Impedance Pneumography for Minimally Invasive Measurement of Heart Rate in Late Stage Invertebrates

Amalia M. Harrington; Holland Haverkamp; Heather  J. Hamlin

doi:10.3791/61096

JoVE Journal > Environment

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Impedance Pneumography for Minimally Invasive Measurement of Heart Rate in Late Stage Invertebrates

Published: April 04, 2020

doi:

10.3791/61096

Amalia M. Harrington^1,2, Holland Haverkamp^1,3, Heather J. Hamlin^1,2

¹School of Marine Sciences,University of Maine, ²Aquaculture Research Institute,University of Maine, ³Ecology and Environmental Sciences Program,University of Maine

Summary

Measuring heart rate during a thermal challenge provides insight into physiological responses of organisms as a consequence of acute environmental change. Using the American lobster (Homarus americanus) as a model organism, this protocol describes the use of impedance pneumography as a relatively noninvasive and nonlethal approach to measure heart rate in late stage invertebrates.

Abstract

Temperatures in oceans are increasing rapidly as a consequence of widespread changes in world climates. As organismal physiology is heavily influenced by environmental temperature, this has the potential to alter thermal physiological performance in a variety of marine organisms. Using the American lobster (Homarus americanus) as a model organism, this protocol describes the use of impedance pneumography to understand how cardiac performance in late stage invertebrates changes under acute thermal stress. The protocol presents a minimally invasive technique that allows for real-time collection of heart rate during a temperature ramping experiment. Data are easily manipulated to generate an Arrhenius plot that is used to calculate Arrhenius break temperature (ABT), the temperature at which heart rate begins to decline with increasing temperatures. This technique can be used in a variety of late stage invertebrates (i.e., crabs, mussels, or shrimps). Although the protocol focuses solely on the impact of temperature on cardiac performance, it can be modified to understand the potential for additional stressors (e.g., hypoxia or hypercapnia) to interact with temperature to influence physiological performance. Thus, the method has potential for wide-ranging applications to further understand how marine invertebrates respond to acute changes in the environment.

Introduction

In recent decades, increased input of greenhouse gases (i.e., carbon dioxide, methane, and nitrous oxide) into the atmosphere has resulted in widespread patterns of environmental change¹. The world’s oceans are rapidly warming²^,³, a trend that may have severe impacts on organismal physiology. Temperature heavily influences physiological rates, and organisms have an optimal temperature range for performance⁴^,⁵^,⁶. As such, individuals may encounter difficulties in maintaining proper oxygen delivery to tissues as temperatures stray outside of this range. This has the potential to lead to declines in aerobic performance in the face of warming ocean temperatures⁵^,⁷.

In a laboratory setting, a method to understand the physiological impacts of environmental change is to examine cardiac performance in the context of thermal stress. This provides insight into how exposure to predicted warming conditions may alter performance curves⁵^,⁶ as well as the potential for acclimation plasticity⁸. A variety of methods have been successfully implemented to previously measure heart rate in marine invertebrates. However, many of these techniques involve surgical removal or major manipulation of the exoskeleton and prolonged implantation of measurement devices⁹^,¹⁰^,¹¹, which introduces additional stress to the test subject and increases the time needed for a successful recovery prior to experimentation. Moreover, less invasive techniques (e.g., visual observation, videography) may be restricted to early life history stages when organisms may be fully or semi-transparent¹². Furthermore, additional challenges may be presented to researchers who are not well-versed in more technologically advanced methodologies (e.g., observations via infrared transducers or Doppler perfusion⁸^,¹¹).

This protocol uses the American lobster (Homarus americanus) as a late stage marine invertebrate model to demonstrate the use of impedance pneumography for assessing changes in heart rate during a temperature ramping experiment. Impedance pneumography involves passing of an oscillating electrical current (AC) across two electrodes positioned on either side of the pericardium to measure changes in voltage as the heart contracts and relaxes¹³^,¹⁴. This technique is minimally invasive, as it employs the use of small electrodes (i.e., 0.10–0.12 mm diameter) that are gently implanted just beneath the exoskeleton. Finally, it provides real-time assessments of both heart rate and water temperature during the ramp through the use of a data logger.

The protocol also provides instructions for calculating Arrhenius break temperature (ABT), the temperature at which heart rate begins to decrease with increasing temperatures¹³^,¹⁵. The ABT serves as a nonlethal indicator of the thermal limit of capacity in test subjects that may be favored over measuring the critical thermal maximum (CT_max, the upper limit of cardiac function⁵^,⁶), as lethal limits are often extreme and rarely encountered in the natural environment⁵.

