Supported by the NIH grant SBC COBRE 1P20GM121310-01.

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Wyoming. All procedures, including electrophysiological recordings, are carried out at room temperature, approximately 23 &#176;C. All methods described here are optimized for recording tectal neurons from tadpoles between developmental stage 42 and 49 (staged according to Neiuwkoop and Faber15).
1. In Vivo Preparation
<ol>
	<li>Anesthetize the tadpole.
	<ol>
		<li>Place the tadpole in a small Petri dish containing Steinberg&#39;s solution with 0.01% MS-222 for approximately 5 min. 
		&#8203;NOTE: MS-222 aka &#34;Tricaine&#34; is a common fish and amphibian anesthetic. The tadpoles are raised in the Steinberg&#39;s solution in mM: 0.067 KCl, 0.034 Ca(NO3)2&#8226;4H2O, 0.083 MgSO4&#8226;7H2O, 5.8 NaCl, 4.9 HEPES.</li>
		<li>Ensure the tadpole is deeply anaesthetized (non-responsive and no longer swimming) before proceeding to step 2.</li>
	</ol>
	</li>
	<li>Secure the anesthetized tadpole to a submerged silicone block on the floor of the dissecting/recording dish.
	<ol>
		<li>Use a disposable transfer pipette to move the anaesthetized tadpole to a dissection/recording dish containing external recording solution (115 mM of NaCl, 2 mM of KCl, 3 mM of CaCl2, 3 mM of MgCl2, 5 mM of HEPES, 1 mM of glucose; adjust pH to 7.25 using 10 N of NaOH, osmolarity 255 mOsm).
		<ol>
			<li>To minimize spontaneous muscle twitches, add the acetylcholine receptor blocker, tubocurarine (100 &#181;M) to the external solution. (Typically, this is done by using a 200 &#181;L pipette to add 100 &#181;L of a 10 mM tubocurarine stock to 10 mL of external solution).</li>
		</ol>
		</li>
		<li>Using insect pins, secure the tadpole, dorsal side up, to a submerged block of silicone elastomer (such as Sylgard 184, see Materials Table for details), which has been glued to the dissecting dish floor. 
		Note: The placement of the pins is critical: place them on either side of the brain, sufficiently caudal to avoid impaling the afferent RGC axons that project from the eye and enter the brain just anterior to the optic tectum (Figure 1A).</li>
	</ol>
	</li>
	<li>Fillet the brain along the midline.
	<ol>
		<li>For a clear view of the brain, remove the skin overlying the brain by making a superficial incision along the midline using a sterile 25-G needle.</li>
		<li>Fillet the brain along the same midline axis by inserting the needle into the neural tube and pulling gently upward (dorsally) such that the dorsal portion of the tube is cleanly cut (severed) while leaving the floor plate intact (Figure 1B). 
		&#8203;NOTE: It is important that the floor plate of the neural tube be left intact because the afferent sensory inputs enter the tectum via the floor plate.</li>
	</ol>
	</li>
	<li>Remove the transparent ventricular membrane that covers the tectal neurons.
	<ol>
		<li>Move the recording dish to the electrophysiology rig and use a broken glass pipette tip to remove the ventricular membrane overlying the tectal neuron cell bodies. Break the tip by lightly dragging it across a delicate task wiper.</li>
		<li>Screw the pipette into the pipette holder and lower it down to the optic tectum via the micromanipulator.</li>
		<li>Use the broken pipette to peel the ventricular membrane away from the tectum. 
		NOTE: It is not necessary to remove the membrane from the entire tectum; a small window will suffice and provide access to plenty of neurons.</li>
	</ol>
	</li>
	<li>Obtain the whole cell patch clamp recordings. 
	NOTE: The approach from this point forward is similar to carrying out whole cell patch clamp recordings from mouse brain slices as described in Segev, et al.16(Figure 1C).</li>
	<li>Record the tectal neuron responses to a whole field flash of light projected onto the retina.
	<ol>
		<li>To project a whole field flash of light onto the retina, place an optic fiber adjacent to the tadpole&#39;s eye. At the other end of the optic fiber is an LED in line with a variable resistor that allows for the light luminance to be controlled. Trigger the LED by the digital output of the amplifier. In this way, whole field flashes of varying intensities of light can be recorded (Figure 4A).</li>
	</ol>
	</li>
</ol>
2. Whole Brain Preparation
<ol>
	<li>Perform steps 1.1 to 1.3. 
	Note: There is no need for tubocurarine in the external solution for this preparation.</li>
	<li>Isolate the brain.
	<ol>
		<li>Use a 25-G needle to sever the hindbrain (Figure 2A).</li>
		<li>To isolate the whole brain from the tadpole, gently run the needle underneath the brain, in a caudal-to-rostral direction to sever all lateral and ventral connective tissue and nerve fibers.</li>
	</ol>
	</li>
	<li>Secure the brain to a block of silicone elastomer.
	<ol>
		<li>Once completely freed, secure the brain to a block of silicon elastomer by placing one pin through one of the olfactory bulbs and another pin through the hindbrain (Figure 2B). This is the optimal configuration for recording from tectal neurons.</li>
	</ol>
	</li>
	<li>Move the dish containing the pinned whole brain preparation from the dissecting scope to the electrophysiology rig. Remove the ventricular membrane using a broken glass pipette as described in step 1.4.</li>
	<li>Place a bipolar stimulating electrode on the optic chiasm (where the axon tracts from each eye cross at the midbrain) to directly activate the RGC axons.
	<ol>
		<li>Place the bipolar stimulating electrode rostral, and almost adjacent to, the large middle ventricle. The optic chiasm is located immediately rostral to the large middle ventricle (Figure 2B).
		<ol>
			<li>Gently lower the stimulating electrode down onto the optic chiasm such that a small dent is formed in the tissue. The bipolar electrode is driven by a pulse stimulator which allows the strength of stimulation to be precisely controlled.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Perform the whole cell patch clamp recording (Figure 2C) as described by Segev, et al.16</li>
</ol>
3. Horizontal Brain Slice Preparation
<ol>
	<li>Perform steps 2.1 to 2.3.</li>
	<li>Use a razor blade to excise the most lateral fourth (which in vivo corresponds to the most dorsal fourth) of one side of one optic tectum. This cut is made parallel to the rostral-caudal plane as shown in Figure 3A.</li>
	<li>Re-pin the brain to the side of the silicone elastomer with the sliced side facing up (so that the somata and neuropil can be directly accessed for recording) and the ventricular surface of the brain facing away from the silicone elastomer block (Figure 3B) (so that a bipolar electrode can be placed on the optic chiasm).</li>
	<li>Perform the whole cell patch clamp or local filed potential recording (Figure 3C) as described by Segev, et al.16</li>
</ol>

