This work was supported by the Australian Research Council under Linkage Project grant LP120100264.

1. Culture of Spiral Ganglion Neurons
<ol>
	<li>Sterilize small round (e.g. 10 mm diameter) glass coverslips and curved forceps in an autoclave. Transfer the sterilized coverslips into individual wells of a sterile 4-ring 35 mm petri dish or 4-well plate, using the sterilized forceps. Apply 150 &mu;l of poly-L-ornithine (500 &mu;g/ml) and mouse laminin (0.01 mg/ml) to the top surface of the coverslip and place in an incubator (37 &deg;C) for up to 48 hr. Ensure that the coverslips do not float away from the bottom of the well.</li>
	<li>Prepare 50 ml sterile Neurobasal media (NBM) for each neural culture: 47.5 ml neurobasal A, 0.5 ml N2 supplement, 1 ml B27 supplement, 0.5 ml L-glutamine, and 0.5 ml penicillin-streptomycin. Note: Supplements can be frozen, stored at -20 &deg;C and added to the media on the day required.</li>
	<li>Dissociate spiral ganglion neurons from post-natal day 4-7 rat pups as previously described 20,21, using both enzymatic (0.025% trypsin and 0.001% DNase I) and mechanical techniques. Refer to Whitlon et al. 22 and Vieira et al. 23 for detailed procedures of spiral ganglion neuron culture, or Parker et al. 24 for a demonstration of modiolus isolation.</li>
	<li>Aspirate any remaining poly-L-ornithine/laminin solution from the coverslips and wash briefly with NBM.</li>
	<li>Add 150-200 &mu;l of the dissociated spiral ganglion neuron suspension to the coverslips and place into an incubator (37 &deg;C, 10% CO2). Note: up to 20 coverslips can be prepared from an average litter of 8 rat pups.</li>
	<li>Four hours after plating neurons, aspirate the solution to remove cell debris and replace with 150-200 &mu;l warmed fresh NBM. Note: media may require daily replenishment to avoid dehydration.</li>
	<li>Return coverslips to the incubator until required for electrophysiological recordings. Note: dissociated spiral ganglion neuron cultures can be used for electrophysiological experiments four hours after dissociation and for up to two days thereafter. Time in vitro should be taken into account during analysis of results. Replenish NBM every 24-48 hr.</li>
</ol>
2. Preparation for Patch Clamp Recordings
<ol>
	<li>Prepare solutions
	<ol>
		<li>Intracellular (micropipette) solution: 115 mM K-gluconate, 7 mM KCl, 10 mM HEPES, 0.05 mM EGTA, 2 mM Na2ATP, 2 mM MgATP, 0.5 mM Na2GTP (adjust to pH 7.3 with KOH; adjust to 295 mOsmol/kg with sucrose). Pass the solution through a sterile filter (0.2 &mu;m) and divide into 200 &mu;l aliquots to be stored at -20 &deg;C until the day of recording.</li>
		<li>Extracellular (bath) solution: 137 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, 10 mM glucose (adjust to pH 7.4 with NaOH; adjust to 300-310 mOsmol/kg with sucrose). This solution is made on the day of recording.</li>
	</ol>
	</li>
	<li>Prepare recording micropipettes with a resistance of 2-6 M&Omega;. We use a CO2 laser puller (P-2000; Sutter Instruments) and borosilicate glass (1.0 mm outer diameter; 0.58 mm inner diameter; 75 mm length).</li>
	<li>Prepare the laser. This protocol is intended for use with a fiber-coupled laser, such as the 1,870 nm Infrared Nerve Stimulator from OptoTech P/L. The optical fiber used for light delivery in our experiments is a 200/220 &mu;m core/cladding diameter silica fiber with a numerical aperture of 0.22 and FC-PC connectors at both ends (AFW Technologies MM1-FC2-200/220-5-C-0.22). The patch cords were cut in half to produce two fiber pigtails (i.e. connectorized at one end and cleaved at the other). The effects of fiber core diameter and numerical aperture on laser induced temperature changes have been discussed in detail by Thompson et al. 25
	<ol>
		<li>Cleave the tip of the light delivery fiber using standard techniques and ensure that the resulting tip is of high quality by observation with an optical microscope (i.e. the tip should be perpendicular to the fiber axis and appear flat upon visual inspection). Connect the light delivery fiber to the fiber-coupled output of the stimulation laser using an appropriate through connector (e.g. Thorlabs ADAFC2).</li>
		<li>Measure the output laser power from the cleaved tip of the light delivery fiber using an appropriate instrument (e.g. Coherent FieldMate with LM-3 detector head). It is recommended to check the laser power every time the light delivery fiber is cleaved or a significant adjustment is made to the laser (e.g. transportation from one lab to another).</li>
		<li>Insert light delivery fiber into a fiber chuck or equivalent device and affix the chuck to the appropriate micropositioner. It is important to be able to accurately determine the angle &theta; that the optical fiber makes with the coverslip. This angle can be measured by taking a photograph of the experimental arrangement and using image processing software (e.g. ImageJ) to obtain the angle. The angle &theta; (or range of possible angles) is primarily constrained by spatial limitations of the experimental arrangement - in particular the microscope. Typical values of &theta; in our experiments (based on an upright microscope) are around 36&deg;, however the optimal angle may differ significantly for alternative setups (e.g. those using an inverted microscope).</li>
