Name: impedance pneumography for minimally invasive measurement of heart rate in late stage invertebrates
Uploaded: 2020-04-04T13:00:00.000+00:00
Duration: 8 min 25 s
Description: Watch this Scientific Journal Video about Impedance Pneumography for Minimally Invasive Measurement of Heart Rate in Late Stage Invertebrates at JoVE.com

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.

1. Equipment setup
<ol>
	<li>Wrap clear, malleable tubing around itself to create a heat-exchanging coil that is approximately 8&#8211;10 cm in diameter and has extensions 40&#8211;70 cm long. Secure the coil using electrical tape.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Fill the acclimation chamber and experimental arena with 7.5 L of artificial sea water (salinity = 35 ppt, pH = 8.1, temperature = ~12 &#176;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.</li>
</ol>
2. Implantation of electrodes
<ol>
	<li>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.</li>
	<li>Carefully secure the lobster&#8217;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.</li>
	<li>Dry off the carapace with a paper towel and clean it with a cotton ball soaked in 70% ethanol.</li>
	<li>Create the holes for the electrodes.
	<ol>
		<li>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.</li>
		<li>Finish each hole by gently inserting a sterile dissecting needle.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Obtain the electrodes (36&#8211;38 G magnetic wire, 0.10&#8211;0.12 mm diameter) and scrape off a small bit of insulation at the wire&#8217;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.</li>
	<li>Secure each wire lead using a small drop of cyanoacrylate glue and allow it to dry for 5&#8211;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.</li>
	<li>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&#8211;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.</li>
	<li>Turn the data logger on and open the LabChart software on the computer. Click New Experiment and leave the Chart View screen open.</li>
	<li>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%&#8211;75% of the full scale. Close the Input Amplifier by clicking OK.</li>
	<li>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.</li>
	<li>On Channel 2, select T-Type pod to record real-time temperature data.</li>
	<li>When both channels are set up properly, click the Start button, and the data logger will begin logging data.</li>
</ol>
3. Temperature ramping
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Adjust the balance as needed and place a comment on the output stating that the trial has begun.</li>
	<li>The output can and should be saved periodically throughout the experiment.
	<ol>
		<li>Click File and select Save As to initially save the output to the computer.</li>
		<li>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&#8211;20 min during the experiment to prevent data loss.</li>
	</ol>
	</li>
	<li>Increase the water temperature of the experimental arena at a rate of ~1.5 &#176;C every 15 min to achieve a ramp from 12 &#176;C to 30 &#176;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 &#176;C thermal gradient, and individuals can acclimate to and survive at temperatures of up to 30 &#176;C16. As such, 30 &#176;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.</li>
	<li>Throughout the temperature ramp, record whenever an adjustment that may impact the output occurs.
	<ol>
		<li>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.</li>
