Special thanks to the Plant Growth Facility and EcoCore analytical facility at Colorado State University and to the many students and teachers who helped on this project. This work is funded by the NSF DEB grant #0918482, USDA National Needs Fellowship, and the Cotrufo-Hoppess fund for soil ecology research.

1. Chamber Construction
<ol>
	<li>Construct the labeling chamber in a greenhouse to allow for maximum natural light potential for plant growth. Make sure that adequate power supply is available to power all chamber components.</li>
	<li>Construct the labeling chamber by mounting 3.175 mm thick transparent acrylic walls (polycarbonate would also be suitable) and a 6.35 mm thick transparent acrylic ceiling on an aluminum frame with a white-painted steel floor to maximize solar reflectance. The dimensions of the chamber can be tailored to suit individual research needs.</li>
	<li>Mount the chamber on &#190; in (19 mm) plywood on cinder blocks.</li>
	<li>Drill holes in the acrylic glass, aluminum frame and steel floor and use screws to fasten all components together.</li>
	<li>Seal all seams with silicone caulk to insure an airtight seal.</li>
	<li>Construct a door by mounting one section of the acrylic paneling on long screws, which can be screwed down using removable wing nuts.</li>
	<li>Seal the door with weather stripping to prevent air leakage.</li>
	<li>Select an area directly adjacent to the chamber as the control center to mount all temperature, humidity, CO2 controls, and monitoring equipment.</li>
	<li>Carefully drill small holes in the chamber wall adjacent to the control center for all electrical wires and gas tubing. Use silicone caulk to seal the holes around all wires and tubing to prevent air leakage.</li>
	<li>Test the chamber for air leakage by filling it with a high level of CO2 (e.g.&#160;800 ppm) and letting it sit overnight. If the CO2 concentration in the chamber is maintained at its original level then it is airtight. If the concentration drops overnight then all seams should be examined and resealed with silicone caulk until an airtight seal is achieved.</li>
	<li>For some plants adapted to high light conditions, add lights connected to a timer to the immediate exterior of the chamber to increase light penetration and plant productivity.</li>
</ol>
2. Temperature and Humidity Controls
<ol>
	<li>Regulate chamber temperature by installing a commercial split type air conditioner with the cooling (evaporator) coils located inside the chamber and the compressor and condenser coils located outside the greenhouse to dissipate the heat. Set the air conditioner to maintain the desired temperature.</li>
	<li>Use a small room dehumidifier to control humidity in the chamber.
	<ol>
		<li>Drill a drainage hole through the floor of the chamber adjacent to the dehumidifier.</li>
		<li>Remove the condensate collector from the dehumidifier, and connect a drainage tube from the dehumidifier through the drainage hole in the floor of the chamber.</li>
		<li>Underneath the chamber, place an open jar filled with water for the dehumidifier to drain directly into. This creates an airtight seal but also allow for pressure equilibration.</li>
	</ol>
	</li>
	<li>Install a controller, such as the Omega iSeries controller, in the control center with a humidity sensor inside the chamber.
	<ol>
		<li>Connect the controller to the dehumidifying system with a solid-state relay and set the humidity controller with a high alarm and swing to maintain optimal growing conditions in the chamber. Optimal temperature and humidity conditions will differ for different plant species.</li>
	</ol>
	</li>
</ol>
3. CO2&#160;Controls
<ol>
	<li>The&#160;13C-CO2 enrichment is achieved using two pure CO2 gas tanks, one of 10 atom%13C-CO2 or higher and one of 1.1 atom%13C-CO2 (natural abundance).</li>
	<li>Monitor CO2 concentration by having a diaphragm pump continuously draw chamber air through an Infrared Gas Analyzer (IRGA) and then pump the air back into the chamber, thus maintaining a closed system (Figure 1).</li>
	<li>Set a low alarm and dead band on the IRGA software to maintain CO2 concentrations within a desired range. Here, we use a low alarm of 360 ppm with a 40 ppm dead band to maintain CO2 concentrations between 360-400 ppm.</li>
	<li>Connect a metering valve to each tank and carefully adjust them to achieve the target&#160;13C enrichment level. Set the tank regulators to 20 psi.</li>
	<li>Insert a solenoid valve between the metering valve and the regulator of each tank. Join the outlets of the two metering valves together and pipe them into the center of the chamber. Wire a solid-state relay to the IRGA output to control the solenoid valves (Figure 1).</li>
</ol>
4. Web-based Remote Monitoring System
<ol>
	<li>Monitor CO2 concentrations from the IRGA software by logging it to a local file once every 30 sec.</li>
	<li>Create a custom utility (e.g.&#160;in perl or another programming language) to pick up entries from the local CO2 logging file, along with the current laptop timestamp, and upload them to a back-end web application.</li>
	<li>Set the web application to query the temperature and humidity sensor data.</li>
	<li>Use a monitoring system to check the status of the temperature in the chamber every five minutes to prevent potential temperature spikes that would destroy the plants if the air conditioning system failed.</li>
	<li>Monitor the CO2, temperature and humidity data on any standard web browser so that any unexpected temperature spikes or CO2 drops can be immediately attended to.</li>
</ol>
5. Irrigation System
<ol>
	<li>Drill one small hole in the wall of the acrylic glass chamber per pot.</li>
	<li>Use irrigation tubing to create one drip irrigation ring per pot and feed the irrigation tubing through the chamber wall to the exterior.</li>
