We thank Melinda Forseth for taking photos of oysters and measuring their gape width in ImageJ. We thank Denise Breitburg for access to the aquaria with diel cycling hypoxia and cyclical pH conditions. We thank the Smithsonian Environmental Research Center, Edgewater, Maryland, for space for the experiments. The hypoxia experiments were funded by a National Oceanic and Atmospheric Administration - Center for Sponsored Coastal Ocean Research grant No. NA10NOS4780138 and the Smithsonian Hunterdon Fund to Denise Breitburg. The valve gape measurements during the hypoxia experiments were funded by a Faculty Enhancement Grant by Washington College to Elka T. Porter.

1. Construction of the Wheatstone Bridge for the Valve Gape Apparatus
NOTE: The strain gauge is nominally 1000 &#937;, so to fully balance and null the bridge, all components should be 1000 &#937;.
<ol>
	<li>As in Figure 1, solder two 1 k&#937; precision resistors to each other and then to a ~976 &#937; resistor and a 100 &#937; 10 turn potentiometer. The typical strain gauge range is a few &#937; off of the 1000 &#937; nominal, so only this resistance needs to be matched. Use a fixed 976 &#937; resistor and an adjustable resistor (10 turn potentiometer) for this purpose.</li>
	<li>Bring one side of the variable arm and one side of the 1 k&#937; resistor to a connector to attach to the bivalve strain gauge as shown in Figure 1.</li>
	<li>Attach the top and the bottom of the bridge to a power supply to provide bias current to the bridge as shown in Figure 1.</li>
	<li>Attach the midpoints of the bridge to the preamplifier and data-recorder, as shown in Figure 1. This will record the bivalve response as its gape movement modulates the resistance of the strain gauge.</li>
	<li>After the bivalve strain gauge is attached, null the bridge by adjusting the 10 turn potentiometer until the voltage at the data recorder input is zero. Use a separate BNC connector and a hand held voltmeter for this purpose. 
	NOTE: Nulling the output means that very high gains can be used with the data recorder without saturating the input, allowing very precise voltages (gape response) to be measured. 
	NOTE: Our multichannel setup uses discrete components hand soldered on a standard piece of electronics &#34;perf-board&#34;. This could easily be formalized on a very inexpensive circuit board with little additional effort.</li>
</ol>
2. General Setting for Diel-Cycling Hypoxia and Cyclical pH
<ol>
	<li>Keep oysters with known low disease prevalence61 in 75-L aquaria under a 14 h light: 10 h dark photoperiod, 7 days per week48, and expose them to cyclical changes in [DO] mimicking diel-cycling hypoxia as found in nature7,8. During 14 h of light, simulate the light levels on a sunny day at 2-m depth in the Rhode River, Maryland61.</li>
	<li>Set up diel-cycling hypoxia and cyclical pH in the laboratory for long-term experiments as described in detail by Burrell et al. (2015)49. Manipulate [DO] and pH in each aquarium and hold at pre-determined levels at all times. To avoid valve closure due to food limitation, add phytoplankton62 to the aquaria. Compensate for any potential DO uptake from algal decomposition by regulating [DO] in each aquarium.</li>
</ol>
3. Bivalve Acclimation
<ol>
	<li>Acclimate oysters to diel-cycling hypoxia for about one week before performing gape experiments.</li>
</ol>
4. Setting Up Diel-Cycling Hypoxia Plateau Stages
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57404/57404fig2.jpg" /> 
Figure 2. [DO] was manipulated experimentally every day to generate diel cycling hypoxia. Here, 2 consecutive diel cycles are shown over which gape measurements were taken. Each diel cycle contains four [DO] phases: a = supersaturation (SS), b = normoxia after supersaturation, c = low plateau (LP), d = normoxia after low plateau. For the Low Plateau, severe hypoxia ([DO] = 0.6 mg/L) with control or cycling pH is indicated with a thick black line, mild hypoxia ([DO] = 1.7 mg/L) with cycling pH with a thin grey line and normoxia ([DO] = 7.3 mg/L) with a dashed line. Grey boxes indicate 10-h dark intervals over two days. <a href="/files/ftp_upload/57404/57404fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>To create diel cycling hypoxia, adjust [DO] over the course of a day in four consecutive phases where [DO] is held constant: 2 h supersaturation, 6 h normoxia after supersaturation, 4 h low plateau (normoxia in the case of the controls) and 2 h normoxia after the low plateau phases.</li>
	<li>During the low plateau phase, keep the normoxic treatment constant at a [DO] of 7.3 mg/L, the mildly hypoxic treatment constant at a [DO] of 1.7 mg/L, and the severe hypoxia treatments constant at a [DO] of 0.6 mg/L.</li>
	<li>For severe hypoxia, run two pH treatments during the low plateau, i.e., severe hypoxia with control pH in which the pH is kept at 7.8 and severe hypoxia with cyclical pH where the pH is cycled from 7.8 to 7.0 during the low plateau. Run the mildly hypoxic treatment with cyclical pH where pH is cycled from 7.8 to 7.0. Run a second mildly hypoxic treatment with constant pH if desired.</li>
</ol>
5. Ramping Up/Down of [DO] Between Plateau Stages
<ol>
	<li>Schedule the ramp up or down of [DO] between the four constant phases to take 2 h each, except for the ramp down from normoxia after supersaturation to the low plateau phase, which takes 3 h. Often, hypoxia has been found to occur late at night or early morning hours in the summer7,8; thus, keep the low plateau phase in the dark as well as the periods ramping down to it or up from it, for a total length of darkness of 10 h.</li>
	<li>Repeat diel-cycling hypoxia 4-5 days a week61.</li>
