Our work on RNA localization is supported by a grant from the NIH (R01GM071049) to KLM.

Part 1: Transcription of Fluorescently Labeled mRNA.
<ol>
 <li>Linearize plasmid DNA containing the RNA localization element or other relevant sequence and resuspend at 1 &mu;g/&mu;l with DEPC-treated H2O. The DNA template must have upstream promoter sites for transcription by T7, SP6, or T3 RNA polymerase.</li>
 <li>Add the following reagents to a sterile 1.5mL tube:
 <table>
 <tbody>
 <tr><td>a. 10X Tx buffer (see M&amp;M)</td><td>2 &mu;l</td></tr>
 <tr><td>b. 20X cap/NTP mix (see M&amp;M)</td><td>1 &mu;l</td></tr>
 <tr><td>c. 1 mM Alexa Fluor 546-14-UTP (Invitrogen)</td><td>1 &mu;l</td></tr>
 <tr><td>d. UTP, [&alpha;-32P](1 &mu;Ci/&mu;l) (Perkin Elmer)</td><td>1 &mu;l</td></tr>
 <tr><td>e. DEPC-H2O</td><td>11 &mu;l</td></tr>
 <tr><td>f. 0.2 M DTT</td><td>1 &mu;l</td></tr>
 <tr><td>g. RNasin (Promega)</td><td>1 &mu;l</td></tr>
 <tr><td>h. linear template DNA</td><td>1 &mu;l</td></tr>
 <tr><td>i. RNA Polymerase (Promega)</td><td>1 &mu;l</td></tr>
 </tbody>
 </table>
 </li> 
 <li>Mix reagents gently and centrifuge briefly (10 sec. in a microcentrifuge).</li>
 <li>Incubate for 2-4 hours at 37&deg; C, covering the tube with aluminum foil to prevent photobleaching of fluorophore.</li>
 <li>Add 1 &mu;l 1 mg/ml RNase-free DNase (Promega) to degrade template DNA.</li>
 <li>Incubate 15 minutes at 37&deg; C.</li>
 <li>Add 79 &mu;l 20 mM EDTA (pH 8.0) to stop the reaction.</li>
 <li>Remove 1 &mu;l to a separate tube, labeled as "input" and save.</li>
 <li>Spin reaction through a 1 ml G50 column (see Materials &amp; Methods), which will include unincorporated nucleotides while excluding full length mRNA.</li>
 <li>Concentrate the RNA by ethanol precipitation:
 <ol class="lalpha">
 <li>Add 250 &mu;l 100% ethanol, 10 &mu;l 7M CH3COONH4, 1 &mu;l carrier RNA (yeast tRNA, 10 &mu;g/&mu;l) or 1 &mu;l glycogen (20 mg/ml).</li>
 <li>Mix well and freeze until solid at -80&deg; C for &gt;30 minutes or on dry ice for 15 minutes. </li>
 <li>Spin at maximum speed in microcentrifuge for 15 minutes, decant supernatant.</li>
 <li>Wash with 150 &mu;l of 75% ethanol.</li>
 <li>Spin for 3 minutes at maximum speed in microcentrifuge, decant supernatant.</li>
 <li>Dry the pellet at 37&deg; C for 3 minutes, ensuring that all ethanol has evaporated.</li>
 </ol>
 </li>
 <li>Resuspend in 11 &mu;l DEPC-H2O and remove 1 &mu;l to a separate tube, labeled as "incorporated".</li>
 <li>Determine the percent incorporation and dilute the RNA to 50 nM (see Materials &amp; Methods for calculation). </li>
 <li>The RNA should be frozen in 5 &mu;l aliquots, for single use to avoid freeze / thaw cycles and can be stored for several weeks at -80&deg; C.</li>
</ol>
Part 2: Preparing for Microinjection.
<ol>
 <li>RNA Preparation: Thaw an aliquot of RNA (50 nM), and denature the RNA at 70&deg; C for 3-5 minutes. Spin for 10 minutes in room temperature microcentrifuge at maximum speed to remove any particulates, and keep on ice.</li>
 <li>Oocyte Preparation:
 <ol class="lalpha">
 <li>Surgically remove ovary from Xenopus laevis females (Nasco). </li>
 <li>Trim pieces of Xenopus laevis ovary into a 50 ml conical tube containing 25 ml of Collagenase solution (see Materials &amp; Methods). </li>
 <li>Shake gently at 18&deg; C for 15 minutes, or until oocytes are visibly separated from ovary. Due to batch to batch variation, collagenase treatment time may vary and should be monitored carefully through visual inspection to ensure defoliculation. </li>
 <li>Allow oocytes to settle in tube, remove solution and wash the oocytes with MBSH buffer (see Materials &amp; Methods). </li>
