This research was supported by grants from the Israel Science Foundation (ISF), and the United States-Israel Binational Science Foundation (BSF). We thank Mr. Anatoly Shapochnikov for the construction of the wax filament heater.

1. Measuring the intensity response relationship, prolonged depolarizing afterpotential (PDA), and the early receptor potential (ERP) using the electroretinogram
<ol>
	<li>Suitable rearing conditions for D. melanogaster preparation
	<ol>
		<li>Raise D. melanogaster flies in bottles containing standard yellow corn containing food in an incubator maintained at a temperature of 24 &#176;C and in a 12 h dark/light cycle</li>
		<li>Keep the fly bottles in the dark at least 24 h prior to the experiment.</li>
	</ol>
	</li>
	<li>General setup
	<ol>
		<li>Prepare recording pipettes by pulling 1 mm x 0.58 mm (O.D x I.D) fiber-filled borosilicate glass capillaries (Figure 6L, O). The resistance of the pipettes should be 5-10 M&#937;; any suitable puller can be used.</li>
		<li>Coat two silver wires with AgCl2, inserting 0.25 mm silver wire into 3 M KCl solution connected to a custom-made 5 V power supply.</li>
		<li>Insert each coated silver wire into the electrode holders (Figure 6N).</li>
		<li>Fill the glass capillary with filtered Ringer&#39;s solution (see Table 1) using an elongated tip syringe (Figure 6M).</li>
		<li>Insert the wire electrode into the glass capillary. Ensure that the solution within the capillary is in contact with the silver wire.</li>
		<li>Insert the electrode holders (Figure 7P, N) into the two electrode micromanipulators (Figure 7G).</li>
	</ol>
	</li>
	<li>Procedure of preparing the fly for electrical recordings 
	NOTE: To keep the fly under dark-adapted conditions, use only dim red light illumination during the following steps.
	<ol>
		<li>Anesthetize the flies in the bottle with CO2 gas using the fly sleeper system (Figure 6A, B) and pour them into the sleeper container.</li>
		<li>Choose one fly and carefully hold it by its wing using a sharp tweezer. Cover the rest of the flies with a Petri dish.</li>
		<li>Place the fly on the fly holder in the proper orientation-lying on its side, with its back toward the hand (Figure 6P).</li>
		<li>Turn ON the power supply of the soldering iron. Set the current to ~2.25 A. This current should heat the 0.25 mm platinum-iridium filament to ~55-56 &#176;C (see Supplemental File).</li>
		<li>Place a drop of wax with a low melting temperature (~55-56 &#176;C) on the soldering iron (Figure 6F).</li>
		<li>Using tweezers, lift the fly from its wings and fix its wings to the fly holder (Figure 6I) using the soldering iron.</li>
		<li>Using the soldering iron, connect the fly&#39;s back to the stand surface with wax (Figure 6P).</li>
		<li>Lower the tip of the soldering iron onto the joining point of the legs and melt the wax to cover all the legs together (Figure 6P).</li>
		<li>Place a small drop of wax between the head and the back in the neck area (Figure 6P). 
		NOTE: Take special care to avoid overheating the fly head. Ensure that the fly is properly fixed and is unable to move during the experiment; minor movements may create artifacts in the recordings. Ensure that the trachea openings (breathing inlets) in the thorax and abdomen are not covered with wax.</li>
		<li>Place the fly holder (Figure 7Q) in a dark Faraday cage on a magnet block (Figure 7I) and ensure that the fly is ~5 mm from the end of the light guide (Figure 7L).</li>
		<li>Place the recording electrode (Figure 7P) above the fly&#39;s eye and the ground electrode (Figure 7N) over the fly&#39;s upper back using the micromanipulators.</li>
		<li>Insert the ground electrode into the back of the fly using the micromanipulators.</li>
		<li>Insert the recording electrode into the outer periphery of the fly&#39;s eye, preferably, using the micromanipulators. 
		NOTE: After inserting the electrode into the eye, a small dimple will be observed; pull the electrode upwards without removing it from the eye until the dimple disappears. The electrodes can also be immersed in small droplets of electrode jelly applied at the torso and eye.</li>
	</ol>
	</li>
	<li>Intensity-response protocol
	<ol>
		<li>Place an orange filter (590 edge filter) in front of the high-pressure Xenon lamp. Use a large (six orders of magnitude) attenuating neutral density (ND) filter.</li>
		<li>Wait 60 s in the dark and give a 5 s light pulse.</li>
		<li>Replace the ND filter with a less attenuating ND filter in one order of magnitude increments.</li>
		<li>Wait 60 s in the dark and give a second 5 s light pulse.</li>
		<li>Repeat Steps 1.4.1.-1.4.4, gradually increasing the light intensity using less attenuating ND filters (the last pulse should be generated with no ND filter at all). Ensure to use the series of ND filters in the proper direction, starting from large attenuation and reaching low attenuation.</li>