	<li>Seal the holes around the irrigation tubing with silicone caulk to prevent air leakage.</li>
	<li>On the exterior of the chamber, connect the irrigation tubing to the tubing for a peristaltic pump.</li>
	<li>Use a small hose clamp to close off all irrigation tubing to prevent air leakage between watering.</li>
</ol>
6. Potting Plants
<ol>
	<li>Select a pot size appropriate for the plants being grown. Here, 40, 15 L pots are used.</li>
	<li>Create a soil-free potting mix by mixing sand, vermiculite, and profile porous ceramic.</li>
	<li>Test the water holding capacity of the potting mix by weighing a filled pot dry, soaking the pot with water and allowing it to drain completely, and weighing the pot wet. Use this maximum water holding capacity to ensure that excess labeled fertilizer and water does not leak from the pots during watering.</li>
	<li>Germinate seeds in potting soil prior to planting them in the pots. This ensures that only successfully germinated seeds are started in the labeling chamber.</li>
	<li>Inoculate the seedlings with fresh soil slurry to introduce beneficial microbes.</li>
	<li>Once the seeds have germinated, carefully transplant seedlings to the pots&#160;in the desired number.</li>
	<li>Once potted, move the pots into the chamber and assemble each pot with an individual irrigation hose.</li>
</ol>
7. Sealing the Chamber
<ol>
	<li>When first sealing the door to the chamber a large mass of external air fills the chamber space. Scrub out this external CO2 by connecting a soda lime scrubber to the air pump to scrub the CO2 concentration down to at least 200-250 ppm before filling the chamber back up to 400 ppm using the&#160;13C-CO2 tank mixture.</li>
	<li>Try to keep the chamber closed through the duration of the growing season to minimize natural abundance CO2 contamination.</li>
	<li>Monitor plant growth visually and adjust fertilization and irrigation according to demand.</li>
</ol>
8. Fertilization and Irrigation
<ol>
	<li>Use a fertilizer solution, such as a modified Hoagland&#39;s solution23, to fertilize the plants through the irrigation system.</li>
	<li>Label the fertilizer with&#160;15N by using a&#160;15N-KNO3 subsolution at the targeted atom%15N level by mixing 98 atom%15N-KNO3with natural abundance&#160;15N-KNO3 (0.37 atom%15N).</li>
	<li>Mix up enough of the fertilizer solution upon each fertilization event for the entire chamber, based on the water holding capacity of the potting mix. Place the proper amount of fertilizer for one pot in a glass jar, and prepare as many jars there are pots.</li>
	<li>Unclamp the irrigation hoses and place each of them in a jar with the fertilizer solution, and then connect them to the peristaltic pump.</li>
	<li>Water plants by pumping water through the peristaltic pump through the individual drip irrigation hoses regularly as the plants need it.</li>
	<li>Fertilization with Hoagland&#39;s solution should follow the plant demand or experimental design, with increasing nutrient demand as the plant productivity increases to maximize productivity.</li>
	<li>First, pump the Hoagland&#39;s solution through the irrigation hose, then pump a water rinse through to minimize algal and bacterial growth in the tubes.</li>
	<li>Reclamp all hoses after fertilization and irrigation to eliminate chamber air leakage.</li>
</ol>
9. Uniform and Differential Labeling
<ol>
	<li>For differential labeling of structural and metabolic components, remove plants from labeling chamber 1-3 weeks prior to harvest. Plants that are to be uniformly labeled can remain in the sealed labeling chamber continuously until harvest and irrigated with&#160;15N-Hoagland&#39;s solution.</li>
	<li>Keep the removed plants in the greenhouse during this time so that they receive adequate light and CO2 at natural abundance&#160;13C.</li>
	<li>For differential&#160;15N labeling, continue to fertilize and irrigate the plants as usual, but use natural abundance&#160;15N-KNO3 in the Hoagland&#39;s fertilizer solution.</li>
</ol>
10. Harvest
<ol>
	<li>Stop watering plants 1 week prior to harvest so plants begin to senesce and potting medium dries out.</li>
	<li>Open the chamber and move the pots out for immediate clipping and harvest of the aboveground biomass</li>
	<li>Pour out the potting mix and roots over a coarse screen.</li>
	<li>Use the screen to separate out the roots from the potting mix and shake the roots free of potting mix.</li>
	<li>Place the roots on a 2 mm sieve and rinse them with water to remove any remaining potting material. Use tweezers to remove any vermiculite that may cling to the roots.</li>
	<li>Allow roots to air-dry in preparation for future experiments.</li>
</ol>
11. Litter Chemistry
<ol>
	<li>Weigh air-dry plant material to determine labeling chamber biomass.</li>
	<li>Grind a subsample for chemical analysis.</li>
	<li>Place 2.0 g of oven dried (60 &#176;C) litter in a 125 ml&#160;acid washed flask and add 50 ml&#160;of deionized water.</li>
	<li>Place the sample on a preheated (60 &#176;C) stirrer plate and place a stirrer bar in the flask. Set the stirring for 200 rpm and allow the sample to heat for 30 min.</li>
	<li>After 30 min, filter the litter solution through a 20 &#956;m Nylon mesh on a vacuum filtration system.</li>
	<li>Transfer the extract to a preweighed acid washed tube and freeze.</li>
	<li>Transfer the solid litter residue to a preweighed aluminum pan and dry at 60 &#176;C. Weigh pan and litter after drying to determine hot water residue mass.</li>
	<li>Freeze dry the hot water extract and weigh to determine the hot-water extract mass.</li>
	<li>Analyze oven-dried (60 &#176;C) litter, freeze-dried hot water extract, and oven-dried hot water residue in an Elemental Analyzer - Isotope Ratio Mass Spectrometer (EA-IRMS).</li>
</ol>

