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Cardiac physiology and function differ between organisms, as you can imagine the physiology is more complex within developed as opposed to that of simpler ones. Fish, for example have simpler hears consisting of two chambers, no valves and have a low-pressure system. Adult amphibians such as frogs or salamander have a more complex hear consisting of three chambers The level of complexity increases as we discuss the heart of adult amphibians. The adult amphibians have three chambers in their heart. Their heart has a spongy vent that avoids the mixing of blood. Reptiles have a higher complexity. Reptile’s hearts consist of vents that are divided into sub chambers. These sub chambers keep oxygen and deoxygenated blood separated. This separation is done by pressure difference. The mammalian heart is the most complex of the structures. The mammalian heart has four chambers. The mammalian heart also circulates in series which is significant because it doesn’t allow for oxygenated blood to mix with deoxygenated. The simplest of these hearts is the Procambarus clarkii.
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While the complex mammalian heart has four chambers and the simple fish heart has two chambers, the P. clarkia, also known as the Crayfish, has a single chambered heart that looks like a sac. The crayfish crustacean heart is made up of muscular leaflets that layer the heart. The crustacean heart is suspended in the pericardial cavity. The crustacean heart is specifically located in the dorsal portion of the cephalothorax. The heart is located there for maximum protection. The heart has many arteries; however, the heart has no veins. The blood, which in the crayfish is known as the haemolymph, flow into the heart through an opening called the ostia. The ostia is an opening that leads the haemolymph into the ventricle. The haemolymph then goes through the ventricle and pumped into the arteries to the rest of the body. What causes the heart beat rhythm in the crayfish is the cardiac ganglion.
There are different methods of manipulating the heart rate for an organism; however, the methods are dependent on what causes the heart to beat. If the heart rate is dependent on the muscle, which is called myogenic and is found in many vertebrates, then manipulation of the cardiac muscles is necessary. The crayfish, which is neurogenic, meaning that contraction of the heart is dependent on neurons, then neurotransmitters are necessary to manipulate the heart rate. Serotonin, acetylcholine, and epinephrine will be used in this experiment to manipulate the heart rate.
Serotonin is a neurotransmitter that the invertebrates are familiar with because serotonin is a natural neurotransmitter in invertebrates that contracts the heart and increases the speed of the heart rate. Acetylcholine is found in to increase heart rate and contraction in invertebrates. Acetylcholine serves a different purpose in vertebrates. In vertebrates, acetylcholine decreases the heart rate. Epinephrine increases the heart rate in both invertebrates and vertebrates. Epinephrine increases the heart rate in both because epinephrine helps regulate blood pressure control and vasoconstriction (Epinephrine, n.d.). Temperature is another manipulation that we can do. A decrease in temperature would decrease the heart rate in invertebrates and an increase of temperature would increase the heart rate.
We know that all the neurons will stimulate an increase in heart rate activity, however, we hypothesize that the serotonin and acetylcholine will increase the heart rate at a constant rate. We also hypothesize that epinephrine will increase the heart rate the most because of its role in the sympathetic nervous system.
Materials and Methods
In this experiment, there are 9 sections that will be done. The sections will be broken up to further organize the methods. All the materials, however, will be given together. The following materials are needed for the entirety of the experiment: Crayfish, Fine insulated copper wire, Matches, Foam pad, Tupperware container, Aluminum foil, 18 gram needle, Cyanoacrylate adhesive, Zip kicker cyanoacrylate accelerator, Cotton tipped applicators, Freshwater crustacean saline, Bulb thermometer, 100 ml graduated cylinder, 1 ml micropipettor, blue micropipettor tips, Impedance converter, 9 V battery, 1 mM Serotonin, 1 mM Acetylcholine, 1 mM of Epinephrine.
All of these materials are necessary for the experiment. There experiment is broke up into 9 sections. These sections include preparing the Impedance converter, the Crayfish preparation, getting the baseline heart rate of the Crayfish, finding the stressed heart rate, manipulating the heart rate using Serotonin, Acetylcholine, Epinephrine, and Temperature, and lastly, collecting the data.
For the Impedance Converter, there are controls that are necessary to know. Those controls include the following, On-Off-Test, Balance, AC-Long/Short, Size, Input, Calib. On-Off-Test turns the instrument on or off and tests the quality of the batter. Balance adjusts the Impedance Converter oscillator by a precise 10-turn potentiometer. For proper operation the needle must hover around the zero. Adjust the instrument to zero to accommodate any major change in impedance. AC-Long provides a coupling constant of 1 second. This means that all signals with frequencies longer than 1 second are cut off. AC-Short provides a coupling constant of 0.1 seconds. AC-Short cuts off all signal frequencies longer than 0.1 seconds. Size is a knob that adjusts amplitude of the signal. Input is our source impedance (i.e. Crayfish) connected by electrodes plus the wires to the green binding posts on the back of the instrument. Lastly, the Calib is a switch connected to a 0.25 ohm resistor. Depressing this switch shorts out the resistor which changes the 0.25 ohm. Normal physiological impedance changes range from 0.2 ohm (cardiograph) to 5 ohm (respiration.)
For use, connect the output of the Impedance Converter to a recording device. Then attach the electrode cables to the green input binding post. Next, place the electrodes on either side of the biological structure of which you are measuring impedance. Try to make them as secure as possible without harming your subject. Lastly, adjust the balance to hover around zero and begin recording data.
To prepare the Crayfish, cut 2 lengths of wire about 12 inches. You may use a ruler to measure the 12 inches. Use a match to burn 2-3 mm of insulation off of one end of the wires. Burn 1 cm of insulation off the other end. Keep track of which end is which for both of the wires. Using the foam pad, secure a Crayfish. Using an 18 g needle, drill 2 small holes just through the carapace of the cephalothorax. When you drill the hole, a small drop of hemolymph will appear. Stop when you see the hemolymph.
Using the wires, put the 2-3 mm stripped end into one of the holes. The wire does not have to be deep into the hole. Hold the wire in place and add a small drop of cyanoacrylate adhesive over the hole. Apply a small amount of the Zip kicker. The wire should secure. Repeat this step with the second wire and hole.
Then, place the animal in a Tupperware container and add 100 ml of freshwater crustacean saline. Record the temperature of the saline. Next, hook the 1 cm stripped ends of the wires into the input posts on the back of the impedance converter. Once this is done, begin the LabTutor module called “Crayfish Heart Rate.”
Baseline Heart Rate
To get the baseline heart rate begin by covering the top of the container with foil. Allow the Crayfish to sit quietly for 10 minutes. After you wait 10 minutes, click Start and examine the trace. Adjust the electrode connection to the impedance converter if the recording is not fine. If the recording continues with bad recording, get a new Crayfish. Record the heart rate for 5 to 10 minutes. Make sure the heart rate is steady. After the 5 to 10 minutes, click Stop and annotate the start and end of the baseline heart rate trace.
Stressed Heart Rate
Uncover the container and Start the trace. Examine the trace and make sure the heart is being recorded. Once this is done, record the heart rate for 5 to 10 minutes. After the five to ten minutes, click Stop and annotate the start and end of the stressed heart rate trace.
Now, obtain the stock solution of 1 mM Serotonin from the refrigerator. Use the micropipettor with a blue tip and transfer 1 ml of solution to the container. Calculate the final concentration of the neurotransmitter in the holding water. Then, return the solution to the refrigerator. After that, allow the animal to sit with the foil cover for 5 minutes so that the neurotransmitter can equilibrate between the holding water and the animal’s hemolymph. After this wait, start the trace and record for 5 to 10 minutes. After the 5 to 10 minutes, click Stop and annotate the start and end of the serotonin heart rate trace. Replace the holding water with 100 ml of fresh Freshwater Crustacean Saline.
Obtain the stock solution of 1 mM Acetylcholine from the refrigerator. Using the micropipettor with a blue tip, transfer 1 ml of solution to the container. Calculate the final concentration of the neurotransmitter in the holding water. Then, return the solution to the refrigerator. Again, allow the animal to sit with the foil cover on for 5 minutes so that the neurotransmitter can equilibrate between the holding water and the animal’s hemolymph. Start the trace and record for 5 to 10 minutes. Click Stop and annotate the start and end of the acetylcholine heart rate trace. Replace the holding water with 100 ml of fresh Freshwater Crustacean Saline.
Now, obtain the stock solution of 1 mM Epinephrine from the refrigerator. Use the micropipettor with a blue tip and transfer 1 ml of solution to the container. Calculate the final concentration of the neurotransmitter in the holding water. Then, return the solution to the refrigerator. After that, allow the animal to sit with the foil cover for 5 minutes so that the neurotransmitter can equilibrate between the holding water and the animal’s hemolymph. After this wait, start the trace and record for 5 to 10 minutes. After the 5 to 10 minutes, click Stop and annotate the start and end of the epinephrine heart rate trace. Replace the holding water with 100 ml of fresh Freshwater Crustacean Saline.
Obtain a white bucket that is 2/3 full of ice. Rest the cover container on the surface of the ice. Allow the animal to equilibrate for 5 minutes. After the 5 minutes, measure the temperature of the holding water. While doing this, avoid disturbing the animal. Start the trace for 5 to 10 minutes. Click Stop and annotate the start and end of the temperature heart treat trace. Remove the container from the ice and set aside.
Find the heart rate for each of the annotate sections to collect your data and compare the baseline heart rate to the heart rate manipulated from each experimental condition.
There results we received were out of five different groups. We will keep the data of the five different results separated for clarity and then combine the results to see if our hypothesized trend was true.
Charts and Graphs
Table 1 are the results our group acquired from our neurotransmitter. Figure 1 shows a graph of the individual results
Table 1: Group results
Figure 1: Individual post results shown in a graph
Figure 2: Average of the results. There is averages for the pre and post. Control 1 is heart rate at rest and heart rate at stress. The following four go in this order: Sero, Ach, Epi, Temp
Figure 3: The average differences of Pre and Post. Control 1 is heart rate at rest and heart rate at stress. The following four go in this order: Sero, Ach, Epi, Temp
When comparing the individual groups, note that group five did not get full data for its heart rate experiment. Group five lacked pre and post results for epinephrine and temperature. Group five also lacked the results for a stressed heart rate. Group four had a huge skew to the right when collecting the pre temperature procedure. The collected data for the pre temperature and pre epinephrine for group four shows a heart rate of 104 and 102 while the other groups had 72, 72, and 64 for pre temp and 72, 72, and 78 for pre epinephrine. These results are in table four. Group three seemed to have a crayfish with a lower heart rate. The average base heart rate for groups 1, 2, 4, and 5 is 71.5 beats; however, group three had a heart rate of 50. The rest of the data follows this trend. This drop of heart rate might be because of the crayfish being a subordinate. In a study done by Heidi Schapker, a dominate crayfish and subordinate crayfish displayed different heart rates. The dominate crayfish had an average base heart rate around 70, while the subordinate crayfish had an average base heart rate at 65 (Schapker, 2002). The study agrees that group three might have had a subordinate crayfish.
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These different data collections could have skewed our total collected results; however, while focusing on the base and stress, the heart rate did not increase as much as expected. The average difference between the average base heart rate and stressed heart rate was 2.8 beats. In a study done by Laura Listerman, the crayfish used would have a heart rate increase between 100-110 heartbeats per minute when stimulated to defend itself (Listerman, 2000). The stressed crayfish in the study could have potentially had a higher heart rate because the cue of defense could require more blood pump than any other response; however, none of the stimulants used in our experiment rose the heart rate anywhere near 105 beats per minute.
