The Authors would first like to acknowledge all students from the freshman Biology 121- Ecology Module lab at Juniata College for their help in generating data used in this study. We would also like to thank Dr. Randy Bennett, Chris Walls, Sherry Isenberg, and Taylor Cox for their assistance in acquiring materials necessary to develop this methodology. Additionally, we would like to thank Dr. Norris Muth and Dr. John Unger for their advice on methodological development and Dr. Jill Keeney and the Biology department for their support of this endeavor. We would also like to thank the anonymous reviewers that have helped to shape and focus this manuscript.&#160; Last but not least, I&#39;d like to thank Hudson Grant for his help with the initial stonefly collection for use in development of this technique

Note: This experiment did not use vertebrates and therefore did not require approval by Juniata College&#39;s Institute for Animal Care and Use Committee. However for individuals adapting this method for use with vertebrates, IACUC approval should be sought.
1. Field Sample Collection
<ol>
	<li>Determine and evaluate potential field sites for the ability to collect, store, and transport stoneflies quickly to minimize time in transit with a maximum recommended time in transit of 1 hr.</li>
	<li>Perform kick-net sampling at the selected field site following standard kick-net procedures enough times to collect at least 35 stoneflies12.</li>
	<li>Collect 50 L of stream water and rocks with a maximum diameter of 2 cm from streams.</li>
	<li>Place aquariums in a refrigerator set to the temperature of the stream site. Distribute rocks collected at the stream site into aquariums and fill with 4 L of stream water per aquarium. Place 20-30 collected stoneflies per aquarium and place a bubbling stone attached to an aquarium bubbler into each tank and turn on bubblers to continuously add room air to the water.</li>
	<li>Allow the stoneflies to adjust to the new environment in the aquariums for a 48 hr period.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/54430/54430fig1.jpg" /> 
Figure 1. Set up for dissolved oxygen manipulation. (A) 1) Fitting for copper pipe to male hose barb 2) Location of stopper seal to examine for ensuring well sealing flask. (B) 1) 2 L side-arm flask filled with 1.9 L of water 2) Gas tube and air bubbler (blue) for use in nitrogen bubbling and room air bubbling, respectively 3) Nitrogen tank and gauged values 4) 2 L flask filled with 0.4 L of water with vacuum tube submerged 5) Dissolved oxygen meter. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54430/54430fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Experimental Set up
<ol>
	<li>On a bench top, connect a standard walled vacuum tube to the side arm of a 2 L side-arm flask as is shown (1 in Figure 1B).</li>
	<li>Fill the flask with 1.9 L of stream water from 3 L plastic containers holding collected stream water in refrigerator set to 12 &#176;C.</li>
	<li>Place the flask and tubing on a tray large enough to hold an ice bath around the side-arm flask without obscuring the view of the flask interior and fill the tray with ice.</li>
	<li>Drill two 3 mm diameter holes in a rubber stopper to allow the passage of 1) a copper pipe to deliver the gas to the vessel and 2) the probe of a DO meter into the 2 L side-arm flask (1 in Figure 1B).</li>
	<li>Make a lateral incision from the edge of the stopper to one of the holes to allow seating of the wire of the DO probe into the stopper.</li>
	<li>Connect a coupler with a 3 mm male hose barb to a piece of 2 mm diameter copper pipe (1 in Figure 1A). Ensure that this pipe is long enough to reach to within 10 cm of the bottom of the flask while reaching through the stopper.</li>
	<li>Place the pipe with coupler though the second hole in the stopper until the length from the bottom of the stopper is enough to reach to within 10 cm of the bottom of the flask.</li>
	<li>Connect a 0.75 m length, thin walled polyethylene gas tube with a diameter of 3 mm to the coupler on the pipe.</li>
	<li>Slide both the DO probe and copper pipe into the flask and seal the flask with the stopper.</li>
	<li>Check for a secure seal between the stopper and the flask, as well as a snug fit between the pipe and the probe wire within the stopper.</li>
	<li>Fill a 1 L flask with 0.4 L of tap water and place adjacent to the tray with the ice bath and vacuum flask.</li>