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The authors have nothing to disclose.

All methods described in this work are optimized for recording tectal neurons from tadpoles between developmental stage 42 and 49 (staged according to Neiuwkoop and Faber15). By stage 42, the tadpoles are sufficiently large and sufficiently developed so that the insect pins can be placed on either side of brain for in vivo recordings and for carrying out the whole brain dissection. At earlier stages, when the tadpoles are essentially two-dimensional (i.e., flat), the approaches described here are not optimal.
Due to the ease in which tectal neurons can be accessed and recorded in whole cell configuration, this model circuit has been a hotbed for fundamental discoveries about how neural circuits form while functioning - and function while forming19. Here we have described how to acquire whole cell patch clamp recordings from tectal neurons, including how to perform the tadpole in vivo and whole brain preparations, the different recording configurations, and examples of the different types of data. The following is a discussion of the strengths and limitations of each preparation, followed by tips for achieving high-quality whole cell recordings.
Xenopus tadpoles begin displaying visual avoidance behaviors by stage 47/48, approximately 7 - 8 days post-fertilization3,13. This means that even at these relatively early stages of development, the retinotectal circuit is functioning and capable of processing visual stimuli. Fundamental questions regarding the development and plasticity of the retinotectal circuit have been directly addressed using the in vivo preparation to record tectal neuron responses to visual stimuli projected onto the retina. In vivo recordings in the tadpole are relatively straightforward compared to the equivalent recording configuration in mammals due to the relative simplicity of the nervous system and lack of a skull. Because individual tectal neuron somas can be visualized in vivo, sealing onto a given neuron and achieving whole cell patch clamp recording configuration is essentially equivalent to carrying out whole cell patch clamp recording in rodent slice preparations. Tadpoles typically survive for several hours on the rig (which can be readily assessed by monitoring the heart beat). One limitation inherent of the tadpole in vivo preparation is that only tectal neurons residing in the deep somatic layer of the optic tectum, the layer that is immediately adjacent to the large middle ventricle, are accessible for recording (Figure 1). This means that tectal neurons residing in the outer somatic layers of the tectum have been largely unexplored in vivo.
The experimental advantage of the whole brain preparation is that it isolates the brain from the peripheral sensory receptors in the eyes and skin, eliminating all spurious sensory-driven activity. While no longer capable of being driven by visual stimuli, the RGC axonal inputs that target the tectum are still present in the whole brain preparation and they can be directly activated in a highly controlled manner via a bipolar stimulating electrode (FHC) placed on the optic chiasm where the two sets of RGC axon tracts enter the brain. The first reported use of the whole brain preparation was in 1996, in an important study by Wu&#160;et al.8, in which RGC axons were activated using a bipolar stimulating electrode such that the authors could quantify the strength of individual RGC axons onto postsynaptic tectal neurons, and in doing so made fundamental discoveries about the progression of synapse maturation. As with the in vivo preparation, the whole brain preparation allows access to only those neurons residing in the deep layer, adjacent to the ventricular membrane.
Although the optic tectum is composed of approximately nine cell body layers17, all previous electrophysiology studies have focused on the deep layer tectal neurons because the traditional whole brain and in vivo preparations (described above) allow access only to the neurons of the deep layer. In order to access tectal neurons across all somatic layers, from deepest to most superficial as well as the entire neuropil where RGC axons form synaptic connections with tectal neuron dendrites, we developed a modified whole brain preparation, referred to as &#34;the horizontal brain slice preparation&#34; (Figure 314). As with the whole brain preparation, the horizontal brain slice preparation maintains the RGC axons within the brain and they can be controlled by placing a bipolar stimulating electrode on the optic chiasm.
Overall, the preparations are designed to optimize visualization of and access to tectal neuron somata to carry out whole cell patch clamp recording. Direct access to the soma is essential for quality whole cell patch clamp recordings: access to the inside of the neuron requires the formation of a very strong (&#62;1 &#215; 109 &#937;) seal between the pipette and the neuron&#39;s plasma membrane, which requires direct contact of the pipette tip onto that membrane. Hence, sufficient removal of the ventricular membrane is key for successful recording. Also, key for successful RGC-evoked recordings is the correct placement of the bipolar stimulating electrode onto the optic chiasm. Essentially all tectal neurons receive direct synaptic input from the RGC axons. If no RGC-evoked response is observed, then this is most likely due to incorrect placement of the bipolar electrode such that it is not on the optic chiasm and not contacting the optic nerves. This can be corrected by simply re-positioning the stimulating electrode. Failure to observe RGC-evoked responses after re-positioning the bipolar electrode could mean that the RGC axon projection was accidentally severed during the dissection. In this case, it is necessary to start over with a new preparation. Finally, failure to observe an RGC-evoked response could be due to recording from a cell that is not a common tectal neuron. For instance, glia typically do not display the normal RGC-evoked monosynaptic response displayed by tectal neurons, and neither do the large mesencephalic neurons that have been found, albeit at very low numbers, residing in the optic tectum20. The likelihood of recording from one of these non-tectal cells is very low due to the sheer number of tectal neurons. Of course, quality recordings also require the tectal neurons to be healthy. Characteristics of a healthy tectal neuron are a compact round or oval-shaped soma that is light-colored and without blemishes. Once whole cell recording configuration has been achieved, the health of the neuron can be further evaluated at the electrophysiological level by measuring fundamental electrical properties such as resting membrane potential, input resistance, and capacitance. The resting membrane potential of healthy tectal neurons between developmental stages 42 and 49 is approximately -45 mV, with an average input resistance of approximately 1 G&#937;, and a capacitance of 10 - 12 pF3,4,5. A small fraction of neurons will inevitably be damaged due to the removal of the ventricular membrane. The somata of damaged or unhealthy neurons appear grainy, bloated, or shriveled. An example of an unhealthy neuron is shown Figure 2C. If the majority of neurons appear unhealthy, this could be a reflection that there is something wrong with the external recording solution, so it is recommended to make up fresh external recording solution. Besides appearing healthy, the best neurons for recording are typically associated with a large number of other neurons, as opposed to being isolated. Also, choose cells that are easily accessible. For example, do not push the tissue too hard to access a cell because this might damage the tissue or move the tissue and thus shift the stimulating electrode out of place. Finally, recordings should be restricted to the middle third of the tectum (Figure 2B) to avoid potential differences due to the developmental gradient that exists across the rostro-caudal axis4,8, unless, of course, the focus of the experiment is the aforementioned developmental gradient. The tectum proper is the area rostral of the proliferative zone and caudal to the caudal edge of large middle ventricle.
The whole cell patch clamp recording technique is not without its limitations. In general, this style of recording involves recording one neuron at a time. This is in contrast to approaches involving imaging of a new generation GCaMP6 calcium indicators, which allow for the activity of large populations of neurons to be imaged simultaneously in a slice preparation and in vivo.
Another drawback inherent to whole cell voltage clamp recordings is the space clamp error (the inability to control the voltage much further beyond the soma), which limits temporal control of voltage4. Overall, these voltage clamp issues are minimal when recording from tectal neurons however, because of their relatively small size, modest dendritic arborizations, and relatively small and slow currents compared to mammalian neurons. These characteristics of tectal neurons render them especially amenable for voltage clamp recording, measurement of many parameters, and construction of profiles for individual neurons5,6.
Future directions include optimizing an in vivo horizontal brain slice preparation to characterize the function of outer layer neurons in the processing of visual stimuli.