 <li>Repeat wash twice more for a total of three washes. This oocyte isolation protocol is similar to that detailed in Cohen et al., 2009. </li>
 <li>Manually sort stage III/IV oocytes (Dumont, 1972) in MBSH buffer under a standard dissecting microscope. Stage III oocytes are completely opaque but white, while stage IV oocytes are slightly larger and speckled with pigment. Oocytes that are slightly transparent are too young and oocytes that fully pigmented or exhibit polarized pigment distribution are too old.</li>
 </ol>
 </li>
 <li>Needle Preparation: Pull and bevel needles to an outer diameter of ~ 0.05 mm. (We use Drummond Scientific 3.5 inch capillaries (order # 3-000-203-G/X), a Sutter Instrument Co. micropipette puller and a WPI Inc. beveller.)</li>
</ol>
Part 3: Microinjection
<ol>
 <li>Calibrate needle with DEPC-H2O to deliver 2 nl per injection. We front load our needle using a gas-driven microinjector (see Materials &amp; Methods), and calibrate drop size using a micrometer. </li>
 <li>Load RNA into needle.</li>
 <li>Place sorted oocytes into an injection dish containing MBSH buffer. We microinject on a dish with a layer of black foam rubber. The pale oocytes stand out well on a white background.</li>
 <li>Carefully inject each oocyte with 2 nl of RNA at 50 nM.</li>
 <li>Expel RNA, rinse needle with DEPC-H2O, and load next RNA for injection.</li>
</ol>
Part 4: Oocyte Culture
<ol>
 <li>Place oocytes in a well of a sterile 24 well plate (Sigma Aldrich). We culture up to a thousand oocytes per well. </li>
 <li>Remove buffer and replace with 400 &mu;l Oocyte Culture Medium (OCM; see Materials &amp; Methods) per well. Place culture plate inside an air-tight plastic container with wet paper towel to maintain moist environment during culture.</li>
 <li>Incubate oocytes at 18&deg; C for time points ranging between 8 and 48 hours.</li>
</ol>
Part 5: Fixation, Dehydration and Storage of Oocytes
<ol>
 <li>Sort out any dead oocytes and place surviving oocytes in glass vials. We routinely observe &gt;90% survival. </li>
 <li>Place oocytes in MEMFA fixative (see Materials &amp; Methods) and rock for 20 minutes. Protect the oocytes from light by covering with aluminum foil.</li>
 <li>Wash oocytes by removing fixative and replacing with equal volume of MBSH buffer. Repeat this wash once more for a total of two washes.</li>
 <li>Wash oocytes into anhydrous methanol:
 <ol class="lalpha">
 <li>Remove half of the volume, replace with methanol.</li>
 <li>Repeat step "a" twice.</li>
 <li>Remove all of the solution, replace with methanol.</li>
 <li>Repeat methanol wash.</li>
 </ol>
 </li>
 <li>Oocytes can be stored at -20&deg; C until ready to image.</li>
</ol>
Part 6: Imaging RNA Localization by Confocal Microscopy
<ol>
 <li>Before imaging, add Murray s Clearing Medium (2:1 benzyl benzoate-benzyl alcohol) to a glass bottom FluoroDish (WPI Inc.) to cover the imaging surface. </li>
 <li>Carefully transfer the oocytes from methanol to FluoroDish, minimizing the volume of methanol transferred.</li>
 <li>Image on an inverted confocal microscope (We have successfully used a Zeiss LSM510 and a Leica TCS SP2).</li>
</ol>
<img src="/files/ftp_upload/1704/1704fig1.jpg" alt="Figure 1" /> Figure 1: Visualization of Subcellular RNA Localization by Microinjection of Fluorescently Labelled Transcripts.&nbsp; A.&nbsp; Following microinjection, RNA labeled with Alexa Fluor 546 is initially restricted to the nucleus. B. After eight hours of culture, a fluorescently labeled control RNA can be seen uniformly distributed in the cytoplasm of the oocyte. C. However, fluorescently labeled RNA containing sequences that recruit the transport machinery can be seen in the process of subcellular localization to the vegetal pole of the oocyte (towards bottom). Scale bar = 50 &mu;m.

