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 &#176;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 &#8220;Segmental linear regression&#8221; 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 &#8220;segmented&#8221; 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&#8217; preferred program for statistical determination of the Arrhenius Break Temperature. The &#8220;Regression Wizard&#8221; 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 &#176;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 &#176;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

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

Preparation and Testing of Impedance-based Fluidic Biochips with RTgill-W1 Cells for Rapid Evaluation of Drinking Water Samples for Toxicity

This manuscript describes how to prepare fluidic biochips with Rainbow trout gill epithelial (RTgill-W1) cells for use in a field portable water toxicity sensor. A monolayer of RTgill-W1 cells forms on the sensing electrodes enclosed within the biochips. The biochips are then used for testing in a field portable electric cell-substrate impedance sensing (ECIS) device designed for rapid toxicity testing of drinking water. The manuscript further describes how to run a toxicity test using the prepared biochips. A control water sample and the test water sample are mixed with pre-measured powdered media and injected into separate channels of the biochip. Impedance readings from the sensing electrodes in each of the biochip channels are measured and compared by an automated statistical software program. The screen on the ECIS instrument will indicate either "Contamination Detected" or "No Contamination Detected" within an hour of sample injection. Advantages are ease of use and rapid response to a broad spectrum of inorganic and organic chemicals at concentrations that are relevant to human health concerns, as well as the long-term stability of stored biochips in a ready state for testing. Limitations are the requirement for cold storage of the biochips and limited sensitivity to cholinesterase-inhibiting pesticides. Applications for this toxicity detector are for rapid field-portable testing of drinking water supplies by Army Preventative Medicine personnel or for use at municipal water treatment facilities.

This manuscript describes how to prepare fluidic biochips with Rainbow trout gill epithelial cells for use in a field portable electric cell-substrate impedance sensor. The protocol for running a rapid drinking water toxicity test with the sensor is also described.

This manuscript describes how to prepare fluidic biochips with Rainbow trout gill epithelial (RTgill-W1) cells for use in a field portable water toxicity ...

Enhancement of the Initial Growth Rate of Agricultural Plants by Using Static Magnetic Fields

Electronic devices and high-voltage wires induce magnetic fields. A magnetic field of 1,300-2,500 Gauss (0.2 Tesla) was applied to Petri dishes containing seeds of Garden Balsam (Impatiens balsamina), Mizuna (Brassica rapa var. japonica), Komatsuna (Brassica rapa var. perviridis), and Mescluns (Lepidium sativum). We applied magnets under the culture dish. During the 4 days of application, we observed that the stem and root length increased. The group subjected to magnetic field treatment (n = 10) showed a 1.4 times faster rate of growth compared with the control group (n = 11) in a total of 8 days (p &#60;0.0005). This rate is 20% higher than that reported in previous studies. The tubulin complex lines did not have connecting points, but connecting points occur upon the application of magnets. This shows complete difference from the control, which means abnormal arrangements. However, the exact cause remains unclear. These results of growth enhancement of applying magnets suggest that it is possible to enhance the growth rate, increase productivity, or control the speed of germination of plants by applying static magnetic fields. Also, magnetic fields can cause physiological changes in plant cells and can induce growth. Therefore, stimulation with a magnetic field can have possible effects that are similar to those of chemical fertilizers, which means that the use of fertilizers can be avoided.

The goal of this protocol is to demonstrate the acceleration of the initial growth rate of plants by applying static magnetic fields with no external energy.

Electronic devices and high-voltage wires induce magnetic fields. A magnetic field of 1,300-2,500 Gauss (0.2 Tesla) was applied to Petri dishes containing ...

Implementation of Portable Emissions Measurement Systems (PEMS) for the Real-driving Emissions (RDE) Regulation in Europe

Vehicles are tested in controlled and relatively narrow laboratory conditions to determine their official emission values and reference fuel consumption. However, on the road, ambient and driving conditions can vary over a wide range, sometimes causing emissions to be higher than those measured in the laboratory. For this reason, the European Commission has developed a complementary Real-Driving Emissions (RDE) test procedure using the Portable Emissions Measurement Systems (PEMS) to verify gaseous pollutant and particle number emissions during a wide range of normal operating conditions on the road. This paper presents the newly-adopted RDE test procedure, differentiating six steps: 1) vehicle selection, 2) vehicle preparation, 3) trip design, 4) trip execution, 5) trip verification, and 6) calculation of emissions. Of these steps, vehicle preparation and trip execution are described in greater detail. Examples of trip verification and the calculations of emissions are given.