	<li>Add oysters with sensors in the ramp-up phase before supersaturation and remove them, after closing them, two days later so they experience two complete diel cycles (Figure 2) with sensors attached.</li>
</ol>
6. Making the Sensor Cable
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57404/57404fig3.jpg" /> 
Figure 3. Bivalve SGM strain gauge valve sensor. (a) Strain gauge sensor, (b) strain gauge attached to an oyster showing a loop of the strain gauge between both valves, (c) strain gauge sensor mounted on an oyster. <a href="/files/ftp_upload/57404/57404fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Construct valve gape sensor cables by twisting two 32 AWG (7/40) wires (see Table of Materials) of desired color (Figure 3a). Cut a length of wire twice the length of the desired gape sensor cable, i.e., twice the length from the aquarium to the SGM plus a little extra.</li>
	<li>Have two people stand across from each other, each holding one end of the cable.</li>
	<li>Slide the handle of a pair of scissors over the cable so that in the middle of the cable a pair of scissors is held by the cable. Keep the cable under tension at all times (it cannot go slack).</li>
	<li>Have both people twist the cable in the opposite direction keeping the cable under tension. As the cable starts to twist, the pair of scissors in the middle (which provides weight) will sink down, and both ends of the cable can twist around each other.</li>
	<li>Put the two ends of the cable together and hold them high up, not allowing the scissors with the cable to touch the ground or go slack. After the cable is finished twisting, cut the cable on the other end so there are 4 cable leads total, two on each end.</li>
	<li>Alternatively, to speed this step up, have one person hold a drill with the wires clamped in on one end, keep tension on the wires, and twist the wires by turning the drill on.</li>
	<li>Strip 2 cm of insulation from the wires at each end of the twisted pair.</li>
</ol>
7. Soldering the Sensor
<ol>
	<li>Tin the exposed cable.</li>
	<li>Solder Dsub connector male crimp pins on the two leads on one end of the sensor cable by pretinning the pins and inserting the cables.</li>
	<li>Then, insulate the leads by putting shrink tubing over each lead. Choose shrink tubing that is not much larger in diameter than the cable leads. Shrink the heat shrink with a heat gun. The shrink tubing should now be tightly wrapped around the leads. 
	NOTE: These two leads get plugged into the Wheatstone bridge during experiments.</li>
	<li>Solder the remaining end of the sensor cable to the two contacts of a 1000 &#937; strain gauge with dimensions of 13.5 mm * 5.5 mm grids on the remaining two leads (Figure 3a).</li>
	<li>Add a sensor identifying tag to the sensor cable (Figure 3a).</li>
</ol>
8. Sealing the Sensor
Note: The strain gauges corrode quickly in seawater and thus must be very well sealed. The quality of the seal affects the useful life of the sensor.
<ol>
	<li>Seal the strain gauges by attaching them to PCT-2A&#160;tape leaving 15 mm of tape at the long ends and 2 mm on the sides of the sensor (Figure 3a). After attaching PCT-2A tape to the underside of the strain gauge, add quick gel on the sensor leads and the attached open ends of the cables.</li>
	<li>Attach a second piece of PCT-2A tape of the same size as the bottom piece to the top side of the sensor and attach both pieces of tape with the strain gauge in the middle together. Do not allow the quick gel to flow anywhere other than over the leads and the sensor cables.</li>
	<li>To improve the seal, add thin strips of PCT-2A&#160;tape to the edges where the two pieces of tape surrounding the sensor meet. Do this for the three exposed edges of the sensors.</li>
	<li>Repeat steps 6.1 to 8.4 eleven times to construct 12 sensor cables with sensors, one for each channel on the Wheatstone bridge. 
	Note:&#160;Successful sealing of the strain gauges was the key variable affecting bivalve gape sensor survival in seawater and for minimizing occasional &#34;jumps&#34; in sensor output. Earlier trials without the sealing technique described above resulted in early sensor corrosion and subsequent failure, or rapid drift beyond the measurement range. Future improvements should include techniques to improve the seal.</li>
</ol>
9. Attaching the Valve Gape Sensor to a Bivalve (Figure 3b,c)
<ol>
	<li>Remove oyster from diel-cycling hypoxia aquarium before the low plateau phase.</li>
	<li>With nail polish write an identifying number on the clean, dry shell of each C. virginica.</li>
	<li>Attach the sealed strain gauges to the clean and dried valves of live C. virginica (Figure 3b, c) using aquarium sealant to allow the sensor to be removed after the run. Attach the gape sensor with the leads on the right (flat) valve of oyster about mid-way on the shell (Figure 3c) and leave a loop of the strain gauge (Figure 3b) across the valves so as not to impede the opening of the valves (Figure 3b). Glue the remaining free end of the sensor on the left valve. 
	NOTE: Observations confirm that oysters open their valves normally with the sensors in place.</li>
	<li>It does not matter which way around the sensor is glued on, however, the direction of the signal will be altered if some leads are glued on the left valve and some on the right valve. The direction of the signal becomes clear from the forced closing of the bivalve at the end of a run. For all bivalves, gluing the strain gauge part with the sensor leads to the same valve side for a consistent direction of change in the gape response signal.</li>
