1. Instrumentation 
<ol>
<li>To measure the response of olfactory sensory neurons to photolysis of caged compounds we use a flash lamp in combination with a typical patch-clamp recording system including: a patch-clamp amplifier, a recording electrode and a reference electrode connected to the head-stage of a patch-clamp amplifier, a digitizer, a computer, software for data acquisition, micro-manipulators, an epifluorescence microscope, a perfusion system, an anti-vibration table and a Faraday cage (Figure 2).</li>
<li>To generate a high-intensity flash of ultraviolet (UV) light we use a Xenon flash lamp (Rapp OptoElectronic JML-C2 Flash Unit, Figure 3A) that functions by discharging a high voltage capacitor across a short-arc flash tube that is filled with Xenon gas6-7. The light flash can be triggered manually or by the computer, using the same software that controls electrophysiological acquisition.</li>
<li>A UV short pass filter optimized for uncaging applications (270-390 nm) is inserted into an output slot of the flash lamp and a quartz fiber optic light guide is connected to the microscope (see 1.5).</li>
<li>The intensity of the light flash and therefore the extent of photolysis can be modified by changing specific parameters: voltage or capacitance6. Indeed, the light intensity is related to the energy stored in the capacitor and depends on the capacitance value and on the voltage across it, according to the standard relation 0.5 CV2. One of three capacitance values can be selected on the front panel of our flash lamp (C1=1000 &mu;F, C2=2000 &mu;F, or C3=3000 &mu;F) and the voltage can be varied between 100 to 385 Volts (Figure 3A), producing a stored energy ranging between 5 and 220 J. The flash duration varies from 0.5 to 1 ms at full half maximal width and depends on the selected capacitance value. Before selecting a different capacitance or setting the voltage to a lower value, it is necessary to discharge the capacitor, by selecting the discharge mode on the front panel of the flash lamp (Figure 3A). 
However the light energy is not simply proportional to the stored energy due to manyfold reasons related to the physics of gas discharges and to non-linear losses in the circuit as the electrical energy is increased6. To measure the energy associated to a light flash, a joulemeter, as the JM10 provided by RappOptoElectronics, can be used. Note that the energy at the exit of the quartz light guide with UV filter in place is in the range 1-15 mJ per flash, as reported in the user guide of the Rapp OptoElectronic Flash Unit. 
A photodiode assembly (Figure 3B) is provided with the flash lamp unit and can be used to estimate variations of light intensities as a function of C and V. Indeed, the output of the photodiode is proportional to the light intensity. Following the user guide instructions, the light guide is connected to the input of the photodiode assembly, while the output is connected to an oscilloscope. Figure 3C shows the peak output voltage of the photodiode in response to light flashes for the three capacitance values C1, C2, or C3, available in our flash lamp unit, versus the applied voltage. These values will change as the flash bulb becomes old. It is a good practice to regularly use the photodiode to check the deterioration of the flash bulb. 
Instead of modifying the capacitance and/or voltage on the front panel, a more precise and faster control of light intensity is obtained by placing neutral density filters into a second filter slot of the flash lamp (Figure 3A). This approach is quicker and additionally the flash duration, that depends on the selected capacitance value, is not modified. With neutral density filters ranging from 100% to 0.1 % transmission the flash intensity can be quickly modified over a large range.</li>
<li> The light flash is directed to the neuron, plated on glass-bottom dishes, through a quartz fiber optic light guide (Figure 2). The light guide is connected to the epifluorescence attachment of the inverted microscope using an adapter. A modified filter cube without excitation filter directs the UV beam to the neuron by a dichroic mirror. </li>
<li>The light is focused through a 100x oil immersion objective. The area covered by the flash is visualized by inserting the light guide into an illuminator that provides a continuous source of visible light. The image (represented in yellow in the video) is viewed on a monitor through a CCD camera connected to the microscope. The contour of the illuminated area is drawn on the monitor to localize the UV flash area for the experiments. Before delivering the UV flash, the ciliary region of the neuron is brought into the area drawn on the monitor.</li>
<li>The dimension of the spot of UV light is controlled by the objective magnification, the diameter of the light guide, and the position of the light guide at the back of the microscope. We use a 100x magnification oil immersion objective. The diameter of the circular spot is measured by directing the light from an illuminator on a micrometer slide placed on the stage of the microscope. The spot diameter in our set-up is approximately 16, 33, 50 or 105 &mu;m for a light guide with a diameter respectively of 200, 400, 600 or 1250 &mu;m.</li>
</ol>
2. Preparing solutions
Dissection
<ol>
<li>Ringer&#x2019;s solution: 140 mM NaCl, 5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, 10 mM glucose, 1 mM sodium pyruvate (pH 7.4 adjusted with 1M NaOH).</li>
<li>Stop mix solution: Ringer&#x2019;s solution with 0.1 mg/ml BSA, 0.3 mg/ml leupeptin, and 0.02 mg/ml DNaseI.</li> 
<li>Zero-divalent Ringer&#x2019;s solution: 140 mM NaCl, 5 mM KCl, 10 mM HEPES, 1 mM EDTA, 10 mM glucose, 1 mM sodium pyruvate (pH 7.4 with 1M NaOH). Add 200 &mu;M cysteine and 2 U/ml papain for enzymatic dissociation. 
Note: Sterile filter the solutions for dissection.</li>
</ol>
Patch-clamp recording solutions