	<li>Submerge the polyethylene tube coming from the large vacuum flask into the water of the 1 L flask. Secure the tube with tape such that it will remain submerged through the experiment.</li>
	<li>Connect the 3 mm diameter gas line from the vacuum flask to an aquarium room-air bubbler. Begin to bubble the water in the 2 L flask by plugging in the aquarium bubbler, which introduces room air and oxygen to the water.</li>
	<li>Monitor the DO concentration and temperature of the water with the DO meter for 5 min or until equilibrium of DO is established within the chamber such that little change in DO is occurring.</li>
</ol>
3. Testing the Stability of the Experimental Set Up
<ol>
	<li>Test each setup for DO stability prior to the addition of stoneflies.</li>
	<li>Add three or four rocks to the 2 L flask so that stoneflies have substrate conducive for pushups.</li>
	<li>Begin a trial manipulation of DO by disconnecting the gas tube from the bubbler and attaching it to the nitrogen gas line.</li>
	<li>Start bubbling nitrogen at 20 cubic feet per hr (CFH) for approximately 40 sec to 1 min.</li>
	<li>Once the DO has dropped to within 0.5 mg/L of the target concentration, reduce the flow to 15 CFH and allow the concentration to decrease to the target.</li>
	<li>Cease flow of nitrogen immediately once the target concentration is reached.</li>
	<li>Use the aquarium room-air bubbler to return the concentration to the target concentration if the DO decreases below the target.</li>
	<li>If the DO is unstable during the testing of a set-up then check the water volume is still at 1.9 L and no water has bubbled out, water temperature is stable and not changing, and seals on all fittings appear to be tight and sealed.</li>
	<li>Once three trials have been performed and the experimenter has confidence in the ability to control DO, attach the gas line to the bubbler and bubble to equilibrium again.</li>
	<li>Bubble to equilibrium by attaching the 3 mm diameter gas line to the aquarium bubbler and starting the addition of room air to the water until the concentration of oxygen in the water does not increase or change for 3 min.</li>
	<li>Once at equilibrium, stop bubbling and unseal the flask.</li>
</ol>
4. Stonefly Push-up Experiment
<ol>
	<li>Divide the total number of stoneflies by the number of observers to determine the number of trials to perform.</li>
	<li>Determine different DO levels between 2 and 10 mg/L to evaluate the behavioral response of stoneflies (number of pushups).</li>
	<li>Set up one flask per trial and add an equal number of stoneflies as there are observers to the flask (4 stoneflies within this design), place the probe and pipe back into the flask, then reseal the flask with the rubber stopper. 
	Note: An initial DO concentration of 10 mg/L was chosen as the first observation point since it was the DO concentration of the stream from where the stoneflies were sampled.</li>
	<li>Once the water is at 10 mg/L by bubbling following steps 2.10-2.11, record the starting water temperature and allow the stoneflies to attach to the rock substrate in the flask.</li>
	<li>Assign only one observer to watch a single stonefly to ensure accurate counting of push-up behavior, which is the up and down body movement exhibited by the stonefly.</li>
	<li>Count and record the number of push-ups observed over the course of a 3 min observation period.</li>
	<li>Manipulate DO to the next experimental DO level and repeat 3 min observation period for the additional experimental levels. 
	Note: Within this experimental design, three different DO levels were evaluated.</li>
</ol>
5. Statistical Analysis
<ol>
	<li>To perform statistical analysis use average number of push-ups across the four stoneflies across a group for a given DO trial.</li>
	<li>Use the free R statistical computing software12 to perform an Analysis of Variance (ANOVA) on the number of push-ups and the DO concentrations using the order of each experimental trial (DO level) and temperature as covariates. Analyzed DO as discrete levels of a single factor.</li>
	<li>Use an Anderson-Darling normality test on residuals to check for normality13.</li>
	<li>Perform a linear regression on the data by plotting the mean number of push-ups against DO concentrations.</li>
</ol>

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The authors declare that they have no competing financial interests.

