The authors would like to acknowledge the support by the &#8220;Animalerie Centrale de Gif-sur-Yvette&#8221; for animal husbandry. This work was supported by funds from ANR (ANR-09-BLAN-0262-02 and ANR-11-JSV2-002-01) to Michael Schubert, by the European Union FP6 grant &#8220;Embryomics&#8221; and by the ANR grant &#8220;ANR-10-BLAN-121801 Dev-Process&#8221; to Jean-Fran&#231;ois Nicolas and Nadine Peyri&#233;ras. Jo&#227;o Emanuel Carvalho is financed by a FCT doctoral fellowship (SFRH/BD/86878/2012).
Requests for the vectors described here can be addressed directly to the authors.

1. Preparation of Instruments and Reagents
<ol style="margin-left: 40px;">
	<li>Transfer Pasteur pipettes
	<ol>
		<li>Generate a series of transfer Pasteur pipettes with different tip diameters by pulling 230 mm long Pasteur pipettes above a flame at different speeds. Ensure that the taper is as long as possible for smooth and fine control of aspiration.</li>
		<li>With a diamond scribe, scratch the pipette along a line perpendicular to the length of the pipette. With both hands, pull the pipette parallel to its length to generate a blunt cut. Swiftly flame-polish the pipette without sealing the tip.</li>
		<li>Vary the diameter of the tip with the stage of the oocytes/embryos to be pipetted: 300-400 &#181;m for unfertilized oocytes (around 150 &#181;m in diameter), 600 &#181;m for fertilized eggs (around 500 &#181;m in diameter), 200-400 &#181;m for hatched neurulae. Use pipettes with a mouth-monitored aspiration tube to transfer oocytes/embryos from one dish to another.</li>
	</ol>
	</li>
	<li>Injection needles
	<ol>
		<li>Manipulate the capillaries with gloves to ensure RNase-free conditions. Use borosilicate glass with filament capillaries with dimensions of OD 1.20 mm, ID 0.94 mm, length 10 mm.</li>
		<li>If capillaries are pulled on the type of heating-filament needle puller described in the Materials List, use the following settings: Heat 600, Pull 50, Velocity 80, Time 60, Pressure 200 or 300. Otherwise, ensure that the shape, which is crucial for successful injections, is as shown in Figure 2 with the following properties: 4-8 &#181;m outer tip diameter, 2 cm taper length. 
		NOTE: Needles can be pulled before the spawning season and used throughout the season.</li>
	</ol>
	</li>
	<li>5% Phenol Red stock solution (4x)
	<ol>
		<li>Prepare fresh before each spawning season.</li>
		<li>In a 0.22 &#181;m-filtration tube, weigh out 25 mg of Phenol Red powder.</li>
		<li>Add 0.5 ml of DNase- and RNase-free water to the tube containing the powder.</li>
		<li>Spin for 3-5 min at 18,000 x g at room temperature to filter-sterilize the solution and remove crystals that could clog the injection needle.</li>
		<li>Store at 4 &#176;C or store 250 &#181;l aliquots at -20 &#176;C.</li>
	</ol>
	</li>
	<li>0.25 mg/ml poly-lysine solution
	<ol>
		<li>Prepare fresh before each spawning season.</li>
		<li>Dissolve 5 mg of poly-L-lysine in 20 ml distilled water. Store 5 ml aliquots at -20 &#176;C.</li>
		<li>Use a defrosted aliquot immediately and only once to ensure reproducible and robust adhesion of the oocytes to the poly-lysine-coated dish.</li>
	</ol>
	</li>
	<li>mRNA synthesis
	<ol>
		<li>Prepare fresh before each spawning season.</li>
		<li>Linearize 5 &#181;g of DNA of interest with the adequate enzyme (usually for 2 hr, at 37 &#176;C). To check the completeness of the digestion, run 2% in volume of the digestion mix on a 1% agarose-TBE gel in TBE buffer at 150 W for 20 min.</li>
		<li>Extract the linearized DNA with 25:24:1 phenol (pH 8.0):chloroform:isoamyl alcohol. Vortex for 20 sec, centrifuge 10 min at 18,000 x g and collect the aqueous (upper) phase.</li>
		<li>Extract the aqueous phase again with 24:1 chloroform:isoamyl alcohol. Vortex for 20 sec, centrifuge 10 min at 18,000 x g and collect the aqueous phase.</li>
		<li>Precipitate the linearized DNA with 100:10:300 linearized DNA:3 M sodium acetate (pH 5.2):100% ethanol overnight at -20 &#176;C.</li>
		<li>Centrifuge 20 min at 18,000 x g at 4 &#176;C. Rinse in 70% ethanol.</li>
		<li>Centrifuge 10 min at 18,000 x g at 4 &#176;C. Let dry and resuspend in RNase-free water at a final concentration of 0.5 &#181;g/&#181;l.</li>
		<li>Transcribe 1 &#181;g of linearized DNA using an mRNA synthesis kit with the appropriate polymerase, according to manufacturer&#8217;s instructions.</li>
		<li>Extract the mRNA with 5:1 phenol (pH 4.7):chloroform and ammonium acetate stop solution provided with the kit.</li>