	<li>During each step of the process of sealing sensors and attaching them to the oysters, check the sensor resistance periodically to make sure it stays within the range of the readout electronics and to check for potential sensor damage during processing.</li>
	<li>Attach 12 oysters to 12 sensors to fill all 12 channels of the SGM.</li>
	<li>Add oysters to the different aquaria just before the supersaturation plateau phase.</li>
</ol>
10. Setup of Datalogger and Multiplexer for the Valve Gape Apparatus
<ol>
	<li>Attach a sensor to the bivalve and then connect the cable to the SGM with individual Wheatstone bridges (Figure 1) for each sensor (a total of 12 bridges) and connected to a datalogger.</li>
	<li>Set the data logger sample period to read once every 2.5 s and then averaged for 15 s. 
	NOTE: The 62k sample memory on the datalogger used here allowed 12 h of unattended operation. Larger dataloggers and faster sampling rates can be selected, if desired.</li>
	<li>Zero the gape signal for each individual sensor after attaching the sensor to a bivalve using the potentiometer dials on the SGM.</li>
	<li>When necessary, as in higher salinities, correct each sensor periodically for drift in sensor response so that the sensor remains in the range of the SGM (-25 to +25 mV) output of the Wheatstone bridge. The SGM range was chosen for maximum sensitivity between the Wheatstone bridge and the input of the datalogger.</li>
</ol>
11. Deployment of Bivalves with Gape Sensors in Diel-Cycling Hypoxia
<ol>
	<li>After the aquarium sealant has cured (ca 30 min) return the bivalves to an oxygenated holding bucket with flow-through water for at least 5 h to allow the aquarium sealant to leach before submersion in the experiment.</li>
	<li>Place oysters in the aquariums during the ramp-up phase to the supersaturation (SS) plateau (Figure 2).</li>
</ol>
12. Recovery of Bivalves With Gape Sensors
<ol>
	<li>When recovering the oyster from the aquarium with diel-cycling hypoxiaat the end of the two day period, use your fingers to trigger the oyster to close and produce a closing signal in the sensor voltage. This way, the direction and the magnitude of the closing signal can be recorded</li>
	<li>Define the oyster as open if the sensor value is larger than &#189; of the closed value. Define the oyster as closed if the sensor value is smaller than &#189; the closing value. 
	NOTE: Eastern oysters do not open much at the intermediate range.</li>
</ol>
13. Linearization of Raw Valve Gape Data
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57404/57404fig4.jpg" /> 
Figure 4.Linearization of the raw gape data of an oyster. (a) 14 h light: 10 h dark phases, (b) [DO] over a two-day period with periods of severe hypoxia ([DO] 0.6 mg/L), (c) Raw gape data time series of one oyster. Linear drift of the data is caused by corrosion of the sensor and has to be removed in post-processing, (d) gape data time series after post-processing, (e) approximate real time. To determine the magnitude and the direction of the closing spike, the oyster is triggered to close at the end of a run. Magnitudes vary between different sensors. <a href="/files/ftp_upload/57404/57404fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Visually identify oyster gape activity, and use the closed (or open, whichever one is longer) phases of the bivalves to determine instrument drift. Use IF statements in spreadsheet software to mark the closed values with one and open values with zero (e.g., =IF (Cell &#62; 5, 1 (true), 0 (false))).</li>
	<li>Remove the shortest phases and run a linear regression on the remaining raw data. Use the regression equation to linearize the experimental data (Figure 4). Linearization has to be done for each sensor individually. 
	Note: Sensor drift is linear over short (minutes) and long (days) time scales and the amount of drift varies from sensor to sensor. Drift is typically negligible over short time spans such as an hour, in which only occasionally linear drift has been noted. Sensor drift necessitates that calibrations of valve gape and sensor gape have to be done for each sensor separately and that a linearization has to be done for each sensor in post-processing of the data.</li>
</ol>
14. Calibration of Sensors: Actual Gape Width Versus Sensor mV
In a side experiment, sensor voltage can be calibrated to the actual gape width of oysters.
<ol>
	<li>In an aquarium, set up an oyster with a gape sensor. Initiate gape recording. Trigger the oyster to close by touching it. Soon after, the oyster will open up again.</li>
	<li>As the oyster is opening, take photographs and write down the sensor voltage of the gape apparatus each time a photograph is taken. Keep track of which photograph goes with which gape apparatus reading. Place a reference scale in the photograph, at the same level of the valves.</li>
	<li>Analyze the distance the oyster is open in each photograph (mm) using an open source image processing program such as ImageJ.</li>
	<li>Compare actual gape width to sensor voltage and run a regression analysis. This gives the calibration for the actual opening status and further sensor measurements can then be related to actual gape width of the bivalve. 
	Note: While this shows gape width is linearly related to sensor voltage, in actuality, this process takes too long and was not performed for the hypoxia gape experiments. For the hypoxia gape experiments it was important to remove the oysters for the least amount of time necessary from their aquaria to equip them with sensors and return them to the aquaria. For some experiments, the gape width measurement may be important and the calibration techniques above should be followed.</li>
</ol>