Critical steps 
This procedure provides a simple and efficient way to manipulate DO in a laboratory setting to perform behavioral studies on aquatic organisms. We found there to be several critical steps/items to be aware of when performing this experiment that directly related to the outcomes. Within a trial, it is critical to maintain the chamber pressure to avoid changes in the partial pressure of gasses above the water, and subsequent DO fluctuations. Following the steps outlined in the &#34;testing the stability of the experimental setup&#34; subsection of the protocol is critical. Inspecting the seal of the stopper with the flask, ensuring complete submersion of the vacuum tubing into the 1 L flask of water (to prevent backflow of room air), and seating the gas line and DO wire seal with the stopper can help maintain a stable chamber environment. Additionally, maximizing the volume of water in the chamber is necessary to improve stability of DO within a trial as we have found that too little volume results in erratic and unstable DO manipulations.
Maintaining water temperature in the chamber is imperative for good control of internal DO levels. While we were able to maintain cold-water temperatures within individual group experiments, some minimal variation in temperature across different trials was evident (Figure 2). Since our overall range of temperatures was low (11.8-13.5&#730;C) and within the typical range for stoneflies, it did not prove to be a significant predictor in our model for stonefly pushups. However, water temperature is known to effect the oxygen saturation potential of water14,15, and failing to maintain chamber water temperature would have direct impacts on DO levels. With a good seal, adequate volume of water, and stable water temperature, internal chamber pressure and DO is easily maintained throughout the experiment and control of DO between trials is stable and reproducible.
Limitations and Modifications 
Two potential limitations of this experiment are the size of the chamber and the length of the observation period. The volume of water (~2 L) and small opening in the neck of the chamber limits the size of organism able to be used. At this scale, the protocol would allow for easy substitution of stoneflies for other macroinvertebrates, amphibians, and smaller fishes, but would not be applicable for larger organisms (i.e., predatory fish, octopi). However, it would be possible to scale this experiment up by using larger glassware, and adapting the overall protocol to meet different learning/research objectives with larger organisms. Additionally, any substrate required in the chamber for larger organisms should be taken into account when choosing glassware due to small neck size on the 2 L flask. In our experiment small river-stones were collected with stoneflies and provided ample chamber substrate for the stoneflies to perform pushups.
Internal chamber conditions over short time observation periods were maintained with minimal fluctuations, however, uncertainties remain about longer observation periods. Utilizing the 3 min observation period outlined in the protocol, chamber water temperature and DO levels were maintained at constant values. We were able to maintain DO and water temperature up to a 5 min observation period, but at a 10 min long observation period, water temperatures in the chamber began to rise. In the current protocol, it is not feasible to extend the observation period beyond 5 min. However, adaptations to the current protocol (such as using a climate controlled room) could allow for increased long-term stability of water temperatures. Further, a more robust and detailed analysis of observation time versus ambient water conditions (temperature, DO, partial pressure) would help determine limiting factors.
This protocol has the ability to be modified (as mentioned above) to meet a number of different needs and objectives. One additional modification to the existing protocol would be a modification to the bubbling system. While we utilized solely a small copper pipe to bubble the water, we did notice that the addition of the large nitrogen gas bubbles often dislodged the stoneflies from their hold on the river stones and sent them floating around the chamber. We attempted to use a bubbling stone for more even dispersal of nitrogen gas, but found that the chamber water was not agitated enough to evenly disperse the nitrogen gas, resulting in a column of hypoxic conditions within the chamber. Further refinement of the nitrogen gas delivery system may provide useful insight, and remove the potential confound of dislodging stoneflies from substrate between DO trials.
Significance and Future Applications 
This experiment was the first of its kind to include a detailed protocol development to manipulate DO levels for animal behavior observation in a laboratory setting. While other published work suggested the use of nitrogen gas to manipulate DO levels7,8,16, insufficient methodological detail is given to allow for replication. Interest in this protocol development stemmed from our desire to manipulate DO levels and observe animal behavior for use in a college-level introductory ecology laboratory at Juniata College. Within the class of 24 students, this protocol proved reproducible across trials with differing DO levels and across groups of students. Moreover, this protocol provides a highly accessible and cost-effective way to manipulate DO levels for laboratory experimentation.
While this protocol was developed specifically for use with stoneflies in a teaching lab, it could easily be adapted for other objectives. More specifically, this protocol could easily be used with other small aquatic macroinvertebrates, fish, and even amphibians depending on the species of interest. For example members of the order Amphipodia that increase their locomotive activity in response to hypoxia17 could be used, or goldfish (Carassius auratus) that exhibit a &#34;gulping&#34; behavior at the water surface during hypoxic conditions16. Additionally, various life-stages of aquatic organisms could also be used with this protocol to help further our understanding of organismal oxygen demands throughout development. This protocol could also be utilized to study biochemical responses to hypoxia by experimenting with amphibians such as mudpuppies (Necturus maculosus)18. Further, this protocol can be scaled up or down in size to meet the needs of larger or smaller organisms and teaching or research applications. While we feel that the protocol and the specific application itself is of broad ecological interest, the greatest strength of this protocol is that it provides a great foundation development across taxonomic groups and experimental objectives.