		<li>Vortex for 20 sec, centrifuge 10 min at 18,000 x g and collect the aqueous phase.</li>
		<li>Extract the aqueous phase again with 24:1 chloroform:isoamyl alcohol. Vortex for 20 sec, centrifuge 10 min at 18,000 x g and collect the aqueous phase.</li>
		<li>Precipitate the mRNA with 100% isopropanol overnight at -20 &#176;C.</li>
		<li>Centrifuge 20 min at 18,000 x g at 4 &#176;C. Rinse in 80% ethanol.</li>
		<li>Centrifuge 10 min at 18,000 x g at 4 &#176;C. Let the pellet dry at room temperature for no longer than 5-10 min as it will then be difficult to resuspend. Resuspend in DNase- and RNase-free water to a final concentration of at least 2 &#181;g/&#181;l to ensure a decent final mRNA concentration in the injection mix.</li>
		<li>To check the quality and size of the transcription product, run 0.5 &#181;l of the mRNA on a RNase-free 1% agarose-TBE gel in TBE buffer at 150 W for 20 min. Store 2 &#181;l aliquots at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Poly-lysine-coated dishes 
	NOTE: Poly-lysine coated dishes are used to immobilize the oocytes during injection.
	<ol>
		<li>For each 35 mm cell-culture Petri dish (5 in total), cover the bottom of the Petri dish with 1 ml of the thawed 0.25 mg/ml poly-lysine solution. Incubate at room temperature for 5 min.</li>
		<li>For each 35 mm cell-culture Petri dish (5 in total), transfer the 0.25 mg/ml poly-lysine solution into another 35 mm cell-culture Petri dish. Incubate at room temperature for 5 min.</li>
		<li>Discard the 0.25 mg/ml poly-lysine solution.</li>
		<li>Let the Petri dishes dry, upside-down at room temperature for 2 hr.</li>
		<li>Store the poly-lysine-coated dishes wrapped in a plastic wrap at 4 &#176;C to avoid contamination for one week maximum.</li>
	</ol>
	</li>
	<li>Agarose-coated dishes 
	NOTE: They are used to culture injected embryos. The agarose provides a cushion for the injected embryos and prevents them from sticking to the bottom of the dish.
	<ol>
		<li>Make artificial seawater (ASW) using 37-38 g/L commercial salts + 0.25 mM NaHCO3 in reverse osmosis water.</li>
		<li>Dissolve agarose to a 1% concentration in 0.22 &#181;m-filtered ASW by heating the solution in a microwave.</li>
		<li>Swiftly pour the warm agarose solution from one 35 mm Petri dish into another one in order to ensure a very thin agarose coating of the dish.</li>
		<li>Store the agarose-coated dishes wrapped in Saran wrap at 4 &#176;C to avoid contamination for one week maximum.</li>
	</ol>
	</li>
	<li>Injection mix and loading of the injection needles
	<ol>
		<li>About 2 hr before starting the injections, make a 2 &#181;l injection mix in RNase- and DNase-free water with final concentrations of 1-1.8 &#181;g/&#181;l of mRNA, 15% glycerol, 1.25% Phenol Red. 
		NOTE: The Phenol Red colors the solution, which allows monitoring of the injection efficiency and the identification of successfully injected embryos. Glycerol favors mRNA diffusion within the oocyte.</li>
		<li>Centrifuge 4 min at 18,000 x g to pellet crystals. Keep on ice until use.</li>
		<li>With a 10 &#181;l pipette, collect 0.5 &#181;l of the injection mix, avoiding the bottom of the tube, where the crystals have been pelleted.</li>
		<li>Backfill at least two injection needles (in case one breaks during the injection) by pipetting the 0.5 &#181;l drop of injection mix at the large opening of the needle.</li>
		<li>Install the needles in a storage jar with liquid at the bottom at 4 &#176;C to prevent evaporation of the injection mix. Let the injection mix slowly travel to the tip of the injection needle for at least 1 hr to minimize the creation of bubbles.</li>
		<li>Store additional injection mix at -80 &#176;C for maximum of three additional uses, after which the mRNA quality deteriorates (data not shown).</li>
	</ol>
	</li>
</ol>
2. Collection of Biological Material, Microinjection and Embryo Culture
<ol style="margin-left: 40px;">
	<li>Oocyte and sperm collection 
	NOTE: See Theodosiou et al.12 for a detailed protocol for inducing spawning and for gamete collection.
	<ol>
		<li>Shock males and females in ASW at 23 &#176;C for 24 hr.</li>
		<li>One to two hours before sunset, transfer the adults into individual cups in ASW at 19 &#176;C because most adults will spawn 1-2 hr after sunset.</li>
		<li>Rinse 35 mm Petri dishes in filtered ASW and let them dry upside-down to prevent the oocytes from sticking to the bottom of the dish.</li>