Extracellular solutions
<ol>
 <li>Extracellular Ringer&#x2019;s solution: 140 mM NaCl, 5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, 10 mM glucose, 1 mM sodium pyruvate (pH 7.4 adjusted with 1M NaOH).</li>
 <li>Low-Ca Ringer&#x2019;s solution: 140 mM NaCl, 5 mM KCl, 10 mM EGTA, 1 mM MgCl2, 10 mM HEPES, 10 mM glucose, 1 mM sodium pyruvate (pH 7.4 adjusted with 1M NaOH).</li>
</ol>
Intracellular solutions
Always prepare and use caged compound solutions in dim light to avoid degradation of caged compounds from ambient light. Protect containers from light using aluminum foil.
Caged cAMP:
<ol>
<li>Make an intracellular solution containing 145 mM KCl, 4 mM MgCl2, 0.5 mM EGTA, 10 mM HEPES, 1 mM MgATP and 0.1 mM NaGTP (pH 7.4 adjusted with 1M KOH).</li>
<li>Make a stock solution dissolving caged cAMP (BCMCM-cAMP4) in DMSO at 50 mM. Make aliquots of 5 &mu;l. Store up to one year at -20&deg;C.</li>
<li>Dissolve an aliquot of the caged cAMP stock solution in the intracellular solution to obtain a final concentration of 50 &mu;M caged cAMP. Mix using a vortex.</li>
<li>Make aliquots of 1 ml of the caged cAMP intracellular solution. Store up to two months at -20&deg;C until use.</li>
</ol>
Caged Ca:
We prepare intracellular solutions containing 3 mM DMNP-EDTA5 50% loaded with 1.5 mM Ca.
<ol>
<li>Make an intracellular solution containing 140 mM KCl (or CsCl), 10 mM HEPES (pH 7.4 adjusted with 1M KOH or CsOH).</li>
<li>Open a vial of 5 mg DMNP-EDTA. Add 52.8 &mu;l of a 0.1 M calcium chloride standard solution and 1 ml of intracellular solution. Mix using a vortex.</li>
<li>To avoid uncaging of caged Ca by ambient light, transfer the solution to a tube protected from light with aluminum foil. Add intracellular solution to reach a final volume of 3.5 ml. Mix using a vortex. Make aliquots of about 1 ml of the caged Ca intracellular solution and store up to two months at -20&deg;C until use.</li> 
</ol>
Notes: During experiments, protect caged compound solutions from light using aluminum foil and keep them on ice.
Sterile filter the intracellular solution.
3. Dissociation of mouse olfactory sensory neurons
Animals were handled in accordance with the Italian Guidelines for the Use of Laboratory Animals (Decreto Legislativo 27/01/1992, no. 116) and European Union guidelines on animal research (No. 86/609/EEC). 
<ol>
<li>Prepare three glass-bottom dishes coated with 10 mg/ml concanavalin-A and 10 mg/ml poly-L-lysine.</li>
<li>To prepare for the dissection, prepare 1 ml of zero-divalent Ringer&#x2019;s solution. Fill two 50 mm Sylgard coated Petri dishes with Ringer&#x2019;s solution and place them on ice. Also chill a papain-cysteine mix, 0.5 ml of the stop mix solution, and a syringe with a 0.22 &mu;m filter containing about 10 ml Ringer&#x2019;s solution.</li>
<li>Sacrifice a mouse by CO2 asphyxiation followed by decapitation. Remove the skin overlying the skull, cut along the jaw lines and place the head in a Sylgard coated Petri dish. </li>
<li>Place the dish under a stereomicroscope. Using a scalpel, sagitally bisect the head along the midline. Turn half of the head with the medial side up. Carefully remove the septum to expose the turbinates (see also Cygnar et al., 201013). </li>
<li>Using number 3 forceps and a scalpel remove turbinates lined with the olfactory epithelium. Transfer to a Sylgard coated Petri dish. </li>
<li> Using number 55 superfine forceps, separate the pieces of olfactory epithelium from the attached parts of turbinates. Transfer them to a tube containing 1 ml zero divalent Ringer&#x2019;s solution. Add the papain and cysteine mix. Using micro-scissors, mince the epithelia and incubate at room temperature for 3 minutes. Gently mince again with micro-scissors and wait for 5 additional minutes.</li>
<li>Using a flame-polished Pasteur pipette, transfer to a tube containing 0.5 ml of the stop mix solution.</li>
<li> Centrifuge at 300 g (1500 RPM) for 5 minutes. Discard the supernatant and add about 1 ml of filtered Ringer&#x2019;s solution. Gently triturate with a flame-polished Pasteur pipette.</li>
<li> Plate about 0.3 ml of the solution containing the cells in the center of the three glass-bottom dishes. Allow cells to settle for about one hour at +4&deg;C. Add Ringer&#x2019;s solution.</li>
</ol>
4. Recording
<ol>
<li>Prepare patch pipettes from borosilicate capillaries to obtain resistances of 3-7 M&Omega; when filled with intracellular solution.</li> 
<li>Place one glass-bottom dish containing the dissociated olfactory sensory neurons on the stage of the inverted microscope. Perfuse with Ringer&#x2019;s solution.</li>
<li>Protect a syringe containing the intracellular solution with caged compounds from light with aluminum foil and keep it refrigerated. Work in dim light to avoid degradation of caged compounds from ambient light.</li>
<li>Look for an isolated olfactory sensory neuron with intact cilia at 100x magnification. Move the stage of the microscope to position the ciliary region inside the circle drawn on the monitor to localize the previously identified UV flash area. </li>
<li>Fill a patch pipette with the intracellular solution containing the caged compound using a flexible needle. Remove bubbles from the pipette tip. Insert the pipette into the holder on the headstage of the patch-clamp amplifier. Approach the soma of the neuron using the micromanipulator and apply gentle suction to obtain a gigaohm seal. Set the voltage at -60 mV and apply additional suction to reach the whole-cell mode using standard patch-clamp techniques12.</li> 
<li>Wait for at least two minutes for the diffusion of the caged compound into the cilia.</li>
<li>On the computer start a stimulation protocol and record the response. Use a low-pass filter at 1 kHz and digitize at a sample frequency of at least 2 kHz.</li>
<li>A typical stimulation protocol clamps the voltage at a constant value and triggers the release of a UV flash from the flash lamp for photolysis of the caged compound in the ciliary region. </li>
