Neuromuscular Junction
Lab Manual
© University of Minnesota
Version: July, 2015
For questions and comments about this lab manual, please contact
Professor Will Durfee, Department of Mechanical Engineering, University of Minnesota, wkdurfee@umn.edu
Table of Contents
1
INTRODUCTION............................................................................................................................... 3
1.1
1.2
1.3
1.4
1.5
1.6
2
EQUIPMENT AND INSTRUMENTATION ................................................................................... 7
2.1
2.2
2.3
2.4
2.5
2.6
3
OVERVIEW OF NEUROPHYSIOLOGY EQUIPMENT ................................................................................ 7
THE NMJ EQUIPMENT RACK ............................................................................................................. 7
OSCILLOSCOPE BASICS ...................................................................................................................... 9
PULSE STIMULATOR BASICS .............................................................................................................. 9
RECORDING ELECTRODE AND AMPLIFIER BASICS .............................................................................. 9
COMPUTER-BASED EXPERIMENT CONTROL ....................................................................................... 9
TECHNIQUES, EXERCISES AND EXPERIMENTS ...................................................................10
3.1
3.2
3.3
3.4
3.5
3.6
3.7
3.8
3.9
3.10
3.11
3.12
3.13
3.14
3.15
3.16
3.17
3.18
3.19
3.20
3.21
4
COURSE OVERVIEW ........................................................................................................................... 3
PREVIEW OF EXPERIMENT ACTIVITIES ............................................................................................... 4
NOTEBOOK ....................................................................................................................................... 5
YOUR RESPONSIBILITIES ................................................................................................................... 5
KEEPING YOUR SANITY ..................................................................................................................... 5
HELP US IMPROVE THIS MANUAL ...................................................................................................... 6
GETTING TO KNOW THE LAB ............................................................................................................10
CABLES AND CONNECTORS ..............................................................................................................10
GETTING TO KNOW YOUR OSCILLOSCOPE.........................................................................................11
WORKING THE STIMULATOR ............................................................................................................17
USING THE ELECTRODE PREAMPLIFIER.............................................................................................22
USING MICROSCOPES .......................................................................................................................24
SIMULATED CELL .............................................................................................................................25
CHECKING ELECTRODE IMPEDANCE .................................................................................................26
MAKING A GLASS MICROELECTRODE ...............................................................................................28
PITHING THE FROG ......................................................................................................................32
DISSECTING THE CUTANEOUS PECTORALIS MUSCLE ....................................................................33
MEASURING MEMBRANE POTENTIAL ...........................................................................................38
MINIATURE END PLATE POTENTIALS (MEPPS)............................................................................42
EFFECT OF K+ CONCENTRATION ON MEMBRANE POTENTIAL.......................................................44
STIMULATING THE NERVE FIBER WITH A SUCTION ELECTRODE ...................................................47
MOTOR UNIT ACTION POTENTIALS ..............................................................................................49
END PLATE POTENTIALS ..............................................................................................................52
MEASURING SARCOMERE LENGTH ..............................................................................................54
FORCE MEASUREMENTS ..............................................................................................................56
FORCE SENSOR SETUP..................................................................................................................57
MUSCLE TWITCH FORCE ..............................................................................................................59
YOUR RESEARCH PROJECT .......................................................................................................63
4.1
4.2
4.3
4.4
PURPOSE ..........................................................................................................................................63
DELIVERABLE ..................................................................................................................................63
PRESENTATION TIPS .........................................................................................................................64
RESEARCH PROJECT SUGGESTIONS ...................................................................................................64
APPENDIX A
TEK SCOPE REFERENCE .....................................................................................65
APPENDIX B
SAMPLE PLOTS ............................................................................................................69
APPENDIX C
STIMULATOR CONTROLS ...................................................................................71
APPENDIX D
FROG RINGER'S SOLUTIONS ..............................................................................73
1
APPENDIX E
NOISE!.............................................................................................................................75
APPENDIX F
BASIC EXPERIMENT DESIGN AND STATISTICS ................................................76
APPENDIX G
FITTING A STRAIGHT LINE TO DATA USING EXCEL .................................77
APPENDIX H
BIPOLAR HOOK ELECTRODES ..........................................................................79
APPENDIX I
FIXING SUCTION ELECTRODES .............................................................................81
APPENDIX J
MEASURING ELECTRODE AND MEMBRANE IMPEDANCE............................82
APPENDIX K
CHANGING SETTINGS ON THE SUTTER MICROPIPETTE PULLER ........86
APPENDIX L
RESISTOR COLOR CODES ........................................................................................87
APPENDIX M
NMJ COURSE SUPPLIES AND DRUGS ...............................................................88
2
1 Introduction
1.1
Course overview
Welcome to the Neuromuscular Junction short course. The purpose of this course is to
provide you with the background and skills needed to understand basic neurophysiology
with an emphasis on nerves, skeletal muscle and the neuromuscular junction. During the
week, there will be a few foundation lectures that cover the basics of nerve, muscle and
muscle disease, and classic electrophysiology measurement methods. The primary
purpose of the course, however, is for you to conduct hands-on experiments using an
isolated frog nerve-muscle preparation.
Lecture topics for the week will include:
• Equipment in the electrophysiology lab
• Biophysical Basis of Membrane Potentials
• Neuromuscular Signal Transmission
• Muscle Contraction
plus lectures on groundbreaking research happening at the University of Minnesota.
The experiment lab stations contain state-of-the-art equipment for exciting nerves and
recording the resulting activity in membrane, nerve and muscle. Enrollment in the course
is kept small to ensure there will only be two or three students per station. This means
you can learn by doing rather than learn by reading or learn by watching.. Some of the
week is structured, but much is not. Take advantage of this unique opportunity to explore
and hone your skills. If you are taking this course as part of the Itasca summer
neurobiology experience, the lab is open 24/7. Dive in and immerse yourself!
If you do not have a technical background, the electronics and equipment will seem
strange, confusing, and sometimes overwhelming. A lot of information will be coming at
you very quickly. Don't let it get to you. The first time you confront the equipment you
will be all thumbs. The next time there will be something that clicks, and the next time
you will start to feel comfortable spinning the dials. The course will immerse you in
neurophysiology. You will be lost at the beginning of the week, but by the end you will
be an expert. Stick with it and you will be rewarded.
This lab manual covers some of the basics of instrumentation found in a neurosciences
wet lab and the experiments and methods you will be learning this week. A companion
Lecture Notes document contains background reading in basic neuromuscular junction
physiology. Read this material on your own because it will help you understand the
lectures and will put the experiments in context.
3
1.2
Preview of experiment activities
The laboratory side of the week starts with an introduction to the basic equipment and
instrumentation that are at the heart of experimental neurophysiology. We start out
reviewing how an oscilloscope works, then move on to electrical pulse stimulators,
isolators and recording amplifiers. Next, you learn how to pull, prepare and test a glass
microelectrode with a tip diameter less than 0.1 microns for insertion directly into an
axon.
The experimental prep you will be using this week is the cutaneous pectoralis muscle of
the Rana pipiens (Grass frog), a superficial muscle located in the chest. This is a
convenient muscle to use because it is formed into a thin, broad sheet making it
particularly easy to stick microelectrodes into single muscle fibers. The muscle sheet is
two or three cells thick and has about 200 cell across its breadth. You'll learn how to
sacrifice the frog by pithing, and then will use careful micro-dissection techniques to
extract an intact muscle and nerve. This is quite challenging because the muscle is waferthin and the nerve so small that it can barely be seen. Successfully passing this hurdle
should give you a real appreciation for the difficulty and complexity of experimental
preps
You will be conducting many experiments with this prep, some of which involve
recording membrane potentials, some of which involve recording muscle force. The
excitation will either be changes in the chemistry of the bathing solution that surrounds
the muscle, or stimulation of the muscle nerve. What you do will depend both on your
interests and how the class proceeds. There are a few experiments every group will do
including measuring the resting membrane potential and determining whether the
theoretical potential expressed in the Goldman-Hodgkin-Katz equation holds as the
extracellular potassium concentration is changed. In another experiment you will measure
end-plate potentials evoked by nerve stimulation and view the all-or-none phenomenon
that characterizes muscle fiber depolarization. You will also search for miniature end
plate potentials that are indicators of the quantal nature of acetylcholine release from the
presynaptic terminal. Other experiments you may get to include measuring the length of a
sarcomere, examining the timing characteristics of a muscle force twitch, and finding the
force-length properties of stimulated muscle.
The end of the course is devoted to independent experimental research projects carried
out by each team. Here is where you can follow your own interests and try out in depth
something that sparked your interest from the earlier experiments, or launch into some
new experiments based on something you read. The instructors can help you formulate a
research question, and you can quickly come up to speed on a topic using the reference
books in the lab and on-line literature searches and article reviews. The course finishes
with a brief presentation by each team on their research results.
There will be a variety of views about using the frogs as sources of material for the
experiments. No matter what your views, please treat these experiments seriously and
make every effort to maximize the amount of learning you receive from every prep. In
designing the course, the instructors felt strongly that the most valuable part of the
4
experience are the experiments using live biological tissue. One can certainly learn
theoretical neuroscience from books, videos and simulations. True understanding,
however, only comes from experience with the real thing.
1.3
Notebook
The lab notebook is the lifeline of a good scientist. Use your notebook as a diary of the
week. Record everything about everything you do in the lab. Add sketches of setups and
commentary on what worked and what did not work. Record your failures along with
your successes. Tape plots or printouts right in the notebook. Write down your
observations, hypotheses and methods. Make note of anything relevant that you read and
where you read it. Sketch out the steps you used in dissecting and sketch your experiment
setups.
Number each page and date major entries. Use ink. You can cross out entries, but don't
erase. Who knows, you may have an idea that later turns into a patent. Your dated
notebook will be the record of invention.
You will have done well if you run out of notebook pages at the end of the week.
1.4
Your responsibilities
This week will be a guided, immersive experience. You will have plenty of support from
the instructors, but you must assume the responsibility for learning. The course is very
different from any cookbook style lab course you may have taken in the past. We will
help you learn how to operate the tools, but using them in a way that maximizes your
learning is up to you. Don't expect the equipment to be perfect, the instructions to be
accurate and all experiments to work. This is the real world. This is how real research
labs operate. Sometimes things will go flawlessly, sometimes not. At some point during
the week, you will get frustrated. When you successfully work past this, you'll be
rewarded with the satisfaction of having mastered and understood some of the most
challenging techniques and principles that form the foundation of experimental
neuroscience.
The equipment you will be using is expensive, thousands of dollars. Treat it with respect.
If something breaks, try and fix it. If you can't, alert an instructor.
There are no parents or custodians to clean up after you. At the end of each day, clean up
your mess. When you leave on the last day, your station should look exactly like it did
when you sat down on the first day. Same with the common areas. Cleanup is a shared
responsibility and everyone must do their part.
1.5
Keeping your sanity
When you get the course schedule, you will notice that it goes all day with no break
except for lunch. Everybody hits the wall after a while. We encourage you to take brief
mental health breaks every so often. You don’t have to ask. Get outside for 5 minutes and
stretch your legs. You will come back refreshed and ready to go.
5
1.6
Help us improve this manual
The NMJ Lab Manual is a work in progress. Please help us to make the manual better for
next year’s students. Alert the instructors about typos and about sections that you found
confusing. Any and all comments are welcome!
6
2 Equipment and instrumentation
2.1
Overview of neurophysiology equipment
The fundamental method of experimental neuroscience is to excite the system of interest
and measure the response (Figure 2-1). No matter how sophisticated (or simple) the
experiment, every piece of associated equipment is there either to (1) excite the system,
(2) monitor the response, or (3) to control the experiment.
Stimulation
equipment
Excitation
System
under test
Response
Measuring
equipment
Figure 2-1: In most neuroscience experiments you stimulate a system and measure the response.
In your experiments, the primary form of stimulation will be the application of electrical
pulses to the nerve through a stimulation electrode. The system can also be excited or
modified chemically by changing the concentrations of ions in the solutions which bathe
the biological tissue.
The primary form of the response is the voltage across a single muscle cell membrane
that you will measure through a microelectrode piercing the membrane, connected to an
amplifier that multiplies the small membrane voltages up to a signal that can be recorded
on an oscilloscope or computer.
A second form of response is the whole muscle force resulting from the application of a
stimulus. If a single pulse is applied, a single muscle force twitch is elicited. If a train of
pulses is applied, the output force is constant. The forces are small and are detected by
tying one end of the muscle through a small suture to the loading pin of a sensitive force
transducer that converts the force to a proportional electrical signal. Gain and offset are
applied to the electrical signal by a force sensor amplifier for recording on an
oscilloscope.
The following sections describe the stimulating and recording equipment used in the
course. Laboratory exercises designed to get you familiar with this equipment appear in
Chapter 3.
2.2
The NMJ equipment rack
The equipment rack used in this course is shown in Figure 2-2.
7
Figure 2-2: Equipment rack.
At the top is an amplifier used to boost small membrane potentials. Next down is a set of
utility instruments, including a bridge amplifier used to condition the signal coming from
the force transducer. Next is an oscilloscope. Below that are two stimulators.
8
2.3
Oscilloscope basics
Will be covered in class.
2.4
Pulse stimulator basics
Will be covered in class.
2.5
Recording electrode and amplifier basics
Membrane potentials are measured using a fluid filled glass micropipette electrode whose
tip is drawn to a fine diameter by heating and pulling rapidly in a precision micropipette
pulling machine. The machine uses an electric coil to heat and soften the glass coupled
with a two-step pulling process. The first pull is slow to gradually taper down the tip. The
second pull is rapid to give the electrode a sub-micron tip diameter. The size and shape of
the tip are determined by the heater temperature and the speed and timing of the two
pulling stages.
2.6
Computer-based experiment control
The modern neurophysiology research lab uses a computer to acquire data and control
experiments. Commercial software is available for running common neurophysiology
experiments, but many labs write their own data acquisition and control applications
using LabVIEW (www.labview.com), while a few write software from scratch using C++
or Visual Basic.
Computers are not used in this course because it is important for you to get your hands
dirty while fighting to control the experiment manually and to get signals to appear on the
scope. If you are staying on at Itasca to take additional modules of the neurosciences, you
will be using LabVIEW applications that have been written for the course that may make
some of the data acquisition tasks simpler.
9
3 Techniques, exercises and
experiments
3.1
Getting to know the lab
Sometime during the day, explore the entire lab (both rooms). Rummage through all the
cabinets and shelves and see what’s there. Who knows, later in the week you may
actually need some of that aluminum foil you have found.
Treat equipment and supplies with care, but don’t worry about breakage because most of
the equipment is reasonably rugged. The only exception is the Olympus microscope. If
you loosen the handles marked with orange tape and the scope comes crashing down,
that’s serious. So think before you leap.
3.2
Cables and connectors
The well-equipped neurophysiology lab is a maze of cables and connectors, all with the
purpose of bringing electrical signals and power from one place to another. Fortunately,
in this lab we have standardized on just a few types of the more common cables and their
corresponding connectors.