We don't have anything to disclose.

Typical studies focus on the continuous, extended time periods of low oxygen conditions and the response, often measured as the survival, of animals. However, at present, our understanding of the behavioral responses of animals to diel-cycling hypoxia is minimal63. Thus, more studies should focus on the behavior of organisms in response to diel-cycling hypoxia which occurs regularly over the summer in many estuaries7,8.
<p class="jove_...

Hypoxia, i.e., dissolved oxygen concentrations [DO] sufficiently low to negatively affect biological and ecological processes but often functionally defined as [DO] of &#60; 2 mg/L1, and anoxia (functionally defined as [DO] of 0.0-0.2 mg/L) are occurring more frequently and severely in the world&#39;s coastal waters, estuaries and the deep ocean2,3 and are often exacerbated by increasing eutrophication4,5. With an increasing areal extent of hypoxia and anoxia, macrofauna are negatively affected and lose habitat extent and habitat quality. Climate change is predicted to worsen hypoxia and anoxia6.
In many stratified, nutrient-enriched estuaries such as Chesapeake Bay, USA, seasonally persistent hypoxia can prevail and can occur year after year2. In addition, diel-cycling of hypoxia is frequent in estuaries such as the Chesapeake Bay and other locations and occurs late during the night or the early morning hours in the summer7,8.
Most studies have focused on the effect of continuous exposure of organisms to low [DO] and on their tolerance to hypoxia and anoxia9,10,11,12,13,14. Moreover, studies have looked at the large-scale shift in species distributions, abundances, and species composition in response to extended low [DO]4,15. Often species that are very sensitive to low [DO], die in masses,16 shifting the remaining species to a younger, smaller-sized, short-lived fauna as, for example, found on the Louisiana-Texas Shelf ecosystem4.
Behavioral changes typically precede community collapse17 and studies have reported on behavioral responses of organisms to extended low [DO]4,16,17,18,19,20,21,22,23,24,25. These studies, however, do not focus on the responses of organisms to diel-cycling exposures of hypoxia and the fluctuating nature of [DO] availability in estuaries.
Diel-cycling hypoxia in shallow estuaries has received increasing awareness as studies monitor [DO] more frequently over the course of days with sondes in estuaries16,26. Water can remain hypoxic for hours at the end of the night or early morning hours in the summer when there is no oxygen-generating photosynthesis during the night but high oxygen-consuming aerobic respiration7,16. It was also found that the tides affected the diel cycling of low DO conditions with the most extreme minima observed when low tides coincided with the end of the night27. Only after several hours of hypoxia does [DO] come back to normoxia7,16,28 in the daily cycle.
To determine the behavioral response of C. virginica to diel-cycling hypoxia and pH we monitored the opening and closing of the valves of oysters exposed to laboratory induced diel cycling of [DO] and cyclical pH. Gape responses of bivalves have been used to detect adverse environmental conditions. Valve closures of bivalves in response to contaminants29,30,31, toxic algae32,33,34, thermal pollution35,36,37, food quantity decrease38,39,40, feeding regime39,41, emersion37,42, photoperiod43,44, pH45,46, and combined pH and dissolved oxygen47 have been measured. Gape techniques have, for example, included direct observations48,49,13, continuous measurements using reed switches and magnets (Dreissena monitor)50, or fiber-optic sensors51 that require clear water. In addition, magnet and magnetic-field strength Hall sensors have been used to study mussel gape angle52,53,54,55, and a high-frequency electromagnetic induction system that can measure the varying distance between two electric coils that are glued on the valves has been used56,57,58,59. A high voltage source is required for the electromagnetic induction system and power has to be delivered to both sides of the shell52. This system is also commercially available as the &#34;MOSSELMONITOR&#34; (http://mosselmonitor.nl/).
On a tight research budget, we constructed an inexpensive strain gauge monitor (SGM) to continuously measure oyster gape over laboratory-induced diel cycling of [DO] and pH, under low visibility conditions. Our system is also much simpler than competing systems, allowing many animals to be instrumented during an experiment. We wanted to determine the behavioral responses of C. virginica to diel cycling severe ([DO] = 0.6 mg/L) hypoxia with control pH (pH = 7.8) and cycling pH (pH = 7.8-7.0), respectively, and gape responses to mild ([DO] = 1.7 mg/L) hypoxia. Moreover, we wanted to determine if oysters are able to rapidly respond to changes in [DO] over the diel cycle and how they respond when normoxia returns after a hypoxic event. Perhaps oysters are optimally adapted to the rapidly fluctuating environment that is found in many estuaries16,27 where they live. While more complex valve gape monitors are available, the SGM offers an inexpensive technique that allows continuous measurements of valve gape in waters even in low visibility conditions.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57404/57404fig1.jpg" /> 
Figure 1.&#160;Wheatstone bridge for the valve gape apparatus. <a href="/files/ftp_upload/57404/57404fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The strain gauge sensors used for monitoring bivalve gape are resistive films in a meander pattern on a polyimide backing. Small amounts of strain modulate the resistance of the sensor. The bivalve flexes the strain gage when it gapes causing a change in the sensor&#39;s resistance. We employed a nulling, balanced, Wheatstone bridge for each bivalve channel as shown in Figure 1 to measure the change in sensor resistance. The Wheatstone bridge is nulled by the potentiometer allowing a fairly high gain to be employed by the datalogger. A Wheatstone bridge is a standard method for accurately measuring an unknown resistance using a ratio to a known resistance standard and a voltmeter. The history of this very old technique is discussed in Ekelof (2001)60. We integrated 12 channels, each with its own Wheatstone bridge and nulling potentiometer, into the Strain Gauge Monitor (SGM) unit.