The retinotectal circuit is the major component of the amphibian visual system. It is comprised of the RGCs in the eye, which project their axons to the optic tectum where they form synaptic connections with postsynaptic tectal neurons. The Xenopus tadpole retinotectal circuit is a popular developmental model to study neural circuit formation and function. There are many attributes of this tadpole&#39;s retinotectal circuit that render it a powerful experimental model1,2,3. One major attribute, and the focus of this article, is the ability to carry out whole cell patch clamp recordings from tectal neurons, in vivo or using a whole brain preparation. With an electrophysiology rig outfitted with an amplifier that supports voltage- and current-clamp recording modes, whole cell patch clamp recordings allow a neuron&#39;s electrophysiology to be characterized at high resolution. As a result, whole cell patch clamp recordings from tectal neurons across the key stages of retinotectal circuit formation have provided a detailed and comprehensive understanding of the development and plasticity of intrinsic4,5,6,7 and synaptic8,9,10,11 properties. Combining whole cell patch clamp tectal neuron recordings, the ability to express genes or morpholinos of interest in these neurons12, and a method to assess visual guided behavior via an established visual avoidance test13 promotes the identification of links between molecules, circuit function, and behavior.
It is important to note that the type of high resolution data acquired from whole cell patch clamp recordings is not possible using newer imaging approaches such as the genetic calcium indicator GCaMP6, because although using calcium indicators permits the imaging of calcium dynamics across large populations of neurons simultaneously, there is no direct or obvious way that the specific electrical parameters can be obtained by measuring delta fluorescence in the somata, and there is no way to voltage clamp the neuron to measure current-voltage relationships. Clearly these two distinct approaches, electrophysiological recordings and calcium imaging, possess non-overlapping strengths and generate different types of data. Thus, the best approach depends on the specific experimental question being addressed.
Here, we describe our method for acquiring whole cell patch clamp recordings from neurons of the tadpole optic tectum using an in vivo preparation, whole brain preparation, and a newer modified whole brain preparation that was developed in our lab14. In the Representative Results section, we demonstrate the experimental advantages of each preparation and the different types of data that can be obtained. The limits and strengths of the different preparations, as well as tips for troubleshooting, are included in the Discussion section.