Dissolved oxygen (DO) is a key physiochemical parameter important in mediating a number of biological and ecological processes within aquatic ecosystems. Exposures to acute and chronic sub-lethal hypoxia reduce growth rates in certain aquatic insects and reduce the survival of insects exposed1. This protocol was developed to provide a controlled method to manipulate DO levels in stream water to observe the effects on animal behavior. Since all aerobic aquatic organisms&#39; survival depends on the oxygen concentration in order to live and reproduce, changes in the concentration of DO are often reflected in behavioral changes by organisms. More mobile aquatic invertebrates and fish have been observed to respond to low oxygen concentrations (hypoxic) by seeking locales with higher DO2,3. For less mobile aquatic organisms, behavioral adaptations to increase intake of DO may be the only viable option. The aquatic macroinvertebrate order of Plecoptera (stonefly) has been noted to perform &#34;push-up&#34; movements to increase the flow of water, and uptake of oxygen, across their external gills4-6. These adaptive behaviors have been observed in natural environments and in laboratory experiments.
Laboratory manipulation of DO in water opens up significant opportunities for animal behavior studies, but significant gaps in methodological deployment exist. For example, one study used large aquaria to evaluate the physiological response time of Largemouth bass (Micropterus salmoides) to hypoxic environments following gassing with nitrogen, but scant detail is given for the methodology7. Another study performed on Zebra fish (Danio rerio) described using nitrogen gas and a porous stone to deliver gas to water and reduce the DO of the water8. For chemistry-based applications, methods for degasification of solvents utilize specialized apparatus9-11 to remove oxygen from solvents, but would not be suitable for animal behavior studies. While these studies employ methods to remove oxygen from water, no descriptive method could be identified that would allow for evaluation of animal behavior in response to DO changes.
This method described hereafter is an attempt to fully describe a protocol for manipulation of DO of water by using nitrogen gas. Further, this method was developed towards observing relationships between stonefly behavior (pushups) and DO that was employed in a freshman-level biology laboratory. One of the main benefits of this method is that it can easily be performed within a laboratory with common glassware and materials accessible to most secondary and higher education institutions. The protocol is also easily adaptable, allowing for individuals to scale the procedure to meet the objectives set forth for research or teaching applications.

<table><tbody><tr><td>Filter flask 2 L</td><td>Pyrex</td><td>5340</td><td></td></tr><tr><td>Rubber Stopper size 6</td><td>Sigma-Aldrich</td><td>Z164534</td><td></td></tr><tr><td>Nalgene 180 Clear Plastic Tubing</td><td>Thermo Scienfitic</td><td>8001-1216</td><td></td></tr><tr><td>Whisper 60 air pump</td><td>Tetra</td><td></td><td></td></tr><tr><td>Standard flexible Air line tubing</td><td>Penn Plax</td><td>ST25</td><td></td></tr><tr><td>0.25 inch Copper tubing</td><td>Lowes Home Improvement</td><td>23050</td><td></td></tr><tr><td>Male hose barb</td><td>Grainger</td><td>5LWH1</td><td></td></tr><tr><td>Female Connector</td><td>Grainger</td><td>20YZ22</td><td></td></tr><tr><td>Heavy Duty Dissolved Oxygen Meter</td><td>Extech</td><td>407510</td><td></td></tr><tr><td>Nitrogen gas</td><td>Matheson TRIGAS</td><td></td><td></td></tr><tr><td>Radnor AF150-580 Regulator</td><td>Airgas</td><td>RAD64003036</td><td></td></tr></tbody></table>

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.

Six trials of the described setup were performed by 24 freshmen undergraduate students in a teaching laboratory setting to quantify the number of push-ups stoneflies perform in response to different DO concentration in water. The average number of push-ups performed within a DO level and within each trial was pooled to plot push-ups against the DO level in Figure 2. An ANOVA was performed initially utilizing DO concentration, sequential order of trials, temperature, as well as the interactions between all variables. Results suggest that only DO concentration significantly influenced the number of push-ups performed by stoneflies (R2 adj. = 0.322, p = 0.004) and no other variable or interaction was a significant predictor of pushups. All data used in this analysis was confirmed for normality using an Anderson-Darling test.

<img alt="Figure 2" src="/files/ftp_upload/54430/54430fig2.jpg" /> 
Figure 2. Mean number of push-ups performed by stoneflies grouped by trial plotted against dissolved oxygen concentration. This shows a significant negative relationship (R2 adj. = 0.322, p =0.004) between push-ups and dissolved oxygen concentration (slope of -6.063). Red numbers indicate water temperature (in &#176;C) for a trial. The temperatures were stable across 3 min trial periods, but varied across the experiment. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54430/54430fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Code File: R code for the statistical analyses. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54430/DO_anova_Script.txt" target="_blank">Please click here to download this file.</a>

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A Simple Approach to Manipulate Dissolved Oxygen for Animal Behavior Observations

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 ...

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9.2K Views.  Juniata College. 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.

Video: A Simple Approach to Manipulate Dissolved Oxygen for Animal Behavior Observations

The Benthic Exchange of O2, N2 and Dissolved Nutrients Using Small Core Incubations

The measurement of sediment-water exchange of gases and solutes in aquatic sediments provides data valuable for understanding the role of sediments in nutrient and gas cycles. After cores with intact sediment-water interfaces are collected, they are submerged in incubation tanks and kept under aerobic conditions at in situ temperatures. To initiate a time course of overlying water chemistry, cores are sealed without bubbles using a top cap with a suspended stirrer. Time courses of 4-7 sample points are used to determine the rate of sediment water exchange. Artificial illumination simulates day-time conditions for shallow photosynthetic sediments, and in conjunction with dark incubations can provide net exchanges on a daily basis. The net measurement of N2 is made possible by sampling a time course of dissolved gas concentrations, with high precision mass spectrometric analysis of N2:Ar ratios providing a means to measure N2 concentrations. We have successfully applied this approach to lakes, reservoirs, estuaries, wetlands and storm water ponds, and with care, this approach provides valuable information on biogeochemical balances in aquatic ecosystems.

The Benthic Exchange of O2, N2 and Dissolved Nutrients Using Small Core Incubations

Here, we present small core incubations for the measurement of sediment-water gas and solute exchange. These will provide reliable measurements of sediment-water exchange that assess the role of sediment in influencing biological and biogeochemical processes in aquatic ecosystems.

The measurement of sediment-water exchange of gases and solutes in aquatic sediments provides data valuable for understanding the role of sediments in ...

Laboratory Simulation of an Iron(II)-rich Precambrian Marine Upwelling System to Explore the Growth of Photosynthetic Bacteria

A conventional concept for the deposition of some Precambrian Banded Iron Formations (BIF) proceeds on the assumption that ferrous iron [Fe(II)] upwelling from hydrothermal sources in the Precambrian ocean was oxidized by molecular oxygen [O2] produced by cyanobacteria. The oldest BIFs, deposited prior to the Great Oxidation Event (GOE) at about 2.4 billion years (Gy) ago, could have formed by direct oxidation of Fe(II) by anoxygenic photoferrotrophs under anoxic conditions. As a method for testing the geochemical and mineralogical patterns that develop under different biological scenarios, we designed a 40 cm long vertical flow-through column to simulate an anoxic Fe(II)-rich marine upwelling system representative of an ancient ocean on a lab scale. The cylinder was packed with a porous glass bead matrix to stabilize the geochemical gradients, and liquid samples for iron quantification could be taken throughout the water column. Dissolved oxygen was detected non-invasively via optodes from the outside. Results from biotic experiments that involved upwelling fluxes of Fe(II) from the bottom, a distinct light gradient from top, and cyanobacteria present in the water column, show clear evidence for the formation of Fe(III) mineral precipitates and development of a chemocline between Fe(II) and O2. This column allows us to test hypotheses for the formation of the BIFs by culturing cyanobacteria (and in the future photoferrotrophs) under simulated marine Precambrian conditions. Furthermore we hypothesize that our column concept allows for the simulation of various chemical and physical environments &#8212; including shallow marine or lacustrine sediments.

Laboratory Simulation of an IronII-rich Precambrian Marine Upwelling System to Explore the Growth of Photosynthetic Bacteria

We simulated a Precambrian ferruginous marine upwelling system in a lab-scale vertical flow-through column. The goal was to understand how geochemical profiles of O2 and Fe(II) evolve as cyanobacteria produce O2. The results show the establishment of a chemocline due to Fe(II) oxidation by photosynthetically produced O2.

A conventional concept for the deposition of some Precambrian Banded Iron Formations (BIF) proceeds on the assumption that ferrous iron [Fe(II)] upwelling ...

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.

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.

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

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 ...

Bioengineering

Anaerobic Protein Purification and Kinetic Analysis via Oxygen Electrode for Studying DesB Dioxygenase Activity and Inhibition

Oxygen-sensitive proteins, including those enzymes which utilize oxygen as a substrate, can have reduced stability when purified using traditional aerobic purification methods. This manuscript illustrates the technical details involved in the anaerobic purification process, including the preparation of buffers and reagents, the methods for column chromatography in a glove box, and the desalting of the protein prior to kinetics. Also described are the methods for preparing and using an oxygen electrode to perform kinetic characterization of an oxygen-utilizing enzyme. These methods are illustrated using the dioxygenase enzyme DesB, a gallate dioxygenase from the bacterium Sphingobium sp. strain SYK-6.

Here we present a protocol for anaerobic protein purification, anaerobic protein concentration, and subsequent kinetic characterization using an oxygen electrode system. The method is illustrated using the enzyme DesB, a dioxygenase enzyme which is more stable and active when purified and stored in an anaerobic environment.

Oxygen-sensitive proteins, including those enzymes which utilize oxygen as a substrate, can have reduced stability when purified using traditional aerobic ...