		<li>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 &#8220;spike&#8221; in the output. If this occurs, make a comment to identify areas of the output that should be removed during the data conversion process.</li>
	</ol>
	</li>
	<li>When the ramp is completed, remove the lobster from the experimental arena and place it into a recovery bath (12 &#176;C) for ~20 min. If desired, continue to monitor the lobster&#8217;s heart rate until it returns to basal levels.</li>
	<li>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 &#176;C every 15 min over the course of an additional 2.5 h.</li>
</ol>
4. Data conversion
<ol>
	<li>Open Data Pad. Set Column A to time by double-clicking on Column A and clicking on Selection &#38; 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.</li>
	<li>Set Column B to the average temperature by double-clicking on Column B and selecting the Statistics 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&#8217;s window. Click OK to close the window.</li>
	<li>Converting the voltage recorded to beats per minute
	<ol>
		<li>Double-click on Column C and select Selection &#38; 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.</li>
		<li>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 &#8220;Sine&#8221; or &#8220;Spikey&#8221; 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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>Select Commands and Multiple Add to Data Pad.</li>
		<li>Select Time from the Find using drop-down menu and pull data every 30 s by checking the Every box and entering &#8220;30&#8221; under the Select menu.</li>
		<li>Click the Current selection option from the Step through menu and click Add.</li>
	</ol>
	</li>
	<li>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.</li>
</ol>
5. Calculation of Arrhenius break temperature
<ol>
	<li>Open the data file in Excel and manipulate the output from the LabChart software.
	<ol>
		<li>Convert the temperature from Celsius to the reciprocal of Kelvin using the following equation: [1000/(temperature &#176;C + 273.15 K)].</li>
		<li>Obtain the natural log of heart rate: ln(BPM).</li>
	</ol>
	</li>
	<li>Generate an Arrhenius plot by plotting heart rate as a function of temperature, expressed as ln(BPM) vs. reciprocal (K)13,15.</li>
	<li>In SigmaPlot, fit the data with a piecewise regression and determine the intersection point, which is the ABT.
	<ol>
		<li>Copy and paste the transformed data into a new workbook. Select the Statistics option from the main menu and Regression Wizard from the drop-down list.</li>
		<li>In the Equation window, select piecewise from the Equation Category menu and 2 segment linear under the Equation Name box. Click Next.</li>
		<li>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.</li>
		<li>After reviewing the Fit Results window, click Next and check the box for Create Report in the Numeric Output Options window. Click Next.</li>
		<li>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.</li>
		<li>On the Results 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).</li>
		<li>Using the parameter values and equations generated, set the two segments equal to each other and solve for the variable &#8220;t&#8221; to determine the ABT. Convert this value back to Celsius using the following equation: &#176;C = (1000/t) - 273.15. 
		NOTE: The ABT can also be calculated in the R statistical computing environment using the package &#8220;segmented&#8221;17 in the program SAS18, or using the &#8220;Segmental linear regression&#8221; routine in Prism819.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