	</ol>
	</li>
	<li>PDA protocol (this protocol can only be performed on white-eyed flies)
	<ol>
		<li>Give a 5 s light pulse of maximal intensity using an orange filter (590 edge filter, to convert maximal photopigment to the R state).</li>
		<li>Replace the orange filter with a broad-band blue (BP450/40 nm) filter and give three 5 s light pulses at maximum intensity. 
		NOTE: It is also possible to give a long continuous maximum intensity blue light pulse until a steady-state voltage response is reached.</li>
		<li>Wait 60 s in the dark, replace the blue filter with the previous orange filter, and give two 5 s light pulses with 60 s intervals.</li>
	</ol>
	</li>
	<li>ERP/M-potential protocol for measuring the photo-equilibrium spectrum of M (this protocol can only be performed on white-eyed flies10,25)
	<ol>
		<li>Give a continuous blue (bandpass (BP) 450/40 nm) light pulse until reaching a steady-state voltage response, which converts the maximal amount of photopigment from the R state to the M state of R1-6 cells.</li>
		<li>Give a brief (&#60;3 ms) intense light flash of a wavelength between 350-700 nm (the known absorption spectrum of R1-6 cells photopigment) using narrow (~20 nm) bandpass filters and measure the peak amplitude of the M1 phase of the M potential response (which reflects the metarhodopsin absorption at this specific wavelength at photo-equilibrium10,25). 
		NOTE: For synchronous activation of a large pool of photopigment molecules, a brief, intense flash of light is required so that the photon content will be packed in a short duration. The M potential is composed of two components: M1 (corneal negative phase), which reflects the charge displacement of the metarhodopsin in the photoreceptor, and M2 (corneal positive phase11,33), which reflects the amplified M1 response of the photoreceptors in the lamina9,10,11. Each of these components can be identified and measured. However, it is preferable to measure the M1 potential since it is a direct linear manifestation of the M levels. If M2 is used, make sure that its amplitude is in the linear range by using mild carotenoid deprivation24,25.</li>
		<li>Give a continuous blue (BP450/40 nm) light pulse again, followed by a brief (&#60;1 ms) intense light flash of a different wavelength.</li>
		<li>Repeat this protocol until the entire absorption spectrum of M at photo equilibrium is covered.</li>
	</ol>
	</li>
</ol>
2. ERC protocol for measuring the action spectrum of R and the M states of R1-6 cells using whole-cell voltage-clamp recordings
NOTE: For a detailed protocol for using whole-cell voltage-clamp recordings, see Katz et al.34. The M-potential uses the ERG to measure the activation of the M state only because the contribution of the R state is suppressed by the membrane capacitance. In contrast, the ERC measures the activation of both R (positive ERC) and M (negative ERC) states because voltage-clamp recordings remove the effect of membrane capacitance (see introduction).
<ol>
	<li>Convert the photopigment to the desired state (R or M). For R to M conversion, first, adapt the flies by a brief (&#60;1 ms) adaptive blue (BP450/40 nm) flash. For M to R conversion, give a short adaptive orange (OG590 edge filter) flash.</li>
	<li>Give a brief light flash (&#60;1 ms) of a wavelength between 350-700 nm and measure the maximal negative or positive amplitude of the ERC response, which reflects the M/R absorption at this specific wavelength. 
	NOTE: For synchronous activation of a large pool of photopigment molecules, a brief, intense flash of light is required to pack the photon content in a short duration. The maximal photopigment conversion from R to M by blue flash can reach ~80% of the total photopigment molecules because of the overlap in the absorption spectrum of R and M (Figure 3A). Therefore, at wavelengths below ~550 nm, the ERC has two components: a negative phase that reflects the response of the metarhodopsin, and a positive phase that reflects the response of the remaining rhodopsin. The ERC depends linearly on light intensity (Figure 5D). Accordingly, to derive the spectral sensitivity of R and M states, each phase of the ERC needs to be normalized at the various wavelengths for equal energy29.</li>
	<li>Repeat Steps 2.1.-2.2. using flashes of different wavelengths.</li>
	<li>Plot the normalized positive and negative ERC as a function of wavelength. 
	NOTE: The strong flash of light induces a massive current through the light-sensitive channels leading to metabolic stress on the photoreceptor, which, in turn, causes light-independent channel opening. The ERC is superimposed on this constitutive current.</li>
</ol>

