K.B. is supported by NIH grant GM54544.

Part 1: Equipment and Supplies
<ol>
<li>Electrophysiological recording from Drosophila embryos first requires proficiency in embryonic dissection techniques, which are described in another JoVE video.</li>
<li>Electrophysiological recording from Drosophila embryos utilizes standard patch clamp recording configurations. Patch clamp recording equipment and software suitable for many other preparations is also suitable for recording from Drosophila embryos. Because Drosophila embryos are glued to a cover slip during dissection, the recording chamber should accept cover slips. The thickness of the preparation and manner in which it is mounted to the coverslip requires an upright microscope with excellent DIC optics and a long working distance lens.</li>
<li>Two types of bath solutions are used for electrophysiological recording; 1) &ldquo;standard&rdquo; (Jan and Jan, 1976a) or &ldquo;modified standard&rdquo; (Broadie, 2000) salines, based on solutions commonly used for recording in other invertebrate systems, and 2) &ldquo;haemolymph-like&rdquo; (HL) salines (Stewart et al 1994), a compromise between the standard saline and the ionic concentrations measured in the Drosophila haemolymph. It should be noted that none of these salines accurately mimic the ionic concentrations measured in the haemolymph, which are unfavorable for recording (Broadie, 2000). Standard saline contains (in mM): 135 NaCl, 5 KCl, 4 MgCl2, 1.8 CaCl2, 72 sucrose, 5 TES, pH 7.2.</li>
</ol>
Part 2: Recording Configuration
<ol>
<li>For physiological experiments, the dissected embryo preparation is placed in a small plexiglass recording chamber (&lt;0.5 ml) and viewed in transmitted light with an upright compound microscope fitted with differential interference contrast (Nomarski) optics and a long-working distance 40-100X (typically 40X) water-immersion objective (Broadie and Bate, 1993a). Recordings are typically made below room temperature (16-22&deg;C), with increasing difficulty at elevated temperatures (22+&deg;C). We achieve this by cooling the room rather than cooling the preapartion.</li>
<li>Patch pipettes can be pulled from a variety of fiber-filled glasses (e.g. borosilicate glass, 1 mm outer diameter) and the tips should be fire-polished to final resistances of 5-10 M&Omega;. A chloride-silver ground wire is placed in the bath. Signals are amplified using a patch-clamp amplifier (e.g. Axopatch-1D, Axon Instruments), filtered with an 8-pole Bessel filter at 2-10 Hz, and either sampled on-line or stored digitally for later analysis.</li>
<li>Conventional patch-clamp recordings are most often made at 18&deg;C with patch pipettes pulled from fiber-filled borosilicate glass (tips heat polished to ~1 &micro;m i.d.). Current records are made from identified multinucleate muscle fibers in voltage-clamp (standard holding potentials &ndash;60 to &ndash;80 mV) (Broadie and Bate, 1993a-d; Broadie et al., 1997; Fergestad et al., 1999; Featherstone et al., 2000). The patch pipette solution contains (in mM): 120 KCl, 20 KOH, 4 MgCl2, 0.25 CaCl2, 5 EGTA, 4 Na2ATP, 36 Sucrose, and 5 TES, buffered at pH 7.2. Recording electrodes are back-filled.</li>
</ol>
Part 3: Muscle Recording
<ol>
<li>In principle, any embryonic muscle may be recorded from in this preparation. In practice, most recording has concentrated on the large, ventral longitudinal muscles in the inner-most muscle layer (Bate, 1990; Broadie and Bate, 1993a-c). Records have also been made from the dorsal and lateral muscles of this muscle layer (Kidokoro and Nishikawa, 1994; Auld et al. 1995; Nishikawa and Kidokoro, 1995; Baumgartner at al. 1996). However, most experiments focus on the group of four longitudinal, ventral muscles (6,7,12,13), particularly muscle 6 in the anterior abdomen (A2-A3).</li>