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The authors have nothing to disclose.

This design for a continuous isotope labeling chamber was used to produce uniformly and differentially&#160;13C and&#160;15N labeled A. gerardii for subsequent field and laboratory experiments. During three growing seasons of operation, the chamber has successfully maintained temperature, humidity, and CO2 concentrations within the set parameters (Figure 2). The reliability of the temperature control system is critical during the peak of the summer when high solar radiation can cause overheating in the air-tight chamber. Eliminating excess humidity caused by transpiration from the growing plants ensures that plant stomata remain open for photosynthetic uptake22 and that water condensation does not inhibit light penetration or damage the structure of the chamber.
The near continuous monitoring of CO2 concentration by the IRGA software maintained continuous&#160;13C labeling of the plants while they were growing in the chamber. Due to the high photosynthetic activity of A. gerardii growing in this chamber, CO2 was injected into the system frequently during daylight hours of the peak growing season when photosynthetic activity drew CO2 concentrations down to 360 ppm, approximately every 15-20 min. The metering of the enriched and natural abundance&#160;13C-CO2 tanks allowed for a controlled 4.4 atom%13C atmosphere through the growing season for uniform plant tissue labeling.&#160;13C-CO2 production can also be achieved by mixing&#160;13C-sodium bicarbonate or&#160;13C-sodium carbonate and hydrochloric acid, however this type of system is more complicated and requires more monitoring and maintenance, so we recommend using&#160;13C-CO2 gas. An important consideration for monitoring CO2 concentrations using an IRGA is that infrared analyzers lose two thirds of their sensitivity when measuring&#160;13C-CO2. This underestimation of approximately 2.9% ppm for our 4.4%13C-CO2 mixture was not of great concern to us, but could become a more significant issue when labeling at higher&#160;13C levels27.
A. gerardii is a warm season perennial tallgrass prairie graminoid species. The design of this chamber was optimized for A. gerardii production (Figure 1). The size and height of the chamber were chosen in consideration for the maximum height productivity of A. gerardii in the field, as well as for the desired plant biomass production for future experiments. A. gerardii is known to be light limited in the field24,25. PAR within a greenhouse can be diminished by 30-47% as compared to exterior levels26. Since our plants were grown in an acrylic glass chamber inside a greenhouse, PAR limitation was a concern. When turned on, the fluorescent lights increased PAR within the chamber by 9.5%, which may have helped to increase productivity in this light sensitive species. When using this chamber design to grow other types of plants specific physiological needs such as size, lighting, nutrient demands, temperature sensitivity, and soil moisture should be carefully tailored to maintain optimal growing conditions for the plants.
When working with expensive isotopically labeled compounds, such as 10 atom%13C-CO2 and 98 atom%15N-KNO3, efficiency of labeling is an important consideration. This chamber design optimizes&#160;13C labeling by making all efforts to seal the chamber and minimize air leakage into and out of the chamber. If this chamber is never opened during the growing season, then none of the&#160;13C labeled CO2 from the chamber is leaked out to the atmosphere. The CO2 build up during nighttime respiration does not appear to damage the growing plants and is quickly taken up after sunrise (Figure 2). During differential labeling, the chamber was briefly opened to remove the differentially labeled pots but this did not appear to dilute the targeted 4.4 atom%13C labeling of the continuously labeled plants (Table 1).&#160;13C labeling was also optimized by scrubbing out the initial atmospheric air trapped in the chamber upon sealing. During preliminary tests on the chamber without an initial scrub of atmospheric CO2, plants were measured to have a diluted&#160;13C level in the first leaves produced than in the later leaves produced. The initial scrub of atmospheric CO2 upon chamber closure appears to eliminate this issue by allowing for continuous&#160;13C labeling from seedling to maturity. Maintaining a soil-free potting mixture of sand, vermiculite, and clay also eliminates natural abundance CO2 contamination from soil respiration. The elimination of soil from the system does require careful fertilization and inoculation considerations, which may be unique to different plant species.&#160;15N labeling through the targeted 7 atom%15N Hoegland&#39;s solution produced highly labeled plant material at 6.7 atom%15N (Table 1). A slight dilution from the targeted&#160;15N label may be caused by some natural abundance N in the potting mix or from the native soil inoculation.
During biosynthesis of compounds, natural discrimination of&#160;13C (or&#160;15N) occurs as a result of kinetic fractionation and nonstatistical isotope distribution in the synthesized compounds. Thus, in the case of C, secondary products (e.g.&#160;lipids, phenol compounds) are generally depleted in&#160;13C as compared to primary products (carbohydrates). This natural&#160;13C discrimination appears to persist also when plants are grown in an enriched&#160;13C atmosphere, as can be seen in the slight difference in atom%13C of the hot water extracts and hot water residues of the uniformly labeled plants (Table 1). This natural kinetic fractionation is very small compared to the enrichment and does not compromise the uniformity of the labeling.
Differential labeling of structural and metabolic plant tissues is a novel technique with potential for advanced studies in litter decomposition, microbial ecology and soil organic matter formation. The difference in&#160;13C and&#160;15N of the hot water extracts and hot water residues indicates a significant dilution of&#160;13C and&#160;15N in the leachable, low molecular weight compounds (hot water extracts) from the structural plant material (hot water residues) after 7, 14, and 22 days of differential labeling (P&#60;0.005). This differential labeling of plant tissues can be used to track the fate of structural and metabolic components separately through an ecosystem. The 13C differential labeling was more extreme than&#160;15N differential labeling. This may be due to the immediacy of&#160;13C dilution when removing the plants from the&#160;13C-CO2 labeled atmosphere, while the&#160;15N label still remains in the potting mix for some time and&#160;15N dilution occurs more slowly. For more drastic differential labeling of&#160;15N, one may consider flushing the pots with water prior to natural abundance fertilization during the final weeks of growth outside the chamber.
The design and operation of this continuous&#160;13C and&#160;15N labeling system for uniform or differential, metabolic and structural, plant tissue labeling provides a novel method for producing isotopically labeled plant material for advanced research. The design and operational details of this chamber have been chosen for 4.4 atom%13C and 7 atom%15N labeling of A. gerardii, but can be tailored to other plant types and isotope labeling levels. The growing conditions described here should be tailored to suit the size, temperature, humidity, light, water and nutrient demands of the particular plant species of interest. Labeling with&#160;18O or&#160;2H can also be achieved by labeling the water used in the irrigation system. The system described here addresses many of the challenges of uniform and differential&#160;13C labeling of plant material. This basic chamber design can be used by other research groups to produce highly labeled plant material for advanced studies in ecosystem biogeochemistry.