</ol>
5. Representative results:
You should be able to produce local uncaging of caged cAMP or of caged Ca in the ciliary region of an isolated olfactory sensory neuron and record the current response in the whole-cell voltage-clamp configuration. 
Figure 4 shows a typical inward current elicited by a UV flash producing photolysis of caged cAMP, recorded at a voltage of -60 mV in the presence of an extracellular low Ca Ringer&#x2019;s solution. In this condition the inward current is due to Na entry through CNG channels. The rising phase of the current was fast and was fitted by a single exponential function with a time constant of 3.4 ms.
Figure 5A-B show the responses of another olfactory sensory neuron in low Ca and in Ringer&#x2019;s solution with 1mM Ca. The rising phase of the current at -60 mV became much slower and multiphasic (Figure 5 A-B). This is due to the action of Ca entering the cilia through CNG channels and activating a secondary Cl current10. The earlier cationic current component, due to activation of CNG, is smaller in 1mM Ca Ringer solution than in low Ca solution because of the block due to the permeating Ca ions that reduce the overall current.
Another way to reduce the increase of Ca in the cilia is to clamp the neuron at +60 mV (Figure 5 C-D). The rising phase of the response due to cAMP uncaging at +60 mV was well described by a single exponential with a time constant of 6.7 ms, indicating the presence of only one current component.
By photoreleasing Ca inside the cilia of an olfactory sensory neuron you should be able to measure a rapidly rising current. This current is carried by Cl ions. Figure 6 A shows inward currents at -50 mV induced by photorelease of caged Ca in response to UV flashes of different intensities. The rising phase of the Ca-activated Cl currents was well described by a single exponential with time constants varying between 3.8 to 5 ms (Figure 6 B).
<img src="/files/ftp_upload/3195/3195fig1.jpg" alt="Figure 1"/> Figure 1. Olfactory transduction in the cilia of olfactory sensory neurons. Odorant molecules bind to odorant receptors (OR) activating a G protein that in turns activates adenylyl cyclase (ACIII) producing an intracellular increase in cAMP. cAMP opens cyclic nucleotide-gated (CNG) channels allowing the entry of Na and Ca ions. The intracellular Ca increase activates Ca-activated Cl channels. Caged cAMP or caged Ca can be introduced in the cilia diffusing through a patch pipette. A flash of UV light produces photolysis of the caged compound (Modified, with permission, from Pifferi et al. 20062).
<img src="/files/ftp_upload/3195/3195fig2.jpg" alt="Figure 2"/> Figure 2. The patch-clamp recording and flash photolysis system. The set-up components include a patch-clamp amplifier, a computer, a digitizer, an epifluorescence microscope, a Xenon flash lamp, a CCD camera and a monitor. Blue and violet lines indicate respectively the visible and UV light path.
<img src="/files/ftp_upload/3195/3195fig3.jpg" alt="Figure 3"/> Figure 3. Xenon flash lamp. (A) Light source used for flash photolysis of caged compounds. (B) Photodiode module used to evaluate the intensity of the light flash. (C) The light guide from the flash lamp was connected to the input of the photodiode and the output was visualized onto an oscilloscope. One of the three available capacitance values (C1, C2 or C3) was selected on the front panel switch of the flash lamp and the voltage was changed turning the knob on the front panel. The output voltage from the photodiode in response to different flash intensities was plotted versus the applied voltage for each capacitance value: C1 = 1000 &mu;F, C2 = 2000 &mu;F, or C3 = 3000 &mu;F. A 600 &mu;m diameter light guide was used.
<img src="/files/ftp_upload/3195/3195fig4.jpg" alt="Figure 4"/> Figure 4. Patch-clamp recording in response to photolysis of caged cAMP in low extracellular Ca solution. (A) Whole-cell current response induced in an isolated olfactory sensory neuron by photolysis of caged cAMP localized to the cilia. A UV flash was released at the time indicated by the arrow. The holding potential was -60 mV. (B) The current rising phase was well fitted with a single exponential function (dotted line) with a time constant of 3.4 ms.
<img src="/files/ftp_upload/3195/3195fig5.jpg" alt="Figure 5"/> Figure 5. Current responses induced by photolysis of caged cAMP in low Ca and in Ringer solutions. (A) An olfactory sensory neuron was bathed in Ringer solution containing 1 mM Ca or in low Ca solution at the holding potential of -60 mV. A UV flash was released at the time indicated by the arrow. (B) Current responses plotted on an expanded timescale showed a multiphasic rising phase in Ringer, while the rising phase was well fitted with a single exponential function (dotted line) with a time constant of 3.5 ms for the response recorded in low Ca solution. (C) Currents responses from the same neuron shown in (A) bathed in Ringer&#x2019;s solution at the holding potential of -60 and +60 mV. (D) Current responses plotted on an expanded timescale displayed a multiphasic rising phase at -60 mV, whereas at +60 mV the rising phase was well fitted by a single exponential with a time constant of 6.7 ms (dotted line). 
<img src="/files/ftp_upload/3195/3195fig6.jpg" alt="Figure 6"/> Figure 6. Responses to photolysis of caged Ca. (A) Whole-cell currents induced by photolysis of caged Ca at -50 mV. UV flashes were released at the time indicated by the arrow. Flash intensities were varied with neutral density filters. (B) Expanded timescale shows the rapid increase in the current after Ca photorelease. Currents were well fitted by a single exponential function (dotted lines), with time constants of 5, 4.8, 3.8 ms. (Reproduced, with permission, from Boccaccio &amp; Menini, 200710).

