pAnap was a kind gift from Dr. Peter Schultz (Scripps Research Institute). This work was funded by the Canadian Institutes for Health Research Grants MOP-102689 and MOP-136894 (to R.B.) and Canadian Foundation for Innovation Grant 950-225005.

Frog manipulations were performed in accordance with the Canadian guidelines and have been approved by the ethics committee (CDEA, protocol #15-042) of University of Montr&#233;al.
1. mRNA Preparation for fUAA Incorporation
<ol>
	<li>Choose a site of interest in the protein where conformational changes are expected to occur. Select an amino acid in this region to be substituted for the fUAA. 
	NOTE: The choice of position is based on the structural rearrangements that are expected. If a high resolution structure exists and a hypothesis of the expected movements, the anap should be placed such that the chemical environment will alter; this might be either a change in the dielectric constant (hydrophobic versus hydrophilic environment) or, more likely, quenching by another amino acid. The best quenchers are tryptophans. Anap should be in contact with the quencher in one state (overlap of the van-der-Waals radii) and free of it in the other. If no high-resolution structures or models exist, one would have to scan the region of interest. In either case, it is advisable to select several nearby locations to increase the probability of obtaining expression and fluorescence signal. In order to minimize steric effects during protein maturation and/or function, one may choose to replace large and aromatic amino acids (Phe, Trp, Tyr). The authors, however, have experienced that scanning a region of interest for fUAA insertion regardless of the substituted amino acid, is more productive.</li>
	<li>Insert an amber stop codon (TAG) at the selected site using site-directed mutagenesis15. Ensure that the protein of interest does not end on an amber stop codon (TAG). If so, mutate to a different one (ochre or opal stop codon). Amplify, isolate and sequence the DNA. Obtain protein mRNA with in vitro transcription16 and store the mRNA at 20 &#176;C or 80 &#176;C.</li>
</ol>
2. Oocyte Preparation and Injection
<ol>
	<li>Surgically obtain Stage V or VI oocytes from Xenopus laevis frogs and defolliculate with collagenase as previously described17.
	<ol>
		<li>Anesthetize frogs with an appropriate anesthetic according to the approved animal protocol (here: 3-aminobenzoic acid ethyl ester). When they fail to respond to a gentle pinch to a toe-tip (loss of withdrawal reflex), then they are suitably anesthetized for surgery.</li>
		<li>Immediately remove the frogs from the anesthetic solution and thoroughly rinse its skin with fresh water. This rinse will prevent the animal from falling into deeper levels of anesthesia by removing unabsorbed chemical from the skin surface.</li>
		<li>Remove ovary nodes from one side surgically and carefully open the nodes using two forceps. Incubate and agitate the oocytes in &#34;standard oocyte solution&#34; (SOS) containing 1% (w/v) collagenase for 20-30 min to defolliculate. Wash three times with SOS solution.</li>
		<li>Select large and healthy oocytes individually and incubate them in Barth&#39;s solution supplemented with antibiotics (100 U/mL penicilin, 100 &#181;g/mL streptomycin, 10 mg/100 mL kanamycin) and 5% horse serum at 18 &#870;C &#160;for at least 4 h before injection. 
		NOTE: After 2 - 4 surgeries with a 4 month delay in between, Xenopus laevis are euthanized by prolonged (&#62;1 h) incubation with 3-aminobenzoic acid ethyl ester.</li>
	</ol>
	</li>
	<li>For nuclear injection of DNA, prepare a long and thin injection tip to be able to reach the nucleus and to avoid damaging the oocyte. Fill the injection tip with oil and mount it on the nanoinjector device.
	<ol>
		<li>Install the nano-injector under a stereo microscope and use forceps to break the end of the tip. Eject oil until there is no air bubbles trapped inside the end of the tip.</li>
	</ol>
	</li>
	<li>Place 1 &#181;L of 0.1 &#181;g/&#181;L pAnap in nuclease-free water containing NaOH (1% of 1 N NaOH) on a piece of parafilm under a stereoscope and fill the injection tip with the DNA.</li>
	<li>Transfer 40 oocytes to a mesh-coated injection dish containing Barth&#39;s solution supplemented with antibiotics. 
	NOTE: To make the mesh-coated injection dish, cut out an appropriately sized piece of 800 &#181;m nylon mesh to fill a polystyrene Petri dish. Add chloroform to the center and then place the mesh on top. Hold the mesh flat until the plastic sets.</li>