<table border="0" cellpadding="0" cellspacing="0" style="width:1012px;" width="1011">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:423px;">Stemi Stereo 508</td>
			<td style="width:143px;">Zeiss</td>
			<td style="width:148px;">495009-0006-000&#160;</td>
			<td style="width:299px;">Dissecting microscope</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MS-222 &#34;Tricane&#34;</td>
			<td>Finquel</td>
			<td>ARF5G</td>
			<td>Amphibian general anesthetic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium Chloride (NaCl)</td>
			<td>Fisher Scientific</td>
			<td>S271-3</td>
			<td>Used to prepare Stienberg&#39;s solution and external solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Potassium Chloride (KCl)</td>
			<td>Fisher Scientific</td>
			<td>P217-500</td>
			<td>Used to prepare Stienberg&#39;s solution and external solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">HEPES</td>
			<td>Sigma-Aldrich</td>
			<td>H3375-1KG</td>
			<td>Used to prepare Stienberg&#39;s solution and external solution</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:26px;">Calcium nitrate tetrahyrate (Ca(NO3)&#8226;4H2O)</td>
			<td>Sigma-Aldrich</td>
			<td>237124-500G</td>
			<td>Used to prepare Stienberg&#39;s solution&#160;&#160;</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:26px;">Magnesium Sulfate (MgSO4)</td>
			<td>Mallinckrodt Chemicals</td>
			<td>6066-04</td>
			<td>Used to prepare Steinberg&#39;s solution</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:26px;">Calcium Chloride (CaCl2)</td>
			<td>Sigma-Aldrich</td>
			<td>C5080-500G</td>
			<td>Used to prepare external recording solution</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:26px;">Magnesium Chloride (MgCl2)</td>
			<td>J.T. Baker</td>
			<td>2444-01</td>
			<td>Used to prepare external recording solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">D-glucose Anhydrous</td>
			<td>Mallinckrodt Chemicals</td>
			<td>6066-04</td>
			<td>Used to prepare external recording solution</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Tubocurarine hydrochloride pentahydrate</td>
			<td>Sigma</td>
			<td>T2379</td>
			<td>Nicotinic acetylcholine receptor antagonist</td>
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			<td height="21" style="height:21px;">Insect Pins</td>
			<td>Fine Science Tools</td>
			<td>26002-10</td>
			<td>0.1mm diameter stainless steel pins</td>
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			<td height="21" style="height:21px;">Sylgard 184 Silicone Elastomer Kit</td>
			<td>Dow Corning</td>
			<td>761028</td>
			<td>Preweighed monomer and curing agent kit</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Sterile Polystyrene Petri Dish - 60x15mm</td>
			<td>Fisher Scientific</td>
			<td>AS4052</td>
			<td>Small petri dishes</td>
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		<tr height="21">
			<td height="21" style="height:21px;">PrecisionGlide Needle 25Gx5/8 (.0.5mm X 16mm)</td>
			<td>BD</td>
			<td>305122</td>
			<td>Syringe needles</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">1mL Slip Tip Tuberculin Syringe&#160;</td>
			<td>BD</td>
			<td>309659</td>
			<td>Disposable, sterile syringes</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Borosilicate pipette glass</td>
			<td>Sutter Instrument</td>
			<td>BF150-86-10HP</td>
			<td>Pulled to desired specifications using pipette pulling machine</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Flaming/Brown Micropipette Puller</td>
			<td>Sutter Instruments</td>
			<td>P-97</td>
			<td>Fabricates micropipettes for electrophysiology recording</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Kimwipes Kimtech wipes</td>
			<td>Kimberly-Clark</td>
			<td>34120</td>
			<td>Delicate task lint-free wipers</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Axon Instruments MultiClamp 700B Headstage CV-7B</td>
			<td>Molecular Devices</td>
			<td>1-CV-7B</td>
			<td>Current clamp and voltage clamp headstage</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MP-285 Motorized Manipulator with Tabletop Controller</td>
			<td>Sutter Instrument</td>
			<td>MP-285/T</td>
			<td>Control for headstage on electrophysiology rig</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fiber-Coupled LED (Green)</td>
			<td>Thorlabs</td>
			<td>M530F2</td>
			<td>Fiber optic cable paired with green LED</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cluster Bipolar Electrode (25&#181;m diameter)</td>
			<td>FHC</td>
			<td>30207</td>
			<td>Bipolar stimulating electrode</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">ISO-Flex Stimulator</td>
			<td>A.M.P.I. (Israel)&#160;</td>
			<td>Contact manufacturer</td>
			<td>Flexible stimulus isolator</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Axon Instruments 700B Multipatch Amplifier</td>
			<td>Molecular Devices</td>
			<td>2500-0157</td>
			<td>Amplifier for voltage- and current-clamp recording&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Digidata 1322A digitizer</td>
			<td>Molecular Devices</td>
			<td>2500-135</td>
			<td>Data acquisition system for electrophysiology recording</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Axio Examiner.A1</td>
			<td>Zeiss</td>
			<td>491404-0001-000&#160;</td>
			<td>Microscope for electrophysiology</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Micro-g Lab Table</td>
			<td>TMC</td>
			<td>63-533</td>
			<td>Air table for electrophysiology microscope</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Inspiron 620 Personal Desktop Computer with Windows 7 64-bit</td>
			<td>Dell</td>
			<td>D06D001</td>
			<td>Computer running electrophysiology software</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">c2400 CCD camera</td>
			<td>Hamamatsu</td>
			<td>70826-5</td>
			<td>Charge-coupled device camera for electrophysiology imaging</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">7 O&#39;Clock Super Platinum Stainless Razorblades</td>
			<td>Gillette</td>
			<td>CMM01049</td>
			<td>Platinum-coated stainless razor blades</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Transfer Pipets</td>
			<td>Fisher Scientific</td>
			<td>13-711-7M</td>
			<td>Disposable Polyethylene transfer pipets</td>
		</tr>
	</tbody>
</table>

preparations and protocols for whole cell patch clamp recording of xenopus laevis tectal neurons