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. More...

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&#8217;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&#8211;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 (CTmax, the upper limit of cardiac function5,6), as lethal limits are often extreme and rarely encountered in the natural environment5.

<table><tbody><tr><td>1.6 mm (1/16 in) drill bit</td><td>Milwaukee Tool at Home Depot</td><td>1001294900</td><td>This is for a 1.6 mm (1/16 in) diameter drill bit. This item can be found at most home-improvement stores.</td></tr><tr><td>38 AWG Copper Magnet Wire</td><td>TEMCo</td><td>MW0093</td><td>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.</td></tr><tr><td>Cyanoacrylate glue</td><td>Loctite</td><td>852882</td><td>This item includes a brush tip, which makes it easier to control the amount of glue used to secure electrodes to the carapace.</td></tr><tr><td>Ethanol, 70% Solution, Molecular Biology Grade</td><td>Fisher BioReagents</td><td>BP82931GAL</td><td>This reagent is used in combination with the sterile cotton balls to disinfect the carapace prior to electrode implantation.</td></tr><tr><td>Excel</td><td>Microsoft</td><td>N/A</td><td>This program is used in the protocol for organizing, manipulating, and analyzing data. It is compatible with both PC and Mac operating systems.</td></tr><tr><td>Fisherbrand 8-Piece Dissection Kit</td><td>Fisher Scientific</td><td>08-855</td><td>This kit includes the forceps, scissors, dissecting knife (and blades), and dissecting needle needed to accomplish the electrode implantation steps in the protocol.</td></tr><tr><td>Fisherbrand Isotemp Refrigerated/Heated Bath Circulators: 5.4-6.5L, 115V/60Hz</td><td>Fisher Scientific</td><td>13-874-180</td><td>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 &amp;deg;C, and the unit heats/cools rapidly and is easy to drain upon conclusion of use.</td></tr><tr><td>Fisherbrand Sterile Cotton Balls</td><td>Fisher Scientific</td><td>22-456-885</td><td>These swabs should be soaked in 70% ethanol before being used to disinfect the carapace prior to electrode implantation.</td></tr><tr><td>Fork Terminal, Red Vinyl, Butted Seam, 22 to 16 AWG, 100 PK</td><td>Grainger</td><td>5WHE6</td><td>Terminals are soldered to the magnetic wire to construct the wire electrodes. These can be purchased from a variety of home-improvement vendors.</td></tr><tr><td>Impedance converter</td><td>UFI</td><td>Model 2991</td><td>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.</td></tr><tr><td>LabChart software</td><td>ADInstruments</td><td>N/A</td><td>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.</td></tr><tr><td>LED Soldering Iron</td><td>Grainger</td><td>28EA35</td><td>This is a generic soldering iron that can be used to solder the magnetic wire to the fork terminals to create the wire electrodes.</td></tr><tr><td>PowerLab datalogger</td><td>ADInstruments</td><td>ML826</td><td>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.</td></tr><tr><td>Prism8</td><td>GraphPad</td><td>N/A</td><td>This program provides an additional option for calculating the Arrhenius Break Temperature through its &amp;ldquo;Segmental linear regression&amp;rdquo; data analysis option. This program does not require any programming and is compatible with both Mac and Windows operating systems.</td></tr><tr><td>R</td><td>R Project</td><td>N/A</td><td>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 &amp;ldquo;segmented&amp;rdquo; package. There are a number of tutorials and user guides available online through the r-project.org website.</td></tr><tr><td>Rosin Core Solder</td><td>Grainger</td><td>331856</td><td>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).</td></tr><tr><td>SAS</td><td>SAS Institute</td><td>N/A</td><td>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.</td></tr><tr><td>SigmaPlot</td><td>Systat Software, Inc.</td><td>N/A</td><td>This is the authors&amp;rsquo; preferred program for statistical determination of the Arrhenius Break Temperature. The &amp;ldquo;Regression Wizard&amp;rdquo; 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.</td></tr><tr><td>T-type Pod</td><td>ADInstruments</td><td>ML312</td><td>Suitable for measurement of temperatures from 0-50 &amp;deg;C using T-type thermocouples.</td></tr><tr><td>T-type Thermocouple Probe</td><td>ADInstruments</td><td>MLT1401</td><td>Compatible with the T-type Pod for connection. Measures temperature up to 150 &amp;deg;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.</td></tr><tr><td>UV Cable Tie, Black</td><td>Home Depot</td><td>295813</td><td>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.</td></tr></tbody></table>

impedance pneumography for minimally invasive measurement of heart rate in late stage invertebrates

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.

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 incre...

Watch this Scientific Journal Video about Impedance Pneumography for Minimally Invasive Measurement of Heart Rate in Late Stage Invertebrates at JoVE.com

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

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.

Temperatures in oceans are increasing rapidly as a consequence of widespread changes in world climates. As organismal physiology is heavily influenced by ...

impedance-pneumography-for-minimally-invasive-measurement-heart-rate

Universidad de Cartagena UDEC

Research

JoVE Journal

Environment

5.9K Views.  University of Maine. 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.

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

Right Ventricular Systolic Pressure Measurements in Combination with Harvest of Lung and Immune Tissue Samples in Mice

