This work was supported by a National Institutes of Health NIDDK grant (K01DK092320) and startup funding from the Department of Pediatrics at the University of Texas McGovern Medical School.

The following protocol has been approved by the University of Texas Health Science Center at Houston&#39;s Center for Laboratory Animal Medicine Animal Welfare Committee, which serves as the Institutional Care and Use Committee (protocol #: HSC-AWC-13-135).
1. Identification and Selection of Blastomeres for Kidney-targeted Injections&#160;
<ol>
	<li>Prior to generating embryos, use the Normal Table of Xenopus Development21 to understand the orientation of the early cell divisions in the embryo. Alternatively, access diagrams of the early developmental stages of Xenopus on Xenbase11.</li>
	<li>Access the interactive Xenopus cell fate maps on Xenbase11 to select which blastomere will be targeted for microinjection.</li>
	<li>Observe that the single cell embryo has a darkly pigmented animal pole and a vegetal pole, which is white and yolky. Note that a protective membrane, known as the vitelline envelope, covers the embryo.</li>
	<li>Notice that the first cleavage typically occurs between the left and right sides of the embryo. These cells contribute equally to the pronephric lineage.</li>
	<li>Note that the second cleavage divides the dorsal and ventral halves of the embryo, leading to a 4-cell embryo. The dorsal cells are smaller and have less pigment than the ventral cells (Figure 2A and Figure 3A).
	<ol>
		<li>Identify the ventral blastomeres (V; the large, dark cells) on the left and right sides, which contribute more to the developing kidney than the dorsal (D; small, light cells) blastomeres (Figure 2A).</li>
		<li>If injecting into a 4-cell embryo, inject the left ventral blastomere to target the left kidney (Figure 3A and Section 3).</li>
	</ol>
	</li>
	<li>The third cleavage bisects the animal and vegetal sides, resulting in an eight-cell embryo. At this point, there are four animal blastomeres [left and right ventral (V1) and dorsal (D1)] and four vegetal blastomeres [left and right ventral (V2) and dorsal (D2)] (Figure 2B and Figure 3B).
	<ol>
		<li>Locate the ventral, vegetal blastomeres (V2). These blastomeres contribute more to the developing kidney any other cells at this stage (Figure 2B). To target the left kidney of an 8-cell embryo, inject into the left V2 blastomere (Figure 3B and Section 3).</li>
	</ol>
	</li>
	<li>Notice that the fourth and fifth cleavages bisect the animal and vegetal blastomeres. Two progeny are generated from each blastomere, resulting in a 16-cell embryo. The cells are named after their predecessor. For example, the V2 blastomere from the 8-cell stage gives rise to V2.1 and V2.2 progeny at the 16-cell stage (Figure 2C). The V2.2 cell at the 16-cell stage provides the majority of the cells contributing to the developing kidney.</li>
	<li>Note the sixth and seventh cleavages result in a 32-cell embryo. Again, two progeny are generated from each blastomere, which are named following their predecessor. For example, the V2.2 blastomere from the 16-cell stage gives rise to V2.2.1 and V2.2.2 at the 32-cell stage. There is an alternative naming system at the 32-cell stage in which cells are identified in four rows as A, B, C, and D (from animal to vegetal), and in four columns as 1, 2, 3, and 4 (from dorsal to ventral). Thus, the V2.2.2 blastomere, which contributes the most&#160;to the developing pronephros, is called C3 under this alternative naming system (Figure 2D).</li>
</ol>
2. Preparation of Embryos
<ol>
	<li>Prepare 50 ml of Dejelly Solution (2% cysteine, NaOH to pH 8.0).</li>
	<li>Isolate both testes from a single male frog according to standard protocols14. Place the testes in a 60 mm Petri dish filled with 10 ml Testes Storage Solution (1x Marc&#39;s Modified Ringers [MMR; 0.1 M NaCl, 2 mM KCl, 1 mM MgSO4, 2 mM CaCl2, 5 mM HEPES pH 8, 0.1 mM EDTA]12, 1% bovine serum albumin, 50 &#181;g/ml gentamycin). Store the testes at 4 &#176;C. 
	Note: Testes can be stored for approximately 7 - 10 days at 4 &#176;C, but fertilization efficiency will decrease the longer the testes have been stored.</li>
	<li>Squeeze a female frog to obtain eggs according to standard protocols14. Collect the eggs in a 100 mm Petri dish. Pour off any excess water.</li>