	<li>As the oocyte nucleus is located in the animal (dark) pole, aim the injection tip at the center of the animal pole and impale such that the tip reaches near the center of the animal hemisphere (or 2-3x the depth compared to RNA injection). Inject 9.2 nL of pAnap into the nucleus of each oocyte. The thin tip and small injection volume may result in irregular injection or blocked tip. Occasionally check whether the injection works by injecting into the air. 
	NOTE: Whether the DNA gets properly injected into the nucleus is uncertain. Expect therefore 10 - 40% of the oocytes to not express the tRNA/synthetase pair. See Discussion for further elaboration.</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/55598/55598fig2.jpg" /> 
Figure 2: Illustration of DNA and RNA Injection into Xenopus&#160;Oocytes for Anap Incorporation. 
First, pAnap is injected into the nucleus of the Xenopus oocyte (1). After 6-24 h, Anap and channel RNA are coinjected into the vegetal pole (2). Anap will be orthogonally aminoacylated with the tRNA bearing an amber stop anti-codon, by the aminoacyl-tRNA synthetase which is encoded by pAnap. This way, the aminoacylated Anap-tRNAs are recognized by the ribosome at the inserted amber stop codon in the channel RNA, resulting in suppression of the stop codon and insertion of Anap. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/55598/55598fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="6">
	<li>Incubate the oocytes in 2 mL Barth&#39;s solution supplemented with antibiotics and 5% Horse serum (HS) at 18 &#176;C for 6-24 h to allow robust expression of Anap specific tRNAs and tRNA-synthetases. 
	NOTE: DNA incubation time can last several days before RNA injection, but it does not increase expression.</li>
	<li>Prepare the nano-injector for RNA injection (same as step 2.2 but the injection tip does not need to be as thin as for DNA injection). Work only under red light from this point to prevent photobleaching of the Anap.</li>
	<li>Mix 1 &#181;L of 1 mM Anap with 1 &#181;L of 1-2 &#181;g/&#181;L mRNA directly on a piece of parafilm and fill the injection tip with the mixed solution. Impale just below the membrane in the vegetal (bright) pole and inject 46 nL in each pAnap-injected oocyte. 
	NOTE: The required mRNA concentration depends on the protein of interest.</li>
	<li>Incubate the oocytes protected from light in a box or wrapped in aluminium foil, in Barth&#39;s solution supplemented antibiotics and 5% horse serum at 18 &#176;C for 2-3 days. Exchange with fresh Barth&#39;s solution every day and remove dead oocytes to avoid contamination.</li>
</ol>
3. VCF Setup
<ol>
	<li>Install the cut-open oocyte voltage-clamp equipment as previously described18.</li>
	<li>Mount the electrophysiology recording system on an upright fluorescence microscope by installing the recording chamber on a slider that allows moving it between the standard stereoscope for placing the oocyte and the microscope to perform the fluorescence measurements (Figure 2c). 
	NOTE: The geometry of the chamber for cut-open oocytes is not suitable to use the normal transmitted light for illumination during manipulation. Therefore, a &#34;gooseneck&#34; halogen lamp with red filter is used to illuminate sideways from the top. The condenser of the microscope can be removed make space to lower the stage for the electrophysiology chamber.</li>
	<li>Connect a photodiode detection system to the C-mount exit port of the fluorescence microscope (Figure 2a). Connect the photocurrent readout to a second input channel in the digital signal processor (DSP, analog/digital &#8211; digital/analog converter).</li>
	<li>Use a 100 W, 12 V halogen lamp as light source for the fluorescence excitation. 
	NOTE: Alternatively, Hg-burners may be used, but have to be reduced in intensity to prevent too rapid photobleaching during recordings. LED illumination is only recommended if the respective LEDs show significant intensity in the excitation range (e.g., ~350 nm for Anap). Most white LEDs do not reach far into the UV spectrum.</li>
	<li>Insert an electrically-controlled shutter between excitation light source and microscope and connect its control (typically TTL-pulse) to a digital output of the DSP. Time the TTL-pulse in the recording software (see manufacturer documentation), such that the shutter opens ~100 ms before the beginning of the recording. This way, any vibration during the opening process does not interfere with the recording. The time depends on the speed and vibration of the shutter. End the pulse 5ms before the end of the recording as shown in Figure 4. This way, the value for the total fluorescence is also recorded.</li>