Understanding the dynamics of plant-soil-atmosphere processes is critical for accurately predicting how the global carbon (C) and nitrogen (N) cycles function under current and future environmental conditions. Stable isotopes are powerful tools in quantitative studies of plant-soil-atmosphere C and N cycling. Tracing rare stable isotopes from plant material through the ecosystem provides sensitive information in studies of biogeochemical cycling, from CO2 fluxes&#160;and soil organic matter formation to small-scale stable-isotope biomarker probing&#160;e.g.4,8,9. Combining&#160;13C labeling with&#160;15N labeling, or other stable isotopes such as&#160;2H or&#160;18O in plant tissue provides a high-detection, traceable, yet complex substrate for use in coupled studies of plant and soil biochemistry. The ability to uniformly or differentially label structural and metabolic plant material adds further ability to address complex questions about C and N cycling through ecosystems. The benefit of using isotope labeled plant material in quantitative studies of C and N accounting, however, depends on the ability to produce 13C and 15N labeled material that is either uniformly or differentially labeled.
Isotope labeling has been used in studies addressing plant C and N assimilation10, allocation11 and rhizodeposition12. Uniformly 13C and 15N labeled plant material provides a complex labeled substrate for studies of litter decomposition1,4, soil organic matter formation2,6, soil CO2 emissions4, soil food web studies13, and studies of soil C residence times2,14. Studies utilizing&#160;13C labeled biochar from labeled plant material are also beginning to reveal new information about formerly overlooked soil char pools15. While 15N, 2H, and 18O labeling are relatively easy to achieve through water and fertilizer treatment, the challenge exists in producing uniformly 13C labeled plant material through 13C-CO2 fixation.
Continuous isotope labeling from seedling to maturity in a sealed chamber produces uniform isotope labeling throughout the plant. Other methods such as repeated pulse labeling16 and foliar application or wicking17,18 do not produce uniformly isotope labeled plant material, nor clear differential labeling of specific C-compounds (e.g.&#160;metabolic vs. structural)19. An important consideration in isotope labeling is labeling efficiency, due to the high cost of rare isotope enriched compounds used in labeling. Although continuous 13C labeling has been used in the past2-4,20, there is not to our knowledge a published detailed technical description of a continuous labeling chamber with evidence of high labeling efficiency and accurate control of the amount and uniformity of isotope labeling.
On the forefront of litter decomposition and soil organic matter formation research is the concept that metabolic plant material (i.e. leachable, labile, low molecular weight compounds) and structural plant material (i.e. lignin, cellulose, hemicellulose) are processed differently in terms of microbial use efficiency, soil organic matter formation, and long term soil C storage5-7. Plant material that is differentially labeled in its structural and metabolic components, therefore, is a useful tool in advancing litter decomposition and soil organic matter formation research. Differential labeling with dual isotopes allows for the tracing of structural and metabolic components separately through the ecosystem using a multiple-pool isotope technique21.
Continuous isotope labeling with 13C and other isotopes in a sealed chamber requires careful attention to plant physiological conditions to maximize plant productivity and isotope labeling efficiency. Daytime temperature spikes must be controlled to prevent plant damage when growing in an airtight chamber. An optimal range of humidity and temperature are required to maintain open plant stomata and CO2 uptake22. High levels of humidity cause fogging of the chamber walls, which minimizes light availability and may damage the chamber structure. Careful consideration to isotope labeling efficiency by eliminating natural abundance isotopes from the chamber (e.g.&#160;coming from potting with soil organic matter) and preventing exposure to external air is important when working with expensive heavy-isotope labeled compounds.
Here, we present a method for building and operating a continuous dual 13C and 15N isotope labeling chamber for the production of plant material that is either uniformly labeled or has its structural and metabolic components labeled at distinct levels. 13C labeling is controlled at the chamber level, while fertilization and 15N labeling is controlled at the individual pot level. Representative results are shown to demonstrate the ability of this method to control temperature, humidity, and CO2 concentration throughout the growing season. Results from growing Andropogon gerardii,&#160;Kaw also demonstrate this method&#39;s ability to produce uniformly or differentially labeled plant material. The specific chamber design and operation scheme described can be modified to grow different plant species, as well as to accommodate 2H or&#160;18O labeling.

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design and operation of a continuous 13c and 15n labeling chamber for uniform or differential, metabolic and structural, plant isotope labeling

Tracing rare stable isotopes from plant material through the ecosystem provides the most sensitive information about ecosystem processes; from CO2 fluxes and soil organic matter formation to small-scale stable-isotope biomarker probing. Coupling multiple stable isotopes such as 13C with 15N, 18O or&#160;2H has the potential to reveal even more information about complex stoichiometric relationships during biogeochemical transformations. Isotope labeled plant material has been used in various studies of litter decomposition and soil organic matter formation1-4. From these and other studies, however, it has become apparent that structural components of plant material behave differently than metabolic components (i.e. leachable low molecular weight compounds) in terms of microbial utilization and long-term carbon storage5-7. The ability to study structural and metabolic components separately provides a powerful new tool for advancing the forefront of ecosystem biogeochemical studies. Here we describe a method for producing 13C and 15N labeled plant material that is either uniformly labeled throughout the plant or differentially labeled in structural and metabolic plant components.
Here, we present the construction and operation of a continuous 13C and 15N labeling chamber that can be modified to meet various research needs. Uniformly labeled plant material is produced by continuous labeling from seedling to harvest, while differential labeling is achieved by removing the growing plants from the chamber weeks prior to harvest. Representative results from growing Andropogon gerardii Kaw demonstrate the system&#39;s ability to efficiently label plant material at the targeted levels. Through this method we have produced plant material with a 4.4 atom%13C and 6.7 atom%15N uniform plant label, or material that is differentially labeled by up to 1.29 atom%13C and 0.56 atom%15N in its metabolic and structural components (hot water extractable and hot water residual components, respectively). Challenges lie in maintaining proper temperature, humidity, CO2 concentration,&#160;and light levels in an airtight 13C-CO2 atmosphere for successful plant production. This chamber description represents a useful research tool to effectively produce uniformly or differentially multi-isotope labeled plant material for use in experiments on ecosystem biogeochemical cycling.