		<li>Upon spawning, immediately collect sperm and oocytes with a 1,000 &#181;l pipette.
		<ol>
			<li>Keep sperm and oocytes separate from adult amphioxus because the contact is detrimental for gamete health (data not shown).</li>
			<li>Furthermore, avoid startling the adults, which leads to movements that dissipate, and hence dilute, both sperm and oocytes. To keep the sperm active as long as possible and to optimize fertilization rate, collect the sperm as concentrated as possible.</li>
		</ol>
		</li>
		<li>Keep sperm on ice in a 1.5 ml tube.</li>
		<li>Transfer oocytes in filtered ASW into the pre-rinsed 35 mm Petri dishes.</li>
		<li>Transfer 100-500 oocytes with a previously pulled 300-400 &#181;m transfer Pasteur pipette to another 35 mm Petri dish to perform the injections.</li>
		<li>Fertilize the remainder of the clutch as a control for sperm and oocyte quality or for other experiments.</li>
	</ol>
	</li>
	<li>Oocyte injection
	<ol>
		<li>Install the injection needle on the micromanipulator at a 50&#176; angle relative to the horizontal plane. 
		NOTE: Angles of less than 50&#176; will push the oocytes around on the dish, while angles of more than 50&#176; will not allow an appropriate monitoring of the needle position relative to the oocyte.</li>
		<li>Under a fluorescent dissecting scope with 25X oculars, transfer 30 oocytes with the 300-400 &#181;m transfer Pasteur pipette on a poly-lysine-coated dish containing filtered ASW.</li>
		<li>Deposit the oocytes along a line to carry out injections in an ordered way and to distinguish injected from non-injected oocytes. Inject small numbers (30 oocytes) to minimize the exposure time of oocytes to poly-lysine, which tends to deform developing embryos (data not shown).</li>
		<li>Use the dark field illumination to render the oocytes as translucent as possible.</li>
		<li>With the coarse movement knob of the micromanipulator, bring the injection needle close to an oocyte.</li>
		<li>With fine forceps, cut open the needle at the level where the tip starts to be curved. By pulsing with the injector, verify that red injection mix is actually flowing out of the needle.</li>
		<li>At 200X magnification and with the fine movement knob of the micromanipulator, gently move the injection needle inside the core of the oocyte. 
		NOTE: If inserted too superficially, the injected solution will not remain inside the oocyte. If inserted too far, the oocyte will be destroyed.</li>
		<li>Inject with 1-3 pulses of 120 msec duration and 1-10 psi pressure. If the needle is fine enough, inject with continuous flow at constant pressure. Ensure that the injection volume corresponds to 1/5 to 1/3 of the volume of a single oocyte.</li>
		<li>Following injection, pull the needle out swiftly to avoid leakage of the oocyte.</li>
		<li>Verify that the injected solution remains within the oocyte and that after a few sec, the injected solution spreads throughout the oocyte.</li>
		<li>Move on to the next oocyte in line.</li>
		<li>Keep some uninjected embryos of each series as negative control to estimate the background fluorescence when scanning for injected embryos.</li>
	</ol>
	</li>
	<li>Fertilization, selection of injected embryos and embryo culture
	<ol>
		<li>Fertilize the oocytes as soon as a series has been injected. As oocyte quality declines with time, inject and fertilize oocytes within 1 hr after spawning12.</li>
		<li>Depending on the sperm concentration, add 1-5 drops of sperm to the oocytes and swirl the dish. 
		NOTE: The fertilization envelope should become apparent on the embryos after about 1 min.</li>
		<li>Allow the embryos to detach from the poly-lysine-coated dish, while injecting another series of oocytes.</li>
		<li>Transfer the embryos with the 600 &#181;m transfer Pasteur pipette into an agarose-coated Petri dish. Remove the embryos from the poly-lysine-coated dish as soon as possible, if at all possible before the 2-cell stage. 
		NOTE: In case of prolonged exposure to poly-lysine, the embryos tend to become densely-packed blastulae, flattened on the side touching the bottom of the dish.</li>
		<li>At the 2-cell to 4-cell stage, select with a fluorescent dissecting scope with DSR filter the successfully-injected embryos, i.e., those with a normal morphology that exhibit a Phenol Red-derived red fluorescent signal.</li>
		<li>Keep the embryos in culture in filtered ASW in agarose-coated Petri dishes at 19 &#176;C until the desired stage for in vivo imaging.</li>
	</ol>
	</li>
</ol>