		<li>Make connections between the laser and the patch clamp data acquisition system (e.g. Digidata 1440A, Molecular Devices) as shown in Figure 2. The digital output from the patch clamp data acquisition system should be connected to the laser via an external function generator, making it possible to specify laser pulse parameters independent of the data acquisition system. Alternatively this output can be connected directly to the laser driver (requiring the pulse length and repetition rate to be set by the data acquisition software). In either case, the signal used to trigger the laser should be connected back to an input of the data acquisition system to ensure that the timing and length of the laser pulses can be recorded concurrently with the electrophysiological signal.</li>
	</ol>
	</li>
</ol>
3. Patch Clamp Recordings for Investigation of INS
<ol>
	<li>Fill the appropriate container of the perfusion system with extracellular solution and adjust the flow rate to provide perfusion of the bath at a rate of 1-2 ml/min. We use a gravity-fed system (aspirator bottle, pinch valve, and PE tubing) with an in-line heater to enable rapid heating of the solution, and a peristaltic pump to remove spent solution by suction.</li>
	<li>Place a coverslip with cultured cells into the recording chamber (bath) of an upright microscope. Using a high magnification water-immersion objective (e.g. 40X) and phase-contrast (e.g. differential interference contrast or Dodt gradient contrast), visually locate the spiral ganglion neurons within the culture. A typical spiral ganglion neuron is phase-bright, round and approximately 15 &mu;m in diameter with a prominent nucleus, as shown in Figure 1a.</li>
	<li>Once a neuron has been located, switch to a lower magnification objective (e.g. 10X) and locate the target neuron within the visual field.</li>
	<li>Move the light delivery optical fiber into position using the following procedure (or equivalent):
	<ol>
		<li>Use the micropositioner to move the output fiber until the tip is close to the target neuron in both the horizontal and vertical planes. The vertical position of the fiber tip can be verified by moving the objective up and down (scanning the focus).</li>
		<li>Switch back to the high magnification objective and position the tip of the optical fiber in its intended position next to the neuron (see Figure 2 inset). When adjusting the vertical position of the fiber, it is important that the bottom edge of the fiber is resting on (i.e. just touching) the coverslip, to minimize uncertainty in the location of the fiber. Optimal alignment in the horizontal plane will depend on the angle &theta; that the fiber makes with the coverslip and the radius of the fiber r. In order to position the fiber so that the center of the target neuron lies along the fiber axis, the horizontal distance between the upper edge of the fiber and the target cell should be 
		&Delta;target = r (cosec &theta; - 2 sin &theta;), 
		where negative values would indicate that the optical fiber overhangs the cell. When aligning the fiber, a distance of &Delta;target between the top edge of the fiber and the center of the neuron can be approximately achieved by visual inspection. However &Delta;target is designed to serve as a rough guideline only. Positioning the fiber such that the actual distance &Delta; between the upper edge of the fiber and the center of the neuron differs measurably from &Delta;target (as in Figure 1) can provide insight into the effect of both radial displacement (i.e. from the center of the beam) and axial displacement (i.e. along the fiber axis) of the beam relative to the target neuron. For example, setting &Delta; &asymp; 0 would be expected to increase the local temperature in the vicinity of the cell - i.e. since the beam profile for typical multimode fibers is well approximated by a top hat distribution 26, the decrease in locally absorbed energy (temperature) due to displacement from the center of the beam is minimal compared to the increase in absorbed energy from moving the fiber end face closer to the cell. 
		While care should be taken to position the fiber as accurately and repeatably as possible using visual cues, precise measurement of the fiber location relative to the target neuron (e.g. &Delta;) is of greater importance than positioning it a predetermined distance away. &Delta; can be determined (with an estimated maximum uncertainty of &plusmn;3 &mu;m) by image analysis as described in step 3.8.2 of the protocol. In some experiments 5,8, white light sources have been coupled through the fiber in order to target the desired cell. This approach was found to be ineffective in the upright microscope setup, as the intensity of light scattered from the target region was relatively low at these shallow angles (&theta;).</li>
		<li>Once the fiber is in position, move it out by a known amount to enable straightforward positioning of the micropipette. The fiber can then be returned to the original position when a neural recording is achieved.</li>