<table><tbody><tr><td>Campbell CR 10x data logger</td><td>Campbell Scientific, Logan, Utah</td><td></td><td>Or other data logger. At Campbell the CR 10X has been replaced with the CR 1000</td></tr><tr><td>Campbell CR 10x multiplexer</td><td>Campbell Scientific, Logan, Utah</td><td></td><td>Data logger needs to have space for 12 channels</td></tr><tr><td>Dsub connector male crimp pins</td><td>TE Connectivity</td><td>205089-1</td><td>pins for gape sensor leads</td></tr><tr><td>PCA tape</td><td>Micro Measurements Corp, NC</td><td></td><td>To seal the strain gauge</td></tr><tr><td>Duro Quick Gel</td><td>Ace Hardware</td><td></td><td>Superglue</td></tr><tr><td>SG13/1000-LY43 or LY41</td><td>Omega Engineering Inc., Stanford, CT</td><td></td><td>Strain gauges</td></tr><tr><td>32 AWG (7/40) teflon Alpha wires</td><td>AlphaWire, Elizabeth, NJ</td><td>2840/7</td><td>Sensor cables, different colors are available</td></tr><tr><td>1/16&quot; heat shrink tubing</td><td>Qualtek</td><td>B01A3QKKO6</td><td>To seal the leads of the sensor cable</td></tr><tr><td>Weller WES51 Analog Soldering Station</td><td>Amazon</td><td></td><td>Lots of soldering, need a good soldering iron. https://www.amazon.com/Weller-WES51-Analog-Soldering-Station/dp/B000BRC2XU/ref=sr_1_23?s=hi&amp;amp;ie=UTF8&amp;amp;qid=1505654295&lt;br/&gt; &amp;amp;sr=1-23&amp;amp;keywords=soldering+iron</td></tr><tr><td>Rosin Soldering Flux Paste</td><td>Amazon</td><td></td><td>Needed for soldering</td></tr><tr><td>60-40 Tin Lead Rosin Core Solder Wire</td><td>Amazon</td><td></td><td>Needed for soldering</td></tr><tr><td>Aquarium sealant</td><td>Home Depot</td><td></td><td>Attach sensors to bivalve</td></tr><tr><td>PC Laptop</td><td></td><td></td><td>Any old PC to run Campbell gape program</td></tr><tr><td>heat gun</td><td>Amazon</td><td></td><td>shrink shrink tubing</td></tr><tr><td>Drill</td><td>Hardware store, Amazon</td><td></td><td>for twisting wires to make sensor cables</td></tr><tr><td>AC to DC power module</td><td>Acopian</td><td>DB15-30</td><td>Wheatstone bridge power supply</td></tr><tr><td>Poteniometer</td><td>Clarostat</td><td>733A</td><td>Wheatsone bridge nulling</td></tr><tr><td>isolating BNC connector</td><td>Sterren Electronics</td><td>&quot;200-148</td><td>Wheatstone bridge output for multimeter</td></tr><tr><td>Fused AC receptical panel module</td><td>Adam technologies</td><td>IEC-GS-1-200</td><td>Wheatstone bridge power supply connector</td></tr><tr><td>976 ohm 1% resistor</td><td>Vishay Dale</td><td>CMF50976R00FHEB</td><td>Wheatstone bridge resistor</td></tr><tr><td>1 kohm 1% resistor</td><td>Vishay Dale</td><td>CMF501K0000FHEB</td><td>Wheatstone bridge resistor</td></tr><tr><td>Potentiometer scale dial</td><td>Kilo International</td><td>462</td><td>10 turn dial for nulling potentiometer</td></tr><tr><td>DB25 male panel connector</td><td>TE connectivity</td><td>1757819-8</td><td>Data logger connector on Wheatstone bridge</td></tr><tr><td>DB25 female panel connector</td><td>TE connectivity</td><td>1757819-8</td><td>Sensor connector to Wheatstone bridge</td></tr><tr><td>perforated circuit board</td><td>Vector electronics</td><td>64P44WE</td><td>circuit board for mounting of bridge components</td></tr><tr><td>enclosure</td><td>Hammond Manufacturing</td><td>1444-29</td><td>Enclosure for sensor readout electronics</td></tr></tbody></table>

a strain gauge monitor (sgm) for continuous valve gape measurements in bivalve molluscs in response to laboratory induced diel-cycling hypoxia and ph

An inexpensive, laboratory-based, strain gauge valve gape monitor (SGM) was developed to monitor the valve gape behavior of bivalve molluscs in response to diel-cycling hypoxia. A Wheatstone bridge was connected to strain gauges that were attached to the shells of oysters (Crassostrea virginica). The recorded signals allowed for the opening and closing of the bivalves to be recorded continuously over two-day periods of experimentally-induced diel-cycling hypoxia and diel-cycling changes in pH. Here, we describe a protocol for developing an inexpensive strain gauge monitor and describe, in an example laboratory experiment, how we used it to measure the valve gape behavior of Eastern oysters (C. virginica), in response to diel-cycling hypoxia and cyclical changes in pH. Valve gape was measured on oysters subjected to cyclical severe hypoxic (0.6 mg/L) dissolved oxygen conditions with and without cyclical changes in pH, cyclical mild hypoxic (1.7 mg/L) conditions and normoxic (7.3 mg/L) conditions. We demonstrate that when oysters encounter repeated diel cycles, they rapidly close their shells in response to severe hypoxia and close with a time lag to mild hypoxia. When normoxia is restored, they rapidly open again. Oysters did not respond to cyclical pH conditions superimposed on diel cycling severe hypoxia. At reduced oxygen conditions, more than one third of the oysters closed simultaneously. We demonstrate that oysters respond to diel-cycling hypoxia, which must be considered when assessing the behavior of bivalves to dissolved oxygen. The valve SGM can be used to assess responses of bivalve molluscs to changes in dissolved oxygen or contaminants. Sealing techniques to better seal the valve gape strain gauges from sea water need further improvement to increase the longevity of the sensors.

Oysters exposed to uninterrupted normoxic estuarine water (no hypoxia during the low plateau phase of the daily cycle) were open most of the time and only briefly closed infrequently (Figure 5). When they closed varied from oyster to oyster. This pattern has also been found by Loosanoff and Nomejko 194644 and Higgins 198039. Oysters also did not respond to the difference in dark and light phases.
<p class="j...

Watch this Scientific Journal Video about A Strain Gauge Monitor SGM for Continuous Valve Gape Measurements in Bivalve Molluscs in Response to Laboratory Induced Diel-cycling Hypoxia and pH at JoVE.co

A Strain Gauge Monitor SGM for Continuous Valve Gape Measurements in Bivalve Molluscs in Response to Laboratory Induced Diel-cycling Hypoxia and pH

Understanding the behavioral responses of bivalve suspension-feeders to environmental variables, such as dissolved oxygen, can explain some ecosystem processes. We have developed an inexpensive, laboratory-based, strain gauge monitor (SGM) to measure valve gape responses of oysters, Crassostrea virginica, to diel-cycling hypoxia and cyclical pH.