The function of the right heart is to pump blood through the lungs, thus linking right heart physiology and pulmonary vascular physiology. Inflammation is a common modifier of heart and lung function, by elaborating cellular infiltration, production of cytokines and growth factors, and by initiating remodeling processes 1.
Compared to the left ventricle, the right ventricle is a low-pressure pump that operates in a relatively narrow zone of pressure changes. Increased pulmonary artery pressures are associated with increased pressure in the lung vascular bed and pulmonary hypertension 2. Pulmonary hypertension is often associated with inflammatory lung diseases, for example chronic obstructive pulmonary disease, or autoimmune diseases 3. Because pulmonary hypertension confers a bad prognosis for quality of life and life expectancy, much research is directed towards understanding the mechanisms that might be targets for pharmaceutical intervention 4. The main challenge for the development of effective management tools for pulmonary hypertension remains the complexity of the simultaneous understanding of molecular and cellular changes in the right heart, the lungs and the immune system.
Here, we present a procedural workflow for the rapid and precise measurement of pressure changes in the right heart of mice and the simultaneous harvest of samples from heart, lungs and immune tissues. The method is based on the direct catheterization of the right ventricle via the jugular vein in close-chested mice, first developed in the late 1990s as surrogate measure of pressures in the pulmonary artery5-13. The organized team-approach facilitates a very rapid right heart catheterization technique. This makes it possible to perform the measurements in mice that spontaneously breathe room air. The organization of the work-flow in distinct work-areas reduces time delay and opens the possibility to simultaneously perform physiology experiments and harvest immune, heart and lung tissues.
The procedural workflow outlined here can be adapted for a wide variety of laboratory settings and study designs, from small, targeted experiments, to large drug screening assays. The simultaneous acquisition of cardiac physiology data that can be expanded to include echocardiography5,14-17 and harvest of heart, lung and immune tissues reduces the number of animals needed to obtain data that move the scientific knowledge basis forward. The procedural workflow presented here also provides an ideal basis for gaining knowledge of the networks that link immune, lung and heart function. The same principles outlined here can be adapted to study other or additional organs as needed.

A specific and rapid protocol to simultaneously investigate right heart function, lung inflammation, and the immune response is described as a learning tool. Video and figures describe physiology and microdissection techniques in an organized team-approach that is adaptable to be used for small to large sized studies.

The function of the right heart is to pump blood through the lungs, thus linking right heart physiology and pulmonary vascular physiology. Inflammation is ...

Immunology and Infection

In utero Measurement of Heart Rate in Mouse by Noninvasive M-mode Echocardiography

Congenital heart disease (CHD) is the most frequent noninfectious cause of death at birth. The incidence of CHD ranges from 4 to 50/1,000 births (Disease and injury regional estimates, World Health Organization, 2004). Surgeries that often compromise the quality of life are required to correct heart defects, reminding us of the importance of finding the causes of CHD. Mutant mouse models and live imaging technology have become essential tools to study the etiology of this disease. Although advanced methods allow live imaging of abnormal hearts in embryos, the physiological and hemodynamic states of the latter are often compromised due to surgical and/or lengthy procedures. Noninvasive ultrasound imaging, however, can be used without surgically exposing the embryos, thereby maintaining their physiology. Herein, we use simple M-mode ultrasound to assess heart rates of embryos at E18.5 in utero. The detection of abnormal heart rates is indeed a good indicator of dysfunction of the heart and thus constitutes a first step in the identification of developmental defects that may lead to heart failure.

In utero Measurement of Heart Rate in Mouse by Noninvasive M-mode Echocardiography

Ultra-high frequency ultrasound is a powerful live imaging tool to examine cardiac abnormalities in small animals. Its noninvasiveness allows maintaining the physiologic state of embryos. Herein, we show the use of M-mode ultrasound to measure the heart rates of embryos at E18.5 in utero.

Congenital heart disease (CHD) is the most frequent noninfectious cause of death at birth. The incidence of CHD ranges from 4 to 50/1,000 births (Disease ...

Medicine

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained and restrained whole larvae. For direct control of the environment around the heart another approach utilizes the dissected larvae and removal of the internal organs in order to bathe the heart in desired compounds. The exposed heart also allows membrane potentials to be monitored which can give insight of the ionic currents generated by the myocytes and for electrical conduction along the heart tube. These approaches have various advantages and disadvantages for future experiments that are discussed. The larval heart preparation provides an additional model besides the Drosophila skeletal NMJ to investigate the role of intracellular calcium regulation on cellular function. Learning more about the underlying ionic currents that shape the action potentials in myocytes in various species, one can hope to get a handle on the known ionic dysfunctions associated to specific genes responsible for various diseases in mammals.

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various ways to monitor heart function in the larva of Drosophila for assessing questions dealing with the function of gap junctions, ion channel mutations, modulation of pacemaker activity and pharmacological studies.

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained  and  ...