Our labeling chamber is 1.2 m&#160;x 2.4 m x 3.6 m in size and holds 40,15 L pots (Figure 1). The computerized IRGA control system maintained CO2 concentrations between our set values of 360 and 400 ppm during the photosynthetically active period of the day (Figure 2a). The low CO2 alarm feature on the IRGA triggered solenoid valves to allow CO2 from the&#160;13C enriched and natural abundance tanks into the chamber when the concentration dropped below the minimum threshold (e.g.&#160;360 ppm). The dead band feature stopped the flow when the concentration reached the upper set point (e.g.&#160;400 ppm). The iSeries temperature and humidity monitoring system connected to the air conditioner and dehumidifier held climate conditions within the set parameters throughout the growing season (Figure 2b). We used a one-ton (3.5 kW) air conditioning unit to keep the chamber cool.
The remote monitoring system allowed the logged data to be viewed at any time by a standard web browser. The CO2 concentrations, temperature and humidity values were down sampled by the web application to display graphs over the past 24-240 hr, in 24 hr increments. This created a quick visual to confirm that the daily fluctuations were within the expected limits. Viewing the web interface also showed the current chamber status, as well as provided alerts to potential problems such as not receiving recent data. At any time the complete dataset could also be downloaded from the web interface.
We measured photosynthetically active radiation (PAR) in the immediate interior and exterior of the chamber at four points with and without the lights on in the middle of the summer and the middle of the day using a quantum sensor. The PAR in the chamber was 31.5% lower than the exterior when the chamber lights were off and 22% lower than the exterior when the lights were on. Thus, the chamber lights help to significantly increase PAR penetration within the chamber by 9.5% (P&#60;0.05).
Our continuous labeling system was able to produce 2759 g of A. gerardii biomass, 37% of which was aboveground biomass and 63% of which was belowground biomass. We achieved a 4.4 atom%13C whole plant label in our uniform plant material by setting the solenoid valves on the two CO2 tanks accordingly (Figure 1, Table 1). We achieved a 6.7 atom%15N whole plant label in our uniform plant material by mixing 98 atom%15N-KNO3 with 0.37 atom%15N-KNO3 in the KNO3 subsolution of a modified Hoagland&#39;s solution23 (Table 1). We watered the A. gerardii weekly with 750 ml total fluid (water plus Hoagland&#39;s solution) throughout the growing season. We fertilized with 200-500 ml of&#160;15N labeled Hoagland&#39;s solution per week depending on plant productivity.
We utilized the hot water extraction method to determine if there were isotopic differences between the uniform and differentially labeled plant material. For the differentially labeled plants, upon harvest we removed any leaves that were completely dead and handled these separately as they were likely not differentially labeled. When looking at&#160;13C content, all four incorporation days were significantly different from each other for the whole plant and the hot water extract, but for the hot water residue day 14 and 22 were not significantly different from each other (Table 1). When comparing the plant tissue fractions within each day, the hot water extract and residue were significantly different from each other for all four days and by day 22 the whole plant, extract, and residue were all significantly different from each other (Table 1). For the&#160;15N incorporation into plant components, there were differences between days of incorporation and plant tissue fractions. For the hot water extract all four of the incorporation days were significantly different from each other for&#160;15N, and for the whole plant and the hot water residue the shorter days of incorporation were significantly different than the longer days of incorporation (Table 1). The plant tissue fractions in the uniform plants were not significantly different from each other in&#160;15N, but the hot water extract and residue were significantly different from each other in&#160;15N for the differentially labeled litter.
All isotopic values are reported using the atom percent (atom %) notation (Equation&#160;1), which is a more accurate notation than %&#160;to use at high levels of heavy isotope enrichment21. For example:
<img alt="" height="54" src="/files/ftp_upload/51117/51117eq2.jpg" width="202" /> (1)
For this study, we ran statistical analyses using SAS version 9.2. We tested differences between the chamber interior and exterior light levels using a paired t-test. We tested differences between&#160;13C and&#160;15N labeling of hot water extracts and hot water residues using one-way analysis of variance (ANOVA) in PROC ANOVA. We used Duncan&#39;s multiple range test for multiple comparisons analysis. Significance was accepted at a P- level of 0.05. We used a Wilcoxon rank sum test to test that the data met the assumptions of the analysis.
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/51117/51117fig1highres.jpg" src="/files/ftp_upload/51117/51117fig1.jpg" /> Figure 1.&#160;Schematic diagram of the 40 pot capacity continuous multi-isotope labeling chamber from a bird&#39;s eye view. Dotted lines represent electrical wiring, while solid lines represent gas or water tubing. <a href="https://www.jove.com/files/ftp_upload/51117/51117fig1highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/51117/51117fig2highres.jpg" src="/files/ftp_upload/51117/51117fig2.jpg" /> Figure 2.&#160;A) Average CO2 concentration (ppm) (+/- SE) over a twenty-four hour period for an entire growing season. B) Average temperature (&#186;C), open circles, and humidity (%), closed circles (+/- SE) over a twenty-four hour period for an entire growing season. 
<a href="https://www.jove.com/files/ftp_upload/51117/51117fig2highres.jpg" target="_blank">Click here to view larger image.</a>
<table border="0" cellpadding="0" cellspacing="0" width="559">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
		<col span="2" />
	</colgroup>
	<tbody>
		<tr height="17">
			<td height="17" style="height:17px;width:77px;"></td>
			<td style="width:85px;"></td>
			<td style="width:92px;">Uniform (0)</td>
			<td style="width:97px;">Differential (7)</td>
			<td style="width:104px;">Differential (14)</td>
			<td style="width:104px;">Differential (22)</td>
		</tr>
		<tr height="19">
			<td height="38" rowspan="2" style="height:38px;width:77px;">Whole Litter</td>
			<td>13C Atom %</td>
			<td>4.46&#177;0.02 Aab</td>
			<td>3.93&#177;0.05 Ba</td>
			<td>3.64&#177;0.03 Ca</td>
			<td>3.35&#177;0.06 Db</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">15N Atom %</td>
			<td>6.69&#177;0.07 Aa</td>
			<td>6.72&#177;0.01 Aa</td>
			<td>6.33&#177;0.06 Ba</td>
			<td>6.41&#177;0.07 Ba</td>
		</tr>
		<tr height="19">
			<td height="38" rowspan="2" style="height:38px;width:77px;">Hot Water Extract</td>
			<td>13C Atom %</td>
			<td>4.59&#177;0.04 Aa</td>
			<td>3.35&#177;0.06 Bb</td>
			<td>2.79&#177;0.06 Cb</td>
			<td>2.37&#177;0.03 Dc</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">15N Atom %</td>
			<td>6.69&#177;0.03 Aa</td>
			<td>6.43&#177;0.01 Bb</td>
			<td>5.89&#177;0.07 Db</td>
			<td>6.16&#177;0.05 Cb</td>
		</tr>
		<tr height="19">
			<td height="38" rowspan="2" style="height:38px;width:77px;">Hot Water Residue</td>
			<td>13C Atom %</td>
			<td>4.37&#177;0.06 Ab</td>
			<td>4.1&#177;0.03 Ba</td>
			<td>3.79&#177;0.10 Ca</td>
			<td>3.66&#177;0.05 Ca</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">15N Atom %</td>
			<td>6.57&#177;0.04 Ba</td>
			<td>6.71&#177;0.02 Aa</td>
			<td>6.45&#177;0.02 Ca</td>
			<td>6.44&#177;0.03 Ca</td>
		</tr>
	</tbody>
</table>
Table 1.&#160;Isotopic composition and litter chemistry for uniform and differentially labeled litter. Days of differential labeling outside the chamber are in parentheses. Comparisons between days of incorporation are in capital letters (across rows) and between litter fractions are in lower case letters (down columns) for each variable.