A Strain Gauge Monitor (SGM) for Continuous Valve Gape Measurements in Bivalve Molluscs in Response to Laboratory Induced Diel-cycling Hypoxia and pH

An inexpensive, laboratory-based, strain gauge valve gape monitor (SGM) was developed to monitor the valve gape behavior of bivalve molluscs in response ...

a-strain-gauge-monitor-sgm-for-continuous-valve-gape-measurements

Research

JoVE Journal

Environment

8.9K Views.  University of Baltimore. Understanding the behavioral responses of bivalve suspension-feeders to environmental variables, such as dissolved oxygen, can explain some ecosystem processes. We have developed an inexpensive, laboratory-based, strain gauge monitor (SGM) to measure valve gape responses of oysters, Crassostrea virginica, to diel-cycling hypoxia and cyclical pH.

Video: A Strain Gauge Monitor SGM for Continuous Valve Gape Measurements in Bivalve Molluscs in Response to Laboratory Induced Diel-cycling Hypoxia and pH

A Simple Approach to Manipulate Dissolved Oxygen for Animal Behavior Observations

The ability to manipulate dissolved oxygen (DO) in a laboratory setting has significant application to investigate a number of ecological and organismal behavior questions. The protocol described here provides a simple, reproducible, and controlled method to manipulate DO to study behavioral response in aquatic organisms resulting from hypoxic and anoxic conditions. While performing degasification of water with nitrogen is commonly used in laboratory settings, no explicit method for ecological (aquatic) application exists in the literature, and this protocol is the first to describe a protocol to degasify water to observe organismal response. This technique and protocol were developed for direct application for aquatic macroinvertebrates; however, small fish, amphibians, and other aquatic vertebrates could be easily substituted. It allows for easy manipulation of DO levels ranging from 2 mg/L to 11 mg/L with stability for up to a 5 min animal-observation period. Beyond a 5 min observation period water temperatures began to rise, and at 10 min DO levels became too unstable to maintain. The protocol is scalable to the study organism, reproducible, and reliable, allowing for rapid implementation into introductory teaching labs and high-level research applications. The expected results of this technique should relate dissolved oxygen changes to behavioral responses of organisms.

This article describes a simple and reproducible protocol to manipulate dissolved oxygen conditions in a laboratory setting for animal behavior studies. This protocol may be used in both teaching and research laboratory settings to evaluate organismal response of macroinvertebrates, fishes, or amphibians to changes in dissolved oxygen concentration.

The ability to manipulate dissolved oxygen (DO) in a laboratory setting has significant application to investigate a number of ecological and organismal ...

Monitoring the Wall Mechanics During Stent Deployment in a Vessel

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the arterial wall lead to tissue injury, which initiates restenosis2-7. This hypothesis needs further investigations including better quantifications of non-uniform strain distribution on the artery following stent implantation. A non-contact surface strain measurement method for the stented artery is presented in this work. ARAMIS stereo optical surface strain measurement system uses two optical high speed cameras to capture the motion of each reference point, and resolve three dimensional strains over the deforming surface8,9. As a mesh stent is deployed into a latex vessel with a random contrasting pattern sprayed or drawn on its outer surface, the surface strain is recorded at every instant of the deformation. The calculated strain distributions can then be used to understand the local lesion response, validate the computational models, and formulate hypotheses for further in vivo study.

Stent-induced arterial strain distributions are characterized using an optical surface strain measurement system. This visualization technique is used to gain insights into the impact of stent implantation on the host vessel.

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the ...

Bioengineering

Bioindication Testing of Stream Environment Suitability for Young Freshwater Pearl Mussels Using In Situ Exposure Methods

Knowledge of habitat suitability for freshwater mussels is an important step in the conservation of this endangered species group. We describe a protocol for performing in situ juvenile exposure tests within oligotrophic river catchments over one-month and three-month periods. Two methods (in both modifications) are presented to evaluate the juvenile growth and survival rate. The methods and modifications differ in value for the locality bioindication and each has its benefits as well as limitations. The sandy cage method works with a large set of individuals, but only some of the individuals are measured and the results are evaluated in bulk. In the mesh cage method, the individuals are kept and measured separately, but a low individual number is evaluated. The open water exposure modification is relatively easy to apply; it shows the juvenile growth potential of sites and can also be effective for water toxicity testing. The within-bed exposure modification needs a high workload but is closer to the conditions of a natural juvenile environment and it is better for reporting the real suitability of localities. On the other hand, more replications are needed in this modification due to its high-hyporheic environment variability.

Bioindication Testing of Stream Environment Suitability for Young Freshwater Pearl Mussels Using In Situ Exposure Methods

In situ bioindications enable determination of the suitability of an environment for endangered mussel species. We describe two methods based on the juvenile exposure of freshwater pearl mussels in cages to oligotrophic river habitats. Both methods are implemented in variants for open water and hyporheic water environments.

Knowledge of habitat suitability for freshwater mussels is an important step in the conservation of this endangered species group. We describe a protocol ...

Quantitative and Temporal Control of Oxygen Microenvironment at the Single Islet Level

Simultaneous oxygenation and monitoring of glucose stimulus-secretion coupling factors in a single technique is critical for modeling pathophysiological states of islet hypoxia, especially in transplant environments. Standard hypoxic chamber techniques cannot modulate both stimulations at the same time nor provide real-time monitoring of glucose stimulus-secretion coupling factors. To address these difficulties, we applied a multilayered microfluidic technique to integrate both aqueous and gas phase modulations via a diffusion membrane. This creates a stimulation sandwich around the microscaled islets within the transparent polydimethylsiloxane (PDMS) device, enabling monitoring of the aforementioned coupling factors via fluorescence microscopy. Additionally, the gas input is controlled by a pair of microdispensers, providing quantitative, sub-minute modulations of oxygen between 0-21%. This intermittent hypoxia is applied to investigate a new phenomenon of islet preconditioning. Moreover, armed with multimodal microscopy, we were able to look at detailed calcium and KATP channel dynamics during these hypoxic events. We envision microfluidic hypoxia, especially this simultaneous dual phase technique, as a valuable tool in studying islets as well as many ex vivo tissues.