Figure 3-1 shows the cables and connectors you will encounter in the lab. "BNC" cables
are what you will use to make connections between various instruments. The signal is
carried by the center conductor while the outside is ground. Make the connection by
pushing on and giving a partial twist to lock into place. An inventory of cables in
standardized lengths is kept in the lab. You will also find male-to-male BNC adapters to
join cables and BNC "Tee's" for connecting more than one cable to an instrument.
Banana leads and banana plugs are also common for interconnects. When using a BNCto-banana adapter, pay attention to orientation. The marked tab on the adapter indicates
ground. Alligator clips are also handy for making connections
Be creative in making your connections. You can get a signal from anyplace to anyplace
if you put your mind to it.
10
Figure 3-1: Cables and connectors used in the lab. On the left is a BNC cable terminated with a BNC
connector. The center shows a BNC-to-banana adapter. On the right is an alligator clip. On
the far right are a set of banana-to-banana test leads (pix from www.jameco.com)
3.3
Getting to know your oscilloscope
The objective of this exercise is for you to become familiar with controlling the
oscilloscope. Viewing electrophysiological signals can be tricky. The more comfortable
you are manipulating the scope, the better you will be at capturing signals from your
prep. You have completed this exercise when you turn in the deliverable at the end of
this section.
The Tektronix TDS 1002 oscilloscope is shown in Figure 3-2. This is a 2-channel, 60
MHz digital storage scope with inputs coming in from the two BNC connectors at the
bottom. The controls let you set the amplitude and offset of the signals, how quickly the
trace sweeps across the screen, the horizontal position of the trace and the trigger settings.
The set of five buttons just to the right of the screen are called the option buttons because
their function changes depending on what menu is being displayed on the screen.
Although the scope is a sophisticated instrument with many settings and options, if you
have a reasonable understanding of how scopes work, you can figure out the Tek scope
by trial and error fiddling with the knobs.
Appendix A contains reference information on the scope, including identifiers for each
on-screen icon.
11
Figure 3-2: Tektronix oscilloscope. Menu area is at the top. Option buttons are to the
right of the display, then going right are vertical controls, horizontal
controls and time base controls..
To play with the scope, you need a signal. Most scopes are equipped with a calibration
signal source, typically a voltage square wave at a specified voltage and frequency 1.
Find the calibration signal on the Tek scope. (Hint: It's located at the bottom right and
labeled PROBE COMP.) What is the voltage and frequency of the signal?
Connect CH1 to the PROBE COMP lug using a BNC, BNC-to-banana adapter and a
alligator clip on the signal side (the side without the GND tab) of the adapter.
1
The calibration jack is used to see if the scope is working and to "tune" a scope probe.
Scope probes have an internal resistance and capacitance circuit and must be matched to
the input impedance characteristics of the scope. If you look carefully at a good-quality
scope probe (in this lab, we don't have probes, just BNC cables to make connections) you
will see a small screw that allows you to adjust the probe capacitance. To tune a probe,
connect to the calibration source and get a stable wave on the screen. Adjust the screw
until the top and bottom of the square wave are perfectly flat.
12
(Figure 3-3). Be careful to connect to the signal side of the BNC adapter, and not the
ground side that has the tab. Also be careful not to short the alligator clip to the ground
lug on the scope.
Power up the scope. Push CH1 MENU, then the option button (one of the buttons just to
the right of the screen) to select Probe = 1X. Push the AUTOSET button (at top right). In
a short while you should see a 1 kHz, 5 V peak-to-peak square wave on the screen. This
confirms that the scope is working and gives you a signal to play with.
Now, spin the VOLTS/DIV, the VERTICAL POSITION, the HORIZONTAL
POSITION, and the SEC/DIV knobs until everyone on the team understands what each
does.
Figure 3-3: Setup for checking operation of the scope. Connect the scope
PROBE COMP signal to the CH1 input. Push AUTOSET to
get a display. Select Probe = 1X. Manipulate the amplitude
control and the time base and trigger controls until you get a
stable trace that looks something like this.
.
Set the vertical position of the trace moving the POSITION knob until the bottom of the
square wave lines precisely with one of the graticule lines on the screen. Note that zero
volts for the trace is marked by the arrow labeled “1” at the left of the screen. Set the
horizontal position so that the start of one pulse lines up with a graticule line.
What is the peak-to-peak voltage of the signal? What is the frequency in Hz (cycles per
second) of the signal? Does it match what is written on the PROBE COMP jack?
13
Press CH1 MENU then cycle through the options under the Coupling button. DC couples
the signal directly to the display. AC removes the average signal by high-pass filtering
before sending the signal to the display. AC coupling is handy when looking at small
transient signals, for example MEPPs, that ride on top of a large static signal such as a
membrane resting potential. Note how the high-pass filter of the AC coupling distorts the
shape of the pulse. GND coupling sets the channel to 0 Volts before passing to the
display. The GND setting is handy when you are trying to find a trace. When done, leave
CH1 DC coupled.
NOTE: Until now, it is likely that one team member has spun the knobs while the
others have sat around and watched. Now is the time for you to shuffle seats and
change who actually touches the knobs. Go ahead. Don't be shy!
The scope has several built in features for measuring signals. We’ll explore them now.
Push the CURSOR button (in the MENUS area at the top), then with the option buttons
(the ones to the right of the display), select Type = Voltage. Two horizontal cursor lines
will appear on the screen. Use the CH1 and CH2 POSITION knobs to set the cursors at
the top and bottom of the square wave. Read off the exact voltage from the display. Now
select Type = Time and adjust the vertical cursors to find the exact period of the square
wave. Select Type = Off to disable the cursors. The cursors may come in handy later on
when measuring the amplitude or width of an AP or a MEPP. Turn the cursors off for
now.
The scope can take automatic measurements of many displayed signals including
frequency, period, amplitude and rise-time. Push the MEASURE button (in the MENUS
area). Push the top option button. Select Source = CH1, and then Type = Pk-Pk for peak
to peak amplitude. Note that the Pk-Pk measure can be fooled by noise spikes so be
careful when using this function to measure noisy signals. Push the second option button
and set Type = Freq for signal frequency. The amplitude and frequency of the displayed
signal are now shown on the screen. You can have up to five automatic measurements at
one time.
Push TRIG MENU button located in the TRIGGER area, then the Slope option button.
Observe what happens to the signal with each push of Slope. Can you explain what you
see? Note that the location of the trigger is marked by the arrow at the top of the display.
Because this is a digital scope, you can display the waveform before and after the trigger,
depending on where you place the trigger using the HORIZONTAL POSITION control.
Under TRIG MENU, push the Mode button and select Normal. Spin the trigger LEVEL
control while watching the trigger level arrow at the right of the screen. What happens
when the trigger level goes above the displayed square wave. Can you explain what you
see? Keep the trigger level high and use the Mode button to select Auto. What happens?
Return to Normal mode with the high level so that triggering stops. Now push the SET
TO 50% button located in the TRIGGER area. This button automatically sets the trigger
level to about the half way point in your signal and is handy when you aren’t being too
14
fussy about triggering and just want to see something on the screen. Usually, you will
operate the scope in Normal mode. Auto (free running) mode is good when you are trying
to find the trace.
Single sweep
Sometimes you just want to capture a single trace, for example when you have
intermittent events such as MEPPs. To enable single sweep, press SINGLE SEQ, located
in the MENUS area. In this mode, the scope will wait for the next trigger event and
sweep once and only once allowing you to capture a single event on the screen. The top
of the display will show “Ready” when the scope is waiting for a trigger and “Acq.
Complete” when one trace is captured. Re-arm the scope by pressing SINGLE SEQ
again. Return to normal sampling by pressing RUN/STOP. Try capturing a single sweep
of your square wave.
Averaging
Averaging events is an excellent way to reduce the noise on the signal. Because the event
is synchronized to the trigger while the noise is random, with successive traces, the event
will add while the noise will cancel. The scope has a built in averaging function that you
will find very handy for getting clean recordings of MEPPs and APs. To enable
averaging, press ACQUIRE, located in the MENUS area, then the Average option button.
Use the Averages option button to select the number of events to average (4, 16, 64 or
128). The more sweeps the more noise goes away, but at a cost of taking longer to
acquire. Start the averaging process by pressing SINGLE SEQ. Averaging is complete
when the trigger message at the top of the display reads “Acq Complete.”
Return to normal display by pushing the Sample button.
Hint: Later in the week, if you notice signals on the scope that are changing slowly when
you don’t expect them to, it may be because your scope was inadvertently left in
averaging mode.
Printing
On the desktop computer, delete any files from last year that are in the ScopeConnect
folder located on the computer desktop. (You only have to do this once.) Start the
ScopeConnect app on the desktop computer. Click PRINT to transfer an image of the
current scope display, which will take about 20 s. The image will open in a viewing
program. Use Print in the viewing program to make a hard copy. To conserve paper, print
two copies of the image on one sheet of paper (this is an option in the viewing program),
one for you and one for your lab partner. Cut the paper to size and tape into your
notebook. In addition, the CAPTURE button on the ScopeConnect app transfers data
from the scope to a CSV file that can be opened with Excel. This might be useful for
analyzing data that you collect for your research project later in the week.
Important
After every print, use a marker to label the plot with a descriptive caption and the date
and time. In addition, make an “L” that lines up with one horizontal and one vertical
15
division, right along the graticule lines. Mark the horizontal line with the time (from the
SEC/DIV knob) and the vertical line with the voltage (from the VOLTS/DIV knob, plus
the gain of the preamp when measuring nerve signals), or force (when measuring muscle
force). Annotate interesting features on the trace. Appendix B has sample plots showing
how to mark the axes.
Deliverable 1
Get a good-looking trace of the calibration signal on the scope, print, add caption, date
and axes labels, and tape into your notebook as evidence that you are the master of your
scope.
Deliverable 2
Run the calibration signal into both CH1 and CH2 of the scope. Set the vertical gain of
CH2 to something different than CH1. Use the CAPTURE button of ScopeConnect to
transfer the scope data to a CSV file. Open the file with Excel and create a plot of the
data using straight black lines connecting data, no markers, white background and no grid
lines. Add appropriate axes labels and a title. Print the plot and tape into your notebook.
Hint
To reduce electrical noise in the scope signal, using the supplied cable, ground the scope
to the large metal plate located under the microscope. Black cable has alligator clip on
one end for attaching to the scope ground tab and a banana plug on the other end for
attaching to the plate.
Help! I can't get a trace
Here are some things to try if you lose your trace:
1.
2.
3.
4.
5.
6.
7.
8.
Set the trigger to Auto (automatic).
Ground the signal (GD button) and spin the position knob until you see it.
If in Normal mode, spin the trigger level control until the trace flashes across.
If in EXT trigger mode, make sure you have a good trigger signal by pressing TRIG
VIEW.
Push RUN/STEP and confirm trace is running by looking at the indicator in the top
left corner of the display area.
Push AUTOSET (top right). The scope will automatically attempt to set itself to
“good” settings.
Ask somebody at the next station.
Ask an instructor.
16
3.4
Working the stimulator
To excite the nerve, you will use the Dagan model S-900 stimulator (Figure 3-4). The
objective of this exercise is to become familiar with controlling the Dagan. The Dagan is
capable of producing one pulse or a train of pulses at each timed interval.
Figure 3-4: The Dagan stimulator sits above the Grass stimulator.
NOTE: Those who handled the scope controls for the scope exercise should now
tie their hands behind their backs and duct tape their lips and let someone else
take over for this exercise.
The Dagan controls
Skim this section quickly so that you can get to the stimulator deliverables as quickly as
you can. All of this will become clear when you view the pulses on the scope while
playing with the controls, You can always come back to this section to use it as a
reference.
Many of the controls have a selector switch with a black vernier dial just above. The
selector switch sets the base value. The vernier adjusts the setting to be between 0 (fully
CCW) and 100% (fully CW) of the base value. For example, if you set the pulse width
selector to 0.1 S and the vernier to 50 (its midpoint), the actual pulse width will be 50
mS.
17
Figure 3-5: Pulse interval
controls.
Figure 3-6: Pulse delay, train,
and pulse width
controls.
Figure 3-7: Output controls.
PULSE INTERVAL Section
The controls in the PULSE INTERVAL section (Figure 3-5) set the timing between
pulses or between pulse trains. The selector switch determines whether the pulses are
output continuously (CONTINUOUS setting) or just one at a time (SINGLE setting).
When in SINGLE mode, a pulse is emitted each time the INITIATE button is pressed.
The SECONDS knob sets the interval between the start of successive pulses or each
successive pulse train when in train mode. The selector knob sets the base rate (from 10
mS to 1000 S) which is scaled by the black vernier knob directly above. For example, if
you set the selector knob to 1, the vernier knob will scale the interval between 0 and 1
second.
PULSE DELAY Section
The PULSE DELAY section (Figure 3-6) allows you to initiate a pulse or a pulse train a
fixed time away from when the pulse is synched. This is useful if you want to position
where an event appears on the scope trace. It is also used to set the spacing between
double pulses. Normally, you will set the SECONDS knob to .001 and spin the vernier
fully CCW to keep the delay at 0.
TRAIN Section
The TRAIN section (Figure 3-6) enables the stimulator to emit a train of pulses at each
time interval. If you want just one pulse, leave the train switch off. If the switch is on,
DURATION controls the length of the train. As you increase DURATION, you will see
more pulses in your train. PULSE INTERVAL controls the interval between each pulse
within the train. The width of each pulse is set by the controls in the PULSE WIDTH
section. The time interval between each train is set by the controls in the PULSE
INTERVAL section.
PULSE WIDTH Section
The PULSE WIDTH section (Figure 3-6) sets the width of each pulse. If the DOUBLE
PULSE switch is on, two pulses are emitted each time with the interval between them set
by the Pulse Delay control.
18
OUTPUT Section
The OUTPUT section (Figure 3-7) is where you set the amplitude of the pulses and has
BNC connectors to bring signals to the stimulating electrode and scope.
You adjust the strength of the pulse using the 1V/10V switch and the LEVEL knob. If
you set the switch to the 1V position and the LEVEL control to 50, the output pulse
amplitude will be 0.5 V. Same thing when the switch is in the 10V position except now
LEVEL scales the output between 0V and 10V.
The POS BNC is the main stimulus pulse output. The SYNCH BNC provides a brief
pulse of fixed amplitude that is used to trigger the scope. If instead you used the POS
output to trigger the scope, when you changed to a very small amplitude you would lose
your trigger. Also, when delivering a train of pulses, SYNCH delivers just one triggering
pulse at the start of the train.
The MONITOR BNC delivers an exact copy of the stimulus train, but at a fixed
amplitude. This is handy for checking the timing of a complex pulse train on the scope.
The PULSE/CONTINUOUS switch should always be in PULSE mode. The other mode
is for emitting a constant voltage. Likewise the NORMAL/Q-STEP switch should always
be in NORMAL.
It is possible that the knobs on the Dagan unit at your station are not calibrated. For the
most accurate settings, connect the stimulus output to the scope and adjust the Dagan
knobs to the proper settings while viewing the output on the scope.
Initial setup
To view the stimulus pulses on the scope, connect the outputs of the stimulator to the
scope using BNC cables as follows:
1. SYNCH to scope EXT TRIG
2. POS to scope CH 1
Set scope triggering (push TRIG MENU) to Type = EDGE, Source = EXT, Slope =
RISING, Mode = NORMAL. The display will be blank until the stimulator actually
delivers pulses. If you lose track of the display, temporarily set the trigger to Auto mode.
To get a set of pulses on the screen, set the Dagan controls as follows:
1. PULSE INTERVAL section: CONTINUOUS, SECONDS knob = 0.1, vernier = 20
(for pulse interval of 20 mS)
2. PULSE DELAY section: SECONDS knob = .001, vernier fully CCW (zero delay)
3. TRAIN section: Switch = OFF
4. PULSE WIDTH section: DOUBLE PULSE switch = OFF. SECONDS knob = .01,
vernier = 50 (for pulse width of 5 mS)
19
5. OUTPUT section: three switches to PULSE, 10V and NORMAL. LEVEL vernier to
50 (for 5 V pulse amplitude)
Set the scope sweep speed (SEC/DIV knob) to 5 ms per division. Check that you have
taken the scope off Averaging mode (ACQUIRE then the SAMPLE option button then
RUN.) Spin the trigger level control of the scope for a stable trace (or press the SET TO
50% button) and adjust the amplitude and position of CH 1 for a nice looking trace. If
you are successful, you should see a set of pulses that are 5V high and 5mS wide with 20
mS between start of each pulse.
Like what you see? Print it. Did you label the printout?
Note: If your pulses are sagging rather than being crisp and square, the likely cause is that
CH 1 is AC coupled. Change to DC coupling.
Playing with the stimulator controls
Play with the INTERVAL, PULSE WIDTH and LEVEL controls (knobs and verniers)
until you have figured out exactly what they do.
Note: if one or more of the red lights on the stimulator goes on, you have dialed up an
incompatible set of stimulation pulse parameters. Typically this occurs when you set a
pulse width that is longer than the pulse interval.
Use the PULSE DELAY controls to position the pulse anywhere on the scope screen. Set
the PULSE DELAY SECONDS knob to .1 and spin the vernier. You can also set the
location of the pulse on the screen by leaving the delay off and setting the horizontal
position of the scope trigger using the scope controls. Note that when the PULSE
DELAY is active, the scope is triggered with the stimulator SYNCH signal, but the actual
pulse comes at a later time as dictated by the PULSE DELAY settings.
Change the pulse width to 1 mS. Flip the DOUBLE PULSE switch up. Set the PULSE
DELAY selector to 0.01 and spin the PULSE DELAY vernier until you understand how
the double pulse feature works.
Note: Now is the time to switch roles and have a new member of the group
manipulate the controls. Everybody else? Tie your hands behind your back!
Turn double off and set the delay to zero. Set the main PULSE INTERVAL controls to
20 mS and set the pulse width to 1 mS. In the TRAIN section, set the switch ON and play
with the DURATION and PULSE INTERVAL controls until you understand how
everything works.
Hint: See Appendix C for pictures showing what each stimulator control sets for
particular stimulus waveforms
20
Deliverable #1: Print a hardcopy scope image of a train that looks exactly like
Figure 3-8. Trim the hard copy and tape into your notebook. Did you add a
caption, the data and axes labels?
0.5 V
1 mS
2 mS
20 mS
Figure 3-8: Make yours look like this pulse train.
Deliverable #2: Set up your scope and your stimulator so that each time you
push INITIATE on the Dagan, one pulse is captured on the scope screen. The
pulse should have width 20 mS, should start 20 mS from where the scope is
triggered (hint: use the Dagan pulse delay function), and should be 6 V in
amplitude.
Find yourself with extra time? Give your lab partner a challenge. Draw a complicated
looking pulse waveform on paper and have your partner reproduce it with the stimulator.
Reverse roles. Continue until all members of your group are comfortable with the
stimulator and scope.
Note that sometimes getting comfortable with experiment apparatus means shooing your
partner and instructors away so that you have the equipment all to yourself without
anyone looking over your shoulder. Don’t be afraid to say, “Go away please!” Or, come
back when the lab is empty. In Itasca, the lab is open 24/7.
Your comments on and corrections to this manual are always welcome. Tell an instructor
so that the next version of the manual can be even better.
21
3.5
Using the electrode preamplifier
The membrane voltages you measure are small and must be amplified to be seen on the
scope or recorded on the computer. Because the microelectrode is so small, it has very
high input impedance (resistance) which means that special amplifier instrumentation
must be used to get a clean, noise-free signal.
The lab has the Dagan 1X2-700 intracellular preamplifier to condition the signal. The
preamplifier is located at the very top of the equipment rack (Figure 3-9). It is sometimes
referred to as a bridge amplifier. Along with amplifying the electrode signal, the Dagan
unit can inject a small signal through the recording electrode to measure its impedance.
Measuring the recording electrode impedance (Z) is the best way of checking that you
have a viable electrode. Get in the habit of doing this "Z-check" often, particularly when
you are having trouble getting clean membrane voltages.
Figure 3-9: Dagan electrode preamp.
The voltage signal generated by the
membrane is detected by the micropipette
electrode. The electrode is connected to a
head stage that has a gain of one and a
high input impedance to eliminate signal
distortion. The output of the head stage
goes to the Dagan preamp where the
signal is amplified and (optionally)
filtered. The output of the preamp goes to
the scope for viewing or to the computer
for digitizing or both. A block diagram of
the complete recording system is shown in
the figure to the right.
Dagan preamplifier
Electrode
Head
stage
Oscilloscope
Cell
The amplified outputs delivered by the
Dagan preamp are 10 Vm (gain = 10) and
100 Vm (gain = 100). For resting
membrane and action potential recordings
you will be using the 10 Vm output. For
MEPPs, you will use the 100 Vm output.
Figure 3-10: Set-up for recording
membrane potentials.
22
Remember these gains when interpreting the voltage levels of biopotential traces you see
on the scope, and always add vertical and horizontal calibration marks on your printouts.
For the vertical mark, it should indicate the voltage at the cell, not the scope voltage..
The MONITOR numeric display at the top center displays the signal in mV (gain = 1).
Dagan preamp controls
Figure 3-11 through Figure 3-13 show the controls for the Dagan preamp.
Figure 3-11 Power and channel
controls
Figure 3-12 DC current, step
current controls and
output BNCs.
Figure 3-13 Readout LCD.
The CHANNEL 1 section has the OFFSET knob for zeroing the readout voltage, the
BALANCE vernier used for an alternative method to find impedances, the Z TEST
switch for testing electrode impedance and the Vm FILTER selector knob for setting the
cut-off frequency of a low pass filter (leave at 10K).
The DC CURRENT and STEP CURRENT sections are used for the alternative methods
to find impedance and for iontophoresis procedures. Both DC CURRENT and STEP
CURRENT switches should be kept in the OFF position. Below these sections are the 10
Vm and 100 Vm output BNCs.
The MONITOR section has the display LCD. When V1 is selected, the readout shows the
average electrode voltage in mV. The meter updates approximately once per second and
is too slow to show the rapid voltage changes of action potentials or MEPPs.
23
3.6
Using microscopes
The Olympus Model SZH10 microscope used in the lab is shown in Figure 3-14. Adjust
the distance between the two eyepieces to give you a single, circular image. Control
knobs allow you to zoom and focus. Sometimes you have to go back and forth between
the two to get a clear picture. If you can't figure out where you are, zoom out and center
the region of interest before going back in. The closer you zoom in, the more light you
need so if the field appears dim, adjust the gooseneck microscope lamps. To prevent
overheating the tissue, turn off the gooseneck lamps when you take a break.
Figure 3-14:
Microscope for viewing nerve and muscle.
Knobs on the frame let you position the scope over your prep. Be VERY careful in using
these knobs. THINK before you loosen, and always support the scope with your other
hand. The last thing you want is for the scope to come crashing down causing damage to
itself and to your prep. These scopes are VERY expensive.
Exercise: Use the Olympus scope to look at your fingernail and skin around your
nail.
A dissecting scope is used for the frog prep. Although it has less magnifying power, the
advantage of a dissecting scope is the greater distance between the lens and the work area
making it easier to use tools.
Note: 1. When not in use, turn off the scope lights using the intensity knob (not
the switch). 2. Cover your scope when not in use; dust is the enemy!
24
3.7
Simulated cell
Before using a real electrode and real cell, you will be measuring the impedance of a
simulated electrode/cell system constructed from a small network of resistors and
capacitors. This circuit is called the "model cell" and is a small piece of hardware that
plugs directly into the head stage in place of the micropipette. A diagram of the model
cell is shown in Figure 3-15. Photographs of the model cell and head stage are in Figure
3-16 through Figure 3-18.
In the model, Rm is the membrane resistance for ions through channels. Typical values
range from 10 M-ohms to over 100 M-ohms. Cm is the membrane capacitance due to the
insulating (dielectric) properties of the membrane with charge conductors (ionic
solutions) on either side. All cell membranes have a capacitance of 1 micro-Farad/cm2.
Myelinated cells have a slightly lower capacitance because the layers of myelin leads to
greater separation of charges. Re is the resistance of the recording electrodes. For micro
pipette electrodes used in the lab, Re for a usable electrode is 20-60 M-ohms.
Figure 3-15: Real and model electrode/membrane.
Figure 3-16 Model cell
Figure 3-17 Head stage
25
Figure 3-18 Model cell coupled
to head stage.
3.8
Checking electrode impedance
Checking the impedance, a "Z-check" is the best and only way of determining if your
recording microelectrode works. A good electrode has an impedance (Z) of 20-50 Mohm.
A higher impedance means the tip is too small or the electrode is clogged or there is a
poor electrical connection. A lower impedance means the tip is too large or the tip is
broken. This section takes you through the steps of measuring electrode or model cell
impedance.
The preamp has several methods for checking electrode impedance. You will use the ZTest method, the simplest. When in Z-Test mode, the preamp injects current pulses of
peak-to-peak amplitude 1 nA (nano or ten to the minus ninth amp) into the electrode. By
Ohms Law (V = I*R), the voltage across the electrode will be 1 mV for every M-ohm of
resistance. For example, if the test shows a voltage waveform with peak-to-peak
amplitude 20 mV, your electrode has 20 M-ohms of resistance.
Connections
Get a model cell and a N=1 (voltage gain of 1) head stage. Connect the head stage cable
to the connector on the back side of the Dagan preamp. Connect the 10 Vm output on the
Dagan to CH 1 on the oscilloscope. Set the scope to trigger off CH 1. Attach the model
cell to the head stage. On the model cell, plug the lead into the black side to short out the
membrane and into the red side to short out the electrode. Also, ground the scope to the
large metal experiment plate.
The membrane voltage in mV is displayed on the LCD digital voltmeter in the center of
the Dagan preamp (useful for precision measurement of DC voltages), and on the
oscilloscope (useful for transient voltages such as the action potential). With the N=1
head stage and the 10 Vm Dagan output, the voltage you read on the scope will be 10
times the voltage seen in the prep and 10 times the voltage displayed on the LCD.
Conducting a Z-Test (model cell or microelectrode)
1. If you are using the model cell, short out the simulated membrane resistance so that
you are just looking at the simulated electrode. Connect either the model cell, or the
glass micro pipette electrode to the head stage, connect the head stage to the preamp,
and connect the 10 Vm output of the preamp to CH 1 of the scope. Set CH1 to 200
mV/div. If you are using a glass electrode, lower into the saline bath.
2. Zero the preamp by spinning the OFFSET knob until the LCD display reads zero.
3. On the preamp, turn off the GATE switch for the step current feature and turn on the
Z TEST switch.
4. Measure the peak-to-peak amplitude (in mV) of the the square wave that appears on
the scope screen. This is easiest to do by eye straight off the scope screen. The scope
automatic peak-peak measuring tool gets fooled by noise spikes and the cursors take
26
too long to set. The electrode impedance is proportional to this amplitude by the ratio
of 1 megohm for every millivolt of amplitude (1 M-ohm/mV).
Note: Because the signal comes to the scope from the 10 Vm output of the preamp, the
voltage you see on the scope is 10 times the voltage at the electrode. Thus if you see 500
mV on the scope, it is 50 mV at the electrode and the electrode impedance would be 50
M-ohm. Also, check that the scope, Chan 1 probe setting is set to x1 (see scope section).
Noise! There may be 60 Hz noise riding on top of the scope signal from the model cell.
This is because the cell is acting like an antenna, picking up 60 Hz electromagnetic
interference from wall-powered lab equipment. Or there may be other noise on the signal.
Now is a good time to minimize the noise. Your goal is to get the noise level at or below
0.5 mV. Try these steps: (1) Rest the head stage on the metal experiment plate and move
you and your hands away. (2) Ground the scope and the microscope to the plate. (3) Cup
one hand around the head stage, without touching it, and touch the plate with your other
hand. This makes you a ground shield surrounding the head stage. (4) Make an aluminum
foil tent that surrounds the head stage, without touching it. Use an alligator clip lead to
ground the tent.
Note: On the preamplifier, adjust the CAP COMP knob. Start all the way CCW. Do a Ztest on the model electrode. Turn the knob CC until the tops of the Z-test square waves
become flat when viewed on the scope.
Appendix J describes other ways of measuring electrode impedance, and also a method
for separately measuring membrane and electrode impedance.
Note: Head stages are picked to match the electrode impedance. The N=1 head stage
is for electrodes of 2-50 Mohm. The N=0.1 head stage is for 20-500 Mohm. Both
have a voltage gain of 1, but scale the current. For this week, only the N=1 head stage
will be used.
Deliverable
Find the value of Re, the resistor in the model cell that simulates electrode impedance,
while shorting out the model membrane resistor Rm. Look up the resistor color code (see
Appendix L) of the Re resistor to confirm your result.
Reminder: Have you been good about using your lab notebook as a running journal,
including recording information such as the resistances in the model cell?
Suggestion: Keep a model cell at your station along with a banana-to-banana cable for
shorting the model cell. Use the cell to troubleshoot your head stage and preamplifier.
27
3.9
Making a glass microelectrode
Membrane voltages are measured with a glass pipette microelectrode. The electrode is
heated and pulled to a fine point in a special, precision machine (Sutter Instruments
Model P-97 micropipette puller, www.sutter.com). Because the electrodes are easily
broken, you will become skilled in fabricating new ones.
You can't see the tip of the microelectrode, even under the microscope because it is
pulled to less than 0.1 microns in diameter. The only way to estimate the tip diameter is
by measuring the electrode impedance.
While one person is pulling the first electrode, the other should prep the lab station.
Arrange a prep dish in a holder under the microscope. Place a micro manipulator nearby.
Clamp a N=1 head stage in the manipulator (Figure 3-22). Fill a squeeze bottle with frog
Ringer’s. Fill the prep dish with Ringer’s.
Note: Because the puller is an expensive instrument, have someone show you how to use
it before attempting your first pull.
Here are some tips for making electrodes:
•
For pipette stock, use Dagan (Prisim) model FSG 12 or Sutter model BF120-6010 (O.D. = 1.2 mm, I.D. = 0.6 mm)
•
Use Program #3 on the Sutter puller (heat = 490-550 2, pull = 100, velocity = 30,
time = 150). If the wrong program is showing, push RESET, then 3 and ENTER
at the program prompt.
•
To pull, place pipette in the guide grooves, release the latch and slide right side in,
fix the pipette to the right side, draw both sides tight using the finger pieces,
tighten the left side. Check that pipette is centered, then close the cover. Start the
pull by pressing PULL.
•
After pulling, fill electrode with 3 molar KCl, using the syringe-mounted filler 3.
Avoid introducing bubbles that might cause poor contact with the electrode.
Store the filler in its protective sheath. Every few hours and at the end of the day,
flush the filler with distilled water to prevent the KCl from crystallizing and
clogging the tube.
2
The Sutter heat setting depends on the heating element installed. Typically, heat values for the FSG 12
pipette stock are between 490 and 550. If the heat is too low, the electrode cannot be pulled. If the heat is
too high, the tip will be too slender and will break. The instructors will set and lock the proper heat.
Instructors: See Appendix K for how to program heat settings.
3
WPI Microfil, 34 AWG, PN MF 34G-S. Attach to a 6 cc syringe.
28
•
Take an electrode holder (Figure 3-19), dip in chloride bleach solution for 5 sec.
(Figure 3-20), then rinse in distilled water.
•
Thread the electrode wire down into the filled micropipette and tighten the
electrode holder around the pipette. Do not overtighten.
•
If you brush the tip of the electrode against anything, it will break.
•
Bleach and rinse a ground electrode (Figure 3-21).
•
Electrodes have about a six hour life before the KCl crystalizes and clogs the
opening.
Figure 3-19: Pulled pipette and
electrode holder.
Figure 3-20: Bleaching the
electrode.
Figure 3-21: Ground electrode
.
Setting up a fresh electrode for testing
•
Half-fill a clean 9 cm Sylgaard culture dish with frog Ringers. Place on stand and
anchor to stand with dab of clay
•
Anchor the spade terminal end of the ground electrode to the head stage. Use
modeling clay (not too much; it’s a pain to clean up) to hold the electrode
submersed in your prep dish filled with Ringer's. Keep the metal ring that
surrounds the prep dish dry and ensure that the ground electrode does not touch
the ring.
•
Connect the electrode holder to a N=1 head stage and then clamp the head stage
into a micromanipulator. While viewing through the microscope, lower the tip of
the electrode into the Ringer’s solution. Shining the microscope lights up through
the bottom of the prep often provides a better view.
•
Figure 3-22 shows a complete setup for measuring using the micropipette
electrode.
29
•
Check the electrode impedance (see Section 3.8). A good electrode has an
impedance of 20-60 Mohm.
In summary, the order is: (1) pull, (2) fill, (3) bleach electrode and rinse in distilled water,
(4) thread into pipette, (5) Z-test.
Figure 3-22: Microelectrode connected to head stage and lowered into a saline bath. The ground electrode
is also in the bath. Note the angle of insertion which is about right for stabbing muscle cells
If the preamp does not zero: Remove the glass electrode from the holder and stick the
silver wire of the electrode straight into the Ringer’s solution in the prep along with the
ground electrode. Still can’t zero? Try touch the ground wire to the electrode wire. If bad,
there is a loose connection somewhere in the ground wire, the electrode holder or the
head stage.
Tip: Anchor the ground wire with care. Place a small square of lab tape on the plastic
prep dish holder for insulation. Anchor the ground wire with a ball of modeling clay. Run
the ground wire up and over the metal retaining ring for the prep dish and place the tip
into the Ringer’s. Make sure the ground wire does not touch the retaining ring and that
there are no wet or dry salt bridges that could short the ground wire to the apparatus.
30
Tip: Set up the micromanipulator so that the electrode moves up and down at a steep
angle. The angle should be like this
not like this
Deliverable: Make a glass microelectrode. Use the Z-check method to find the
impedance of the electrode. Continue making electrodes until you get one in the 20-50
Mohm range. Report your Z to an instructor
Deliverable: Measure the impedance of an electrode. Purposely break the tip. (Can you
now see the end under the microscope?) Measure the new impedance. Explain what you
observe.
Deliverable: Practice your micromanipulator skills. Pretend you have a muscle prep in
the dish (Figure 3-25). Practice moving the electrode into a cell, back out, over to a new
cell and back in. Then practice going in the orthogonal direction along the line of a cell.
Watch through the microscope at all times. Becoming adept at this skill will save you
time later.
Clean, Quiet Recording Signals
Now, work to get your electrode signal so that it has the lowest electrical noise of any
group working in the lab. You goal is a super-quiet signal with no ripples. With the
electrode in Ringers, fuss with ground leads and positioning equipment until you reach
your goal. Measure and record the peak-to-peak voltage of the noise. You should be at -.5
mV p-p or better. Compare with other groups. Is yours the best?
Deliverable (if you have time): Measure the membrane resistance Rm and
membrane capacitance Cm of the model cell. Appendix J has the details.
Congratulations! You have survived day #1 of the course.
Find yourself with extra time? Go back and do the optional exercises you skipped.
Tonight: Helpful if you (1) read ahead in the manual, (2) read the lecture notes on the
resting membrane potential, (3) read the Adrian and Magleby articles in module reading
list.
31
3.10 Pithing the frog
(This section is adapted from "Experimental Neurobiology: A Laboratory Manual" by Oakley and Schafer,
University of Michigan Press, 1978.)
"Pithing" means to sacrifice an animal by destroying the spinal cord. "Double pithing"
means destroying both the brain and the spinal cord. Pithing is as humane as injecting an
anesthetic, provided it is done rapidly and firmly.
Method
(Note: Your instructors may teach you an alternate method for pithing.)
Hold the frog in one hand just under the shoulders with the arms inside your grip. Grip
firmly but without excessive squeezing.
Stun the frog by whacking on the head with the heavy metal ruler or the pipe.
Open a pair of blunt, heavy dissecting scissors and work one side inside the mouth going
as far in as you can. Position the scissors so that the other blade is posterior to (behind)
the eyes.
Using a swift motion, cut off the upper part of the head. This disconnects the brain from
the rest of the animal.
Scramble the brain (the part you cut off) with the pith needle.
Find the exposed spinal cord and run a dissecting needle down the spinal canal. The
needle will meet some resistance, but force the needle into the canal while twisting until
the frog's hind limbs hyperextend forcefully. This is the critical sign because at this point
you have completely destroyed the spinal cord. If the legs fail to extend, probe for the
cord again.
The frog will be completely limp. Lay in a pan and wait a few seconds. Test for the
withdrawal reflex by pinching the largest toe of one hind foot between your fingernails or
with forceps. If a withdrawal reflex is elicited, repeat the spinal pithing procedure.
Rinse your tools, and dispose biological tissue properly.
Curious about where the frogs are from?
Northern Grass Frogs (Rana pipiens pipiens)
Medium size, 2-1/2” – 3”
Item # L 5300, $4.50 each
Connecticut Valley Biological Supply Company
http://www.ctvalleybio.com/
32
3.11 Dissecting the cutaneous pectoralis muscle
Note: Consult the frog dissection video if things get confusing. The video is loaded on each lab station
computer. However, we prefer that you dive right in and do, rather than watch.
Suggestion: While one person dissects, the other should get the culture dish ready,
should pull and Z-test a microelectrode, should read Section 3.12 and get the equipment
ready for measuring membrane potential.
Equipment: dissecting tools, dissecting tray, large pins, clean 9 cm Sylgaard culture dish,
large and small insect pins.
Sacrifice the frog by pithing. Place on a dissecting tray, ventral side up. Extend the limbs
and use T-pins to attach to the tray. Move the prep over to the dissecting microscope and
arrange yourself, the scope, the light and your tools for clear viewing and easy access.
During the operation, soak the frog with Ringers (Appendix D) on a regular basis to keep
the tissues from drying.
Your scissors and forceps are very delicate. Do not let the tips touch each other. Rest
your wrists while dissecting. You will have better control with less tremor and your arms
won’t get tired.
Visualize the origin of the cutaneous pectoralis (CP) which makes shallow dimples in the
skin near the level of the clavicle marking where the CP connects to the skin. Be careful
not to cut through the CP when you make skin cuts.
With small scissors and fine forceps (#5), cut the skin down the midline (Cut 1 in Figure
3-23).
33
3
1
2
CP attachment
5
Figure 3-23:
4
Skin cuts for removing the left CP muscle.
Make a small access incision in the armpit area (Cut 2). Then do a lateral cut to the center
line (Cut 3) making sure you are just above the CP attachment point by working the
scissors under the skin to the CP attachment before cutting.
Make vertical Cut 4 along the lateral side down to the abdomen. For this cut, go as far
lateral as you can to avoid damaging the muscle. If you can, make the cut to the lateral
side of the large muscle that runs top to bottom along the side.
Lateral Cut 5 should be just below the CP attachment (work your scissors up under the
skin) after which you will be left with a small rectangular patch of skin that holds the
muscle.
Gently lift the skin patch by its superior edge and flip it down. You should be able to just
see the start of the CP muscle which is a thin sheet containing visible longitudinal
strands.
Using blunt dissection from the superior side, gently work the skin and the CP muscle
loose from the underlying tissue. Take care and use the fine scissors to scrape and cut the
connective tissue as you lift. Connective tissue appears transparent and homogeneous in
texture, unlike the muscle whose fibers form visible strands. Do not cut the muscle.
34
Continue the dissection of the muscle along its medial edge down to its insertion on the
xyphoid process, the small piece of bony cartilage on the midline at the posterior end of
the sternum. You may see a faint white line where the muscle inserts.
Next, free up the nerve. The nerve enters the muscle along its lateral edge, about midway
between origin and insertion (Figure 3-24). Blood vessels can be distinguished by their
dark pigment while the nerve is shiny and white. Be very careful during the dissection not
to damage the nerve. To fully expose the nerve, note the long narrow muscle lateral to
and thicker than the CP. The nerve runs close to this muscle and into the axilla (armpit).
Use blunt dissection to expose an opening into the axilla. To enlarge the opening, cut
through the lateral muscle, first ensuring you are cutting neither the CP nor its nerve.
Trace the nerve back from the CP into the axilla. Sever the nerve at a point as far from
the CP as you can, being careful not to cut the blood vessels that run near it.
Once the nerve is free, fold it on top of the CP, then gently dissect the CP muscle free
along its lateral edge
Cut through the sternum laterally anterior to the muscle insertion, going no further than
midline. Then turn your scissors and cut down right along the midline, continuing
through the xyphoid until the xyphoid is free. If you have done this correctly, you will
end up with a small piece of cartilage, free from the body that contains the CP insertion
point. If you stayed on the midline, you will also be able to come back later and use the
CP on the contralateral side.
skin flap
nerve
muscle
xyphoid process
Figure 3-24: Muscle, skin flap, xyphoid process and nerve. All of these
should come with your preparation when you remove it from
the body.
Once the xyphoid is severed, cut down and around to free the entire muscle/nerve
assembly from the body. Take along a generous chunk of distal connective tissue from
the belly; it can be cleaned once the prep is in the dish.
Remove the entire muscle/nerve assembly and place in a clean 9 cm Sylgaard culture
dish. Drape the rest of the frog with a Kim wipe, soak with Ringers and store in fridge.
35
With the prep skin flap down, anchor to the dish first using large (#0 or #1) insect pins,
then replacing with tiny pins that won't get in the way of your electrodes. Place pins in
the top two corners of the skin flap and then one or two through the xyphoid process,
stretching out the muscle. Stretching the muscle tight puts you on the back side of the
muscle length-tension curve which means the muscle will move less during stimulation.
A tightly stretched prep also makes it easier to impale the cells with the microelectrode.
Advice: Stretch your prep tight.
Clean off excess connective tissue to that you have a clear sheet of muscle.
Use one more pins to stretch out the nerve, taking care not to pull it taut. When you are
done, you should have something that looks like Figure 3-25.
Important: When the prep is ready, using a disposable transfer pipette, suck the Ringer’s
out of your dish and replace with fresh.
A good prep should last four hours. A great prep can last 24 hours. If you leave your prep
for more than 60 minutes, wet and store in the refrigerator.
Figure 3-25: A finished prep, all pinned out and ready for an experiment.
Place the prep in the holder stand under the Olympus microscope.
36
Hint: Position your scope lights so they shine up from under the prep. Generally, this
makes it easier to see and it keeps the lights out of the way.
Zoom in to examine the muscle fibers, nerve branches and end plates. Do you see any
damage or are the fibers in good shape?
Cleanup
Cover the frog body in a tissue soaked in Ringers. If both muscles have been removed
from the frog, remove the frog from the dissection tray and place in a plastic bag marked
for biological tissue waste. (At the end of the day, make sure that bag gets sealed, labeled
and placed in the appropriate freezer.)
Required: At the end of the day, clean your tray, pins and surgical instruments.
37
3.12 Measuring membrane potential
Prepare the standard CP muscle prep, all pinned out, and bathe in Ringer's. For this
experiment, only use sections of the muscle that are not nicked or torn. Each muscle cell
spans the length of the whole muscle. If the muscle is cut at any point, the cells will be
cut in half and will not hold the normal membrane voltage.
Prepare a glass micro recording electrode, connect to a head stage and mount in a micromanipulator. Place the ground lead from the head stage into the bath and anchor with
some modeling clay. Lower the recording electrode into the solution and do a Z-check to
confirm proper electrode impedance of 20-50 Mohm. Figure 3-26 shows this setup.
With the electrode in the bath, use the OFFSET knob on the bridge amplifier to zero the
voltage.
Help, I can’t zero the voltage! Try these steps: (1) Confirm that ground lead tip is in
the bath. (2) Remove the electrode from the bath. Remove the electrode holder from the
head stage and replace with a model cell. Confirm that you can zero the signal. If can’t,
consult an instructor. (3) Clean ground lead with emory cloth. (4) Confirm that ground
lead is secure in the head stage. (4) Pull a new electrode.
38
Figure 3-26: Prep to measure membrane voltage. Glass microelectrode is connected to a head stage
and mounted in a micro manipulator. Ground lead from the head stage goes into the bath
and is anchored with a dab of clay. Microscope lamp can either shine up through the
bottom or down from the top. Experiment to see which works best.
Scope settings (Figure 3-27):
• 10 Vm preamp output to CH 1
• 100 Vm preamp output to CH 2
• CH 1 vertical gain = 200 mV/div (because of the 10 times gain in the preamp, this
means that each division on the scope is 20 mV of electrode voltage)
• CH 2 vertical gain = 100 mV/div (so that each division on the scope is 1 mV of
electrode voltage)
• CH 1 is DC coupled
• CH 2 is AC coupled
• Trigger mode = AUTO, Trigger source = CH 2
• Sweep speed = 5 mS/DIV
• Adjust the vertical position of the CH 1 trace so that zero volts (the arrow labeled “1”
on the left side of the display) lines up with the second to the top graticule line
• Adjust the vertical position of CH 2 so that zero volts (the arrow labeled “2”) lines up
with the second to the bottom graticule line
39
TRIG
CH 1
CH 1 = 200 mV/DIV, DC
CH 2 = 100 mV/DIV, AC
TRIG
LEVEL
CH 2
SWEEP = 5 mS/DIV, TRIG MODE = AUTO
Figure 3-27: Scope display for seeing membrane resting potentials (CH 1) and capturing MEPPs
(CH 2).
Maneuver the microelectrode over the muscle. While viewing through the microscope,
slowly lower into a muscle fiber. Focus on the top layer of cells. You are inside a cell
when the voltage suddenly drops to about -90 mV.
Tip: Set up the micromanipulator so that the electrode moves up and down at a steep
angle. The angle should be like this
not like this
It is difficult to recognize the cells and their boundaries through the microscope. Until
this is mastered, one person should lower the electrode while a partner is watching the
scope display for the negative deflection. Try changing the position and angle of the light
for a better view.
40
Should the electrode enter too far or encounter mechanical resistance, the voltage may
become positive or read a small negative value like -25 mV. Going too far and ramming
the electrode into the bottom of the dish may clog (infinite resistance) or break (zero
resistance) the electrode. Replace with a new one.
If your membrane voltage slowly drifts up, your cell is leaking, most likely around the
electrode puncture site.
If you think you are poking cells, but are not able to get the -90 mV jump, check the
electrode impedance. Try pushing the BUZZ button on the preamplifier for one or two
seconds to free up a clogged electrode.
Deliverable: Record the membrane voltage in 10 different cells and document in your
notebook.
41
3.13 Miniature end plate potentials (MEPPs)
The synaptic transmitter acetylcholine (ACh) is released from the nerve terminal
(presynaptic terminal) and diffuses across the synaptic cleft to the muscle fiber
(postsynaptic terminal). ACh is stored in the presynaptic terminal in quantal packets
called vesicles. Each vesicle contains approximately 5,000 molecules of ACh. During an
action potential (AP) event, about 300 vesicles release their ACh that bind to receptor
channels on the postsynaptic terminal, opening ionic channels that result in the EPP.
Occasionally, in the absence of an AP, a single ACh vesicle is released. This spontaneous
release can cause a depolarization of the muscle membrane of approximately 0.5 mV.
This minute voltage change is called a miniature end plate potential or MEPP. MEPPs are
additive so if more than one vesicle is released at a time, the amplitude of the MEPP will
be in multiples of 0.5 mV. Figure 13-3 in the Magleby reading shows a MEPP recording.
Take a look at that figure now and read enough of the text so that you understand the
physiology behind MEPPs. MEPPs are often visible immediately following a successful
cell penetration, but only if the penetration is very close to an endplate.
Setup
Standard frog muscle prep.
Scope settings: Same as in Section 3.12. Set to trigger off CH 2. The voltage magnitude
of the MEPP is tiny which is why the CH 2 gain is large. CH 2 is AC coupled to
eliminate the slow voltage drift that occurs when recording for long periods of time. 4
To get a clean signal, you will have to minimize electrical noise in the system. If you find
noise is a problem, consult Appendix E for noise reducing methods.
Procedure
Impale a muscle fiber as near to an endplate as possible. The endplate is the very faint
oval structure seen at the distal end of the nerve where it forms the synapse with the
muscle. The endplates on a muscle tend to cluster in one region. Since the fibers are large
and have only one endplate, this may take some effort. But, if luck holds out, you should
hit an endplate in a few tries. Concentrate your search around the visible ends of axons in
the muscle.
Wait for MEPPs.
Once you have a reasonable frequency of MEPPs, use your scope to capture some nice
traces. Set the horizontal position so the scope trigger is in the middle of the screen (the
SET TO ZERO button is a convenient way to do this). Increase the CH 2 vertical gain so
4
AC coupling will slightly distort the shape of your MEPP. However, with DC coupling,
you would not be able to see the 0.5 mV MEPP on top of the 90 mV resting potential.
42
that your MEPPs will fill a good part of the screen. Set the sweep speed to 5 mS/DIV. Set
trigger mode to Normal, the trigger source to CH 2 and set the trigger level to just above
the noise level in CH 2. Now you should be able to capture every MEPP.
Push SINGLE SEQ to capture just one. Or, use the scope Averaging feature to average a
number of MEPPs to reduce the noise in the traces.
An example MEPP recording is shown in Appendix B.
Deliverable: Printouts of some nice MEPPs. Remember to add a caption and to label
the axes.
Hint MEPPs can be viewed on damaged fibers whose membrane potential is not at -90
mV. This means that you can use a heavily damaged prep with fibers that are severed to
hunt for MEPPs.
Challenges
These are potential research or late-night projects. Complete Section 3.14 before
attempting these.
1. Find the frequency of MEPPs occurrences in number per minute or number per hour.
You can slow down the scope trace and count by hand the number of MEPP’s you see
in one minute, or you can time how long it takes the scope to average 64 MEPP’s.
2. Extracellular Ca++ is needed at the nerve terminal to release ACh in response to a
presynaptic AP. Try replacing the normal Ringer's with Ca++ free Ringer's. What
effect does this have on the shape and frequency of MEPP's? For a research project
later in the week, you could examine how MEPP single, double and triple frequency
is changed by Ca++ concentration.
3. Curare is a nicotine specific blocker that will block the ACh receptors on the
postsynaptic membrane. Add curare to the bathing medium. What effect does this
have on the shape and frequency of MEPP's?
4. Increase the osmotic pressure gradient of the bathing solution by adding sucrose. A
drop or two of 1M sucrose solution should be sufficient. What effect does this have
on the shape and frequency of MEPP's?
43
3.14 Effect of K+ concentration on membrane potential
Introduction
In this exercise, you will reproduce the results report in Figure 5 of the 1956 J. Physiol.
article by RH Adrian (see the Readings section of the Lecture Notes). You will be testing
the hypothesis that changing the external potassium ion concentration changes the resting
potential across the membrane (Vm). The sign and magnitude of Vm is determined by the
relative permeability of ion species. The resting potential of a normal frog muscle cell is
approximately -90mV. You can calculate the theoretical approximation for this potential
using the Goldman-Hodgkin-Katz (GHK) equation
 P [ K ]o + PNa [ Na ]o 

Vm = ( RT / nF ) * ln k
 Pk [ K ]i + PNa [ Na ]i 
(Eq. 1)
where R is the universal gas constant (8.31 Joules/Moles*°K), T is the temperature (°K),
F is the charge per mole of electrons (9.6486E4 C/Mole) and n is the ionic valence (n = 1
for Na+ and K+). Pk and PNa are the permeabilities of K and Na through the membrane
and [K] and [Na] are the concentrations of K+ and Na+ outside (o) and inside (i). Since
the Cl- permeability is small, its gradient can be neglected in the GHK equation.
By considering just the ratio of permeabilities, b = PNa /Pk, and noting that RT/nF*ln(10)
at body temperature (310°K) is (8.31*310/96485)*2.3026 = 61.5 mV, then
 [ K ]o + b[ Na ]o 

Vm = 61.5 log10 
 [ K ]i + b[ Na ]i 
Normal reference values are [K]o = 2.5 mM, [K]i = 140 mM, [Na]o = 120 mM, [Na]i = 23
mM, b = .02. The GHK equation that relates membrane voltage in mV to the external
potassium concentration is
 [ K ] o + (.02)(120) 

Vm = 61.5 log 10 
 140 + (.02)(23) 
that approximates to
 [ K ] o + 2.4 
Vm = 61.5 log 10 

 140

(Eq. 2)
For normal potassium [K]o = 2.5 mM and using Eq. 2, this means the normal membrane
resting potential is Vm = -90 mV.
44
Exercise: Using Eq. 2, calculate the theoretical resting membrane potential Vm for the
following external concentrations of potassium: [K]o = 1, 2.5, 5, 10, 25, 50, 100 mM.
Write the answers as a table in your lab notebook. (Hint: Log10 in Excel is LOG10().
Note that when the external potassium level is 138 mM or greater, the membrane voltage
will reverse.
For this experiment you will measure Vm for seven levels of external potassium
concentration, and then plot the results to see if your data is similar to the data in Figure 5
of the Adrian article.
Quiz question: As you raise the external potassium concentration, will the membrane
voltage go up or will it go down? Why?
Setup
Standard frog CP prep with a glass microelectrode to measure membrane voltage.
Procedure
Standard [K]o solutions are available in the lab. Transfer small amounts of the
concentrations you will be using (see below) into a set of beakers, perhaps four
concentrations at a time so that the lab does not run out of beakers.
Bathe an intact CP muscle prep, all pinned out, in normal Ringer's ([K]o = 2.5 mM). Be
sure that the muscle is intact from origin to insertion. The muscle cells run end-to-end in
the CP so nicks and cuts will cause the cells to leak their internal solution and will not be
able to support a membrane voltage.
Stick a cell with the microelectrode and measure the membrane voltage, which should be
around -90 mV. Repeat for 10 cells. Record the data in your notebook.
Suck the solution out with a transfer pipette or a piece of small tubing connected to a
syringe and replace with a solution of the next [K]o. Repeat the rinse four times to be sure
all the solution is new. Allow five minutes for the prep to come to equilibrium before
taking measurements. Record the membrane voltage of 10 cells. Repeat for the next
concentration. Because you are sticking so many cells, it is likely that you will break a
few electrodes. For each new electrode, run a Z check.
Use these [K]o solutions in this order:
2.5, 1, 5, 10, 25, 50, 100, 2.5 mM
The last measurement at 2.5 mM, normal Ringer’s, is a control, taken to verify that the
tissue is still viable.
45
As you proceed, enter the data into your notebook and into the GHK Excel spreadsheet
located on the lab computers. The Excel application will plot the data along with a line
showing the theoretical level of membrane voltage.
If you are short on time, collect data from five cells per concentration rather than 10.
Deliverable: A printout of the GHK Excel plot showing the theoretical membrane
voltage curve and the average and standard deviation of your data. Don’t forget to mark
the printout with the date
Looking for something else to do? Measure the membrane resistance and capacitance
using the methods described in Appendix J.
46
3.15 Stimulating the nerve fiber with a suction electrode
Action potentials can be excited in the nerve branch using a suction electrode that sucks
the cut nerve bundle up into a tube in close proximity to a coiled, platinum wire
stimulating electrode. The electrode used in the lab is Model 728000 from AM Systems
(www.a-msystems.com) shown in Figure 3-28. You will be using this electrode for the
next several lab exercises. Be gentle; this electrode is fragile.
Figure 3-28: Suction electrode.
What you need
(1) Suction electrode. (2) Suction electrode cable (has BNC on one end and two banana
plugs on the other end.) (3) Dagan stimulus isolation box Model S-910.
Connect one end of the BNC cable to the suction electrode and the banana plugs (use
yellow and blue plugs) on the other end of the suction electrode cable to the front of the
stimulus isolation box. With a BNC cable, connect the BNC on the back of the isolation
box to the ISOLATOR channel on the Dagan stimulator The remaining connections are
discussed in the next section.
Using
Draw Ringers up into the electrode, sucking it up by spinning the syringe knob until you
see liquid just above the white plastic tip.
Clamp the business end of a filled suction electrode to a micromanipulator and position
over your prep with the electrode tip touching the nerve and the reference electrode (the
wire coiled around the outside) in the bath. Spin the knob to gently suck the end or side
of the nerve into the electrode (Figure 3-29). Have the nerve completely plug the tube so
47
that current cannot exit the tube without going through the nerve. The complete setup for
measuring APs with suction and recording electrodes is shown in Figure 3-30.
Figure 3-29: Suction electrode technique showing side suck of the nerve (left) and end suck of the nerve
(right).
Figure 3-30: Suction electrode in place for a membrane potential experiment. View of layout at right
Tip: If your suction electrode is broken, repair instructions are in Appendix I.
48
3.16 Motor unit action potentials
Objective: View the motor unit action potential (MUAP), the action potential (AP) that
travels down the muscle fiber in response to a nerve stimulation pulse.
Prep: Standard frog cutaneous pectoralis prep, but this time with an intact nerve. If you
can, when pinning, overstretch the muscle to minimize movement artifact.
Connections: (1) Stimulating electrode to the banana plugs on the front of the stimulus
isolation box using the yellow and blue plugs on the special BNC-to-banana cable. (2)
BNC on the back of the isolation box to ISOLATOR on the Dagan stimulator. (3)
MONITOR on the Dagan to CH 2 on the scope so that you can see the stim pulse on the
scope. (4) SYNC on the stimulator to EXT TRIG on the scope.
Stimulating electrode: Suck the nerve up into a suction electrode.
Recording electrode: Standard glass micropipette recording electrode on a
micromanipulator. Dagan preamp 10 Vm output to CH 1 on the scope.
Stimulator settings
1. Pulse interval: 1.0 sec
2. Train: off, Double pulse off, Delay = 0
3. Pulse width: 0.1 msec
4. OUTPUT LEVEL vernier knob: 0%
Isolator settings
1. Range knob: 100Volts. Leave on this setting and use the LEVEL knob on the Dagan
stimulator to control stimulus strength. The LEVEL knob full CCW applies 0 volts
and full CW applies 100 V. The 1V/10V switch on the Dagan has no effect. Also note
that the MONITOR output on the Dagan will not change in amplitude as you adjust
the LEVEL control. .
2. Polarity: Normal (+).
Scope settings
1. Sweep speed: 1 ms/div
2. CH 1 gain = 200 mV/div
3. CH 2 gain so that the stim pulse is about one division high
4. Vertical positions: CH 2 stim pulse trace along the bottom. CH 1 membrane voltage
trace lined up with a graticule near the top when CH 1 is at zero volts (GND
coupling)
5. Trigger mode = Normal
6. Trigger source = Ext
7. Trigger slope = (+)
8. Trigger level to get a reliable trigger on each stim pulse.
9. Horizontal position: adjust so that the action potential ends up in the middle of the
screen.
49
Procedure:
1. While viewing through the microscope, turn the stimulator amplitude up using the
LEVEL knob until you observe the muscle fibers just twitching with each stim pulse.
Don’t be afraid to spin the knob all the way to 100%. Flip the Normal/Reverse
polarity switch on the isolation box and leave it in the position that gives you the
biggest twitch. If you see twitches when the LEVEL control is at 10 or less, change
the range knob on the isolator from 100 Volts to 10 Volts. Once you see the twitch,
turn the stimulator selection dial (PULSE INTERVAL section) to SINGLE. Now you
can twitch the muscle each time you push the stimulator INITIATE button.
2. Insert the microelectrode into a muscle fiber near an endplate where the muscle and
nerve meet. Sometimes the pipette electrode will jump out of the cell due to the
motion artifact. You can always tell whether you are inside by looking at the resting
voltage. In this experiment, you will be re-poking cells all the time.
3. Stimulate the nerve by pushing the simulator INITIATE button. If all went well, you
should see an action potential (AP) on the scope with each push. Don't confuse the
AP with the stimulus artifact, a large, brief spike resulting from the stimulus pulse
itself. Move the scope horizontal position to get the action potential centered on the
scope screen. Capture a good looking pulse on the scope. Print and admire your
work. (Did you title the printout and label the axes?). Try the scope averaging
function to get very clean traces.
An example AP recording is in Appendix B.
Hint: If you can’t tell whether you are looking at an AP or a stimulus artifact, flip the
polarity switch on the stimulus isolation box. The artifact will flip but not the AP.
Hint: Once you have an AP, lower the stimulus amplitude as far as possible without
losing the AP. This will minimize the stimulus and motion artifacts.
Warning: If you see bubbles forming in the prep bath, turn off the stimulus isolator
immediately as something is causing large DC currents to flow through your prep.
No response to stimulation?
(1) The nerve or muscle is damaged. (2) Stimulator is off. (3) Stimulator settings are
wrong. (4) The electrode is filled with distilled water rather than Ringers. (5) You have
sucked up something other than a nerve. (6) No suction. (7) The reference electrode is out
of the bath.
Deliverable: Some nice AP printouts.
50
Other things you can do
1. Measure timing and voltage changes within the AP. Relate features on the AP to
membrane channel changes.
2. Find the stimulation threshold (in volts) to generate an AP.
3. Convince yourself that the AP is an “all-or-nothing” phenomenon by slowly varying
the stimulus amplitude while stimulating once per second in CONTINUOUS mode.
4. Repeated muscle twitches may cause tearing of the cell around the electrode, leading
to leakage of cell contents and a gradual reduction in membrane voltage Vm. Watch
Vm over time and note any change in AP shape, amplitude or duration. Can you
explain what you see?
5. Measure the time delay between the stimulus artifact and the peak of the AP. Poke the
same cell several places further away from the endplate and measure the delays. Use
the data to estimate AP conduction velocity (m/sec) in the muscle fiber. What could
cause errors in this estimate?
6. Find the value of the AP refractory period, the time period just following one AP
when you cannot elicit a second. Set the stimulator in double pulse mode. Starting at
a spacing of 5 mS, reduce the spacing in 1 mS increments until the second AP
disappears. Home in on the refractory period by fine tuning the spacing.
7. Find the stimulus amplitude that just produces an AP. Change the stimulus pulse
width and repeat. Plot the curve of threshold settings of PW and AP. What are the
chronaxie and rheobase? (Look the terms up in a reference text.)
8. Show how the timing and amplitude of the AP change with resting membrane
potential level. (Hint: You already know how to change the resting potential.)
51
3.17 End plate potentials
Background: Neuromuscular transmission starts with an action potential propagating
down an axon to the end where it opens voltage-gated Ca channels. The entry of Ca++
causes exocytosis of acetylcholine (Ach) vesicles from the presynaptic terminal into the
synaptic cleft between the terminal of the axon and the muscle membrane end-plate. Ach
binds to receptor sites (AchR molecules) on the end-plate causing their channels to open.
These non-specific channels are permeable to Na+, K+, Ca++ and other ions. Since these
channels are more permeable to Na+ then to K+ and the driving force across the
membrane is greater for Na+, there will be a net influx of positive charge. This leads to a
depolarization, which is called the end-plate potential (EPP) or post-synaptic potential
(PSP). The EPP occurs only in the end-plate region but can be seen further down the fiber
through passive charge spread. Contrast this behavior to the action potential which travels
by active depolarization of the cell membrane. The EPP is normally 10-40 mV, which
exceeds the threshold for the action potentials that are then generated and propagated in
both directions along the muscle fiber.
To study the EPP without generating an action potential, it is necessary to reduce the
amplitude of the EPP by reducing the number of active channels. One could apply curare
which binds to AchR preventing channel opening in some fraction of the Ach channels,
or tetrodotoxin (TTX) to block the voltage gated channels. Reducing the calcium
concentration also reduces Ach release and will reduce the EPP amplitude to below AP
threshold levels. In the frog prep, the calcium concentration can be lowered by changing
the bathing solution to a low-calcium Ringer’s. By gradually replacing the bathing
solution, you can observe the quantal nature of synaptic transmission by observing step
changes in EPP amplitude. As the number of vesicles releasing ACh decreases, you can
observe the incremental jumps in EPP amplitude due to the quantal nature of vesicle
release. You will also be reducing the frequency (but not the amplitude) of MEPPs.
For this experiment, your setup will be almost exactly like Figure 13-1 in the reading by
KL Magleby titled "Neuromuscular Transmission". Look at that figure now and read
enough of the text so that you understand the basic physiology that drives the EPP
process.
Prep and equipment and connections: Same as the MUAP experiment, except this
time try to insert your recording microelectrode near an endplate. In a frog muscle, the
shape of this junction resembles a brush more than a plate. To see it, look carefully using
the highest magnification.
Hint: You are in a good location is you see MEPPs. No MEPPs means likely no EPP
because the electrode is too far from an endplate.
Procedure
1. Get a nice looking AP showing on the scope. The first part of the AP, which has a
slower rise, is the EPP. It will come right after the stimulus artifact. Depending on the
52
location of your electrode with respect to the endplate, you may or may not see a
distinct EPP. A typical EPP has an amplitude of about 30 mV and lasts about 7 ms.
2. Change the stimulator back to CONTINUOUS. Gradually replace the bathing
solution with the low Ca++ Ringer's. Watch the muscle and the EPP as the low Ca++
takes effect. Stop applying the low Ca++ solution when the muscle almost stops
twitching in response to each nerve stimulation. Return the simulator to SINGLE
mode.
3. Poke a muscle fiber near the end plate. Pulse the stimulator. If you don't see a
response on the scope, increase the simulation amplitude. If you still don't get a
response, try another fiber, perhaps close to one of the fibers that is twitching. You
don't want to be in a twitching fiber because those fibers still have AP's. If you still
can't get a response, slowly add back a small amount (just a few drops) of normal
Ringer's to the bathing solution. Your goal is to record from a cell that responds to
stimulation with a stable EPP and no AP.
4. Note the amplitude and shape of the EPP. Try penetrating the same muscle fiber in a
different location along its length. Is the shape and amplitude of the EPP the same?
Are you closer or further away from the endplate?
How to tell the difference between an EPP and an AP: (1) EPP is 10-40 mV, AP is 70110 mV. (2) EPP lasts 5-10 ms, AP lasts 2-4 ms.
Tip: You will only see EPPs if your recording electrode is near the NMJ.
Tip: Sometimes you can get EPPs in depolarized membranes which is what happens with
cut fibers or fibers that have ripped due to electrode poking.. Try impaling fibers that
look damaged and do not twitch. Probe near the end plates.
Deliverable: Some nice EPP printouts (labeled of course)
Challenge
1. Find an AP that has an EPP as an initial shoulder. Print and show to an instructor.
2. Plot the amplitude of the EPP as a function of distance along the fiber. Charge decay
theory says EPP amplitude should decay exponentially with distance from the junction.
Does your plot show this?
Double Challenge
As you lower Ca++, fewer and fewer vesicles contribute to the EPP. The minimum EPP
is caused by one vesicle, just the same as for the MEPP. If your are very careful with
Ca++ application and have a noise-free recording prep, you can see the quantal nature of
the EPP as additional vesicles are added. Can you deliver some printouts that show this?
53
3.18 Measuring sarcomere length
The length of the muscle sarcomeres can be estimated by taking advantage of the fact that
sarcomeres occur at very regular spacing along a muscle fiber. Shining a laser beam
through a thin sheet of muscle causes a diffraction pattern with the width of the first
dominant fringe being directly proportional to sarcomere length.
This measurement is surprisingly easy to do with a laser (Figure 3-31), the muscle and a
sheet of plain white paper. Figure 3-32 shows the setup with a laser shining up through
the central portion of the muscle sheet with the paper located a distance L from the
muscle. Start by holding the paper about 6 inches (not exact) above the muscle and look
down through the paper. You will see a bright central line and also a pair of fainter lines
equally spaced on either side of the center line. You might have to turn off the room
lights to see the fainter side lines. These are the first pair of diffraction lines and the
distance between them is 2x in the figure. (Challenge question: The lines run
perpendicular to the line of muscle fibers. Why?)
2x
θ
L
muscle
laser
Figure 3-31: Laser used for measuring sarcomere
length. Shine up through prep
Figure 3-32: Laser diffraction set-up to measure
sarcomere length.
Once you confirm that you can see the diffraction lines, come up with a clever way of
holding the paper fixed at a distance of 12 or more inches over the muscle. For example,
you could tape the paper to the ceiling, but that might be too far away too make out the
faint lines. With the paper fixed, measure the distance between the paper and the muscle
and the distance between the diffraction lines. (An easy way to do this is to mark the
paper where the lines are and later measure the spacing with a ruler.)
To compute the sarcomere length, use Bragg's law for diffraction
nλ = d sin θ
where n = diffraction order (=1 for the first diffraction pair), λ = the wavelength of the
laser (0.635 microns), θ = arctan(x/L = the angle of diffraction, d = sarcomere length.
54
For example, if you measure a diffraction spacing of 2x = 5.4 cm and L = 10 cm, then θ =
arctan(0.27) = 15.11, therefore d = (1*0.635)/sin(15.11) = 2.44 microns. At full overlap,
sarcomeres are 1.6 to 2.4 microns long while for zero overlap they are about 3.8 microns.
Deliverable
Find the sarcomere length (in microns) for the pinned out prep.
55
3.19 Force measurements
The whole point of muscle is to generate force causing parts of the body to move. Up
until now, you have measured the electrical properties of the muscle in response to
chemical and stimulation excitation. Now it is time to measure the mechanical properties
of the muscle.
Measuring force requires an excitation, in this case a stimulation pulse, and a means to
sense the force. The latter is achieved with a force transducer (sometimes called a force
gauge, force sensor or load cell), a device that converts an applied force to an electrical
signal that can be measured on the scope.
Measuring force also requires a means to easily change the overall muscle length. In your
prep, you will keep the skin flag anchored but release the xyphoid process. By tying a
thread through the xyphoid and attaching the other end to a manipulator, you can set the
muscle to whatever length you chose.
Each time a stimulus pulse is applied, the muscle twitches. When measuring single cell
electrical properties, you wanted as little of the muscle to be activated as possible to
prevent motion artifacts. When measuring force, you want all of the muscle to be active
so that the force can be read. (Our force transducers are not sensitive enough to pick up
the force from a single cell).
Because muscles are elastic, they resist passive stretching. This passive force, roughly
exponential with length should be contrasted with the active force generated from
excitation. The total force is the active force plus the passive force. Because the passive
force can be considerable with a stretched muscle, sometimes the best way to read the
active force is to zero out the force transducer each time you set the muscle to a new
length.
The first step in measuring force is to calibrate the force transducer. After that you will
prep the muscle so that length can be changed. Next will be measuring force twitches in
response to stimulation. Once you have these basic techniques down, there are a wide
range of experiments you can perform that reveal various characteristics of the
mechanical output of muscle.
56
3.20 Force sensor setup
The Grass Instruments Model FT03 force sensor (Figure 3-33) is used to measure
isometric muscle force from the frog CP setup. The sensor will be mounted on a
manipulator stage to make it easy to change muscle length. The following table shows
the basic specs for the sensor when it has no added springs, the most sensitive setting 5.
Max load (gm)
Min resolvable load (mg)
Resonant frequency (Hz)
50
2
85
The FT03 has an internal strain gage full Wheatstone bridge that converts changes in
strain of the loading beam to changes in resistance of the bridge which in turn results in
changes in voltage at the bridge output. The output of the force sensor is a small voltage
that must be amplified to be seen on the scope. This is the job of the force sensor
amplifier, the Grass P11T Strain Gage Amplifier (Figure 3-34).
WARNING: The sensor is delicate and does not like to be hit or dropped. Treat it with
respect.
Figure 3-33: Grass FT03 force sensor.
Figure 3-34: Grass P11T strain gage amplifier.
Before measuring muscle force, the sensor and amplifier must be calibrated.
Tip: Setup and calibration of the force sensor takes time. We suggest that one person
does the calibration while the other prepares the muscle as described in Section 3.21.
5
By adding springs, the max load can be changed to 200, 1000 or 2000 gms. To change springs, see
Section 5.1 of the Grass instruction manual for the FT03.
57
Connections: FT03 sensor to P11T amplifier IN. P11T amplifier OUT (the BNC
connector) to scope CH1. Grass RPS210 power supply brick to P11T amplifier POWER
IN.
Calibration: The sensor must be calibrated the first time you use it. For calibration, find
and weigh on the Mettler balance a small alligator clip that weighs between 0.5 and 2
gms. Clamp the sensor in the horizontal position with the cable facing up. Do the
following to calibrate the force sensor.
1. Adjust the GAIN knob on the P11T to about the middle of its range.
2. Adjust the BALANCE VOLTAGE knob on the P11T to get close to 0.00 V on the
P11T display.
3. Record the output value, which will be near, but probably not exactly 0.00 V
4. Clip alligator clip onto the top loading stalk of the sensor so that it loads down the
sensor (Figure 3-35). Record the output value on the P11T display. Remove clip.
5. Repeat the last two steps three or four times to get a set of no load and load
readings.
6. For each set of readings, find the difference between the no load and load reading.
Compute the average difference and divide by the weight of the clip. The result is
the calibration gain of the sensor in volts/gram. You should get a calibration gain
in the range of 1.0 V/g to 2.0 V/g. If not, change the gain knob and recalibrate.
After calibration, you can zero the sensor with the BALANCE VOLTAGE knob as often
as you like. If you change the GAIN knob, you must recalibrate the sensor. If the display
reads over 6 V, the amplifier is saturated, which means the gain is too high.
Figure 3-35: Loading force sensor stalk with a small alligator clip to calibrate the sensor.
58
3.21 Muscle twitch force
The amount of force generated by a muscle depends on the overall length of its fibers, or
more precisely, the length of the sarcomeres. In this experiment you will generate a
muscle twitch and will measure the stimulated muscle force as the overall length of the
muscle is changed.
Prep
.
Pin the proximal (skin) end of the muscle to the prep dish, close to the side of the dish.
Leave the distal end with the associated chunk of xyphoid process free. Using a length of
5-0 suture thread, lasso the muscle right where it joins the xyphoid. Tugging on the
thread now gives you a way to stretch the muscle.
Attach the force sensor to a micromanipulator so that you can adjust muscle length
accurately and can record length changes in millimeters. Tie the other end of the suture
thread through the hole in the loading stalk of the force sensor. Use the stalk on the side
facing away from the force sensor cable. Keep the suture slack while tying to avoid
accidentally tearing the muscle. Keep the distance between the muscle and the force
sensor under 10 cm.
Align the force sensor and manipulator so that when you change one axis on the
manipulator, the thread and sensor loading stalk are in axial alignment and the sensor
moves back and forth along the direction of the thread with the thread just clearing the
bath.
Note: It is important that the thread be as close to parallel to the table as possible without
touching the lip of the prep dish and that the thread moves axially when you turn the
knob on the manipulator.
Arrange a suction electrode to stimulate the muscle motor nerve. Stretch the muscle out a
bit and confirm that you can see twitches when you stimulate the nerve.
Your prep should look something like Figure 3-36 and Figure 3-37.
59
Figure 3-36: Prep for measuring muscle force
Figure 3-37: Close-up
60
Connect the strain gage amplifier OUT to scope CH1. Set the stimulator to SINGLE.
Capture a nice looking force twitch on the scope It should be about 100 mS long. Print
and label axes, where the vertical axis is labeled in gms using the data collected when
calibrating the sensor. Measure the amplitude and duration (time from 50% of peak
amplitude on the rising side to 50% of peak on the falling side) of the twitch.
If you can't get a nice looking twitch, but you see the muscle twitching, try raising the
gain on the strain gage amplifier (recalibrate following the experiment). Or try stretching
the muscle. If the force twitch is saturating the amplifier, turn the gain down.
Hint: If the nerve keeps pulling out of the suction electrode, or if you are missing a
length of nerve, use a hook electrode with one leg on each side of the prep. See Appendix
H.
Hint: If there is no twitch, confirm that the stimulator is working by placing a recording
electrode in the bath (not in a cell) and confirming the presence of a stimulus artifact.
Deliverable: A printout of a nice looking muscle force twitch. Label the vertical axis in
gms.
Once you get reliable twitches, there are many options for experiments using this prep,
some of which are described below. These are also possibilities for your independent
research project. For a research project about muscle force, consider using the frog
sartorius, which is easier to dissect and has large force.
Length-tension (overall muscle/tendon length)
Collect twitch data at a variety of overall muscle-tendon lengths where length is
measured right off the micromanipulator. After setting each new length, zero the force
sensor so that your reading will be the active force. Stretch the muscle by about 1 mm
each time. Plot the muscle active length-tension curve using units of mm and gms. If you
want the muscle total and passive length tension curve, keep track of what the resting
force is each time you change muscle length.
Length-tension (sarcomere length)
Do a length-tension experiment, but this time measure the actual length of the sarcomeres
using the laser light method described previously. Position the laser underneath the
muscle chamber, and direct the light beam through the central region of the preparation
Position a calibration grid above the preparation, such that the laser beam diffraction
pattern is displayed on it. Stimulate the muscle preparation and record total force:
determine both the preload force and the active (peak twitch forces). By recording total
force and determining the peak force, it is possible to determine the length-tension
relationships for both the passive and active forces. Lengthen or shorten the preparation
and repeat above. Note, you should be able to also estimate the passive changes in
61
sarcomere spacing by noting changes in the diffraction patterns. Deliverable: A plot of
the passive sarcomere length vs. overall muscle length.
Recruitment curve
As the stimulus pulse amplitude is increased, more and more axons are activated, more
and more motor units take part in the twitch and the twitch peak force increases. Collect
peak twitch force as you change the stimulus amplitude. Plot force as a function of
stimulus amplitude to get the recruitment curve.
Force-frequency
See what happens as you increase the frequency of stimulation. Plot a force versus
stimulation frequency curve. (WARNING: When stimulating at anything greater than
once per second, stimulate for very brief periods only to minimize muscle fatigue.)
Doublets
Set the stimulator to double pulse mode. See what happens to the force twitch as you
lower the spacing between doublets from 100 mS to 1 mS. Plot the twitch amplitude
versus doublet spacing.
62
4 Your research project
4.1
Purpose
Now that you have learned the theory of NMJ and muscle contraction and have mastered
basic laboratory techniques, it's time to launch off on your own. For the remainder of the
week, your team will conduct an independent experimental research project on a topic
that interests you. You can explore one of the experiments you covered earlier in the
week in more depth, or something completely new. For ideas, develop your own idea,
look through this manual for ideas, look at the course readings for ideas or consult the
course instructors.
Once you have chosen a topic, use the reference books in the lab, journal articles in the
readings package or the internet (PubMed) to come up to speed quickly on the topic.
Only spend enough time on background to figure out what you are doing; this is a lab
course and we want you to be doing experiments.
While free thinking is encouraged, please have some reasonable idea or hypothesis that
forms the basis for your experiment. The idea is to do hypothesis-driven research.
4.2
Deliverable
Your deliverable is a 12 minute oral presentation on your project, done in a style you
would use for an oral presentation at a conference. Presentations should follow this
format:
Slide 1:
Title of project, names of investigators, date. Have the title state the results of
the experiment, e.g. “Muscle Force Increases with Stimulation Frequency,”
rather than “The Effect of Stimulation Frequency on Muscle Force.”
Slide 2: Slide title = “Background” The basic physiology relevant to your experiment
and anything you found out in the literature. Keep it brief. One slide.
Slide 3: Slide title = “Specific Aims” State the aim of your project and the formal
hypothesis that drives your experiment.
Slide 4: Slide title = “Methods”. Details on your prep. One slide. You may add one
additional slide if your methods are particularly unusual or complex.
Slides 5-n: Results. The data you collected. Use tables and plots and present your data as
clearly as you can. The title of each slide should make a statement about the
result, rather than the title being “Result”. Use as many slides as you need,
keeping in mind time limits for the whole presentation. We recommend one or
two results slides, keeping in mind that you can have more than one plot per
slide.
63
Slide n+1: Slide title = “Discussion”. Interpret the data for the audience. What does it all
mean? Did it confirm or refute your hypothesis? Did your data match what
others have found in the past? One slide
Hint: Impress us with your data, not with your ability to master PowerPoint.
4.3
•
•
•
•
•
•
•
•
4.4
Presentation tips
Avoid stories of how long it took or how hard it was to collect the data. Instead you
want to present as if you were at a conference.
Slides should be clean, with images and data charts dominating over text
Report numeric data to two significant digits, unless you have reason to believe your
data is accurate to better than 1%.
Use statistical tests (t-test, ANOVA) to test hypothesis.
For plots, use white background and no grid lines.
On plots, label axes, including units (e.g. “Force (gms)” and “Memb voltage (mV)”,
and add a descriptive title.
Use nice fat dots to mark data points and connect with straight lines (no fitting or
smooth lines).
Use a bar chart to compare two or more treatment conditions. For repeated data,
height of the bar marks the mean. Add whiskers to indicate standard deviation.
Research project suggestions
There are many possible topics you can pursue for your research project. Here are some
ideas to get you going, but you are by no means restricted to this list. Be creative and
explore what interests you.
Frequency of single, double and triple MEPPs
Measure refractory period
Effect of temperature on anything
Recruitment curve
Doublets
Effect of NMJ blockers on twitch time and
amplitude
Pharmacology of NMJ
Effect of X on EPP
Human muscle force assessment
How MEPPs amplitude and frequency vary with
temp, [Ca++], ....
Length-tension curve (overall and sarcomere)
Effect of pH, glucose, temperature
Force-frequency relation
Effect of X on muscle force fusion frequency
AP threshold curve for pulse width, amplitude
combinations
Effect of X on MEPP
Any of this week’s experiments, but done more
carefully
Your idea goes here
64
Appendix A TEK SCOPE
REFERENCE
Display area
1
2
3
4
5
SCOPE DISPLAY AREA
Icon shows acquisition mode
Icon shows trigger status. "Ready" means scope is waiting for a trigger.
"Trig'd" means scope is being triggered. "Acq. Complete" means the scope
has finished acquiring a single sweep.
Marker shows horizontal trigger position. Turn HORIZONTAL POSITION
knob to adjust
Readout shows time between trigger marker and center graticule
Marker shows trigger voltage level
65
6
7
8
9
10
11
12
13
14
15
16
Markers show zero voltage level of displayed channels
Down arrow indicates waveform is inverted
Readouts show vertical scale factors of displayed channels
Bw icon means channel has been bandwidth limited
Readout shows horizontal (timebase) scale
Readout shows window time base setting, if zoom window is in use
Readout shows trigger source
Icon shows trigger slope
Readout shows trigger level
Various operation messages
Readout shows trigger frequency
CH1 (or CH2) MENU
Controls appearance of channel traces. Hit twice to turn channel off.
Coupling
BW Limit
Volts/Div
Probe
Invert
Selects how channel is coupled to scope. Use DC for most signals. Use
AC if trying to see small signal on top of a large offset. Use GND to
flatline trace
Always leave OFF
Always leave Coarse
1X if using BNC cables. 10X if using scope probe
Leave Off
TRIG MENU
Controls how scope is triggered
Type
Source
Slope
Mode
Coupling
Use Edge
CH1 or CH2 to trigger off signal being viewed on scope. EXT for
triggering off external signal (e.g. when using stimulator as a trigger), AC
LINE to trigger off wall current (handy to see if getting AC noise on
signal)
Dictates edge of trigger
Use Normal or Auto
Use DC. Some special situations use AC
VERTICAL CONTROLS
Use POSITION to position trace on the screen and VOLTS/DIV to set the vertical scale.
66
HORIZONTAL CONTROLS
Use POSITION to position trigger. If you want to see lots of the wave post-trigger,
position trigger at left. If you want to see what happened just before the trigger event,
position trigger at right. If the trigger is far to the left or right and you want to get it back
in the center, push SET TO ZERO. The readout at the top of the display shows the time
at the center of the screen with the trigger representing zero time.
To zoom in, push HORIZ MENU, then the Window Zone option button. Two cursors
appear to define a window zone. Adjust with the HORIZONTAL POSITION and
SEC/DIV controls, then press the Window option button to zoom in. Press again to
restore.
TRIGGER CONTROLS
LEVEL sets level of trigger. SET TO 50% is an easy way to set a trigger level if you
have a signal present. TRIG VIEW is handy to see what the trigger signal looks like
when you are using EXT trigger.
RUN/STOP
Push to freeze the signal. Push again to unfreeze.
SINGLE SEQ
Push to initiate a single sweep or to start an averaging operation. Handy for capturing an
intermittent event. After pushing, trigger icon will say "Ready" when waiting for a
trigger, and "Acq Complete" after the sweep or the averaging operation is over.
PRINT
Use the ScopeConnect app on the station PC to capture screen images and to transfer
scope data to Excel.
AVERAGE
Use averaging to scan a number of times and average the waveform. This is an excellent
way to reduce the noise on a repetitive signal. The Averages soft button lets you select
4/16/64/128 sweeps for averaging. The more sweeps, the more noise goes away, but at a
cost of taking longer. To Average: ACQUIRE then AVERAGE then SINGLE SEQ.
Averaging is complete when message at top reads “Acq complete.” ACQUIRE then
SAMPLE to disable averaging.
MEASURE
67
This is a way to measure time and amplitude features of stored trace or continuous signal.
Push MEASURE, then one of the five soft buttons. Select the Source for what will be
measured, and then the Type. Type choices are: frequency, period, mean, peak-to-peak
amplitude, RMS amplitude of one cycle, min, max, rise time, fall time, width of positive
pulse, width of negative pulse. Use Type = Mean to turn the scope into a digital
voltmeter.
CURSOR
Excellent way to measure time and amplitude features of a stored trace. To measure
voltage, select Type = Voltage, then use the CH1 and CH2 VERTICAL POSITION
controls to position cursors on the screen. To measure time, select Type = Time.
STORING WAVEFORMS
The scope can store any displayed waveform for later recall. Push SAVE/RECALL (in
the MENUS area), select the source to store, Ref to store the trace in either memory A or
memory B, then Save to actually store the waveform.
To display a stored waveform, push SAVE/RECALL, select a waveform with the Ref
option button, then display on the screen with Ref(x) = On. Remove the stored waveform
from the screen by selecting Ref(x) = Off. A stored wave can be superimposed on a
current trace, which is one way of getting three traces on the screen.
FFT
Find the spectral content of a signal using the FFT function. Center the signal in the
display, and adjust VOLTS/DIV until the signal fills the screen. Turn SEC/DIV to set the
FFT frequency resolution (faster setting means the FFT shows a larger frequency range).
Push MATH MENU (in the VERTICAL section), set the Operation option button to FFT
and select the Math FFT Source channel. Once the spectrum is acquired and displayed,
use the HORIZONTAL POSITION knob to sweep through the spectrum. The readout at
the top of the display shows the frequency at the center graticule line.
68
Appendix B Sample plots
0.2
mV
5 ms
MEPPs. Averaged 4 times. Note amplitude and time. Also note how time and amplitude
are annotated with the “L” marker. This is important because the scope readings do not
take amplifier gain into account. Also note how the scope CURSOR function was used to
find the MEPP amplitude (see the DELTA readout at the right, and note that it is off by
the amplifier gain of 100).
69
20
mV
1 ms
A motor unit action potential. Averaged to eliminate noise. Note the amplitude and
timing. The AP is on scope CH1 and the stimulus signal is on CH2. Note where zero
volts is for CH1 (marked by the arrow labeled “1”). What was the resting membrane
potential for this cell?
70
Appendix C Stimulator controls
The figures below show what each stimulator control sets.
Single pulses
PA (4)
Synch
PW (3)
Del (2)
1.
2.
3.
4.
PI (1)
PULSE INTERVAL section, SECONDS control
PULSE DELAY section, SECONDS control
PULSE WIDTH section, SECONDS control
OUTPUT section, LEVEL control
Double pulses
PA (4)
PW (3)
Del (2)
PI (1)
1.
2.
3.
4.
PULSE INTERVAL section, SECONDS control
PULSE DELAY section, SECONDS control
PULSE WIDTH section, SECONDS control
OUTPUT section, LEVEL control
71
Pulse trains
PA (5)
PW (4)
TPI (3)
TD (2)
PI (1)
1.
2.
3.
4.
5.
PULSE INTERVAL section, SECONDS control
TRAIN section, DURATION control
TRAIN section, PULSE INTERVAL control
PULSE WIDTH section, SECONDS control
OUTPUT section, LEVEL control
72
Appendix D Frog Ringer's solutions
Composition of normal frog Ringer's (2.5 mM KCl)
Compound
KCl
NaCl
Na2HPO4
NaH2PO4
CaCl2 *
MgCl2
mM
2.5
115.0
4.00
0.85
1.8
1.0
gms for 5L
0.94
33.58
2.84
0.59
1.32
1.02
*Add after spinning to oxygenate.
To make a 5L solution of normal frog Ringer's, prepare as follows:
1. Add to 1 L distalled (boxed) H2O all ingredients above (use “gms for 5L” column)
except CaCl2.
2. Add distilled H2O to 5 L.
3. Spin to oxygenate.
3. Add CaCl2
4. Adjust Ph to 7.4 with 1 M NaOH if needed.
For the low Ca, high Mg solution used to create a partial synaptic block during EPP
measurements (Section 3.17), do not add the CaCl2, but instead add 0.47g CaCl2 to 4L of
the solution and 0.37g MgCl2 to 1L of the solution, then mix together.
73
For the effect of external K+ concentration on the membrane potential experiment
(Section 3.14), need solutions with the following concentrations, along with normal (2.5
mM [K]) Frog Ringers.
[K]
1.0
5.0
10.0
25.0
50.0
100.0
[Na]
116.5
112.5
107.5
92.5
67.5
17.5
To mix the six 1L stock solutions (1/2L of each is enough for NMJ week):
1. Obtain six 1 L containers. Label with [K+] concentration.
2. In another container, add 1 L distilled (boxed) H2O .
3. Add: 5.7 gms Na2HPO4, 1.2 gms NaH2PO4, 2.03 gms MgCl2
4. Ph should be about 7.4. If not, add 1 M NaOH to adjust.
5. Fill each of the six containers with 900 mL distilled (boxed) H2O.
6. Add 100 mL of the solution made in Step 3 to each container.
7. Add NaCl and KCl using this table
[K]
1.0
5.0
10.0
25.0
50.0
100.0
gms NaCl
6.8
6.6
6.3
5.4
4.0
1.0
gms KCl
0.07
0.37
0.74
1.86
3.72
7.45
8. Oxygenate by spinning.
9. Add 0.26 gms CaCl2 to each container.
10. Store overnight in refrigerator.
74
Appendix E Noise!
Membrane voltages are very small. A MEPP event has an amplitude of 0.5 mV. These
tiny changes can be swamped in the electrical noise radiated by the room lights, wallpowered instruments and computers, or by improper grounding. To get nice, clean
voltage waveforms, you may have to fuss with your setup. Eliminating noise is somewhat
of a black art, but here are two things you can try if noise is a problem.
First, make sure your experiment plate is grounded to the scope and equipment rack
(below, left). Try connecting and disconnecting the various ground leads attached to the
base plate while watching noise on the scope. Try grounding the front face of the
microscope in addition to or in place of the microscope rear ground connection. You can
also try grounding yourself, grounding all pieces of equipment, or pulling a lower
impedance electrode. If the signal slowly drifts, try a new prep dish ground wire. There
are no rules here; go with whatever works.
Second, if you still have noise and want to measure MEPPs, shield your head stage
and/or other low-voltage parts of your setup by making an aluminum foil tent to go over
your prep, and/or by wrapping the head stage in foil. Using an alligator clip, ground the
foil to the plate (below, right). Try placing the foil in different positions: front, back, left,
right or all. Only try foil this after you have determined all other noise reduction methods
don’t work.
Grounding the plate
Shielding the setup with aluminum foil. Ground the
foil.
75
Appendix F Basic Experiment
Design and Statistics
Use good experiment design methods to plan a study protocol and the appropriate
statistical methods to test hypotheses and report scientific results. This can either be
descriptive statistics (e.g. means and standard deviations) to report data trends and
scatter, or statistical inference tests (e.g. t-test and ANOVA) to test a hypothesis. Here is
a crash course including how to use Excel to do the calculations.
Experiment Design
Text coming… (single factor, 2-level, single factor N-level, run order, replications,
number of samples or subjects to use)
Descriptive Statistics
Text coming…
T-Test
Text coming…
ANOVA
Text coming…
76
Appendix G Fitting a straight line to
data using Excel
Excel can be used to simplify the task of finding the force sensor gain from calibration
data by taking advantage of its ability to fit a straight line to a set of points. The slope of
the line fit to the calibration data is the sensor gain. Here is how to do it.
Assume that you have eight data points, four from the no load and four from the loaded
condition, and that the alligator clip you used to load the sensor stalk weighed 1.23 gms.
Enter your load/voltage data point pairs into columns A and B of a new spreadsheet file.
It will look like this.
B
A
1 Load (gms) Sensor (Volts)
0
0.292
2
0
0.221
3
0
0.245
4
0
0.189
5
0.876
1.23
6
1.23
0.799
7
0.853
1.23
8
1.23
0.825
9
Highlight the data and use the Chart Wizard to make a plot. Plot type is XY Scatter Plot
without lines connecting the points. After formatting, your plot will look like something
like this.
77
Force sensor calibration
1
Sensor (volts)
0.8
0.6
0.4
0.2
0
0
0.5
1
1.5
Load (gm s)
Right click on any data point, select Add Trendline. Under the Type tab, select Linear.
Under the Options tab, check Display equation on chart and Display R-squared value on
chart. Your chart will now look like this.
Force sensor calibration
1
y = 0.489x + 0.2368
R2 = 0.9877
Sensor (volts)
0.8
0.6
0.4
0.2
0
0
0.5
1
1.5
Load (gm s)
The resulting line and equation is the least-squares, best-fit straight line to your data. The
calibration of your force sensor is the slope of line which can be read straight off the
displayed equation. For this sensor, the calibration is 0.489 volts per gm.
78
Appendix H Bipolar Hook Electrodes
A backup method for activating muscle fibers is a bipolor hook electrode, basically two
platinum wires stuck in the solution so that the current passing between them flows
through the muscle tissue and nerve terminal branches (Figure H-1, Figure H-2). The
threshold of stimulation for muscle fibers is about ten times that for nerve fibers, so even
though you are passing the current through the muscle tissue, unless you have applied an
NMJ blocking agent such as curare, you will still be generating action potentials and
force twitches indirectly by activating the nerve.
There are three reasons for using hook electrodes: (1) the muscle is not fully twitching
with the suction electrode, (2) you have no nerve or a damaged nerve in the prep, (3) you
are doing experiments with an NMJ blocking agent, (4) you are getting frustrated with
the suction electrode.
Figure H-1: Bipolar hook electrode.
Figure H-2: Tip of hook electrode.
Place the stimulating electrode in another micromanipulator (Figure H-3). For a full
setup, you need three manipulators, one to hold the recording electrode, one to hold the
stimulating electrode and one to hold the force sensor. The hook electrode connects to the
stimulus isolation using its attached cable.
Position the electrode so that the hooks are in the bath on either side of the muscle sheet,
close to, but not touching the tissue (Figure H-4). Place so that current passing between
the two hooks will pass through the muscle in the nerve endplate region.
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Figure H-3: Biplor hook electrode for stimulating.
Figure H-4: Place hooks in bath on either side of
muscle.
Note
To truly isolate the stimulus current to the nerve, a suction electrode is the better choice.
Suction electrodes suck the nerve up inside a small tube that contains the stimulating
electrode wire. Current passing from the wire goes through the nerve to activate axons.
80
Appendix I Fixing Suction
Electrodes
Read the AM Systems suction electrode instructions.
If the glass tube is broken:
Use Prism FSG12 capillary tubing (1.2 mm o.d., 0.6 mm i.d.) for replacing. Score tube
with emery cloth or a file and snap off to produce a length of about 5 cm. When
assembling the replacement tube, put the polished (non-cut) end of the capillary distal so
that a smooth edge is presented to the nerve. Or, polish by rotating the end of the tube
over a flame.
If the electrode is not sucking:
Test by removing the suction electrode rotary syringe and replace with a standard 6 cc
syringe. This makes it easier to see what, if anything is coming in and out with suction. If
no suction, tighten the screw-in connections that secure the tube and the glass capillary to
the body. If the rotary syringe is misbehaving, try loosening the locking ring on top and
resetting the syringe zero position. Check that the BNC connector is tight. Check that all
O-rings are present and not twisted.
Repair is all about finding and fixing leaks.
If the center or ground wire is broken:
Solder on a new silver wire, .010” dia, using silver solder.
Note:
If needed, can use FMG12 tubing (1.2 mm o.d., 0.7 mm i.d.).
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Appendix J Measuring electrode and
membrane impedance
The Dagan preamplifier can be used in two other ways to not only measure the resistance
of the electrode with much greater accuracy, but also to measure the impedance (both
resistance and capacitance) of the cell membrane. For these measurements the preamp
injects current into the electrode/membrane system and monitors the resulting voltage
response.
Make these cable connections:
1. Stimulator MONITOR to amplifier GATE
2. Stimulator SYNCH to scope EXT TRIG
3. Amp 10 Vm to scope CH 1
4. Amp I MON to scope CH 2
Implement these amplifier settings:
1. Z TEST switch off,
2. STEP CURRENT switches to (+) and OFF
3. MONITOR display V1 button to read voltage on the display
Resistance measurement method #1: Bridge Balance Method
In this method, the amplifier injects a small steady-state current into the electrode. The
resistance of the load can then be measured by monitoring the voltage. “Load” means
whatever is plugged into the head stage. If it is the model cell, the load could be the
simulated membrane, the simulated electrode, or both depending how you have the
jumper placed. If the head stage is connected to a microelectrode lowered into Ringer’s,
the load is the electrode resistance. If that microelectrode is poked into a cell, the load is
the electrode resistance plus membrane resistance.
To measure load resistance:
1. Spin the amplifier BALANCE control to zero (fully CCW)
2. Use the amplifier OFFSET knob to precisely zero the voltage as read on the LCD
display.
3. Switch the MONITOR display to read current by pushing the I1 button below the
display.
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4. Flip the STEP CURRENT switch to CONT to inject a continuous current, and
spin the STEP CURRENT vernier until the display reads about 50 nA. You
should see the voltage across the cell (CH 1 on the scope) change in response.
5. Switch the MONITOR display back to read voltage (the V1 button below the
display)
6. Use the amplifier BALANCE knob to bring the voltage back to zero. The reading
on the BALANCE knob that zeros the voltage equals the resistance of the load.
(Full CW is 100 M-ohms)
Challenge:
Using the model cell, use this method to measure resistance with the membrane shorted,
the electrode shorted and with both in the circuit. How does this compare with what you
got using the Z TEST method?
Resistance measurement method #2: Impedance Calculation Method
In this method, the preamp injects a step change of current into the load using the
stimulator for timing. Examination of the resulting voltage waveform reveals the
electrode resistance, the membrane resistance and the membrane capacitance.
Amplifier settings: BALANCE knob to zero (full CCW), Z TEST switch off, STEP
CURRENT switches to (+) and OFF, STEP CURRENT current knob zeroed (full CCW),
MONITOR display V1 button (to read voltage), Spin OFFSET to zero display.
Stimulator settings: PULSE INTERVAL: Selector switch to CONTINUOUS, Interval =
500 mS. DELAY = 0, TRAIN off, WIDTH = 300 mS. You should see the red LED
labeled ACTIVE flashing twice a second.
Scope trigger settings: Type = Edge, Source = Ext, Slope = Rising, Mode = Normal,
Coupling = DC. Pres SET TO 50% or play with the trigger level control until you see the
“Trig’d” label at the top of the scope display flashing twice a second in synchrony with
the stimulator.
Scope time base: 50 msec/DIV.
Model cell: Unplug the shorting cable so that the full simulated electrode membrane
combination is presented as the load.
To measure impedance:
1. MONITOR button I1 to measure current on the display, STEP CURRENT switch
to CONT, adjust vernier until you get about 50 nA going to the electrode (read off
display).
2. STEP CURRENT switch to GATE. Fuss with the scope amplitude and position
controls and the horizontal position controls until you get the voltage display on
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CH 1 to look something like the figure below. Position CH 2, which shows the
step of current going into the electrode, below and out of the way of CH 1. Print;
there is a lot to look at on this figure.
Examine the wave form which should be something like the figure below. You will see
an initial rapid rise (labeled Is*Re in the figure). Is is the current being sent through the
load and Re is the electrode resistance. By Ohm’s law, V=IR, the voltage is the current
times the resistance. The rise is rapid because there is no capacitance associated with the
electrode. Since you can read the voltage off the scope (don’t forget there is a gain of 10
in the amplifier), and you know Is from the monitor display, you can compute Re. Or, spin
the BALANCE knob until the rapid rise phase disappears. The value of the BALANCE
knob will be the electrode resistance.
The exponential rise portion of the waveform is labeled Is * Rm in the figure. This part is
caused by the membrane resistance and capacitance. To compute the membrane
resistance, use Ohms law as you did for the electrode resistance. Or, use the BALANCE
control to zero out this portion of the waveform. (There will be spikes at the start and end
of the pulse; you are zeroing out the flat portion in the middle.) The additional resistance
read off the BALANCE control is the membrane resistance.
Now reset BALANCE fully CCW. Turn off CH 2 and expand the CH 1 trace so that it
fills the screen horizontally and vertically with the start of the exponential rise phase due
to the membrane lined up with division markers. Save and print this trace.
Use a straightedge to draw a line tangent to the initial slope of the membrane rise as
shown in the figure. The time at which this line crosses the final value line is one time
constant, marked as Rm * Cm in the figure. Since you already know the membrane
resistance, use these facts to find Cm. For this calculation, use time in seconds and
resistance in mega-ohms. The resulting capacitance units will be in microfarads.
Challenge:
For the model cell, use these methods to find the precise electrode and membrane
resistance and your best estimate of the membrane capacitance.
84
Is*Rm
V
Is*Re
Rm*Cm
t
Voltage seen when a step change in current is applied to an electrode/membrane combination, or to the
model cell.
85
Appendix K Changing settings on the
Sutter micropipette puller
This section is for instructor reference. Only instructors should change the heat setting.
Use FSG 12 glass micropipette stock (1.2 mm O.D., 0.6 mm I.D.)
Electrode impedance is determined by pipette tip geometry. The puller dictates a precise
and repeatable heat, delay time to pull, pull force and pull velocity. These are the
numbers you see on the displayed puller program.
Leave the following settings alone:
PULL = 100
VELOCITY = 30
TIME = 150
HEAT = 490-550
The HEAT setting dictates tip diameter and therefore electrode impedance. Higher
settings means the glass becomes very ductile and long slender tips with high impedance
will be pulled. Lower settings results in shorter tips with larger diameters and lower
impedance.
Each time the heating element is replaced or each time the puller is transported or each
time the puller is used after being idle for several months, the HEAT setting should be
checked to see if good electrodes are being pulled. If you are not getting electrodes
somewhere close to 40 Mohms (20-60 is acceptable), it is time to change the HEAT
setting. Change in increments of 5 and iterate until good electrodes come out. For the
FSG 12 glass micropipette stock, useful HEAT values are between 490 and 550. In July
2006, 515 worked well, but your values may be different.
To change the setting:
1. Unlock the program by CLR > 0 > 7 > 0
2. Enter new heat number without hitting ENTR
3. Lock the program by CLR > 0 > 7 > 1
Need more info? Wade through the Sutter manual.
86
Appendix L Resistor Color Codes
Resistors are marked with colored bands that indicate the resistance value and an optional
value tolerance.
The value is a set of three bands. Band 1 is the first significant digit, Band 2 is the second
significant digit and Band 3 is the number of zeros that follows the number. The code is
Color
Black
Brown
Red
Orange
Yellow
Green
Blue
Violet
Grey
White
Number
0
1
2
3
4
5
6
7
8
9
For example, a resistor marked with green-blue-green is 56 followed by 5 zeros or 5.6
mega-ohms. A resistor marked with red-red-orange is 22 followed by 3 zeros or 22 kiloohms.
The final band is either gold or silver and indicates the value tolerance with gold meaning
plus or minus 5% and silver meaning plus or minus 10%. A resistor marked green-bluegreen-silver is 5.6 ± 0.56 M-ohms and could be anywhere between 5.0 and 6.2 M-ohms.
87
Appendix M NMJ Course Supplies
and Drugs
88