<ol>
	<li>Cohen, S., Au, S., Pant, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+of+Xenopus+laevis+oocytes.">Microinjection of Xenopus laevis oocytes.</a> JoVE. 24, (2009).</li><li>Dumont, J. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oogenesis+in+Xenopus+laevis+(Daudin).+I.+Stages+of+oocyte+development+in+laboratory+maintained+animals.">Oogenesis in Xenopus laevis (Daudin). I. Stages of oocyte development in laboratory maintained animals.</a> J. Morphol. 136, 153-179 (1972).</li><li>Gagnon, J. A., Mowry, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+transport+in+ovo:+Simultaneous+visualization+of+two+RNAs.">RNA transport in ovo: Simultaneous visualization of two RNAs.</a> Mol Reprod Dev. 76, 1115-1115 (2009).</li><li>Gautreau, D., Cote, C. A., Mowry, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+copies+of+a+subelement+from+the+Vg1+RNA+localization+sequence+are+sufficient+to+direct+vegetal+localization+in+Xenopus+oocytes.">Two copies of a subelement from the Vg1 RNA localization sequence are sufficient to direct vegetal localization in Xenopus oocytes.</a> Development. 124, 5013-5020 (1997).</li><li>Lewis, R. A., Gagnon, J. A., Mowry, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PTB/hnRNP+I+is+required+for+RNP+remodeling+during+RNA+localization+in+Xenopus+oocytes.">PTB/hnRNP I is required for RNP remodeling during RNA localization in Xenopus oocytes.</a> Mol Cell Biol. 28, 678-6786 (2008).</li><li>Messitt, T. J., Gagnon, J. A., Kreiling, J. A., Pratt, C. A., Yoon, Y. J., Mowry, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiple+kinesin+motors+coordinate+cytoplasmic+RNA+transport+on+a+subpopulation+of+microtubules+in+Xenopus+oocytes.">Multiple kinesin motors coordinate cytoplasmic RNA transport on a subpopulation of microtubules in Xenopus oocytes.</a> Dev Cell. 15, 426-436 (2008).</li><li>Mowry, K. L., Melton, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vegetal+messenger+RNA+localization+directed+by+a+340-nt+RNA+sequence+element+in+Xenopus+oocytes.">Vegetal messenger RNA localization directed by a 340-nt RNA sequence element in Xenopus oocytes.</a> Science. 255, 991-994 (1992).</li><li>Yoon, Y. J., Mowry, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus+Staufen+is+a+component+of+a+ribonucleoprotein+complex+containing+Vg1+RNA+and+kinesin.">Xenopus Staufen is a component of a ribonucleoprotein complex containing Vg1 RNA and kinesin.</a> Development. 131, 3035-3045 (2004).</li></ol>

Experiments using laboratory animals were performed in accordance with the guidelines and regulations set forth by Brown University.

Here we have presented a protocol for visualizing mRNA localization in Xenopus oocytes. This method, using fluorescently labeled RNA transcripts has higher signal to noise ratio than previously obtained with digoxigenin-labeled transcripts and is simpler and faster than in-situ based approaches (Mowry and Melton, 1992, Gautreau et al., 1997). Using this method, we can engineer RNA sequence mutations and rapidly test for in vivo function. Further, by using additional Alexa-UTP fluorophores, multiple RNA species can be visualized in the same oocyte. We have found that in Xenopus oocytes Alexa Fluor 546-14-UTP gives superior results as compared with Alexa Fluor 488-5-UTP because of autofluorescence in the 488 nm wavelength range; however, dual imaging of two RNAs is nonetheless possible (Gagnon and Mowry, 2009). Also, this technique works well in conjunction with immunostaining protocols to detect protein colocalization, as a wide variety of compatible fluorescent secondary antibodies are available (Yoon and Mowry, 2006, Messitt et al., 2008). Finally, disruption of the RNA localization process through over-expression of dominant negative factors or small molecule interference can be combined with this assay to visualize subtle defects in the mRNA localization pathway (Messitt et al., 2008). This technique has proven to be a relatively simple and robust assay for detecting mRNA distribution in vivo and can be integrated easily into existing biochemical, molecular and cell biological techniques for probing mechanisms of mRNA transport.