	<li>Cut off &#188; of a testis while it is in Testes Storage Solution using forceps and a razor blade. Transfer the piece of testis to the Petri dish containing eggs. Adjust the size of the testis portion used to account for the size of the testis, how long the testis has been stored, and how many eggs are to be fertilized. Generally use &#188; to 1/3 of a freshly dissected testis to fertilize one clutch of eggs.</li>
	<li>Cut the testis portion into small pieces using forceps and a razor blade. Add enough 0.3x&#160;MMR + 30 mg/ml gentamycin to the Petri dish to cover the eggs. Swirl the MMR in the dish to mix.</li>
	<li>Wait approximately 30 min for fertilization to take place at room temperature. Note that the animal hemisphere (the pigmented side of the embryo) will sit on top of the embryo upon effective fertilization. Then, remove the MMR from the Petri dish using a transfer pipette. Add enough Dejelly Solution to the dish to cover the embryos.</li>
	<li>Over the next few minutes, gently swirl the dish intermittently. Vigorous shaking of the dish at this time can cause axis defects.&#160;The jelly coat on the embryos will dissolve, and the embryos will congregate in the center of the dish during swirling. Once the embryos are closely touching each other in the center of the dish, remove the Dejelly Solution with a transfer pipette. Do not leave embryos in the Dejelly Solution for longer than 5 minutes, or the embryos may be damaged.</li>
	<li>Wash the dejellied embryos 3 - 5 times in 0.3x MMR + 30 mg/ml gentamycin by carefully pouring or pipetting off the MMR and filling the dish with new MMR. Do not remove all of the MMR from the dish, or the embryos may be damaged.</li>
	<li>Remove any unfertilized eggs or pieces of testis from the Petri dish using a transfer pipette.</li>
	<li>Incubate embryos between 14 and 22 &#176;C. 
	Note: Embryos grown at lower temperatures develop more slowly than embryos grown at higher temperatures. Timing of developmental stages can be found on Xenbase22.
	<ol>
		<li>To space out their development, place half of the embryos from a single fertilization in a Petri dish kept at 14 &#176;C, and the other half of the embryos in a Petri dish kept at 18 &#176;C. This allows for two sets of injections into 4-cell or 8-cell embryos from a single fertilization.</li>
	</ol>
	</li>
</ol>
3. Preparation of Injection Solutions and Microinjection of Embryos
<ol>
	<li>Prepare the injection solution containing 0.01 ng/nl membrane-bound red fluorescent protein (MEM-RFP) mRNA9 while the embryos are developing to the 4-cell or 8-cell stage. Store the injection solution on ice until ready to inject.</li>
	<li>Load a 7&#34; replacement glass capillary tube into a needle puller, with the top of the replacement tube aligned with the top of the needle puller case. Set the heat #2 value to 800, and the pull value to 650. Press the &#34;pull&#34; button to pull the needle. This will create 2 needles from a single 7&#34; glass capillary tube.</li>
	<li>Snip off the tip of a pulled needle with a pair of Dumont forceps. 
	Note: After pulling the needle, the tip is sealed shut and must be cut open. The closer to the point of the needle that it is cut, the smaller the diameter the needle tip will be. Although the diameter of the tip will not affect the injection volume with the microinjection system used here, a tip with a larger diameter is more likely to damage the embryo.</li>
	<li>Slip the micropipette collet onto the back of the needle. Next, slip the large hole O-ring onto the back of the needle behind the collet.</li>
	<li>Fill the needle with mineral oil using a 27 gauge hypodermic needle, being careful not to get air bubbles in the needle.</li>
	<li>Slip the needle onto the plunger of the microinjector, seating the needle into the large hole of the white plastic spacer installed on the plunger. The plunger should have a small O-ring nearest to the body of the microinjector, followed by the white spacer, large hole O-ring, and collet. Secure the needle by tightening the collet. Pull gently on the needle to make sure that it is properly secured.</li>
	<li>Press and hold the &#34;empty&#34; button on the microinjector control box until there are two beeps.</li>
	<li>Pipette 3 &#181;l&#160;of injection solution onto a piece of Parafilm. Insert the tip of the needle into the bead of injection solution on the Parafilm. Press and hold the &#34;fill&#34; button on the microinjector control box to draw the injection solution into the needle.</li>
	<li>Fill a 60 mm Petri dish lined with 500 micron polyester mesh with 5% Ficoll in 0.3x MMR + 30 mg/ml gentamycin. Carefully pipette 20 - 30 4-cell or 8-cell embryos into the dish.</li>
