The overall goals of the following experiments are to understand the properties of excitable membranes and the ionic basis of the resting membrane potential, and to learn methods to measure the membrane potential as well as to demonstrate properties of synaptic transmission. In one part of the experiment, the resting membrane potential will be measured while altering extracellular potassium. In another part of the experiment, mechanisms of synaptic differentiation will be investigated by recording, stimulated synaptic responses in various motor units.
Next, a neural circuit is presented that is easy to maintain. This preparation can be used for teaching as well as researching various aspects of a sensory to CNS to motor neuron to muscle circuit. This set of experiments show the ease of using the crayfish model to address issues relating to membrane potential synaptic integration and transmission, and structural differences of muscle fiber types.
The main advantages of using the crayfish preparation in teaching electrophysiological concepts and techniques are the ease of dissection, the preparation's viability of minimal saline, the ease of obtaining synaptic responses, and the ease of discerning the overall anatomical arrangement of the muscles. These techniques can also be applied to other preparations and will further our understanding of synaptic physiology and modulation and biochemical and structural differences in the demonstrated preparation. To begin the surgery, select a crayfish approximately six to 10 centimeters in body length and place it in crushed ice for five minutes to anesthetize it to decapitate the crayfish, hold it from the back of the head, making sure to keep your fingers out of reach of the cray fish's claws and mouth using large scissors.
Quickly remove the head by making a clean cut from behind the eyes of the crayfish, remove the legs and claws of the crayfish to avoid injury. Remove the styles on males and the swimmer ettes on both males and females. Dispose of the head and appendages.
Next, separate the abdomen from the thorax by making a cut along the articulating membrane, which joins the abdomen and thorax. Save the abdominal portion of the crayfish and dispose of the thorax, pointing the scissors slightly down towards the ventral side, and at an angle, make a cut in the shell along the lower lateral border of each side of the abdomen. Be careful not to cut too deeply into the crayfish.
Follow the natural shell pattern of lines of the crayfish that run the length of each segment Using forceps. Remove the ventral portion of the shell by pulling it up and back. Taking care not to destroy the abdominal muscles.
Remove the flexor muscles. At this point, a white mass of tissue can be seen on top of the deep flexor muscles. Remove this tissue carefully with forceps.
The GI tract is now visible as a small tube running along the midline of the deep flexor muscles. Remove it by pinching it with forceps at the top and pulling it away from the abdomen. Cut the bottom of the GI tract at the end of the tail.
Rinse the dissection with saline to ensure that fecal waste does not interfere with the preparation. Using dissection pins pin the top and bottom corners of the preparation to a S guard coated Petri dish. The abdominal extensor muscles are now exposed.
Pour saline solution into the Petri dish to cover the preparation until intracellular recordings are performed. Move the Petri dish to the microscope and use wax underneath the dish to prevent it from moving. The apparatus used for intracellular recordings is shown in this figure.
The appropriate connections and settings are given in the accompanying text. Set up the lab chart software according to the instructions. In the accompanying text, place the ground wire into the saline bath.
Place the tip of a glass micro pipette filled with three molar potassium chloride into the saline bath and test the electrode resistance. The electrode resistance should be 20 to 60 mega ohms. Use the coarse knob on the amplifier to move the line on the lab chart to zero under high intensity illumination.
Look through the microscope and identify the longitudinal DEM muscles. Use the electrode probe to insert the intracellular electrode into a muscle fiber of the DEM muscle. The electrode should barely be inserted into the muscle fiber.
Be careful not to penetrate all the way through the cell. Measure the amplitude of the resting potential. Drag the M cursor onto the trace from the location on the lower left hand corner of the chart window.
Place the marker M on the baseline and move the cursor to the lowest point on the recorded signal. The difference between the marker and the active cursor is displayed on the upper right side of the screen. This value is the voltage of the resting membrane potential.
Divide the recorded voltage by the amount of amplification in this case 10 x amplification was used. Then convert the voltage from volts to millivolts. After recording the resting potential for that muscle fiber, look through the microscope and carefully remove the electrode from the muscle fiber.