Implementation of Portable Emissions Measurement Systems PEMS for the Real-driving Emissions RDE Regulation in Europe

The European Commission has developed a Real-Driving Emissions (RDE) test procedure to verify pollutant emissions during real-world vehicle operation using the Portable Emissions Measurement Systems (PEMS). This paper presents the experimental procedures required by the newly-adopted RDE test.

Vehicles are tested in controlled and relatively narrow laboratory conditions to determine their official emission values and reference fuel consumption. ...

Production and Measurement of Organic Particulate Matter in a Flow Tube Reactor

Organic particulate matter (PM) is increasingly recognized as important to the Earth's climate system as well as public health in urban regions, and the production of synthetic PM for laboratory studies have become a widespread necessity. Herein, experimental protocols demonstrate approaches to produce aerosolized organic PM by &#945;-pinene ozonolysis in a flow tube reactor. Methods are described for measuring the size distributions and morphology of the aerosol particles. The video demonstrates basic operations of the flow tube reactor and related instrumentation. The first part of the video shows the procedure for preparing gas-phase reactants, ozonolysis, and production of organic PM. The second part of the video shows the procedures for determining the properties of the produced particle population. The particle number-diameter distributions show different stages of particle growth, namely condensation, coagulation, or a combination of both, depending on reaction conditions. The particle morphology is characterized by an aerosol particle mass analyzer (APM) and a scanning electron microscope (SEM). The results confirm the existence of non-spherical particles that have grown from coagulation for specific reaction conditions. The experimental results also indicate that the flow tube reactor can be used to study the physical and chemical properties of organic PM for relatively high concentrations and short time frames.

This paper describes the operation procedure for the flow tube reactor and related data collection. It shows the protocols for setting the experiments, recording data and generating the number-diameter distribution as well as the particle mass information, which gives useful information about chemical and physical properties of the organic aerosols.

Organic particulate matter (PM) is increasingly recognized as important to the Earth's climate system as well as public health in urban regions, and the ...

Determination of the Settling Rate of Clay/Cyanobacterial Floccules

The mechanisms underpinning the deposition of fine-grained, organic-rich sediments are still largely debated. Specifically, the impact of the interaction of clay particles with reactive, planktonic cyanobacterial cells to the sedimentary record is under studied. This interaction is a potentially major contributor to shale depositional models. Within a lab setting, the flocculation and sedimentation rates of these materials can be examined and measured in a controlled environment. Here, we detail a protocol for measuring the sedimentation rate of cyanobacterial/clay mixtures. This methodology is demonstrated through the description of two sample experiments: the first uses kaolin (a dehydrated form of kaolinite) and Synechococcus sp. PCC 7002 (a marine coccoid cyanobacteria), and the second uses kaolin and Synechocystis sp. PCC 6803 (a freshwater coccoid cyanobacteria). Cyanobacterial cultures are mixed with varying amounts of clay within a specially designed tank apparatus optimized to allow continuous, real-time video and photographic recording. The sampling procedures are detailed as well as a post-collection protocol for precise measurement of chlorophyll a from which the concentration of cyanobacterial cells remaining in suspension can be determined. Through experimental replication, a profile is constructed that displays sedimentation rate.

The interaction and sedimentation of the clay and bacterial cells within the marine realm, observed in natural environments, can be best investigated in a controlled lab environment. Here, we describe a detailed protocol, which outlines a novel method for measuring the sedimentation rate of clay and cyanobacterial floccules.

The mechanisms underpinning the deposition of fine-grained, organic-rich sediments are still largely debated. Specifically, the impact of the interaction ...

Double-stranded RNA Oral Delivery Methods to Induce RNA Interference in Phloem and Plant-sap-feeding Hemipteran Insects

Phloem and plant sap feeding insects invade the integrity of crops and fruits to retrieve nutrients, in the process damaging food crops. Hemipteran insects account for a number of economically substantial pests of plants that cause damage to crops by feeding on phloem sap. The brown marmorated stink bug (BMSB), Halyomorpha halys (Heteroptera: Pentatomidae) and the Asian citrus psyllid (ACP), Diaphorina citri Kuwayama (Hemiptera: Liviidae) are hemipteran insect pests introduced in North America, where they are an invasive agricultural pest of high-value specialty, row, and staple crops and citrus fruits, as well as a nuisance pest when they aggregate indoors. Insecticide resistance in many species has led to the development of alternate methods of pest management strategies. Double-stranded RNA (dsRNA)-mediated RNA interference (RNAi) is a gene silencing mechanism for functional genomic studies that has potential applications as a tool for the management of insect pests. Exogenously synthesized dsRNA or small interfering RNA (siRNA) can trigger highly efficient gene silencing through the degradation of endogenous RNA, which is homologous to that presented. Effective and environmental use of RNAi as molecular biopesticides for biocontrol of hemipteran insects requires the in vivo delivery of dsRNAs through feeding. Here we demonstrate methods for delivery of dsRNA to insects: loading of dsRNA into green beans by immersion, and absorbing of gene-specific dsRNA with oral delivery through ingestion. We have also outlined non-transgenic plant delivery approaches using foliar sprays, root drench, trunk injections as well as clay granules, all of which may be essential for sustained release of dsRNA. Efficient delivery by orally ingested dsRNA was confirmed as an effective dosage to induce a significant decrease in expression of targeted genes, such as juvenile hormone acid O-methyltransferase (JHAMT) and vitellogenin (Vg). These innovative methods represent strategies for delivery of dsRNA to use in crop protection and overcome environmental challenges for pest management.

This article demonstrates novel techniques developed for oral delivery of double-stranded RNA (dsRNA) through the vascular tissues of plants for RNA interference (RNAi) in phloem sap feeding insects.

Phloem and plant sap feeding insects invade the integrity of crops and fruits to retrieve nutrients, in the process damaging food crops. Hemipteran ...

Detached Leaf Assays to Simplify Gene Expression Studies in Potato During Infestation by Chewing Insect Manduca sexta

The multitrophic nature of gene expression studies of insect herbivory demands large numbers of biological replicates, creating the need for simpler, more streamlined herbivory protocols. Perturbations of chewing insects are usually studied in whole plant systems. While this whole organism strategy is popular, it is not necessary if similar observations can be replicated in a single detached leaf. The assumption is that basic elements required for signal transduction are present within the leaf itself. In the case of early events in signal transduction, cells need only to receive the signal from the perturbation and transmit that signal to neighboring cells which are assayed for gene expression.
The proposed method simply changes the timing of the detachment. In whole plant experiments, larvae are confined to a single leaf which is eventually detached from the plant and assayed for gene expression. If the order of excision is reversed, from last in whole plant studies, to first in the detached study, the feeding experiment is simplified.
Solanum tuberosum var. Kennebec is propagated by nodal transfer in a simple tissue culture medium and transferred to soil for further growth if desired. Leaves are excised from the parent plant and relocated to Petri dishes where the feeding assay is conducted with the larval stages of M. sexta. Damaged leaf tissue is assayed for the expression of relatively early events in signal transduction. Gene expression analysis identified infestation-specific Cys2-His2 (C2H2) transcription factors, confirming the success of using detached leaves in early response studies. The method is easier to perform than whole plant infestations and uses less space.

The presented method creates natural herbivore damaged plant tissue through the application of Manduca sexta larvae to detached leaves of potato. The plant tissue is assayed for expression of six transcription factor homologs involved in early responses to insect herbivory.

The multitrophic nature of gene expression studies of insect herbivory demands large numbers of biological replicates, creating the need for simpler, more ...