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A., Tsacopoulos, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+recovery+from+a+transient+inhibition+of+the+oxidative+metabolism+of+the+photoreceptors+of+the+drone+(Apis+mellifera).">The recovery from a transient inhibition of the oxidative metabolism of the photoreceptors of the drone (Apis mellifera).</a> Journal of Experimental Biology. 119, 165-181 (1985).</li><li>Agam, K., Frechter, S., Minke, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+the+Drosophila+TRP+and+TRPL+channels+requires+both+Ca+2++and+protein+dephosphorylation.">Activation of the Drosophila TRP and TRPL channels requires both Ca 2+ and protein dephosphorylation.</a> Cell Calcium. 35 (2), 87-105 (2004).</li></ol>

The authors declare no conflict of interest.

The major advantage of using the Drosophila photoreceptor preparation is its accessibility, the ease and accuracy of light stimulation, and, most importantly, the ability to apply the power of molecular genetics7. Extensive genetic studies have established Drosophila as an extremely useful model system for the genetic dissection of complex biological processes7. The relatively simple structure of the Drosophila genome (consisting of only four chro...

The light-activated rhodopsin (R), which is a G-protein-coupled receptor (GPCR), is composed of a 7 transmembrane protein (opsin) and a chromophore. In Drosophila melanogaster (fruit fly), photon absorption induces isomerization of the 11-cis-3-OH-retinal chromophore to all-trans-3-OH-retinal1, promoting the conformational change of the rhodopsin to metarhodopsin (M, Figure 1A). Unlike vertebrate rhodopsin, the predominant fraction of invertebrate&#160;chromophore does not dissociate from the opsin, resulting in the physiologically active dark-stable pigment state M. In turn, additional photon absorption by the all-trans-3-OH-retinal chromophore induces isomerization of the chromophore2,3, generating the R pigment state with the 11-cis-3-OH-retinal chromophore. The R state is a dark, stable, and physiologically non-active photopigment. In addition to the extremely fast photon regeneration route of the chromophore4, much like vertebrate&#160;photopigments, an alternative enzymatic slow route for chromophore regeneration exists in invertebrates, in which some of the stages are performed in retinal cells surrounding the photoreceptors cells5,6.
Drosophila&#160;entails great advantages as a model organism for studying invertebrate photoreceptors. In particular, the accessibility of the preparation and the ability to apply molecular genetics have made Drosophila a powerful model system7. Hence, several in vivo and ex vivo experimental methods for studying phototransduction in general and photopigment levels, in particular, have been established. The simplest in-vivo method exploits the relatively large extracellularly recorded voltage response to light of the Drosophila eye. Accordingly, light stimulation evokes an electrical voltage response in the entire eye that can be measured using extracellular electroretinogram (ERG) recording, which is ~3 orders of magnitude larger than the ERG response to light of vertebrate eyes8,9. The Drosophila ERG response is robust and easily obtained, which makes it a convenient method for identifying abnormalities in light response due to mutations. The ERG response to light arises mainly from the photoreceptors, pigment (glia) cells, and secondary neurons of the lamina (see Figure 1B). The main components of the ERG are (i) the extracellular voltage response of the photoreceptors, (ii) the &#34;on&#34; and &#34;off&#34; transients at the beginning and end of the light stimulus that arise from the lamina neurons (Figure 2A,&#160;inset, ON, OFF), (iii) the slow response of the glia cells (Figure 2A, inset, arrows), and (iv) the brief and transient response, resulting from charge displacement during photopigment activation that precedes the ON transient10 (Figure 2C [inset],&#160;D,&#160;E). This brief response is composed of two phases (M1 and M2, Figure 2C [inset]) and can be induced only by extremely strong light stimulation, which activates simultaneously millions of photopigment molecules. It is neither observed under blue stimulation (Figure 2D, blue trace) nor in mutants with highly reduced photopigment levels (Figure 2E, red trace), but its amplitude is mildly enhanced in a mutant that abolishes PLC activity (Figure 2E, orange trace). The M1 phase is a typical ERP of the fly arising from the activation of M in the photoreceptors. The M1 phase, which has a positive polarity (intracellularly), releases a neurotransmitter in the normal way in a sign-inverting synapse and activates the lamina neurons, which respond to the photoreceptor depolarization by generating the synaptically amplified M2 phase. Thus, both M1 and M2 phases reflect M activation10,11.
The depolarization of the photoreceptor generates the corneal-positive &#34;on&#34; transient, arising from the sign-inverting synapse between the photoreceptor axon and the monopolar neurons of the lamina10,11 (Figure 1B). The slow rise and decay of the ERG arise from the depolarization of the pigment cells (Figure 2A, inset, arrows) mainly due to K+ efflux from the photoreceptor cells12 via the transient receptor potential (TRP) and TRP-like (TRPL) channels13,14,15. These slow kinetic components largely mask and distort the waveform of the photoreceptor response when compared to intracellular or whole-cell recordings of the photoreceptor response to light9,10. In addition, at very strong illuminations, an additional transient response, which precedes and partially fuses with the &#34;on&#34; transient, may be observed (Figure 2C [inset],D,E). This signal originates directly from the massive activation of the photopigment10.
Several light regime protocols using neutral density (ND) and color filters, as well as strong illuminating flashes, have been developed to investigate the eye in general and the phototransduction cascade in particular. These protocols have also been used to investigate the properties of the photopigment.
The intensity-response protocol measures the peak amplitude of the ERG voltage response of the entire eye to increasing light intensities (Figure 2A,B). This protocol assists in detecting changes in the sensitivity of the photoreceptor cells to light9.