	<li>Using a hair loop14, manipulate the embryos so that the blastomere to be injected is facing the needle. To target the left kidney, line up the embryos so that the left ventral blastomeres of 4-cell embryos or the left V2 blastomeres of 8-cell embryos face the needle.</li>
	<li>Inject 10 nl of injection solution into the selected blastomere of each embryo in the dish. 
	Note: The mesh at the bottom of the Petri dish stabilizes the embryos and prevents them from rolling, allowing them to be injected without the use of a hair loop for stabilization.</li>
	<li>Transfer injected embryos into wells of a culture plate that have been filled with 5% Ficoll in 0.3x MMR + 30 mg/ml gentamycin. Incubate the injected embryos at 16 &#176;C for at least one hour to allow the injected blastomeres to heal.</li>
	<li>Transfer the healed embryos into wells of a new culture plate that have been filled with 0.3x MMR + 30 mg/ml gentamycin by stage 9 (prior to gastrulation).</li>
	<li>Incubate the embryos at 14 - 22 &#176;C until the embryos reach stage 38 - 4021.</li>
</ol>
4. Fixation and Immunostaining of Embryos
<ol>
	<li>Prepare 50 ml of MOPS/EGTA/Magnesium sulfate/Formaldehyde Buffer [MEMFA: 100 mM MOPS (pH 7.4), 2 mM EGTA, 1 mM MgSO4, 3.7% (v/v) formaldehyde].</li>
	<li>Using a transfer pipette, put 10 - 20 stage 38 - 40 embryos in a glass vial. Add 10 &#181;l 5% benzocaine in 100% ethanol to the vial and invert vial to mix. Wait 10 min to anesthetize the embryos.</li>
	<li>Remove the MMR from the vial using a glass pipette. If processing multiple vials at the same time, the vials can be held upright in a 24-well cell culture plate.</li>
	<li>With a glass pipette, fill the vial with MEMFA. Place the vial on a three-dimensional rocking platform for 1 hr at room temperature.</li>
	<li>Remove the MEMFA from the vial using a glass pipette. Fill the vial with 100% methanol. Place the vial on a three-dimensional rocking platform for 10 min at room temperature. Repeat this wash step one more time, and store the embryos in 100% methanol overnight at -20 &#176;C.</li>
	<li>Prepare 1x Phosphate Buffered Saline-Bovine Serum Albumin-Triton (PBT): 1x PBS, 2 mg/ml bovine serum albumen, 0.1% Triton X-100.</li>
	<li>Prepare the primary antibody solution: 1x PBT with 10% goat serum with a 1:5 dilution of mouse monoclonal 4A6 antibody (to label the membranes of the intermediate, distal and connecting tubules20), a 1:30 dilution of mouse monoclonal antibody 3G8 (to label lumen of the proximal tubules20), and a 1:250 dilution of rabbit polyclonal RFP antibody (to label the MEM-RFP tracer). Store at 4 &#176;C.</li>
	<li>Prepare the secondary antibody solution: 1x PBT with 10% goat serum, 1:500 Alexa 488 goat anti-mouse IgG (stock concentration 2 mg/ml; to label 4A6 and 3G8), and 1:500 Alexa 555 goat anti-rabbit IgG (stock concentration 2 mg/ml; to label the MEM-RFP tracer). Store at 4 &#176;C, covering the tube in foil to protect from light.</li>
	<li>Alternatively, collect the primary and secondary antibodies after staining, and save at 4 &#176;C for reuse in future experiments. If the antibody is to be saved for reuse, add 0.01% sodium azide.</li>
	<li>Immunostain the embryos using established protocols18.</li>
</ol>
5. Visualization of Embryos and Analysis of Targeted Pronephric Tissue&#160;
<ol>
	<li>Screen the immunostained embryos to verify that the correct blastomere was injected by viewing the fluorescence of the tracer under a fluorescent stereomicroscope at 1X (to view whole embryo) and 5X (to view kidney) magnification. Place embryos in a multi-well glass plate with the wells filled with 1x PBT using a transfer pipette with the tip cut off. Manipulate the embryos with a hair loop. Use only embryos which have the co-injected tracer present in the pronephros on the left side of the embryo (Figure 3C, F and Figure 4C, F) for gene overexpression or knockdown analysis.</li>
	<li>Alternatively, clear the embryos in Murray&#39;s Clear (2 parts benzyl benzoate: 1 part benzyl alcohol) by placing the embryos in a glass vial and filling the vial with Murray&#39;s Clear. Visualize the embryos using a glass well plate. 
	Note: Murray&#39;s Clear is an organic solvent, and should be handled with care. Wear gloves, and only use glass vials and pipettes with Murray&#39;s Clear.</li>
	<li>Store embryos at 4 &#176;C in 1x PBT for 2 - 3 weeks. For long-term storage of embryos, dehydrate the embryos by washing two times in 100% methanol at room temperature for 10 min. Store the embryos at -20 &#176;C in 100% methanol.</li>
</ol>

