Membrane Transport

Overview

Most cells have a potential difference across their membrane. The potential inside is negative with respect to the outside and the magnitude of the potential is between 40mV and 100mV--depending upon the cell and the environment. Generally the membrane potential is produced by two factors:

The sodium-potassium pump uses energy from ATP hydrolysis to transport three sodium ions out of the cell in exchange for two potassium ions into the cell. This exchange of potassium and sodium ions helps produce an asymmetric distribution of ions across the membrane, so that sodium is at a higher level outside the cell while potassium is at a higher level inside the cell (See The distribution of ions and charged proteins across the membrane of the squid giant axon, the equilibrium and membrane potentials.). Thus these ions tend to diffuse down their concentration gradients and then return to their original location by the pump. However, this process of diffusion is interfered with by a membrane that is not equally permeable to these ions and by the presence of negatively charged proteins inside the cell.

 

  1. The distribution of ions and charged proteins across the membrane of the squid giant axon, the equilibrium and membrane potentials.

Ion

Intracellular concentration (mM)

Extracellular concentration (mM)

Equilibrium potential

Sodium

50

440

+55MV

Potassium

400

20

-76mV

A- proteins

345

0

 

 

 

Membrane potential = -65 mV

Proteins are large molecules that usually do not move across the cell membrane. Large quantities of negatively charged amino acids, like aspartate and isothionate, may be incorporated into intracellular proteins and produce an electrostatic gradient that attracts cations into the cell. In the case of sodium, the concentration gradient (See The distribution of ions and charged proteins across the membrane of the squid giant axon, the equilibrium and membrane potentials.) and the electrostatic gradient attract sodium ions into the cell. However, the low permeability of the resting membrane to sodium causes only a few sodium ions to enter the cell. The two gradients for potassium work in opposite directions: the concentration gradient (See The distribution of ions and charged proteins across the membrane of the squid giant axon, the equilibrium and membrane potentials.) pushes potassium out of the cell, while the electrostatic gradient attracts potassium into the cell. In the resting cell, the relatively high permeability of the membrane to potassium causes these ions to leave the cell. In the resting cell, these displaced ions are picked up by the sodium-potassium pump and transported back across the membrane to maintain status quo. Thus, there is a constant flux of cations across the membrane.

A passing knowledge of Ohms' Law (voltage = current x resistance) tells you that the movement of ions (current) across the membrane (resistance) produces a voltage. The voltage produced by the flow of any particular ion is called the equilibrium potential and is given by the Nernst Equation, which for K+ (at 20 degrees Celsius) is:

 

EK+ (millivolts) = 57.5 log10 [K+]out

[K+] in

 

While this equation can be used to derive the equilibrium potential for any ion, the values cannot be used to predict the membrane potential (See The distribution of ions and charged proteins across the membrane of the squid giant axon, the equilibrium and membrane potentials.). Instead a second equation, the constant field or Goldman equation, must be used to predict the resting membrane potential.

 

Em (millivolts) = 57.5 log10 (PK[K]out + PNa[Na]out + PCl[Cl]in)

PK[K]in + PNa[Na]in + PCl[Cl]out

 

This equation assumes that the membrane potential is produced by the combined effect of all ions and that the contribution of each ion is determined by its relative permeability across the membrane. In most resting membranes, the permeability of the membrane is highest to potassium, so the membrane potential is close to (but not the same as) the equilibrium potential for potassium (See The distribution of ions and charged proteins across the membrane of the squid giant axon, the equilibrium and membrane potentials.).

Experiment 4: Membrane Potentials

Overview

The aim of the present laboratory exercise is to record resting potentials across the membranes of fast extensor muscle fibers in the tail of crayfish (Procambarus Clarkii). Microelectrodes are glass capillary tubes which have been melted and then pulled to produce a very fine (<0.5 um diameter) tip at one end. The tip is placed through the membrane and is so fine that the membrane seals around the tip. The microelectrode is filled with potassium chloride (plus a dye) so that it acts as a "saline bridge" between the inside of the cell and the recording equipment.

The Goldman equation can be used to predict the membrane potential.

 

Em (millivolts) = 57.5 log10 (PK[K]out + PNa[Na]out + PCl[Cl]in)

PK[K]in + PNa[Na]in + PCl[Cl]out

 

According to this equation, the membrane potential depends upon the concentration of the different ions across the membrane and the relative permeability of the membrane to these ions. Since you will record from muscle fibers that are functionally identical--they all contract to rapidly extend the tail--is it reasonable to assume that the fibers within this muscle will be the same? You will test this idea by measuring the membrane potential from several fibers in the same muscle and in the muscle in different abdominal segments.

Finally, the above equation indicates that the membrane potential is dependent upon the concentration gradients of the different ions. Since the permeability of the resting membrane is highest to potassium, it would seem reasonable that changing the potassium gradient across the membrane would have the greatest effect. You will test the notion that changing extracellular potassium concentration will change the membrane potential by measuring the membrane potential in modified crayfish saline with different potassium concentrations.

Equipment Required

PC computer

iWorx/214 and USB cable

Preparation dish

Dissection microscope and light source

Microelectrode probe and indifferent electrode

Microelectrode manipulator

Glass microelectrodes plus adapter

BNC-BENCH cable; grounding lead

Crayfish and modified salines (Appendix)

Equipment Setup

  1. Place the preparation dish on the microscope stage and orient the light source so that it shines onto the center of the dish.
  2. Mount the probe in the microelectrode manipulator and place it near the dissection microscope.
  3. Connect the iWorx/214 to the power outlet and to the computer (described in Chapter 1).
  4. Push the plug on the end of the microelectrode probe cable into channel three of the iWorx/214 unit (°See : Diagram to show the equipment used to record resting membrane potentials from fast abdominal extensor muscles.).
 
  1. : Diagram to show the equipment used to record resting membrane potentials from fast abdominal extensor muscles.

The Dissection

  1. Place a crayfish on ice for 10 minutes.
  2. Cut off the head and cut between the tail and thorax.
  3. Hold the tail and cut longitudinal through the shell and flexor muscles; cut along the series of indentations in the abdomen on both sides--see diagram.
  4. Begin at the anterior end of the abdomen and pull the two halves of the shell apart. It may be necessary to cut (use small forceps) the connections of the segmental flexor muscles onto the dorsal shell.
  5. Discard the ventral portion of the shell (See : Diagram to show the dissection of the crayfish tail.).
 
  1. : Diagram to show the dissection of the crayfish tail.

To expose the fast extensor muscles:

  1. Leave the telson ("tail flukes") attached to the dorsal shell.
  2. Place the dorsal shell in the preparation dish and quickly fill the dish with crayfish saline.
  3. Push one pin through the shell in the first abdominal segment and a second pin through the telson.
  4. Place the dish under the dissection microscope, focus on the preparation and use small forceps to pull out (and then cut away) the gut (the green tube in the midline) and any connective tissue.
 
  1. : Diagram to show the lateral (L) and medial (M) bundles of the fast flexor muscle in each hemisegment. The abdominal flexor muscles (F) have been removed.
  2. Examine the preparation, compare with See : Diagram to show the lateral (L) and medial (M) bundles of the fast flexor muscle in each hemisegment. The abdominal flexor muscles (F) have been removed. and identify:
  3. The six abdominal segments.
  4. A pair of fast extensor muscles in one segment--one muscle on either side of the mid-line.
  5. The medial and lateral bundles of the fast extensor muscle in each hemisegment.

The Preparation

Mount a glass microelectrode in the adapter as follows:

  1. Unscrew the end of the adapter and remove it from the barrel.
  2. Use a syringe to fill the barrel of the adapter with saline.
  3. Place the removed end of the adapter over the blunt end of an electrode so that the thread faces away from the tip.
  4. Push the end of the electrode through the orange gasket and screw the two parts of the adapter together.
  5. Push the adapter's metal plug into the probe socket.
  6. Carefully position the microelectrode tip over the preparation.
  7. Use the micromanipulator's vertical controls to move the microelectrode down until the tip touches the meniscus of the saline overlying the preparation.
  8. Connect the indifferent electrode to the alligator clip attached to the cable from the microelectrode probe. Place the indifferent electrode in the saline.
  9. Check that the microelectrode tip and the indifferent electrode are in the crayfish saline.

Start the Software

  1. Click the (Windows) Start menu, move the cursor to Programs and then to the iWorx folder and select LabScribe.
  2. When the program opens, select Load from the Settings menu.
  3. When the dialog box appears, select AK214 and then click OK.
  4. Click on the Settings menu again and select the "Membrane #1" settings file.
  5. After a short time, LabScribe will appear on the computer screen with the Membrane #1 settings.

Theory

LabScribe displays time in the horizontal direction and voltage in the vertical direction. In this exercise, you will push the tip of a microelectrode through the membrane of a muscle fiber and see the membrane potential as a downward deflection of the trace.

Important Notes

  1. If you have a problem with trace stability, check that the electrodes are in the saline.
  2. You should get into the routine of checking the resistance of your electrode--press and then release the button on the microelectrode probe cable and watch the trace deflection; the calibration is 10mV/megaOhm. A good electrode for this preparation has a resistance between five and 15 megaOhms.
  3. The trace may have a "ripple" in it. This noise may be reduced by using a cable fitted with two alligator clips--connect any piece of exposed metal on the dissection scope to any grounded point on the iWorx unit. Also, turn off and even unplug and remove the light source.

Exercise 1: Impaling Muscle Fibers

Aim: To measure the membrane potentials in different muscle fibers.

 

Procedure

  1. Click Start and then click AutoScale in the channel three title area.
  2. Look down the microscope and use the micromanipulator's controls to move the tip of the microelectrode over a bundle of muscle fibers. At this stage appreciate that:
  3. You cannot see the tip of the microelectrode!
  4. It makes no difference which fiber you penetrate.
  5. Use the micromanipulator's controls to gradually lower the tip of the microelectrode. At this stage you should look at the trace on the computer screen, not down the microscope
  6. When the electrode tip hits the membrane you will see a small deflection of the trace--up or down--the tip is now on the muscle membrane.
  7. At this stage, penetrate the muscle fiber by either:
  8. Continuing to push the electrode tip through the membrane or
  9. By gently tapping the base of the micromanipulator--this will create a small amount of vibration in the electrode tip, which will penetrate the membrane--like a pin going through a balloon.
  10. Watch the trace and notice that it will rapidly deflect downward (See : A trace viewed in the Main window to show the downward deflection when a muscle fiber is impaled. The cursors are used to measure voltage values 0.526 volts (top right of screen) or 526 millivolts.). When this happens, the tip of the microelectrode will be inside the muscle fiber so do not touch the manipulator! If necessary, click AutoScale to view the entire trace.
 
  1. : A trace viewed in the Main window to show the downward deflection when a muscle fiber is impaled. The cursors are used to measure voltage values 0.526 volts (top right of screen) or 526 millivolts.

Data Analysis

  1. Scroll to the beginning of your trace.
  2. Click the 2 cursor icon (See : The LabScribe toolbar), so that two blue vertical lines appear over the recording window.
  3. Move the mouse to place the pointer on one cursor, click the mouse and drag it to a location prior to cell penetration. Repeat with the second cursor, placing it on the plateau after cell penetration (See : A trace viewed in the Main window to show the downward deflection when a muscle fiber is impaled. The cursors are used to measure voltage values 0.526 volts (top right of screen) or 526 millivolts.).
  4. Read the value (in Volts) in the right channel title area; divide this number by 10 to obtain the membrane potential.
 
  1. : The LabScribe toolbar

Exercise 2: Membrane Potentials from Different Fibers

Aim: To measure any variations in the membrane potentials between muscle fibers.

 

Procedure

  1. Click Start.
  2. Penetrate fibers on the surface of the muscle in the same half segment, on the contralateral side and in other segments.
  3. Make measurements and convert your voltage values as described above.

Questions

  1. Were the resting potential values of fibers in any one muscle bundle identical or different?
  2. Were the average values for the resting potential the same for the medial and lateral bundle in any one hemisegment?
  3. Were the average values for the resting potential the same or different for bundles in different segments?
  4. Why was there variation in the values for the resting potential recorded from different fibers? Think of as many reasons as you can, ranging from recording error to the molecular basis of the resting potential.

Exercise 3: Membrane Potentials and Extracellular Potassium

Aim: To measure membrane potential differences created by changing [K+]out.

 

Procedure

  1. Bathe the preparation in the modified saline containing 5mM KCl--when compared with normal saline, 45mM NaCl has been replaced with 45mM choline chloride or sucrose--these two molecules do not pass through sodium or potassium channels.
  2. Wait five minutes and take measurements from a single bundle of muscle fibers that you previously identified since they have similar membrane potentials.
  3. Repeat the above using a saline with 50mM KCl--the choline chloride or sucrose has been omitted from this saline. After five minutes, impale the same muscle fibers as before.

Data Analysis

Make membrane potential measurements from your traces (See : A trace viewed in the Main window to show the downward deflection when a muscle fiber is impaled. The cursors are used to measure voltage values 0.526 volts (top right of screen) or 526 millivolts.).

Questions

  1. What happens to the resting potential if you change the level of potassium in the crayfish saline?
  2. Why does an increase in [K+]outside create the observed changes in resting potential?

Experiment 5: The Sciatic Nerve

Overview

The inside of a cell is negatively charged with respect to the outside, and the previous experiment showed that the magnitude is usually between 50 and 80 mV. This membrane potential is produced by an asymmetric distribution of charged ions across the membrane, which is created by high levels of intracellular charged proteins, a membrane with a high permeability to potassium and the sodium pump. Some cells, like nerves and muscles, can transiently reverse their membrane potential. This event, where the membrane goes from negative to positive and back to negative, takes place within milliseconds and is called an action potential. This laboratory examines the conduction of action potentials along axons in the frog sciatic nerve.

In the resting cell the permeability of the membrane to potassium (PK) is greater than that to sodium (PNa). In many cases, synaptic activity coming from other nerve cells can open channels and change the membrane potential. If the membrane potential becomes less negative (i.e. depolarized) to or beyond a certain level (called threshold) an action potential is produced. Sodium channels in the cell membrane are sensitive to membrane depolarization and respond by quickly opening. This increase in PNa allows sodium ions to diffuse into the cell and further depolarize the membrane, which increases the likelihood of additional voltage-sensitive sodium channels opening. This positive feedback system continues as the membrane potential becomes positive as it is driven towards the sodium equilibrium potential.

An inherent property of the voltage-sensitive sodium channels is that they close soon after they open. At this point the depolarized membrane potential opens voltage-sensitive potassium channels, which allows potassium ions to leave the cell and the membrane to repolarize. This process of membrane hyperpolarization closes the voltage-sensitive potassium channels and reprimes the sodium channels so that they are ready to open once more.

Action potential conduction is a process whereby the action potential moves from the site of initiation to another location in the nerve cell. When one region of the axon has an action potential, the positive charge leaks to the adjacent (unstimulated) region to depolarize it and produce an action potential. In this way the signal moves from one region to the next, and ultimately from one end of the axon to the other. Some axons are myelinated; the axon is covered with a Schwann cell--a type of glial cells which electrically insulates the axon. The spaces between the Schwann cells are called the nodes of Ranvier and they are the only regions where the axon membrane is exposed to the extracellular fluid. The myelin insulation allows the currents associated with the action potential to leak from one node to the next, so that action potentials take place only at the nodes.

In this laboratory you will record evoked action potentials from the Sciatic nerve of a frog. You will ligate the nerve at each end, remove it from the animal and place it across metal electrode in a special nerve bath chamber. You will apply brief electrical shocks to one end of the nerve to produce action potentials and record the response extracellularly from the other end of the nerve as the action potentials travel over the electrodes. You will examine certain principles associated with nerve conduction:

  1. The compound action potential--observing one or more populations of axons, each with similar conduction velocities.
  2. Stimulus-response/axon recruitment--how the response changes with increased stimulus voltage.
  3. The conduction velocity--you will measure how fast action potentials are conducted down the axons.
  4. The effects of temperature--how cooling the nerve changes the conduction velocity.
  5. Bidirectionality--whether axons conduct in both directions.

Equipment Required

PC computer

iWorx/214 and USB cable

Nerve Chamber

Leads and cables

Frog Ringer's solution (Appendix)--have two solutions:

  • most at room temperature.
  • some (about 100 ml) chilled on ice.

Equipment Setup

  1. Connect the iWorx/214 unit to the computer (described in Chapter 1).
  2. Attach the AMI connector on one end of the cable to the channel one and two input on the iWorx/214 unit.
  3. Attach one end of each of the three electrode cables to the ground and channel one inputs on the lead pedestal; use the alligator clips to connect the other ends to the electrodes on the nerve bath (See : The equipment setup to record from the sciatic nerve.).
  4. Use the mode control on the iWorx/214 to select the 3-10KHz filter. .
  5. Attach the stimulating electrodes to the iWorx/214 output and the electrodes on the nerve bath (See : The equipment setup to record from the sciatic nerve.).
 
  1. : The equipment setup to record from the sciatic nerve.

Start the Software

  1. Click the (Windows) Start menu, move the cursor to Programs and then to the iWorx folder and select LabScribe.
  2. When the program opens, select Load from the Settings menu.
  3. When the dialog box appears, select AK214 and then click OK.
  4. Click on the Settings menu again and select the "Membrane #2" settings file.
  5. After a short time, LabScribe will appear in Scope mode on the computer screen with the Membrane #2 settings.
  6. By default, the stimulus (0.1 ms and 0.25 Volts) is applied when you click Start to initiate the trace, which will take 16,000 samples--these setting can be checked by selecting Preferences in the Edit menu.

The Dissection

  1. Double-pith a frog.
  2. Remove the skin from the legs by making an incision through the skin around the entire lower abdomen. Cut the connections between the skin and the body--especially around the base of the pelvic girdle. Use stout forceps to pull the skin off the frog in one piece (like removing its pants).
  3. Moisten the exposed limbs of the frog with Ringer's solution.
  4. Locate the sciatic nerve in the abdomen: with the frog dorsal surface uppermost, use a pair of forceps to hold the urostyle. Cut the soft tissue with scissors (keep the tips up) and cut the length of the urostyle.
  5. Select the sciatic nerve on one side and use a short piece of thread to ligate the nerve as close to the vertebral column as possible. Tie the knot tightly (the limb should jump) and cut the nerve between the knot and the vertebral column. Keep the exposed nerves moist at all times with Ringer's solution. Do not pinch or stretch the nerve.
  6. Locate the nerve in the thigh by separating the muscles. You should be able to follow the nerve along the length of the thigh; a blood vessel runs parallel with the nerve.
  7. Ligate the nerve just above the knee; cut the nerve between the knot and the knee--the calf should jump.
  8. Dissect the nerve from the connective tissue in the thigh, the abdomen and around the pelvis.
  9. When the entire nerve has been exposed, grasp the threads at either end and lift the nerve from the body and place it in the nerve bath, which should be quickly filled with Ringer's solution, to immerse the nerve.

Important Notes

  1. The end of the nerve that was connected to the spinal column should be over the stimulating electrodes and the end from the knee region over the recording electrodes.
  2. The ligature (knot) should be located between the two recording electrodes in the nerve bath.
  3. Each thread should be wound around the end electrode and then draped over the edge of the bath--this will prevent movement of the nerve when the bath is drained.

Exercise 1: The Compound Action Potential

Aim: To apply a brief shock to one end of the nerve and record a compound action potential from the other.

 
  1. : The LabScribe toolbar

Procedure

  1. Click the stimulator icon in the LabScribe toolbar (See : The LabScribe toolbar) to display the stimulator panel. Check that the amplitude of the shock is 0.25 volt and the pulse width is 0.1 ms.
  2. Drain the frog Ringer's solution from the nerve chamber--if necessary, carefully dry any large drops of saline from the recording electrodes with a corner of a wipe.
  3. Click Start to stimulate the nerve with a single shock and notice the stimulus artifact at the beginning (left) of the trace and the compound action potential. At this point click Stop--your trace may look like See : The compound action potential.
 
  1. : The compound action potential

Exercise 2: Stimulus and Response

Aim: To quantify the relationship between stimulus amplitude and response amplitude.

Procedure

  1. Click the stimulator icon in the LabScribe toolbar (See : The LabScribe toolbar) to display the stimulator panel; set the amplitude to zero Volts.
  2. Drain the frog Ringer's solution from the nerve chamber--if necessary, carefully dry any large drops of saline from the recording electrodes with a corner of a wipe.
  3. Click Start to stimulate the nerve with zero Volts--observe a (flat) trace.
  4. Click the single cursor in the LabScribe toolbar (See : The LabScribe toolbar), type the voltage value into the Marks field and hit Enter on the keyboard.
  5. Click the stimulator icon in the LabScribe toolbar to display the stimulator panel; set the amplitude to 0.05 Volts. Click Start and then Stop when the trace appears--you may see a small blip on the screen at the left side of the screen, which is the stimulus artifact. Enter the voltage as a mark as before.
  6. Continue increasing the voltage in 0.05 Volt increments until a compound action potential is observed.
  7. Continue increasing the voltage and notice that the compound action potential increases in size until it reaches a maximum.
  8. Select Save in the File menu, select Desktop, name your file and click Save.
  9. Fill the nerve chamber with fresh frog Ringer's solution--to prevent desiccation of the nerve.

Data Analysis

  1. Starting with your last recording, click the 2 cursors in the LabScribe toolbar (See : The LabScribe toolbar), click and drag them horizontally to place one on the flat portion of the trace and the second on the peak of the compound action potential, as shown in See : The compound action potential.
  2. Measure the absolute voltage difference between the 2 cursors in the title bar--the value for V2-V1 in See : The compound action potential is 0.508 Volts.
  3. Scroll back through your recordings repeating the above process, matching the measured amplitude of the compound action potential with the voltage applied--type the numbers into the Marks field.
  4. Graph or tabulate your results.

Questions

  1. In single axons, does action potential amplitude change?
  2. How many axons are producing an action potential at a zero Volts shock?
  3. If the size of a single action potential does not change and you are recording from many axons, how many axons are firing after threshold is reached for the compound response?
  4. Why does the compound action potential increase in size to a maximum response?
  5. How many axons are firing when the maximum response is recorded?

Exercise 3: Conduction Velocity

Aim: To measure the velocity of action potential conduction

Procedure

  1. Click the stimulator icon in the LabScribe toolbar (See : The LabScribe toolbar) to display the stimulator panel; set the amplitude to 0.25 Volts--or a voltage that produces a compound action potential with a maximum amplitude.
  2. Drain the frog Ringer's solution from the nerve chamber--if necessary, carefully dry any large drops of saline from the recording electrodes with a corner of a wipe.
  3. Click Start to stimulate the nerve and observe the compound action potential--type "long pathway" into the Marks field and hit Enter on the keyboard.
  4. Use the clips on the stimulating electrodes and move each by an increment of one electrode bar towards the recording electrodes.
  5. Click Start to stimulate the nerve and observe the compound action potential--type "pathway - 10 mm" into the Marks field and hit Enter on the keyboard.
  6. Fill the nerve chamber with chilled Ringers solution. Select Save in the File menu.

 

 
  1. : Conduction velocity--two stacked traces.

Data Analysis

  1. Click the Analysis icon in the toolbar (See : The LabScribe toolbar) and deselect all sweeps except the last two and select stacked--your traces may look like See : Conduction velocity--two stacked traces..
  2. Click the 2 cursor icon in the toolbar (See : The LabScribe toolbar); click and drag them to the peaks of each response (See : Conduction velocity--two stacked traces.).
  3. Read off the T2-T1 value (0.0052s or 0.52ms in See : Conduction velocity--two stacked traces.).
  4. Calculate the conduction velocity (in m/s) as (10 mm/T2-T1 value in ms).

Theory

You moved the stimulating electrodes by one electrode--a distance of 10 mm--and you measured the change in the time between the two peaks. Thus, you can calculate conduction velocity

Exercise 4: Conduction Velocity and Temperature

Aim: To examine the effects of cooling on the velocity of action potential conduction

Procedure

  1. Repeat the above procedure after draining the chilled Ringer solution from the nerve bath.
This part of the experiment must be done quickly since the nerve will begin to warm as soon as the bath is drained.
  1. Fill the bath with Ringers at room temperature after the experiment and allow to warm as you perform the data analysis as described above.

Questions

  1. Does the conduction velocity change when the nerve is cooled?
  2. What channel parameters may change with temperature?

Exercise 5: Bidirectionality

Aim: To examine whether an action potential travels in the wrong direction and if so, at what velocity.

Procedure

  1. Use forceps to remove the threads from the electrode posts, turn the nerve 180 degrees and reattach the threads. You will now stimulate and record from the nerve at opposite ends.
  2. Drain the frog Ringer's solution from the nerve chamber--if necessary, carefully dry any large drops of saline from the recording electrodes with a corner of a wipe.
  3. Click Start to stimulate--if necessary increase the amplitude of the shock to produce a compound action potential.
  4. Use the above procedure to measure the conduction velocity.

Questions

  1. Do you record an action potential?
  2. What is the conduction velocity? Is this similar to the first value you recorded when the nerve was in the original orientation?
  3. If these are the same axons, how can an axon conduct as action potentials in both directions?
Where are the cell bodies and synapses in this preparation?

Experiment 6: Reflexes and Reaction Times

Overview

During our day-to-day lives we detect changes in the environment and react appropriately. An external stimulus is detected by one or more neurons, which send the sensory information to the central nervous system, where it is processed. If a motor response is initiated, it usually involves a series of action potentials that produce a muscle contraction and a movement of one or more parts of the body. A simple reflex is perhaps the easiest of this type of stimulus-response reaction. A loud sound or something flying at your eye makes you blink, while a tap on the tendon under the knee cap produces the knee-jerk (or myotactic) reflex.

 
  1. : A cross section of the spinal cord showing the single synapse between the sensory and the motor neurons involved in the myotactic reflex.

A simple reflex like the myotactic reflex is produced via single synapses between sensory axons and motor neurons. The required circuitry for this reflex is confined to the spinal cord, as shown in See : A cross section of the spinal cord showing the single synapse between the sensory and the motor neurons involved in the myotactic reflex.. Sensory information also ascends to higher centers, but the brain is not necessary or required to perform the reflex. More complex reflexes usually involve additional (inter-) neurons and more than one population of motor neurons. Thus, more neurons and synapses are involved, which usually results in a longer delay between stimulus and response and often a more complex response. One example of such a complex response is the flexion withdrawal reflex, where a noxious stimulus to one leg causes withdrawal of the stimulated leg and extension of the other.

In this lab you will study the time taken between a stimulus and the response. These reaction time measurements will be made from a volunteer subjected to harmless visual and sound stimuli. In addition, the effect of priming and prediction will be examined.

Equipment Required

PC computer

IWorx/214 and USB cable

Event marker

Plethysmograph

Equipment Setup

  1. Connect the iWorx/214 unit to the computer (described in Chapter 1).
  2. Plug the event marker into the DIN socket on channel three.
  3. Plug the plethysmograph into the DIN socket on channel four. The equipment should look like See : The equipment used to measure reaction times from a volunteer..
 
  1. : The equipment used to measure reaction times from a volunteer.

Start the Software

  1. Click the (Windows) Start menu, move the cursor to Programs and then to the iWorx folder and select LabScribe.
  2. When the program opens, select Load from the Settings menu.
  3. When the dialog box appears, select AK214 and then click OK.
  4. Click on the Settings menu again and select the "Membrane #3" settings file.
  5. After a short time, LabScribe will appear on the computer screen with the Membrane #3 settings.

Exercise 1: Reaction Time and Sound

Aim: To measure the reaction time of a volunteer to a sound.

 

Procedure

  1. Ask the volunteer to sit in a chair placed in a location so that their back is facing the computer screen with the keyboard immediately behind them.
  2. Ask the volunteer to relax and listen as another student taps the white surface of the plethysmograph with a pencil. Ask the volunteer if they can hear the tapping sound.
  3. Ask the volunteer to click the event marker as soon as they hear the tap.
  4. Click Start.
  5. Present a total of 10 taps, but make sure that the taps are delivered so that the volunteer cannot predict when the stimulus will be presented.
  6. Click Stop to halt recording.
  7. Select Save As in the File menu, type a name for the file and save it in an appropriate place on the hard drive.

Data Analysis

  1. Click the 2 cursor icon (See : The LabScribe toolbar), so that two blue vertical lines appear over the recording window.
  2. Drag the cursors left and right so that the large spike on the plethysmograph channel and the signal from the event marker are located between the two blue lines.
  3. Click the Analysis icon (See : The LabScribe toolbar) to open the Analysis window.
 
  1. : The LabScribe toolbar
  2. Use the mouse to click and drag one cursor to the beginning of the spike on the plethysmograph channel and the second cursor to the onset of the signal from the event marker (See : Data produced by tapping the plethysmograph, which entered a large spike on the lower trace and produced a sound, which the volunteer used as a cue to press the event marker (upper trace). The data are displayed in the Analysis window and the two cursors are positioned to measure the reaction time (T2-T1).).
  3. Enter the time (T2-T1) difference into the Journal either by typing the values directly, or by right clicking in the Analysis window and selecting send data/titles to the Journal.
  4. Use the scroll bars to scroll through your data in the Analysis window and repeat the measurements for all 10 trials.
  5. Omit the longest and shortest values and average the remaining eight values to give the mean reaction time.
 
  1. : Data produced by tapping the plethysmograph, which entered a large spike on the lower trace and produced a sound, which the volunteer used as a cue to press the event marker (upper trace). The data are displayed in the Analysis window and the two cursors are positioned to measure the reaction time (T2-T1).

Exercise 2: Reaction Time and Prompted Sounds

Aim: To measure the reaction time of a volunteer to sounds delivered immediately after a verbal prompt.

 

Procedure

Repeat exercise #1, but ask the volunteer if they are ready immediately prior to tapping the plethysmograph.

Data Analysis

Measure the time interval (T2-T1) between stimulus and response for each event (See : Data produced by tapping the plethysmograph, which entered a large spike on the lower trace and produced a sound, which the volunteer used as a cue to press the event marker (upper trace). The data are displayed in the Analysis window and the two cursors are positioned to measure the reaction time (T2-T1).).

Exercise 3: Reaction Time and Predictable Sounds

Aim: To measure the reaction time of a volunteer to sounds delivered at a predictable interval.

 

Procedure

Repeat exercise #1, but tap the plethysmograph at a predictable interval.

Data Analysis

Measure the time interval (T2-T1) between stimulus and response for each event.

Questions

  1. Is the average reaction time the same for all three conditions?
  2. Does the time interval decrease during exercises two and three?

Exercise 4: Reaction Time and Visual Cues

Aim: To measure the reaction time of a volunteer to a visual cue.

 

Procedure

  1. Ask the volunteer to sit in a chair and face the computer screen.
  2. Ask the volunteer to watch the screen and (quickly) press the event marker button as soon as they see the trace deflection.
  3. Click Start; a second student (out of site) should tap the plethysmograph (and the volunteer should respond).
  4. Present a total of 10 trials, but make sure that the events are delivered so that the volunteer cannot predict when the visual cue will be presented.
  5. Click Stop to halt recording.
  6. Select Save from the File menu.

Data Analysis

Your result may look like See : Data produced using a visual cue from the plethysmograph). The volunteer used this visual cue on the screen (lower trace) to press the event marker (upper trace). The data are displayed in the Analysis window and the two cursors are positioned to measure the reaction time (T2-T1)..

 
  1. : Data produced using a visual cue from the plethysmograph). The volunteer used this visual cue on the screen (lower trace) to press the event marker (upper trace). The data are displayed in the Analysis window and the two cursors are positioned to measure the reaction time (T2-T1).
  2. Use the cursor and the marker to measure the time delay between visual stimulus and response.
  3. Repeat the measurements for all 10 trials.
  4. Omit the longest and shortest values and average the remaining eight values to give the mean reaction time.

Questions

  1. Is the average reaction time comparable to the data from exercise #1? What does this tell you about the time taken to react to oral and visual cues?

Exercise 5: Reflexes

Aim: To examine different reflexes.

 

The Knee-Jerk Reflex

  1. Ask the volunteer to sit in a chair and cross their legs.
  2. Firmly strike the tendon below the knee cap and watch the knee jerk.
  3. Ask the volunteer to cup and link their hands and then pull outwards (this is the Jendrassil maneuver). Repeat step #2.

 

What effect does the Jendrassil maneuver have on the knee-jerk reflex? Can you explain why?

The Papillary Reflex

  1. Shade the eyes for 30 seconds.
  2. Shine a light into one eye. What is the response of the pupil?
  3. Repeat steps 1 and 2, but note the response of the unstimulated eye.

 

What is the effect of light on the eyes? Can you explain why this reflex would be beneficial to you?

The Spinociliary Reflex

  1. A student should look into one of the volunteer's eyes.
  2. The student should gently stroke the hair in the hairline behind one ear. What is the effect on the pupil?
  3. Repeat step #2, but stroke the hair behind the other ear.

 

Can you explain why this reflex would be beneficial to you?