Protocol

1. Equipment setup

Wrap clear, malleable tubing around itself to create a heat-exchanging coil that is approximately 8–10 cm in diameter and has extensions 40–70 cm long. Secure the coil using electrical tape.
Attach the heat-exchanging coil to the external supply and return fittings of a cooling/heating circulating water bath. Ensure the connection is secure using hose clamps.
Fill the well of the cooling/heating circulating water bath with reverse osmosis (RO) water and plug the power cord into an outlet. Turn the water bath on and make sure there are no leaks in its connection to the heat exchanging coil.
Set up the impedance convertor by plugging in the black BNC cable to the AC output on the unit and connecting it to the data logger (Table of Materials) using the Channel 1 port.
Plug the thermocouple probe (temperature recorder) into the T-type pod, then plug the T-type pod into the Channel 2 port of the data logger.
Plug the power cord of the data logger into a power supply and connect the data logger to a PC computer using the USB cable connector.
Fill the acclimation chamber and experimental arena with 7.5 L of artificial sea water (salinity = 35 ppt, pH = 8.1, temperature = ~12 °C).
NOTE: The volume, temperature, and chemistry of the water needed for the acclimation chamber and starting conditions in the experimental arena are dependent upon experimental design. Importantly, these containers must be large enough to comfortably submerge the test subject.

2. Implantation of electrodes

Place the lobster on a plastic grate that fits easily into the experimental arena such that the body comfortably makes a Y-shape at one end of the rectangle.
Carefully secure the lobster’s claws and abdomen to the plastic grate using small cable ties. The cable ties should be tight enough to prevent movement but allow room for surgical scissors to remove them upon completion of the experiment.
Dry off the carapace with a paper towel and clean it with a cotton ball soaked in 70% ethanol.
Create the holes for the electrodes.
1. Using a small drill bit (e.g., 1.6 mm), slowly and carefully hand-drill two small holes (nearly) through the carapace on either side of the pericardium.
2. Finish each hole by gently inserting a sterile dissecting needle.
3. If the needle does not easily go through the carapace, continue to slowly hand-drill before trying the needle again.
  NOTE: To minimize stress in experimental animals, practicing this technique prior to experimentation is highly recommended. Over time, users can easily determine by feeling when the drill bit is nearly though the carapace and switch to the needle. Hand-drilling is appropriate for lobsters and crabs, especially if the exoskeleton is soft (i.e., animal has recently molted). However, if the test subject has a thicker exoskeleton or shell (i.e., a bivalve), a Dremel tool is more appropriate.
Obtain the electrodes (36–38 G magnetic wire, 0.10–0.12 mm diameter) and scrape off a small bit of insulation at the wire’s tip using a dissecting knife blade. Carefully bend the tip of each wire into a small hook using forceps and insert one into each of the newly drilled holes.
Secure each wire lead using a small drop of cyanoacrylate glue and allow it to dry for 5–10 min.
NOTE: It is crucial to use the glue sparingly, as adding too much will reinsulate the wire and prevent the signal from being recorded.
Once the glue is dry, attach the wire leads to the impedance convertor and turn it on. Place the lobster into the acclimation chamber and allow it to acclimate to the implanted electrodes for 15–20 min.
NOTE: Quick or jarring movements, as well as incompletely dried glue, may cause the electrodes to become detached from the carapace. If this happens, return to step 2.6.
Turn the data logger on and open the LabChart software on the computer. Click New Experiment and leave the Chart View screen open.
In Chart View, locate the Channel Function menu for Channel 1 from the right-hand section of the screen. Choose Input Amplifier from the menu and select AC Coupling. The incoming signal from the test subject will now appear on the screen in real-time.
NOTE: The sensitivity of the channel can be adjusted by selecting the Range pop-up menu. Adjust the range until the signal peaks are 25%–75% of the full scale. Close the Input Amplifier by clicking OK.
On the impedance convertor, adjust the Gain (size) and Balance until a strong signal is observed on the data logger output, aiming to keep the Balance near zero.
On Channel 2, select T-Type pod to record real-time temperature data.
When both channels are set up properly, click the Start button, and the data logger will begin logging data.