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No conflicts of interest declared.

Flash photolysis of caged compounds combined with patch-clamp recordings is a useful technique to obtain rapid and local jumps in the concentration of physiologically active molecules both inside and outside cells. Several types of caged compounds1 have been synthesized, and this technique can be applied to various types of cells, including cultured cells expressing ion channels that can be activated or modulated by photolysis of some of the available caged compounds11.
Photolysis o...

<table><tbody><tr><td>Adapter module flash lamp to microscope</td><td>Rapp OptoElectronic</td><td>FlashCube 70</td><td></td></tr><tr><td>Air table</td><td>TMC</td><td>MICRO-g 63-534</td><td></td></tr><tr><td>Digitizer</td><td>Axon Instruments</td><td>Digidata 1322A</td><td></td></tr><tr><td>Data Acquisition Software</td><td>Axon Instruments</td><td>pClamp 8</td><td></td></tr><tr><td>Data Analysis Software</td><td>WaveMetrics</td><td>Igor</td><td></td></tr><tr><td>Mirror for adapter module</td><td>Rapp OptoElectronic</td><td>M70/100</td><td></td></tr><tr><td>Electrode holder</td><td>Axon Instruments</td><td>1-HL-U</td><td></td></tr><tr><td>Faraday&amp;rsquo;s cage</td><td>Custom Made</td><td></td><td></td></tr><tr><td>Filter cube</td><td>Olympus Corporation</td><td>U-MWU</td><td>Excitation filter removed</td></tr><tr><td>Flash lamp</td><td>Rapp OptoElectronic</td><td>JML-C2</td><td></td></tr><tr><td>Forceps Dumont #55 </td><td>World Precision Instruments, Inc.</td><td>14099</td><td></td></tr><tr><td>Glass capillaries </td><td>World Precision Instruments, Inc.</td><td>PG10165-4</td><td></td></tr><tr><td>Glass bottom dish</td><td>World Precision Instruments, Inc.</td><td>FD35-100</td><td></td></tr><tr><td>Illuminator</td><td>Olympus Corporation</td><td>Highlight 3100</td><td></td></tr><tr><td>Inverted microscope</td><td>Olympus Corporation</td><td>IX70</td><td></td></tr><tr><td>Micromanipulators </td><td>Luigs &amp;amp; Neumann</td><td>SM I</td><td></td></tr><tr><td>Micropipette Puller </td><td>Narishige International</td><td>PP-830</td><td></td></tr><tr><td>Monitor</td><td>HesaVision</td><td>MTB-01</td><td></td></tr><tr><td>Neutral density filters</td><td>Omega Optical</td><td>varies</td><td></td></tr><tr><td>Objective 100X</td><td>Carl Zeiss, Inc.</td><td>Fluar 440285</td><td>Either Zeiss or Olympus </td></tr><tr><td>Objective 100X</td><td>Olympus Corporation</td><td>UPLFLN 100XOI2</td><td>Either Zeiss or Olympus</td></tr><tr><td>Optical UV shortpass filter </td><td>Rapp OptoElectronic</td><td>SP400</td><td></td></tr><tr><td>Patch-clamp amplifier</td><td>Axon Instruments</td><td>Axopatch 200B</td><td></td></tr><tr><td>Photo Diode Assembly </td><td>Rapp OptoElectronic</td><td>PDA</td><td></td></tr><tr><td>Quartz light guide</td><td>Rapp OptoElectronic</td><td>varies</td><td>We use 600 &amp;#x03BC;m diameter</td></tr><tr><td>Silver wire</td><td>World Precision Instruments, Inc.</td><td>AGT1025</td><td></td></tr><tr><td>Silver ground pellet</td><td>Warner Instruments</td><td>64-1309</td><td></td></tr><tr><td>Xenon arc lamp</td><td>Rapp OptoElectronic</td><td>XBL-JML</td><td></td></tr><tr><td>Reagent</td><td>Company</td><td>Catalogue number</td><td></td></tr><tr><td>BCMCM-caged cAMP</td><td>BioLog</td><td>B016</td><td></td></tr><tr><td>Bovine serum albumin (BSA)</td><td>Sigma-Aldrich</td><td>A8806</td><td></td></tr><tr><td>CaCl2 standard solution 0.1 M</td><td>Fluka</td><td>21059</td><td></td></tr><tr><td>Caged Ca: DMNP-EDTA</td><td>Invitrogen</td><td>D6814</td><td></td></tr><tr><td>Cysteine</td><td>Sigma-Aldrich</td><td>C9768</td><td></td></tr><tr><td>Concanavalin A type V (ConA)</td><td>Sigma-Aldrich</td><td>C7275</td><td></td></tr><tr><td>CsCl</td><td>Sigma-Aldrich</td><td>C4036</td><td></td></tr><tr><td>DMSO</td><td>Sigma-Aldrich</td><td>D8418</td><td></td></tr><tr><td>DNAse I</td><td>Sigma-Aldrich</td><td>D4527</td><td></td></tr><tr><td>EDTA</td><td>Sigma-Aldrich</td><td>E9884</td><td></td></tr><tr><td>EGTA</td><td>Sigma-Aldrich</td><td>E4378</td><td></td></tr><tr><td>Glucose </td><td>Sigma-Aldrich</td><td>G5767</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H3375</td><td></td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>P3911</td><td></td></tr><tr><td>KOH</td><td>Sigma-Aldrich</td><td>P1767</td><td></td></tr><tr><td>Leupeptin </td><td>Sigma-Aldrich</td><td>L0649</td><td></td></tr><tr><td>MgCl2</td><td>Fluka</td><td>63020</td><td></td></tr><tr><td>Papain</td><td>Sigma-Aldrich</td><td>P3125</td><td></td></tr><tr><td>Poly-L-lysine </td><td>Sigma-Aldrich</td><td>P1274</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>S9888</td><td></td></tr><tr><td>NaOH</td><td>Sigma-Aldrich</td><td>S5881</td><td></td></tr><tr><td>NaPyruvate</td><td>Sigma-Aldrich</td><td>P2256</td><td></td></tr></tbody></table>