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Genet. Genomics. 41, 215-219 (2014).</li><li>Zhang, Q. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Continuous+culture+of+two+lancelets+and+production+of+the+second+filial+generations+in+the+laboratory.">Continuous culture of two lancelets and production of the second filial generations in the laboratory.</a> J. Exp. Zool. B. 308, 464-472 (2007).</li><li>Yasui, K., Igawa, T., Kaji, T., Henmi, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stable+aquaculture+of+the+Japanese+lancelet+Branchiostoma+japonicum+for+7+years.">Stable aquaculture of the Japanese lancelet Branchiostoma japonicum for 7 years.</a> J. Exp. Zool. B. 320, 538-547 (2013).</li><li>Benito-Guti&#233;rrez, E., Weber, H., Bryant, D. V., Arendt, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+generating+year-round+access+to+amphioxus+in+the+laboratory.">Methods for generating year-round access to amphioxus in the laboratory.</a> PLoS One. 8, e71599 (1371).</li></ol>

The authors have nothing to disclose.

In this article, we present, for the first time, a detailed and reproducible protocol for the injection of B. lanceolatum oocytes, which, after B. floridae13,14 and B. belcheri15, is thus the third amphioxus species, for which such a technique has been described. Importantly, the protocol described here also includes the description of vector systems suited for the production of fluorescent proteins in B. lanceolatum from injected mRNA produced in vitro (...

During development, a single cell gives rise to an entire organism in a highly complex process that involves both cell divisions and movements. To better understand the biological principles underlying the dynamics of cell behavior, developmental biologists have started to use fluorescence-based in vivo imaging techniques. Specific compartments of cells, such as cell membranes, can either be labeled by treatments with fluorescent dyes, an approach hampered by a lack of specificity and of tissue penetration1, or by the specific introduction into the embryo of exogenous mRNAs encoding fluorescent proteins2. Different techniques can be used for the efficient delivery of exogenous compounds, such as mRNAs. These include, but are not limited to, microinjection, electroporation, bombardment with microparticles, lipofection and transduction3,4. Although all of these approaches can be used to introduce exogenous compounds into a developing embryo, only microinjection allows the application of predefined and precise quantities into each cell3. Microinjection techniques have been described for all major developmental model systems4 (e.g., fruit flies, nematode worms, zebrafish, frogs, mice) as well as for some alternative models4, including those used for comparative studies aimed at understanding the evolution of developmental mechanisms (e.g., sea anemones, annelid worms, sea urchins, ascidian tunicates, the cephalochordate amphioxus).
Cephalochordates, which together with tunicates and vertebrates establish the chordate phylum, are particularly well-suited models to study the evolution of chordates and the diversification of vertebrates from an invertebrate ancestor5-8. The cephalochordate lineage diverged very early during chordate evolution; and extant cephalochordates, which are subdivided into three genera (Branchiostoma, Asymmetron and Epigonichthys), resemble vertebrates both in terms of overall anatomy and genome architecture5-8. Of the about 30 species of cephalochordates that have been described so far, five are available for embryological and developmental studies6,9: Asymmetron lucayanum (the Bahama lancelet), Branchiostoma floridae (the Florida amphioxus), Branchiostoma lanceolatum (the European amphioxus), Branchiostoma belcheri (the Chinese amphioxus) and Branchiostoma japonicum (the Japanese amphioxus). Ripe adults of three of these species (B. lanceolatum, B. belcheri and B. japonicum) can be induced to spawn on-demand during the breeding season10,11. In addition, at least for B. lanceolatum, efficient spawning can also be induced in artificial sea water12, thereby making this particular cephalochordate species accessible for laboratories that do not have access to natural seawater. The combination, in B. lanceolatum, of a convenient and reliable access to embryos with an efficient delivery method, such as microinjection, so far the only delivery technique developed in amphioxus (in both B. floridae and B. belcheri)13-15, will enable the development of a novel suite of manipulative techniques, including lineage tracing- and dynamic cell behavior-based approaches.
A protocol for the efficient microinjection of mRNAs to express fluorescent proteins in the B. lanceolatum embryo was hence developed. Furthermore, to provide a basic toolkit for live imaging of B. lanceolatum embryos, vector systems were developed that allow membrane-associated and nuclear expression of fluorescent proteins. For membrane targeting, enhanced green fluorescent protein (eGFP) was fused to the human HRAS CAAX box and nuclear localization of mCherry and eGFP was obtained by fusion to the zebrafish histone 2B (H2B) exon (Figure 1, Supplementary File 1). Furthermore, with the goal to optimize protein translation, the Kozak sequences and codons of the constructs have been modified and adapted to usage in B. lanceolatum. Taken together, the injection method and expression vectors presented here will serve as a basis for the generation of new experimental approaches for cephalochordates, notably analyses using the latest fluorescence-based in vivo imaging techniques.