A correction to the step 2 of Part 1 for article Visualizing RNA Localization in Xenopus Oocytes.

Erratum: Visualizing RNA Localization in Xenopus Oocytes

10X Tx buffer
<ul>
<li class="none">60 mM MgCl2</li>
<li class="none">400 mM Tris-HCl (pH 7.5)</li>
<li class="none">20 mM spermidine-HCl</li>
</ul>
20x cap/NTP mix
<ul>
<li class="none">10 mM CTP</li>
<li class="none">10 mM ATP</li>
<li class="none">9 mM UTP</li>
<li class="none">2 mM GTP</li>
<li class="none">20 mM G(ppp)G Cap Analog (New England Biolabs)</li>
</ul>
G-50 column
<ul>
<li class="none">Hydrate 5 g Sephadex G-50 beads (Sigma Aldrich) in 100 ml deionized H2O. DEPC-treat for 30 min. and autoclave. Store incomplete stock at room temperature. Before use, add the following RNase-free solutions:</li>
<li class="none">0.5 ml 0.2 M EDTA</li>
<li class="none">1 ml 1 M Tris pH 8.0</li>
<li class="none">0.5 ml 20% SDS</li>
<li class="none">Store complete G-50 solution at 4&deg; C.</li>
<li class="none">Remove and discard the plunger from a 3 ml syringe (BD Biosciences) and place the barrel of the syringe into a 15 ml conical tube (Corning). Plug the syringe with a small amount of glass wool (a plug about half the size of a penny).</li>
<li class="none">Swirl complete G-50 solution to resuspend beads.</li>
<li class="none">Add 2 ml G-50 solution to the empty column.</li>
<li class="none">Spin for 1 minute at 1,000 x g in benchtop centrifuge.</li>
<li class="none">Add 200 &mu;l DEPC-treated deionized H2O to each column. Spin.</li>
<li class="none">Repeat wash twice more for a total of three washes.</li>
<li class="none">Remove syringe barrel to a fresh 15 ml conical tube.</li>
</ul>
Collagenase solution
<ul>
<li class="none">75 mg collagenase from Clostridium histolyticum (Sigma Aldrich)</li>
<li class="none">25 ml 0.1 M KPO3+ (pH 7.4)</li>
</ul>
MBSH buffer
<ul>
<li class="none">88 mM NaCl</li>
<li class="none">1 mM KCl</li>
<li class="none">2.4 mM NaHCO3</li>
<li class="none">0.82 mM MgSO4 X 7H2O</li>
<li class="none">0.33 mM Ca(NO3)2 X 4H2O</li>
<li class="none">0.41 mM CaCl2 X 6H2O</li>
<li class="none">10 mM HEPES (pH 7.6)</li>
</ul>
Oocyte Culture Medium
<ul>
<li class="none">50% L15 medium</li>
<li class="none">15 mM HEPES (pH 7.6)</li>
<li class="none">1 mg/ml insulin</li>
<li class="none">100 mg/ml gentamicin</li>
<li class="none">50 U/ml nystatin</li>
<li class="none">50 U/ml penicillin</li>
<li class="none">50 mg/ml streptomycin</li>
</ul>
MEMFA solution
<ul>
<li class="none">0.1 M MOPS (pH 7.4)</li>
<li class="none">2 mM EGTA</li>
<li class="none">1 mM MgSO4</li>
<li class="none">3.7% formaldehyde</li>
</ul>
Computing RNA yield
<ul>
<li class="none">Determine CPM in "input" and "incorporated" samples using a standard scintillation counter.</li>
<li class="none">incorporation = ("incorporated") / (10 x "input")</li>
<li class="none">Typical incorporation values range between ~0.03 and 0.10.</li>
<li class="none">Maximum theoretical yields for different polymerases: 
T7, T3, SP6 - 2.64 &mu;g</li>
<li class="none">Reaction yield in &mu;g = (maximum yield of polymerase used) X (incorporation)</li>
<li class="none">Dilute RNA to 50 nM = (&mu;g RNA) / 320 / (length of RNA in bases) / (5X10-8)</li>
<li class="none">The reaction usually yields ~50-100 &mu;l of RNA at 50 nM.</li>
</ul>