	<li>Insert an appropriate filter cube (excitation filter, dichroic mirror and emission filter) in the filter cube turret. For Anap, use Ex: 377/50 nm bandpass, dichroic 409 nm longpass, and Em: 470/40 nm bandpass.</li>
</ol>
<img alt="Figure 3" src="/files/ftp_upload/55598/55598fig3.jpg" /> 
Figure 3: VCF setup. (A) Side view of the VCF setup showing the light path inside the microscope. The filter cube contains an excitation filter, dichroic mirror and an emission filter. (B) Selected oocyte chamber dimensions are 3.4 cm for upper chamber radius (1), 5.5 cm for bottom chamber length (2), 1.4 cm for bottom chamber width (3) and 1.7 cm for middle chamber width (4). (C) Front view of the VCF setup. The first ocular on the left is for mounting the oocyte in the cut-open voltage clamp chamber and for permeabilization. Then, the chamber is slid under the microscope at the second ocular to the right. Here, the V1 electrode is inserted into the oocyte using the 4X objective, and fluorescence is recorded using the water-immersion 40X objective. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/55598/55598fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. VCF Recording
<ol>
	<li>Follow preparation steps for cut-open oocyte voltage-clamp as previously described and visualized18 (agar bridge preparation, mounting the oocyte, saponin permeabilization). However, work under red light at all time to avoid bleaching the fluorophore prior to recordings. When placing the oocyte, make sure that the animal pole faces upwards. The pigmentation under the animal pole membrane shields against autofluorescence originating from the cytosol and therefore reduces background fluorescence.</li>
	<li>Slide the chamber over to the microscope and focus using a 4X objective.</li>
	<li>Impale the oocyte with the voltage sensing V1 electrode (3 M KCl), switch to the 40X water-immersion (NA 0.8 - 0.9) objective. Focus on the animal pole which is facing upwards.</li>
	<li>Turn off the red light. Select the right filter cube by turning the filter cube turret and the optical exit port connected to the photodiode. Turn on the halogen lamp at highest intensity and shortly switch the shutter open for 2-5 s to read the background fluorescence intensity originating from the oocyte. With the described setup the value should be around 50-200 pA for Anap.</li>
	<li>Turn on the clamp, flip the bath/guard switch to active and adjust the membrane potential (V1 - V2) to the command potential by turning the knob on the I headstage.</li>
	<li>Select holding potential, step protocol, number and length of pulses etc. in the recording software. Record voltage-dependent currents and Anap fluorescence intensities.</li>
</ol>
5. Two-color VCF
<ol>
	<li>To monitor two locations in the same protein simultaneously, mutate an extracellular and accessible amino acid into cysteine, and remove other cysteines to ensure specific labeling with thiol-chemistry.</li>
	<li>Perform step 2.1-2.5.</li>
	<li>Prior to VCF recordings, incubate oocytes in 5 &#181;M TMR-maleimide in labeling solution for 15 min (or other dye with non-overlapping spectra compared to Anap).</li>
	<li>Wash the oocytes with labeling solution three times to remove excess dye.</li>
	<li>Perform step 4.1-4.6.</li>
	<li>Insert an appropriate filter cube for TMR (excitation filter, dichroic mirror and emission filter) in the filter cube turret. Switch to the TMR filter cube by turning the filter turret.</li>
	<li>Read the background fluorescence for TMR as described for Anap in step 4.4. 
	NOTE: Labeling with thiol-chemistry results in high background fluorescence due to unspecific labeling in the membrane. Therefore, the TMR background fluorescence may saturate the amplifier (&#62;2,000 pA). In that case, do not decrease the light intensity, but simply subtract the background fluorescence by adding an offset current to the photodiode. In commercially available systems use the &#34;sample-and-hold&#34; feature on the detector system. Note the background fluorescence value (using a 10X neutral density filter) in a laboratory journal, as this value will not be recorded (saturation).</li>
	<li>Record voltage-dependent currents and TMR fluorescence intensities simultaneously as in step 4.6.</li>
</ol>