flash photolysis of caged compounds in the cilia of olfactory sensory  neurons

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds1. Caged compounds are molecules made physiologically inactive by a chemical cage that can be broken by a flash of ultraviolet light. Here, we show how to obtain patch-clamp recordings combined with photolysis of caged compounds for the study of olfactory transduction in dissociated mouse olfactory sensory neurons. The process of olfactory transduction (Figure 1) takes place in the cilia of olfactory sensory neurons, where odorant binding to receptors leads to the increase of cAMP that opens cyclic nucleotide-gated (CNG) channels2. Ca entry through CNG channels activates Ca-activated Cl channels. We show how to dissociate neurons from the mouse olfactory epithelium3 and how to activate CNG channels or Ca-activated Cl channels by photolysis of caged cAMP4 or caged Ca5. We use a flash lamp6,7 to apply ultraviolet flashes to the ciliary region to uncage cAMP or Ca while patch-clamp recordings are taken to measure the current in the whole-cell voltage-clamp configuration8-11.

Watch this Scientific Journal Video about Flash Photolysis of Caged Compounds in the Cilia of Olfactory Sensory  Neurons at JoVE.com

Flash Photolysis of Caged Compounds in the Cilia of Olfactory Sensory  Neurons

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds. Here, we show how to obtain patch-clamp recordings combined with photolysis of caged cAMP or caged Ca for the study of olfactory transduction in dissociated mouse olfactory sensory neurons.

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds1. ...

flash-photolysis-caged-compounds-cilia-olfactory-sensory

Research

JoVE Journal

Neuroscience

12.7K Views.  International School for Advanced Studies. Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds. Here, we show how to obtain patch-clamp recordings combined with photolysis of caged cAMP or caged Ca for the study of olfactory transduction in dissociated mouse olfactory sensory neurons.

Video: Flash Photolysis of Caged Compounds in the Cilia of Olfactory Sensory  Neurons

Preparation of Living Isolated Vertebrate Photoreceptor Cells for Fluorescence Imaging

In the vertebrate retina, phototransduction, the conversion of light to an electrical signal, is carried out by the rod and cone photoreceptor cells1-4. Rod photoreceptors are responsible for vision in dim light, cones in bright light. Phototransduction takes place in the outer segment of the photoreceptor cell, a specialized compartment that contains a high concentration of visual pigment, the primary light detector. The visual pigment is composed of a chromophore, 11-cis retinal, attached to a protein, opsin. A photon absorbed by the visual pigment isomerizes the chromophore from 11-cis to all-trans. This photoisomerization brings about a conformational change in the visual pigment that 
initiates a cascade of reactions culminating in a change in membrane potential, and bringing about the transduction of the light stimulus to an electrical signal. The 
recovery of the cell from light stimulation involves the deactivation of the intermediates activated by light, and the reestablishment of the membrane potential. Ca2+ modulates the activity of several of the enzymes involved in phototransduction, and its concentration is reduced upon light stimulation. In this way, Ca2+ plays an important role in the recovery of the cell from light stimulation and its adaptation to background light.
Another essential part of the recovery process is the regeneration of the visual pigment that has been destroyed during light-detection by the 
photoisomerization of its 11-cis chromophore to all-trans5-7. This regeneration begins with the release of all-trans retinal by 
the photoactivated pigment, leaving behind the apo-protein opsin. The released all-trans retinal is rapidly reduced in a reaction utilizing NADPH to all-
trans retinol, and opsin combines with fresh 11-cis retinal brought into the outer segment to reform the visual pigment. All-trans retinol is 
then transferred out of the outer segment and into neighboring cells by the specialized carrier Interphotoreceptor Retinoid Binding Protein (IRBP).
Fluorescence imaging of single photoreceptor cells can be used to study their physiology and cell biology. Ca2+-sensitive fluorescent dyes can be used to examine in detail the interplay between outer 
segment Ca2+ changes and response to light8-12 as well as the role of inner segment Ca2+ stores in Ca2+ homeostasis13,14. 
Fluorescent dyes can also be used for measuring Mg2+ concentration15, pH, and as tracers of aqueous and membrane compartments16. 
Finally, the intrinsic fluorescence of all-trans retinol (vitamin A) can be used to monitor the kinetics of its formation and removal in single 
photoreceptor cells17-19.