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic tectum, is a popular model to study how neural circuits self-assemble. The ability to carry out whole cell patch clamp recordings from tectal neurons and to record RGC-evoked responses, either in vivo or using a whole brain preparation, has generated a large body of high-resolution data about the mechanisms underlying normal, and abnormal, circuit formation and function. Here we describe how to perform the in vivo preparation, the original whole brain preparation, and a more recently developed horizontal brain slice preparation for obtaining whole cell patch clamp recordings from tectal neurons. Each preparation has unique experimental advantages. The in vivo preparation enables the recording of the direct response of tectal neurons to visual stimuli projected onto the eye. The whole brain preparation allows for the RGC axons to be activated in a highly controlled manner, and the horizontal brain slice preparation allows recording from across all layers of the tectum.

To record light-evoked responses a whole field flash of light is projected onto the retina while the resulting response is recorded from individual tectal neurons (Figure 4A). This particular protocol is designed to measure both the response of the neuron to the light turning on (&#34;On&#34; response) and then turning off 15 s later to measure the &#34;Off response.&#34;&#160;Tectal neurons typically exhibit robust On and Off responses (shown here recorded in voltage clamp mode, with the neuron clamped to -60mV to measure synaptic current (Figure 4B)). The amount of synaptic drive received by any one neuron can be quantified by recording spontaneous synaptic events (Figure 4C). Current-voltage relationships (I-V curves) can be generated from the same neuron by stepping the neuron to increasingly depolarized potentials and recording the resulting voltage-activated Na+ and K+ currents, referred to as &#34;mixed current recordings&#34; (Figure 4D, E). It has been well established that for these amphibian neurons, peak Na+ and K+ currents can be quantified via mixed current recordings because the peaks do not temporally overlap with one another4,7.
Direct activation of RGC axons at the optic chiasm bypasses all upstream computation and processing of visual stimuli that take place in the eye, and allows for studying the synaptic transmission between presynaptic RGC axons and postsynaptic tectal neurons in its most pure and reduced form. Tectal neuron responses to RGC activation are generated from RGCs stimulated by light or directly using a bipolar stimulating electrode consisting of two components: a monosynaptic component (the direct synaptic input from activated RGCs) and a polysynaptic component (the local recurrent network activity feeding back onto the neuron being recorded). In the case of the light evoked responses, the monosynaptic and polysynaptic components overlap with one another by some undeterminable amount. This temporal overlap limits what can be concluded about synaptic transmission between the retina and optic tectum. Using a whole brain preparation and activating the RGC axon tract directly via an electrode placed on the optic chiasm (Figure 5A), however, produces a mono- and polysynaptic component that are essentially non-overlapping in time. The stimulating strength of the bipolar electrode can be adjusted. As the stimulating strength is increased, there is a greater number of RGC axons activated (Figure 5B). A minimal stimulation protocol determines the synaptic strength of an individual RGC axon onto an individual tectal neuron; a maximal stimulation protocol reflects the total, or maximum, synaptic input from all the RGC axons combined (Figure 5C). The proper ratio of excitatory to inhibitory synaptic input (E:I ratio) received by an individual tectal neuron is critical for normal visual function18. Figure 5D shows an example of the RGC-evoked excitatory and inhibitory current traces recorded from an individual tectal neuron. The probability of the transmitter release from RGC presynaptic axon terminals is measured by paired pulse recordings. For this, the postsynaptic tectal neuron is recorded in the voltage clamp mode to measure synaptic currents, while the RGC axons are activated successively, separated by a 50 ms time interval (Figure 5E).
Whole cell patch clamp recordings can be carried out from tectal neurons residing in any somatic layer. Similar to the whole brain preparation, RGC axons can be activated by placing a bipolar recording electrode on the optic chiasm (Figure 6A). All recordings carried out in the whole brain preparation can also be performed using this horizontal brain slice preparation. Figure 6B shows an example trace of an RGC-evoked response recorded from a neuron in the superficial layer. This preparation also allows for the spatial pattern of RGC synaptic input onto the tectal dendrites to be measured. For this, RGC-evoked field potentials are recorded every 10 &#181;m along the medial-lateral axis (i.e., the distal-proximal axis) of the neuropil (Figure 6C). This generates a high-resolution profile of the pattern of functional RGC input. The field potentials can then be transformed into current source densities by calculating the second spatial derivative14 and the current source densities can in turn be transformed into an image plot. The image plot in Figure 6C shows that the strongest RGC input targets the more distal area of the neuropil.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57465/57465fig1.jpg" /> 