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The authors have nothing to disclose.

The in vivo aminoacylation of tRNAs which are continuously being transcribed together with the tRNA-synthetase, makes it possible to obtain high expression levels for fluorescence measurements. For efficient fUAA incorporation, it is critical that pAnap is correctly injected into the nucleus. Due to the uncertainty of the exact location of the nucleus, 10-40% of the DNA injections are expected to fail, resulting in non-expressing (or leak-expressing) oocytes. Therefore, it is important to check expression in abs...

Over 200 unnatural amino acids of various chemical and physical properties have been genetically incorporated into proteins in E. coli, yeast and mammalian cells3. The unnatural amino acid is incorporated in response to a specific stop codon via an orthogonal engineered tRNA/synthetase pair. The genetical approach to modify proteins has provided valuable insights into protein structure and function. Here, we present a protocol for using Voltage-Clamp Fluorometry (VCF) in combination with a fluorescent UAA.
In VCF, the simultaneous observation of functional data and structural rearrangements localized around the fluorescent probe (~5 &#197;) allows us to obtain dynamic information with millisecond resolution1. The fluorescent probes alter their quenching state upon localized movement of the protein. A movement of only 1-2 &#197; is sufficient to lead to significant changes in the fluorescence intensity4. After identification of the site of interest in the target protein, the site is mutated by point mutation. Classically, the residue had been mutated to a cysteine whereas now, an amber stop codon (TAG) is introduced for genetic fUAA incorporation. The protein is then in vitro transcribed.
While other expression systems (e.g.,&#160;mammalian cells) can be used5,6,7, Xenopus oocytes are preferable for structure-function studies because of their larger size, leading to easier manipulation and higher fluorescence intensity (more fluorophores) and, therefore, to-noise ratio. Furthermore, Xenopus oocytes have low background from endogenous proteins2,8, and the dark pigmentation on the animal pole shields against background fluorescence from the cytosol. The Xenopus oocytes are surgically removed and DNA encoding the orthogonal tRNA/tRNA-synthetase pair specific for the fUAA is injected into the nucleus of the oocytes. After a 6-24 h incubation time, the protein RNA is co-injected with the fUAA into the cytosol of the oocytes, followed by a 2-3 days incubation period. In order to prevent any damage to the fUAA (photobleaching), the procedures including Anap have to be carried out under red light to avoid fluorophore excitation.
Oocytes are studied on a cut-open oocyte voltage-clamp setup, which is mounted on an upright fluorescence microscope, and electrical current and fluorescence changes are simultaneously recorded9,10. Alternatively, two-electrode voltage clamp1 or patch-clamp configurations11can be used. Fluorescence is excited by appropriate wavelengths with low RMS noise and emission recorded using a photodiode linked to an amplifier with high amplification.
There are several advantages of using fluorescent unnatural amino acids (fUAAs) in voltage-clamp fluorometry. One is access to the cytosolic side of the membrane proteins; many regulatory processes are located here (e.g., Ca2+- or nucleotide binding sites, fast and closed-state inactivation of voltage-gated ion channels, pore opening, module coupling). All these processes are now accessible for fluorescent labeling.
Another advantage is the small size of the probe leading to less disturbance of the protein. So far, two orthogonal tRNA/tRNA synthetase pairs for fUAAs have been engineered12,13, where 3-(6-acetylnaphthalen-2-ylamino)-2-aminopropanoic acid (Anap) is the only fUAA which has been used in Xenopus oocytes2,8. Anap is an environmentally sensitive fluorophore with a molecular weight of 272.3 g/mol and is only slightly bigger than tryptophan12 (Figures 1A, 1B). Due to its small size, fewer steric effects are likely to be introduced by the fluorophore compared to conventional fluorophores attached via a linker (typically more than 500 g/mol). Moreover, in the case of Anap, the fluorophore is located closer to the protein backbone than those linked to cysteines, and consequently, Anap is probing more localized rearrangements. Finally, removal of endogenous cysteines in conventional VCF in order to ensure site-specific labeling is no longer a requirement in UAA-VCF and therefore (i) leaves the proteins in (almost) their native state and (ii) allows VCF to be applied to study a wider range of proteins in which function may be altered by cysteine substitution.
<img alt="Figure 1" src="/files/ftp_upload/55598/55598fig1.jpg" /> 
Figure 1: Anap and Fluorescence Spectra. (A) Chemical structure of Anap. (B) Normalized absorption spectrum and emission spectra for 1 nM Anap, demonstrating the sensitivity of Anap fluorescence to the solvent hydrophobicity. Emission spectra were obtained by exciting at 350 nm. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/55598/55598fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
A disadvantage of using fluorescent UAAs is that a heterogeneous population of proteins may result from stop codon readthrough, translational reinitiation, C-terminal truncated proteins or crosstalk with endogenous aminoacylation if the amount of aminoacylated tRNAs is scarce. Such leak expression should always be checked for in the absence of the fUAA and the tRNA/tRNA synthetase pair. We addressed the issue of translational reinitiation and how to circumvent it for N-terminal insertion sites previously14. However, when the fUAA, tRNA and tRNA synthetase are present in saturated amounts, there only remains a low probability of leak expression.
The key procedural difference between fUAA-VCF and conventional VCF is the injection and handling of the oocytes; the injection of DNA encoding the tRNA and tRNA synthetase (pAnap) is followed by the introduction of Anap, which is either co-injected with the protein mRNA or alternatively added to the incubation solution as an acetoxymethyl (AM) ester.