Watch this Scientific Journal Video about Design and Operation of a Continuous 13C and 15N Labeling Chamber for Uniform or Differential, Metabolic and Structural, Plant Isotope Labeling at JoVE.com

Design and Operation of a Continuous 13C and 15N Labeling Chamber for Uniform or Differential, Metabolic and Structural, Plant Isotope Labeling

This method explains how to build and operate a continuous 13C and 15N isotope labeling chamber for uniform or differential plant tissue labeling. Representative results from metabolic and structural labeling of Andropogon gerardii are discussed.

Design and Operation of a Continuous 13C and 15N Labeling Chamber for Uniform or Differential, Metabolic and Structural, Plant Isotope Labeling

Tracing rare stable isotopes from plant material through the ecosystem provides the most sensitive information about ecosystem processes; from CO2 fluxes ...

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JoVE Journal

Environment

21.5K Views.  Colorado State University. This method explains how to build and operate a continuous 13C and 15N isotope labeling chamber for uniform or differential plant tissue labeling. Representative results from metabolic and structural labeling of Andropogon gerardii are discussed.

Video: Design and Operation of a Continuous 13C and 15N Labeling Chamber for Uniform or Differential, Metabolic and Structural, Plant Isotope Labeling

A Rapid and Efficient Method for Assessing Pathogenicity of Ustilago maydis on Maize and Teosinte Lines

Maize is a major cereal crop worldwide. However, susceptibility to biotrophic pathogens is the primary constraint to increasing productivity. U. maydis is a biotrophic fungal pathogen and the causal agent of corn smut on maize. This disease is responsible for significant yield losses of approximately $1.0 billion annually in the U.S.1 Several methods including crop rotation, fungicide application and seed treatments are currently used to control corn smut2. However, host resistance is the only practical method for managing corn smut. Identification of crop plants including maize, wheat, and rice that are resistant to various biotrophic pathogens has significantly decreased yield losses annually3-5. Therefore, the use of a pathogen inoculation method that efficiently and reproducibly delivers the pathogen in between the plant leaves, would facilitate the rapid identification of maize lines that are resistant to U. maydis. As, a first step toward indentifying maize lines that are resistant to U. maydis, a needle injection inoculation method and a resistance reaction screening method was utilized to inoculate maize, teosinte, and maize x teosinte introgression lines with a U. maydis strain and to select resistant plants.
Maize, teosinte and maize x teosinte introgression lines, consisting of about 700 plants, were planted, inoculated with a strain of U. maydis, and screened for resistance. The inoculation and screening methods successfully identified three teosinte lines resistant to U. maydis. Here a detailed needle injection inoculation and resistance reaction screening protocol for maize, teosinte, and maize x teosinte introgression lines is presented. This study demonstrates that needle injection inoculation is an invaluable tool in agriculture that can efficiently deliver U. maydis in between the plant leaves and has provided plant lines that are resistant to U. maydis that can now be combined and tested in breeding programs for improved disease resistance.

A Rapid and Efficient Method for Assessing Pathogenicity of Ustilago maydis on Maize and Teosinte Lines

The use of a needle injection method to inoculate maize and teosinte plants with the biotrophic pathogen Ustilago maydis is described. The needle injection inoculation method facilitates the controlled delivery of the fungal pathogen in between the plant leaves where the pathogen enters the plant through the formation of appresoria. This method is highly efficient, enabling reproducible inoculations with U. maydis.

Maize is a major cereal crop worldwide. However, susceptibility to biotrophic pathogens is the primary constraint to increasing productivity. U. maydis is ...