A method is described for the preparation of single living photoreceptor cells from different vertebrate species for fluorescence imaging. The method can be used to image the fluorescence of endogenous fluorophores, such as NADH or vitamin A, or that of exogenously added fluorescent dyes sensitive to Ca2+ or other factors.

In the vertebrate retina, phototransduction, the conversion of light to an electrical signal, is carried out by the rod and cone photoreceptor cells1-4.  ...

Calcium Imaging of Odor-evoked Responses in the Drosophila Antennal Lobe

The antennal lobe is the primary olfactory center in the insect brain and represents the anatomical and functional equivalent of the vertebrate olfactory bulb1-5. Olfactory information in the external world is transmitted to the antennal lobe by olfactory sensory neurons (OSNs), which segregate to distinct regions of neuropil called glomeruli according to the specific olfactory receptor they express. Here, OSN axons synapse with both local interneurons (LNs), whose processes can innervate many different glomeruli, and projection neurons (PNs), which convey olfactory information to higher olfactory brain regions.
Optical imaging of the activity of OSNs, LNs and PNs in the antennal lobe - traditionally using synthetic calcium indicators (e.g. calcium green, FURA-2) or voltage-sensitive dyes (e.g. RH414) - has long been an important technique to understand how olfactory stimuli are represented as spatial and temporal patterns of glomerular activity in many species of insects6-10. Development of genetically-encoded neural activity reporters, such as the fluorescent calcium indicators G-CaMP11,12 and Cameleon13, the bioluminescent calcium indicator GFP-aequorin14,15, or a reporter of synaptic transmission, synapto-pHluorin16 has made the olfactory system of the fruitfly, Drosophila melanogaster, particularly accessible to neurophysiological imaging, complementing its comprehensively-described molecular, electrophysiological and neuroanatomical properties2,4,17. These reporters can be selectively expressed via binary transcriptional control systems (e.g. GAL4/UAS18, LexA/LexAop19,20, Q system21) in defined populations of neurons within the olfactory circuitry to dissect with high spatial and temporal resolution how odor-evoked neural activity is represented, modulated and transformed22-24.
Here we describe the preparation and analysis methods to measure odor-evoked responses in the Drosophila antennal lobe using G-CaMP25-27. The animal preparation is minimally invasive and can be adapted to imaging using wide-field fluorescence, confocal and two-photon microscopes.

Calcium Imaging of Odor-evoked Responses in the Drosophila Antennal Lobe

We describe an established technique to measure and analyze odor-evoked calcium responses in the antennal lobe of living Drosophila melanogaster.

The antennal lobe is the primary olfactory center in the insect brain and represents the anatomical and functional equivalent of the vertebrate olfactory ...

Using Affordable LED Arrays for Photo-Stimulation of Neurons

Standard slice electrophysiology has allowed researchers to probe individual components of neural circuitry by recording electrical responses of single cells in response to electrical or pharmacological manipulations1,2. With the invention of methods to optically control genetically targeted neurons (optogenetics), researchers now have an unprecedented level of control over specific groups of neurons in the standard slice preparation. In particular, photosensitive channelrhodopsin-2 (ChR2) allows researchers to activate neurons with light3,4. By combining careful calibration of LED-based photostimulation of ChR2 with standard slice electrophysiology, we are able to probe with greater detail the role of adult-born interneurons in the olfactory bulb, the first central relay of the olfactory system. Using viral expression of ChR2-YFP specifically in adult-born neurons, we can selectively control young adult-born neurons in a milieu of older and mature neurons. Our optical control uses a simple and inexpensive LED system, and we show how this system can be calibrated to understand how much light is needed to evoke spiking activity in single neurons. Hence, brief flashes of blue light can remotely control the firing pattern of ChR2-transduced newborn cells.

Adult-born neurons expressing ChR2 can be manipulated in slice electrophysiological preparations in order to examine their contribution towards the function of olfactory neural circuits.

Standard slice electrophysiology has allowed researchers to probe individual components of neural circuitry by recording electrical responses of single ...