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			<td></td>
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			<td height="20" style="height:20px;">35 mm Petri dishes</td>
			<td>Falcon</td>
			<td>353001</td>
			<td>culture-treated</td>
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			<td height="21" style="height:21px;width:297px;">Filtration unit (Stericup 1L)</td>
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			<td>0.22 micron filtration</td>
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			<td height="21" style="height:21px;width:297px;">Spin-X tubes</td>
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			<td>0.22 micron filtration tube</td>
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			<td height="21" style="height:21px;width:297px;">Phenol Red</td>
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			<td height="21" style="height:21px;width:297px;">Poly-L-Lysine hydrobromide</td>
			<td style="width:156px;">SIGMA</td>
			<td style="width:143px;">P9155</td>
			<td></td>
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			<td height="21" style="height:21px;width:297px;">H2O Dnase, Rnase-free</td>
			<td style="width:156px;">Gibco</td>
			<td style="width:143px;">10977-035</td>
			<td></td>
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			<td height="21" style="height:21px;width:297px;">mMessage mMachine SP6 Transcription kit</td>
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			<td style="width:143px;">AM1340</td>
			<td>mRNA synthesis kit</td>
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			<td height="21" style="height:21px;width:297px;">Phenol pH8</td>
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			<td style="width:156px;">SIGMA</td>
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			<td style="width:156px;">Europrix&#160;</td>
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			<td>Commercial salts</td>
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			<td style="width:143px;"></td>
			<td></td>
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			<td height="21" style="height:21px;width:297px;">Equipment</td>
			<td style="width:156px;"></td>
			<td style="width:143px;"></td>
			<td></td>
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		<tr height="40">
			<td height="40" style="height:40px;width:297px;">Fluorescent dissecting scope with 200x magnification&#160;</td>
			<td style="width:156px;">Leica</td>
			<td style="width:143px;">MZ16F</td>
			<td>25x oculars, DSR and GFP2 filters</td>
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			<td height="21" style="height:21px;width:297px;">Micromanipulator</td>
			<td style="width:156px;">Marzhauzer</td>
			<td style="width:143px;">M-33</td>
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			<td height="21" style="height:21px;width:297px;">Injector</td>
			<td style="width:156px;">Picospritzer</td>
			<td style="width:143px;">model II or III</td>
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			<td height="21" style="height:21px;width:297px;">Needle puller</td>
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			<td style="width:143px;">P97</td>
			<td>heating-filament needle puller</td>
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			<td height="20" style="height:20px;width:297px;">Fine forceps</td>
			<td>FINE SCIENCE TOOLS GMBH</td>
			<td>11252-30</td>
			<td>Dumont #5</td>
		</tr>
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expression of fluorescent proteins in branchiostoma lanceolatum by mrna injection into unfertilized oocytes

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

The protocol detailed above provides the basis for the microinjection of B. lanceolatum oocytes and hence for the introduction into developing B. lanceolatum embryos of mRNA encoding fluorescent proteins for in vivo imaging. Although the technique is certainly robust and reliable, the rate of successful injections using this protocol remains variable (Table 1). The very likely explanation for this intriguing fact is the extreme variability of oocyte clutches: different egg batc...

Watch this Scientific Journal Video about Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes at JoVE.com

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

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

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

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

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Developmental Biology

10.9K Views.  Institut Pasteur. We report here the robust and efficient expression of fluorescent proteins after mRNA injection into unfertilized oocytes of Branchiostoma lanceolatum. The development of the microinjection technique in this basal chordate will pave the way for far-reaching technical innovations in this emerging model system, including in vivo imaging and gene-specific manipulations.

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

Manipulation and In Vitro Maturation of Xenopus laevis Oocytes, Followed by Intracytoplasmic Sperm Injection, to Study Embryonic Development

Amphibian eggs have been widely used to study embryonic development. Early embryonic development is driven by maternally stored factors accumulated during oogenesis. In order to study roles of such maternal factors in early embryonic development, it is desirable to manipulate their functions from the very beginning of embryonic development. Conventional ways of gene interference are achieved by injection of antisense oligonucleotides (oligos) or mRNA into fertilized eggs, enabling under- or over-expression of specific proteins, respectively. However, these methods normally require more than several hours until protein expression is affected, and, hence, the interference of gene functions is not effective during early embryonic stages. Here, we introduce an experimental system in which expression levels of maternal proteins can be altered before fertilization. Xenopus laevis oocytes obtained from ovaries are defolliculated by incubating with enzymes. Antisense oligos or mRNAs are injected into defolliculated oocytes at the germinal vesicle (GV) stage. These oocytes are in vitro matured to eggs at the metaphase II (MII) stage, followed by intracytoplasmic sperm injection (ICSI). By this way, up to 10% of ICSI embryos can reach the swimming tadpole stage, thus allowing functional tests of specific gene knockdown or overexpression. This approach can be a useful way to study roles of maternally stored factors in early embryonic development.

Manipulation and In Vitro Maturation of Xenopus laevis Oocytes, Followed by Intracytoplasmic Sperm Injection, to Study Embryonic Development

We describe methods of manipulating Xenopus laevis immature oocytes, in vitro maturation of oocytes to eggs, and intracytoplasmic sperm injection. This protocol allows degradation of some maternal proteins and overexpression of genes of interest at fertilization, and hence is valuable to study roles of specific factors in early embryonic development.