Using Deuterium Oxide as a Non-Invasive, Non-Lethal Tool for Assessing Body Composition and Water Consumption in Mammals

Body condition scoring systems and body condition indices are common techniques used for assessing the health status or fitness of a species. Body condition scoring systems are evaluator dependent and have the potential to be highly subjective. Body condition indices can be confounded by foraging, the effects of body weight, as well as statistical and inferential problems. An alternative to body condition scoring systems and body condition indices is using a stable isotope such as deuterium oxide to determine body composition. The deuterium oxide dilution method is a repeatable, quantitative technique used to estimate body composition in humans, wildlife, and domestic species. Additionally, the deuterium oxide dilution technique can be used to determine the water consumption of an individual animal. Here, we describe the adaption of the deuterium oxide dilution technique for assessing body composition in big brown bats (Eptesicus fuscus) and for assessing water consumption in cats (Felis catis).

This article describes the deuterium oxide dilution technique in two mammals, an insectivore and carnivore, to determine total body water, lean body mass, body fat mass, and water consumption.

Body condition scoring systems and body condition indices are common techniques used for assessing the health status or fitness of a species. Body ...

PARbars: Cheap, Easy to Build Ceptometers for Continuous Measurement of Light Interception in Plant Canopies

Ceptometry is a technique used to measure the transmittance of photosynthetically active radiation through a plant canopy using multiple light sensors connected in parallel on a long bar. Ceptometry is often used to infer properties of canopy structure and light interception, notably leaf area index (LAI) and effective plant area index (PAIeff). Due to the high cost of commercially available ceptometers, the number of measurements that can be taken is often limited in space and time. This limits the usefulness of ceptometry for studying genetic variability in light interception, and precludes thorough analysis of, and correction for, biases that can skew measurements depending on the time of day. We developed continuously logging ceptometers (called PARbars) that can be produced for USD $75 each and yield high quality data comparable to commercially available alternatives. Here we provide detailed instruction on how to build and calibrate PARbars, how to deploy them in the field and how to estimate PAI from collected transmittance data. We provide representative results from wheat canopies and discuss further considerations that should be made when using PARbars.

Here, we present detailed instructions on how to build and calibrate research quality ceptometers (light sensors that integrate light intensity across many sensors arrayed linearly along a horizontal bar).

Ceptometry is a technique used to measure the transmittance of photosynthetically active radiation through a plant canopy using multiple light sensors ...

Laser-Induced Fluorescence Emission (L.I.F.E.) as Novel Non-Invasive Tool for In-Situ Measurements of Biomarkers in Cryospheric Habitats

Global warming affects microbial communities in a variety of ecosystems, especially cryospheric habitats. However, little is known about microbial-mediated carbon fluxes in extreme environments. Hence, the methodology of sample acquisition described in the very few studies available implies two major problems: A) high resolution data require a large number of samples, which is difficult to obtain in remote areas; B) unavoidable sample manipulation such as cutting, sawing, and melting of ice cores that leads to a misunderstanding of in situ conditions. In this study, a prototype device that requires neither sample preparation nor sample destruction is presented. The device can be used for in situ measurements with a high spectral and spatial resolution in terrestrial and ice ecosystems and is based on the Laser-Induced Fluorescence Emission (L.I.F.E.) technique. Photoautotrophic supraglacial communities can be identified by the detection of L.I.F.E. signatures in photopigments. The L.I.F.E. instrument calibration for the porphyrin derivates chlorophylla (chla) (405 nm laser excitation) and B-phycoerythrin (B-PE) (532 nm laser excitation) is demonstrated. For the validation of this methodology, L.I.F.E. data were ratified by a conventional method for chla quantification that involved pigment extraction and subsequent absorption spectroscopy. The prototype applicability in the field was proven in extreme polar environments. Further testing on terrestrial habitats took place during Mars analog simulations in the Moroccan dessert and on an Austrian rock glacier. The L.I.F.E. instrument enables high resolution scans of large areas with acceptable operation logistics and contributes to a better understanding of the ecological potential of supraglacial communities in the context of global change.

Laser-Induced Fluorescence Emission L.I.F.E. as Novel Non-Invasive Tool for In-Situ Measurements of Biomarkers in Cryospheric Habitats

Carbon fluxes in the cryosphere are hardly assessed yet but are crucial regarding climate change. Here we show a novel prototype device that captures the phototrophic potential in supraglacial environments based on laser-induced fluorescence emission (L.I.F.E.) technology offering high spectral and spatial resolution data under in situ conditions.