Biology

Characterization of the Isolated, Ventilated, and Instrumented Mouse Lung Perfused with Pulsatile Flow

The isolated, ventilated and instrumented mouse lung preparation allows steady and pulsatile pulmonary vascular pressure-flow relationships to be measured with independent control over pulmonary arterial flow rate, flow rate waveform, airway pressure and left atrial pressure. Pulmonary vascular resistance is calculated based on multi-point, steady pressure-flow curves; pulmonary vascular impedance is calculated from pulsatile pressure-flow curves obtained at a range of frequencies. As now recognized clinically, impedance is a superior measure of right ventricular afterload than resistance because it includes the effects of vascular compliance, which are not negligible, especially in the pulmonary circulation. Three important metrics of impedance - the zero hertz impedance Z0, the characteristic impedance ZC, and the index of wave reflection RW - provide insight into distal arterial cross-sectional area available for flow, proximal arterial stiffness and the upstream-downstream impedance mismatch, respectively. All results obtained in isolated, ventilated and perfused lungs are independent of sympathetic nervous system tone, volume status and the effects of anesthesia. We have used this technique to quantify the impact of pulmonary emboli and chronic hypoxia on resistance and impedance, and to differentiate between sites of action (i.e., proximal vs. distal) of vasoactive agents and disease using the pressure dependency of ZC. Furthermore, when these techniques are used with the lungs of genetically engineered strains of mice, the effects of molecular-level defects on pulmonary vascular structure and function can be determined.

The following protocol outlines the process of isolating, ventilating and instrumenting mouse lungs to measure steady or pulsatile pulmonary vascular pressure-flow relationships in order to quantify the effects of blood flow, airflow, airway changes and vascular changes on right ventricular afterload.

The isolated, ventilated and instrumented mouse lung preparation allows steady and pulsatile pulmonary vascular pressure-flow relationships to be measured ...

Intraperitoneal Glucose Tolerance Test, Measurement of Lung Function, and Fixation of the Lung to Study the Impact of Obesity and Impaired Metabolism on Pulmonary Outcomes

Obesity and respiratory disorders are major health problems. Obesity is becoming an emerging epidemic with an expected number of over 1 billion obese individuals worldwide by 2030, thus representing a growing socioeconomic burden. Simultaneously, obesity-related comorbidities, including diabetes as well as heart and chronic lung diseases, are continuously on the rise. Although obesity has been associated with increased risk for asthma exacerbations, worsening of respiratory symptoms, and poor control, the functional role of obesity and perturbed metabolism in the pathogenesis of chronic lung disease is often underestimated, and underlying molecular mechanisms remain elusive. This article aims to present methods to assess the effect of obesity on metabolism, as well as lung structure and function. Here, we describe three techniques for mice studies: (1) assessment of intraperitoneal glucose tolerance (ipGTT) to analyze the effect of obesity on glucose metabolism; (2) measurement of airway resistance (Res) and respiratory system compliance (Cdyn) to analyze the effect of obesity on lung function; and (3) preparation and fixation of the lung for subsequent quantitative histological assessment. Obesity-related lung diseases are probably multifactorial, stemming from systemic inflammatory and metabolic dysregulation that potentially adversely influence lung function and the response to therapy. Therefore, a standardized methodology to study molecular mechanisms and the effect of novel treatments is essential.

The incidence of obesity is rising and increases the risk of chronic lung diseases. To establish the underlying mechanisms and preventive strategies, well-defined animal models are needed. Here, we provide three methods (glucose-tolerance-test, body plethysmography, and lung fixation) to study the effect of obesity on pulmonary outcomes in mice.

Obesity and respiratory disorders are major health problems. Obesity is becoming an emerging epidemic with an expected number of over 1 billion obese ...