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Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blastomeres+show+differential+fate+changes+in+8-cell+Xenopus+laevis+embryos+that+are+rotated+90+degrees+before+the+first+cleavage.">Blastomeres show differential fate changes in 8-cell Xenopus laevis embryos that are rotated 90 degrees before the first cleavage.</a> Dev. Growth Differ. 40 (2), 189-198 (1998).</li><li>Colman, A., Drummond, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+stability+and+movement+of+mRNA+in+Xenopus+oocytes+and+embryos.">The stability and movement of mRNA in Xenopus oocytes and embryos.</a> J. Embryol. Exp. Morph. 97, 197-209 (1986).</li></ol>

The authors declare that they have no competing financial interests.

Targeting the pronephros of developing Xenopus embryos relies on identifying and injecting the correct blastomere. Injection of the V2 blastomere of 8-cell embryos targets the left pronephros18. This leaves the contralateral right pronephros as an internal control. If morpholino knockdown or RNA overexpression is used to alter kidney development, the contralateral right pronephros can be used to compare the effects of gene knockdown or overexpression on the left pronephros. In this case, proper contro...

The Xenopus embryonic kidney, the pronephros, is a good model for studying kidney development and disease. The embryos develop externally, are large in size, can be produced in large numbers, and are easily manipulated through microinjection or surgical procedures. In addition, the genes governing kidney development in mammals and amphibians are conserved. Mammalian kidneys progress through three stages: the pronephros, mesonephros, and metanephros1, while embryonic amphibians have a pronephros and adult amphibians have a metanephros. The basic filtering unit of these kidney forms is the nephron, and both mammals and amphibians require the same signaling cascades and inductive events to undergo nephrogenesis2, 3. The Xenopus pronephros contains a single nephron composed of proximal, intermediate, distal and connecting tubules, and&#160;a glomus (analogous to the mammalian glomerulus)1, 4-6 (Figure 1). The single, large nephron present in the Xenopus pronephros makes it suitable as a simple model for the study of genes involved in kidney development and disease processes.
Cell fate maps have been established for early Xenopus embryos, and are freely available online at Xenbase7-11. Here, we describe a technique for microinjection of lineage tracers to target the developing Xenopus pronephros, although the same technique can be adapted to target other tissues such as the heart or eyes. Lineage tracers are labels (including vital dyes, fluorescently labeled dextrans, histochemically detectable enzymes, and mRNA encoding fluorescent proteins) that can be injected into an early blastomere, allowing the visualization of the progeny of that cell during development. This protocol utilizes MEM-RFP mRNA, encoding membrane targeted red fluorescent protein12, as a lineage tracer. The targeted microinjection techniques for individual blastomeres in 4- and 8-cell embryos described here can be utilized for injection with morpholinos to knock down gene expression, or with exogenous RNA to overexpress a gene of interest. By injecting into the ventral, vegetal blastomere, primarily the pronephros of the embryo will be targeted, leaving the contralateral pronephros as a developmental control. Co-injection of a tracer verifies that the correct blastomere was injected, and shows which tissues in the embryo arose from the injected blastomere, verifying targeting of the pronephros. Immunostaining of the pronephros allows visualization of how well the pronephric tubules have been targeted. Overexpression and knockdown effects can then be scored against the contralateral side of the embryo, which serves as a developmental control, and can be used to calculate the pronephric index13. The availability of cell fate maps allows this targeted microinjection technique to be used to target tissues other than the pronephros, and co-injection of a fluorescent tracer allows the targeted microinjection to each tissue to be verified prior to analysis.
During embryo microinjection, developmental temperature should be regulated tightly, given that the rate of Xenopus development is highly dependent upon it14. Embryos should be incubated at cooler temperatures (14 - 16 &#176;C) for 4- and 8-cell injections because the development time is slowed down. At 22 &#176;C, development time from stage 1 (1 cell) to stage 3 (4 cells) is approximately 2 hours, while at 16 &#176;C development time to stage 3 is approximately 4 hours. It takes approximately 15 minutes to go from a 4-cell embryo to an 8-cell (stage 4) embryo at 22 &#176;C, but takes approximately 30 minutes at 16 &#176;C. Similarly, at 22 &#176;C, it only takes 30 minutes for an 8-cell embryo to progress to a 16-cell embryo (stage 5). This time is increased to 45 minutes at 16 &#176;C. Therefore, it is useful to slow the development rate of the embryos to enable enough time for injections at the 8-cell stage before the embryos progress to the 16-cell stage. Additionally, growth temperatures can be modulated to speed or slow embryonic development until the kidney has fully developed.
The epidermis of tadpole-stage Xenopus embryos is relatively transparent, allowing for easy imaging of the developing pronephros without dissection or clearing of the tissue15. Due to the relative transparency of Xenopus embryos, live cell imaging is also feasible16,17. Whole-mount immunostaining to visualize the pronephros is possible with established antibodies that label the proximal, intermediate, distal and connecting tubules of stage 38 - 40 embryos that allow for assessment of pronephric development after targeted manipulation of gene expression in Xenopus embryos18-20.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sodium chloride</td>