<table><tbody><tr><td>&lt;strong&gt;Solutions&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Barth&#039;s solution&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>NaCl&amp;nbsp;</td><td>Sigma-Aldrich</td><td>S7653</td><td>90 mM</td></tr><tr><td>KCl</td><td>Fisher Scientific</td><td>BP366-500</td><td>3 mM</td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>M-9397</td><td>0.82 mM</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>C-7902</td><td>0.41 mM</td></tr><tr><td>Ca(NO&lt;sub&gt;3&lt;/sub&gt;)&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>C-1396</td><td>0.33 mM</td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H4034</td><td>5 mM</td></tr><tr><td>NaOH hydrate</td><td>BDH</td><td>BDH7225-4</td><td>pH 7.6</td></tr><tr><td>Penicilin</td><td>Invitrogen</td><td>15140122</td><td>100 U/mL</td></tr><tr><td>Streptomycin</td><td>Invitrogen</td><td>15140122</td><td>100 &amp;micro;g/mL</td></tr><tr><td>Kanamycin</td><td>Invitrogen</td><td>15160054</td><td>10 mg/100mL</td></tr><tr><td>Horse Serum (HS)</td><td>Invitrogen</td><td>16050122</td><td>5%</td></tr><tr><td>&lt;strong&gt;SOS Standard Oocyte Solution&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>NaCl&amp;nbsp;</td><td>Sigma-Aldrich</td><td>746398</td><td>102 mM</td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>746436</td><td>3 mM</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>M9272</td><td>1 mM</td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H4034</td><td>5 mM</td></tr><tr><td>&lt;strong&gt;External recording solution&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>N-methyl-D-glucamine (NMDG)</td><td>Alfa Aesar</td><td>L14282</td><td>115 mM</td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H4034</td><td>10 mM</td></tr><tr><td>Calcium hydroxide</td><td>Sigma-Aldrich</td><td>239232</td><td>2 mM</td></tr><tr><td>MES hydrate</td><td>Sigma-Aldrich</td><td>258105</td><td>pH 7.2</td></tr><tr><td>&lt;strong&gt;Internal recording solution&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>N-methyl-D-glucamine (NMDG)</td><td>Alfa Aesar</td><td>L14282</td><td>115 mM</td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H4034</td><td>10 mM</td></tr><tr><td>Ethylenediamine Tetraacetic Acid (EDTA)</td><td>Fisher Scientific</td><td>E478-500</td><td>2 mM</td></tr><tr><td>MES hydrate</td><td>Sigma-Aldrich</td><td>258105</td><td>pH 7.2</td></tr><tr><td>&lt;strong&gt;Labeling solution&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>KOH</td><td>Fisher Scientific</td><td>P250-1</td><td>115 mM</td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H4034</td><td>10 mM</td></tr><tr><td>Calcium hydroxide</td><td>Sigma-Aldrich</td><td>239232</td><td>2 mM</td></tr><tr><td>MES hydrate</td><td>Sigma-Aldrich</td><td>258105</td><td>pH 7.2</td></tr><tr><td>&lt;strong&gt;TMR stock solution&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Tetramethylrhodamine-5-maleimide (TMR)</td><td>Molcular Probes by Life Technologies</td><td>T6027</td><td>5 mM in DMSO</td></tr><tr><td>&lt;strong&gt;Anap stock solution&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Anap</td><td>ABZENA (TCRS)</td><td>Custom synthesis TCRS-170</td><td>1 mM in nuclease-free water and 1% NaOH 1 N</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Material/Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>pAnap</td><td>Addgene</td><td>48696</td><td></td></tr><tr><td>High Performance Oocyte Clamp</td><td>Dagan Corporation</td><td>CA-1B</td><td></td></tr><tr><td>Gpatch Acquisition software</td><td>Department of Anesthesiology, University of California, Los Angeles</td><td></td><td></td></tr><tr><td>Analysis software</td><td>Department of Anesthesiology, University of California, Los Angeles</td><td></td><td></td></tr><tr><td>Recording Chamber</td><td>Custom machined</td><td></td><td></td></tr><tr><td>Photo diode detection system</td><td>Dagan Corporation</td><td>PhotoMax-200/PIN</td><td></td></tr><tr><td>Electrical shutter driver</td><td>UNIBLITZ</td><td>VCM-D1</td><td></td></tr></tbody></table>