The prolonged depolarizing afterpotential (PDA) protocol&#160;exploits the differences in the absorption spectra of rhodopsin and metarhodopsin that allows, in Drosophila, a massive photopigment conversion of R to its physiologically active and dark-stable intermediate M state2. In the ERG voltage response, a relatively short pulse of saturating light is given, and the resulting voltage response is recorded. Under this condition, a ceiling (reversal potential) is reached by the depolarization signal because activation of a fraction of a percent of the huge amount of rhodopsin molecules (~1 x 108) is sufficient to reach the ceiling. The presence of the phototransduction components in great abundance ensures that this ceiling will be reached even in mutants with a significant reduction in concentration or subtle malfunction of the phototransduction components. This situation precludes the isolation of these mutants. Pak et al. introduced the PDA screening7 seeking a reliable and revealing test to isolate visual mutants. In Drosophila, the PDA response is brought about by genetically removing the red screening pigment, which allows photopigment conversion, and the application of blue light, which is preferentially absorbed by rhodopsin (Figure 3A) and, thus, results in a large net conversion of the R to the M photopigment state. Phototransduction termination is disrupted at the level of the photopigment by a large net conversion of R to M, which, in turn, results in sustained excitation long after the light is turned off (Figure 2C, Figure 4A [top]). During the PDA period, the photoreceptors are less sensitive to subsequent test lights and are partially desensitized (inactivated). The PDA detects even minor defects in rhodopsin biogenesis and tests the maximal capacity of the photoreceptor cell to maintain excitation for an extended period. Since it strictly depends on the presence of high concentrations of rhodopsin, it easily scores for deficient replenishment of the phototransduction components. Remarkably, the PDA screen has yielded many new and very important visual mutants (reviewed in Pak et al.7). Thus, the PDA mutants isolated by Pak et al.7 are still extremely useful for analyzing the Drosophila visual system.
The PDA is induced in Drosophila by saturating blue light, resulting in continuous depolarization long after light offset (Figure 4A [top]). After saturating PDA-inducing blue light, the peripheral photoreceptors (R1-6) remain continuously active in the dark at their maximal capacity, reaching saturation. Additional saturating blue lights during the PDA do not produce any additional response in R1-6 cells for many seconds but induce a response in R7-8 cells that is superimposed on the PDA. The superimposed responses are explained by the different absorption spectra of the photopigments expressed in these cells (R7-8)16. The PDA can be suppressed by the photoconversion of M back to R with saturating orange light (Figure 4A [top]). The ability of the PDA to bring the photoreceptor cells to their maximal active capacity, a situation that cannot be achieved by intense white light, explains why it has been a major tool to screen for visual mutants of Drosophila. This is because it allows the detection of even minor defects in proteins involved in the biogenesis of normal photopigment levels17,18. Two groups of PDA defective mutants have been isolated: neither inactivation nor afterpotential (nina) mutants and inactivation but not afterpotential (ina) mutants. The phenotype of the former is a lack of a PDA and the associated inactivation arising from a large reduction in the photopigment levels (Figure 4A [middle]). The phenotype of the latter shows inactivation but no dark depolarization after blue light due to a still-unknown mechanism in the mutant with normal rhodopsin levels but lacking proteins interacting with the TRP channels (Figure 4A [bottom]).
The PDA arises from the difference in the amount of photopigment relative to arrestin (ARR2), which binds and terminates M activity19,20,21 (Figure 1A). In Drosophila photoreceptors, the amount of the photopigment is about fivefold larger than the amount of ARR219. Thus, ARR2 levels are insufficient to inactivate all the M molecules generated by a large net photoconversion of R to M, leaving an excess of M constantly active in the dark17,19,20,22,23. This mechanism explains the elimination of the PDA response by mutations or by carotenoid deprivation24,25, causing a reduction in photopigment level, but does not affect arrestin levels. Moreover, this explanation also accounts for the phenotypes of null ARR2 (arr23)&#160;mutant allele21, in which PDA could be achieved at ~10 fold dimmer blue light intensities19,20,21 (Figure 4B,C). The PDA is not a unique feature of fly photoreceptors, and it appears in every tested species that has dark stable M with an absorption spectrum different from that of the R state, allowing sufficient photoconversion of the photopigment from the R to the M state. A thoroughly investigated species in which the PDA phenomenology was discovered is the barnacle (Balanus) photoreceptor, in which the absorption spectrum of the R state is in a longer wavelength than the M state2 (Figure 3B). Accordingly, unlike the situation in the fly, in the barnacle, orange-red light induces a PDA, while blue light suppresses the PDA2.
The early receptor potential (ERP) protocol exploits the charge displacement occurring during R or M activation. The visual pigment is an integral part of the surface membrane of the signaling compartment of both vertebrate&#160;and invertebrate membranes3. Accordingly, the activation process in which the photopigment molecules change from one intermediate state to the next is accompanied by a charge displacement4,26. As the photopigment molecules are electrically aligned in parallel with the membrane capacitance4, a rapid synchronized conformational change generates a fast polarization change of the surface membrane, which, in flies, occurs in the signaling compartment composed of a stack of ~30,000-50,000 microvilli called rhabdomere. This polarization then discharges passively through the membrane capacitance of the cell body until the cell membrane is equally polarized. The ERP is the extracellular recording of the charge displacement. The intracellularly recorded ERP manifests the extracellular ERP integrated by the time constant of the cell membrane4,27,28. The current activated by the visual pigment charge displacement could also be measured in whole-cell voltage-clamp recordings29,30 (Figure 5A-D), with the major advantage (in early receptor current (ERC) recordings) of minimizing the effect of membrane capacitance on the kinetics of the signal.
The protocol section describes how to perform ERG measurements from Drosophila eye9 and ERC measurements by whole-cell recordings from Drosophila isolated ommatidia31,32. We also describe specific protocols that are used to investigate phototransduction in general and photopigments in particular.