visualizing rna localization in xenopus oocytes

RNA localization is a conserved mechanism of establishing cell polarity. Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to spatially restrict gene expression of Vg1 protein. Tight control of Vg1 distribution in this manner is required for proper germ layer specification in the developing embryo. RNA sequence elements in the 3' UTR of the mRNA, the Vg1 localization element (VLE) are required and sufficient to direct transport. To study the recognition and transport of Vg1 mRNA in vivo, we have developed an imaging technique that allows extensive analysis of trans-factor directed transport mechanisms via a simple visual readout. 
To visualize RNA localization, we synthesize fluorescently labeled VLE RNA and microinject this transcript into individual oocytes. After oocyte culture to allow transport of the injected RNA, oocytes are fixed and dehydrated prior to imaging by confocal microscopy. Visualization of mRNA localization patterns provides a readout for monitoring the complete pathway of RNA transport and for identifying roles in directing RNA transport for cis-acting elements within the transcript and trans-acting factors that bind to the VLE (Lewis et al., 2008, Messitt et al., 2008). We have extended this technique through co-localization with additional RNAs and proteins (Gagnon and Mowry, 2009, Messitt et al., 2008), and in combination with disruption of motor proteins and the cytoskeleton (Messitt et al., 2008) to probe mechanisms underlying mRNA localization.

Watch this Scientific Journal Video about Visualizing RNA Localization in Xenopus Oocytes at JoVE.com

Visualizing RNA Localization in Xenopus Oocytes

Visualization of in vivo RNA transport is accomplished by microinjection of fluorescently labeled RNA transcripts into Xenopus oocytes, followed by confocal microscopy.

Visualizing RNA Localization in Xenopus Oocytes

RNA localization is a conserved mechanism of establishing cell polarity.  Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to ...

visualizing-rna-localization-in-xenopus-oocytes

Nanyang Technological University

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JoVE Journal

Biology

11.5K Views.  Brown University. Visualization of in vivo RNA transport is accomplished by microinjection of fluorescently labeled RNA transcripts into Xenopus oocytes, followed by confocal microscopy.

Video: Visualizing RNA Localization in Xenopus Oocytes

Patch Clamp Recording of Ion Channels Expressed in  Xenopus Oocytes

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological measurement of single or multiple ion channels in cells. This technique can be applied to ion channels in both their native environment and expressed in heterologous cells, such as oocytes harvested from the African clawed frog, Xenopus laevis. Here, we describe the well-established technique of patch clamp recording from Xenopus oocytes. This technique is used to measure the properties of expressed ion channels either in populations (macropatch) or individually (single-channel recording). We focus on techniques to maximize the quality of oocyte preparation and seal generation. With all factors optimized, this technique gives a probability of successful seal generation over 90 percent. The process may be optimized differently by every researcher based on the factors he or she finds most important, and we present the approach that have lead to the greatest success in our hands.

This is intended as an introduction to patch clamp recording from Xenopus laevis oocytes. It covers vitelline membrane removal, formation of a gigaohm seal (gigaseal), and the optional conversion of the patch to the outside-out topology.

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological ...

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 ...