voltage-clamp fluorometry in xenopus oocytes using fluorescent unnatural amino acids

Voltage-Clamp Fluorometry (VCF) has been the technique of choice to investigate the structure and function of electrogenic membrane proteins where real-time measurements of fluorescence and currents simultaneously report on local rearrangements and global function, respectively1. While high-resolution structural techniques such as cryo-electron microscopy or X-ray crystallography provide static images of the proteins of interest, VCF provides dynamic structural data that allows us to link the structural rearrangements (fluorescence) to dynamic functional data (electrophysiology). Until recently, the thiol-reactive chemistry used for site-directed fluorescent labeling of the proteins restricted the scope of the approach because all accessible cysteines, including endogenous ones, will be labeled. It was thus required to construct proteins free of endogenous cysteines. Labeling was also restricted to sites accessible from the extracellular side. This changed with the use of Fluorescent Unnatural Amino Acids (fUAA) to specifically incorporate a small fluorescent probe in response to stop codon suppression using an orthogonal tRNA and tRNA synthetase pair2. The VCF improvement only requires a two-step injection procedure of DNA injection (tRNA/synthetase pair) followed by RNA/fUAA co-injection. Now, labelling both intracellular and buried sites is possible, and the use of VCF has expanded significantly. The VCF technique thereby becomes attractive for studying a wide range of proteins and &#8211; more importantly &#8211; allows investigating numerous cytosolic regulatory mechanisms.

Figure 4 shows an example of VCF recordings obtained from an oocyte expressing Shaker channels with fast inactivation removed (IR), L382stop-W434F in presence of pAnap and Anap. The W434F mutation blocks the ionic potassium currents, which makes it possible to measure the transient gating charge displacements (gating currents). The simultaneous recordings of gating currents (upper trace) and Anap fluorescence intensity changes (lower trace) upon depolarization demonstrate...

Watch this Scientific Journal Video about Voltage-clamp Fluorometry in Xenopus Oocytes Using Fluorescent Unnatural Amino Acids at JoVE.com

Voltage-clamp Fluorometry in Xenopus Oocytes Using Fluorescent Unnatural Amino Acids

This article describes an enhancement of conventional Voltage-Clamp Fluorometry (VCF) where Fluorescent Unnatural Amino Acids (fUAA) are used instead of maleimide dyes, to probe structural rearrangements in ion channels. The procedure includes Xenopus oocyte DNA injection, RNA/fUAA coinjection, and simultaneous current and fluorescence measurements.

Voltage-clamp Fluorometry in Xenopus Oocytes Using Fluorescent Unnatural Amino Acids

Voltage-Clamp Fluorometry (VCF) has been the technique of choice to investigate the structure and function of electrogenic membrane proteins where ...