Repeat the intracellular recording on other muscle fibers. The next part of this experiment will monitor the effect of raising the extracellular potassium concentration on the resting membrane. Potential six crayfish saline solutions will be used with potassium concentrations of 5.4 20, 40, 60, 80, and 100 millimolar.
Pick a muscle that is not twitching for doing the next intracellular recording between recordings. Replace the bath solution with the next higher concentration of potassium chloride solution. Being sure to cover the preparation completely each time.
Record the changes in membrane potential for each solution. The effect of raising the extracellular potassium concentration, the membrane potential is shown in this screen capture plot the resting membrane potentials for each concentration of potassium on a semi log plot of extracellular potassium concentration versus membrane potential. Line up the resting potential at 5.4 millimolar extracellular potassium.
For the hypothetical and observed curves, compare the slope of the observed line to the slope of the hypothetical line. After finishing these electrophysiologic recordings, the next step is to examine the anatomy of the muscle fibers and their innervation pattern. Transfer the preparation to the standing dish and add methylene blue.
After five minutes, remove the methylene blue and add fresh crayfish saline. Look for the main nerve that innervates the muscles within a segment. Sketch the innervation pattern to the S-E-M-D-E-L 2D EL one and DEM muscles in a segment.
Next, move the preparation underneath a fume hood. Remove the saline and add the fixative. The fixative used here is a poin solution.
The purpose of the fume hood is to avoid the vapors of the fixative. Be very careful not to get this solution on your skin or in your eyes. If your eyes start to burn, wash your eyes immediately at the eyewash station.
Let the boin solution remain on the preparation for about 10 minutes, and then use a pipette to exchange the solution for saline. Cut out a thin segment of DEL one or DEL two muscle. Place it on a glass slide.
Label the slide. Repeat the procedure for the SEM muscle. Use the compound microscope to view the sarcomere banding pattern in both tissue preparations.
The next series of experiments will examine how to record synaptic responses in crayfish abdominal muscles. Select another crayfish, decapitate it. Remove the thorax and appendages and expose the abdominal extensor muscles as shown in the surgery.
In the first section of this video, using the microscope, find the nerve carrying the motor axons to the extensor muscles. Look for the segment with the most accessible nerve. The nerve is white and can be seen by using a pipette to spray saline on the preparation or by lightly blowing on the preparation.
This causes the nerve to move and thus makes it easier to identify. Place the suction electrode held by the microm manipulator to directly over the nerve. Gently pull on the syringe to draw the nerve into the electrode.
You can see the nerve being sucked into the electrode through the microscope. Adjust the settings in the lab chart software as specified in the accompanying article. Fill a three molar potassium chloride glass micro electrode and connect it to the head stage and power lab.
Use the course knob on the amplifier to move the line on the lab chart to zero. Before inserting the electrode, the toggle knob should be turned on and then off several times. In order to test the electrode resistance, measure the amplitude of the resulting values.
Look through the microscope and use the electrode probe to insert the electrode into one of the longitudinal muscle fibers, either DEM or DL one or DL two. Be careful not to penetrate through the muscle fiber. The preparation is now ready for the experiment.
The suction electrode is used to stimulate the segmental nerve bundle and the glass micro electrode is used to record the synaptic response from the muscle fiber trigger the graft stimulator to stimulate the nerve at one hertz for phasic responses. Next, place the electrode into an SEL muscle fiber. The SEL is a tonic muscle.
Stimulate the nerve with short bursts of pulses at 10 hertz for tonic responses. Compare the recorded responses of the DEL and SEL muscles. Note that the phasic DEL muscles have larger amplitude synaptic responses than the tonic SEL muscles.
This next set of experiments focus on the crayfish flexor muscles rather than the extensor muscles. Starting with a new crayfish, isolate the abdomen as shown in this surgery. In the first section of this video, place the isolated tail preparation in a saline filled Petri dish, pin down the tail and upper portion of the preparation.
