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TEACHING IN THE LABORATORY
Department of Biological Sciences, Idaho State University, Pocatello, Idaho
Address for correspondence and present address of P. A. Cotter: Dept. of Biological Sciences, 3211 Providence Dr., Anchorage, AK 99508 (e-mail: afpac1{at}uaa.alaska.edu)
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Abstract |
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Key words: fish; cardiac physiology; electrocardiography; electrocardiogram
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Introduction |
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A review of the literature shows that while cardiac physiology receives much attention in teaching laboratories, the number of nonmammalian, vertebrate models that permit experimentation is limited. As the use of living specimens provides unique insights into physiology (10), it is important to develop effective laboratory protocols to study cardiac physiology. Knight et al. (7) detailed techniques for studying electrical and mechanical activity of the frog heart in vitro, and Yoshida (18) described a wide range of cardiac physiology studies using an in situ preparation of frog hearts. Mammalian models are often restricted to graduate or medical school laboratories due to cost, student apprehension and preparedness, and growing resistance to the use of mammalian subjects (9, 13). Furthermore, most of the mammalian cardiac physiology preparations published in Advances in Physiology Education are designed for graduate- or medical school-level demonstration, not inquiry-based undergraduate laboratory exercises (10, 13).
In undergraduate laboratories, cardiac function and regulation are often studied using invertebrate (e.g., clams or other available invertebrates) or student subjects. Although many important physiological principles can be explored with these systems, they do have limitations. Students may not relate well to invertebrate animals and may not understand their anatomy, physiology, and natural history. Therefore, despite the fact that some invertebrate responses to cardiac manipulation are consistent with mammalian themes of cardiovascular regulation, some fundamental aspects of invertebrate cardiac function may not be clear in students’ minds. Furthermore, collecting electrocardiograms (ECGs) from invertebrates may present technical challenges beyond the scope of the undergraduate laboratory. The use of human subjects has the opposite problem and an added challenge: students typically relate well to their own bodies, tend to be quite interested in how their bodies work, and data collection is straightforward, but only rudimentary manipulations can be done, and it is difficult to control variables.
A desirable animal model for studying vertebrate cardiovascular physiology should 1) be easy to handle and anesthetize; 2) allow for the easy administration of drugs that may or may not have specific cardiovascular effects; 3) be manipulated by means other than drug administration (temperature, salinity, hypoxia, etc.); and 4) have a well-documented natural history to make relevant conclusions concerning evolution, development, ecology, and physiology. Furthermore, survival of the experimental animal is often a concern for students and instructors. Fish possess many of these qualities, yet there are only a few published physiology laboratory protocols involving fishes (3, 11, 12). Extensive scientific literature exists that characterizes the cardiovascular responses of trout (and other fishes) to a variety of environmental, drug, and anesthetic perturbations. These publications make the use of this laboratory exercise attractive in courses stressing small research projects with publication-style laboratory write-ups.
At Idaho State University, we used farm-reared (Clear Springs Foods, Buhl, ID) rainbow trout (Oncorhynchus mykiss Walbaum) in our undergraduate animal physiology laboratory to study cardiac function and regulation. Trout are relatively easy to maintain, and they readily adapt to laboratory handling and treatment. This laboratory supplements a more-traditional human ECG laboratory, allows a broad range of student-directed experimental manipulations, and promotes both active and cooperative learning environments. Students have been surprised to learn that ECG tracings from trout are quite similar to mammalian tracings; this similarity helps students recognize the relevance to humans/mammals. In addition, students gain experience with animal care and how to safely and effectively anesthetize and recover an animal. All protocols were approved by the Idaho State University Animal Care and Welfare Committee.
Materials and Methods
Idaho is currently the nation's largest commercial producer of rainbow trout, growing 75% of the estimated 70 million pounds for the food fish market. In our studies, we used domesticated rainbow trout across age, sex, and size classes (>100 g). Trout were reared in constant flow spring water (14°C) raceways at Clear Springs Foods before being transferred to the Aquatics Research Facility at Idaho State University. Before being studied, fish were held in 1,000-liter circular tanks containing recirculating, filtered, dechlorinated, and UV-sterilized water at 14°C and were fed every other day (1% of body wt). Further details on the maintenance of fish for teaching and research can be found in Stickney and Kohler (14).
Student profile and background. This laboratory was part of an upper-division Animal Physiology course consisting mostly of biology and zoology majors at the junior or senior level. Student backgrounds were diverse, but all students possessed a foundation in biological, chemical, and physiological principles. Few students had prior experience in handling animals in a research or laboratory setting. Prior to this laboratory, students were introduced to the fundamentals of cardiac function, electrophysiology, and ECG waveforms in nonlaboratory class meetings. Principles of electrocardiography were reinforced in the first of the two-session sequence, when students could explore practical and theoretical aspects of obtaining ECGs from "practice" fish (see ECG laboratory schedule). Each laboratory section had 10–20 students and was successfully taught by one instructor and a teaching assistant.
ECG laboratory schedule. Our ECG exercise lasted two 3-h laboratory periods. During period one, students learned cardiac anatomy; ECG theory and practice; ECG software; how to handle, anesthetize, and recover fish; how to place electrodes for optimal data acquisition; and data analysis. Additionally, students were presented with a selection of variables (see below) from which they generated questions to address in the subsequent laboratory period. Each small group (3–4 students) wrote a one- to two-page proposal outlining the question and experimental methods; the proposal was reviewed by the instructor prior to the experiment. In the second laboratory period, students conducted their experiments. Appropriate laboratory protocols, behavior, and attire were taught to, and expected of, students in both laboratory sessions.
General setup.
There are a variety of ways to set up the procedure. For all, be sure to 1) adequately oxygenate or aerate the recirculating water source, 2) maintain adequate water flow across the fish's gills [
600 ml·min–1·kg body wt–1 is sufficient in trout (3)], and 3) keep the fish's skin moist using wet paper or a continuous stream of water whenever fish are maintained out of water. We used a plastic utility cart in which we drilled a hole in the top to allow for return drainage to the anesthetic bath reservoir (Fig. 1). We affixed a small drain or funnel to the underside and attached a short section of tubing to prevent spills. Below the drain sits a refrigerated anesthetic reservoir. Trout were acclimated to 14°C, and experiments always began at the acclimation temperature. Alternatively, we have used plastic/fiberglass tanks with or without a standpipe to record ECGs for immersed fish. For either setup, we used a small submersible pump (model NK-1, Little Giant Pump, Oklahoma City, OK) and attached flexible tubing to bring water from the anesthetic bath reservoir to the fish (Fig. 1). Tubing size and type should be chosen to match the size of fish and the arrangement of the experimental setup. To promote a stable ECG, sufficient and continuous irrigation of gills should be maintained, and bright red gills should be observed. We typically split the water line with Y-connectors and supply two fish (for 2 groups of students) on each table from each anesthetic reservoir. This system reduces material investment and provides replication of treatments (e.g., anesthetic concentration, water quality, etc.).
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Anesthetic administration and fish recovery. This laboratory is straightforward and, depending on resources, many substitutions and modifications are possible. Regardless of what modifications are made, it is important that 1) appropriate anesthetic doses are established for the species and conditions prior to using the lab in the classroom, 2) continuous and visible flow of water across the gills is maintained and gill color is noted, and 3) at least one student in each group is designated to recover fish. We have found that enthusiasm for the fish's welfare is high, and students readily volunteer to be responsible for recovering fish.
Approximately 20 liters of water-buffered anesthetic solution should be prepared prior to the experiment (see Table 2 for anesthetic measures), and its temperature should be stabilized. Our trout were held at 14°C; however, the choice of experimental temperature will depend on the acclimation temperature of the fish used. A submersible or in-line pump and associated tubing (enough tubing to reach from the anesthetic receptacle to the fish's mouth when on the table) should be plugged in to circulate the anesthetic solution within the anesthetic container. After pumps and tubing are in place, place a fish in the anesthetic bath and observe the fish as it becomes anesthetized. Using the anesthetic concentrations listed in Table 2, trout are fully anesthetized within 1–3 min. After the fish has lost equilibrium and opercular movements have slowed, but not stopped, transfer fish to the V-shaped holder, ventral side up, and immediately insert the open end of the tubing into the fish's mouth. The elapsed time to notable stages of anesthesia (e.g., loss of equilibrium and loss of response to stimuli; Tables 3 and 4) should be recorded; these may be important data points for discussion with other groups or used to compare values between fish undergoing anesthesia. To keep tubing secured in the mouth, simply cut a slit in a small square of open-cell foam, slide the tubing through the foam, and place the foam in the fish's mouth. Flow volume can be regulated using Y-connectors and/or tube clamps.
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ECG leads and placement.
Three bipolar shielded electrodes are used to record ECGs. Various placement regimens can be used, and some preliminary experimentation may be required. We obtained the best results with the animal's right side electrode (+) midway and on the extreme lateral aspect of the isthmus. The left electrode (–) was inserted 1 cm posterior to the right electrode but on the opposite side of the isthmus. A third "ground" electrode was inserted 
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down the body of the fish (Fig. 2). Before each electrode is inserted, it is helpful to pierce the skin with a 27- or 30-gauge (preferred) needle and then insert the electrode into the needle hole subcutaneously (do not insert the electrode vertically or you may pierce the pericardium and heart). After only a few trials, electrode placement and insertion becomes quite easy. Changing electrode locations may be necessary to obtain a clean ECG signal (Fig. 3). Some signal filtering can be performed to smooth the tracing; however, students may not have the adequate background or comfort to take advantage of the signal-processing capabilities of the data-acquisition system. The above procedure is sufficient for ECGs recorded from fish positioned ventral side up (submerged or emerged). If you wish to turn the fish dorsal side up, we recommend suturing each needle electrode in place (see below).
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Procedures and Data Collection
An organized approach to data collection should be encouraged and facilitated. Although groups will perform different experiments, there is much value in collecting data that can be compared among groups (e.g., time course of anesthetic administration and recovery). We have included a sample data collection sheet that could be easily adapted for classroom use (see Table 4).
There are several possible treatments to use and experiments to perform with this protocol. The following is not an exhaustive list but represents variables we have used or suggested in the student and/or research laboratory. For each variable below, we provide possible questions to address experimentally in an undergraduate laboratory; the dependent variable chosen might depend on your learning outcome for the laboratory and the experience of the students.
Anesthetics. There are a wide variety of anesthetics available, and Summerfelt and Smith (15) provide an excellent review of their characteristics and use guidelines. We have used clove oil (prior to restrictions of its use by the United States Food and Drug Administration), tricaine methanesulfonate (MS-222, Crescent Chemical, Phoenix, AZ), and benzocaine (Sigma Chemical, St. Louis, MO). We determined effective doses by selecting concentrations resulting in both 1) an induction of deep anesthesia within 100–200 s and 2) maintenance of anesthesia at the same dosage. Varying the anesthetic concentration can be easily accomplished.
Sample anesthetic questions are as follows:
Temperature. Temperature trials are quite successful and provide a great opportunity for students to see how ectothermic animals respond to acute temperature fluctuations. Our trout were acclimated to 14°C; we do not permit students to warm them higher than 20°C. We used ice packs to maintain/modify temperature. It is important to use ice packs (or plastic containers filled with ice) rather than raw ice to avoid diluting the anesthetic bath or adding chlorinated water and thus confounding the results. We have successfully used a circulating water bath fitted with a closed system heat-exchange apparatus (an aluminum coil fitted to a temperature-controlled recirculating water bath) in the research laboratory, but due to time and financial constraints we have not used it in the teaching laboratory. If temperature is to remain constant, care must be taken to monitor temperature and respond to slight fluctuations by the removal and addition of ice packs.
Sample temperature questions are as follows:
Oxygen supply. Alterations in cardiac electrophysiology due to hypoxia can be explored. The oxygen concentration of the anesthetic bath can be manipulated by bubbling nitrogen, air, or oxygen through the bath. An oxygen probe could be employed to closely monitor the anesthetic bath oxygen concentration. Alternatively, recording ECGs during brief periods of mouth tube removal may also induce hypoxic effects. A comparison of cardiac responses to these two techniques may be used to distinguish environmental hypoxia from water withdrawal. Care must be taken, and the welfare of the fish should be the primary concern when experimenting with hypoxia. Compared with other fishes such as the carp, catfish, goldfish, or eel, trout (and their hearts) are hypoxia intolerant.
Sample oxygen concentration questions are as follows:
Emersion/submersion. Fish can be maintained indefinitely out of water as long as a constant flow of water passes across the gills and the skin surface is kept moist with flowing water or a wet paper towel. If the procedure is performed in a container, a standpipe can be added and the water level can be adjusted to assess whether submersion/emersion affects ECG. Once again, it is very important to artificially irrigate the gills via the aeration hose even during submersion; deeply anesthetized fish may not be able to adequately pump water across their gills.
Sample emersion/submersion questions are as follows:
Orientation. Fish orientation in submerged or emerged states can easily be studied. Extensive manipulation of the anesthetized fish often necessitates suturing electrodes to the body surface. We used 1-0 metric Prolene monofilament suture (Ethicon, Somerville, NJ), but many other sutures are sufficient. Scales can make piercing fish skin very difficult, especially in larger animals. It is often necessary to remove scales, by gentle scraping, in the suturing area before the skin of the fish is pierced.
Sample orientation questions are as follows:
Salinity/osmotic variation. Moving freshwater-acclimated brook char (Salvelinus fontinalis) into a higher salinity environment has been has been used to achieve slight osmotic stress (3). Although we have yet to conduct such experiments, we suggest that osmotic variation experiments be performed using a range of salinities appropriate for the species and its osmotic history. Pure NaCl could be added to the anesthetic bath to achieve concentrations that are within the accepted osmoregulatory abilities of the fish species studied. Salinity can be measured using electrical conductivity, water density, or vapor osmometry.
Sample salinity/osmotic variation questions are as follows:
Environmental pH. Exposing the anesthetized animal to varying environmental pH while recording ECGs can be done using 0.1 N H2SO4 for lowering pH and 0.1 N NaOH or KOH for increasing pH (5). The reversal of pH or returning pH to neutral/beginning values can be easily done by back titrating with H2SO4 or NaOH/KOH; this may be of interested to test whether electrocardiographic changes correlate with pH modifications or time under anesthesia. Modifications in pH should be done slowly and monitored carefully so as not to overshoot the intended pH. Note that the presence of a buffer in both the tricaine and benzocaine solutions may influence the ability to alter pH.
Sample pH modification questions are as follows:
Electrical axis: Two-dimensional axis determination can be performed. This exercise helps students conceptualize not only electrophysiology of the heart but also the methods employed to understand it. It is crucial that students understand the basics of the ECG and the physiological/analytical basis of positive and negative deflections in the ECG (18). See Ueno et al. (16) and Yoshikawa et al. (19) for details of axis determination in fish.
A sample electrical axis question is as follows:
Analysis
Analysis of recorded ECGs depends on many factors including the capabilities of the recording/analysis equipment, student/faculty preparedness and willingness, and focus of the laboratory. Certainly, how heart rate varies under various treatments is of interest. New computer-based ECG systems, however, permit detailed analysis of interval (e.g., QT), segments (P-R and S-T), wave durations (P and QRS), heart rate variability, and signal voltage (e.g., QRS amplitude). It may be of interest to study how some of these dependent variables influence others. For example, the mammalian QT interval is typically inversely related to heart rate, but not in a linear fashion (1). Our study (2), however, has suggested that the QT interval and heart rate are independent in anesthetized rainbow trout.
This laboratory can be used for students at all experience levels. It provides a basis for developing experimental design skills, statistical and physiological analysis and hypothesis testing, detailed investigations in comparative cardiovascular physiology, and learning how to apply nonmammalian experimental models to mammals. For more advanced students, a vast biomedical literature base of ECG analysis allows students to study novel questions and compare their results with other studies on fish and/or other vertebrates.
Trout heart anatomy. Similar to the mammalian heart, the heart of teleost fish is enclosed within a pericardium, consists of contractile cells (cardiomyocytes), and develops a characteristic impulse that propagates across the chambers. Unlike the mammalian heart, the trout heart comprises four chambers in series. The thin-walled sinus venosus receives venous blood and contains pacemaker cells, which are responsible for initiating the heartbeat. Blood leaves the sinus venosus and enters the single atrium, a slightly muscular chamber capable of generating force when electrically stimulated. The thick-walled ventricle provides the force to propel blood to the gills and through the circulation. Its anatomy shows considerable variability across fish species. The last chamber, the bulbus arteriosus, is a heavy-walled elastic chamber that attenuates the pulsatile blood flow leaving the ventricle. All chambers are separated from one another by passive, one-way valves.
Many students are surprised to observe the similarities of fish and mammalian ECGs, especially when the anatomical differences of the two hearts are noted. It should be explained that despite the paired atria and ventricles in mammals, each of these pairs produce a single waveform on the ECG (i.e., a P wave for the combined depolarization of the atria and a QRS complex for the combined depolarization of the ventricles). In fish, the single atrium and ventricle produce a single P wave and QRS complex, respectively.
Concluding Remarks
In a department capable of maintaining captive fish and performing ECGs, this laboratory requires no real complex or expensive equipment and is applicable to cardiovascular studies across a wide spectrum of complexity. It is easily modified to meet the instructor's desired outcomes and can be used in single or multisession laboratories. Unlike ECG laboratory exercises involving humans, this laboratory facilitates well-controlled experimental manipulations and promotes both active and cooperative learning environments. Furthermore, scientific inquiry, study design, following and/or developing scientific protocols, coordination of a research team, and animal care are also emphasized.
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Acknowledgments |
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Received for publication September 17, 2006. Accepted for publication February 23, 2007.
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