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Universidad de Cartagena UDEC

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Biochemistry

10.4K Views.  University of Montréal. This article describes an enhancement of conventional Voltage-Clamp Fluorometry (VCF) where Fluorescent Unnatural Amino Acids (fUAA) are used instead of maleimide dyes, to probe structural rearrangements in ion channels. The procedure includes Xenopus oocyte DNA injection, RNA/fUAA coinjection, and simultaneous current and fluorescence measurements.

Video: Voltage-clamp Fluorometry in Xenopus Oocytes Using Fluorescent Unnatural Amino Acids

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 ...

Biology

Use of pHluorin to Assess the Dynamics of Axon Guidance Receptors in Cell Culture and in the Chick Embryo

During development, axon guidance receptors play a crucial role in regulating axons sensitivity to both attractive and repulsive cues. Indeed, activation of the guidance receptors is the first step of the signaling mechanisms allowing axon tips, the growth cones, to respond to the ligands. As such, the modulation of their availability at the cell surface is one of the mechanisms that participate in setting the growth cone sensitivity. We describe here a method to precisely visualize the spatio-temporal cell surface dynamics of an axon guidance receptor both in vitro and in vivo in the developing chick spinal cord. We took advantage of the pH-dependent fluorescence property of a green fluorescent protein (GFP) variant to specifically detect the fraction of the axon guidance receptor that is addressed to the plasma membrane. We first describe the in vitro validation of such pH-dependent constructs and we further detail their use in vivo, in the chick spinal chord, to assess the spatio-temporal dynamics of the axon guidance receptor of interest.

We describe here the use of a pH-sensitive green fluorescent protein variant, pHluorin, to study the spatio-temporal dynamics of axon guidance receptors trafficking at the cell surface.&#160;The pHluorin-tagged receptor is expressed both in cell culture and in vivo, using electroporation of the chick embryo.

During development, axon guidance receptors play a crucial role in regulating axons sensitivity to both attractive and repulsive cues. Indeed, activation ...

Neuroscience

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

Spontaneous intracellular calcium activity can be observed in a variety of cell types and is proposed to play critical roles in a variety of physiological processes. In particular, appropriate regulation of calcium activity patterns during embryogenesis is necessary for many aspects of vertebrate neural development, including proper neural tube closure, synaptogenesis, and neurotransmitter phenotype specification. While the observation that calcium activity patterns can differ in both frequency and amplitude suggests a compelling mechanism by which these fluxes might transmit encoded signals to downstream effectors and regulate gene expression, existing population-level approaches have lacked the precision necessary to further explore this possibility. Furthermore, these approaches limit studies of the role of cell-cell interactions by precluding the ability to assay the state of neuronal determination in the absence of cell-cell contact. Therefore, we have established an experimental workflow that pairs time-lapse calcium imaging of dissociated neuronal explants with a fluorescence in situ hybridization assay, allowing the unambiguous correlation of calcium activity pattern with molecular phenotype on a single-cell level. We were successfully able to use this approach to distinguish and characterize specific calcium activity patterns associated with differentiating neural cells and neural progenitor cells, respectively; beyond this, however, the experimental framework described in this article could be readily adapted to investigate correlations between any time-series activity profile and expression of a gene or genes of interest.

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

We present a two-part protocol that combines fluorescent calcium imaging with in situ hybridization, allowing the experimenter to correlate patterns of calcium activity with gene expression profiles on a single-cell level.

Spontaneous intracellular calcium activity can be observed in a variety of cell types and is proposed to play critical roles in a variety of physiological ...

Developmental Biology

Imaging Membrane Potential with Two Types of Genetically Encoded Fluorescent Voltage Sensors

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate goal is to image neuronal activity in vivo, one must be able to image activity of a single cell to ensure successful in vivo preparations. This procedure will describe how to image membrane potential in a single cell to provide a foundation to eventually image in vivo. Here we describe methods for imaging GEVIs consisting of a voltage-sensing domain fused to either a single fluorescent protein (FP) or two fluorescent proteins capable of F&#246;rster resonance energy transfer (FRET) in vitro. Using an image splitter enables the projection of images created by two different wavelengths onto the same charge-coupled device (CCD) camera simultaneously. The image splitter positions a second filter cube in the light path. This second filter cube consists of a dichroic and two emission filters to separate the donor and acceptor fluorescent wavelengths depending on the FPs of the GEVI. This setup enables the simultaneous recording of both the acceptor and donor fluorescent partners while the membrane potential is manipulated via whole cell patch clamp configuration. When using a GEVI consisting of a single FP, the second filter cube can be removed allowing the mirrors in the image splitter to project a single image onto the CCD camera.

