Cold Spring Harb Protoc 2010 Zhang Pdb.prot5487

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doi: 10.1101/pdb.prot5487Cold Spring Harb Protoc;

Bing Zhang and Bryan Stewart

Larval Body-Wall MusclesDrosophilaElectrophysiological Recording from

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Electrophysiological Recording from DrosophilaLarval Body-Wall Muscles

Bing Zhang and Bryan Stewart

INTRODUCTION

The Drosophilalarval neuromuscular junction (NMJ) shares many structural and functional similaritiesto synapses in other animals, including humans. These include the basic feature of synaptic transmis-sion, as well as the molecular mechanisms regulating the synaptic vesicle cycle. Because of its largesize, easy accessibility, and the well-characterized genetics, the fly NMJ remains an excellent modelsystem for dissecting the cellular and molecular mechanisms of synaptic transmission. This protocoldescribes the steps for performing intracellular recording from fly larval body-wall muscles andexplains how to record and analyze spontaneous and evoked synaptic potentials. Methods usedinclude larval dissection (filleting), identification of muscle fibers and their innervating nerves,

the use of a micromanipulator and microelectrode in penetrating the muscle membrane, and nervestimulation to evoke synaptic potentials.

RELATED INFORMATION

Details of how to assemble an electrophysiology rig can be found in Equipment Setup for DrosophilaElectrophysiology (Zhang and Stewart 2010a). This protocol includes equipment used in the ColdSpring Harbor Laboratory (CSHL) course, Neurobiology of Drosophila, but equipment fromreputable commercial suppliers other than those mentioned here should work equally well. A method

for preparing microelectrodes is described in Fabrication of Microelectrodes, Suction Electrodes,and Focal Electrodes (Zhang and Stewart 2010b). Those who are unfamiliar with the operation ofthe amplifier and the software used for voltage-clamp experiments should see ElectrophysiologicalRecording from a Model Cell (Zhang and Stewart 2010c). Details for dissection of third-instarlarvae are found in Dissection of DrosophilaLarval Body-Wall Muscles (Ramachandran and Budnik2010). The Drosophila larval body wall muscle preparation was first used for electrophysiologicalanalysis in the 1970s (Jan and Jan 1976a,b). This preparation has become the gold standard forstudying neuronal excitability as well as synaptic transmission.

2010 Cold Spring Harbor Laboratory Press 1 Vol. 2010, Issue 9, September

Adapted from Drosophila Neurobiology(ed. Zhang et al.). CSHL Press,Cold Spring Harbor, NY, USA, 2010.

Cite as: Cold Spring Harb Protoc; 2010; doi:10.1101/pdb.prot5487 www.cshprotocols.org

Protocol

MATERIALS

CAUTIONS AND RECIPES: Please see Appendices for appropriate handling of materials marked with , andrecipes for reagents marked with .

Reagents

Drosophila melanogaster, wild-type strains (Canton-S or Oregon-R) and synaptic transmission

mutants (e.g., Syx3-69, snap-25ts, syt I, and lap), neuronal excitability mutants (e.g., eag, Sh

double mutant), or muscle excitability mutants (e.g., larvae that overexpress the inward

rectifier potassium channel Kir2.1 in body-wall muscles [referred to as the K

irmutant])

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METHOD

Dissecting Third-Instar Larvae

1. Dissect third-instar larvae as described in Dissection of Drosophila Larval Body-Wall Muscles(Ramachandran and Budnik 2010). Use either a Sylgard-coated Petri dish and small insect pins ora magnetic dish with paper clip pins. Perform the dissection in ice-cold, Ca 2+-free HL-3 saline(Stewart et al. 1994).

2. Using a pair of fine scissors, transect the segmental nerves loose from the base of the ventral nervecord so that the nerve end can be picked up with a suction electrode (Step 20).

3. Immediately following dissection, rinse the NMJ preparation at least three times with roomtemperature HL-3 saline containing the desired level of Ca2+.

The purpose of these washes is to ensure that the bathing solution is free of any material that could clog thesuction electrode. We recommend using 0.8 mM Ca2+ in the HL-3 saline.

4. Position the NMJ preparation:

i. Place the dish on the center of the microscope stage.

ii. Bring the preparation into focus and take note of the orientation, segments, and musclepatterns.

iii. Place the preparation so that the posterior end of the larva points toward you.

This orientation gives enough space on the two lateral sides so that the electrodes can access the NMJwithout bumping into the insect pins.

Recording Membrane Potentials

5. Backfill a microelectrode with 3 M KCl solution:

i. Fill the electrode approximately two-thirds full.

ii. Add one drop of the KCl solution at the end of the glass pipette so that the solution can bedrawn into the fine tip of the electrode through the built-in capillary (see Fabrication ofMicroelectrodes, Suction Electrodes, and Focal Electrodes [Zhang and Stewart 2010b]).

Maintain and raise all flies on cornmeal-based fly food at room temperature. Avoid crowding in the vials, so

that larvae can grow healthy and large.

HL-3 saline (variable Ca2+)

Also prepare HL-3 saline without Ca2+, ice cold.

KCl, 3 M

Equipment

Dissection instruments

Electrophysiology rig (as described in Equipment Setup for DrosophilaElectrophysiology

[Zhang and Stewart 2010a])

Microelectrodes, intracellular (as prepared in Fabrication of Microelectrodes, Suction

Electrodes, and Focal Electrodes [Zhang and Stewart 2010b])

Microscope

Petri dish (Sylgard-coated) and small insect pins

Alternatively, use a magnetic dish with paper clip pins (see Step 1).

Scissors (fine)

Suction electrode (as prepared in Fabrication of Microelectrodes, Suction Electrodes, and

Focal Electrodes [Zhang and Stewart 2010b])

Syringe, 1-mL or 5-mL

Tissue (e.g., Kimwipe)



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iii. Use a piece of soft tissue (e.g., Kimwipe) to remove any liquids outside of the electrodeso that salt crystals do not accumulate inside the electrode holder.

6. Position the microelectrode and microelectrode holder:

i. Carefully insert the microelectrode into the holder.

ii. Tighten the screw cap slightly so that the microelectrode is held firmly in place.

Do not screw the cap on too tightly or the glass electrode may break.

iii. Make sure that the silver wire from the electrode holder is in contact with the KCl solutioninside the electrode lumen.

iv. Screw the electrode holder onto the head stage.

7. Place the reference (ground) wire into the bath saline (HL-3 saline).

8. Adjust the focal plane above your preparation and lower the microelectrode into contact withthe HL-3 saline.

The first sign that the electrode has entered the saline is that the digital voltmeter begins to display a steadyvoltage.

See Troubleshooting.

9. Turn the Input offset knob in the ME1 panel until Vm

reads 0 mV. Lock the knob if necessary toavoid accidentally changing the setting.

10. Visually inspect the tip of the electrode for any obvious damage under the microscope. Replace itif the electrode is broken or contains an excessive amount of debris or dirt.

11. Check the input resistance of the electrode by turning the toggle switch to the on position onthe Step Command panel and manually injecting 1 nA of current.

A good electrode for intracellular recording usually has an input resistance ranging from 15 to 25 M. Recordthe electrode input resistance in a notebook to serve as a reference on which electrode to use in the future.

12. Perform bridge balance to cancel out the resistance of the electrode until the square wavedisappears (similar to those conducted in Fig. 1A-D). Use the Capacitance compensation knob(with a yellow top) to increase the rise time.

Avoid overcompensation (i.e., when the trace will be ringing with high-frequency oscillations).

13. Lower the focal plane first and then the electrode so that the tip of the electrode is always withinthe visual field. Repeat the focus down first and lower electrode next motion until the electrode

tip is just above the preparation.A common mistake that most novices make is to lower the electrode too fast and too far, with the result that theelectrode is driven through the preparation!

14. Once the electrode is close to the muscle surface, switch the manipulator to the fine controlmovement mode to position the microelectrode above a muscle fiber (e.g., muscle 6 in abdomi-nal segment 4). Carefully lower the electrode above the preparation while adjusting the focal planetoward the preparation.

15. Penetrate the muscle membrane slowly and gently. If you are using the Sutter P-250 motorizedmanipulator, switch to the diagonal mode so that the electrode moves diagonally toward themuscle, which makes membrane penetration easier.

Two hints that your electrode is touching or pressing on the muscle surface are that (1) the digital voltmeterbegins to show negative potentials up to 4 to 5 mV and (2) you may notice a dent on the muscle surfaceat the tip of the electrode. The potential readings are a more reliable and sensitive indicator, especially if it is

difficult to clear the electrode tip. At this point, a slight advance or vibration of the electrode is sufficient to enterthe cell. There should be a rapid voltage drop to at least 45 mV.

16. Wait for the membrane potential to become more hyperpolarized. It should stabilize at 65 to75 mV.

i. If the resting potential remains at -45 mV or is even more depolarized, withdraw theelectrode and try inserting it into the adjacent muscle or try a muscle cell in a differentsegment.




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ii. Continue this trial-and-error approach until a reasonably good resting potential is obtained.

Before inserting the electrode into another muscle cell, check its input resistance again, and replace theelectrode if its input resistance is dramatically altered.

See Troubleshooting.

17. Once a suitable and stable resting potential is obtained, cancel out the electrode resistance byadjusting the bridge balance again:

i. Inject a -1-nA current pulse for 400 msec duration at 1 Hz (Fig. 2A; Fig. 3A,B).

ii. Determine and record the muscle-input resistance (Rm) and the membrane time

constant . Record the Rm

and values in the notebook.

Fly muscleRm

varies between 5 and 12 M, and is ~30 msec.

iii. Try a new muscle fiber if the input resistance of the recorded muscle is


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Positioning the suction electrode

18. Immerse the stimulating wire in saline:

i. Lower the suction electrode just below the surface of the bath saline (HL-3 saline).

ii. Use a 1- or 5-mL syringe to gently apply negative pressure (suction) so that the HL-3 salinefills one-half of the length of the suction electrode.

This ensures that the stimulating silver wire inside the electrode is deeply immersed in the saline. Avoid usinga larger (e.g., 10-mL) syringe because the suction it generates is too strong and difficult to control.


FIGURE 2. B* '' ;9 9. " >'9 9= ' ';' ** ='


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24. Design a stimulation paradigm that can deliver two pulses separated by 100 msec (10 Hz),50 msec (20 Hz), 20 msec (50 Hz), and 10 msec (100 Hz), respectively. Observe the relativeamplitude of two evoked EJPs in HL-3 saline containing 0.6 mM Ca2+.

This is a classic exercise that is often referred to as twin-pulse or two-pulse facilitation (TPF).

25. Repeat the above experiments in NMJ preparations based in HL-3 saline containing 1, 1.5, and2 mM Ca2+, respectively.

Examples of short-term synaptic plasticity at 30 Hz at 0.6 mM Ca2+ and 1.5 mM Ca2+ are shown in Figure 4Aand Figure 4B, respectively.

ELECTRODE CLEANUP

26. At the end of the day, rinse the suction electrode thoroughly with H2O and leave the electrode

to air-dry to prevent bacterial growth. When rinsing the suction electrode, always suck H2O in

and do not push the liquid out, because small bits of debris may clog the tip of the electrode,rendering it useless.

A suction electrode, properly cared for, can be used repeatedly for as long as it works (6 mo or longer).

27. Discard the intracellular microelectrode, and turn off the electronics.

DATA ANALYSIS

28. Using Clampfit, analyze the input resistance and time constant of the muscle membrane, and theamplitude of EJPs.

29. Use Mini Analysis to analyze the mini frequency and amplitude.

30. Estimate quantal content of a synapse by calculating the ratio of the average EJP amplitude andthe average mini amplitude, provided that the resting potential of different recorded muscles issimilar and that the [Ca2+] is relatively low.

Otherwise, nonlinear summation should be taken into consideration in determining quantal content (Martin1976; Stevens 1976; Feeney et al. 1998).

TROUBLESHOOTING

Problem: No voltage can be detected when the microelectrode is placed in the HL-3 saline bathsolution.[Step 8]

Solution: There may be an open circuit. Consider the following:

1. Look for broken wires, and replace any that are found.

2. Confirm that the ground wire is in the bath.


FIGURE 4. !- 9?' '9? ' '* ;, ','9, ** '; >';' %C'2+& '*


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3. Look for a large air bubble inside the electrode tip that got there when the microelectrode wasbackfilled.

4. Determine whether the silver wire inside the electrode holder is contacting the golden pin.

Problem: Resting potentials of the impaled muscle cell never hyperpolarize beyond 35 to 45 mV.[Step 16]

Solution:Wild-type larval body-wall muscles have a resting potential from 65 to 75 mV. A poor rest-ing potential usually means that the muscle has been damaged during dissection or penetration

with the microelectrode. Consider the following:

1. Healthy muscles have a translucent appearance, whereas degenerating muscles are brownish orvisibly tearing apart from the muscle attachment points in the denticle belt area. This tends tooccur if the preparation is left at warm temperatures. Keep and work with muscle preps in a coolerroom or cooled microscope stage (18C to 20C).

2. Physical tearing or damage to muscles should be avoided at all costs during dissection or electrodeimpalement. However, electrode-caused damage is difficult to detect by eye. One way to detectany damage is to measure the muscle input resistance and time constant. Both of these parame-ters will reduce dramatically in a damaged muscle. Thus, it is always a good idea to routinely mon-itor muscle input resistance and time constants in all of your experiments.

Problem: Although the resting potentials of the muscle and minis were normal, no evoked synapticpotentials were detected when the segmental nerve was stimulated.

[Step 22]

Solution: Consider the following when evoking synaptic potentials:

1. The stimulator must be turned on and the voltage output level must be high enough to evokeaction potentials. Gradually increase the amplitude of the stimulus strength until an evoked poten-tial is seen.

2. Check to see if the polarity of the stimulus was reversed. Switch the polarity of the stimulus isola-tor and test again.

3. Determine if the suction electrode contains enough HL-3 saline so that the stimulating wire is incontact with the saline. (If the wire is not in contact with the saline, then no current can passthrough the suction electrode.) Gently push the nerve ending off the suction electrode, raise thesuction electrode away and above the NMJ, and refill it with HL-3 saline. It is important to raisethe suction electrode so that no nerves are picked up during refilling. Lower the electrode and pickup the nerve ending again.

4. Inspect the nerve for damage that may have occurred during dissection. If the forceps havepinched the nerve, you will not get normal EJPs. Start over with a new dissection.

5. If too much pressure is applied to the suction electrode when picking up the nerve, it can causedisruption of the synaptic contact. Switch the suction electrode to a different segment and try togently pick up the nerve.

6. Verify that the HL-3 saline contains Ca2+, which is essential to trigger exocytosis. Prepare HL-3saline containing Ca2+ and repeat the experiment.

Problem: Instead of an EJP with peak amplitude of 25 mV, an odd-looking EJP was obtained havingmuch smaller amplitude and unusually slow rising and falling rates.

[Step 22]

Solution: Most likely the motor nerve from a neighboring segment was stimulated. Larval body-wallmuscles are electrically coupled by gap junctions with homologous muscles in the neighboring seg-ments (Gho 1994). EJPs as well as minis can passively invade the neighboring muscles resulting inEJPs or minis with much slower kinetics. The minis with slow rising and falling phases should beexcluded in determining mini frequency and amplitude.



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ACKNOWLEDGMENTS

We thank the members of our laboratories for their important contributions to our research programs.B.Z. especially thanks Hong Bao for her expert assistance with the Neurobiology of Drosophilacourse and for providing some of the figures in the chapter from which this article was adapted.Research in the Zhang laboratory was supported by grants from the National Science Foundation(NSF), National Institutes of Health (NIH), Oklahoma Center for the Advancement of Science andTechnology (OCAST), and research funds from the University of Oklahoma. B.Z. also thanks Phillip

Vanlandingham, Rudhof Bohm, and Hong Bao for their constructive comments. B.S.s research was

supported by a Canada Research Chair and grants from the Canadian Institute for Health Research,the Natural Science and Engineering Research Council of Canada, and the Canadian Foundation

for Innovation. An extensive citation of all publications related to the topics covered in this protocolwas not possible because of space limitations. We therefore focused on select initial publications andapologize to our colleagues whose publications were not cited.

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