Fertilization of Xenopus oocytes using the Host Transfer Method

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to generate maternal-effect mutant animals. Additionally, many of the techniques to overexpress or inhibit gene function in organisms such as Xenopus and zebrafish fail to sufficiently target critical maternal signaling pathways, such as Wnt signaling. In Xenopus, manipulating gene function in cultured oocytes and subsequently fertilizing them can ameliorate these problems to some extent. Oocytes are manually defolliculated from donor ovary tissue, injected or treated in culture as desired, and then stimulated with progesterone to induce maturation. Next, the oocytes are introduced into the body cavity of an ovulating host female frog, whereupon they will be translocated through the host's oviduct and acquire modifications and jelly coats necessary for fertilization. The resulting embryos can then be raised to the desired stage and analyzed for the effects of any experimental perturbations. This host-transfer method has been highly effective in uncovering basic mechanisms of early development and allows a wide range of experimental possibilities not available in any other vertebrate model organism.

Fertilization of Xenopus oocytes using the Host Transfer Method

Procedure for fertilizing Xenopus oocytes by the host transfer method.

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

In vitro Transcription and Capping of Gaussia Luciferase mRNA Followed by HeLa Cell Transfection

In vitro transcription is the synthesis of RNA transcripts by RNA polymerase from a linear DNA template containing the corresponding promoter sequence (T7, T3, SP6) and the gene to be transcribed (Figure 1A). A typical transcription reaction consists of the template DNA, RNA polymerase, ribonucleotide triphosphates, RNase inhibitor and buffer containing Mg2+ ions.
Large amounts of high quality RNA are often required for a variety of applications. Use of in vitro transcription has been reported for RNA structure and function studies such as splicing1, RNAi experiments in mammalian cells2, antisense RNA amplification by the "Eberwine method"3, microarray analysis4 and for RNA vaccine studies5. The technique can also be used for producing radiolabeled and dye labeled probes6. Warren, et al. recently reported reprogramming of human cells by transfection with in vitro transcribed capped RNA7. The T7 High Yield RNA Synthesis Kit from New England Biolabs has been designed to synthesize up to 180 &mu;g RNA per 20 &mu;l reaction. RNA of length up to 10kb has been successfully transcribed using this kit. Linearized plasmid DNA, PCR products and synthetic DNA oligonucleotides can be used as templates for transcription as long as they have the T7 promoter sequence upstream of the gene to be transcribed.
Addition of a 5' end cap structure to the RNA is an important process in eukaryotes. It is essential for RNA stability8, efficient translation9, nuclear transport10 and splicing11. The process involves addition of a 7-methylguanosine cap at the 5' triphosphate end of the RNA. RNA capping can be carried out post-transcriptionally using capping enzymes or co-transcriptionally using cap analogs. In the enzymatic method, the mRNA is capped using the Vaccinia virus capping enzyme12,13. The enzyme adds on a 7-methylguanosine cap at the 5' end of the RNA using GTP and S-adenosyl methionine as donors (cap 0 structure). Both methods yield functionally active capped RNA suitable for transfection or other applications14 such as generating viral genomic RNA for reverse-genetic systems15 and crystallographic studies of cap binding proteins such as eIF4E16.
In the method described below, the T7 High Yield RNA Synthesis Kit from NEB is used to synthesize capped and uncapped RNA transcripts of Gaussia luciferase (GLuc) and Cypridina luciferase (CLuc). A portion of the uncapped GLuc RNA is capped using the Vaccinia Capping System (NEB). A linearized plasmid containing the GLuc or CLuc gene and T7 promoter is used as the template DNA. The transcribed RNA is transfected into HeLa cells and cell culture supernatants are assayed for luciferase activity. Capped CLuc RNA is used as the internal control to normalize GLuc expression.

In vitro Transcription and Capping of Gaussia Luciferase mRNA Followed by HeLa Cell Transfection

This method describes high yield in vitro synthesis of both capped and uncapped mRNA from a linearized plasmid containing the Gaussia luciferase (GLuc) gene. The RNA is purified and a fraction of the uncapped RNA is enzymatically capped using the Vaccinia virus capping enzyme. In the final step, the mRNA is transfected into HeLa cells and cell culture supernatants are assayed for luciferase activity.

In vitro transcription is the synthesis of RNA transcripts by RNA polymerase from a linear DNA template containing the corresponding promoter sequence ...