Amphibian eggs have been widely used to study embryonic development. Early embryonic development is driven by maternally stored factors accumulated during ...

Gene Transfer into the Chicken Auditory Organ by In Ovo Micro-electroporation

Chicken embryos are ideal model systems for studying embryonic development as manipulations of gene function can be conducted with relative ease in ovo. The inner ear auditory sensory organ is critical for our ability to hear. It houses a highly specialized sensory epithelium that consists of mechano-transducing hair cells (HCs) and surrounding glial-like supporting cells (SCs). Despite structural differences in the auditory organs, molecular mechanisms regulating the development of the auditory organ are evolutionarily conserved between mammals and aves. In ovo electroporation is largely limited to early stages at E1 - E3. Due to the relative late development of the auditory organ at E5, manipulations of the auditory organ by in ovo electroporation past E3 are difficult due to the advanced development of the chicken embryo at later stages. The method presented here is a transient gene transfer method for targeting genes of interest at stage E4 - E4.5 in the developing chicken auditory sensory organ via in ovo micro-electroporation. This method is applicable for gain- and loss-of-functions with conventional plasmid DNA-based expression vectors and can be combined with in ovo cell proliferation assay by adding EdU (5-ethynyl-2&#180;-deoxyuridine) to the whole embryo at the time of electroporation. The use of green or red fluorescent protein (GFP or RFP) expression plasmids allows the experimenter to quickly determine whether the electroporation successfully targeted the auditory portion of the developing inner ear. In this method paper, representative examples of GFP electroporated specimens are illustrated; embryos were harvested 18 - 96 hr&#160;after electroporation and targeting of GFP to the pro-sensory area of the auditory organ was confirmed by RNA in situ hybridization. The method paper also provides an optimized protocol for the use of the thymidine analog EdU to analyze cell proliferation; an example of an EdU based cell proliferation assay that combines immuno-labeling and click EdU chemistry is provided.

Gene Transfer into the Chicken Auditory Organ by In Ovo Micro-electroporation

The auditory organ mediates hearing. Here we present a modified in ovo micro-electroporation method optimized for studying auditory progenitor cell proliferation and differentiation in the developing chicken auditory organ.

Chicken embryos are ideal model systems for studying embryonic development as manipulations of gene function can be conducted with relative ease in ovo. ...

Induction of Acute Skeletal Muscle Regeneration by Cardiotoxin Injection

Skeletal muscle regeneration is a physiological process that occurs in adult skeletal muscles in response to injury or disease. Acute injury-induced skeletal muscle regeneration is a widely used, powerful model system to study the events involved in muscle regeneration as well as the mechanisms and different players. Indeed, a detailed knowledge of this process is essential for a better understanding of the pathological conditions that lead to skeletal muscle degeneration, and it aids in identifying new targeted therapeutic strategies. The present work describes a detailed and reproducible protocol to induce acute skeletal muscle regeneration in mice through a single intramuscular injection of cardiotoxin (CTX). CTX belongs to the family of snake venom toxins and causes myolysis of myofibers, which eventually triggers the regeneration events. The dynamics of skeletal muscle regeneration is evaluated by histological analysis of muscle sections. The protocol also illustrates the experimental procedures for dissecting, freezing, and cutting the Tibialis Anterior muscle, as well as the routine Hematoxylin &#38; Eosin staining that is widely used for subsequent morphological and morphometric analysis.

This manuscript describes a detailed protocol to induce acute skeletal muscle regeneration in adult mice and subsequent manipulations of the muscles, such as dissection, freezing, cutting, routine staining, and myofiber cross-sectional area analysis.

Skeletal muscle regeneration is a physiological process that occurs in adult skeletal muscles in response to injury or disease. Acute injury-induced ...

Immuno-fluorescent Labeling of Microtubules and Centrosomal Proteins in Ex Vivo Intestinal Tissue and 3D In Vitro Intestinal Organoids

