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><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&amp;nbsp;</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 &lt;em&gt;et al.&lt;/em&gt;, 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&quot; 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>27 G 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 intermediate, distal and connecting tubules.</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 kidney tubules.</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

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Developmental Biology

11.2K 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

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 ...

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The efficient in vivo visualization of blood vessels and the reliable quantification of their complexity are key to understanding the biology and disease of the vascular network. Here, we provide a detailed method to visualize blood vessels with a commercially available fluorescent dye, human plasma acetylated low density lipoprotein DiI complex (DiI-AcLDL), and to quantify their complexity in Xenopus tropicalis. Blood vessels can be labeled by a simple injection of DiI-AcLDL into the beating heart of an embryo, and blood vessels in the entire embryo can be imaged in live or fixed embryos. Combined with gene perturbation by the targeted microinjection of nucleic acids and/or the bath application of pharmacological reagents, the roles of a gene or of a signaling pathway on vascular development can be investigated within one week without resorting to sophisticated genetically engineered animals. Because of the well-defined venous system of Xenopus and its stereotypic angiogenesis, the sprouting of pre-existing vessels, vessel complexity can be quantified efficiently after perturbation experiments. This relatively simple protocol should serve as an easily accessible tool in diverse fields of cardiovascular research.

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

This protocol demonstrates a fluorescence-based method to visualize the vasculature and to quantify its complexity in Xenopus tropicalis. Blood vessels can be imaged minutes after the injection of a fluorescent dye into the beating heart of an embryo after genetic and/or pharmacological manipulations to study cardiovascular development in vivo.

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The ...

Microinjection of Xenopus Laevis Oocytes

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic transport. Because of its large nucleus and high density of nuclear pore complexes (NPCs), nuclear transport can be easily visualized in the Xenopus oocyte. Much insight into the mechanisms of nuclear import and export has been gained through use of this system (reviewed by Pant&eacute;, 2006). In addition, we have used microinjection of Xenopus oocytes to dissect the nuclear import pathways of several viruses that replicate in the host nucleus. 
Here we demonstrate the cytoplasmic microinjection of Xenopus oocytes with a nuclear import substrate. We also show preparation of the injected oocytes for visualization by thin-sectioning EM, including dissection, dehydration, and embedding of the oocytes into an epoxy embedding resin. Finally, we provide representative results for oocytes that have been microinjected with the capsid of the baculovirus Autographa californica nucleopolyhedrovirus (AcMNPV) or the parvovirus Minute Virus of Mice (MVM), and discuss potential applications of the technique.

Here we demonstrate cytoplasmic microinjection of Xenopus laevis oocytes with a nuclear import substrate, as well as preparation of the injected oocytes for visualization by thin-sectioning electron microscopy.

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic ...

Biology

Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos.

An essential tool for investigating the role of a gene during development is the ability to perform gene knockdown, overexpression, and misexpression studies. In zebrafish (Danio rerio), microinjection of RNA, DNA, proteins, antisense oligonucleotides and other small molecules into the developing embryo provides researchers a quick and robust assay for exploring gene function in vivo. In this video-article, we will demonstrate how to prepare and microinject in vitro synthesized EGFP mRNA and a translational-blocking morpholino oligo against pkd2, a gene associated with autosomal dominant polycystic kidney disease (ADPKD), into 1-cell stage zebrafish embryos. We will then analyze the success of the mRNA and morpholino microinjections by verifying GFP expression and phenotype analysis. Broad applications of this technique include generating transgenic animals and germ-line chimeras, cell-fate mapping and gene screening. Herein we describe a protocol for overexpression of EGFP and knockdown of pkd2 by mRNA and morpholino oligonucleotide injection.

Microinjection is a well-established and effective method for introducing foreign substances into fertilized zebrafish embryos. Here, we demonstrate a robust microinjection technique for performing mRNA overexpression, and morpholino oligonucleotide gene knockdown studies in zebrafish.

An essential tool for investigating the role of a gene during development is the ability to perform gene knockdown, overexpression, and misexpression ...

Neural Explant Cultures from Xenopus laevis

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During axon outgrowth, the growth cone must integrate multiple sources of guidance cue information to modulate its cytoskeleton in order to propel the growth cone forward and accurately navigate to find its specific targets1. How this integration occurs at the cytoskeletal level is still emerging, and examination of cytoskeletal protein and effector dynamics within the growth cone can allow the elucidation of these mechanisms. Xenopus laevis growth cones are large enough (10-30 microns in diameter) to perform high-resolution live imaging of cytoskeletal dynamics (e.g.2-4 ) and are easy to isolate and manipulate in a lab setting compared to other vertebrates. The frog is a classic model system for developmental neurobiology studies, and important early insights into growth cone microtubule dynamics were initially found using this system5-7 . In this method8, eggs are collected and fertilized in vitro, injected with RNA encoding fluorescently tagged cytoskeletal fusion proteins or other constructs to manipulate gene expression, and then allowed to develop to the neural tube stage. Neural tubes are isolated by dissection and then are cultured, and growth cones on outgrowing neurites are imaged. In this article, we describe how to perform this method, the goal of which is to culture Xenopus laevis growth cones for subsequent high-resolution image analysis. While we provide the example of +TIP fusion protein EB1-GFP, this method can be applied to any number of proteins to elucidate their behaviors within the growth cone.

Neural Explant Cultures from Xenopus laevis

Culturing neural explants from dissected Xenopus laevis embryos that express fluorescent fusion proteins allows for imaging of growth cone cytoskeletal dynamics.

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During ...

Neuroscience

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 ...