<li>No enzymatic treatment is required prior to muscle recording in young embryos (&lt;16 hrs AEL). However, a muscle sheath develops in the later embryo (&gt;16 hrs AEL) and must be removed with collagenase (collagenase type IV, 1 mg/ml in divalent cation-free saline, 0.5-2 min at RT) prior to patch recording. Following enzyme treatment, seal resistances on the muscle are typically &gt;1 G&Omega;.</li>
<li>Patch-clamp recordings can be made from the embryonic muscles using a number of standard techniques (Broadie and Bate 1993a-c; Broadie et al. 1994, 1995, 1997; Nishikawa and Kidokoro, 1995). Any standard patch-clamp variations -- cell-attached, inside-out, outside-out -- can be employed to record single channel activity in the muscle or isolated muscle membrane patches. Single glutamate receptor channel activity (~12 pA at -60 mV) are generally observable on the falling phase of spontaneous excitatory junction currents (sEJCs) recorded in whole cell mode.</li>
<li>Most recording is done in whole-cell recording configuration, achieved easily from the cell-attached state with slight suction or an electrical &ldquo;buzz&rdquo;. The muscles can be recorded from either in voltage-clamp (typical holding potentials of -60 to -80 mV) or current-clamp configurations. Muscles in younger embryos (&lt;13 hrs AEL) cannot be effectively voltage-clamped owing to extensive coupling between the embryonic myotubes (Broadie and Bate, 1993a); this is usually not a problem during later (14+ hrs AEL) developmental stages (Ueda and Kidokoro, 1996).</li>
<li>Nystatin-perforated patch-clamp recording can be used to verify records obtained with standard whole-cell techniques (Broadie and Bate, 1993a). The perforation solution is made as follows: 10 mg pluronic F-127 (Molecular Probes p-1572) is dissolved in 20 ml DMSO, and 10 mg Nystatin dissolved in this solution to make the stock. 100 &mu;l of the stock is added to 5 ml filtered pipette solution to make the recording solution. Whole-cell currents are usually obtained &lt;3 mins after seal formation. Series resistance is markedly increased in most instances and cells can develop an increased leakage current (Broadie and Bate, 1993a).</li>
<li>The whole-cell voltage-gated current in mature embryonic (and larval) muscle is composed of five prominent components (Wu and Haugland, 1985; Zagotta et al. 1988; Broadie and Bate, 1993b): an inward calcium current (ICa) and four outward potassium currents (Tsunoda and Salkoff, 1995), including two fast, inactivating currents, voltage-gated (IA) and calcium-dependent (ICF), and two delayed, non-inactivating currents, voltage-gated (IK) and calcium-dependent (ICS). Ion-substitution experiments can be used to dissect these currents from the whole-cell response during voltage-clamp (Broadie and Bate, 1993b).</li>
</ol>
Part 4: Neuromuscular Junction Stimulation and Recording
<ol>
<li>The most common and reliable method of NMJ stimulation uses a glass suction electrode on the segmental peripheral nerve. Suction electrodes can be pulled from a variety of fiber-filled electrode glass (1-1.5 &mu;m outer diameter) and should be fire-polished to achieve the desired configuration (~5 &mu;m inner diameter). A small segment (&lt;50 &mu;m) of the appropriate segmental motor nerve is drawn into the pipette with gentle suction to form a tight seal.</li>
<li>It is common to stimulate the intact nerve by drawing in a loop of the nerve near the CNS exit point. This approach allows easy stimulation of the intact preparation, but has the disadvantage that spontaneous CNS-evoked activity can occur superimposed on the applied stimulation paradigm. An alternative is to stimulate the cut motor nerve. This can be achieved if the CNS is removed, although this procedure tends to stretch the nerve and may cause damage. The nerve may also be cut with a sharp glass microelectrode, but this procedure is tedious and difficult. It is recommended that records be made from the intact preparation.</li>