<table><tbody><tr><td>1 mL syringe with elongated tip</td><td></td><td></td><td>Figure 6M</td></tr><tr><td>1 rough tweezers</td><td></td><td>Dumont #5, Standard</td><td>0.1 mm x 0.06 mm, length 110 mm, Inox (Figure 6H)</td></tr><tr><td>2 condenser lenses</td><td></td><td></td><td></td></tr><tr><td>A/D converter</td><td>Molecular Device</td><td>Digidata 1200</td><td>Possible replacement: any digidata from molecular devices (e.g 1440A) -Figure 7C</td></tr><tr><td>Amplifier</td><td>Almost perfect electronics</td><td></td><td>Possible replacement: Warner instruments- IE251A or IE-210 (comes with headstage)- Figure 7D</td></tr><tr><td>Anti-vibration Table</td><td>Newport</td><td>VW-3036-OPT-01</td><td>Figure 7H</td></tr><tr><td>Capillaries</td><td>Harvard Apparatus</td><td>Borosilicate glass capillaries</td><td>1 mm x 0.58 mm (Figure 6O)</td></tr><tr><td>Clampex</td><td>Molecular Device</td><td></td><td>Software</td></tr><tr><td>CO&lt;sub&gt;2 &lt;/sub&gt;tank</td><td></td><td></td><td></td></tr><tr><td>Cold light source</td><td>Schott</td><td>KL1500 LCD</td><td>Figure 6C</td></tr><tr><td>Delicate wipers</td><td>Kimtech</td><td></td><td>Kimwipes (Figure 6K)</td></tr><tr><td>Electrode holder</td><td></td><td></td><td>Suitable for capillary O.D. 1 mm (Figure 6N, Figure 7N, and Figure 7P)</td></tr><tr><td>Faraday cage</td><td>Home made</td><td></td><td>Electromagnetic noise shielding and black front curtain (Figure 7K)</td></tr><tr><td>Filter (Color)</td><td>Schott</td><td>OG590, Edge filter</td><td>Figure 7S</td></tr><tr><td>Filter (Color)</td><td>Schott</td><td>BP450/40 nm</td><td>Figure 7S</td></tr><tr><td>Filter (Color)</td><td>Blazers</td><td>550 nm</td><td>Figure 7S</td></tr><tr><td>Filter (Color) for cold light source</td><td>Schott</td><td>RG630</td><td>Figure 6C</td></tr><tr><td>Filter (Heat)</td><td>Schott</td><td>KG3</td><td>Figure 7S</td></tr><tr><td>Filters (Neutral density filter)</td><td>Chroma</td><td>6,5,4,3,2,1,0.5,0.3</td><td>Figure 7S</td></tr><tr><td>Flash Lamp system</td><td>Honeywell</td><td></td><td>Figure 7U</td></tr><tr><td>Fly sleeper system with injector</td><td>Inject + matic</td><td></td><td>Figure 6A-B</td></tr><tr><td>Lamp power supply</td><td>PTI</td><td>LPS-220</td><td>Figure 7W</td></tr><tr><td>Light detector</td><td>Home made</td><td></td><td>Phototransistor (Figure 7O)</td></tr><tr><td>Light guide</td><td></td><td></td><td>3 mm diameter, 1.3 m long (Figure 7L,M)</td></tr><tr><td>Light source</td><td></td><td></td><td>High-pressure ozone-free 75 W Xenon lamp (operating on 50 W), possible replacement: Cairn research- OptoLED (Figure 7R)</td></tr><tr><td>Low temperature melting wax</td><td>Home made</td><td></td><td>Composed of mixture of beeswax (T&lt;sub&gt;m&lt;/sub&gt;&amp;asymp;62 &amp;deg;C) and paraffin at ~3:1 to reach a melting temperature of ~55&amp;ndash;56 &amp;deg;C (Figure 6J)</td></tr><tr><td>Magnetic stand for flies</td><td>Home made</td><td></td><td>Figure 6I and Figure 7Q</td></tr><tr><td>Microelectrode preamplifier system with head-stage</td><td>Almost perfect electronics</td><td></td><td>Impedance tester (Figure 7G)</td></tr><tr><td>Micromanipulator (mechanical coarse)</td><td>Tritech Research, Narishige</td><td>M-2</td><td></td></tr><tr><td>Micromanipulator (mechanical fine)</td><td>Leitz Microsystems</td><td>Leitz Mechanical Micromanipulator</td><td>Figure 7F</td></tr><tr><td>pCLAMP</td><td>Molecular Device</td><td></td><td>Software</td></tr><tr><td>Petri dish</td><td></td><td></td><td>60 mm</td></tr><tr><td>Pulse generator</td><td>AMPI</td><td>Master 8</td><td>Figure 7A</td></tr><tr><td>Redux cream for electrocardiography</td><td>Parker Laboratories</td><td>Redux Electrolyte Cr&amp;egrave;me</td><td></td></tr><tr><td>Shutter driver</td><td>Uniblitz, Vincent Associates</td><td>VCM-D1 Single Channel Uni-stable</td><td>Figure 7V</td></tr><tr><td>Shutter system</td><td>Uniblitz, Vincent Associates</td><td>LS2 2 mm Uni-stable Shutters</td><td>Figure 7V</td></tr><tr><td>Silver Wire</td><td>Warner Instruments</td><td></td><td>0.25&amp;ndash;1 mm diameter, needs to be chloridized</td></tr><tr><td>Soldering iron composed of a platinum-iridium filament</td><td></td><td></td><td>0.25 mm diameter (Figure 6F)</td></tr><tr><td>Stereoscopic zoom Microscope</td><td>Nikon</td><td>SMZ-2B</td><td>Figure 6D</td></tr><tr><td>Stereoscopic zoom Microscope</td><td>Wild</td><td>Wild M5</td><td>With 6, 12, 25 and 50 magnification settings (Figure 7E)</td></tr><tr><td>Syringe filters</td><td>Millex</td><td></td><td>22 &amp;micro;m PVDF filter</td></tr><tr><td>Vertical pipette puller</td><td>Sutter/ Narishige</td><td>Model P-97/PP-830</td><td>Use either vertical or horizontal puller, as preferred (Figure 6L)</td></tr><tr><td>Wax filament heater</td><td>Home made</td><td></td><td>See figure S1 (Figure 6E-G)</td></tr><tr><td>Xenon Flash Lamp system</td><td>Dr. Rapp OptoElectronic</td><td>JML-C2</td><td>Figure 7X</td></tr></tbody></table>

electrophysiological methods for measuring photopigment levels in drosophila photoreceptors

The Drosophila G-protein-coupled photopigment rhodopsin (R) is composed of a protein (opsin) and a chromophore. The activation process of rhodopsin is initiated by photon absorption-inducing isomerization of the chromophore, promoting conformational changes of the opsin and resulting in a second dark-stable photopigment state (metarhodopsin, M). Investigation of this bi-stable photopigment using random mutagenesis requires simple and robust methods for screening mutant flies. Therefore, several methods for measuring reductions in functional photopigment levels have been designed. One such method exploits the charge displacements within the photopigment following photon absorption and the huge amounts of photopigment molecules expressed in the photoreceptors. This electrical signal, named the early receptor potential (or early receptor current), is measured by a variety of electrophysiological methods (e.g., electroretinogram and whole-cell recordings) and is linearly proportional to functional photopigment levels. The advantages of this method are the high signal-to-noise ratio, direct linear measurement of photopigment levels, and independence of phototransduction mechanisms downstream to rhodopsin or metarhodopsin activation. An additional electrophysiological method called prolonged depolarizing afterpotential (PDA) exploits the bi-stability of Drosophila photopigment and the absorption-spectral differences of fly R and M pigment states. The PDA is induced by intense blue light, converting saturating amounts of rhodopsin to metarhodopsin, resulting in the failure of light-response termination for an extended time in darkness, but it can be terminated by metarhodopsin to rhodopsin conversion using intense orange light. Since the PDA is a robust signal that requires massive photopigment conversion, even small defects in the biogenesis of the photopigment lead to readily detected abnormal PDA. Indeed, defective PDA mutants led to the identification of novel signaling proteins important for phototransduction.

Figure 2&#160;exemplifies the robustness and ease of using the ERG technique. It is robust because it is recorded in the virtually intact fly by a simple technique of extracellular voltage recordings that require a simple electrophysiological setup. The robustness is manifested by obtaining recordings of light responses with relatively large amplitudes (in the millivolt range) even when mutations strongly reduce or distort the light response. Therefore, even an inexperienced experimenter can...

Watch this Scientific Journal Video about Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors at JoVE.com

Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors

We present a protocol to electrophysiologically characterize bi-stable photopigments: (i) exploiting the charge displacements within the photopigment molecules following photon-absorption and their huge amount in the photoreceptors, and (ii) exploiting the absorption-spectra differences of rhodopsin and metarhodopsin photopigment states. These protocols are useful to screen for mutations affecting bi-stable photopigment systems.&#160;

Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors

The Drosophila G-protein-coupled photopigment rhodopsin (R) is composed of a protein (opsin) and a chromophore. The activation process of rhodopsin is ...

electrophysiological-methods-for-measuring-photopigment-levels

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Neuroscience

2.1K Views.  Hebrew University. We present a protocol to electrophysiologically characterize bi-stable photopigments: (i) exploiting the charge displacements within the photopigment molecules following photon-absorption and their huge amount in the photoreceptors, and (ii) exploiting the absorption-spectra differences of rhodopsin and metarhodopsin photopigment states. These protocols are useful to screen for mutations affecting bi-stable photopigment systems. 

