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, 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.
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 Hardie1 and independently by Ranganathan and colleagues2 ~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 Ca2+ buffer called calphotin, located between the signaling compartment and the cell body3,4, and a high expression level of the Na+/Ca2+ exchanger (CalX) in the microvilli5. 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 (Periplaneta americana)6 and cricket (Gryllus bimaculatus)7 ommatidia were performed similarly to that described for D. melanogaster. In addition, patch clamp experiments on dissociated photoreceptors of the file clam, (Lima scabra) and scallop (Pecten irradians) were performed in a slightly different manner from that conducted on D. melanogaster, allowing both whole-cell8 and single-channel measurements9. Here, the major achievements obtained in D. melanogaster using this technique are described. The Discussion includes the description of some pitfalls and limitations of this technique.
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 channels27,28,29 and INAD30,31,32 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 composition19,28.
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 production33. 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 Ca2+ and the abolishment of the normal response to light34,35. 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 mitochondria18,36. 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 rhabdomere37. However, this channel activity differs from that of TRPL channels heterologously expressed in tissue culture cells38 and from TRP channel activity derived from shot noise analysis obtained from isolated ommatidia34. Presumably, the dissection procedure greatly damaged the photoreceptor cells when using this method.
The authors have nothing to disclose.
The experimental part of this research was supported by grants from the US-Israel Bi National Science Foundation (to B.M. and I.L.), the Israel Science Foundation (ISF), the Deutsch-Israelische Projektkooperation (DIP) (to B.M.), and the Biotechnology and Biological Sciences Research Council (BBSRC Grant numbers: BB/M007006/1 and BB/D007585/1) to R.C.H.
10 ml syringe | |||
5 ml syringe | |||
1 ml syringe with elongated tip | |||
Petri dish | 60 mm | ||
Silicone dissection dish/block | Dow Corning | Sylguard 184 Silicone Elastomer Kit | Throughly mix both components, pour into mould and cure for 2 hours at 60ᵒC |
Syringe filters | Millex | 22 µm PVDF filter | |
Capillaries (for omatidia separation) | Glass, 1.2 x 0.68 mm (~7.5 cm each) | ||
Polyethylene Tubing | Becton Dickinson | 1.57 x 1.14 mm (35 cm) | |
2 small beakers | 50 ml or less | ||
2 paraffin film sheets | Parafilm M | ~5 x 5 cm | |
Bath chamber | home made | ||
Cover slips | 22 x 22 mm No. 0 | ||
Paraplast Plus | Sigma | Paraffin – polyisobutylene mixture | To glue the coverslip onto the bottom of the bath chamber |
Ground | Warner Instruments | 64-1288 | Hybrid Assembly Ag-AgCl Wire Assembly |
Headless micro dissection needle | Entomology, 12 mm | ||
Micro dissecting needle holder | |||
Vise | |||
2 fine tweezers + 1 rough tweezers | Dumont #5, Biology | 0.05 x 0.02 mm, length 110 mm, Inox | |
Stereoscopic zoom Microscope | Nikon | SMZ-2B | |
Cold light source | Schott | KL1500 LCD | |
Filter (Color) for cold light source | Schott | RG620 | |
Delicate wipers | Kimtech | Kimwipes | |
Electrode holder | Warner Instruments | QSW-T10P | Q series Holders compatible with Axon amplifiers, straight body style |
Silver Wire | Warner Instruments | 0.25 mm diameter, needs to be chloridized | |
Micromanipulator | Sutter Instruments | MP 85 | Huxley-Wall Style Micromanipulator |
Faraday cage | home made | Electromagnetic noise shielding and black front curtain | |
Anti-vibration Table | Newport | VW-3036-OPT-01 | |
Osilloscope | GW | GOS-622G | |
Perfusion system | Warner Instruments | VC-8P | Pinch valve control system |
Perfusion valve controller | Scientific instruments | BPS-8 | |
Suction system | |||
Amplifier | Molecular Device | Axopatch-1D | |
Head-stage | Molecular Device | CV – 4 | Gain: x 1/100 |
A/D converter | Molecular Device | Digidata 1440A | |
Clampex | Molecular Device | 10 | software |
pCLAMP | Molecular Device | 10 | software |
Light source (Xenon Arc lamp) | Sutter Instruments | Lambda LS | |
Light detector | home made | phototransistor | |
Filter wheel and shutter controller | Sutter Instruments | Lambda 10-2 with a Uniblitz shutter | |
Filters (Natural density filter) | Chroma | 6,5,4,3,2,1,0.5,0.3 | |
Filter (Color) | Schott | OG590, Edge filter | |
Xenon Flash Lamp system | Dr. Rapp OptoElecftronic | JML-C2 | |
Light guide | Quartz | ||
Pulse generator | AMPI | Master 8 | |
Microscope | Olympus | IX71, Inverted | |
Red illumination filter (Microscope) | RG630 / RG645 ø45mm | ||
Microscope objective | Olympus | X60/0.9 UplanFL N air or X60/1.25 UplanFI oil | |
CCD Camera | Andor | iXon DU885K | |
NIS Element | Nikon | AR | software |
Ca+2 indicator | Invitrogen | Calcium green 5N | |
Excitation & emission filters and dichroic mirror | Chroma | 19002 – AT – GFP/FITC Longpass set | |
Vertical pipette puller | Narishige | Model PP83 | Use either vertical or horizontal puller, as preferred. |
Horizontal pipette puller | Sutter Instrument | Model P-1000 Flaming/Brown Micropipette Puller | |
Filament | Sutter Instrument | 3 mm trough or square box | |
Capillaries | Harvard Apparatus | borosilicate glass capillaries | 1 x 0.58 mm |