	</ol>
	</li>
	<li>Fill the micropipette with intracellular solution and securely fit into place on the headstage of the amplifier (e.g. Multiclamp 700B, Molecular Devices). Using tubing attached to the side of the microelectrode holder, apply a small amount of positive pressure to prevent clogging of the micropipette.</li>
	<li>Using a micromanipulator, move the micropipette into position just above the target neuron.</li>
	<li>Protocol for whole cell recordings:
	<ol>
		<li>Apply a 10 msec, +10 mV square wave pulse (e.g. Seal Test) in voltage-clamp mode of the amplifier and adjust the micropipette potential (pipette offset) of the baseline signal to 0 nA. The seal test should also be used to determine whether the resistance of the microelectrode is within the desired range (e.g. 2 - 6 M&Omega;).</li>
		<li>While monitoring the resistance, use fine adjustments of the micromanipulator to gently position the micropipette onto the surface of the target neuron. When the micropipette is in contact with the neuron, the resistance will increase (by ~ 0.5 to 1 M&Omega;). Immediately following the resistance increase, remove the positive fluid pressure and apply a small negative pressure. Once resistance has passed 10 - 20 M&Omega;, fix the membrane potential to a holding level (e.g. around -60 mV).</li>
		<li>The electrode resistance should continue to increase until it exceeds 1 G&Omega;, signifying that an effective seal (termed 'gigaseal') has been formed between the cell membrane and micropipette. At this point, remove all fluid pressure from the micropipette.</li>
		<li>Once the gigaseal has been formed, use short current pulses (25 to 100 &mu;sec; +1 V) or brief pulses of negative fluid pressure to rupture the cell membrane and achieve whole cell configuration.</li>
		<li>Record the membrane capacitance, series resistance and input resistance as determined from the exponential curve fitted to the current during the seal test pulse. Minimize the capacitance transients by adjusting the CpFast and CpSlow controls on the amplifier, then switch the amplifier into whole cell mode, and compensate capacitance and resistance until a flat current is observed during the seal test. Apply series resistance compensation (~70% correction; 70% prediction), adjusting capacitance and resistance controls to maintain a flat test seal response.</li>
		<li>Switch to current-clamp mode in the amplifier. Take note of the resting membrane potential (in the absence of current injection). Set a holding current to stabilize the membrane potential at the desired level (e.g. -60 mV). Neutralize the pipette capacitance and adjust the bridge balance to balance the voltage drop.</li>
		<li>Check the firing properties of the neuron by stimulating with depolarizing current (+10 to +200 pA in +10 pA steps; 300 msec duration).</li>
	</ol>
	</li>
	<li>
	<ol>
		<li>Move the optical fiber back into position next to the neuron. Using imaging software coupled to a CCD camera, capture images of the position of the fiber with respect to the target neuron, focusing on the plane of the neuron initially, and then on the top edge of the optical fiber (see Figure 1).</li>
		<li>By subsequent analysis of the resulting images (e.g. Figures 1b and 1c) it is possible to precisely determine &Delta; (the position of the upper edge of the optical fiber relative to the center of the target neuron). Once &Delta; is known, parameters such as the distance from the fiber end face to the target neuron (along the fiber axis) z and the radial displacement of the neuron from the center of the beam &delta;r can be calculated using simple trigonometric relations: 
		z = r sin 2&theta; + &Delta; cos &theta; 
		&delta;r = ((r cos 2&theta; - &Delta; sin &theta;)2 + &delta;y2)1/2, 
		where &delta;y is the distance between the fiber axis and the center of the neuron as seen from above (e.g. see Figure 1). 
		Analysis should take into account positional variations from cell to cell, as accurate knowledge of these location parameters may be required to resolve possible differences between stimulation processes 25.</li>
	</ol>
	</li>
</ol>
4. INS Experiments
<ol>
	<li>While recording electrophysiological data in either current clamp or voltage clamp configurations, run the stimulation laser at the desired parameters (e.g. power, pulse length, repetition rate etc.). With our laser, the optical power is controlled via a direct input to the laser driver and is specified manually before each recording. Pulse length and repetition rate can be controlled by either an external signal generator or the data acquisition software (as described in step 2.3.4). Ensure that data are recorded from both the patch clamp channel and the laser trigger channel.</li>
</ol>
Laser pulses with lengths ranging from around 500 &mu;sec to 15 msec and energies of ~0.25-5 mJ per pulse typically yield measurable electrical responses. Setting the repetition rate of laser pulses to be 1 Hz or less may be useful for initial experiments, since it will minimize the effects of this parameter. Typical results showing the change in the recorded signal are presented in the following Section.