A method for imaging changes in membrane potential using genetically encoded voltage indicators is described.

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically carried out in low throughput. Rapid and ongoing expansion of induced pluripotent stem cell (iPSC) technology presents a new standard in cardiovascular research and alternate methods are now necessary to increase throughput of electrophysiological data at a single cell level. VF2.1Cl is a recently derived voltage sensitive dye which provides a rapid single channel, high magnitude response to fluctuations in membrane potential. It possesses kinetics superior to those of other existing voltage indicators and makes available functional data equivalent to that of traditional microelectrode techniques. Here, we demonstrate simplified, non-invasive action potential characterization in externally paced human iPSC derived cardiomyocytes using a modular and highly affordable photometry system.

Here we describe optical acquisition and characterization of action potentials from induced pluripotent stem cell derived cardiomyocytes using a high-speed modular photometry system.

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically ...

Bioengineering

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system. They provide multiple experimental advantages, such as accessibility to numerous imaging approaches, electrophysiological techniques and behavioral assays. The Xenopus tadpole olfactory system is particularly well suited to investigate the function of synapses established during normal development or reformed after injury. Here, we describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae. We outline a combination of in vivo measurements of presynaptic calcium responses in glomeruli of the olfactory bulb with olfactory-guided behavior assays. Methods can be combined with the transection of olfactory nerves to study the rewiring of synaptic connectivity. Experiments are presented using both wild-type and genetically modified animals expressing GFP reporters in central nervous system cells. Application of the approaches described to genetically modified tadpoles can be useful for unraveling the molecular bases that define vertebrate behavior.

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system in vivo. We describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae in normal rearing conditions or after injury.

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system. They provide multiple experimental advantages, such as ...

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 ...

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 ...

Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe dementia (FTLD) is dysfunction of communication between neurons and motor neurons and muscle. The underlying process of synaptic transmission involves membrane depolarization-dependent synaptic vesicle fusion and the release of neurotransmitters into the synapse. This process occurs through localized calcium influx into the presynaptic terminals where synaptic vesicles reside. Here, the protocol describes fluorescence-based live-imaging methodologies that reliably report depolarization-mediated synaptic vesicle exocytosis and presynaptic terminal calcium influx dynamics in cultured neurons.
Using a styryl dye that is incorporated into synaptic vesicle membranes, the synaptic vesicle release is elucidated. On the other hand, to study calcium entry, Gcamp6m is used, a genetically encoded fluorescent reporter. We employ high potassium chloride-mediated depolarization to mimic neuronal activity. To quantify synaptic vesicle exocytosis unambiguously, we measure the loss of normalized styryl dye fluorescence as a function of time. Under similar stimulation conditions, in the case of calcium influx, Gcamp6m fluorescence increases. Normalization and quantification of this fluorescence change are performed in a similar manner to the styryl dye protocol. These methods can be multiplexed with transfection-based overexpression of fluorescently tagged mutant proteins. These protocols have been extensively used to study synaptic dysfunction in models of FUS-ALS and C9ORF72-ALS, utilizing primary rodent cortical and motor neurons. These protocols easily allow for rapid screening of compounds that may improve neuronal communication. As such, these methods are valuable not only for the study of ALS but for all areas of neurodegenerative and developmental neuroscience research.

Two related methods are described to visualize subcellular events required for synaptic transmission. These protocols enable the real-time monitoring of the dynamics of presynaptic calcium influx and synaptic vesicle membrane fusion using live-cell imaging of in vitro cultured neurons.

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe ...

Voltage-clamp Fluorometry in Xenopus Oocytes Using Fluorescent Unnatural Amino Acids

Summary

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Chapters in this video

Patch Clamp Recording of Ion Channels Expressed in Xenopus Oocytes

Use of pHluorin to Assess the Dynamics of Axon Guidance Receptors in Cell Culture and in the Chick Embryo

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

Imaging Membrane Potential with Two Types of Genetically Encoded Fluorescent Voltage Sensors

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

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

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

Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis