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. 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.


Introduction
Extensive genetic studies of the fruit fly, Drosophila melanogaster (D. melanogaster), initiated more than 100 years ago, have established the D. melanogaster fly as an extremely useful experimental model for the genetic dissection of complex biological processes. The methodology described below combines the accumulated power of D. melanogaster molecular genetics with the high signal-to-noise ratio of whole-cell patch clamp recordings. This combination allows for the study of D. melanogaster phototransduction as a model of inositol-lipid signaling and TRP channel regulation and activation, both in the native environment and at the highest resolution of single molecules.
Application of the whole-cell recording method to D. melanogaster photoreceptors has revolutionized the study of invertebrate phototransduction. This method was developed by Hardie 1 and independently by Ranganathan and colleagues 2 ~26 years ago and was designed to exploit the extensive genetic manipulation tools of D. melanogaster and use them to uncover mechanisms of phototransduction and inositol-lipid signaling. At first, this technique suffered from a rapid reduction in light sensitivity and a low yield of ommatidia during the dissection process, which prevented detailed quantitative studies. Later, the addition of ATP and NAD to the patch pipette dramatically increased the suitability of the preparation for prolonged quantitative recordings. Thereafter, extensive characterization of the signal-transduction mechanism at the molecular level was realized.
Currently, D. melanogaster phototransduction is one of the few systems in which phosphoinositide signaling and TRP channels can be studied ex vivo at single-molecule resolution. This makes D. melanogaster phototransduction and the methodology developed to study this mechanism a highly sensitive model system. This protocol describes how to dissect the D. melanogaster retina and mechanically strip the isolated ommatidia from the surrounding pigment (glia) cells. This enables the formation of a giga-seal and a whole-cell patch clamp on the photoreceptor cell bodies. Fortunately, most signaling proteins are confined to the rhabdomere and do not diffuse. In addition, there is an immobile Ca 2+ buffer called calphotin, located between the signaling compartment and the cell body 3,4 , and a high expression level of the Na + /Ca 2+ exchanger (CalX) in the microvilli 5 . Together, the protein confinement to the rhabdomere, the calphotin buffer, and the high expression of the CalX allow for relatively prolonged (i.e. up to ~20 min) whole-cell recordings, without the loss of essential components of the phototransduction process and while maintaining high sensitivity to light. The following protocol describes how to obtain isolated ommatidia and perform whole-cell recordings that appear to preserve the native properties of the phototransduction cascade. Whole-cell patch clamp experiments on dissociated cockroach 1. Raise D. Melanogaster flies at a low population density (i.e. ~20 flies in a 6 oz. bottle) in bottles containing standard cornmeal food at 19-24°C . NOTE: It is preferable to work on dark-adapted flies. To maintain high sensitivity to light, reduce the diversity and prevent retinal degeneration in mutant flies. 2. Rear the flies in the dark for at least 24 h prior to the experiment. NOTE: The flies used for the experiments should be recently eclosed (<2 h) and still soft, pale, and display the meconium. Ommatidia can also readily be prepared from pupae, though their sensitivity to light is then steeply dependent on age 10 .

Retina Dissection and Ommatidia Isolation: Option 1
NOTE: Perform all of the following steps under the stereoscopic zoom microscope using an amplification suitable for properly viewing the preparation (See Figure 1).
1. Place four drops of ES-0Ca 2+ and one drop of TS solution on a 60-mm Petri dish that has been turned over.
2. Using rough tweezers, catch a newly eclosed (<2 h after eclosion) fly by its wings or body. From this point on, perform all procedures rapidly and under dim red illumination at 20 ±1 °C. 3. While still grasping the fly with the rough tweezers, use the first pair of fine tweezers to detach the fly head from the body. Submerge the head in the first ES-0Ca 2+ drop.
4. Dissect the head in half along the sagittal plane using the second pair of fine tweezers. Ensure that, at the end of this step, both eyes are still intact. 5. Transfer one half of the head to the second ES-0Ca 2+ drop and the other half to the third ES-0Ca 2+ drop.
6. Using the fine tweezers, remove as much of the tissue surrounding the eye as possible and ensure that no harm is caused to the retina. 7. Firmly grasp the edge of one cornea with the fine tweezers and scoop out the retina using the scooper. NOTE: Upon completion of this step, the cornea will be left empty and intact, separated from an intact retina. 8. Rinse the trituration pipette connected to the tubing with DDW and fill the pipette with a small amount of ES-0Ca 2+ from the fourth drop.
NOTE: This step must be performed every time a new ommatidia-separating pipette is used and removed from the ethanol beaker (the solution filling the pipette should match the solution in which the retina is submerged). 9. Gently aspirate by mouth to draw the isolated retina into the pipette. Use extreme caution not to aspire air bubbles into the pipette. 10. Transfer the isolated retina to the drop of TS. Perform steps 4.6-4.10 on the second eye as well. 11. Wipe away the drops of ES-0Ca 2+ using delicate wipes, leaving only the drop of TS containing both retinae on the Petri dish. Add six more drops of TS to the top of the Petri dish. Transfer both retinas to one of the other TS drops. 12. Replace the trituration pipette with a pipette of a smaller-diameter opening. Rinse it as described in step 4.8, using TS as the solution to fill the pipette. 13. Rapidly and repeatedly aspirate and expirate both retinae in the solution to begin the separation of isolated ommatidia stripped of pigment cells from the whole retina. NOTE: The isolated ommatidia are visible in the TS drop, and as the isolation process progresses, the TS drop becomes less translucent. 14. Transfer the remaining retinae to the next TS drop. Fill the pipette with the entire former TS drop (containing the isolated ommatidia) and expirate the drop into the bath chamber. 15. Repeat steps 4.12-4.14 to achieve maximal isolated ommatidia. Wait approximately 1 min to allow the isolated ommatidia to sink and bind to the bottom of the bath chamber. 16. Using the perfusion system, start the flow of ES-0Ca 2+ with 1.5 mM Ca 2+ into the bath chamber. Ensure that the chamber is completely filled with the solution, from bottom to top, and that the ground is completely submerged in the solution. Continue to wash the bath 4-5x.

Retina Dissection and Ommatidia Isolation: Option 2
Note: Perform all of the following steps under the stereoscopic zoom microscope, using an amplification suitable for properly viewing the preparation (See Figure 1).
3. For the dissection, create a large drop (<0.5 mL) of ES solution on the silicone dissection block. Add two "reservoir" drops (~50 µL each) of TS solution to a 60 mm Petri dish 4. Immobilize a newly eclosed (<2 h post-eclosion) fly in a glass tube on ice and pick it up by its wings using tweezers. From this point on, perform all procedures rapidly and under dim red illumination at 20 ±1 °C. 5. Holding the fly with tweezers, cut off the fly's head using a razor blade chip mounted in a holder. Pick up an insect pin (12 mm long, 0.1 mm in diameter) with the tweezers and pierce the head between the eyes. 6. Briefly submerge the head in 70% ethanol; this prevents air bubbles from forming on the head/eye surface. Pin the head under the ES drop on the silicone dissection dish. 7. Cut off both eyes using the razor blade chip by using a sawing motion along the line of the frontal margin of the eye. 8. Firmly grasp the edge of one cornea with the fine tweezers. 9. Scoop out the retina using the scooper.
NOTE: Upon completion of this step, the cornea will be left empty and intact, separated from an intact retina. 10. Without damaging the retina, use the tweezers and scooper to gently remove adhering air sacs and excess brain tissue.
NOTE: The preparation of isolated retinae is also useful for Western blot analyses of non-specific retinae proteins 11 , whole-mount histology, and whole-retina imaging. Patch clamp recordings can also be conducted on photoreceptors from the whole retina 12 . 11. Take the trituration pipette with the largest diameter, connect it to the tubing, and backfill the pipette with a small amount of TS from one of the reservoir drops in the Petri dish by gentle suction (i.e. by mouth). Perform this step every time a new ommatidia trituration pipette is used. 12. Gently blow the TS over the two retinae by mouth and then draw the isolated retinae into the pipette using gentle suction. Use caution to ensure that no air bubbles enter the pipette. 13. Transfer the isolated retinae to the Petri dish, forming a small drop (~20 µL), and wash them once or twice with TS from one of the reservoir drops. 14. Incubate the retinae in the dark for 20-25 min. 15. Replace the ommatidia trituration pipette with a pipette with a smaller-diameter opening (using fresh TS from one of the reservoir drops as in step 4.6 to backfill the pipette). 16. Rapidly aspirate and expirate both retinae in a small drop (~20 µL) to begin the separation of the isolated ommatidia.
NOTE: At the first stage, the surrounding pigmented glia should disintegrate, leaving visible small debris in the solution. 17. After substantial small debris has accumulated, but before many ommatidia have separated, use fresh TS from one of the reservoir drops and transfer the retinae to a new small drop. 18. Select a smaller-diameter trituration pipette, backfill, and continue to triturate.
NOTE: As ommatidia now begin to separate, their elongated forms should be clearly visible under the high power of the stereomicroscope. If necessary, keep changing the trituration pipettes to smaller diameters until a good yield of ommatidia is visible 19. Once a reasonable yield of ommatidia are visible and the drop is no longer translucent, fill the pipette with the entire drop containing the isolated ommatidia and expirate the drop gently into the bottom of the bath chamber pre-filled with ES. 20. Wait approximately 1 min to allow the isolated ommatidia to sink and settle on the bottom of the bath chamber. 21. Using the perfusion system, start the flow of ES into the bath chamber. Ensure that the chamber is completely full of the solution, from bottom to top, and that the ground is totally submerged in the solution. Continue to wash the bath 4-5x. NOTE: Thereafter, continuous perfusion is not required, though the bath should be briefly flushed before introducing a new patch pipette. NOTE: During all of the following steps, use only dim red light illumination and ensure that the exposure of the ommatidia to light is minimal (i.e. work rapidly and turn off the dissecting light source and chamber red illumination used for viewing the ommatidia when performing tasks that do not demand viewing the ommatidium). In addition, perform all of the following steps according to standard electrophysiological protocol.

Whole-Cell Recording
1. Under an inverted microscope (40X objective), carefully inspect all of the ommatidia in the bath and choose a suitable ommatidium for the experiment.
1. Ensure that the outer membrane of the ommatidium is smooth and intact, that the long axis is approximately at a right angle relative to the electrode approach direction (as seen in Figure 3A), and that the distal section of the ommatidium is not surrounded by any excess tissue. Place the chosen ommatidium at the optical axis of the objective lens (in the center of the field of vision) to ensure uniform illumination. 4. Blow into the pipette by mouth, through the tube connected to the electrode holder, causing it to fill with positive pressure. Close the tube valve to maintain the pressure. 5. Using the micromanipulator, insert the electrode into the bath chamber. 6. Guide the electrode close to the distal section of the ommatidium, such that there is no contact between the electrode and the ommatidium, until a small dimple (due to the positive pressure in the patch pipette) can be observed in the ommatidium. 7. Open the recording software (see the Table of Materials). Open the "membrane test module" to apply continuous square voltage pulses of 2 mV at a rate of 100 Hz. 8. Set the junction potential to "zero," by adjusting the appropriate knob in the patch clamp amplifier to set the base of the square pulse to "zero" current NOTE: The electrophysiological setup includes a head stage (i.e. first-stage amplification) connected to an amplifier (i.e. second-stage amplification). The amplified analogue signal is converted to a digital signal using the A/D converter, which is controlled by software installed on a PC computer. 9. Release the positive pressure in the pipette by opening the valve of the tube connected to the electrode holder. Gently create negative pressure in the pipette by sucking out of the tube, leading to the association of the pipette to the cell membrane. Close the tube valve to maintain the pressure. 10. Ensure that the electrode resistance viewed on the computer screen is elevated to 100 -150 MΩ. Release the negative pressure in the pipette by manually opening the valve of the tube connected to the electrode holder. 11. Ensure that the electrode resistance is elevated to at least 1-2 GΩ. NOTE: At this point, a seal has been formed between the electrode and the photoreceptor. 12. Offset the capacitive currents of the pipette by adjusting the appropriate knob in the patch clamp amplifier. 13. Create rapid, short, and forceful bouts of negative pressure in the electrode, sucking by mouth out of the tube connected to the electrode holder to "break" into the photoreceptor membrane and create a whole-cell configuration. Alternatively, use the "Zap button" to apply short, rectangular electrical pulses, starting with a duration of "0.1 ms," or apply a combination of both methods. NOTE: The generation of the whole-cell configuration is revealed by a sudden increase in pipette capacitance (typically ~60 pF for a wildtype R1-6 photoreceptor; a capacitance of only ~20 pF indicates a recording from an R7 photoreceptor; a capacitance above ~90 pF indicates a recording from two photoreceptors). 14. Set the holding potential of the photoreceptor to the required voltage (usually, -70 mV), manually using the appropriate knob in the patch clamp amplifier. NOTE: It is possible to perform this step after a seal has been obtained (step 6.11) and before the whole-cell configuration has been achieved. 15. Offset the capacitive currents and series resistance (a measured series resistance value greater than 25 MΩ indicates that the electrode pipette has been clogged) and, if required (i.e. for larger currents), apply series resistance compensation using the appropriate knobs in the patch clamp amplifier. 16. Close the black front curtain of the Faraday cage to obtain maximal darkness and electrical isolation. 17. Begin the recording process using the software and administer light stimuli and/or pharmacological substances according to the desired experimental procedure.

Simultaneous Whole-Cell Recordings and Ca 2+ Imaging
1. For genetically encoded Ca 2+ indicators, isolate the ommatidia as described above using D. melanogaster flies expressing GCaMP6f 13 . Use a CCD camera (see the Table of Materials) for the fluorescence measurement and ensure that the microscope is equipped with proper excitation and emission filters and a dichroic mirror (see the  (Figure 4; see the Table of Materials), isolate the ommatidia as described above. In addition, ensure that the pipette solution includes a 20-100 µM calcium indicator. 3. Use the imaging software (see the Table of  ).

Representative Results
The described method has enabled the accurate recording of the fundamental unitary currents that generate spontaneous and light-evoked quantum bumps, which sum to produce the macroscopic response to light, under defined conditions. It also allowed the comparison between wildtype and mutant flies that have defects in critical signaling molecules (Figures 3 and 5) 14,15,16,17,18 . In addition, the ability to measure reversal potential under bi-ionic conditions revealed fundamental biophysical properties of the TRP and TRP-like (TRPL) channels 18,19 . It also enabled the measurement of the effects of amino acid substitutions in the pore region of TRP that modified its Ca 2+ permeability 20 .
The light response obtained by patch clamp whole-cell recordings depends linearly on light intensity for at least 4 orders of magnitude. This could not be resolved by using ERG and intracellular recording methods. Accordingly, a series of responses to brief flashes of increasing intensity and a plot of the intensity response function revealed a strict linearity of the flash response with increasing light intensity. The strict linearity holds up to at least several hundred pA, but it is debatable whether thereafter it is linearity or clamp control that breaks down (Figure 6). These results suggest that the macroscopic responses to light are a linear summation of the unitary responses to light (i.e. quantum bumps).
It has been well established using voltage recordings that dim light stimulation induces discrete voltage fluctuations (i.e. quantum bumps) in most invertebrate species. The D. melanogaster quantum bumps result from the concerted opening of ~15 TRP channels and ~2 TRPL channels at the peak of the bump 18 . Each bump is generated by the absorption of a single photon, while the macroscopic response to more intense lights is the summation of these elementary responses 14,21     Store at -20 °C. Store at -20 °C. Table 4: Intracellular Solution (IS2). Chemical description and the specific quantities required to produce Intracellular Solution IS2, which is mostly used for reversal potential measurements of the light-induced current.

Discussion
The application of whole-cell recordings to D. melanogaster photoreceptors allowed for the discovery and the functional elucidation of novel signaling proteins, such as TRP channels 27,28,29 and INAD 30,31,32 scaffold protein. Ever since the initial introduction of this technique, it enabled the resolution of long-term basic questions regarding the ionic mechanism and voltage dependence of the light response. This occurred because of the conferred ability to accurately control the membrane voltage and extracellular and intracellular ionic composition 19,28 .
A major obstacle of the patch clamping technique in D. melanogaster has been the fragility of the isolated ommatidia preparation. Detailed studies have revealed that the integrity of the phototransduction machinery critically depends on the continuous supply of ATP, especially during light exposure, which leads to a large consumption of ATP. Unfortunately, the mechanical striping of the pigment (i.e. glia) cells, which is required to reach the photoreceptor membrane with the patch pipette, eliminates the main source of metabolites necessary for ATP production 33 . Application of exogenous ATP into the recording pipette only partially fulfills the requirement for large quantities of ATP. A short supply of ATP leads to spontaneous activation of the TRP channels and to the dissociation of the phototransduction machinery from the light-activated channels, causing a large increase in cellular Ca 2+ and the abolishment of the normal response to light 34,35 . This sequence of events is not due to damage of the photoreceptors by the dissection procedure, but rather to the cellular depletion of ATP. To prevent this sequence of events from occurring and to maintain normal light responses, the photoreceptors should not be exposed to intense lights, which consume large quantities of ATP. Also, NAD must be included in the recording pipette, presumably to facilitate ATP production in the mitochondria 18,36 . For measurements of spontaneous and quantum bumps, the above difficulty is minimal because only dim lights are used. In practice, a stable whole-cell recording can be maintained for ~20-25 min, although there is a tendency for response kinetics to slow down over this period. A single preparation of dissociated ommatidia may remain viable for up to 2 h.
An additional shortcoming of the isolated ommatidia preparation is the inaccessibility of the microvilli, which translates to the inaccessibility of the TRP and TRPL channels to the recording pipette, preventing single-channel recordings. Using a method they developed, Bacigalupo and colleagues succeeded at directly recording single-channel activity from the rhabdomere 37 . However, this channel activity differs from that of TRPL channels heterologously expressed in tissue culture cells 38 and from TRP channel activity derived from shot noise analysis obtained from isolated ommatidia 34 . Presumably, the dissection procedure greatly damaged the photoreceptor cells when using this method.

Disclosures
The authors have nothing to disclose.