In this experiment, the data shows that serotonin had the biggest difference between pre and post results. Serotonin also had the most error-free results. This is mostly due to serotonin being the first stimulant to manipulate the heart rate of the crayfish. Being the first means there would be no error in the water like the other stimulants could have had. Serotonin had an average pre serotonin heart rate of 65.6 beats and average post serotonin heart rate of 82.4 beats. The difference between the two is 16.8 beats. This is a huge jump compared to the other stimulants. This could be likely due to, as stated in the introduction, serotonin being endogenous to this organism when dealing with heart rate. As a natural stimulant to the neurogenic heart rate of the crayfish, serotonin could have the most affect to the heart rate. Our hypothesis of epinephrine being the neurotransmitter to cause the highest change is not supported by this data. The final concentration calculated for serotonin after it has been placed in the water with the crayfish was 9.9 x 10^-6 M.
Acetylcholine seemed to have the highest heart rate compared to the others; however, the difference between pre and post did not take first place. ACh had an average pre ACh of 74 beats and average post ACh of 85.2 beats. The average difference between the two was 11.2 beats. Note that group three did not have a consistent “pre” result compared to the rest of their “pre”. The pre ACh heart rate for group three was 72 beats while the others exhibited 58, 58, and 64 beats. There might have been an error while doing calculating the heart rate for pre ACh for group three; however, this pre ACh being 72 beats was more consistent with the other groups. This data supported our hypothesis of ACh increasing the heart rate. The final concentration calculated for ACh after it has been placed in the water with the crayfish was 9.9 x 10^-6 M.
It was hypothesized that epinephrine would cause the highest increase compared to the other neurotransmitters. Epinephrine did have the second highest difference. The data not does meet what actually happens because of the human error in the pre epinephrine; however, epinephrine still had a huge effect to the heart rate. This data supported our hypothesis of epinephrine increasing the heart rate. The final concentration calculated for epinephrine after it has been placed in the water with the crayfish was 9.9 x 10^-6 M.
The lower temperature affected the heart rate of the crayfish as expected. The heart rate decreased when put in colder water. This is because of the crayfish being a poikilotherm. As a poikilotherm, the body temperature of the crayfish is variable to its environment. As the environmental temperature decreases, it is expected that so will the body temperature. This could cause the heart rate to decrease since all movement is slower. This data supports our hypothesis of cold temperature dropping the heart rate.
In conclusion, our hypothesis was supported about which stimulants would increase or decrease the heart rate; however, our hypothesis about epinephrine being the neurotransmitter causing the biggest heart rate increase was not supported. As stated before, this could possibly due to human error. The human error could have been the result of an error in calculating the heart rate or not using the proper amount of solution. There was also a lack of data from one of the groups. The equipment used could have also given us inaccurate data. Because of this, a redo of the experiment is necessary to get better results.
It is important to notice that the neurotransmitters used to stimulate the neurogenic heart all increased the heart rate. This shows that even if there is a lack of one neurotransmitter that there can potentially be a backup neurotransmitter in its place. In addition, the fact that colder water decreases the heart rate is important because it must mean that an increase in temperature could cause the heart rate to increase. With climate having a rise in temperature, this could affect the crayfish and many other poikilotherm organisms. A constant increase in heart rate to acclimate towards temperature could be detrimental to these organisms.
For future experiments, I would gather crayfish with similar base heart rates and categorize them. Having a crayfish with a low heart rate be experimented with crayfish with relatively normal heart rates skews the data. However, if crayfish were categorized when being experimented, the data could be more organized in terms of their heart rates. The use of several buckets with Freshwater crustacean saline already prepared could help prevent any human errors when it came to reusing the same bucket and trying to avoid contamination. If several buckets were prepared ahead of time, the experiment would run more efficiently and would have less human error. This might change the results by having data that are more accurate.
In the end, it is important to study this to understand the neurogenic heart rate. As stated before, knowing which neurotransmitters work best for the heart could allow us to better understand the roles of neurotransmitters and how a neurogenic heart functions. With this knowledge, we could potentially use the information for medical uses and environmental uses for animals with similar functions.
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