The advent of 3D in vitro organoids that mimic the in vivo tissue architecture and morphogenesis has greatly advanced the ability to study key biological questions in cell and developmental biology. In addition, organoids together with recent technical advances in gene editing and viral gene delivery promises to advance medical research and development of new drugs for treatment of diseases. Organoids grown in vitro in basement matrix provide powerful model systems for studying the behavior and function of various proteins and are well suited for live-imaging of fluorescent-tagged proteins. However, establishing the expression and localization of the endogenous proteins in ex vivo tissue and in in vitro organoids is important to verify the behavior of the tagged proteins. To this end we have developed and modified tissue isolation, fixation, and immuno-labeling protocols for localization of microtubules, centrosomal, and associated proteins in ex vivo intestinal tissue and in in vitro intestinal organoids. The aim was for the fixative to preserve the 3D architecture of the organoids/tissue while also preserving antibody antigenicity and enabling good penetration and clearance of fixative and antibodies. Exposure to cold depolymerizes all but stable microtubules and this was a key factor when modifying the various protocols. We found that increasing the ethylenediaminetetraacetic acid (EDTA) concentration from 3 mM to 30 mM gave efficient detachment of villi and crypts in the small intestine while 3 mM EDTA was sufficient for colonic crypts. The developed formaldehyde/methanol fixation protocol gave very good structural preservation while also preserving antigenicity for effective labeling of microtubules, actin, and the end-binding (EB) proteins. It also worked for the centrosomal protein ninein although the methanol protocol worked more consistently. We further established that fixation and immuno-labeling of microtubules and associated proteins could be achieved with organoids isolated from or remaining within the basement matrix.

Immuno-fluorescent Labeling of Microtubules and Centrosomal Proteins in Ex Vivo Intestinal Tissue and 3D In Vitro Intestinal Organoids

We present protocols for isolation of intestinal 3D structures from in vivo tissue and in vitro basement matrix embedded organoids, and detail different fixation and staining protocols optimized for immuno-labeling of microtubule, centrosomal, and junctional proteins as well as cell markers including the stem cell protein Lgr5.

The advent of 3D in vitro organoids that mimic the in vivo tissue architecture and morphogenesis has greatly advanced the ability to study key biological ...

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

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

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

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

Hemogenic Reprogramming of Human Fibroblasts by Enforced Expression of Transcription Factors

The cellular and molecular mechanisms underlying specification of human hematopoietic stem cells (HSCs) remain elusive. Strategies to recapitulate human HSC emergence in vitro are required to overcome limitations in studying this complex developmental process. Here, we describe a protocol to generate hematopoietic stem and progenitor-like cells from human dermal fibroblasts employing a direct cell reprogramming approach. These cells transit through a hemogenic intermediate cell-type, resembling the endothelial-to-hematopoietic transition (EHT) characteristic of HSC specification. Fibroblasts were reprogrammed to hemogenic cells via transduction with GATA2, GFI1B and FOS transcription factors. This combination of three factors induced morphological changes, expression of hemogenic and hematopoietic markers and dynamic EHT transcriptional programs. Reprogrammed cells generate hematopoietic progeny and repopulate immunodeficient mice for three months. This protocol can be adapted towards the mechanistic dissection of the human EHT process as exemplified here by defining GATA2 targets during the early phases of reprogramming. Thus, human hemogenic reprogramming provides a simple and tractable approach to identify novel markers and regulators of human HSC emergence. In the future, faithful induction of hemogenic fate in fibroblasts may lead to the generation of patient-specific HSCs for transplantation.

This protocol demonstrates the induction of a hemogenic program in human dermal fibroblasts by enforced expression of the transcription factors GATA2, GFI1B and FOS to generate hematopoietic stem and progenitor cells.

The cellular and molecular mechanisms underlying specification of human hematopoietic stem cells (HSCs) remain elusive. Strategies to recapitulate human ...

Fluorescent Immunolocalization of Arabinogalactan Proteins and Pectins in the Cell Wall of Plant Tissues

Plant development involves constant adjustments of the cell wall composition and structure in response to both internal and external stimuli. Cell walls are composed of cellulose and non-cellulosic polysaccharides together with proteins, phenolic compounds and water. 90% of the cell wall is composed of&#160;polysaccharides (e.g., pectins) and arabinogalactan proteins (AGPs). The fluorescent immunolocalization of specific glycan epitopes in plant histological sections remains a key tool to uncover remodeling of wall polysaccharide networks, structure and components.
Here, we report an optimized fluorescent immunolocalization procedure to detect glycan epitopes from AGPs and pectins in plant tissues. Paraformaldehyde/glutaraldehyde fixation was used along with LR-White embedding of the plant samples, allowing for a better preservation of the tissue structure and composition. Thin sections of the embedded samples obtained with an ultra-microtome were used for immunolocalization with specific antibodies. This technique offers great resolution, high specificity, and the chance to detect multiple glycan epitopes in the same sample. This technique allows subcellular localization of glycans and detects their level of accumulation in the cell wall. It also permits the determination of spatio-temporal patterns of AGP and pectin distribution during developmental processes. The use of this tool may ultimately guide research directions and link glycans to specific functions in plants. Furthermore, the information obtained can complement biochemical and gene expression studies.

This protocol describes in detail how the plant material for immunolocalization of Arabinogalactan proteins and pectins is fixed, embedded in a hydrophilic acrylic resin, sectioned and mounted on glass slides. We show cell wall related epitopes will be detected with specific antibodies.

Plant development involves constant adjustments of the cell wall composition and structure in response to both internal and external stimuli. Cell walls ...