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L., Trauner, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+photochemical+tools+for+controlling+neuronal+activity.">New photochemical tools for controlling neuronal activity.</a> Current Opinion in Neurobiology. 19, 544-552 (2009).</li><li>Boyden, E. S., Zhang, F., Bamberg, E., Nagel, G., Deisseroth, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Millisecond-timescale+genetically+targeted+optical+control+of+neural+activity.">Millisecond-timescale genetically targeted optical control of neural activity.</a> Nature Neuroscience. 8, 1263-1268 (2005).</li><li>Albert, E. 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The authors have nothing to disclose.

Using the protocols outlined in this paper it is possible to extract and culture spiral ganglion neurons and to investigate laser-evoked electrical activity by performing whole cell patch clamp experiments. When used in vitro, the patch clamp technique provides a level of control over experimental parameters that is not achievable in vivo. Laser stimulation parameters such as wavelength, pulse energy, pulse length, pulse shape, and pulse repetition sequences can be studied in a reproducible setting. In ...

The fields of neurophysiology and medical bionics rely heavily on techniques that allow controllable stimulation of electrical responses in neural tissue. While electrical stimulation remains the gold standard in neural excitation, it suffers from a number of drawbacks such as the presence of stimulation artifacts when recording neural responses, and a lack of stimulation specificity due to the spread of current into surrounding tissue 1.
The last two decades have seen the development of optically mediated stimulation techniques 2. Several of these techniques require modification of the target tissue, either via the addition of a particular molecule (e.g. caged molecules) 3 or some form of genetic manipulation (e.g. optogenetics) 4, neither of which are easy to apply outside of a research setting. Of particular interest therefore is infrared neural stimulation (INS), whereby neural tissue is excited by pulsed infrared laser light. INS has the potential to overcome many of the shortcomings of electrical stimulation by enabling highly specific, non-contact stimulation of neural tissue 2. However, while INS has been successfully demonstrated in a variety of settings in vivo, the precise mechanism of excitation remains uncertain.
Some recent publications have shown progress towards uncovering the mechanism behind INS 5-7. Rapid heating due to absorption of the laser light by water appears to play a key role. However, beyond this a consensus is yet to be reached. Shapiro et al. 7 propose a highly general mechanism whereby rapid heating causes a perturbation in the distribution of charged particles adjacent to the cell membrane, leading to a change in the capacitance of the cell membrane and subsequent depolarization. In addition, Albert et al. 5 assert that laser induced heating activates a specific class of temperature sensitive ion channels (transient receptor potential vanilloid channels), allowing ions to pass through the cell membrane. At this stage it is unclear how these mechanisms combine, or indeed whether there are further factors that are yet to be identified.
Although a small number of publications (references 5,7-9) have investigated INS in vitro, the vast majority of work published in this field has been carried out in vivo (e.g. references 1,6,10-18). Infrared stimulation of auditory neurons has been an area of particular interest, owing to the potential applications in cochlear implants 10,14-18. While in vivo experiments are important to verify the effectiveness of the technique in various settings, the increased level of control afforded by in vitro studies is expected to lead to a more detailed understanding of the mechanism responsible for INS. This report describes the preparation of cultured spiral ganglion neurons for patch clamp investigations, as these can be used to study fundamental mechanisms while also linking to the large body of existing data from the auditory system.