Microfluidic oxygen control confers more than just convenience and speed over hypoxic chambers for biological experiments. Especially when implemented via diffusion through a membrane, microfluidic oxygen can provide simultaneous liquid and gas phase modulations at the microscale-level. This technique enables dynamic multi-parametric experiments critical for studying islet pathophysiology.

Simultaneous oxygenation and monitoring of glucose stimulus-secretion coupling factors in a single technique is critical for modeling pathophysiological ...

Design and Use of an Apparatus for Quantifying Bivalve Suspension Feeding at Sea

As shellfish aquaculture moves from coastal embayments and estuaries to offshore locations, the need to quantify ecosystem interactions of farmed bivalves (i.e., mussels, oysters, and clams) presents new challenges. Quantitative data on the feeding behavior of suspension-feeding mollusks is necessary to determine important ecosystem interactions of offshore shellfish farms, including their carrying capacity, the competition with the zooplankton community, the availability of trophic resources at different depths, and the deposition to the benthos. The biodeposition method is used to quantify feeding variables in suspension-feeding bivalves in a natural setting and represents a more realistic proxy than laboratory experiments. This method, however, relies upon a stable platform to satisfy the requirements that water flow rates supplied to the shellfish remain constant and the bivalves are undisturbed. A flow-through device and process for using the biodeposition method to quantify the feeding of bivalve mollusks were modified from a land-based format for shipboard use by building a two-dimensional gimbal table around the device. Planimeter data reveal a minimal pitch and yaw of the chambers containing the test shellfish despite boat motion, the flow rates within the chambers remain constant, and operators are able to collect the biodeposits (feces and pseudofeces) with sufficient consistency to obtain accurate measurements of the bivalve clearance, filtration, selection, ingestion, rejection, and absorption at offshore shellfish aquaculture sites.

A flow-through device for using the biodeposition method to quantify filtration and feeding behavior of bivalve mollusks was modified for shipboard use. A two-dimensional gimbal table built around the device isolates the apparatus from boat motion, thereby allowing the accurate quantification of bivalve filtration variables at offshore shellfish aquaculture sites.

As shellfish aquaculture moves from coastal embayments and estuaries to offshore locations, the need to quantify ecosystem interactions of farmed bivalves ...

Fabrication and Implantation of Miniature Dual-element Strain Gages for Measuring In Vivo Gastrointestinal Contractions in Rodents.

Gastrointestinal dysfunction remains a major cause of morbidity and mortality. Indeed, gastrointestinal (GI) motility in health and disease remains an area of productive research with over 1,400 published animal studies in just the last 5 years. Numerous techniques have been developed for quantifying smooth muscle activity of the stomach, small intestine, and colon. In vitro and ex vivo techniques offer powerful tools for mechanistic studies of GI function, but outside the context of the integrated systems inherent to an intact organism. Typically, measuring in vivo smooth muscle contractions of the stomach has involved an anesthetized preparation coupled with the introduction of a surgically placed pressure sensor, a static pressure load such as a mildly inflated balloon or by distending the stomach with fluid under barostatically-controlled feedback. Yet many of these approaches present unique disadvantages regarding both the interpretation of results as well as applicability for in vivo use in conscious experimental animal models. The use of dual element strain gages that have been affixed to the serosal surface of the GI tract has offered numerous experimental advantages, which may continue to outweigh the disadvantages. Since these gages are not commercially available, this video presentation provides a detailed, step-by-step guide to the fabrication of the current design of these gages. The strain gage described in this protocol is a design for recording gastric motility in rats. This design has been modified for recording smooth muscle activity along the entire GI tract and requires only subtle variation in the overall fabrication. Representative data from the entire GI tract are included as well as discussion of analysis methods, data interpretation and presentation.

Fabrication and Implantation of Miniature Dual-element Strain Gages for Measuring In Vivo Gastrointestinal Contractions in Rodents.

The in vivo measurement of smooth muscle contractions along the gastrointestinal tract of laboratory animals remains a powerful, though underutilized, technique. Flexible, dual element strain gages are not commercially available and require fabrication. This protocol describes the construction of reliable, inexpensive strain gages for acute or chronic implantation in rodents.

Gastrointestinal dysfunction remains a major cause of morbidity and mortality. Indeed, gastrointestinal (GI) motility in health and disease remains an ...

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.

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 ...

The ex vivo Isolated Skeletal Microvessel Preparation for Investigation of Vascular Reactivity

The isolated microvessel preparation is an ex vivo preparation that allows for examination of the different contributions of factors that control vessel diameter, and thus, perfusion resistance1-5. This is a classic experimental preparation that was, in large measure, initially described by Uchida et al.15 several decades ago. This initial description provided the basis for the techniques that was extensively modified and enhanced, primarily in the laboratory of Dr. Brian Duling at the University of Virginia6-8, and we present a current approach in the following pages. This preparation will specifically refer to the gracilis arteriole in a rat as the microvessel of choice, but the basic preparation can readily be applied to vessels isolated from nearly any other tissue or organ across species9-13. Mechanical (i.e., dimensional) changes in the isolated microvessels can easily be evaluated in response to a broad array of physiological (e.g., hypoxia, intravascular pressure, or shear) or pharmacological challenges, and can provide insight into mechanistic elements comprising integrated responses in an intact, although ex vivo, tissue. The significance of this method is that it allows for facile manipulation of the influences on the integrated regulation of microvessel diameter, while also allowing for the control of many of the contributions from other sources, including intravascular pressure (myogenic), autonomic innervation, hemodynamic (e.g., shear stress), endothelial dependent or independent stimuli, hormonal, and parenchymal influences, to provide a partial list. Under appropriate experimental conditions and with appropriate goals, this can serve as an advantage over in vivo or in situ tissue/organ preparations, which do not readily allow for the facile control of broader systemic variables. 
The major limitation of this preparation is essentially the consequence of its strengths. By definition, the behavior of these vessels is being studied under conditions where many of the most significant contributors to the regulation of vascular resistance have been removed, including neural, humoral, metabolic, etc. As such, the investigator is cautioned to avoid over-interpretation and extrapolation of the data that are collected utilizing this preparation. The other significant area of concern with regard to this preparation is that it can be very easy to damage cellular components such as the endothelial lining or the vascular smooth muscle, such that variable source of error can be introduced. It is strongly recommended that the individual investigator utilize appropriate measurements to ensure the quality of the preparation, both at the initiation of the experiment and periodically throughout the course of a protocol.