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic development 1, 2. One of the signaling pathways involved in this regulation is the Wnt pathway, which is used many times during embryogenesis and critical for human disease3, 4, 5. Multiple molecular components of this pathway coordinately regulate signaling in a spatially-restricted manner, but the underlying mechanisms are not fully understood. Xenopus embryonic epithelial cells is an excellent system to study subcellular localization of various signaling proteins. Fluorescent fusion proteins are expressed in Xenopus embryos by RNA microinjection, ectodermal explants are prepared and protein localization is evaluated by epifluorescence. In this experimental protocol we describe how subcellular localization of Diversin, a cytoplasmic protein that has been implicated in signaling and cell polarity determination6, 7 is visualized in Xenopus ectodermal cells to study Wnt signal transduction8. Coexpression of a Wnt ligand or a Frizzled receptor alters the distribution of Diversin fused with red fluorescent protein, RFP, and recruits it to the cell membrane in a polarized fashion 8, 9. This ex vivo protocol should be a useful addition to in vitro studies of cultured mammalian cells, in which spatial control of signaling differs from that of the intact tissue and is much more difficult to analyze.

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Xenopus embryonic ectoderm has become an attractive model for studies of cell polarity. An assay is described, in which subcellular distribution of fluorescent proteins is assessed in ectoderm cells. This protocol will help address questions related to spatial control of signaling.

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic ...

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid decline in renal function and development of the clinical syndrome known as acute kidney injury (AKI). Pharmacological agents used to treat medical circumstances ranging from bacterial infection to cancer, when administered individually or in combination with other drugs, can initiate AKI. Zebrafish are a useful animal model to study the chemical effects on renal function in vivo, as they form an embryonic kidney comprised of nephron functional units that are conserved with higher vertebrates, including humans. Further, zebrafish can be utilized to perform genetic and chemical screens, which provide opportunities to elucidate the cellular and molecular facets of AKI and develop therapeutic strategies such as the identification of nephroprotective molecules. Here, we demonstrate how microinjection into the zebrafish embryo can be utilized as a paradigm for nephrotoxin studies.

Renal injuries incurred from nephrotoxins, which include drugs ranging from antibiotics to chemotherapeutics, can result in complex disorders whose pathogenesis remains incompletely understood. This protocol demonstrates how zebrafish can be used for disease modeling of these conditions, which can be applied to the identification of renoprotective measures.

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid ...

Xenopus Oocytes: Optimized Methods for Microinjection, Removal of Follicular Cell Layers, and Fast Solution Changes in Electrophysiological Experiments

The Xenopus oocyte as a heterologous expression system for proteins, was first described by Gurdon et al.1 and has been widely used since its discovery (References 2 - 3, and references therein). A characteristic that makes the oocyte attractive for foreign channel expression is the poor abundance of endogenous ion channels4. This expression system has proven useful for the characterization of many proteins, among them ligand-gated ion channels.
The expression of GABAA receptors in Xenopus oocytes and their functional characterization is described here, including the isolation of oocytes, microinjections with cRNA, the removal of follicular cell layers, and fast solution changes in electrophysiological experiments. The procedures were optimized in this laboratory5,6 and deviate from the ones routinely used7-9. Traditionally, denuded oocytes are prepared with a prolonged collagenase treatment of ovary lobes at RT, and these denuded oocytes are microinjected with mRNA. Using the optimized methods, diverse membrane proteins have been expressed and studied with this system, such as recombinant GABAA receptors10-12, human recombinant chloride channels13, Trypanosome potassium channels14, and a myo-inositol transporter15, 16.
The methods detailed here may be applied to the expression of any protein of choice in Xenopus oocytes, and the rapid solution change can be used to study other ligand-gated ion channels.

Xenopus Oocytes: Optimized Methods for Microinjection, Removal of Follicular Cell Layers, and Fast Solution Changes in Electrophysiological Experiments

Optimized procedures for the isolation of single follicles, cytoplasmic RNA microinjections, the removal of surrounding cell layers, and protein expression in Xenopus oocytes are described. In addition, a simple method for fast solution changes in electrophysiological experiments with ligand-gated ion channels is presented.

The Xenopus oocyte as a heterologous expression system for proteins, was first described by Gurdon et al.1 and has been widely used since its discovery ...

Hydrodynamic Renal Pelvis Injection for Non-viral Expression of Proteins in the Kidney

Hydrodynamic injection creates a local, high-pressure environment to transfect various tissues with plasmid DNA and other substances. Hydrodynamic tail vein injection, for example, is a well-established method by which the liver can be transfected. This manuscript describes an application of hydrodynamic principles by injection of the mouse kidney directly with plasmid DNA for kidney-specific gene expression. Mice are anesthetized and the kidney is exposed by a flank incision followed by a fast injection of a plasmid DNA-containing solution directly into the renal pelvis. The needle is kept in place for ten seconds and the incision site is sutured. The following day, live animal imaging, Western blot, or immunohistochemistry may be used to assay gene expression, or other assays suited to the transgene of choice are used for detection of the protein of interest. Published methods to prolong gene expression include transposon-mediated transgene integration and cyclophosphamide treatment to inhibit the immune response to the transgene.

This protocol describes a method to inject plasmid DNA into the mouse kidney via the renal pelvis to produce transgene expression specifically in the kidney.

Hydrodynamic injection creates a local, high-pressure environment to transfect various tissues with plasmid DNA and other substances. Hydrodynamic tail ...

Technique to Target Microinjection to the Developing Xenopus Kidney

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

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Microinjection of Xenopus Laevis Oocytes

Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos.

Neural Explant Cultures from Xenopus laevis

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

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

Xenopus Oocytes: Optimized Methods for Microinjection, Removal of Follicular Cell Layers, and Fast Solution Changes in Electrophysiological Experiments

Hydrodynamic Renal Pelvis Injection for Non-viral Expression of Proteins in the Kidney