Chromatin Isolation by RNA Purification (ChIRP)

Long noncoding RNAs are key regulators of chromatin states for important biological processes such as dosage compensation, imprinting, and developmental gene expression 1,2,3,4,5,6,7. The recent discovery of thousands of lncRNAs in association with specific chromatin modification complexes, such as Polycomb Repressive Complex 2 (PRC2) that mediates histone H3 lysine 27 trimethylation (H3K27me3), suggests broad roles for numerous lncRNAs in managing chromatin states in a gene-specific fashion 8,9. While some lncRNAs are thought to work in cis on neighboring genes, other lncRNAs work in trans to regulate distantly located genes. For instance, Drosophila lncRNAs roX1 and roX2 bind numerous regions on the X chromosome of male cells, and are critical for dosage compensation 10,11. However, the exact locations of their binding sites are not known at high resolution. Similarly, human lncRNA HOTAIR can affect PRC2 occupancy on hundreds of genes genome-wide 3,12,13, but how specificity is achieved is unclear. LncRNAs can also serve as modular scaffolds to recruit the assembly of multiple protein complexes. The classic trans-acting RNA scaffold is the TERC RNA that serves as the template and scaffold for the telomerase complex 14; HOTAIR can also serve as a scaffold for PRC2 and a H3K4 demethylase complex 13.
Prior studies mapping RNA occupancy at chromatin have revealed substantial insights 15,16, but only at a single gene locus at a time. The occupancy sites of most lncRNAs are not known, and the roles of lncRNAs in chromatin regulation have been mostly inferred from the indirect effects of lncRNA perturbation. Just as chromatin immunoprecipitation followed by microarray or deep sequencing (ChIP-chip or ChIP-seq, respectively) has greatly improved our understanding of protein-DNA interactions on a genomic scale, here we illustrate a recently published strategy to map long RNA occupancy genome-wide at high resolution 17. This method, Chromatin Isolation by RNA Purification (ChIRP) (Figure 1), is based on affinity capture of target lncRNA:chromatin complex by tiling antisense-oligos, which then generates a map of genomic binding sites at a resolution of several hundred bases with high sensitivity and low background. ChIRP is applicable to many lncRNAs because the design of affinity-probes is straightforward given the RNA sequence and requires no knowledge of the RNA's structure or functional domains.

Chromatin Isolation by RNA Purification ChIRP

ChIRP is a novel and rapid technique to map genomic binding sites of long noncoding RNAs (lncRNAs). The method takes advantage of the specificity of anti-sense tiling oligonucleotides to allow the enumeration of lncRNA-bound genomic sites.

Long noncoding RNAs are key regulators of chromatin states for important biological processes such as dosage compensation, imprinting, and developmental ...

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

Whereas cation transport by the electrogenic membrane transporter Na+,K+-ATPase can be measured by electrophysiology, the electroneutrally operating gastric H+,K+-ATPase is more difficult to investigate. Many transport assays utilize radioisotopes to achieve a sufficient signal-to-noise ratio, however, the necessary security measures impose severe restrictions regarding human exposure or assay design. Furthermore, ion transport across cell membranes is critically influenced by the membrane potential, which is not straightforwardly controlled in cell culture or in proteoliposome preparations. Here, we make use of the outstanding sensitivity of atomic absorption spectrophotometry (AAS) towards trace amounts of chemical elements to measure Rb+ or Li+ transport by Na+,K+- or gastric H+,K+-ATPase in single cells. Using Xenopus oocytes as expression system, we determine the amount of Rb+ (Li+) transported into the cells by measuring samples of single-oocyte homogenates in an AAS device equipped with a transversely heated graphite atomizer (THGA) furnace, which is loaded from an autosampler. Since the background of unspecific Rb+ uptake into control oocytes or during application of ATPase-specific inhibitors is very small, it is possible to implement complex kinetic assay schemes involving a large number of experimental conditions simultaneously, or to compare the transport capacity and kinetics of site-specifically mutated transporters with high precision. Furthermore, since cation uptake is determined on single cells, the flux experiments can be carried out in combination with two-electrode voltage-clamping (TEVC) to achieve accurate control of the membrane potential and current. This allowed e.g. to quantitatively determine the 3Na+/2K+ transport stoichiometry of the Na+,K+-ATPase and enabled for the first time to investigate the voltage dependence of cation transport by the electroneutrally operating gastric H+,K+-ATPase. In principle, the assay is not limited to K+-transporting membrane proteins, but it may work equally well to address the activity of heavy or transition metal transporters, or uptake of chemical elements by endocytotic processes.