The patch clamp technique is an excellent tool for investigations of electrophysiological phenomena, providing a means of recording electrical activity in single cells and studying the contribution of the individual underlying currents19. When this technique is applied to a stable in vitro preparation of primary neurons, such as cultured spiral ganglion neurons, it offers the opportunity to study in depth the mechanisms by which neural activity is controlled and manipulated.
The protocols specified in this work outline methods for investigating the effect of laser stimulation on the electrical properties of spiral ganglion neurons through patch clamp recordings. The approach is based on a fiber-coupled laser rather than a free-space laser, allowing safer operation as well as easier and more repeatable alignment without the need to modify the standard microscope configuration. On the basis of these protocols, it should be possible to conduct a wide range of experiments in order to more clearly determine the mechanism or mechanisms behind INS.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Cell culture materials and equipment </td>
 </tr>
 <tr>
 <td>Glass coverslips</td>
 <td>Lomb Scientific</td>
 <td>CSC 10 1 GP</td>
 <td></td>
 </tr>
 <tr>
 <td>4-ring cell culture dish </td>
 <td>VWR International</td>
 <td>82050-542</td>
 <td></td>
 </tr>
 <tr>
 <td>Poly-L-ornithine solution</td>
 <td>Sigma-Aldrich</td>
 <td>P4957</td>
 <td></td>
 </tr>
 <tr>
 <td>Laminin</td>
 <td>Invitrogen</td>
 <td>23017-015</td>
 <td></td>
 </tr>
 <tr>
 <td>Curved forceps</td>
 <td>WPI</td>
 <td>14101</td>
 <td>Dumont #5 tweezers (45&deg; angle tip)</td>
 </tr>
 <tr>
 <td>CO2 Incubator</td>
 <td>ThermoScientific</td>
 <td>Heracell 150i</td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Table 1. Cell culture materials and equipment.</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Neurobasal media</td>
 </tr>
 <tr>
 <td>Neurobasal A</td>
 <td>Gibco</td>
 <td>10888-022</td>
 <td></td>
 </tr>
 <tr>
 <td>N-2 supplement</td>
 <td>Invitrogen</td>
 <td>17502-048</td>
 <td></td>
 </tr>
 <tr>
 <td>B27 serum-free supplement</td>
 <td>Invitrogen</td>
 <td>17504-044</td>
 <td></td>
 </tr>
 <tr>
 <td>Penicillin-Streptomycin</td>
 <td>Invitrogen</td>
 <td>15140-148</td>
 <td></td>
 </tr>
 <tr>
 <td>L-Glutamine</td>
 <td>Invitrogen</td>
 <td>25030-149</td>
 <td></td>
 </tr>
 <tr>
 <td>Intracellular solution</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Potassium chloride</td>
 <td>Sigma-Aldrich</td>
 <td>P4504</td>
 <td></td>
 </tr>
 <tr>
 <td>HEPES</td>
 <td>Sigma-Aldrich</td>
 <td>H4034</td>
 <td></td>
 </tr>
 <tr>
 <td>Potassium D-gluconate</td>
 <td>Sigma-Aldrich</td>
 <td>G4500</td>
 <td></td>
 </tr>
 <tr>
 <td>EGTA</td>
 <td>Sigma-Aldrich</td>
 <td>E3889</td>
 <td></td>
 </tr>
 <tr>
 <td>Na2ATP</td>
 <td>Sigma-Aldrich</td>
 <td>A2383</td>
 <td></td>
 </tr>
 <tr>
 <td>MgATP</td>
 <td>Sigma-Aldrich</td>
 <td>A9187</td>
 <td></td>
 </tr>
 <tr>
 <td>NaGTP</td>
 <td>Sigma-Aldrich</td>
 <td>G8877</td>
 <td></td>
 </tr>
 <tr>
 <td>Potassium hydroxide</td>
 <td>LabServ</td>
 <td>BSPPL738.500</td>
 <td></td>
 </tr>
 <tr>
 <td>Sucrose</td>
 <td>Sigma-Aldrich</td>
 <td>S8501</td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Extracellular solution </td>
 </tr>
 <tr>
 <td>Sodium chloride</td>
 <td>Sigma-Aldrich</td>
 <td>310166</td>
 <td></td>
 </tr>
 <tr>
 <td>Potassium chloride</td>
 <td>Sigma-Aldrich</td>
 <td>P4504</td>
 <td></td>
 </tr>
 <tr>
 <td>HEPES</td>
 <td>Sigma-Aldrich</td>
 <td>H4034</td>
 <td></td>
 </tr>
 <tr>
 <td>Calcium chloride</td>
 <td>Sigma-Aldrich</td>
 <td>383147</td>
 <td></td>
 </tr>
 <tr>
 <td>Magnesium chloride</td>
 <td>Sigma-Aldrich</td>
 <td>M8266</td>
 <td></td>
 </tr>
 <tr>
 <td>D-Glucose</td>
 <td>Sigma-Aldrich</td>
 <td>G8270</td>
 <td></td>
 </tr>
 <tr>
 <td>Sodium hydroxide</td>
 <td>LabServ</td>
 <td>BSPSL740.500</td>
 <td></td>
 </tr>
 <tr>
 <td>Sucrose</td>
 <td>Sigma-Aldrich</td>
 <td>S8501</td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Table 2. Solutions for cell culture and patch clamp. a) Neurobasal media. b) Intracellular solution. c) Extracellular solution.</td>
 </tr>
 <tr>
 <td>Upright microscope</td>
 <td>Zeiss </td>
 <td>AxioExaminerD1</td>
 <td>Equipped with Dodt contrast </td>
 </tr>
 <tr>
 <td>Water-immersion objective</td>
 <td>Zeiss</td>
 <td>W Plan-APOCHROMAT 40x/0.75</td>
 <td></td>
 </tr>
 <tr>
 <td>Platform and X-Y stage </td>
 <td>ThorLabs</td>
 <td>Burleigh Gibraltar</td>
 <td></td>
 </tr>
 <tr>
 <td>Recording chamber</td>
 <td>Warner Instruments</td>
 <td>RC-26G</td>
 <td></td>
 </tr>
 <tr>
 <td>Vibration isolation table</td>
 <td>TMC</td>
 <td>Micro-g 63-532</td>
 <td></td>
 </tr>
 <tr>
 <td>CCD Camera</td>
 <td>Diagnostic Instruments</td>
 <td>RT1200</td>
 <td></td>
 </tr>
 <tr>
 <td>Camera software</td>
 <td>Diagnostic Instruments</td>
 <td>SPOT Basic</td>
 <td></td>
 </tr>
 <tr>
 <td>In-line solution heater</td>
 <td>Warner</td>
 <td>SH-27B</td>
 <td></td>
 </tr>
 <tr>
 <td>Temperature controller</td>
 <td>Warner</td>
 <td>TC-324B</td>
 <td></td>
 </tr>
 <tr>
 <td>Patch clamp amplifier</td>
 <td>Molecular Devices</td>
 <td>Multiclamp 700B</td>
 <td></td>
 </tr>
 <tr>
 <td>Patch clamp data acquisition system</td>
 <td>Molecular Devices</td>
 <td>Digidata 1440A</td>
 <td></td>
 </tr>
 <tr>
 <td>Micromanipulator</td>
 <td>Sutter Instruments</td>
 <td>MPC-325</td>
 <td></td>
 </tr>
 <tr>
 <td>Micropipette glass</td>
 <td>Sutter Instruments</td>
 <td>GBF100-58-15</td>
 <td>Borosilicate glass with filament</td>
 </tr>
 <tr>
 <td>Micropipette Puller</td>
 <td>Sutter Instruments</td>
 <td>P2000</td>
 <td></td>
 </tr>
 <tr>
 <td>Recording Software</td>
 <td>AxoGraph</td>
 <td></td>
 <td>Lab pack and electrophysiology tools</td>
 </tr>
 <tr>
 <td>Aspirator bottle</td>
 <td>Sigma-Aldrich</td>
 <td>CLS12201L</td>
 <td>1 L Pyrex aspirator bottle, with outlet for tubing</td>
 </tr>
 <tr>
 <td>PE Tubing</td>
 <td>Harvard</td>
 <td>PolyE #340</td>
 <td></td>
 </tr>
 <tr>
 <td>Masterflex peristaltic pump</td>
 <td>Cole-Parmer</td>
 <td>HV-07554-85</td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Table 3.Patch clamp equipment.</td>
 </tr>
 <tr>
 <td>1,870 nm laser diode</td>
 <td>Optotech</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>200/220 &mu;m diameter multimode optical fiber patch cord (FC/PC)</td>
 <td>AFW Technologies</td>
 <td>MM1-FC2-200/220-5-C-0.22</td>
 <td>Light delivery optical fiber, silica core and cladding, 0.22 NA</td>
 </tr>
 <tr>
 <td>Optical fiber through connector (FC/PC)</td>
 <td>Thorlabs</td>
 <td>ADAFC2</td>
 <td></td>
 </tr>
 <tr>
 <td>Optical fiber cleaver</td>
 <td>EREM</td>
 <td>FO1 </td>
 <td></td>
 </tr>
 <tr>
 <td>Optical fiber stripping tool (0.25 - 0.6 mm) </td>
 <td>Siemens</td>
 <td></td>
 <td>For removing optical fiber jacket</td>
 </tr>
 <tr>
 <td>Optical fiber stripping tool (0.6 - 1.0 mm)</td>
 <td>Siemens</td>
 <td></td>
 <td>For removing outer coating of patch cord</td>
 </tr>
 <tr>
 <td>Signal generator</td>
 <td></td>
 <td></td>
 <td>Any signal generator that can output the necessary pulse shapes and is capable of being externally triggered</td>
 </tr>
 <tr>
 <td>Optical fiber positioner</td>
 <td></td>
 <td></td>
 <td>Custom made positioner. Could substitute with standard micropositioner used for patch clamp experiments</td>
 </tr>
 <tr>
 <td>Optical fiber chuck</td>
 <td>Newport</td>
 <td>FPH-DJ</td>
 <td></td>
 </tr>
 <tr>
 <td>Laser power meter and detector head</td>
 <td>Coherent</td>
 <td>FieldMate (power meter) with LM-3 (detector head)</td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Table 4. Laser equipment.</td>
 </tr>
 </tbody>
</table>

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.

Spiral ganglion neurons respond to laser illumination with repeatable waveforms in both voltage-clamp and current-clamp recording configurations. Figure 3a shows typical changes in current flow across a cell membrane in response to a 2.5 msec, 0.8 mJ laser pulse (average response from 6 laser pulses, repeated at 1 sec intervals) with the membrane potential held at -70 mV, -60 mV and -50 mV. Net inward currents are consistently evoked in response to laser pulses, returning to initial values after illumina...

Watch this Scientific Journal Video about Whole Cell Patch Clamp for Investigating the Mechanisms of Infrared Neural Stimulation at JoVE.com

Whole Cell Patch Clamp for Investigating the Mechanisms of 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 ...

whole-cell-patch-clamp-for-investigating-mechanisms-infrared-neural

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Neuroscience

20.8K Views.  Swinburne University of Technology. 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.

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

Investigating the Neural Mechanisms of Aware and Unaware Fear Memory with fMRI

Pavlovian fear conditioning is often used in combination with functional magnetic resonance imaging (fMRI) in humans to investigate the neural substrates of associative learning 1-5. In these studies, it is important to provide behavioral evidence of conditioning to verify that differences in brain activity are learning-related and correlated with human behavior. 
Fear conditioning studies often monitor autonomic responses (e.g. skin conductance response; SCR) as an index of learning and memory 6-8. In addition, other behavioral measures can provide valuable information about the learning process and/or other cognitive functions that influence conditioning. For example, the impact unconditioned stimulus (UCS) expectancies have on the expression of the conditioned response (CR) and unconditioned response (UCR) has been a topic of interest in several recent studies 9-14. SCR and UCS expectancy measures have recently been used in conjunction with fMRI to investigate the neural substrates of aware and unaware fear learning and memory processes 15. Although these cognitive processes can be evaluated to some degree following the conditioning session, post-conditioning assessments cannot measure expectations on a trial-to-trial basis and are susceptible to interference and forgetting, as well as other factors that may distort results 16,17 .
Monitoring autonomic and behavioral responses simultaneously with fMRI provides a mechanism by which the neural substrates that mediate complex relationships between cognitive processes and behavioral/autonomic responses can be assessed. However, monitoring autonomic and behavioral responses in the MRI environment poses a number of practical problems. Specifically, 1) standard behavioral and physiological monitoring equipment is constructed of ferrous material that cannot be safely used near the MRI scanner, 2) when this equipment is placed outside of the MRI scanning chamber, the cables projecting to the subject can carry RF noise that produces artifacts in brain images, 3) artifacts can be produced within the skin conductance signal by switching gradients during scanning, 4) the fMRI signal produced by the motor demands of behavioral responses may need to be distinguished from activity related to the cognitive processes of interest. Each of these issues can be resolved with modifications to the setup of physiological monitoring equipment and additional data analysis procedures. Here we present a methodology to simultaneously monitor autonomic and behavioral responses during fMRI, and demonstrate the use of these methods to investigate aware and unaware memory processes during fear conditioning.

A methodology to investigate the neural mechanisms that support aware and unaware memory processes during fear conditioning is described. This method monitors blood oxygen level dependent (BOLD) functional magnetic resonance imaging, skin conductance response, and unconditioned stimulus expectancy during Pavlovian fear conditioning to assess the neural correlates of distinct memory processes.

Pavlovian fear conditioning is often used in combination with functional magnetic resonance imaging (fMRI) in humans to investigate the neural substrates ...

A Computer-assisted Multi-electrode Patch-clamp System

The patch-clamp technique is today the most well-established method for recording electrical activity from individual neurons or their subcellular compartments. Nevertheless, achieving stable recordings, even from individual cells, remains a time-consuming procedure of considerable complexity. Automation of many steps in conjunction with efficient information display can greatly assist experimentalists in performing a larger number of recordings with greater reliability and in less time. In order to achieve large-scale recordings we concluded the most efficient approach is not to fully automatize the process but to simplify the experimental steps and reduce the chances of human error while efficiently incorporating the experimenter's experience and visual feedback. With these goals in mind we developed a computer-assisted system which centralizes all the controls necessary for a multi-electrode patch-clamp experiment in a single interface, a commercially available wireless gamepad, while displaying experiment related information and guidance cues on the computer screen. Here we describe the different components of the system which allowed us to reduce the time required for achieving the recording configuration and substantially increase the chances of successfully recording large numbers of neurons simultaneously.

Multi-electrode patch-clamp recordings constitute a complex task. Here we show how, by automating of many of the experimental steps, it is possible to accelerate the process leading to qualitative improvement in performance and number of recordings.

The patch-clamp technique is today the most  well-established method for recording electrical activity from individual  neurons or their subcellular ...

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 ...

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 ...

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.

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate ...

Time-dependent Increase in the Network Response to the Stimulation of Neuronal Cell Cultures on Micro-electrode Arrays

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network dynamics in neuronal cultures. In contrast with more traditional methods, such as patch-clamping, which can only record activity from a single cell, MEAs can record simultaneously from multiple sites in a network, without requiring the arduous task of placing each electrode individually. Moreover, numerous control and stimulation configurations can be easily applied within the same experimental setup, allowing for a broad range of dynamics to be explored. One of the key dynamics of interest in these in vitro studies has been the extent to which cultured networks display properties indicative of learning. Mouse neuronal cells cultured on MEAs display an increase in response following training induced by electrical stimulation. This protocol demonstrates how to culture neuronal cells on MEAs; successfully record from over 95% of the plated dishes; establish a protocol to train the networks to respond to patterns of stimulation; and sort, plot, and interpret the results from such experiments. The use of a proprietary system for stimulating and recording neuronal cultures is demonstrated. Software packages are also used to sort neuronal units. A custom-designed graphical user interface is used to visualize post-stimulus time histograms, inter-burst intervals, and burst duration, as well as to compare the cellular response to stimulation before and after a training protocol. Finally, representative results and future directions of this research effort are discussed.