Minimum Volume Vitrification of Immature Feline Oocytes

In wild animals&#8217; conservation programs, gamete banking is crucial to safeguard genetic resources of valuable individuals and rare species and to promote biodiversity preservation. In felids, most species are threatened with extinction, and domestic breeds are used as a model to increase the efficiency of protocols for germplasm banking. Among oocyte cryopreservation techniques, vitrification is more and more popular in human and veterinary assisted reproduction. Cryotop vitrification, which was at first developed for human oocytes and embryos, has demonstrated to be well-suited for cat oocytes. This method offers several advantages, such as the feasibility in field conditions and the speed of the procedure, which can be helpful when several samples need to be processed. However, the efficiency is strongly dependent on the operator&#8217;s skills, and intra- and inter-laboratory standardization are needed, as well as personnel training. This protocol describes minimum volume vitrification of immature feline oocytes on a commercial support in a step by step field-friendly protocol, from oocyte collection to warming. Following the protocol, preservation of oocyte integrity and viability at warming (as high as 90%) can be expected, although there is still room for improvement in post-warming maturation and embryonic development outcomes.

This manuscript describes a protocol for the minimum volume vitrification of immature cat oocytes with laboratory-made media on commercial supports. It covers every step from oocyte isolation from ex vivo gonads to vitrification and warming.

In wild animals&#8217; conservation programs, gamete banking is crucial to safeguard genetic resources of valuable individuals and rare species and to ...

Separation of Follicular Cells and Oocytes in Ovarian Follicles of Zebrafish

Zebrafish has become an ideal model to study the ovarian development of vertebrates. The follicle is the basic unit of the ovary, which consists of oocytes and surrounding follicular cells. It is vital to separate both follicular cells and oocytes for various research purposes such as for primary culture of follicular cells, analysis of gene expression, oocyte maturation and in vitro fertilization, etc. The conventional method uses forceps to separate both compartments, which is laborious, time consuming and has high damage to the oocyte. Here, we have established a simple method to separate both compartments using a pulled glass capillary. Under a stereomicroscope, oocytes and follicular cells can be easily separated by pipetting in a pulled fine glass capillary (the diameter depends on the follicle diameter). Compared with the conventional method, this new method has high efficiency in separating both oocytes and follicular cells and has low damage to the oocytes. More importantly, this method can be applied to early-stage follicles including at the pre-vitellogenesis stage. Thus, this simple method can be used to separate follicular cells and oocytes of zebrafish.

Here, we present a simple method for separating follicular cells and oocytes in zebrafish ovarian follicles, which will facilitate investigations of ovarian development in zebrafish.

Zebrafish has become an ideal model to study the ovarian development of vertebrates. The follicle is the basic unit of the ovary, which consists of ...

Evaluation of the Spindle Assembly Checkpoint Integrity in Mouse Oocytes

Aneuploidy is the leading genetic abnormality causing early miscarriage and pregnancy failure in humans. Most errors in chromosome segregation that give rise to aneuploidy occur during meiosis in oocytes, but why oocyte meiosis is error-prone is still not fully understood. During cell division, cells prevent errors in chromosome segregation by activating the spindle assembly checkpoint (SAC). This control mechanism relies on detecting kinetochore (KT)-microtubule (MT) attachments and sensing tension generated by spindle fibers. When KTs are unattached, the SAC is activated and prevents cell-cycle progression. The SAC is activated first by MPS1 kinase, which triggers the recruitment and formation of the mitotic checkpoint complex (MCC), composed of MAD1, MAD2, BUB3, and BUBR1. Then, the MCC diffuses into the cytoplasm and sequesters CDC20, an anaphase-promoting complex/cyclosome (APC/C) activator. Once KTs become attached to microtubules and chromosomes are aligned at the metaphase plate, the SAC is silenced, CDC20 is released, and the APC/C is activated, triggering the degradation of Cyclin B and Securin, thereby allowing anaphase onset. Compared to somatic cells, the SAC in oocytes is not as effective because cells can undergo anaphase despite having unattached KTs. Understanding why the SAC is more permissive and if this permissiveness is one of the causes of chromosome segregation errors in oocytes still needs further investigation. The present protocol describes the three techniques to comprehensively evaluate SAC integrity in mouse oocytes. These techniques include using nocodazole to depolymerize MTs to evaluate the SAC response, tracking SAC silencing by following the kinetics of Securin destruction, and evaluating the recruitment of MAD2 to KTs by immunofluorescence. Together these techniques probe mechanisms needed to produce healthy eggs by providing a complete evaluation of SAC integrity.

Error in chromosome segregation is a common feature in oocytes. Therefore, studying the spindle assembly checkpoint gives important clues about the mechanisms needed to produce healthy eggs. The present protocol describes three complementary assays to evaluate spindle assembly checkpoint integrity in mouse oocytes.

Aneuploidy is the leading genetic abnormality causing early miscarriage and pregnancy failure in humans. Most errors in chromosome segregation that give ...