of Live Caenorhabditis elegans

The nematode Caenorhabditis elegans (C. elegans) is a model system that is widely used to study longevity and developmental pathways. Such studies are facilitated by the transparency of the animal, the ability to do forward and reverse genetic assays, the relative ease of generating fluorescently labeled proteins, and the use of fluorescent dyes that can either be microinjected into the early embryo or incorporated into its food (E. coli strain OP50) to label cellular organelles (e.g. 9-diethylamino-5H-benzo(a)phenoxazine-5-one and (3-{2-[(1H,1&#39;H-2,2&#39;-bipyrrol-5-yl-kappaN(1))methylidene]-2H-pyrrol-5-yl-kappaN}-N-[2-(dimethylamino)ethyl]propanamidato)(difluoro)boron). Here, we present the use of a fluorescent pH-sensitive dye that stains intestinal lysosomes, providing a visual readout of dynamic, physiological changes in lysosomal acidity in live worms. This protocol does not measure lysosomal pH, but rather aims to establish a reliable method of assessing physiological relevant variations in lysosomal acidity. cDCFDA is a cell-permeant compound that is converted to the fluorescent fluorophore 5-(and-6)-carboxy-2&#39;,7&#39;-dichlorofluorescein (cDCF) upon hydrolysis by intracellular esterases. Protonation inside lysosomes traps cDCF in these organelles, where it accumulates. Due to its low pKa of 4.8, this dye has been used as a pH sensor in yeast. Here we describe the use of cDCFDA as a food supplement to assess the acidity of intestinal lysosomes in C. elegans. This technique allows for the detection of alkalinizing lysosomes in live animals, and has a broad range of experimental applications including studies on aging, autophagy, and lysosomal biogenesis.

Assessing Lysosomal Alkalinization in the Intestine of Live Caenorhabditis elegans

A step-by-step guide to probe loss of lysosomal acidity in the intestine of C. elegans using the pH-sensitive vital dye 5(6)-carboxy-2&#39;,7&#39;-dichlorofluorescein diacetate (cDCFDA)

The nematode Caenorhabditis elegans (C. elegans) is a model system that is widely used to study longevity and developmental pathways. Such studies are ...

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to understand morphogenetic processes is to directly measure cellular forces and mechanical properties in vivo during embryogenesis. Here, we present a setup of optical tweezers coupled to a light sheet microscope, which allows to directly apply forces on cell-cell contacts of the early Drosophila embryo, while imaging at a speed of several frames per second. This technique has the advantage that it does not require the injection of beads into the embryo, usually used as intermediate probes on which optical forces are exerted. We detail step by step the implementation of the setup, and propose tools to extract mechanical information from the experiments. By monitoring the displacements of cell-cell contacts in real time, one can perform tension measurements and investigate cell contacts&#39; rheology.

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

We present a setup of optical tweezers coupled to a light sheet microscope, and its implementation to probe cell mechanics without beads in the Drosophila embryo.

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to ...

Studying Oxidative Stress Caused by the Mitis Group Streptococci in Caenorhabditis elegans

Caenorhabditis elegans (C. elegans), a free-living nematode, has emerged as an attractive model to study host-pathogen interactions. The presented protocol uses this model to determine the pathogenicity caused by the mitis group streptococci via the production of H2O2. The mitis group streptococci are an emerging threat that cause many human diseases such as bacteremia, endocarditis, and orbital cellulitis. Described here is a protocol to determine the survival of these worms in response to H2O2 produced by this group of pathogens. Using the gene skn-1 encoding for an oxidative stress response transcription factor, it is shown that this model is important for identifying host genes that are essential against streptococcal infection. Furthermore, it is shown that activation of the oxidative stress response can be monitored in the presence of these pathogens using a transgenic reporter worm strain, in which SKN-1 is fused to green fluorescent protein (GFP). These assays provide the opportunity to study the oxidative stress response to H2O2 derived by a biological source as opposed to exogenously added reactive oxygen species (ROS) sources.

Studying Oxidative Stress Caused by the Mitis Group Streptococci in Caenorhabditis elegans

The nematode Caenorhabditis elegans is an excellent model to dissect host-pathogen interactions. Described here is a protocol to infect the worm with members of the mitis group streptococci and determine activation of the oxidative stress response against H2O2 produced by this group of organisms.

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

Summary

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Chapters in this video

Comprehensive Assessment of Germline Chemical Toxicity Using the Nematode Caenorhabditis elegans

Imaging Subcellular Structures in the Living Zebrafish Embryo

Application of Impermeable Barriers Combined with Candidate Factor Soaked Beads to Study Inductive Signals in the Chick

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

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

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

Assessing Lysosomal Alkalinization in the Intestine of Live Caenorhabditis elegans

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

Studying Oxidative Stress Caused by the Mitis Group Streptococci in Caenorhabditis elegans