Mouse neuronal cells cultured on multi-electrode arrays display an increase in response following electrical stimulation. This protocol demonstrates how to culture neurons, how to record activity, and how to establish a protocol to train the networks to respond to patterns of stimulation.

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network ...

Auditory Brainstem Response and Outer Hair Cell Whole-cell Patch Clamp Recording in Postnatal Rats

The outer hair cell is one of the two types of sensory hair cells in the mammalian cochlea. They alter their cell length with the receptor potential to amplify the weak vibration of low-level sound signal. The morphology and electrophysiological property of outer hair cells (OHCs) develop in early postnatal ages. The maturation of outer hair cell may contribute to the development of the auditory system. However, the process of OHCs development is not well studied. This is partly because of the difficulty to measure their function by an electrophysiological approach. With the purpose of developing a simple method to address the above issue, here we describe a step-by-step protocol to study the function of OHCs in acutely dissociated cochlea from postnatal rats. With this method, we can evaluate the cochlear response to pure tone stimuli and examine the expression level and function of the motor protein prestin in OHCs. This method can also be used to investigate the inner hair cells (IHCs).

This protocol describes methods to record the auditory brainstem response from postnatal rat pups. To examine the functional development of outer hair cells, the experimental procedure of whole-cell patch clamp recording in isolated outer hair cells is described step-by-step.

The outer hair cell is one of the two types of sensory hair cells in the mammalian cochlea. They alter their cell length with the receptor potential to ...

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.

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.

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

Preparation of Acute Spinal Cord Slices for Whole-cell Patch-clamp Recording in Substantia Gelatinosa Neurons

Recent whole-cell patch-clamp studies from substantia gelatinosa (SG) neurons have provided a large body of information about the spinal mechanisms underlying sensory transmission, nociceptive regulation, and chronic pain or itch development. Implementations of electrophysiological recordings together with morphological studies based on the utility of acute spinal cord slices have further improved our understanding of neuronal properties and the composition of local circuitry in SG. Here, we present a detailed and practical guide for the preparation of spinal cord slices and show representative whole-cell recording and morphological results. This protocol permits ideal neuronal preservation and can mimic in vivo conditions to a certain extent. In summary, the ability to obtain an in vitro preparation of spinal cord slices enables stable current- and voltage-clamp recordings and could thus facilitate detailed investigations into the intrinsic membrane properties, local circuitry and neuronal structure using diverse experimental approaches.

Here, we describe the essential steps for whole-cell patch-clamp recordings made from substantia gelatinosa (SG) neurons in the in vitro spinal cord slice. This method allows the intrinsic membrane properties, synaptic transmission and morphological characterization of SG neurons to be studied.

Recent whole-cell patch-clamp studies from substantia gelatinosa (SG) neurons have provided a large body of information about the spinal mechanisms ...

Evaluation of Synaptic Multiplicity Using Whole-cell Patch-clamp Electrophysiology

In the central nervous system, a pair of neurons often form&#160;multiple synaptic contacts and/or functional neurotransmitter release sites (synaptic multiplicity). Synaptic multiplicity is plastic and changes throughout development and in different physiological conditions, being an important determinant for the efficacy of synaptic transmission. Here, we outline experiments for estimating the degree of multiplicity of synapses terminating onto a given postsynaptic neuron using whole-cell patch clamp electrophysiology in acute brain slices. Specifically, voltage-clamp recording is used to compare the difference between the amplitude of spontaneous excitatory postsynaptic currents (sEPSCs) and miniature excitatory postsynaptic currents (mEPSCs). The theory behind this method is that afferent inputs that exhibit multiplicity will show large, action potential-dependent sEPSCs due to the synchronous release that occurs at each synaptic contact. In contrast, action potential-independent release (which is asynchronous) will generate smaller amplitude mEPSCs. This article outlines a set of experiments and analyses to characterize the existence of synaptic multiplicity and discusses the requirements and limitations of the technique. This technique can be applied to investigate how different behavioral, pharmacological or environmental interventions in vivo affect the organization of synaptic contacts in different brain areas.

Here, we present a protocol for evaluating the functional synaptic multiplicity using whole-cell patch clamp electrophysiology in acute brain slices.

In the central nervous system, a pair of neurons often form&#160;multiple synaptic contacts and/or functional neurotransmitter release sites (synaptic ...