The ex vivo Isolated Skeletal Microvessel Preparation for Investigation of Vascular Reactivity

An ex vivo preparation is described for isolation of the largest gracilis muscle resistance arterioles for interrogation of both vascular responses to vasoactive stimuli and the assessment of basic structural properties via passive wall mechanics.

The isolated  microvessel preparation is an ex vivo preparation that allows for examination of the different contributions of  factors that control vessel ...

Biology

Mechano-Node-Pore Sensing: A Rapid, Label-Free Platform for Multi-Parameter Single-Cell Viscoelastic Measurements

Cellular mechanical properties are involved in a wide variety of biological processes and diseases, ranging from stem cell differentiation to cancer metastasis. Conventional methods for measuring these properties, such as atomic force microscopy (AFM) and micropipette aspiration (MA), capture rich information, reflecting a cell's full viscoelastic response; however, these methods are limited by very low throughput. High-throughput approaches, such as real-time deformability cytometry (RT-DC), can only measure limited mechanical information, as they are often restricted to single-parameter readouts that only reflect a cell's elastic properties. In contrast to these methods, mechano-node-pore sensing (mechano-NPS) is a flexible, label-free microfluidic platform that bridges the gap in achieving multi-parameter viscoelastic measurements of a cell with moderate throughput. A direct current (DC) measurement is used to monitor cells as they transit a microfluidic channel, tracking their size and velocity before, during, and after they are forced through a narrow constriction. This information (i.e., size and velocity) is used to quantify each cell's transverse deformation, resistance to deformation, and recovery from deformation. In general, this electronics-based microfluidic platform provides multiple viscoelastic cell properties, and thus a more complete picture of a cell's mechanical state. Because it requires minimal sample preparation, utilizes a straightforward electronic measurement (in contrast to a high-speed camera), and takes advantage of standard soft lithography fabrication, the implementation of this platform is simple, accessible, and adaptable to downstream analysis. This platform's flexibility, utility, and sensitivity have provided unique mechanical information on a diverse range of cells, with the potential for many more applications in basic science and clinical diagnostics.

Presented here is a method to mechanically phenotype single cells using an electronics-based microfluidic platform called mechano-node-pore sensing (mechano-NPS). This platform maintains moderate throughput of 1-10 cells/s while measuring both the elastic and viscous biophysical properties of cells.

Cellular mechanical properties are involved in a wide variety of biological processes and diseases, ranging from stem cell differentiation to cancer ...

Physiological Characterization of the Coral Holobiont Using a New Micro-Respirometry Tool

Metabolic activity, defined as the sum of organismal processes that involve energy, is of critical importance in understanding the function and evolution of life on earth. Measuring organismal metabolic rates is, therefore, at the center of explaining the physiological states of organisms, their ecological roles, and the impact of environmental change on species within terrestrial and aquatic ecosystems. On coral reefs, measures of metabolism have been used to quantify symbiosis functioning between corals and their obligate algal symbionts (Symbiodiniaceae), as well as assess how environmental stressors, including climate change, will impact coral health. Despite this significance, there is a lack of methods, and therefore data, relating to metabolic rate measurements in coral offspring, likely due to their small size. To address this gap, this study aimed to develop a custom setup for measuring the respiration of small (millimeter size range) marine animal ecologies. This low cost and easy setup should allow for the improved measurement of metabolic rate. This will be essential for applied ecological research utilizing the sexual production of corals for reef restoration.

This protocol describes the setup and running of a micro-respirometry system that can be employed to investigate the physiological features of the coral holobiont.

Metabolic activity, defined as the sum of organismal processes that involve energy, is of critical importance in understanding the function and evolution ...

A Strain Gauge Monitor (SGM) for Continuous Valve Gape Measurements in Bivalve Molluscs in Response to Laboratory Induced Diel-cycling Hypoxia and pH

Summary

Explore More Videos

Chapters in this video

A Simple Approach to Manipulate Dissolved Oxygen for Animal Behavior Observations

Monitoring the Wall Mechanics During Stent Deployment in a Vessel

Bioindication Testing of Stream Environment Suitability for Young Freshwater Pearl Mussels Using In Situ Exposure Methods

Quantitative and Temporal Control of Oxygen Microenvironment at the Single Islet Level

Design and Use of an Apparatus for Quantifying Bivalve Suspension Feeding at Sea

Fabrication and Implantation of Miniature Dual-element Strain Gages for Measuring In Vivo Gastrointestinal Contractions in Rodents.

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

The ex vivo Isolated Skeletal Microvessel Preparation for Investigation of Vascular Reactivity

Mechano-Node-Pore Sensing: A Rapid, Label-Free Platform for Multi-Parameter Single-Cell Viscoelastic Measurements

Physiological Characterization of the Coral Holobiont Using a New Micro-Respirometry Tool