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

We describe a method to quantify the activity of K+-countertransporting P-type ATPases by heterologous expression of the enzymes in Xenopus oocytes and measuring Rb+ or Li+ uptake into individual cells by atomic absorption spectrophotometry. The method is a sensitive and safe alternative to radioisotope flux experiments facilitating complex kinetic studies.

Whereas cation transport by the electrogenic membrane transporter Na+,K+-ATPase can be measured by electrophysiology, the electroneutrally operating ...

Facial Transplants in Xenopus laevis Embryos

Craniofacial birth defects occur in 1 out of every 700 live births, but etiology is rarely known due to limited understanding of craniofacial development. To identify where signaling pathways and tissues act during patterning of the developing face, a 'face transplant' technique has been developed in embryos of the frog Xenopus laevis. A region of presumptive facial tissue (the "Extreme Anterior Domain" (EAD)) is removed from a donor embryo at tailbud stage, and transplanted to a host embryo of the same stage, from which the equivalent region has been removed. This can be used to generate a chimeric face where the host or donor tissue has a loss or gain of function in a gene, and/or includes a lineage label. After healing, the outcome of development is monitored, and indicates roles of the signaling pathway within the donor or surrounding host tissues. Xenopus is a valuable model for face development, as the facial region is large and readily accessible for micromanipulation. Many embryos can be assayed, over a short time period since development occurs rapidly. Findings in the frog are relevant to human development, since craniofacial processes appear conserved between Xenopus and mammals.

Facial Transplants in Xenopus laevis Embryos

A technique for transplanting "Extreme Anterior Domain" facial tissue between Xenopus laevis embryos has been developed. Tissue can be moved from one gene expression background into another, allowing the study of local requirements for craniofacial development and for signaling interactions between facial regions.

Craniofacial birth defects occur in 1 out of every 700 live births, but etiology is rarely known due to limited understanding of craniofacial development. ...

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion channels in oocytes. Recordings from the cut-open setup are particularly useful for resolving low magnitude gating currents, rapid ionic current activation, and deactivation. The main benefits over the two-electrode voltage clamp (TEVC) technique include increased clamp speed, improved signal-to-noise ratio, and the ability to modulate the intracellular and extracellular milieu.
Here, we employ the human cardiac sodium channel (hNaV1.5), expressed in Xenopus oocytes, to demonstrate the cut-open setup and protocol as well as modifications that are required to add voltage clamp fluorometry capability.
The properties of fast activating ion channels, such as hNaV1.5, cannot be fully resolved near room temperature using TEVC, in which the entirety of the oocyte membrane is clamped, making voltage control difficult. However, in the cut-open technique, isolation of only a small portion of the cell membrane allows for the rapid clamping required to accurately record fast kinetics while preventing channel run-down associated with patch clamp techniques.
In conjunction with the COVG technique, ion channel kinetics and electrophysiological properties can be further assayed by using voltage clamp fluorometry, where protein motion is tracked via cysteine conjugation of extracellularly applied fluorophores, insertion of genetically encoded fluorescent proteins, or the incorporation of unnatural amino acids into the region of interest1. This additional data yields kinetic information about voltage-dependent conformational rearrangements of the protein via changes in the microenvironment surrounding the fluorescent molecule.

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open Vaseline gap approach is used to obtain low noise recordings of ionic and gating currents from voltage-dependent ion channels expressed in Xenopus oocytes with high resolution of fast channel kinetics. With minor modification, voltage clamp fluorometry can be coupled to the cut-open oocyte protocol.

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion ...

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 ...

Visualizing RNA Localization in Xenopus Oocytes

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Patch Clamp Recording of Ion Channels Expressed in Xenopus Oocytes

Microinjection of Xenopus Laevis Oocytes

Fertilization of Xenopus oocytes using the Host Transfer Method

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

In vitro Transcription and Capping of Gaussia Luciferase mRNA Followed by HeLa Cell Transfection

Chromatin Isolation by RNA Purification (ChIRP)

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

Facial Transplants in Xenopus laevis Embryos

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

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