Figure 1. In vivo preparation procedures. (A) Tadpole pinned down on a piece of silicone elastomer. Pins are indicated by the white arrows. Note the more caudal pin positioning to avoid impaling the optic tracts. The white dashed line indicates the midline. (B) Zoomed-in view of the entire brain after cutting along the dorsal midline and removing overlaying skin. (C) Whole-cell patch-clamp recording in vivo. (L: Lateral. M: Medial. R: Rostral. C: Caudal. D: Dorsal. V: Ventral. OB: Olfactory Bulb. OT: Optic Tectum. HB: Hindbrain). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57465/57465fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57465/57465fig2.jpg" /> 
Figure 2. Whole brain preparation procedures. (A) After the brain is filleted along the midline, the hindbrain is severed. (B) The entire brain is dissected out and pinned down to the silicone elastomer. The white asterisk indicates the pin in the left olfactory bulb. The red box indicates the middle third of the tectum, i.e., the target area for recording. (C) Whole cell patch clamp recording using whole brain preparation. The black asterisk indicates the area of un-scraped ventricular membrane where cells cannot be accessed for recording. Where the membrane has been successfully removed, the somata appear more distinct and defined. The arrow indicates an unhealthy neuron. (PZ: Proliferative Zone. LMV: Large Medial Ventricle. OB: Olfactory Bulb. OT: Optic Tectum. HB: Hindbrain. L: Lateral. M: Medial. R: Rostral. C: Caudal. D: Dorsal. V: Ventral). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57465/57465fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57465/57465fig3.jpg" /> 
Figure 3. Horizontal brain slice preparation procedures. (A) The most lateral portion of one side of the optic tectum will be sliced (indicated by the dashed line) after whole brain preparation. (B) Horizontal brain slice preparation after being pinned down to the side of the silicone elastomer. Double white lines represent a stimulating electrode placed on the optic chiasm. (C) Zoomed-in view of the soma layers and neuropil using a horizontal brain slice preparation. Dashed curve indicates the boundary between soma layers and neuropil. This figure has been modified from Hamodi, et al.14 (L: Lateral. M: Medial. R: Rostral. C: Caudal. D: Dorsal. V: Ventral. OB: Olfactory Bulb. OC: Optic Chiasm. OT: Optic Tectum. HB: Hindbrain). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57465/57465fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57465/57465fig4.jpg" /> 
Figure 4. Recordings from an in vivo preparation. (A) Schematic showing the typical configuration of an in vivo recording. (B) Light evoked responses including light on and light off responses. Inset traces are zoomed-in view of on- and off-response, respectively. (C) Spontaneous excitatory postsynaptic currents recorded. (D) Na+ and K+ currents in response to voltage clamp steps from -60 mV to 30 mV from a single neuron. (E) IV plots generated from traces in (D). All recordings were done with the neuron held at -60 mV except in (D) to generate IV-plots; neurons are stepped to increasingly more depolarized potentials, in 10 mV increments, to activate voltage-dependent currents. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57465/57465fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/57465/57465fig5.jpg" /> 
Figure 5. Recordings from a whole brain preparation. (A) Schematic showing the typical configuration of whole brain recording including a stimulating electrode placed on the optic chiasm (green and red lines represent RGC axons). (B) Overlaying traces of the RGC responses to changes of RGC stimulation strength. (C) Maximum stimulation response (left) and minimum stimulation response (right). (D) An RGC-evoked excitatory and inhibitory response onto a single neuron. (E) Paired pulse facilitation of the RGC-evoked response of an individual neuron. (OB: Olfactory Bulb. OC: Optic Chiasm. OT: Optic Tectum). All recordings were done with the neuron held at -60 mV except in (D) to measure the excitatory and inhibitory inputs: the excitatory input is recorded by holding the neuron at the reversal potential of &#947;-Aminobutyric acid (GABA)-mediated chloride currents (-45 mv); the inhibitory input by holding the neuron at the reversal potential of &#945;-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA)-mediated currents (+5 mv). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57465/57465fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/57465/57465fig6.jpg" /> 
Figure 6. Recordings from a horizontal brain slice preparation. (A) Schematic showing the configuration of a horizontal brain slice preparation. Note that although the stimulating electrode remains at the optic chiasm, the recording pipette is now positioned to access cells across all the tectal layers exposed by the horizontal brain slice preparation. (B) An RGC response from a neuron residing in the superficial layer of the tectum. (C) Recorded field potentials across the neuropil and the converted current source densities (CSDs) shown by image plot heat maps. (OC: Optic Chiasm. OT: Optic Tectum. HB: Hindbrain). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57465/57465fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons at JoVE.com