Transcript and Metabolite Profiling for the Evaluation of Tobacco Tree and Poplar as Feedstock for the Bio-based Industry

The global demand for food, feed, energy, and water poses extraordinary challenges for future generations. It is evident that robust platforms for the exploration of renewable resources are necessary to overcome these challenges. Within the multinational framework MultiBioPro we are developing biorefinery pipelines to maximize the use of plant biomass. More specifically, we use poplar and tobacco tree (Nicotiana glauca) as target crop species for improving saccharification, isoprenoid, long chain hydrocarbon contents, fiber quality, and suberin and lignin contents. The methods used to obtain these outputs include GC-MS, LC-MS and RNA sequencing platforms. The metabolite pipelines are well established tools to generate these types of data, but also have the limitations in that only well characterized metabolites can be used. The deep sequencing will allow us to include all transcripts present during the developmental stages of the tobacco tree leaf, but has to be mapped back to the sequence of Nicotiana tabacum. With these set-ups, we aim at a basic understanding for underlying processes and at establishing an industrial framework to exploit the outcomes. In a more long term perspective, we believe that data generated here will provide means for a sustainable biorefinery process using poplar and tobacco tree as raw material. To date the basal level of metabolites in the samples have been analyzed and the protocols utilized are provided in this article.

Plant biomass offers a renewable resource for multiple products, including fuel, feed, food, and a variety of materials. In this paper we investigate the properties of tobacco tree (Nicotiana glauca) and poplar as suitable sources for a biorefinery pipeline.

The global demand for food, feed, energy, and water poses extraordinary challenges for future generations. It is evident that robust platforms for the ...

Design and Construction of an Urban Runoff Research Facility

As the urban population increases, so does the area of irrigated urban landscape. Summer water use in urban areas can be 2-3x winter base line water use due to increased demand for landscape irrigation. Improper irrigation practices and large rainfall events can result in runoff from urban landscapes which has potential to carry nutrients and sediments into local streams and lakes where they may contribute to eutrophication. A 1,000 m2 facility was constructed which consists of 24 individual 33.6 m2 field plots, each equipped for measuring total runoff volumes with time and collection of runoff subsamples at selected intervals for quantification of chemical constituents in the runoff water from simulated urban landscapes. Runoff volumes from the first and second trials had coefficient of variability (CV) values of 38.2 and 28.7%, respectively. CV values for runoff pH, EC, and Na concentration for both trials were all under 10%. Concentrations of DOC, TDN, DON, PO4-P, K+, Mg2+, and Ca2+ had CV values less than 50% in both trials. Overall, the results of testing performed after sod installation at the facility indicated good uniformity between plots for runoff volumes and chemical constituents. The large plot size is sufficient to include much of the natural variability and therefore provides better simulation of urban landscape ecosystems.

This paper describes the design, construction, and function of a 1,000 m2 facility containing 24 individual 33.6 m2 field plots equipped for measuring total runoff volumes with time and collection of runoff subsamples at selected intervals for quantification of chemical constituents in the runoff water from simulated home lawns.

As the urban population increases, so does the area of irrigated urban landscape. Summer water use in urban areas can be 2-3x winter base line water use ...

Operation of a 25 KWth Calcium Looping Pilot-plant with High Oxygen Concentrations in the Calciner

Calcium looping (CaL) is a post-combustion CO2 capture technology that is suitable for retrofitting existing power plants. The CaL process uses limestone as a cheap and readily available CO2 sorbent. While the technology has been widely studied, there are a few available options that could be applied to make it more economically viable. One of these is to increase the oxygen concentration in the calciner to reduce or eliminate the amount of recycled gas (CO2, H2O and impurities); therefore, decreasing or removing the energy necessary to heat the recycled gas stream. Moreover, there is a resulting increase in the energy input due to the change in the combustion intensity; this energy is used to enable the endothermic calcination reaction to occur in the absence of recycled flue gases. This paper presents the operation and first results of a CaL pilot plant with 100% oxygen combustion of natural gas in the calciner. The gas coming into the carbonator was a simulated flue gas from a coal-fired power plant or cement industry. Several limestone particle size distributions are also tested to further explore the effect of this parameter on the overall performance of this operating mode. The configuration of the reactor system, the operating procedures, and the results are described in detail in this paper. The reactor showed good hydrodynamic stability and stable CO2 capture, with capture efficiencies of up to 70% with a gas mixture simulating the flue gas of a coal-fired power plant.

Operation of a 25 KWth Calcium Looping Pilot-plant with High Oxygen Concentrations in the Calciner

This manuscript describes a procedure for operating a calcium looping pilot-plant for post-combustion carbon capture with high oxygen concentrations in the calciner in order to reduce or eliminate the flue gas recycle.

Calcium looping (CaL) is a post-combustion CO2 capture technology that is suitable for retrofitting existing power plants. The CaL process uses limestone ...

Continuous Hydrologic and Water Quality Monitoring of Vernal Ponds

Vernal ponds, also referred to as vernal pools, provide critical ecosystem services and habitat for a variety of threatened and endangered species. However, they are vulnerable parts of the landscapes that are often poorly understood and understudied. Land use and management practices, as well as climate change are thought to be a contribution to the global amphibian decline. However, more research is needed to understand the extent of these impacts. Here, we present methodology for characterizing a vernal pond's morphology and detail a monitoring station that can be used to collect water quantity and quality data over the duration of a vernal pond's hydroperiod. We provide methodology for how to conduct field surveys to characterize the morphology and develop stage-storage curves for a vernal pond. Additionally, we provide methodology for monitoring the water level, temperature, pH, oxidation-reduction potential, dissolved oxygen, and electrical conductivity of water in a vernal pond, as well as monitoring rainfall data. This information can be used to better quantify the ecosystem services that vernal ponds provide and the impacts of anthropogenic activities on their ability to provide these services.

Understanding the ecosystem services and processes provided by vernal ponds and the impacts of anthropogenic activities on their ability to provide these services requires intensive hydrologic monitoring. This sampling protocol using in-situ monitoring equipment was developed to understand the impact of anthropogenic activities on water levels and quality.

Vernal ponds, also referred to as vernal pools, provide critical ecosystem services and habitat for a variety of threatened and endangered species. ...