			<td>Fisher</td>
			<td>S271-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Fisher</td>
			<td>P217-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium sulfate&#160;</td>
			<td>Fisher</td>
			<td>M63-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride</td>
			<td>Fisher</td>
			<td>C79-500</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Fisher</td>
			<td>BP310-500</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Fisher</td>
			<td>S311-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Gentamycin solution, 50 mg/mL</td>
			<td>Amresco</td>
			<td>E737-20ML</td>
			<td></td>
		</tr>
		<tr>
			<td>L-Cysteine, 99%+</td>
			<td>Acros Organics</td>
			<td>52-90-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Ficoll</td>
			<td>Fisher</td>
			<td>BP525-100</td>
			<td></td>
		</tr>
		<tr>
			<td>100 mm x 15 mm Petri dish</td>
			<td>Fisher</td>
			<td>FB0875712</td>
			<td></td>
		</tr>
		<tr>
			<td>Mini Fridge II</td>
			<td>Boekel</td>
			<td>260009</td>
			<td>Benchtop incubator for embryos.</td>
		</tr>
		<tr>
			<td>Gap43-RFP plasmid</td>
			<td></td>
			<td></td>
			<td>For making tracer RNA. Ref: Davidson et al., 2006.</td>
		</tr>
		<tr>
			<td>Rhodamine dextran, 10,000 M.W.</td>
			<td>Invitrogen</td>
			<td>D1817</td>
			<td>Tracer.</td>
		</tr>
		<tr>
			<td>Molecular biology grade, USP sterile purified water</td>
			<td>Corning</td>
			<td>46-000-CI</td>
			<td></td>
		</tr>
		<tr>
			<td>7&#34; Drummond replacement tubes</td>
			<td>Drummond</td>
			<td>3-000-203-G/XL</td>
			<td>Microinjection needles.</td>
		</tr>
		<tr>
			<td>Needle puller</td>
			<td>Sutter Instruments</td>
			<td>P-30</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine forceps</td>
			<td>Dumont</td>
			<td>11252-30</td>
			<td>For breaking microinjection needle tip.</td>
		</tr>
		<tr>
			<td>Nanoject II</td>
			<td>Drummond</td>
			<td>3-000-204</td>
			<td>Microinjector.</td>
		</tr>
		<tr>
			<td>Mineral oil, heavy</td>
			<td>Fisher</td>
			<td>CAS 8042-47-5</td>
			<td>Oil for needles.</td>
		</tr>
		<tr>
			<td>27G monoject hypodermic needle</td>
			<td>Covidien</td>
			<td>8881200508</td>
			<td>For loading mineral oil into microinjection needle.</td>
		</tr>
		<tr>
			<td>5 mL Luer-Lok syringe</td>
			<td>BD</td>
			<td>309646</td>
			<td>For loading mineral oil into microinjection needle.</td>
		</tr>
		<tr>
			<td>60 mm x 15 mm Petri dish</td>
			<td>Fisher</td>
			<td>FB0875713A</td>
			<td></td>
		</tr>
		<tr>
			<td>800 micron polyester mesh</td>
			<td>Small Parts</td>
			<td>CMY-0800-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Transfer pipets</td>
			<td>Fisher</td>
			<td>13-711-7M</td>
			<td>For transferring embryos. Cut off the tip so that the embryos are easily taken up by the pipette.</td>
		</tr>
		<tr>
			<td>6-well cell culture plate</td>
			<td>Nest Biotechnology</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MOPS</td>
			<td>Fisher</td>
			<td>BP308-500</td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Acros Organics</td>
			<td>67-42-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Formaldehyde</td>
			<td>Fisher</td>
			<td>BP531-500</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mm x 45 mm screw thread vial</td>
			<td>Fisher</td>
			<td>03-339-25B</td>
			<td>For fixing, staining, and storing embryos.</td>
		</tr>
		<tr>
			<td>24-well cell culture plate</td>
			<td>Nest Biotechnology</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Benzocaine</td>
			<td>Spectrum</td>
			<td>BE130</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Fisher</td>
			<td>BP2818-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Fisher</td>
			<td>A412-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (PBS) 1X powder</td>
			<td>Fisher</td>
			<td>BP661-50</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine serum albumen</td>
			<td>Fisher</td>
			<td>BP1600-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Fisher</td>
			<td>BP151-500</td>
			<td></td>
		</tr>
		<tr>
			<td>3D mini rocker, model 135</td>
			<td>Denville Scientific</td>
			<td>57281</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat serum, New Zealand origin</td>
			<td>Invitrogen</td>
			<td>16210064</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium azide</td>
			<td>Fisher</td>
			<td>S227I-25</td>
			<td></td>
		</tr>
		<tr>
			<td>Monoclonal 3G8 antibody</td>
			<td>European Xenopus Resource Centre</td>
			<td></td>
			<td>Primary antibody to label proximal tubules.</td>
		</tr>
		<tr>
			<td>Monoclonal 4A6 antibody</td>
			<td>European Xenopus Resource Centre</td>
			<td></td>
			<td>Primary antibody to label distal tubule and duct.</td>
		</tr>
		<tr>
			<td>anti-RFP pAb, purified IgG/rabbit</td>
			<td>MBL</td>
			<td>PM005</td>
			<td>Primary antibody to label Gap43-RFP tracer.</td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 goat anti-mouse IgG</td>
			<td>Life Technologies</td>
			<td>A11001</td>
			<td>Secondary antibody to label distal tubule and duct.</td>
		</tr>
		<tr>
			<td>Alexa Fluor 555 goat anti-rabbit IgG</td>
			<td>Life Technologies</td>
			<td>A21428</td>
			<td>Secondary antibody to label Gap43-RFP tracer.</td>
		</tr>
		<tr>
			<td>9 cavity spot plate</td>
			<td>Corning</td>
			<td>7220-85</td>
			<td></td>
		</tr>
		<tr>
			<td>Benzyl benzoate</td>
			<td>Fisher</td>
			<td>105862500</td>
			<td>Optional - for clearing embryos</td>
		</tr>
		<tr>
			<td>Benzyl alcohol</td>
			<td>Fisher</td>
			<td>A396-500</td>
			<td>Optional - for clearing embryos</td>
		</tr>
	</tbody>
</table>

technique to target microinjection to the developing xenopus kidney

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus embryos are large, develop externally, and can be easily manipulated by microinjection or surgical procedures. In addition, fate maps have been established for early Xenopus embryos. Targeted microinjection into the individual blastomere that will eventually give rise to an organ or tissue of interest can be used to selectively overexpress or knock down gene expression within this restricted region, decreasing secondary effects in the rest of the developing embryo. In this protocol, we describe how to utilize established Xenopus fate maps to target the developing Xenopus kidney (the pronephros), through microinjection into specific blastomere of 4- and 8-cell embryos. Injection of lineage tracers allows verification of the specific targeting of the injection. After embryos have developed to stage 38 - 40, whole-mount immunostaining is used to visualize pronephric development, and the contribution by targeted cells to the pronephros can be assessed. The same technique can be adapted to target other tissue types in addition to the pronephros.

Microinjections of 4- and 8-cell Xenopus embryos with MEM-RFP mRNA show different levels of targeting to the pronephros. Figure 4 shows stage 40 embryos with correct MEM-RFP mRNA expression patterns. Embryos were injected in the left ventral blastomere (Figure 4A), and sorted for the proper expression pattern of MEM-RFP mRNA. In addition to expressing MEM-RFP in the proximal, intermediate, distal and connecting tubules of the kidney, properly inj...

Watch this Scientific Journal Video about Technique to Target Microinjection to the Developing Xenopus Kidney at JoVE.com

Technique to Target Microinjection to the Developing Xenopus Kidney

Here, we present a protocol to use fate maps and lineage tracers to target injections into individual blastomeres that give rise to the kidney of Xenopus laevis embryos.

Technique to Target Microinjection to the Developing Xenopus Kidney

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus ...

technique-to-target-microinjection-to-the-developing-xenopus-kidney

Research

JoVE Journal

Developmental Biology

11.1K Views.  University of Texas McGovern Medical School. Here, we present a protocol to use fate maps and lineage tracers to target injections into individual blastomeres that give rise to the kidney of Xenopus laevis embryos. 

Video: Technique to Target Microinjection to the Developing Xenopus Kidney

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 ...

Using Confocal Analysis of Xenopus laevis to Investigate Modulators of Wnt and Shh Morphogen Gradients

This protocol describes a method to visualise ligands distributed across a field of cells. The ease of expressing exogenous proteins, together with the large size of their cells in early embryos, make Xenopus laevis a useful model for visualising GFP-tagged ligands. Synthetic mRNAs are efficiently translated after injection into early stage Xenopus embryos, and injections can be targeted to a single cell. When combined with a lineage tracer such as membrane tethered RFP, the injected cell (and its descendants) that are producing the overexpressed protein can easily be followed. This protocol describes a method for the production of fluorescently tagged Wnt and Shh ligands from injected mRNA. The methods involve the micro dissection of ectodermal explants (animal caps) and the analysis of ligand diffusion in multiple samples. By using confocal imaging, information about ligand secretion and diffusion over a field of cells can be obtained. Statistical analyses of confocal images provide quantitative data on the shape of ligand gradients. These methods may be useful to researchers who want to test the effects of factors that may regulate the shape of morphogen gradients.

Using Confocal Analysis of Xenopus laevis to Investigate Modulators of Wnt and Shh Morphogen Gradients

The manuscript here provides a simple set of methods for analysing the secretion and diffusion of fluorescently tagged ligands in Xenopus. This provides a context for testing the ability of other proteins to modify ligand distribution and allowing experiments that may give insight into mechanisms regulating morphogen gradients.

This protocol describes a method to visualise ligands distributed across a field of cells. The ease of expressing exogenous proteins, together with the ...

Grafting of Beads into Developing Chicken Embryo Limbs to Identify Signal Transduction Pathways Affecting Gene Expression

Using chicken embryos it is possible to test directly the effects of either growth factors or specific inhibitors of signaling pathways on gene expression and activation of signal transduction pathways. This technique allows the delivery of signaling molecules at precisely defined developmental stages for specific times. After this embryos can be harvested and gene expression examined, for example by in situ hybridization, or activation of signal transduction pathways observed with immunostaining.
In this video heparin beads soaked in FGF18 or AG 1-X2 beads soaked in U0126, a MEK inhibitor, are grafted into the limb bud in ovo. This shows that FGF18 induces expression of MyoD and ERK phosphorylation and both endogenous and FGF18 induced MyoD expression is inhibited by U0126. Beads soaked in a retinoic acid antagonist can potentiate premature MyoD induction by FGF18.
This approach can be used with a wide range of different growth factors and inhibitors and is easily adapted to other tissues in the developing embryo.

By grafting beads soaked in growth factors or specific inhibitors of signaling pathways into developing embryos it is possible to directly test their effects in vivo. In this protocol beads are grafted into the limb bud to determine the effects of these molecules on gene expression and signal transduction.

Using chicken embryos it is possible to test directly the effects of either growth factors or specific inhibitors of signaling pathways on gene expression ...

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, cells from the roof of the blastocoel are pluripotent. These cells can be isolated, and programmed to generate various tissues through manipulation of genes expression or induction by morphogens. In this manuscript protocols are described for the use of Xenopus laevis blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development. These protocols allow the investigation of fate acquisition, cell migration behaviors, and cell autonomous and non-autonomous properties. The blastocoel roof explants can be cultured in a serum-free defined medium and grafted into host embryos. This transplantation into an embryo allows the investigation of the long-term lineage commitment, the inductive properties, and the behavior of transplanted cells in vivo. These assays can be exploited to investigate molecular mechanisms, cellular processes and gene regulatory networks underlying neural development. In the context of regenerative medicine, these assays provide a means to generate neural-derived cell types in vitro that could be used in drug screening.

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

In Xenopus embryos, cells from the roof of the blastocoel are pluripotent and can be programmed to generate various tissues. Here, we describe protocols to use amphibian blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development.

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, ...

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

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

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

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

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

A 5-mC Dot Blot Assay Quantifying the DNA Methylation Level of Chondrocyte Dedifferentiation In Vitro

The dedifferentiation of hyaline chondrocytes into fibroblastic chondrocytes often accompanies monolayer expansion of chondrocytes in vitro. The global DNA methylation level of chondrocytes is considered to be a suitable biomarker for the loss of the chondrocyte phenotype. However, results based on different experimental methods can be inconsistent. Therefore, it is important to establish a precise, simple, and rapid method to quantify global DNA methylation levels during chondrocyte dedifferentiation.
Current genome-wide methylation analysis techniques largely rely on bisulfite genomic sequencing. Due to DNA degradation during bisulfite conversion, these methods typically require a large sample volume. Other methods used to quantify global DNA methylation levels include high-performance liquid chromatography (HPLC). However, HPLC requires complete digestion of genomic DNA. Additionally, the prohibitively high cost of HPLC instruments limits HPLC&#39;s wider application.
In this study, genomic DNA (gDNA) was extracted from human chondrocytes cultured with varying number of passages. The gDNA methylation level was detected using a methylation-specific dot blot assay. In this dot blot approach, a gDNA mixture containing the methylated DNA to be detected was spotted directly onto an N+ membrane as a dot inside a previously drawn circular template pattern. Compared with other gel electrophoresis-based blotting approaches and other complex blotting procedures, the dot blot method saves significant time. In addition, dot blots can detect overall DNA methylation level using a commercially available 5-mC antibody. We found that the DNA methylation level differed between the monolayer subcultures, and therefore could play a key role in chondrocyte dedifferentiation. The 5-mC dot blot is a reliable, simple, and rapid method to detect the general DNA methylation level to evaluate chondrocyte phenotype.