Video: Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors

Quantifying Abdominal Pigmentation in Drosophila melanogaster

Pigmentation is a morphologically simple but highly variable trait that often has adaptive significance. It has served extensively as a model for understanding the development and evolution of morphological phenotypes. Abdominal pigmentation in Drosophila melanogaster has been particularly useful, allowing researchers to identify the loci that underlie inter- and intraspecific variations in morphology. Hitherto, however, D. melanogaster abdominal pigmentation has been largely assayed qualitatively, through scoring, rather than quantitatively, which limits the forms of statistical analysis that can be applied to pigmentation data. This work describes a new methodology that allows for the quantification of various aspects of the abdominal pigmentation pattern of adult D. melanogaster. The protocol includes specimen mounting, image capture, data extraction, and analysis. All the software used for image capture and analysis feature macros written for open-source image analysis. The advantage of this approach is the ability to precisely measure pigmentation traits using a methodology that is highly reproducible across different imaging systems. While the technique has been used to measure variation in the tergal pigmentation patterns of adult D. melanogaster, the methodology is flexible and broadly applicable to pigmentation patterns in myriad different organisms.

Quantifying Abdominal Pigmentation in Drosophila melanogaster

This work presents a method to quickly and precisely quantify the abdominal pigmentation of Drosophila melanogaster using digital image analysis. This method streamlines the procedures between phenotype acquisition and data analysis and includes specimen mounting, image acquisition, pixel value extraction, and trait measurement.

Pigmentation is a morphologically simple but highly variable trait that often has adaptive significance. It has served extensively as a model for ...

Genetics

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular recording has historically been a common technique used to study sensitivities of individual photoreceptor cells to different light stimuli that is still being used today. However, there remains a dearth of detailed methodology in the literature for researchers wishing to replicate intracellular recording experiments in the eye. Here we present the insect as a model for examining eye physiology more generally. Insect photoreceptor cells are located near the surface of the eye and are therefore easy to reach, and many of the mechanisms involved in vision are conserved across animal phyla. We describe the basic procedure for in vivo intracellular recording of photoreceptor cells in the eye of a butterfly, with the goal of making this technique more accessible to researchers with little prior experience in electrophysiology. We introduce the basic equipment needed, how to prepare a live butterfly for recording, how to insert a glass microelectrode into a single cell, and finally the recording procedure itself. We also explain the basic analysis of raw response data for determining spectral sensitivity of individual cell types. Although our protocol focuses on determining spectral sensitivity, other stimuli (e.g., polarized light) and variations of the method are applicable to this setup.

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

The electrophysiological technique of intracellular recording is demonstrated and used to determine spectral sensitivities of single photoreceptor cells in the compound eye of a butterfly.

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at the synaptic level. The Active Zone (AZ) is the region at the presynaptic membrane that mediates neurotransmitter release and is composed of a dense collection of scaffold proteins. AZs of Drosophila melanogaster (Drosophila) photoreceptors undergo molecular remodeling after prolonged exposure to natural ambient light. Thus the level of neuronal activity can rearrange the molecular composition of the AZ and contribute to the regulation of the functional output.
Starting from the light exposure set-up preparation to the immunohistochemistry, this protocol details how to quantify the number, the spatial distribution, and the delocalization level of synaptic molecules at AZs in Drosophila photoreceptors. Using image analysis software, clusters of the GFP-fused AZ component Bruchpilot were identified for each R8 photoreceptor (R8) axon terminal. Detected Bruchpilot spots were automatically assigned to individual R8 axons. To calculate the distribution of spot frequency along the axon, we implemented a customized software plugin. Each axon&#39;s start-point and end-point were manually defined and the position of each Bruchpilot spot was projected onto the connecting line between start and end-point. Besides the number of Bruchpilot clusters, we also quantified the delocalization level of Bruchpilot-GFP within the clusters. These measurements reflect in detail the spatially resolved synaptic dynamics in a single neuron under different environmental conditions to stimuli.

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

Here we show how to quantify the number and spatial distribution of synaptic active zones in Drosophila melanogaster&#160;photoreceptors, highlighted with a genetically encoded molecular marker, and their modulation after prolonged exposure to light.

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at ...

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, ...

Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of Drosophila guttifera

Diversified species of Drosophila (fruit fly) provide opportunities to study mechanisms of development and genetic changes responsible for evolutionary changes. In particular, the adult stage is a rich source of morphological traits for interspecific comparison, including wing pigmentation comparison. To study developmental differences among species, detailed observation and appropriate staging are required for precise comparison. Here we describe protocols for staging of pupal periods and quantification of wing pigmentation in a polka-dotted fruit fly, Drosophila guttifera. First, we describe the method for detailed morphological observation and definition of pupal stages based on morphologies. This method includes a technique for removing the puparium, which is the outer chitinous case of the pupa, to enable detailed observation of pupal morphologies. Second, we describe the method for measuring the duration of defined pupal stages. Finally, we describe the method for quantification of wing pigmentation based on image analysis using digital images and ImageJ software. With these methods, we can establish a solid basis for comparing developmental processes of adult traits during pupal stages.

Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of Drosophila guttifera

Protocols for staging pupal periods and measurement of wing pigmentation of Drosophila guttifera are described. Staging and quantification of pigmentation provide a solid basis for studying developmental mechanisms of adult traits and enable interspecific comparison of trait development.

Diversified species of Drosophila (fruit fly) provide opportunities to study mechanisms of development and genetic changes responsible for evolutionary ...