Biochemistry

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

Developmental Biology

A Low-Cost Method of Measuring the In Situ Primary Productivity of Periphyton Communities of Lentic Waters

Measuring the in situ primary productivity of periphyton during the growing season gradient can elucidate the quantitative effect of environmental drivers (mainly phosphorus concentration and light intensity) and species composition on primary productivity. Primary productivity is mainly driven by light intensity, temperature, availability of nutrients, and distribution of the ionic species of the carbonate system in the respective depths of the euphotic zone. It is a complex system that is very difficult to simulate in the laboratory. This cheap, transportable, and easy-to-build floating barge allows measuring the primary productivity accurately-directly under the actual natural conditions. The methodology is based on measuring the primary productivity in real time using noninvasive oxygen sensors integrated into tightly sealed glass jars, enabling online oxygen flux monitoring and providing new insights into metabolic activities. Detailed seasonal in situ measurements of gross primary productivity of microbial mats (or other benthic organisms) can improve current knowledge of the processes controlling primary productivity dynamics in lentic waters.

A Low-Cost Method of Measuring the In Situ Primary Productivity of Periphyton Communities of Lentic Waters

Presented here is a cost-effective and transportable method/facility for measuring the primary productivity of microbial mats under actual in situ environmental temperature and light conditions. The experimental setup is based on widely available materials and can be used under various conditions while offering the advantages of laboratory-based models.

Measuring the in situ primary productivity of periphyton during the growing season gradient can elucidate the quantitative effect of environmental drivers ...

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, Drosophila research lacks popularity in developing countries, possibly due to the misinformed idea that establishing a lab and performing relevant experiments with such tiny insects is difficult and requires expensive, specialized apparatuses. Here, we describe how to build an affordable flylab to quantitatively analyze a myriad of behavioral parameters in D. melanogaster, by 3D-printing many of the necessary pieces of equipment. We provide protocols to build in-house vial racks, courtship arenas, apparatuses for locomotor assays, etc., to be used for general fly maintenance and to perform behavioral experiments using adult flies and larvae. We also provide protocols on how to use more sophisticated systems, such as a high resolution oxygraph, to measure mitochondrial oxygen consumption in larval samples, and show its association with behavioral changes in the larvae upon the xenotopic expression of the mitochondrial alternative oxidase (AOX). AOX increases larval activity and mitochondrial leak respiration, and accelerates development at low temperatures, which is consistent with a thermogenic role for the enzyme. We hope these protocols will inspire researchers, especially from developing countries, to use Drosophila to easily combine behavior and mitochondrial metabolism data, which may lead to information on genes and/or environmental conditions that may also regulate human physiology and disease states.

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

We provide protocols for anyone with a "maker culture" mind to start building a flylab for quantitative analysis of a myriad of behavioral parameters in Drosophila melanogaster, by 3D-printing many of the necessary pieces of equipment. We also describe a high resolution respirometry protocol using larvae to combine behavioral and mitochondrial metabolism data.

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, ...

Behavior

Creating Defined Gaseous Environments to Study the Effects of Hypoxia on C. elegans

Oxygen is essential for all metazoans to survive, with one known exception1. Decreased O2 availability (hypoxia) can arise during states of disease, normal development or changes in environmental conditions2-5. Understanding the cellular signaling pathways that are involved in the response to hypoxia could provide new insight into treatment strategies for diverse human pathologies, from stroke to cancer. This goal has been impeded, at least in part, by technical difficulties associated with controlled hypoxic exposure in genetically amenable model organisms.
The nematode Caenorhabditis elegans is ideally suited as a model organism for the study of hypoxic response, as it is easy to culture and genetically manipulate. Moreover, it is possible to study cellular responses to specific hypoxic O2 concentrations without confounding effects since C. elegans obtain O2 (and other gasses) by diffusion, as opposed to a facilitated respiratory system6. Factors known to be involved in the response to hypoxia are conserved in C. elegans. The actual response to hypoxia depends on the specific concentration of O2 that is available. In C. elegans, exposure to moderate hypoxia elicits a transcriptional response mediated largely by hif-1, the highly-conserved hypoxia-inducible transcription factor6-9. C .elegans embryos require hif-1 to survive in 5,000-20,000 ppm O27,10. Hypoxia is a general term for "less than normal O2". Normoxia (normal O2) can also be difficult to define. We generally consider room air, which is 210,000 ppm O2 to be normoxia. However, it has been shown that C. elegans has a behavioral preference for O2 concentrations from 5-12% (50,000-120,000 ppm O2)11. In larvae and adults, hif-1 acts to prevent hypoxia-induced diapause in 5,000 ppm O212. However, hif-1 does not play a role in the response to lower concentrations of O2 (anoxia, operational definition &lt;10 ppm O2)13. In anoxia, C. elegans enters into a reversible state of suspended animation in which all microscopically observable activity ceases10. The fact that different physiological responses occur in different conditions highlights the importance of having experimental control over the hypoxic concentration of O2.
Here, we present a method for the construction and implementation of environmental chambers that produce reliable and reproducible hypoxic conditions with defined concentrations of O2. The continual flow method ensures rapid equilibration of the chamber and increases the stability of the system. Additionally, the transparency and accessibility of the chambers allow for direct visualization of animals being exposed to hypoxia. We further demonstrate an effective method of harvesting C. elegans samples rapidly after exposure to hypoxia, which is necessary to observe many of the rapidly-reversed changes that occur in hypoxia10,14. This method provides a basic foundation that can be easily modified for individual laboratory needs, including different model systems and a variety of gasses.

Creating Defined Gaseous Environments to Study the Effects of Hypoxia on C. elegans

This paper details how to use continuous-flow hypoxia chambers to generate atmospheres with defined concentrations of O2 to understand biological responses to decreased O2. This system is easy to setup and maintain, and flexible enough to suit a wide range of O2 concentrations and model systems

Oxygen is essential for all metazoans to survive, with one known exception1. Decreased O2 availability (hypoxia) can arise during states of disease, ...

Biology

Adapting Microrespirometry for Drosophila - Insights into CASK Mutant Metabolism

Measuring O2 Consumption in Drosophila melanogaster Using Coulometric Microrespirometry

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable environment. A coulometric microrespirometer consists of an airtight chamber in which O2 is consumed and the CO2 produced by the organism is removed by an absorbent medium. The resulting pressure decrease triggers electrolytic O2 production, and the amount of O2 produced is measured by recording the amount of charge used to generate it. In the present study, the method has been adapted to Drosophila melanogaster tested in small groups, with the sensitivity of the apparatus and the environmental conditions optimized for high stability. The amount of O2 consumed by wildtype flies in this apparatus is consistent with that measured by previous studies. Mass-specific O2 consumption by CASK mutants, which are smaller and known to be less active, was not different from congenic controls. However, the small size of CASK mutants resulted in a significant reduction in O2 consumption on a per-fly basis. Therefore, the microrespirometer is capable of measuring O2 consumption in D. melanogaster, can distinguish modest differences between genotypes, and adds a versatile tool for measuring metabolic rates.

Author Spotlight: Influence of Temperature on Drosophila melanogaster and Desert-Adapted Beetles 

Coulometric respirometry is ideal for measuring the metabolic rate of small organisms. When adapted for Drosophila melanogaster in the present study, measured O2 consumption was within the range reported for wildtype D. melanogaster by previous studies. Per-fly O2 consumption by CASK mutants, which are smaller and less active, was significantly lower than the wildtype.

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable ...

A Simple Approach to Manipulate Dissolved Oxygen for Animal Behavior Observations

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The Benthic Exchange of O₂, N₂ and Dissolved Nutrients Using Small Core Incubations

Laboratory Simulation of an Iron(II)-rich Precambrian Marine Upwelling System to Explore the Growth of Photosynthetic Bacteria

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

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

Anaerobic Protein Purification and Kinetic Analysis via Oxygen Electrode for Studying DesB Dioxygenase Activity and Inhibition

Induction of Hypoxia in Living Frog and Zebrafish Embryos

A Low-Cost Method of Measuring the In Situ Primary Productivity of Periphyton Communities of Lentic Waters

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

Creating Defined Gaseous Environments to Study the Effects of Hypoxia on C. elegans

Measuring O₂ Consumption in Drosophila melanogaster Using Coulometric Microrespirometry