Assessment of Respiratory Function in Conscious Mice by Double-chamber Plethysmography

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to non-invasively assess some features of the respiratory function in humans as well as in laboratory animals. The present article focuses on the application of the double-chamber plethysmography (DCP) in small animals. It provides background information on the methodology as well as a detailed step-by-step procedure to successfully assess respiratory function in conscious, spontaneously breathing animals in a non-invasive manner. The DCP can be used to monitor the respiratory function of multiple animals in parallel, as well as to identify changes induced by aerosolized substances over a chosen time period and in a repeated manner. Experiments on control and allergic mice are used herein to demonstrate the utility of the technique, explain the associated outcome parameters, as well as to discuss the related advantages and shortcomings. Overall, the DCP provides valid and theoretically sound readouts that can be trusted to evaluate the respiratory function of conscious small animals both at baseline and after challenges with aerosolized substances.

The objective of the present article is to provide a detailed description of the recommended procedures to evaluate respiratory function in conscious mice by double-chamber plethysmography.

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to ...

Invasive Hemodynamic Assessment for the Right Ventricular System and Hypoxia-Induced Pulmonary Arterial Hypertension in Mice

Pulmonary arterial hypertension (PAH) is a chronic and severe cardiopulmonary disorder. Mice are a popular animal model used to mimic this disease. However, the evaluation of right ventricular pressure (RVP) and pulmonary artery pressure (PAP) remains technically challenging in mice. RVP and PAP are more difficult to measure than left ventricular pressure because of the anatomical differences between the left and right heart systems. In this paper, we describe a stable right heart hemodynamic measurement method and its validation using healthy and PAH mice. This method is based on open-chest surgery and mechanical ventilation support. It is a complicated procedure compared to closed chest procedures. While a well-trained surgeon is required for this surgery, the advantage of this procedure is that it can generate both RVP and PAP parameters at the same time, so it is a preferable procedure for the evaluation of PAH models.

Here, we present a protocol to perform an invasive hemodynamic assessment of the right ventricle and pulmonary artery in mice using an open-chest surgery approach.

Pulmonary arterial hypertension (PAH) is a chronic and severe cardiopulmonary disorder. Mice are a popular animal model used to mimic this disease. ...

Optocardiography and Electrophysiology Studies of Ex Vivo Langendorff-perfused Hearts

Small animal models are most commonly used in cardiovascular research due to the availability of genetically modified species and lower cost compared to larger animals. Yet, larger mammals are better suited for translational research questions related to normal cardiac physiology, pathophysiology, and preclinical testing of therapeutic agents. To overcome the technical barriers associated with employing a larger animal model in cardiac research, we describe an approach to measure physiological parameters in an isolated, Langendorff-perfused piglet heart. This approach combines two powerful experimental tools to evaluate the state of the heart: electrophysiology (EP) study and simultaneous optical mapping of transmembrane voltage and intracellular calcium using parameter sensitive dyes (RH237, Rhod2-AM). The described methodologies are well suited for translational studies investigating the cardiac conduction system, alterations in action potential morphology, calcium handling, excitation-contraction coupling and the incidence of cardiac alternans or arrhythmias.

The objective of this study was to establish a method for investigating cardiac dynamics using a translational animal model. The described experimental approach incorporates dual-emission optocardiography in conjunction with an electrophysiological study to assess electrical activity in an isolated, intact porcine heart model.

Small animal models are most commonly used in cardiovascular research due to the availability of genetically modified species and lower cost compared to ...

Anesthesia-free Heartbeat Measurements in Freely Moving Zebrafish

Zebrafish (Danio rerio) is a widely used model organism in physiological, pharmacological, and toxicological research due to its genetic similarity to humans and transparent embryonic stage, which facilitates non-invasive cardiovascular studies. However, current methods for heart rate assessment in zebrafish often rely on anesthesia to immobilize the subject, introducing physiological alterations that compromise data accuracy and reproducibility. This study presents a novel, anesthesia-free technique for measuring heartbeat in freely moving zebrafish larvae, addressing a critical limitation in cardiovascular research. The proposed approach integrates shortwave-infrared imaging with machine-learning-based heart tracking, allowing for precise and continuous cardiac activity monitoring in non-immobilized specimens. A convolutional neural network was trained to detect the heart region, and a photoplethysmographic signal was extracted from image sequences to determine heart rate. Experimental validation demonstrated the method's reliability and consistency across multiple test conditions. A key benefit of the methodology is its ability to preserve the natural physiological state of zebrafish, minimizing stress-induced artifacts. This non-invasive, label-free technique offers significant advantages for studying cardiovascular physiology, drug cardiotoxicity, and environmental toxicology, expanding the potential applications of zebrafish as a model for biomedical research.

The study aims to develop technology for anesthesia-free heartbeat measurements in moving zebrafish. Our approach combines shortwave-infrared imaging and machine-learning-based tracking of the heart. It is a non-invasive, label-free, and user-friendly technique that suits a wide range of studies on the zebrafish model.

Zebrafish (Danio rerio) is a widely used model organism in physiological, pharmacological, and toxicological research due to its genetic similarity to ...

Bioengineering

Electrical Impedance Tomography: Enhancing Pulmonary Health and Treatment

Monitoring Lung Function with Electrical Impedance Tomography in the Intensive Care Unit

Electrical Impedance Tomography (EIT) is a groundbreaking, non-invasive, and radiation-free imaging technique for continuous, real-time ventilation monitoring. It also has an application in pulmonary perfusion monitoring. EIT quantifies ventilation and perfusion patterns across the lung from the measurement and processing of impedance changes in the thorax. It is a powerful tool for clinicians to visualize breath-by-breath changes in pulmonary function. 
An innovative application of EIT is its ability to assess pulmonary perfusion using the kinetic analysis of a hypertonic solution injection during a breath-hold. The solution generates an impedance change in the thorax as it circulates through the pulmonary vasculature. This indirect method allows for the estimation of perfusion patterns, contributing significantly to our understanding of pulmonary blood flow dynamics at the bedside.
EIT is not just a tool for monitoring but also can be critical for the diagnosis of respiratory pathologies such as pneumothorax and bronchial intubation. It can help identify the etiology of ventilation/perfusion (V/Q) mismatch in patients receiving invasive mechanical ventilation, which is not possible with other diagnostic tools. Moreover, EIT can assist in the individual optimization of ventilator settings, such as Positive End-Expiratory Pressure (PEEP) titration and tidal volume improving oxygenation and lung health in critical care.
In summary, EIT represents a paradigm shift in bedside pulmonary monitoring and diagnostics. Its non-invasive nature and immediacy of data make EIT an indispensable tool in modern respiratory medicine. With its growing applications, EIT will be pivotal in advancing our understanding of and approach to respiratory care, particularly in intensive care settings.

Electrical Impedance Tomography is a non-invasive, radiation-free, real-time pulmonary ventilation monitoring tool. By measuring impedance changes in the thorax, it can visualize the distribution of air on a breath-by-breath basis. Initially intended for ventilation monitoring, electrical impedance tomography can also measure perfusion via intravenous injection of a saline solution.

Electrical Impedance Tomography (EIT) is a groundbreaking, non-invasive, and radiation-free imaging technique for continuous, real-time ventilation ...

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

Summary

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Chapters in this video

Right Ventricular Systolic Pressure Measurements in Combination with Harvest of Lung and Immune Tissue Samples in Mice

In utero Measurement of Heart Rate in Mouse by Noninvasive M-mode Echocardiography

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

Characterization of the Isolated, Ventilated, and Instrumented Mouse Lung Perfused with Pulsatile Flow

Intraperitoneal Glucose Tolerance Test, Measurement of Lung Function, and Fixation of the Lung to Study the Impact of Obesity and Impaired Metabolism on Pulmonary Outcomes

Assessment of Respiratory Function in Conscious Mice by Double-chamber Plethysmography

Invasive Hemodynamic Assessment for the Right Ventricular System and Hypoxia-Induced Pulmonary Arterial Hypertension in Mice

Optocardiography and Electrophysiology Studies of Ex Vivo Langendorff-perfused Hearts

Anesthesia-free Heartbeat Measurements in Freely Moving Zebrafish

Monitoring Lung Function with Electrical Impedance Tomography in the Intensive Care Unit