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic development, cellular signaling and adult physiology. In this respect, the use of multi-photon microscopy enables quantitative photoactivation of a given light responsive agent in deep tissues at a single cell resolution. As zebrafish embryos are optically transparent, their development can be monitored in vivo. These traits make the zebrafish a perfect model organism for controlling the activity of a variety of chemical agents and proteins by focused light. Here we describe the use of two-photon microscopy to induce the activation of chemically caged fluorescein, which in turn allows us to follow cell's destiny in live zebrafish embryos. We use embryos expressing a live genetic landmark (GFP) to locate and precisely target any cells of interest. This procedure can be similarly used for precise light induced activation of proteins, hormones, small molecules and other caged compounds.

Multiphoton microscopy allows control of low energy photons with deep optical penetration and reduced phototoxicity. We describe the use of this technology for live cell labeling in zebrafish embryos. This protocol can be readily adapted for photo-induction of various light-responsive molecules.

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic ...

Biology

Multi-photon Intracellular Sodium Imaging Combined with UV-mediated Focal Uncaging of Glutamate in CA1 Pyramidal Neurons

Multi-photon fluorescence microscopy has enabled the analysis of morphological and physiological parameters of brain cells in the intact tissue with high spatial and temporal resolution. Combined with electrophysiology, it is widely used to study activity-related calcium signals in small subcellular compartments such as dendrites and dendritic spines. In addition to calcium transients, synaptic activity also induces postsynaptic sodium signals, the properties of which are only marginally understood. Here, we describe a method for combined whole-cell patch-clamp and multi-photon sodium imaging in cellular micro domains of central neurons. Furthermore, we introduce a modified procedure for ultra-violet (UV)-light-induced uncaging of glutamate, which allows reliable and focal activation of glutamate receptors in the tissue. To this end, whole-cell recordings were performed on Cornu Ammonis subdivision 1 (CA1) pyramidal neurons in acute tissue slices of the mouse hippocampus. Neurons were filled with the sodium-sensitive fluorescent dye SBFI through the patch-pipette, and multi-photon excitation of SBFI enabled the visualization of dendrites and adjacent spines. To establish UV-induced focal uncaging, several parameters including light intensity, volume affected by the UV uncaging beam, positioning of the beam as well as concentration of the caged compound were tested and optimized. Our results show that local perfusion with caged glutamate (MNI-Glutamate) and its focal UV-uncaging result in inward currents and sodium transients in dendrites and spines. Time course and amplitude of both inward currents and sodium signals correlate with the duration of the uncaging pulse. Furthermore, our results show that intracellular sodium signals are blocked in the presence of blockers for ionotropic glutamate receptors, demonstrating that they are mediated by sodium influx though this pathway. In summary, our method provides a reliable tool for the investigation of intracellular sodium signals induced by focal receptor activation in intact brain tissue.

We describe the combination of focal UV-induced photo-activation of neuro-active compounds with whole-cell patch-clamp and multi-photon imaging of intracellular sodium transients in dendrites and spines of hippocampal neurons in acute tissue slices of the mouse brain.

Multi-photon fluorescence microscopy has enabled the analysis of morphological and physiological parameters of brain cells in the intact tissue with high ...

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

Whole-cell voltage clamp recordings from Drosophila melanogaster photoreceptors have revolutionized the field of invertebrate visual transduction, enabling the use of D. melanogaster molecular genetics to study inositol-lipid signaling and Transient Receptor Potential (TRP) channels at the single-molecule level. A handful of labs have mastered this powerful technique, which enables the analysis of the physiological responses to light under highly controlled conditions. This technique allows control over the intracellular and extracellular media; the membrane voltage; and the fast application of pharmacological compounds, such as a variety of ionic or pH indicators, to the intra- and extracellular media. With an exceptionally high signal-to-noise ratio, this method enables the measurement of dark spontaneous and light-induced unitary currents (i.e.&#160;spontaneous and quantum bumps) and macroscopic Light-induced Currents (LIC) from single D. melanogaster photoreceptors. This protocol outlines, in great detail, all the key steps necessary to perform this technique, which includes both electrophysiological and optical recordings. The fly retina dissection procedure for the attainment of intact and viable ex vivo isolated ommatidia in the bath chamber is described. The equipment needed to perform whole-cell and fluorescence imaging measurements are also detailed. Finally, the pitfalls in using this delicate preparation during extended experiments are explained.

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

Whole-cell recordings from Drosophila melanogaster photoreceptors enable the measurement of spontaneous dark bumps, quantum bumps, macroscopic responses to light, and current-voltage relationships under various conditions. In combination with D. melanogaster genetic manipulation tools, this method enables the study of the ubiquitous inositol-lipid signaling pathway and its target, the TRP channel.

Whole-cell voltage clamp recordings from Drosophila melanogaster photoreceptors have revolutionized the field of invertebrate visual transduction, ...

FM Dye Cycling at the Synapse: Comparing High Potassium Depolarization, Electrical and Channelrhodopsin Stimulation