3. Temperature ramping

After the acclimation period, place the plastic grate with the attached lobster carefully into the experimental arena and set the heat-exchanging coil on top of the grate.
Place the thermocouple probe near the lobster, ensuring it is fully submerged before placing the lid on the experimental arena to reduce visual stress to the test subject.
Adjust the balance as needed and place a comment on the output stating that the trial has begun.
The output can and should be saved periodically throughout the experiment.
1. Click File and select Save As to initially save the output to the computer.
2. When saving during the experiment, click File and select Save.
  NOTE: Although the LabChart software can recover files in the event of an accidental program shutdown (e.g., a power outage), it is recommended to save active files every 15–20 min during the experiment to prevent data loss.
Increase the water temperature of the experimental arena at a rate of ~1.5 °C every 15 min to achieve a ramp from 12 °C to 30 °C over a 2.5 h period by adjusting the temperature of the recirculating water bath.
NOTE: The geographic distribution of the American lobster spans a 25 °C thermal gradient, and individuals can acclimate to and survive at temperatures of up to 30 °C¹⁶. As such, 30 °C was chosen as the upper limit for this temperature ramp, as it ensures that lobsters experience a stressful scenario that does not reach the critical thermal maximum¹³, which could lead to mortality. The specific rate of warming was selected because it falls within a range of warming rates implemented in studies using other species⁸^,¹⁴ as well as previous research on the American lobster¹³^,²⁷. Prior to implementing this protocol, it is important to 1) determine the appropriate range of temperatures for a given experiment and 2) conduct a pretrial temperature ramp with an empty experimental arena, as this will help to determine the necessary temperature adjustment of the water bath to achieve the desired ramp. This may also differ depending on the volume of water in the arena.
Throughout the temperature ramp, record whenever an adjustment that may impact the output occurs.
1. Note that the balance on the impedance convertor will likely need to be adjusted throughout the experiment, and doing so may cause an unintentional spike in the output.
2. As the temperature in the experimental arena begins to reach levels outside of the preferred thermal range of the test subject, involuntary muscle contractions may result in an erroneous “spike” in the output. If this occurs, make a comment to identify areas of the output that should be removed during the data conversion process.
When the ramp is completed, remove the lobster from the experimental arena and place it into a recovery bath (12 °C) for ~20 min. If desired, continue to monitor the lobster’s heart rate until it returns to basal levels.
After 20 min, hit the Stop button on the PowerLab output and save the file. Carefully remove the electrodes and cut the cable ties with surgical scissors before returning the test subject to its holding tank.
NOTE: Rather than placing a lobster directly into the recovery bath, another option is to slowly return the experimental arena to its starting temperature. This is accomplished by cooling the experimental arena by ~1.5 °C every 15 min over the course of an additional 2.5 h.

4. Data conversion

Open Data Pad. Set Column A to time by double-clicking on Column A and clicking on Selection & Active Point on the left-hand side of the Data Pad Column A Setup menu. Select Time from the right-hand side of the menu and close the window by clicking OK.
Set Column B to the average temperature by double-clicking on Column B and selecting the Estatística option from the left-hand side of the Data Pad Column B Setup menu. Select Mean from the right-hand side of the menu and Channel 2 as the Calculation source at the bottom of the menu’s window. Click OK to close the window.
Converting the voltage recorded to beats per minute
1. Double-click on Column C and select Selection & Active Point on the left-hand side of the menu. Select Selection Duration from the right-hand side of the menu and click OK to close the window.
2. Double-click on Column D and select Cyclic Measurements on the left-hand side of the menu. Select Event Count from the right-hand side of the menu, and Channel 1 as the Calculation source. Click OK to close the window. This will count the peaks of the data to determine heart rate across a selected portion of data.
  NOTE: If needed, select the Options button from the bottom of the menu and adjust the Detection Settings to more accurately read the data. Scan through the data file and determine if the “Sine” or “Spikey” shape options result in counts of only the major peaks of the heartbeat output. Additionally, adjust the Detection Adjustment threshold on the right-hand side of the menu to ignore noise in the output file.
3. Double-click on Column E and select Cyclic Measurements on the left-hand side of the menu. Select Average Cyclic Rate, and Channel 1 as the Calculation source. Adjust the Detection Settings and Detection Adjustment to match the settings for Column D (if manipulated in step 4.4.2). Click OK to close the window. This provides the final estimation of heart rate (as beats per minute) over a selected portion of data.
When the columns are set up, return to the data file and highlight the desired sections of the output, omitting areas of erroneous data as identified by comments in section 3.6.
1. Select Commands and Multiple Add to Data Pad.
2. Select Time from the Find using drop-down menu and pull data every 30 s by checking the Every box and entering “30” under the Select menu.
3. Click the Current selection option from the Step through menu and click Add.
Return to the Data Pad screen and select File and Save As to save the output as an Excel file.
NOTE: Here, heart rate is reported (in beats per minute) every 30 s as opposed to every minute based on previous research⁸^,²⁷. This also helps to more accurately capture changes in the real-time collected voltage data. It is possible to select data at shorter or longer time intervals based on individual preference.

5. Calculation of Arrhenius break temperature

Open the data file in Excel and manipulate the output from the LabChart software.
1. Convert the temperature from Celsius to the reciprocal of Kelvin using the following equation: [1000/(temperature °C + 273.15 K)].
2. Obtain the natural log of heart rate: ln(BPM).
Generate an Arrhenius plot by plotting heart rate as a function of temperature, expressed as ln(BPM) vs. reciprocal (K)¹³^,¹⁵.
In SigmaPlot, fit the data with a piecewise regression and determine the intersection point, which is the ABT.
1. Copy and paste the transformed data into a new workbook. Select the Estatística option from the main menu and Regression Wizard from the drop-down list.
2. In the Equation window, select piecewise from the Equation Category menu and 2 segment linear under the Equation Name box. Click Next.
3. In the Variables window, select the transformed temperature data to be the t variable and the transformed heart rate data to be the y variable, using the drop-down options in the Variable Columns menu. Make sure that XY Pair is selected in the Data From menu before clicking Next.
4. After reviewing the Fit Results window, click Next and check the box for Create Report in the Numeric Output Options window. Click Next.
5. In the Graph Options window, check the Create new graph option under the Fit Results Graph section, and Add equation to graph title under the Graph Features Section. Click Finish.
6. On the Resultados output page, retrieve the equations and parameter values for the two regions of the piecewise regression, as well as the statistical output for the regression (e.g., R², F-statistic, and p-value).
7. Using the parameter values and equations generated, set the two segments equal to each other and solve for the variable “t” to determine the ABT. Convert this value back to Celsius using the following equation: °C = (1000/t) – 273.15.
  NOTE: The ABT can also be calculated in the R statistical computing environment using the package “segmented”¹⁷ in the program SAS¹⁸, or using the “Segmental linear regression” routine in Prism8¹⁹.

Representative Results

This protocol describes the use of impedance pneumography to obtain real-time data for heart rate (in voltage) and temperature during a temperature-ramping experiment. When perforing this technique, the amplitude of the voltages and temperatures recorded will vary based on experimental design and focal species. However, the voltage output displayed in real-time follows a generic sine distribution when the protocol is implemented correctly (Figure 1A). As the temperature in the arena is increased, the real-time distribution of voltage changes to reflect an increased frequency of voltage peaks (i.e., heart beats; Figure 1B). As the arena temperature continues to increase to levels outside of the test subject’s optimal performance window, the distribution changes to depict a reduced frequency of voltage peaks with a sine-like shape interrupted by sporadic peaks and/or moments of “flat-lining” (Figure 1C).

Once raw data are converted using the Data Pad component of the LabChart software, the resulting distribution of heart rate (in beats per minute) over the course of the temperature ramp follows a parabolic distribution if the experiment is successful (Figure 2). As the temperature in the arena is increased, the heart rate of the test subject also increases to meet elevated energetic demands associated with warmer temperatures. However, as temperature continues to increase and the test subject begins to experience moderate to extreme thermal stress, heart rate begins to decline or becomes erratic as the subject begins to exhibit passive thermal tolerance (e.g., onset of anaerobic respiration, metabolic rate suppression, and reduced activity⁵^,⁷). When heart rate and temperature data are transformed and an Arrhenius plot is generated, the point at which the heart rate begins to decline (ABT) can be calculated (Figure 3). The Arrhenius plot is then fit with a piecewise regression using statistical software in which the intersection of the two lines represents the ABT.

Figure 1: Representative output from LabChart data logger. Real-time change in voltage across electrodes of the test subject is displayed in red, and the concomitant real-time output of the arena temperature (°C) is displayed in blue. In the beginning of the experiment under cooler temperatures (e.g., 13.1 °C), voltage should follow a generic sine-like distribution (A). As temperature is increased (e.g., 23 °C), the frequency of voltage peaks should increase, but the distribution should remain sine-like (B). Finally, as the test subject is pushed outside of its optimal thermal performance window (e.g., 28.5 °C), the voltage peaks should become erratic as the frequency decreases (C). Please click here to view a larger version of this figure.

Figure 2: Expected distribution of heart rate over the temperature ramp course. Voltage data collected by the data logger are converted to heart rate in beats per minute (BPM) using the Data Pad component of the software. When the ramp is conducted correctly, a parabolic distribution of heart rate over the temperature range tested is displayed. Please click here to view a larger version of this figure.

Figure 3: Example of an Arrhenius plot. Once data have been converted in the Data Pad and exported, they are transformed to generate an Arrhenius plot. In this example, data are fit with a piecewise nonlinear regression in SigmaPlot, generating equations for the left- and right-hand segments (region 1 and region 2, respectively) of the regression line, as well as goodness-of-fit metrics. The intersection of the two regression lines is solved as the ABT (red star). Please click here to view a larger version of this figure.

Discussion

This protocol describes the use of impedance pneumography to measure changes in heart rate of late stage invertebrates during a temperature ramping experiment. The primary benefit of this technique compared to other laboratory-based approaches⁹^,¹⁰^,¹¹ is that it is minimally invasive and does not involve major surgical manipulation of the exoskeleton, thus reducing the amount of recovery time needed prior to experimentation. Moreover, the equipment is easy to use, and resulting data can be simply manipulated and interpreted in the suggested software program. While the American lobster is used here as a model subject, this technique has been successfully implemented in blue mussels (Mytilus spp.¹⁴) and can be easily modified for use in other late stage invertebrates (i.e., crabs, shrimps, and other bivalves).

An additional benefit of the protocol is that it focuses on calculating the ABT as a nonlethal indicator of thermal limits. Although numerous studies present the CT_max as the significant endpoint when determining thermal physiological performance⁵^,⁸^,²⁰^,²¹^,²²^,²³, organisms rarely encounter temperatures in this range in the natural environment⁵. Moreover, as the CT_max is often a lethal temperature, using this metric as the preferred endpoint precludes the use of test subjects in additional or follow-on experimentation post-thermal stress²³. When aiming to calculate the ABT using this protocol, it is crucial to increase the temperature in the experimental arena to the point of pushing the test subject to its physiological limit without inducing death. Therefore, it is recommended to determine the potential thermal limits of the focal species via a pilot study (when possible) prior to determining the full range of the experimental temperature ramp.

It is also recommended that researchers determine and observe natural variations in basal heart rate of a focal species when temperature in the experimental arena is maintained at a constant and non-stressful level prior to the ramping experiment. This is particularly helpful for focal species in which resting heart rate information is not available in the published literature. It also serves as ample practice of electrode implantation techniques. This may also help researchers determine the appropriate acclimation time required to ensure that no false spikes in heart rate are due to handling stress at the beginning of an experiment.

Although the protocol discusses the use of impedance pneumography in the context of thermal stress alone, it can also be utilized to explore the potential interactive effects of other stressors on thermal physiology. Organismal performance may be reduced in the presence of environmental stressors (i.e., hypoxia, hypercapnia, pollutants, and/or changes in salinity), which may also compress optimal temperature ranges for performance⁷^,²⁴^,²⁵^,²⁶. As such, this protocol can be modified to explore how exposure to various stressors prior to temperature ramping may impact performance.

For example, Harrington and Hamlin²⁷ exposed juvenile H. americanus to current or predicted end-century pH conditions (8.0 and 7.6, respectively) for 2 months prior to assessing cardiac performance during a temperature ramp. Lobsters pre-exposed to more acidic environments exhibited a significant reduction in mean ABT compared to those held under current pH conditions. This suggests that a low pH environment reduces thermal performance and may increase the risk of cellular damage due to heat stress at lower temperatures²⁷. Future efforts could expand on the method presented here to include pre-exposure to any combination of environmental stressors prior to following this protocol. Moreover, this protocol can be modified to measure changes in cardiac performance during exposure to biotic stressors as well as how thermal limits can change according to ontogeny⁴^,⁵.

A major limitation of this protocol is that the equipment as described is restricted for use in a laboratory setting, potentially limiting its applicability for field-based experiments that require more specialized equipment⁸. This technique also requires the restraint of highly motile test subjects (e.g., lobsters and crabs) to reduce the production of false data points resulting from non-cardiac muscle movements. Although this may restrict natural behaviors during a temperature ramp, the impact of restraints are consistent across all test subjects. Most importantly, there is the potential for tissue damage or death in test subjects if aggressive or careless drilling during electrode implantation is implemented. This contrasts sharply with infrared photoplethysmography, a truly noninvasive technique that utilizes an external infrared transducer to pass light through the pericardium and record heart function by converting reflected light energy to voltage⁸^,²⁸.

Although infrared photoplethysmography reduces the risk of handling stress compared to impedance pneumography, correctly implanting electrodes using the described method results in minimal trauma, allows for a quick acclimation time, and leads to rapid recovery without inducing mortality in test subjects following the ramping experiment²⁷. As there is no significant difference in the cardiac output recorded by both methods²⁸, it is concluded that impedance pneumography is a reliable and minimally invasive technique for assessing cardiac performance. Finally, the numerous benefits and flexibility of the protocol have the potential to elucidate how various environmental factors interact with temperature to impact physiological performance in late stage crustaceans.

Declarações

The authors have nothing to disclose.

Acknowledgements

The authors thank Paul Rawson for laboratory assistance and the National Science Foundation award IIA-1355457 to Maine EPSCoR at the University of Maine for funds to purchase equipment. This project was supported by the USDA National Institute of Food and Agriculture, Hatch project number MEO-21811 through the Maine Agricultural and Forest Experiment Station, as well as NOAA National Marine Fisheries Service Saltonstall Kennedy Grant #18GAR039-136. The authors also thank three anonymous reviewers for their comments on a previous version of this manuscript. Maine Agricultural and Forest Experiment Station Publication Number 3733.

Materials

1.6 mm (1/16 in) drill bit	Milwaukee Tool at Home Depot	1001294900	This is for a 1.6 mm (1/16 in) diameter drill bit. This item can be found at most home-improvement stores.
38 AWG Copper Magnet Wire	TEMCo	MW0093	This wire is used to make the wire electrode leads that are implanted into the test subjects. This listing is for a 4 oz coil of 38-gauge magnetic wire. TemCo also has 36-gauge magnetic wire that is also suitable for use in constructing wire electrodes.
Cyanoacrylate glue	Loctite	852882	This item includes a brush tip, which makes it easier to control the amount of glue used to secure electrodes to the carapace.
Ethanol, 70% Solution, Molecular Biology Grade	Fisher BioReagents	BP82931GAL	This reagent is used in combination with the sterile cotton balls to disinfect the carapace prior to electrode implantation.
Excel	Microsoft	N/A	This program is used in the protocol for organizing, manipulating, and analyzing data. It is compatible with both PC and Mac operating systems.
Fisherbrand 8-Piece Dissection Kit	Fisher Scientific	08-855	This kit includes the forceps, scissors, dissecting knife (and blades), and dissecting needle needed to accomplish the electrode implantation steps in the protocol.
Fisherbrand Isotemp Refrigerated/Heated Bath Circulators: 5.4-6.5L, 115V/60Hz	Fisher Scientific	13-874-180	This is a complete system that consists of an immersion circulator and a bath. It can be used as a temperature controlled bath or to circulate fluid externally to an application. Temperature range of this water bath is -20 to +100 °C, and the unit heats/cools rapidly and is easy to drain upon conclusion of use.
Fisherbrand Sterile Cotton Balls	Fisher Scientific	22-456-885	These swabs should be soaked in 70% ethanol before being used to disinfect the carapace prior to electrode implantation.
Fork Terminal, Red Vinyl, Butted Seam, 22 to 16 AWG, 100 PK	Grainger	5WHE6	Terminals are soldered to the magnetic wire to construct the wire electrodes. These can be purchased from a variety of home-improvement vendors.
Impedance converter	UFI	Model 2991	Measures impedance changes correlated with very small voltage changes, ranging from 0.2 ohm to over 5 ohms. This model can convert impedance changes that stem from resistance, capacitance, or inductance variations, as well as a combination of all three.
LabChart software	ADInstruments	N/A	Purchase of the PowerLab datalogger includes the LabChart software, but a license for the software can also be directly downloaded online. LabChart allows the user to record data, open and read LabChart files, analyze data, as well as save and export files. There is a free version of the software, LabChart Reader, but users can only open and read LabChart files and analyze them (i.e., it cannot be used to record, save, or export data files). One also has the option of selecting LabChart Pro, which includes LabChart teaching modules that can be used for educational purposes.
LED Soldering Iron	Grainger	28EA35	This is a generic soldering iron that can be used to solder the magnetic wire to the fork terminals to create the wire electrodes.
PowerLab datalogger	ADInstruments	ML826	There are a variety of models of the PowerLab. This catalog number is for the 2/26 model that is a 2 channel, 16 bit resolution recorder with two analog input channels, independently selectable input sensitivities, two independent analog outputs for stimulation or pulse generation and a trigger input. The PowerLab features a wide range of low-pass filters, AC or DC coupling and adaptive mains filter. This unit has a USB interface for connection to Windows or Mac OS computers and a sampling rate of 100,000 samples/s per channel.
Prism8	GraphPad	N/A	This program provides an additional option for calculating the Arrhenius Break Temperature through its “Segmental linear regression” data analysis option. This program does not require any programming and is compatible with both Mac and Windows operating systems.
R	R Project	N/A	This is free software for statistical computing that is compatible with UNIX platforms, as well as Windows and Mac operating systems. This program can also be used to calculate the Arrhenius Break Temperature using the “segmented” package. There are a number of tutorials and user guides available online through the r-project.org website.
Rosin Core Solder	Grainger	331856	This product has a diameter of 0.031 in (0.76 mm) and is ideal for use in soldering speaker wire (similar gauge as magnetic wire used for electrodes).
SAS	SAS Institute	N/A	This program provides an additional option for calculating the Arrhenius Break Temperature. However, it does require programming and is not compatible with Mac operating systems.
SigmaPlot	Systat Software, Inc.	N/A	This is the authors’ preferred program for statistical determination of the Arrhenius Break Temperature. The “Regression Wizard” is easy to use and does not require any programming. One can obtain a free 30-day trial license before purchase. However, it is compatible only with PC computers.
T-type Pod	ADInstruments	ML312	Suitable for measurement of temperatures from 0-50 °C using T-type thermocouples.
T-type Thermocouple Probe	ADInstruments	MLT1401	Compatible with the T-type Pod for connection. Measures temperature up to 150 °C, and is suitable for immersion in various solutions, semi-solids, and tissue (includes a needle for implantation). This product is a 0.6 mm diameter isolated probe that is sheathed in chemical-resistant Teflon and a lead length of 1.0 m.
UV Cable Tie, Black	Home Depot	295813	This is for a 100-pack of 8-inch (20.32 cm), black cable ties. However, based on the size of test subjects, smaller or larger cable ties may be needed. This item, and others like it, can be purchased at any home-improvement store.

Referências

Stocker, T. F., et al. . Climate Change 2013: The Physical Basis. Contribution of Working Group I to the Fifth Assessment Report of the Intergovernmental Panel on Climate Change. , (2013).
Pershing, A. J., et al. Slow adaptation in the face of rapid warming leads to collapse of the Gulf of Maine cod fishery. Science. 350 (6262), 809-812 (2015).
Smale, D. A., et al. Marine heat waves threaten global biodiversity and the provision of ecosystem services. Nature Climate Change. 9 (4), 306-316 (2019).
Pörtner, H. O., Farrell, A. P. Physiology and climate change. Science. 322 (5902), 690-692 (2008).
Pörtner, H. O., Bock, C., Mark, F. C. Oxygen- and capacity-limited thermal tolerance: bridging ecology and physiology. Journal of Experimental Biology. 220 (15), 2685-2696 (2017).
Somero, G. N., Lockwood, B. L., Tomanek, L. . Biochemical adaptation: response to environmental challenges, from life’s origins to the Anthropocene. , (2017).
Sokolova, I. M., Frederich, M., Bagwe, R., Lanning, G., Sukhotin, A. A. Energy homeostasis as an integrative tool for assessing limits of environmental stress tolerance in aquatic invertebrates. Marine Environmental Research. 79, 1-15 (2012).
Tepolt, C. K., Somero, G. N. Master of all trades: thermal acclimation and adaptation of cardiac function in a broadly distributed marine invasive species, the European green crab, Carcinus maenas. Journal of Experimental Biology. 217 (7), 1129-1138 (2014).
Frederich, M., Pörtner, H. O. Oxygen limitation of thermal tolerance defined by cardiac and ventilatory performance in spider crab, Maja squinado. American Journal of Physiology-Regulatory, Integrative and Comparative Physiology. 279 (5), 1531-1538 (2000).
Metzger, R., Sartoris, F. J., Langenbuch, M., Pörtner, H. O. Influence of elevated CO2 concentrations on thermal tolerance of the edible crab Cancer pagurus. Journal of Thermal Biology. 32, 144-151 (2007).
Walther, K., Sartoris, F. J., Bock, C., Pörtner, H. O. Impact of anthropogenic ocean acidification on thermal tolerance of the spider crab Hyas araneus. Biogeosciences. 6 (10), 2207-2215 (2009).
Styf, H. K., Sköld, H. N., Eriksson, S. P. Embryonic response to long-term exposure of the marine crustacean Nephrops norvegicus to ocean acidification. Ecology and Evolution. 3 (15), 5055-5065 (2013).
Camacho, J., Qadri, S. A., Wang, H., Worden, M. K. Temperature acclimation alters cardiac performance in the lobster Homarus americanus. Journal of Comparative Physiology A. 192 (12), 1327-1334 (2006).
Braby, C., Somero, G. N. Ecological gradients and relative abundance of native (Mytilus trossulus) and invasive (Mytilus galloprovincialis) blue mussels in the California hybrid zone. Marine Biology. 148 (6), 1249-1262 (2006).
Stenseng, E., Braby, C. E., Somero, G. N. Evolutionary and acclimation-induced variation in the thermal limits of heart function in congeneric marine snails (Genus Tegula): implications for vertical zonation. Biological Bulletin. 208 (2), 138-144 (2005).
Factor, J. . Biology of the Lobster: Homarus americanus. , (1995).
Muggeo, V. M. Segmented: an R package to fit regression models with broken-lin relationships. R News. 8 (1), 20-25 (2008).
Ryan, S. E., Porth, L. S. A tutorial on the piecewise regression approach applied to bedload transport data. General Technical Report RMS-GTR-189. , (2007).
. . Prism8 Statistics Guide. , (2020).
Cuculescu, M., Hyde, D., Bowler, K. Thermal tolerance of two species of marine crab, Cancer pagurus and Carcinus maenas. Journal of Thermal Biology. 23 (2), 107-110 (1998).
Stillman, J. H. A comparative analysis of plasticity of thermal limits in porcelain crabs across latitudinal and intertidal zone clines. International Congress Series. 1275, 267-274 (2004).
Maderia, D., et al. cellular and biochemical thermal stress response of intertidal shrimps with different vertical distributions: Palaemon elegans and Palaemon serratus. Comparative Biochemistry and Physiology, Part A. 183, 107-115 (2015).
Padilla-Ramirez, S., et al. The effects of thermal acclimation on the behavior, thermal tolerance, and respiratory metabolism in a crab inhabiting a wide range of thermal habitats (Cancer antennarius Stimpson, 1856, the red shore crab). Marine and Freshwater Behaviour and Physiology. 48 (2), 89-101 (2017).
Pörtner, H. O. Ecosystem effects of ocean acidification in times of ocean warming: a physiologist’s view. Marine Ecology Progress Series. 373, 203-217 (2008).
Pörtner, H. O. Oxygen- and capacity-limitation of thermal tolerance: a matrix for integrating climate-related stressor effects in marine ecosystems. Journal of Experimental Biology. 213 (6), 881-893 (2010).
Zittier, Z. M. C., Hirse, T., Pörtner, H. O. The synergistic effects of increasing temperature and CO2 levels on activity capacity and acid-base balance in the spider crab, Hyas araneus. Marine Biology. 160 (8), 2049-2062 (2013).
Harrington, A. M., Hamlin, H. J. Ocean acidification alters thermal cardiac performance, hemocyte abundance, and hemolymph chemistry in subadult American lobsters Homarus americanus H. Milne Edwards, 1837 (Decapoda: Malcostraca: Nephropidae). Journal of Crustacean Biology. 39 (4), 468-476 (2019).
Depledge, M. H. Photoplethysmography – a non-invasive technique for monitoring heart beat and ventilation rate in decapod crustaceans. Comparative Biochemistry and Physiology Part A: Physiology. 77 (2), 369-371 (1984).

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Harrington, A. M., Haverkamp, H., Hamlin, H. J. Impedance Pneumography for Minimally Invasive Measurement of Heart Rate in Late Stage Invertebrates. J. Vis. Exp. (158), e61096, doi:10.3791/61096 (2020).