<li>Stimulation can be applied with a variety of paradigms. Brief stimulation (0.2-1 msec) works best and stimulation intensity (usually 1-5 V) will depend on the suction configuration and must be experimentally determined for each cell. Suprathreshold stimulation of the motor nerve is best achieved by setting the stimulus strength slightly above (+1-2 volts) threshold. Evoked NMJ responses are recorded from the patch-clamped muscle as described above. The shock artifact can be substantially decreased with the use of an isolated virtual ground.</li>
<li>A less reliable alternative to the standard suction electrode nerve stimulation is direct stimulation of the central nervous system (Nishikawa and Kidokoro, 1995). A microelectrode filled with 3-4 M KCl (or KAc) is inserted into the middle of the ventral ganglion and positive pulses of ~2 microamp in intensity and ~2 msec in duration are delivered (Dietcher et al. 1998). The electrode tip should be in the neuropil nearest the hemisegment where stimulation is desired. Synaptic transmission is recorded in the patch-clamped muscle in the standard configuration.</li>
<li>The embryonic NMJ is accessible to whole-cell patch-clamp recording throughout its differentiation. For excitatory junctional current (EJC) recordings, signals are recorded with an standard patch-clamp amplifier, filtered (usually at 1-2 KHz), converted to a digital signal using an analog-to-digital interface, and acquired with a computer using appropriate software.</li>
</ol>
Part 5: Neuromuscular Junction Chemical and Drug Applications
<ol>
<li>Any small, charged molecule can be effectively iontophoresed onto the embryonic NMJ (Aravamudan et al, 1999; Fergestad et al. 1999, 2001). Iontophoretic pipettes can be pulled for specific configurations, with applications for variable periods, using pulses of negative or positive current. It is vital to prevent leakage between pulses with an appropriate opposing backing current (Broadie et al. 1993d; Featherstone et al. 2002, 2005; Rohrbough et al. 2007). The magnitude of this backing current will vary with pipette configuration and should be experimentally determined.</li>
<li>Many experiments have been done with the NMJ neurotransmitter, L-glutamate (Jan and Jan, 1976b), using a 100mM stock solution (Monosodium salt; pH 8-9)(Broadie and Bate, 1993c). High resistance iontophoretic pipettes (100-200 M&Omega;) have been used for glutamate receptor (gluR) mapping (Broadie and Bate, 1993d), and lower resistance pipettes (20-50 M&Omega;) used to estimate the total gluR response (Broadie et al. 1995, 1997; Featherstone et al. 2002, 2005; Rohrbough et al. 2007). Short pulses (0.1-1 ms) of negative current (~10 nA) work well, with a small, positive backing current to prevent leakage.</li>
<li>Non-charged and charged molecules can be applied by pressure ejection (Fergestad and Broadie, 2001; Featherstone et al. 2001). For application, a pressure ejection pipette (&lt;5 &mu;m inner diameter) containing the solution is positioned close (&lt;10 &mu;m) from the nerve terminal. The solution to be ejected can be applied using a picospritzer (5-10 psi) or other pressurized air source (Featherstone et al., 2002). During prolonged experiments, the recording chamber (volume &lt;0.5 ml) should be perfused continuously (0.1 - 0.2 ml/sec) with normal bath saline.</li>
<li>Many experiments have been done with hyperosmotic saline to trigger fusion of synaptic vesicles (Broadie et al. 1995; Aravamudan, et al., 1999; Fergestad et al., 1999; Featherstone, et al., 2000, 2002). Other experiments have been done with toxins, such as argiotoxin 636 against glutamate receptors and black widow spider venom containing latroinsectotoxin (Broadie et al. 1995).</li>
</ol>
Part 6: Complementary Neuromuscular Junction Vital Imaging Techniques
<ol>
<li>Synaptic vesicle kinetics may be monitored with fluorescent lipophilic stryl dyes (Fergestad and Broadie, 2001; Rohrbough et al. 2004). The distribution of vesicles can be directly observed, and rates of endocytosis and exocytosis measured. Several variants have been made with different fluorescent properties (FM1-43, fluorescein-like; FM4-64, rhodamine-like). Modifications of the original dye have yielded products with altered hydrophobic properties, and therefore with different washout kinetics; e.g., FM2-10 dissociates much faster from the membrane.</li>
<li>To label the embryonic NMJ with stryl dyes, 10 &micro;M FM1-43 can be loaded for 1-5 minutes using elevated extracellular potassium (50-90 mM) (Fergestad and Broadie, 2001). A more physiological approach may be achieved via stimulation by a nerve suction electrode. Preparations are then washed repeatedly in Ca2+-free buffer for 5 minutes, and retained fluorescence (representing dyed synaptic vesicles) measured. To measure exocytosis, one can unload the dye using either elevated potassium or nerve electrical stimulation.</li>
<li>Fusion proteins with GFP variants have been produced allowing multicolor labeling of the embryonic NMJ synapse (Brand, 1999; Zhang et al. 2002). Many transgenics expressing GFP-tagged proteins are available including general membrane markers (e.g. mCD8-GFP), postsynaptic markers (e.g. Shaker-GFP, GluRIIA-GFP), cytoskeletal (e.g. actin-GFP, moesin-GFP, tubulin-GFP) and mitochondrial markers (e.g. Cytochrome C-GFP) and synaptic vesicle markers (e.g. synaptobrevin-GFP, synaptotagmin-GFP; Zhang et al., 2002).</li>
<li>GFP reporters label compartments that are highly dynamic and can be used to assess developmental features or mutant phenotypes (Featherstone and Broadie, 2000). Bleaching and then observing the recovery of the fluorescence (FRAP) has been shown to be feasible, at least at the larval Drosophila NMJ (Zhang et al., 2002; Yan and Broadie, 2007). A word of caution, however: some of these markers do not fully rescue null mutants and may even alter normal NMJ physiology.</li>
</ol>
Part 7: Recording from Central Motor Neurons
<ol>
<li>Many embryonic neurons can be individually identified based on soma position and projection morphology (Bate and Martinez-Arias, 1993; Goodman and Doe, 1993; Landgraf et al 1997). Further, the GAL4/UAS system (Brand and Perrimon, 1993) can target GFP expression to visualize identifiable neuron populations (Baines et al., 1998). Recording studies to date have concentrated on a group of five superficial, dorsal neurons within the embryonic ventral nerve cord (VNC); four motor neurons (aCC and RP1-4) and one interneuron pCC (Baines et al., 1998, 1999, 2001; Rohrbough and Broadie, 2002; Featherstone et al. 1995).</li>
<li>Cell bodies of identified neurons are accessible to whole-cell patch-clamp recording electrodes. The VNC neurolemma sheath must first be removed with brief focal treatment with protease (Baines and Bate, 1998; Baines et al., 1999; Rohrbough and Broadie, 2002). Dorsal neuronal cell bodies are exposed using a large diameter (~20 &micro;m) patch pipette containing 0.5-1% protease (type XIV (Sigma) in recording saline). The CNS sheath material is drawn by suction into the pipette tip for 1-2 minutes until the sheath ruptures (Featherstone et al. 1995). To confirm cellular identity during recordings, lucifer yellow (dipotassium salt, 1-2 mg/ml) can be added to the patch solution in order to visualize the position and morphology of the cell.</li>
<li>Standard whole-cell voltage- and current-clamp recordings can be made (Rohrbough and Broadie, 2002; Featherstone et al., 2005). The external recording saline contains (in mM): 140 NaCl, 3 KCl, 2 CaCl2, 4 MgCl2, 5 HEPES, 10 sucrose, 2 NaOH (pH 7.2). The patch solution contains (in mM): 140 K-Acetate, 2 MgCl2, 0.1 CaCl2, 10 HEPES, 1.1 EGTA, 2 Na2-ATP, ~6 KOH (pH 7.2). Voltage-clamp recordings are made at holding potentials of -60 mV at 18 &deg;C. Typical unadjusted resting potentials are in the range of -45 to &ndash;65 mV (-52.9 &plusmn; 7.5 mV, mean &plusmn; S.D., n = 78).</li>
<li>Available assays reveal voltage-gated ionic currents (Baines and Bate, 1998), repetitive action potentials, synaptically-mediated activity and agonist-gated responses (Baines et al., 2001). Synaptic input to motor neurons is recorded in the form of both fast AP-mediated fast excitatory synaptic currents (ESCs), and periodic, sustained (up to 1 sec duration) episodes of excitatory input occurring several times or more per minute (Baines et al., 1999; Baines et al., 2001).</li>
<li>Cell soma respond to iontophoretically applied acetylcholine (ACh) and the inhibitory transmitter GABA (Baines and Bate, 1998; Featherstone et al. 2005). ACh can be applied iontophoretically to the neuronal soma via a sharp microelectrode containing 100mM ACh in dH20, at pH 4-5 to favor agonist iontophoresis by positive current.</li>
</ol>
Representative Results
Ventral longitudinal muscle 6 is segments A2-A3 is the most commonly utilized target for recording (Broadie and Bate, 1993a-d; Harrison et al. 1994; Sweeney at al. 1995; Renden et al. 2001; Huang et al. 2006; Mohrmann et al. 2008). In the mature embryo and newly-hatched first instar larva (20-21 hrs AEL at 25&deg;C = hatching), muscle 6 whole cell capacitance is typically 25-30 pF, and input resistances 20-40 M&Omega;, changing with developmental age. Maximal NMJ synaptic currents range from tens of picoamps (13-14 hrs AEL) to 1-2nA (20-21 hrs AEL), rapidly increasing as a function of age. Direct measure of glutamate-gated currents yield maximal responses of hundreds of picoamps (13-14 hrs AEL) to 1.5-4nA just prior to hatching. Series resistance errors (total current X series resistance) are usually &lt;5 mV but, in mature, pre-hatching embryos, may be as high as 10+ mV and thus series resistance compensation is advised. Embryonic muscles with an average diameter of 10-30 &mu;m, and average length of 40-80 &mu;m, appear to show nearly ideal space clamp parameters. Cell clamp time constants (Rseries X Ccell) average less than 0.25 msec.
In the mature embryo and newly-hatched larva, the dorsal motor neuron population has been subject to more limited recording. Neuronal soma recordings reveal whole cell capacitance in the range of 1.8-2.5 pF (2.0 pF average), series resistance of 40-60 M&Omega; and input resistance of ~5 G&Omega; (Baines and Bate, 1998; Featherstone et al 2005). These cells clearly grow in size and complexity during embryonic maturation, and this growth continues throughout larval development. In third instars, whole cell capacitance is 17-20 pF, series resistance is 25-37 M&Omega; and input resistance averages ~900 M&Omega; (Rohrbough and Broadie, 2002; Richard Baines, personal communication). Ionic conductances increase throughout development, but current density (pA/pF) remains remarkably constant. In mature embryos, synaptic currents average 75-100pA in amplitude, with both rapid ESC events and prolonged currents averaging ~500 msec (Baines and Bate, 1998; Featherstone et al 2005). In embryos, these sustained events average 3-5 per min, increasing to ~20 per min in the first instar (Richard Baines, personal communication). Note that the morphological complexity and membrane properties of Drosophila neurons makes it virtually impossible to properly voltage clamp much of the cell beyond the soma. One should therefore interpret recordings with care.

The authors have nothing to disclose.

Electrophysiological recording from Drosophila embryos requires manual manipulation and dissection. The health of the preparation, and consequent quality of the recordings, depends on one being able to quickly and neatly prepare the fragile embryonic tissues for recording, and then execute the experiment. Experimenters should be proficient in both embryonic dissections and patch clamp electrophysiology before trying to tackle both at once.
Recordings can be done in one of several var...

electrophysiological recording in the drosophila embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Watch this Scientific Journal Video about Electrophysiological Recording in the Drosophila Embryo at JoVE.com

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

electrophysiological-recording-in-the-drosophila-embryo

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11.4K Views.  University of Illinois. Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

Video: Electrophysiological Recording in the Drosophila Embryo

Assay for Neural Induction in the Chick Embryo

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying the formation and initial patterning of the nervous system. This video demonstrates how to graft organizer tissue into a host, a method by which Hensen s node (the organizer in the chick embryo) is grafted to a host competent ectoderm. The organizer graft instructs overlying na ve tissue to adopt a neural fate via neural inducing signals. This mechanism is referred to as neural induction, and constitutes the initial step in the formation of brain and spinal cord in amniotes. This method is essentially used for the characterization of putative neural inducing molecules in chick. This video demonstrates the different steps in the assay for neural induction; First, the donnor embryo is explanted and pinned on a dish. Then, the host embryo is prepared for New culture. The graft is excised and transplanted to the host area pellucida margin. The host is cultured for 18-22 hrs. The assembly is fixed and processed for further applications (e.g. in situ hybridization). This method was originally devised by Waddington 1,2 and Gallera 3,4.

Neural induction is the first step in the formation of the brain. It is a mechanism by which Hensen's node (organizer), instructs adjacent tissue to adopt a neural fate, i.e. to give rise to the nervous system. This video demonstrates an assay for neural induction in chick embryo.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP&#8217;s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP&#8217;s and minature excitatory junction potentials (mEJP&#8217;s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2 (HL3) that contains 5 mM Mg2+ and 1.5 mM Ca2+. Changes in the amplitude of evoked EJP&#8217;s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.

Here we describe electrophysiological methods for measuring synaptic transmission at the neuromuscular junction of Drosophila larva. Evoked release is initiated artificially by stimulating the motor neuron axons, and transmission through the NMJ can be measured by the postsynaptic response evoked in the muscle.

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. ...

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

This technique exposes the Drosophila embryonic neuromusculature for immunohistochemistry or electrophysiological recording. It is useful for studying early events in neuromuscular development or performing electrophysiology in mutants that cannot hatch.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

Visualizing the Beating Heart in Drosophila

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in myogenic hearts. A key step in examining heart function in the fly is finding a way to access the heart in a manner that preserves its myogenic function while still allowing the beating heart organ to be observed and recorded. Two different methods for observing and recording the beating heart in both larva and adult Drosophila are described here. Our semi-intact preparation using adult flies allows clear visualization of the abdominal heart without interference from the pigmented cuticle and overlying fat bodies. To record larval heart beats it is necessary to immobilize the larva, which minimizes body wall movements thereby reducing heart movements that are not associated with myocardial contractions. Our methodologies produce stable adult and larval heart preparations that can beat for hours at rates of 1-3 Hz.

Visualizing the Beating Heart in Drosophila

Technique required for visualizing the beating heart in larval and adult Drosophila are presented. Each life stage requires a different methodology.

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Ambulatory ECG Recording in Mice

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although the surface ECG is utilized for short-term measurements of waveform intervals, it is not practical for long-term studies of heart rate variability or the capture of rare episodes of arrhythmias. Implantable ECG telemeters offer the advantages of simple surgical implantation, long-term recording of electrograms in ambulatory mice, and scalability with simultaneous recordings of multiple animals. Here, we present a step-by-step guide to the implantation of telemeters for ambulatory ECG recording in mice. Careful adherence to aseptic technique is required for favorable survival results with the possibility of implantation and recording from weeks to months. Thus, implantable ECG telemetry is a valuable tool for detection of critical information on cardiac electrophysiology in ambulatory animal models such as the mouse. 

Telemetric ECG has emerged as an essential tool in evaluating animal models for cardiac arrhythmias and sudden cardiac death. Here, we present a stepwise guide to telemetric ECG recordings for application in long-term ambulatory ECG monitoring in mice.

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although ...

Optimized Transfection Strategy for Expression and Electrophysiological Recording of Recombinant Voltage-Gated Ion Channels in HEK-293T Cells

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) is a ubiquitous research strategy. HEK-293T cells must be plated onto glass coverslips at low enough density so that they are not in contact with each other in order to allow for electrophysiological recording without confounding effects due to contact with adjacent cells. Transfected channels must also express with high efficiency at the plasma membrane for whole-cell patch clamp recording of detectable currents above noise levels. Heterologous ion channels often require long incubation periods at 28&deg;C after transfection in order to achieve adequate membrane expression, but there are increasing losses of cell-coverslip adhesion and membrane stability at this temperature. To circumvent this problem, we developed an optimized strategy to transfect and plate HEK-293T cells. This method requires that cells be transfected at a relatively high confluency, and incubated at 28&deg;C for varying incubation periods post-transfection to allow for adequate ion channel protein expression. Transfected cells are then plated onto glass coverslips and incubated at 37&deg;C for several hours, which allows for rigid cell attachment to the coverslips and membrane restabilization. Cells can be recorded shortly after plating, or can be transferred to 28&deg;C for further incubation. We find that the initial incubation at 28&deg;C, after transfection but before plating, is key for the efficient expression of heterologous ion channels that normally do not express well at the plasma membrane. Positively transfected, cultured cells are identified by co-expressed eGFP or eGFP expressed from a bicistronic vector (e.g. pIRES2-EGFP) containing the recombinant ion channel cDNA just upstream of an internal ribosome entry site and an eGFP coding sequence. Whole-cell patch clamp recording requires specialized equipment, plus the crafting of polished recording electrodes and L-shaped ground electrodes from borosilicate glass. Drug delivery to study the pharmacology of ion channels can be achieved by directly micropipetting drugs into the recording dish, or by using microperfusion or gravity flow systems that produce uninterrupted streams of drug solution over recorded cells.

Reliable method for highly efficient in vitro expression and subsequent electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T).

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) ...

Tracking Morphogenetic Tissue Deformations in the Early Chick Embryo

Embryonic epithelia undergo complex deformations (e.g. bending, twisting, folding, and stretching) to form the primitive organs of the early embryo. Tracking fiducial markers on the surfaces of these cellular sheets is a well-established method for estimating morphogenetic quantities such as growth, contraction, and shear. However, not all surface labeling techniques are readily adaptable to conventional imaging modalities and possess different advantages and limitations. Here, we describe two labeling methods and illustrate the utility of each technique. In the first method, hundreds of fluorescent labels are applied simultaneously to the embryo using magnetic iron particles. These labels are then used to quantity 2-D tissue deformations during morphogenesis. In the second method, polystyrene microspheres are used as contrast agents in non-invasive optical coherence tomography (OCT) imaging to track 3-D tissue deformations. These techniques have been successfully implemented in our lab to studythe physical mechanisms of early head fold, heart, and brain development, and should be adaptable to a wide range morphogenetic processes.

This article describes surface labeling and ex ovo tissue culture in the early chick embryo. Techniques amenable to time-lapse bright field, fluorescence, and optical coherence tomography imaging are presented. Tracking surface labels with high spatiotemporal resolution enables kinematic quantities such as morphogenetic strains (deformations) to be calculated in both two and three dimensions.

Embryonic epithelia undergo complex deformations (e.g. bending, twisting, folding, and stretching) to form the primitive organs of the early embryo. ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...