FM dyes are used to study the synaptic vesicle (SV) cycle. These amphipathic probes have a hydrophilic head and hydrophobic tail, making them water-soluble with the ability to reversibly enter and exit membrane lipid bilayers. These styryl dyes are relatively non-fluorescent in aqueous medium, but insertion into the outer leaflet of the plasma membrane causes a &#62;40X increase in fluorescence. In neuronal synapses, FM dyes are internalized during SV endocytosis, trafficked both within and between SV pools, and released with SV exocytosis, providing a powerful tool to visualize presynaptic stages of neurotransmission. A primary genetic model of glutamatergic synapse development and function is the Drosophila neuromuscular junction (NMJ), where FM dye imaging has been used extensively to quantify SV dynamics in a wide range of mutant conditions. The NMJ synaptic terminal is easily accessible, with a beautiful array of large synaptic boutons ideal for imaging applications. Here, we compare and contrast the three ways to stimulate the Drosophila NMJ to drive activity-dependent FM1-43 dye uptake/release: 1) bath application of high [K+] to depolarize neuromuscular tissues, 2) suction electrode motor nerve stimulation to depolarize the presynaptic nerve terminal, and 3) targeted transgenic expression of channelrhodopsin variants for light-stimulated, spatial control of depolarization. Each of these methods has benefits and disadvantages for the study of genetic mutation effects on the SV cycle at the Drosophila NMJ. We will discuss these advantages and disadvantages to assist the selection of the stimulation approach, together with the methodologies specific to each strategy. In addition to fluorescent imaging, FM dyes can be photoconverted to electron-dense signals visualized using transmission electron microscopy (TEM) to study SV cycle mechanisms at an ultrastructural level. We provide the comparisons of confocal and electron microscopy imaging from the different methods of Drosophila NMJ stimulation, to help guide the selection of future experimental paradigms.

Synaptic vesicle (SV) cycling is the core mechanism of intercellular communication at neuronal synapses. FM dye uptake and release are the primary means of quantitatively assaying SV endo- and exocytosis. Here, we compare all the stimulation methods to drive FM1-43 cycling at the Drosophila neuromuscular junction (NMJ) model synapse.

FM dyes are used to study the synaptic vesicle (SV) cycle. These amphipathic probes have a hydrophilic head and hydrophobic tail, making them ...

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. Learning-induced changes in synaptic transmission are often distributed across many neurons and levels of processing in the brain. Therefore, methods to visualize learning-dependent synaptic plasticity across neurons are needed. The fruit fly Drosophila melanogaster represents a particularly favorable model organism to study neuronal circuits underlying learning. The protocol presented here demonstrates a way in which the processes underlying the formation of associative olfactory memories, i.e., synaptic activity and their changes, can be monitored in vivo. Using the broad array of genetic tools available in Drosophila, it is possible to specifically express genetically encoded calcium indicators in determined cell populations and even single cells. By fixing a fly in place, and opening the head capsule, it is possible to visualize calcium dynamics in these cells whilst delivering olfactory stimuli. Additionally, we demonstrate a set-up in which the fly can be subjected, simultaneously, to electric shocks to the body. This provides a system in which flies can undergo classical olfactory conditioning - whereby a previously na&#239;ve odor is learned to be associated with electric shock punishment - at the same time as the representation of this odor (and other untrained odors) is observed in the brain via two-photon microscopy. Our lab has previously reported the generation of synaptically localized calcium sensors, which enables one to confine the fluorescent calcium signals to pre- or postsynaptic compartments. Two-photon microscopy provides a way to spatially resolve fine structures. We exemplify this by focusing on neurons integrating information from the mushroom body, a higher-order center of the insect brain. Overall, this protocol provides a method to examine the synaptic connections between neurons whose activity is modulated as a result of olfactory learning.

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Here we present a protocol with which pre- and/or postsynaptic calcium can be visualized in the context of Drosophila learning and memory. In vivo calcium imaging using synaptically localized calcium sensors is combined with a classical olfactory conditioning paradigm such that the synaptic plasticity underlying this type of associative learning may be determined.

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. ...

Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and spatial resolution on the order of ms and nm, respectively. We have developed a time-resolved electron microscopy technique, &#34;flash-and-freeze,&#34; that induces cellular events with optogenetics and visualizes the resulting membrane dynamics by freezing cells at defined time points after stimulation. To demonstrate this technique, we expressed channelrhodopsin, a light-sensitive cation channel, in mouse hippocampal neurons. A flash of light stimulates neuronal activity and induces neurotransmitter release from synaptic terminals through the fusion of synaptic vesicles. The optogenetic stimulation of neurons is coupled with high-pressure freezing to follow morphological changes during synaptic transmission. Using a commercial instrument, we captured the fusion of synaptic vesicles and the recovery of the synaptic vesicle membrane. To visualize the sequence of events, large datasets were generated and analyzed blindly, since morphological changes were followed in different cells over time. Nevertheless, flash-and-freeze allows the visualization of membrane dynamics in electron micrographs with ms temporal resolution.

We developed a novel technique in electron microscopy, &#34;flash-and-freeze,&#34; that enables the visualization of membrane dynamics with ms temporal resolution. This technique combines the optogenetic stimulation of neurons with high-pressure freezing. Here, we demonstrate the procedures and describe the protocols in detail.

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and ...

Flash Photolysis of Caged Compounds in the Cilia of Olfactory Sensory Neurons

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Preparation of Living Isolated Vertebrate Photoreceptor Cells for Fluorescence Imaging

Calcium Imaging of Odor-evoked Responses in the Drosophila Antennal Lobe

Using Affordable LED Arrays for Photo-Stimulation of Neurons

In vivo Neuronal Calcium Imaging in C. elegans

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

Multi-photon Intracellular Sodium Imaging Combined with UV-mediated Focal Uncaging of Glutamate in CA1 Pyramidal Neurons

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

FM Dye Cycling at the Synapse: Comparing High Potassium Depolarization, Electrical and Channelrhodopsin Stimulation

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy