The lab presented in this paper utilizes a proven four-step pedagogical framework (McLaughlin & Coyle, 2016) to redesign a classic Association of Biology Laboratory Education (ABLE) undergraduate lab (McLaughlin & McCain, 1999) into an authentic research experience on vertebrate four-chambered heart development and physiology. The model system is the chicken embryo. Through their research, students are also exposed to the embryonic anatomy and physiology of the vertebrate heart, the electrical circuitry of the developing heart, and the effects of pharmacological drugs on heart rate and contractility. Classical embryological micro-techniques, explantation of the embryo, surgical removal of the beating heart, isolation of the heart chambers, and more advanced tissue culture methods are also conducted. In this laboratory paradigm, students work in pairs to ask their own questions concerning the effects of two human cardiovascular drugs, denopamine™ and acebutolol™ on both in vivo and in vitro chicken embryonic heart rate and contractility, develop testable hypotheses based on information gathered from relevant scientific literature, devise and carry out a controlled experiment, and present the data in a professional scientific manner pertaining to a topic of clinical significance.

Introduction

Course-based undergraduate research experiences (CUREs) provide a way to increase and broaden participation of students in authentic research (Auchincloss et al., 2014; Bangera & Brownell, 2014), and train them in the essential elements of authentic research, which include reading scientific literature, generating questions (that don't currently have answers), forming hypotheses, designing experiments, collecting and analyzing data, working toward significant findings, and presenting results in both oral and written forms (Spell et al., 2014; Lopatto, 2003; Seago, 1992). A four-step pedagogical framework that includes all essential elements of authentic research has been developed that can serve to simplify and streamline the development and implementation process in an introductory biology laboratory setting (McLaughlin & Coyle, 2016). This framework has been shown to improve student perceptions of science and research, skills and knowledge levels in scientific research practices set forth by the National Research Council (2000), and confidence to effectively engage in scientific research practices including the ability to write strong scientific lab reports and read scientific papers (McLaughlin et al., 2017). These results are consistent with other studies which measured students’ perceptions and attitudes (Brownell et al., 2012; Harrison et al., 2011; Caruso et al., 2009; Weaver et al., 2008; Howard & Miskowski, 2005), ability to design experiments and interpret data (Myers & Burgess, 2003), and ability to develop information fluency (Gehring & Eastman, 2008) in laboratory courses that integrated authentic research-based experiences into the curriculum.

The chicken embryo is a classic model organism used to illustrate the principles of basic vertebrate embryology primarily because it is easily accessible, inexpensive, and direct observation of a living embryo is possible even at the early stages of embryogenesis. With the aid of a dissecting microscope, the key stages of heart development and the circulatory patterns can be observed in vivo, even when the chicken embryo is still attached to the yolk. Another advantage to using the live chicken embryo to demonstrate the principles of heart development is that the embryo can be removed from the shell and maintained in vitro in a temperature controlled environment for several hours while the beating of the heart, flow of blood, and the developmental stages accompanying the formation of the four chambers are even more clearly visible. In addition, the chicken embryonic heart is large enough for properly trained students to surgically isolate it from the embryo. Thus, students who work with this particular embryo in their development and/or physiology laboratories not only learn principles of vertebrate morphogenesis, organogenesis, and cardiac physiology, but are also exposed to simple, yet useful micro-manipulative techniques. These particular dexterous manipulations allow many of those who aspire to become physicians to try their hands (no pun intended) at surgical technique.

We thoroughly redesigned a classic cookbook Association of Biology Laboratory Education (ABLE) undergraduate lab on vertebrate heart development (McLaughlin and McCain, 1999) to incorporate the above highlighted four-step laboratory pedagogical framework. This allows students to carry-out a higher level inquiry-based, authentic research experience (McLaughlin & Coyle, 2016; Goedhart & McLaughlin, 2016; McLaughlin et al., 2017) using present-day cardiovascular drugs and updated tissue culture techniques and equipment. Students also master more advanced developmental biology, cardiovascular physiology, and pharmacology concepts.

Lab Protocol

Developmental and Physiological Aspects of the Chicken Embryonic Heart: Chronotropic and Inotropic Effects of Select Drugs

Objectives

The key objectives in this lab are three-fold: (1) to think critically about, and understand, how the four-chambered vertebrate heart develops using the chicken embryo as a model system; (2) to think critically about specific pharmacological agent(s) and their modes of action(s) at both the cellular and organ (heart) levels; and (3) to carry out a scientific research experiment. To accomplish these objectives, you will have eight weeks to carry out a four-step process wherein you will design and execute an authentic research project. This experimental framework is consistent with the manner in which professional research scientists design, execute, interpret, and communicate their experimental results. The over-arching experimental question that you will be attempting to answer through your research is the following: How do the cardiovascular drugs, denopamine™ and acebutolol™, affect the in vitro heart rate (chronotropic effect) and force of contraction (inotropic effect) of the developing isolated chicken heart?

Background Knowledge

The development of the four-chambered vertebrate heart involves a series of cellular migrations, fusions, and specific differentiations—that is, a multitude of morphogenetic events. Although gestation of the human and chick differ, their four-chambered hearts develop in a very similar fashion, and because of this the chicken embryo is used as a model system for human heart development. The hearts of the human and chicken embryos develop from the fusion of paired precardiac mesodermal tubes (aka “endocardial tubes”) located on either side of the developing foregut, on the ventral surface. These paired tubes begin to fuse at the anterior end (head) and continue to fuse posteriorly to form one continuous tube known as the tubular heart (Figure 1a). Once formed, the tubular heart is completely ventral to the foregut, and it has five distinct regions that can be identified from anterior to posterior: truncus arteriosus, bulbus cordis, primitive ventricle, primitive atrium, and sinus venosus (Figure 1b). Blood flows anteriorly, from the sinus venosus to the truncus arteriosus (future ascending aorta and pulmonary trunk). The heart begins to beat just after the paired heart tubes begin to fuse. It is the sinus venosus that becomes the future embryonic pacemaker.

Figure 1.

Illustration of human heart organogenesis from (A) a “tubular heart” to (B) an S-shaped tube with the prominent ventricle bulging to the right (Gray, 1918/2000). © 2000 copyright Bartleby.com, Inc.

Figure 1.

Illustration of human heart organogenesis from (A) a “tubular heart” to (B) an S-shaped tube with the prominent ventricle bulging to the right (Gray, 1918/2000). © 2000 copyright Bartleby.com, Inc.

The tubular heart then elongates on the right side, looping and bending to form an S shape, with the prominent ventricle bulging to the right (Figures 1b and 2). The heart continues to fold upon itself, moving the sinus venosus and atrium to a position anterior and dorsal to the ventricle and the bulbus cordis (Figure 3). The ventricle is now U-shaped and in the medial ventral position. The blood flows posteriorly and then makes a sharp turn to flow anteriorly.

Figure 2.

Head and S-shaped heart of a developing chick embryo at 1.5 days (~36 hours), viewed from the ventral surface (Gray, 1918/2000). © 2000 copyright Bartleby.com, Inc.

Figure 2.

Head and S-shaped heart of a developing chick embryo at 1.5 days (~36 hours), viewed from the ventral surface (Gray, 1918/2000). © 2000 copyright Bartleby.com, Inc.

Figure 3.

The developmental stages of a human heart beginning with the fusion of two endocardial tubes up to specific compartmentalization. Illustration of development of heart by OpenStax is licensed under CC by 3.0.

Figure 3.

The developmental stages of a human heart beginning with the fusion of two endocardial tubes up to specific compartmentalization. Illustration of development of heart by OpenStax is licensed under CC by 3.0.

The atrium then expands to the left in preparation for its division into the right and left atria (Figure 3). Although the heart still has two chambers at this time, communication between the sinus venosus and the atrium is via the right side of the atrium. This is the first step toward the sinus venosus becoming part of the future right atrium (SA node). The bulbus cordis will eventually give rise to parts of the ventricles, while the truncus arteriosus will become the future ascending aorta and pulmonary trunk. Eventually, when the atrium and ventricle have each divided into a pair of chambers, a typical four-chambered heart is present (Figure 3).

Laboratory Assignment

Below is a description of the four steps you will be following to carry out authentic research and to address the question at hand (Figure 4). Each step will require at least two weeks.

Figure 4.

The four-step pedagogical framework for authentic scientific research (McLaughlin & Coyle, 2016).

Figure 4.

The four-step pedagogical framework for authentic scientific research (McLaughlin & Coyle, 2016).

Step 1. Learn Essential Lab Techniques

During step one, you will learn essential and relevant in vivo and in vitro experimental techniques associated with both chicken embryo and heart manipulation via instructor(s) demonstrations and hands-on training. This step will include a small-scale and instructor-led exercise (use  Appendix 1 worksheet).

Step 2. Design an Experiment

You will work with your lab partner to research, select, and read primary literature articles (at least three) related to the above guided question in order to devise a more specific, self-directed research question. You will then formulate a hypothesis directed toward your specific question that is substantiated by the reference literature you select. Your next step is to design an experiment that attempts to test your hypotheses and incorporates the knowledge you have gained on your selected drug(s), using an appropriate control. Your experimental design must be written up as a traditional research protocol, including detailed procedures and data interpretation. You should consider the following when writing your protocol: (a) how you will administer the drug to an isolated chicken heart; (b) what concentrations you will use (you must have at least three different concentrations); (c) what will be your control; (d) what kind of data you will collect; (e) how you will collect the data; and (f) how you will represent these data in their final form. Your instructor(s) must review your protocol to ensure that it is adequately written, relevant, referenced, and executable within the allotted time frame before you are allowed to proceed with your experiment.

Step 3. Conduct the Experiment

Conduct this experiment using learned techniques while gathering data in your notebook. This will require coordinated work with your lab partner, and you will be expected to make good use of open laboratory time, in addition to scheduled class time. Your instructor(s) will be available for assistance or questions, but the expectation is that your work will be self-directed and paced appropriately.

Step 4. Interpret Data & Communicate Results

Review your results and interpret your data. If your data are puzzling or unclear, your instructor(s) can help you make sense of it. Once you are able to draw the conclusions of your experiment, you will present it to your instructor(s) in the form of a formal research paper or an oral presentation.

Techniques

Materials and Equipment (per student pairs)

  • chicken eggs: 3, 4, or 5 days old (half dozen eggs per week)*

  • chicken egg humidified incubator

  • CMRL (Connaught Medical Research Laboratories) media 1066 [Ward's Science]

  • glass dish (110 mm diameter) lined with cotton (2)

  • stock solution of R(–) denopamine™ [0.1mg/ml] Sigma Aldrich D7815

  • stock solution of acebutolol HCl™ [0.1mg/ml] Sigma Aldrich A3669

  • Falcon centrifuge tubes, 15 mL (6)

  • water bath

  • incubator for glass dishes and drugs

  • dissecting microscope (illumination from above and below the specimen) (2)

  • gooseneck lamp with 100 W bulb (2)

  • Syracuse culture dishes, 65 mm (2)

  • Delta T-EDU Culture Dish Control System [Bioptechs]

  • 20 G needle attached to a syringe

  • iris fine scissors (2)

  • iris micro-dissecting scissors

  • fine forceps (2)

  • micro-knife (hatchet)§

  • embryo spoon§

  • filter paper doughnuts§

  • transfer (beral) pipettes, 1.5 mL (5)

  • Beaker for egg waste (400 mL)

  • DI water squirt bottle for rinsing off embryological tools

  • Scotch Magic tape

  • stopwatch (digital or cell phone)

  • biohazard waste disposal

Notes for Instructors

*Fertilized eggs at specific days of development can be ordered and shipped in-state or out-of-state through Moyer's Chick Farm, Quakertown, PA (215-536-3155).

Any cardiovascular drugs with agonist/antagonist effect can be used in this experiment.

Contact Bioptechs, Butler, PA (420-050-4212) directly, if you would like to try an EDU system using chick embryos in your classroom and to discuss pricing.

§See  Appendix 2 for detailed instructions for making micro-knives, embryo spoons, and filter paper doughnuts.

  • Preparation of Chicken Egg and Bioptechs™ Equipment

    1. Order eggs (in advance) from a chicken farm to receive fertilized eggs of the appropriate day(s) (i.e., ages). Following pick-up, store eggs in a chicken egg incubator at 41°C until use.

    2. Place CMRL media in a 45°C water bath two hours prior to use. (CMRL media has been shown to be the optimal media for in vitro chicken heart experimentation in our lab.)

    3. Place all the glassware in incubator at 41°C two hours prior to use.

    4. Set up Bioptechs EDU Culture Dish Control System on one of the two dissecting scopes. Verify that EDU dish is inserted correctly and system is attached to the power supply.

    5. Add a small amount of warm CMRL media to the Delta T culture dish. Allow the culture dish about two minutes to reach selected 41°C (avian) temperature. Close the lid of the dish.

  • Windowing an Egg (modified from Cruz, 1993)

    1. Obtain an egg (day 3, 4, or 5) from the incubator and place horizontally on a glass dish lined with cotton (under a gooseneck lamp).

    2. Use Scotch Magic tape to tape along the center of the long axis of the egg, covering most of the top surface of the egg. Place two more pieces of tape on either side of the center piece (Figure 5a).

    3. Puncture the rounded end of the egg using a 20 G needle. Insert the needle pointing down into the egg. Withdraw 1–2 mL of albumen. This allows the embryo to move away from the upper surface of the egg, where you will be cutting out the window. Discard the albumen and rinse syringe with water.

    4. CAREFULLY puncture the tape-covered surface of the egg with the tip of your fine scissors. The location of the puncture should be about half an inch off-center (Figure 5a).

    5. Cut out an oval opening while pulling up with your scissors to keep them as far away as possible from the embryo and vitelline membrane (membrane around yolk). The size of the opening depends on the size of the egg, but it should be about 1.5–2.0 inches in diameter. Remove the shell cap with forceps, exposing the window (Figure 5b).

    6. If you are going to observe the embryo for more than a couple minutes while it is still in the egg, add several drops of CMRL media to the surface of the embryo to prevent dehydration. Do not add any CMRL media if you are going to immediately explant the embryo. This will prevent the filter paper doughnut from adhering to the vitelline membrane.

    7. Determine the in vivo heart rate of the embryo three times at 15-second intervals. Record the heart rate in beats per minute (bpm) in your lab notebook. [It is recommended that one student record the time using a stop-watch while another student counts heart beats of the embryo looking through the microscope.]

  • Explantation of a Chicken Embryo (modified from Cruz, 1993)

    1. Explantation refers to removal of an embryo from its normal in vivo environment and placement in a new location; in this case the embryo is transferred to a Bioptechs EDU Culture Dish. This is a useful method because it is much easier to manipulate or operate on an embryo when it is in a dish, rather than in an egg. To begin this process, add about quarter inch of warm CMRL media into a pre-warmed Syracuse dish. Then, place the dish on the stage of your first dissecting microscope and angle the gooseneck lamp as close to the dish as possible in order to keep the media warm. This is a critical step since the chicken embryo's normal body temperature needs to remain at 41°C.

    2. *Using forceps, gently place a filter paper doughnut around the embryo such that it frames the embryo. The filter paper will stick to the vitelline membrane, and the embryo will be exposed through the hole.

    3. Using surgical scissors, cut along the edges of the filter paper (vitelline membrane) off the surface of the egg while holding it with forceps. As you release the filter paper, the embryo will remain attached to it.

    4. Quickly transfer the filter paper doughnut along with attached embryo into a Syracuse culture dish already placed on the microscope.

    5. If the embryo doesn't stay attached to the filter paper, use your embryo spoon to collect it from the underlying yolk. Remember to replace the yolk that transferred over from this process with fresh CMRL media several times (Figure 5c).

    6. Remove the embryo from all attached extra-embryonic membranes and extraneous yolk using fine forceps (Figure 5d).

    7. Then, carefully transfer the embryo from the Syracuse dish into the Bioptechs EDU dish placed on your second dissecting microscope using fine forceps. Check to make sure the temperature is set to 41°C (avian). Add warm CMRL media, if needed.

    8. Determine the in vitro heart rate of the explanted embryo three times at 15-second intervals. Record the heart rate in bpm in your lab notebook. How does this heart rate compare with the in vivo heart rate?

    9. Use Atlas of Descriptive Embryology to identify the exact stage of development of your embryo. Record this in your laboratory notebook.

  • Isolation of the Heart (modified from McLaughlin & McCain, 1999)

    1. The explanted chicken embryo should be oriented ventral side up; this allows you to access the beating heart.

    2. Surgically remove the beating heart by cutting it above the bulbus cordis and below the sinus venosus of the atrium (Figure 5e and 1b). Take a picture of the isolated heart.

    3. Determine the in vitro heart rate of the isolated heart three times at 15-second intervals. Record the heart rate in bpm in your lab notebook.

  • Administration of Cardiovascular Drugs

    1. Prepare the serial dilutions of the drug(s) that you chose to work with. Make each dilution in a separate Falcon centrifuge tube using the warm CMRL media as your solvent. Store your labeled tubes in the rack at 45°C water bath or incubator.

    2. Design table(s) similar to the ones in  Appendix 1, wherein you adequately record all of your heart rate data per your experimental design. Remember to include both in vivo and in vitro embryonic heart rates (control baseline heart rates) before you exogenously expose your embryos to any drugs.

    3. Remove as much CMRL media bathing the isolated heart as possible using transfer pipette, and add 1 mL of the lowest concentration of the drug to the isolated heart in the dish (Figure 5e). Wait at least 30 seconds for it to equilibrate, then record the heart rate (three times at 15-second intervals), noting any arrhythmias (such as tachycardia, bradycardia, atrial flutter, fibrillation, etc.) or changes in heart contractility (variation in force of contraction in any or all chambers). Remove the drug as best as you can from the Bioptechs EDU dish using a transfer pipette, then immediately add the next higher dilution of the drug. Repeat the procedures in this step to obtain in vitro heart rate for all your drug dilutions.

    4. It is imperative that you work up at least five embryos per the age group(s) selected. For example, if you chose to observe the effect of acebutolol™ on the 4- and 5-day old isolated chicken hearts, you will need to work up five 4-day-old eggs and five 5-day-old eggs. Work up your embryos one at a time, repeating Steps II–VI and recording the control heart rates at all stages.

    5. Carry out your experiment!

Figure 5.

The isolation of heart from a 5-day-old chicken embryo illustrated in a series of six steps: (A) preparing to “window an egg” using three vertical strips of tape placed across the long axis of an egg; (B) cutting an oval opening using fine scissors to expose the embryo; (C) “explantation of the embryo” within its amniotic membrane using embryo spoon; (D) removal of amniotic and allantois membranes with fine forceps and/or micro-dissecting scissors in Bioptechs equipment; (E) surgical “isolation of the heart” from the embryo using fine forceps and micro-dissecting scissors/micro-knife; and, (F) “administration of drug” to the isolated heart using plastic transfer pipette.

Figure 5.

The isolation of heart from a 5-day-old chicken embryo illustrated in a series of six steps: (A) preparing to “window an egg” using three vertical strips of tape placed across the long axis of an egg; (B) cutting an oval opening using fine scissors to expose the embryo; (C) “explantation of the embryo” within its amniotic membrane using embryo spoon; (D) removal of amniotic and allantois membranes with fine forceps and/or micro-dissecting scissors in Bioptechs equipment; (E) surgical “isolation of the heart” from the embryo using fine forceps and micro-dissecting scissors/micro-knife; and, (F) “administration of drug” to the isolated heart using plastic transfer pipette.

Notes for Students

*Use a filter paper donut on 3- and 4-day-old embryos; use an embryo spoon only on 5-day-old embryos.

This step is preformed following the approval of your lab protocol by your instructor(s).

Important: Discard egg shells, embryonic waste and embryos in appropriate biohazard containers.

Conclusion

The simplicity and flexibility involved in the four-step pedagogical laboratory framework allows it to be easily adopted for use within the unique infrastructure and resourceful environments at a variety of institutions and at different levels of biological study, effectively increasing student access to authentic scientific research. For example, this pedagogical framework was used to successfully transform a sophomore-level, introductory, cell biology laboratory for majors at a 4-year college branch campus (McLaughlin & Coyle, 2016) and an honors, introductory biology laboratory for non-majors at a 2-year college (Goedhart & McLaughlin, 2016) into authentic research experiences. For the former, students investigated various factors affecting the growth and viability of a mammalian cell culture line, Vero cells, and related their findings to a current issue in the field of cell biology, nutritional and/or physical stress of cells. For the latter, students selected and investigated factors affecting microalgae cell growth and related their findings to a real-life application of social significance. A short video documenting the use of the pedagogical framework from both instructor and student perspectives, From Cookbook to Critical Thinking (https://vimeo.com/118326855), was created by Citrus College and published on Vimeo, Inc. In both laboratory paradigms, students worked within groups to learn modern cellular biology techniques, ask their own questions, develop testable hypotheses based on information gathered from relevant scientific literature, devise and carry out a controlled experiment, and present the data in a professional scientific manner.

The lab presented in this paper also utilizes the four-step pedagogical framework, but this time to transform a sophomore-level, introductory, developmental biology laboratory for biology majors. Students investigate the development of the vertebrate four-chambered heart using the chicken embryo as a model system, while simultaneously being exposed to the embryonic anatomy and physiology of vertebrate heart, the electrical circuitry of the developing heart, and the effects of pharmacological drugs on heart rate and contractility. Classical embryological micro-techniques and procedures are conducted as well. Student, working in pairs, devise a hypothesis concerning the effects of two human cardiovascular drugs, denopamine™ and acebutolol™, on in vitro heart rate and heart contractility. These drugs were selected for their known agonist or antagonist effects on the human heart rate. Other drugs that have been successfully used in this lab to substitute these drugs include dobutamine, verapamil, ractopamine, epinephrine, nicotine, and pseudoephedrine. Indeed, the list of drug substitutions is endless, and as such, allows the instructor(s) semester-to-semester versatility while maintaining the authentic research nature and, as such, enhanced performance and engagement of students (lack of plagiarism). To best exemplify the authentic research nature and enhanced performance of students who have experienced the chicken lab in particular is this reality: Many students have presented their work at peer-reviewed conferences or have published their research (e.g., Gonzalez et al., 2015).

It is important to point out the following when utilizing the framework: Instructors should act as research chaperones, guiding student scientists through each step, providing constant feedback and environments that allow time for student self-reflection, mistakes, and dialogue over assignments before their final submission for grading (i.e., protocol, notebook, scientific paper, poster, etc.). Additionally, the four steps outlined herein should scaffold the scientific process, allowing students who are novices to the scientific process to progressively gain familiarity and comfort with the essential elements of scientific inquiry. Lastly, it is recommended that instructor(s) read how this framework was used by other instructors to rework the semester with regard to time and workload (McLaughlin & Coyle, 2016; Goedhart & McLaughlin, 2016).

We would like to extend our gratitude to Michelle Lynn for sketching a realistic illustrations of the chicken heart isolation procedure. We would also thank Dr. Elizabeth McCain, who worked tirelessly with Dr. Jacqueline McLaughlin on publishing the original version (ABLE) of this lab.

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APPENDIX 1 Examining the Heart Rate of the Embryonic Heart

  1. In your lab notebook, using an embryological atlas like Atlas of Descriptive Embryology, diagram and thoroughly label 3-day-old and 4-day-old chicken hearts (whole mounts).

  2. Window a 3- or 4-day-old egg, then determine and record the in vivo heart rate three times at 15-second intervals in Table 1.

  3. Explant the embryo out of the egg, then determine and record the in vitro heart rate of the embryo three times at 15-second intervals in Table 1.

    • How does this heart rate compare to that of in vivo heart rate?

  4. Surgically isolate the heart. Observe the beating of the isolated heart, then determine and record the in vitro heart rate of the heart three times at 15-second intervals in Table 1.

    • Where does the beating begin and end? Be precise.

    • Is the isolated heart rate similar or different from the heart beat of the explanted, in vitro embryo?

    • Draw a picture of the isolated heart and label each region.

  5. Use the micro-knife or iris micro-dissecting scissors to separate the four regions of the isolated heart (bulbus cordis, sinus venosus, atrium, and ventricle). Observe each isolated region of the heart, then determine and record the in vitro heart rate of each region three times at 15-second intervals in Table 2.

    • Does each region have an intrinsic heart beat? If so, are they synchronous?

    • Does each region beat at the same rate as the in vivo or in vitro explanted heart?

    • Draw a picture of the isolated regions of heart and appropriate labels.

Table 1.
Summary of in vivo and in vitro heart rates of embryo and isolated heart.
In vivoIn vitro
EmbryoExplanted EmbryoIsolated heart
HR (15 sec)HR (bpm)HR (15 sec)HR (bpm)HR (15 sec)HR (bpm)
Trial 1       
Trial 2       
Trial 3       
Average       
In vivoIn vitro
EmbryoExplanted EmbryoIsolated heart
HR (15 sec)HR (bpm)HR (15 sec)HR (bpm)HR (15 sec)HR (bpm)
Trial 1       
Trial 2       
Trial 3       
Average       
Table 2.
Summary of in vitro heart rates in each separated region of heart.
In vitro
Bulbus CordisSinus VenosusAtriumVentricle
HR (15 sec)HR (bpm)HR (15 sec)HR (bpm)HR (15 sec)HR (bpm)HR (15 sec)HR (bpm)
Trial 1         
Trial 2         
Trial 3         
Average         
In vitro
Bulbus CordisSinus VenosusAtriumVentricle
HR (15 sec)HR (bpm)HR (15 sec)HR (bpm)HR (15 sec)HR (bpm)HR (15 sec)HR (bpm)
Trial 1         
Trial 2         
Trial 3         
Average         

APPENDIX 2 Directions for Making Embryological Tools

Micro-knives (modified from Tyler, 1994)

  1. Break a single-edge razor blade into small fragments with the pliers or tin snips, making sure that each fragment has the cutting edge on it. Avoid damaging this edge. The fragment should be between 5 and 10 mm long.

  2. Soak one end of the wooden applicator stick (2 mm diameter; 100–150 mm long) in water for about 5 minutes to soften the wood. Make a 5-mm split down the center of the moistened end of the stick with a fresh razor blade. Do not splay out the two sides of the split; just create a crack in the stick large enough for the razor blade fragment to fit in.

  3. Carefully insert the rough edge of the razor blade fragment into the split at 45° angle to the axis of the stick. You can't use the pliers to do this because they will damage the cutting edge. You must use your fingers—be careful! The cutting edge of the blade should not be embedded in the wood at all, and you should be able to comfortably hold the stick like a pencil and have the razor's edge almost flat with a desk surface.

  4. Let the wood completely dry. Paint the split edges with super glue to firm the placement of the razor blade fragment.

  5. Store the micro-knives such that their edges are protected. One method is to stick Styrofoam peanuts on the razor edge.

Note: Micro-knives can only be used for one lab period; they dull quickly.

Embryo Spoon (modified from Tyler, 1994)

  1. Heat an insect or dissecting pin over a flame and puncture 8–10 holes in the bowl area of the plastic spoon (ice cream spoon), as shown in the picture.

  2. Use the sandpaper (220 or 400 grit) to remove the sharp plastic edges.

Filter Paper Doughnuts

  1. Fold a piece of filter paper (Whatman #1, 3.2 cm diameter) in half, then cut out a semi-circle in center of the filter paper, keeping 0.5 cm distance between two circles.

  2. The end result should be an oval-shaped doughnut.