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

In this paper, we discuss three brain preparations used for whole cell patch clamp recording to study the retinotectal circuit of Xenopus laevis tadpoles. Each preparation, with its own specific advantages, contributes to the experimental tractability of the Xenopus tadpole as a model to study neural circuit function.

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic ...

preparations-protocols-for-whole-cell-patch-clamp-recording-xenopus

Nanyang Technological University

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JoVE Journal

Neuroscience

9.3K Views.  University of Wyoming. In this paper, we discuss three brain preparations used for whole cell patch clamp recording to study the retinotectal circuit of Xenopus laevis tadpoles. Each preparation, with its own specific advantages, contributes to the experimental tractability of the Xenopus tadpole as a model to study neural circuit function.

Video: Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

Neural Explant Cultures from Xenopus laevis

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

Neural Explant Cultures from Xenopus laevis

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

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

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

The process by which the anterior region of the neural plate gives rise to the vertebrate retina continues to be a major focus of both clinical and basic research. In addition to the obvious medical relevance for understanding and treating retinal disease, the development of the vertebrate retina continues to serve as an important and elegant model system for understanding neuronal cell type determination and differentiation1-16. The neural retina consists of six discrete cell types (ganglion, amacrine, horizontal, photoreceptors, bipolar cells, and M&uuml;ller glial cells) arranged in stereotypical layers, a pattern that is largely conserved among all vertebrates 12,14-18.
While studying the retina in the intact developing embryo is clearly required for understanding how this complex organ develops from a protrusion of the forebrain into a layered structure, there are many questions that benefit from employing approaches using primary cell culture of presumptive retinal cells 7,19-23. For example, analyzing cells from tissues removed and dissociated at different stages allows one to discern the state of specification of individual cells at different developmental stages, that is, the fate of the cells in the absence of interactions with neighboring tissues 8,19-22,24-33. Primary cell culture also allows the investigator to treat the culture with specific reagents and analyze the results on a single cell level 5,8,21,24,27-30,33-39. Xenopus laevis, a classic model system for the study of early neural development 19,27,29,31-32,40-42, serves as a particularly suitable system for retinal primary cell culture 10,38,43-45.
Presumptive retinal tissue is accessible from the earliest stages of development, immediately following neural induction 25,38,43. In addition, given that each cell in the embryo contains a supply of yolk, retinal cells can be cultured in a very simple defined media consisting of a buffered salt solution, thus removing the confounding effects of incubation or other sera-based products 10,24,44-45.
However, the isolation of the retinal tissue from surrounding tissues and the subsequent processing is challenging. Here, we present a method for the dissection and dissociation of retinal cells in Xenopus laevis that will be used to prepare primary cell cultures that will, in turn, be analyzed for calcium activity and gene expression at the resolution of single cells. While the topic presented in this paper is the analysis of spontaneous calcium transients, the technique is broadly applicable to a wide array of research questions and approaches (Figure 1).

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

Xenopus laevis provides an ideal model system for studying cell fate specification and physiological function of individual retinal cells in primary cell culture. Here we present a technique for dissecting retinal tissues and generating primary cell cultures that are imaged for calcium activity and analyzed by in situ hybridization.

The process by which  the anterior region of the neural plate gives rise to the vertebrate retina  continues to be a major focus of both clinical and ...

Whole Cell Patch Clamp for Investigating the Mechanisms of Infrared Neural Stimulation

It has been demonstrated in recent years that pulsed, infrared laser light can be used to elicit electrical responses in neural tissue, independent of any further modification of the target tissue. Infrared neural stimulation has been reported in a variety of peripheral and sensory neural tissue in vivo, with particular interest shown in stimulation of neurons in the auditory nerve. However, while INS has been shown to work in these settings, the mechanism (or mechanisms) by which infrared light causes neural excitation is currently not well understood. The protocol presented here describes a whole cell patch clamp method designed to facilitate the investigation of infrared neural stimulation in cultured primary auditory neurons. By thoroughly characterizing the response of these cells to infrared laser illumination in vitro under controlled conditions, it may be possible to gain an improved understanding of the fundamental physical and biochemical processes underlying infrared neural stimulation.

Infrared nerve stimulation has been proposed as an alternative to electrical stimulation in a range of nerve types, including those associated with the auditory system. This protocol describes a patch clamp method for studying the mechanism of infrared nerve stimulation in a culture of primary auditory neurons.

It has been  demonstrated in recent years that pulsed, infrared laser light can be used to  elicit electrical responses in neural tissue, independent of ...

Isolation of Sensory Neurons of Aplysia californica for Patch Clamp Recordings of Glutamatergic Currents

The marine gastropod mollusk Aplysia californica has a venerable history as a model of nervous system function, with particular significance in studies of learning and memory. The typical preparations for such studies are ones in which the sensory and motoneurons are left intact in a minimally dissected animal, or a technically elaborate neuronal co-culture of individual sensory and motoneurons. Less common is the isolated neuronal preparation in which small clusters of nominally homogeneous neurons are dissociated into single cells in short term culture. Such isolated cells are useful for the biophysical characterization of ion currents using patch clamp techniques, and targeted modulation of these conductances. A protocol for preparing such cultures is described. The protocol takes advantage of the easily identifiable glutamatergic sensory neurons of the pleural and buccal ganglia, and describes their dissociation and minimal maintenance in culture for several days without serum.

Isolation of Sensory Neurons of Aplysia californica for Patch Clamp Recordings of Glutamatergic Currents

We describe the dissection of the nervous system of the marine sea hare Aplysia after anesthesia, the isolation of neurons for short term-tissue culture, and recordings of single cell ion currents via the patch clamp technique.

The marine gastropod mollusk Aplysia californica has a venerable  history as a model of nervous system function, with particular significance in  studies ...

Whole-cell Patch-clamp Recordings from Morphologically- and Neurochemically-identified Hippocampal Interneurons

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal population (10-20%), but show a high level of physiological, morphological, and neurochemical heterogeneity, reflecting their diverse functions. Therefore, investigation of interneurons provides important insights into the organization principles and function of neuronal circuits. This, however, requires an integrated physiological and neuroanatomical approach for the selection and identification of individual interneuron types. Whole-cell patch-clamp recording from acute brain slices of transgenic animals, expressing fluorescent proteins under the promoters of interneuron-specific markers, provides an efficient method to target and electrophysiologically characterize intrinsic and synaptic properties of specific interneuron types. Combined with intracellular dye labeling, this approach can be extended with post-hoc morphological and immunocytochemical analysis, enabling systematic identification of recorded neurons. These methods can be tailored to suit a broad range of scientific questions regarding functional properties of diverse types of cortical neurons.

Cortical networks are controlled by a small, but diverse set of inhibitory interneurons. Functional investigation of interneurons therefore requires targeted recording and rigorous identification. Described here is a combined approach involving whole-cell recordings from single or synaptically-coupled pairs of neurons with intracellular labeling, post-hoc morphological and immunocytochemical analysis.