Note that dec Caro experiments on the tonic flexor muscles, the ventral nerve cord must be left intact. Use a scalpel to start a longitudinal cut through the articulating membrane between two ribs on the ventral side of the preparation. Be very careful not to cut too deeply when making this cut as a deep cut could transect the motor nerve root to ensure that the superficial muscles are not damaged.
Do the next step. Looking through a microscope using scissors or a scalpel cut horizontally from the longitudinal cut in the thin membrane layer covering the muscle. Enlarge the cut to make a flap of articulating membrane and lift the flap upward.
Use scissors to cut off the flap exposing the superficial flexor muscles for intracellular recording from the superficial flexor muscles. Use the same instrumentation and setup as was used for recording the resting membrane potential and the synaptic responses from the abdominal extensors. As before, use an intracellular electrode filled with three molar potassium chloride.
Insert the electrode into a muscle fiber of the superficial flexor muscles, being careful not to penetrate through the muscle. Record the spontaneous activity of the EPSPs. Note the different sizes of the EPSPs, and if any IPSPs are present.
Take a small paintbrush and carefully brush along the cuticle edge within the same segment containing the recording electrode. Note any change in frequency or size of the EPSPs due to the stimulation if a greater response is observed. This would indicate recruitment of additional motor neurons responding to the change in sensory nerve firing.
Carefully exchange the saline bath solution to one containing a neuromodulator such as one micromolar serotonin or saline bubbled with carbon dioxide. Then repeat the paintbrush stimulation. Note the effect that changing the bath solution has on the recorded activity.
Change the bath saline back to normal saline and observe that the muscle fiber activity returns to baseline. This figure shows EPSPs recorded in a superficial flexor muscle fiber. Note the different sizes of the EPSPs.
The next stage of this experiment uses an extracellular electrode to monitor action potentials from motor neurons responding to activity in the sensory to CNS to motor neuron circuit. Start by making a suction electrode to stimulate the sensory nerves. Pull a piece of plastic tubing over a flame.
The next step is to trim the tubing back to the desired size. To do this, you will have to look at the nerve in your preparation to judge the desired opening diameter for your suction electrode. Depending on the size of the crayfish, the nerve bundles can vary in size from about one millimeter to a few millimeters in diameter.
Cut the stretched section of tubing to make the opening in the tip the correct size to hold the nerve. If the opening is too large, the nerve could fall out. If the opening is too small, the nerve could be damaged by the pressure of the electrode.
Place, the plastic tubing on the tip of a glass suc electrode. The electrophysiology and software setup is slightly different from monitoring extracellular responses as compared to intracellular recordings set up. The equipment as specified in the accompanying article, suction the third route that innervates the superficial flexor muscle into the suction electrode.
You can now begin to record action potentials. This route has five excitor motor neurons and one inhibitor motor neuron. After recording baseline activity, stimulate the cuticle with a brush to observe the change in generated action potentials.
Following this, replace the bath solution with neuromodulators such as serotonin as in the previous experiment, and record any change in response. Additional exercises are given in the accompanying text. This recording shows the action potentials recorded from the third route before and during stimulation of the cuticle with a brush, compare the frequency of action potentials prior to brushing to their frequency during stimulation.
This recording shows the action potentials recorded from the third route in normal saline and in a serotonin solution. Compare the frequency of action potentials in normal saline to their frequency and serotonin As just shown. Brushing increases the firing frequency over baseline.
Activity also demonstrated is that serotonin increases the firing frequency over normal saline. This figure shows that when brushing and serotonin are combined, there is an even greater increase in firing frequency. The comparison of externally and theoretically derived effects of external potassium concentration on resting membrane potential indicates the influence of ions on membrane potential.
We've also showed you how to record ntic responses at neuromuscular junctions of both phasic and tonic muscles. These techniques can be used for investigative student laboratories to teach fundamental concepts and physiology. The techniques obtained in this laboratory exercise can be used to answer questions related to other laboratory preparations, as well as physiological applications related to medicine and health.