Biology

Electrophysiological Recording of Voltage Responses of Drosophila Retinal Photoreceptors to Light Stimuli

Source: Juusola, M., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/54142">Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo.</a> J. Vis. Exp. (2016).
This video outlines the protocol for electrophysiological recording of photoreceptor activity in the eye of an immobilized Drosophila, enabling precise measurement of voltage changes in response to light stimuli.

1. Recording from R1-R6 Photoreceptors 

	Always be grounded when operating the microelectrode amplifier (for example by touching the metal surface of the ...

JoVE Encyclopedia of Experiments

Neurophysiology

Electrophysiology Techniques

Investigating Prolonged Depolarizing Afterpotential (PDA) in Drosophila Photoreceptors

Source: Gutorov, R., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/63514">Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors.</a> J. Vis. Exp. (2022).
This video demonstrates an experimental protocol to investigate the prolonged depolarizing afterpotential (PDA) in white-eyed Drosophila. The fly&#39;s eye is exposed to intense blue light pulses to convert the photopigment rhodopsin to metarhodopsin, initiating a signaling cascade that opens positive ion channels and causes membrane depolarization. Because blue light prevents the reconversion of metarhodopsin to rhodopsin, the continuous influx of positive ions results in sustained depolarization, known as the prolonged depolarizing afterpotential (PDA).

1. Measuring the prolonged depolarizing afterpotential (PDA) using the electroretinogram

	Suitable rearing conditions for Drosophila melanogaster ...

Stimulation and Modulation Techniques

Whole-Cell Voltage Clamp Recordings from Drosophila Photoreceptors

Source:&#160; Katz, B., et al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/55627">Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors.</a> J. Vis. Exp. (2017)
The video demonstrates recording electrical activity in the photoreceptor neurons of Drosophila&#39;s ommatidium by establishing a whole-cell configuration. It highlights the activation of transient receptor potential channels by light pulses, resulting in measurable quantum bumps.

NOTE: During all of the following steps, use only dim red light illumination and ensure that the ommatidia&#39;s exposure to light is minimal (i.e., work ...

Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells

To prepare a fly in a non-lethal variation, transfer the ice anesthetized fly headfirst into a 200-microliter pipette tip, and carefully push the fly toward the tip with compressed air. Then, using a scalpel, cut off the pipette tip just in front of the head. And using tweezers, carefully push the fly a few millimeters into the pipette tip.Cut off the pipette tip again. And push the fly back toward the tip with the compressed air, so that only the head of the fly protrudes from the pipette tip. After adhering a piece of plasticine onto an object slide, press the pipette tip into it so that the left or the right eye faces upward. Ensure that the eye is oriented correctly under the microscope. For image acquisition, select a water-immersion objective.In the case of a non-lethal variation, use a laboratory pipette to adhere a large drop of chilled water to the underside of the water immersion objective. Carefully place the object slide with the prepared fly onto the microscope stage. Then, lower the water immersion objective manually until it contacts the water surface, or the fly's eye touches the drop.Then, switch the light path toward the microscope camera, and generate a live image. Readjust the focus for the camera and evaluate the orientation of the eye, considering that the eye must face the microscope objective radially. Use the lookup table software to detect oversaturation.In the case of non-pigmented flies, adjust the exposure time such that the brightest pixels are just below the saturation limit for every image. Record an image and save it as a raw file to archive all corresponding metadata of the recording. Then, export the image in a dot TIFF format.To quantify relative EGFP fluorescence in the rhabdomeres of water-immersion micrographs, adjust ImageJ settings by clicking on Analyze. Then, set measurements, and check only the box for mean gray value. Import the dot TIFF image by clicking on File, then Open. Choose a representative region of the image in focus, and enlarge it to 200% to 300% by repeatedly pressing Control and Plus together.Next, select the Oval tool. And while pressing the Shift key, generate a circular selection in the image that is significantly smaller than one fluorescent rhabdomere. Then, look for the exact size displayed below the toolbar in the ImageJ main window. To measure the fluorescence intensities within the circular selection, move the circle to the first rhabdomere with the arrow keys on the keyboard, and click Analyze, then measure.A result window listing the measured gray value will pop up. Continue with measurements of rhabdomeres 2 to 6, and also measure the background signal. In the case of non-pigmented flies, make additional measurements of the corresponding cell body areas. Measure the fluorescence of two more ommatidia, resulting in three technical replicates. Mark the analyzed ommatidia by using the pencil tool and save this image for documentation.Select and copy the measured gray values from the result window and paste them into spreadsheet software for further calculations. Sort the fluorescence intensity values according to their origin into the category's rhabdomere, cell body, and background, and calculate the mean intensity from each category. Then calculate the relative amount of EGFP present in the rhabdomere, using the first formula for non-pigmented eyes, and the second for pigmented eyes.

Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors

Summary

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Chapters in this video

Quantifying Abdominal Pigmentation in Drosophila melanogaster

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

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

Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of Drosophila guttifera

Electrophysiological Recording of Voltage Responses of Drosophila Retinal Photoreceptors to Light Stimuli

Investigating Prolonged Depolarizing Afterpotential (PDA) in Drosophila Photoreceptors

Whole-Cell Voltage Clamp Recordings from Drosophila Photoreceptors

Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells