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 COURSE OVERVIEW ........................................................................................................................... 3 1.2 PREVIEW OF EXPERIMENT ACTIVITIES ............................................................................................... 4 1.3 NOTEBOOK ....................................................................................................................................... 5 1.4 YOUR RESPONSIBILITIES ................................................................................................................... 5 1.5 KEEPING YOUR SANITY ..................................................................................................................... 5 1.6 HELP US IMPROVE THIS MANUAL ...................................................................................................... 6

2 EQUIPMENT AND INSTRUMENTATION ................................................................................... 7 2.1 OVERVIEW OF NEUROPHYSIOLOGY EQUIPMENT ................................................................................ 7 2.2 THE NMJ EQUIPMENT RACK ............................................................................................................. 7 2.3 OSCILLOSCOPE BASICS ...................................................................................................................... 9 2.4 PULSE STIMULATOR BASICS .............................................................................................................. 9 2.5 RECORDING ELECTRODE AND AMPLIFIER BASICS .............................................................................. 9 2.6 COMPUTER-BASED EXPERIMENT CONTROL ....................................................................................... 9

3 TECHNIQUES, EXERCISES AND EXPERIMENTS ...................................................................10 3.1 GETTING TO KNOW THE LAB ............................................................................................................10 3.2 CABLES AND CONNECTORS ..............................................................................................................10 3.3 GETTING TO KNOW YOUR OSCILLOSCOPE .........................................................................................11 3.4 WORKING THE STIMULATOR ............................................................................................................17 3.5 USING THE ELECTRODE PREAMPLIFIER .............................................................................................22 3.6 USING MICROSCOPES .......................................................................................................................24 3.7 SIMULATED CELL .............................................................................................................................25 3.8 CHECKING ELECTRODE IMPEDANCE .................................................................................................26 3.9 MAKING A GLASS MICROELECTRODE ...............................................................................................28 3.10 PITHING THE FROG ......................................................................................................................32 3.11 DISSECTING THE CUTANEOUS PECTORALIS MUSCLE ....................................................................33 3.12 MEASURING MEMBRANE POTENTIAL ...........................................................................................38 3.13 MINIATURE END PLATE POTENTIALS (MEPPS) ............................................................................42 3.14 EFFECT OF K+ CONCENTRATION ON MEMBRANE POTENTIAL.......................................................44 3.15 STIMULATING THE NERVE FIBER WITH A SUCTION ELECTRODE ...................................................47 3.16 MOTOR UNIT ACTION POTENTIALS ..............................................................................................49 3.17 END PLATE POTENTIALS ..............................................................................................................52 3.18 MEASURING SARCOMERE LENGTH ..............................................................................................54 3.19 FORCE MEASUREMENTS ..............................................................................................................56 3.20 FORCE SENSOR SETUP ..................................................................................................................57 3.21 MUSCLE TWITCH FORCE ..............................................................................................................59

4 YOUR RESEARCH PROJECT .......................................................................................................63 4.1 PURPOSE ..........................................................................................................................................63 4.2 DELIVERABLE ..................................................................................................................................63 4.3 PRESENTATION TIPS .........................................................................................................................64 4.4 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

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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

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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.

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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 wafer-thin 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

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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.

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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!

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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.

Systemunder test

ResponseExcitation Measuringequipment

Stimulationequipment

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.

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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.

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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.

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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 BNC-to-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.

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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.

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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 frequency1. 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.

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(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?

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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

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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

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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. Set the trigger to Auto (automatic). 2. Ground the signal (GD button) and spin the position knob until you see it. 3. If in Normal mode, spin the trigger level control until the trace flashes across. 4. If in EXT trigger mode, make sure you have a good trigger signal by pressing TRIG

VIEW. 5. Push RUN/STEP and confirm trace is running by looking at the indicator in the top

left corner of the display area. 6. Push AUTOSET (top right). The scope will automatically attempt to set itself to

“good” settings. 7. Ask somebody at the next station. 8. Ask an instructor.

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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.

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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.

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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)

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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

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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?

2 mS20 mS

0.5 V

1 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.

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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. 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.

Headstage

Dagan preamplifier

Oscilloscope

Ele

ctro

de

Cell

Figure 3-10: Set-up for recording membrane potentials.

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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.

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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!

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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

Figure 3-18 Model cell coupled

to head stage.

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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 Z-Test 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

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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 Z-test 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.

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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 BF120-60-10 (O.D. = 1.2 mm, I.D. = 0.6 mm)

• Use Program #3 on the Sutter puller (heat = 490-5502, 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 filler3.

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.

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• 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.

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• 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.

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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.

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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/

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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).

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1

3

5 4

CP attachment

2

Figure 3-23: 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.

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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.

nerve

muscle

skin flap

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.

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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.

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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.

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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 micro-manipulator. 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.

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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

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.

TRIG

TRIG LEVEL

CH 1

CH 2

CH 1 = 200 mV/DIV, DC

CH 2 = 100 mV/DIV, AC

SWEEP = 5 mS/DIV, TRIG MODE = AUTO

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

++

=iNaik

oNaok

NaPKPNaPKPnFRTVm

][][][][ln*)/( (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

++

=ii

oo

NabKNabKVm

][][][][log5.61 10

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

++

=)23)(02(.140)120)(02(.][

log5.61 10oK

Vm

that approximates to

+=

1404.2][

log5.61 10oK

Vm (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.)

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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 70-110 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?)

Figure 3-31: Laser used for measuring sarcomere

length. Shine up through prep

L

laser

muscle

2x

θ

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

θλ sindn = 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.

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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 setting5.

Max load (gm) 50 Min resolvable load (mg) 2 Resonant frequency (Hz) 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.

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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.

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Figure 3-36: Prep for measuring muscle force

Figure 3-37: Close-up

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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

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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.

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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.

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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 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.

4.4 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 How MEPPs amplitude and frequency vary with

temp, [Ca++], .... Measure refractory period Length-tension curve (overall and sarcomere) Effect of temperature on anything Effect of pH, glucose, temperature Recruitment curve Force-frequency relation Doublets Effect of X on muscle force fusion frequency Effect of NMJ blockers on twitch time and amplitude Pharmacology of NMJ

AP threshold curve for pulse width, amplitude combinations Effect of X on MEPP

Effect of X on EPP Any of this week’s experiments, but done more carefully

Human muscle force assessment Your idea goes here

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Appendix A TEK SCOPE REFERENCE

Display area

SCOPE DISPLAY AREA 1 Icon shows acquisition mode 2 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.

3 Marker shows horizontal trigger position. Turn HORIZONTAL POSITION knob to adjust

4 Readout shows time between trigger marker and center graticule 5 Marker shows trigger voltage level

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6 Markers show zero voltage level of displayed channels 7 Down arrow indicates waveform is inverted 8 Readouts show vertical scale factors of displayed channels 9 Bw icon means channel has been bandwidth limited 10 Readout shows horizontal (timebase) scale 11 Readout shows window time base setting, if zoom window is in use 12 Readout shows trigger source 13 Icon shows trigger slope 14 Readout shows trigger level 15 Various operation messages 16 Readout shows trigger frequency CH1 (or CH2) MENU Controls appearance of channel traces. Hit twice to turn channel off. Coupling 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

BW Limit Always leave OFF Volts/Div Always leave Coarse Probe 1X if using BNC cables. 10X if using scope probe Invert Leave Off TRIG MENU Controls how scope is triggered Type Use Edge Source 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)

Slope Dictates edge of trigger Mode Use Normal or Auto Coupling 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.

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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

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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.

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Appendix B Sample plots

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).

5 ms

0.2 mV

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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?

1 ms

20 mV

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Appendix C Stimulator controls

The figures below show what each stimulator control sets. Single pulses

PI (1)

PA (4)

PW (3)

Del (2)

Synch

1. PULSE INTERVAL section, SECONDS control 2. PULSE DELAY section, SECONDS control 3. PULSE WIDTH section, SECONDS control 4. OUTPUT section, LEVEL control Double pulses

PI (1)

PA (4)

PW (3)

Del (2)

1. PULSE INTERVAL section, SECONDS control 2. PULSE DELAY section, SECONDS control 3. PULSE WIDTH section, SECONDS control 4. OUTPUT section, LEVEL control

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Pulse trains

TPI (3)

PI (1)

PA (5)

PW (4)

TD (2)

1. PULSE INTERVAL section, SECONDS control 2. TRAIN section, DURATION control 3. TRAIN section, PULSE INTERVAL control 4. PULSE WIDTH section, SECONDS control 5. OUTPUT section, LEVEL control

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Appendix D Frog Ringer's solutions

Composition of normal frog Ringer's (2.5 mM KCl) Compound mM gms for 5L KCl 2.5 0.94 NaCl 115.0 33.58 Na2HPO4 4.00 2.84 NaH2PO4 0.85 0.59 CaCl2 * 1.8 1.32 MgCl2 1.0 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.

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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] [Na] 1.0 116.5 5.0 112.5 10.0 107.5 25.0 92.5 50.0 67.5 100.0 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] gms NaCl gms KCl 1.0 6.8 0.07 5.0 6.6 0.37 10.0 6.3 0.74 25.0 5.4 1.86 50.0 4.0 3.72

100.0 1.0 7.45 8. Oxygenate by spinning. 9. Add 0.26 gms CaCl2 to each container. 10. Store overnight in refrigerator.

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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, wall-powered 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.

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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…

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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.

123456789

A BLoad (gms) Sensor (Volts)

0 0.2920 0.2210 0.2450 0.189

1.23 0.8761.23 0.7991.23 0.8531.23 0.825

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.

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Force sensor calibration

0

0.2

0.4

0.6

0.8

1

0 0.5 1 1.5

Load (gms)

Sens

or (v

olts

)

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

y = 0.489x + 0.2368R2 = 0.9877

0

0.2

0.4

0.6

0.8

1

0 0.5 1 1.5

Load (gms)

Sens

or (v

olts

)

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.

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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.

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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.

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Is*Rm

Is*ReRm*Cm

t

V

Voltage seen when a step change in current is applied to an electrode/membrane combination, or to the

model cell.

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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.

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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 Number Black 0 Brown 1 Red 2 Orange 3 Yellow 4 Green 5 Blue 6 Violet 7 Grey 8 White 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 kilo-ohms. 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-blue-green-silver is 5.6 ± 0.56 M-ohms and could be anywhere between 5.0 and 6.2 M-ohms.

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Appendix M NMJ Course Supplies and Drugs

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