A 5-mC Dot Blot Assay Quantifying the DNA Methylation Level of Chondrocyte Dedifferentiation In Vitro

We present a method to quantify DNA methylation based on the 5-methylcytosine (5-mC) dot blot. We determined the 5-mC levels during chondrocyte dedifferentiation. This simple technique could be used to quickly determine the chondrocyte phenotype in ACI treatment.

The dedifferentiation of hyaline chondrocytes into fibroblastic chondrocytes often accompanies monolayer expansion of chondrocytes in vitro. The global ...

Precise Cellular Ablation Approach for Modeling Acute Kidney Injury in Developing Zebrafish

Acute Kidney Injury (AKI) is a common medical condition with a high mortality rate. With the repair abilities of the kidney, it is possible to restore adequate kidney function after supportive treatment. However, a better understanding of how nephron cell death and repair occur on the cellular level is required to minimize cell death and to enhance the regenerative process. The zebrafish pronephros is a good model system to accomplish this goal because it contains anatomical segments that are similar to the mammalian nephron. Previously, the most common model used to study kidney injury in fish was the pharmacological gentamicin model. However, this model does not allow for precise spatiotemporal control of injury, and hence it is difficult to study cellular and molecular processes involved in kidney repair. To overcome this limitation, this work presents a method through which, in contrast to the gentamicin approach, a specific Green Fuorescent Protein (GFP)-expressing nephron segment can be photoablated using a violet laser light (405 nm). This novel model of AKI provides many advantages that other methods of epithelial injury lack. Its main advantages are the ability to &#34;dial&#34; the level of injury and the precise spatiotemporal control in the robust in vivo animal model. This new method has the potential to significantly advance the level of understanding of kidney injury and repair mechanisms.

This work presents a new in vivo model of segmental kidney injury using kidney GFP transgenic zebrafish. The model allows for the induction of the targeted ablation of kidney epithelial cells to show the cellular mechanisms of nephron injury and repair.

Acute Kidney Injury (AKI) is a common medical condition with a high mortality rate. With the repair abilities of the kidney, it is possible to restore ...

Generation of Genetically Modified Mice through the Microinjection of Oocytes

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear injection of DNA expression constructs into fertilized oocytes remains the most commonly used technique to generate transgenic mice for overexpression. With the introduction of CRISPR technology for gene targeting, pronuclear injection into fertilized oocytes has been extended to the generation of both knockout and knockin mice. This work describes the preparation of DNA for injection and the generation of CRISPR guides for gene targeting, with a particular emphasis on quality control. The genotyping procedures required for the identification of potential founders are critical. Innovative genotyping strategies that take advantage of the "multiplexing" capabilities of CRISPR are presented herein. Surgical procedures are also outlined. Together, the steps of the protocol will allow for the generation of genetically modified mice and for the subsequent establishment of mouse colonies for a plethora of research fields, including immunology, neuroscience, cancer, physiology, development, and others.

The microinjection of mouse oocytes is commonly used for both classic transgenesis (i.e., the random integration of transgenes) and CRISPR-mediated gene targeting. This protocol reviews the latest developments in microinjection, with a particular emphasis on quality control and genotyping strategies.

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear ...

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C (UV-C) Damage

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational health hazard. Here, we demonstrate the exposure of primary stem cells from the mouse ocular surface to UV-C radiation. Reactive oxygen species (ROS) formation is the readout of the extent of oxidative stress/damage. In an experimental in vitro setting, it is also essential to assess the percentage of dead cells generated due to oxidative stress. In this article, we will demonstrate the 2&#39;,7&#39;-Dichlorofluoresceindiacetate (DCFDA) staining of UV-C exposed mouse primary ocular surface stem cells and their quantification based on the fluorescent images of DCFDA staining. DCFDA staining directly corresponds to ROS generation. We also demonstrate the quantification of dead and live cells by simultaneous staining with propidium iodide (PI) and Hoechst 3332 respectively and the percentage of DCFDA (ROS positive) and PI positive cells.

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C UV-C Damage

This protocol demonstrates the simultaneous detection of reactive oxygen species (ROS), live cells, and dead cells in live primary cultures from mouse ocular surface cells. 2&#39;,7&#39;-Dichlorofluoresceindiacetate, propidium iodide, and Hoechst staining are used to assess the ROS, dead cells, and live cells, respectively, followed by imaging and analysis.

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational ...

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

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

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

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

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