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal ...

Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. Retinal neurons in the network comprise of numerous subtypes. More than 10 subtypes of bipolar cells, ganglion cells, and amacrine cells have been identified by morphological studies. Multiple subtypes of retinal neurons are thought to encode distinct features of visual signaling, such as motion and color, and form multiple neural pathways. However, the functional roles of each neuron in visual signal processing are not fully understood. The patch clamp method is useful to address this fundamental question. Here, a protocol to record light-evoked synaptic responses in mouse retinal neurons using patch clamp recordings in dark-adapted conditions is provided. The mouse eyes are dark-adapted O/N, and retinal slice preparations are dissected in a dark room using infrared illumination and viewers. Infrared light does not activate mouse photoreceptors and thus preserves their light responsiveness. Patch clamp is used to record light-evoked responses in retinal neurons. A fluorescent dye is injected during recordings to characterize neuronal morphological subtypes. This procedure enables us to determine the physiological functions of each neuron in the mouse retina.

We will demonstrate how to prepare retinal slices from the mouse eye and record light responses in retinal neurons. The entire procedure is conducted in dark-adapted conditions.

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. ...

Perforated Patch-clamp Recording of Mouse Olfactory Sensory Neurons in Intact Neuroepithelium: Functional Analysis of Neurons Expressing an Identified Odorant Receptor

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of olfactory-driven behaviors and their modulation. These coding properties depend heavily on the initial interaction between odor molecules and the olfactory receptor (OR) expressed in the OSNs. The identity, specificity and ligand spectrum of the expressed OR are critical. The probability to find the ligand of the OR expressed in an OSN chosen randomly within the epithelium is very low. To address this challenge, this protocol uses genetically tagged mice expressing the fluorescent protein GFP under the control of the promoter of defined ORs. OSNs are located in a tight and organized epithelium lining the nasal cavity, with neighboring cells influencing their maturation and function. Here we describe a method to isolate an intact olfactory epithelium and record through patch-clamp recordings the properties of OSNs expressing defined odorant receptors. The protocol allows one to characterize OSN membrane properties while keeping the influence of the neighboring tissue. Analysis of patch-clamp results yields&#160;a precise quantification of ligand/OR interactions, transduction pathways and pharmacology, OSNs&#39; coding properties and their modulation at the membrane level.&#160;

Analyzing the physiological properties of olfactory sensory neurons still faces technical limitations. Here we record them through perforated patch-clamp in an intact preparation of the olfactory epithelium in gene-targeted mice. This technique allows the characterization of membrane properties and responses to specific ligands of neurons expressing defined olfactory receptors.

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. However, there is piling evidence that olfaction is not solely limited to chemical sensitivity, but also includes temperature sensitivity. Premetamorphic Xenopus laevis are translucent animals, with protruding nasal cavities deprived of the cribriform plate separating the nose and the olfactory bulb. These characteristics make them well suited for studying olfaction, and particularly thermosensitivity. The present article describes the complete procedure for measuring temperature responses in the olfactory bulb of X. laevis larvae. Firstly, the electroporation of olfactory receptor neurons (ORNs) is performed with spectrally distinct dyes loaded into the nasal cavities in order to stain their axon terminals in the bulbar neuropil. The differential staining between left and right receptor neurons serves to identify the &#947;-glomerulus as the only structure innervated by contralateral presynaptic afferents. Secondly, the electroporation is combined with focal bolus loading in the olfactory bulb in order to stain mitral cells and their dendrites. The 3D brain volume is then scanned under line-illumination microscopy for the acquisition of fast calcium imaging data while small temperature drops are induced at the olfactory epithelium. Lastly, the post-acquisition analysis allows the morphological reconstruction of the thermosensitive network comprising the &#947;-glomerulus and its innervating mitral cells, based on specific temperature-induced Ca2+ traces. Using chemical odorants as stimuli in addition to temperature jumps enables the comparison between thermosensitive and chemosensitive networks in the olfactory bulb.

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

Here we describe a protocol for measuring and analyzing temperature responses in the olfactory bulb of Xenopus laevis. Olfactory receptor neurons and mitral cells are differentially stained, after which calcium changes are recorded, reflecting a sensitivity of some neural networks in the bulb to temperature drops induced at the nose.

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. ...

Whole-cell Patch-clamp Recordings for Electrophysiological Determination of Ion Selectivity in Channelrhodopsins

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate electrical processes in target cells. In this context, ion selectivity of a channelrhodopsin is of particular importance. This article describes the investigation of chloride selectivity for a recently identified anion-conducting channelrhodopsin of Proteomonas sulcata via electrophysiological patch-clamp recordings on HEK293 cells. The experimental procedure for measuring light-gated photocurrents demands a fast switchable &#8211; ideally monochromatic &#8211; light source coupled into the microscope of an otherwise conventional patch-clamp setup. Preparative procedures prior to the experiment are outlined involving preparation of buffered solutions, considerations on liquid junction potentials, seeding and transfection of cells, and pulling of patch pipettes. The actual recording of current-voltage relations to determine the reversal potentials for different chloride concentrations takes place 24 h to 48 h after transfection. Finally, electrophysiological data are analyzed with respect to theoretical considerations of chloride conduction.

This article describes how the ion selectivity of channelrhodopsin is determined with electrophysiological whole-cell patch-clamp recordings using HEK293 cells. Here, the experimental procedure for investigating chloride selectivity of an anion-selective channelrhodopsin is demonstrated. However, the procedure is transferable to other channelrhodopsins of distinct selectivity.