The Plant Infection Test: Spray and Wound-Mediated Inoculation with the Plant Pathogen Magnaporthe Grisea

Plants possess a powerful system to defend themselves against potential threats by pathogenic fungi. For agriculturally important plants, however, current measures to combat such pathogens have proved too conservative and, thus, not sufficiently effective, and they can potentially pose environmental risks. Therefore, it is extremely necessary to identify host-resistance factors to assist in controlling plant diseases naturally through the identification of resistant germplasm, the isolation and characterization of resistance genes, and the molecular breeding of resistant cultivars. In this regard, there is need to establish an accurate, rapid, and large-scale inoculation method to breed and develop plant resistance genes. The rice blast fungal pathogen Magnaporthe grisea causes severe disease symptoms and yield losses. Recently, M. grisea has emerged as a model organism for studying the mechanisms of plant-fungal pathogen interactions. Hence, we report the development of a plant virulence test method that is specific for M. grisea. This method provides for both spray inoculation with a conidial suspension and wounding inoculation with mycelium cubes or droplets of conidial suspension. The key step of the wounding inoculation method for detached rice leaves is to make wounds on plant leaves, which avoids any interference caused by host penetration resistance. This spray/wounding protocol contributes to the rapid, accurate, and large-scale screening of the pathotypes of M. grisea isolates. This integrated and systematic plant infection method will serve as an excellent starting point for gaining a broad perspective of issues in plant pathology.

The Plant Infection Test: Spray and Wound-Mediated Inoculation with the Plant Pathogen Magnaporthe Grisea

Here, we present a protocol to test plant virulence with the plant pathogen Magnaporthe grisea. This report will contribute to the large-scale screening of the pathotypes of fungi isolates and serve as an excellent starting point for understanding the resistant mechanisms of plants during molecular breeding.

Plants possess a powerful system to defend themselves against potential threats by pathogenic fungi. For agriculturally important plants, however, current ...

Quantifying Plant Soluble Protein and Digestible Carbohydrate Content, Using Corn (Zea mays) As an Exemplar

Elemental data are commonly used to infer plant quality as a resource to herbivores. However, the ubiquity of carbon in biomolecules, the presence of nitrogen-containing plant defensive compounds, and variation in species-specific correlations between nitrogen and plant protein content all limit the accuracy of these inferences. Additionally, research focused on plant and/or herbivore physiology require a level of accuracy that is not achieved using generalized correlations. The methods presented here offer researchers a clear and rapid protocol for directly measuring plant soluble proteins and digestible carbohydrates, the two plant macronutrients most closely tied to animal physiological performance. The protocols combine well characterized colorimetric assays with optimized plant-specific digestion steps to provide precise and reproducible results. Our analyses of different sweet corn tissues show that these assays have the sensitivity to detect variation in plant soluble protein and digestible carbohydrate content across multiple spatial scales. These include between-plant differences across growing regions and plant species or varieties, as well as within-plant differences in tissue type and even positional differences within the same tissue. Combining soluble protein and digestible carbohydrate content with elemental data also has the potential to provide new opportunities in plant biology to connect plant mineral nutrition with plant physiological processes. These analyses also help generate the soluble protein and digestible carbohydrate data needed to study nutritional ecology, plant-herbivore interactions and food-web dynamics, which will in turn enhance physiology and ecological research.

Quantifying Plant Soluble Protein and Digestible Carbohydrate Content, Using Corn Zea mays As an Exemplar

The protocols described herein provide a clear and approachable methodology for measuring soluble protein and digestible (non-structural) carbohydrate content in plant tissues. The ability to quantify these two plant macronutrients has significant implications for advancing the fields of plant physiology, nutritional ecology, plant-herbivore interactions and food-web ecology.

Elemental data are commonly used to infer plant quality as a resource to herbivores. However, the ubiquity of carbon in biomolecules, the presence of ...

Colletotrichum fioriniae Development in Water and Chloroform-based Blueberry and Cranberry Floral Extracts

To accurately monitor the phenology of the bloom period and the temporal dynamics of floral chemical cues on fungal fruit rotting pathogens, floral extraction methods and coverslip bioassays were developed utilizing Colletotrichum fioriniae. In blueberry and cranberry, this pathogen is optimally controlled by applying fungicides during the bloom period because of the role flowers play in the initial stages of infection. The protocol detailed here describes how floral extracts (FE) were obtained using water-, chloroform-, and field rainwater-based methods for later use in corresponding glass coverslip bioassays. Each FE served to provide a different set of information: response of C. fioriniae to mobilized floral chemical cues in water (water-based), pathogen response to flower and fruit surface waxes (chloroform-based), and field-based monitoring of collected floral rainwater, moving in vitro observations to an agricultural setting. The FE is broadly described as either water- or chloroform-based, with an appropriate bioassay described to compensate for the inherent differences between these two materials. Rainwater that had run off flowers was collected in unique devices for each crop, alluding to the flexibility and application of this approach for other crop systems. The bioassays are quick, inexpensive, simple, and provide the ability to generate spatiotemporal and site-specific information about the presence of stimulatory floral compounds from various sources. This information will ultimately better inform disease management strategies, as FE decrease the time needed for infection to occur, thus providing insight into changing risks for pathogen infection over the growing season.

Colletotrichum fioriniae Development in Water and Chloroform-based Blueberry and Cranberry Floral Extracts

Here, bioassays designed to monitor the development of a fungal pathogen, Colletotrichum fioriniae, in the presence of blueberry or cranberry floral extracts on glass coverslips are described. Water-, chloroform-, and field rainwater- based floral extraction techniques are detailed as well as insight into how this information can be applied.

To accurately monitor the phenology of the bloom period and the temporal dynamics of floral chemical cues on fungal fruit rotting pathogens, floral ...

A Uniaxial Compression Experiment with CO2-Bearing Coal Using a Visualized and Constant-Volume Gas-Solid Coupling Test System

Injecting carbon dioxide (CO2) into a deep coal seam is of great significance for reducing the concentration of greenhouse gases in the atmosphere and increasing the recovery of coalbed methane. A visualized and constant-volume gas-solid coupling system is introduced here to investigate the influence of CO2 sorption on the physical and mechanical properties of coal. Being able to keep a constant volume and monitor the sample using a camera, this system offers the potential to improve instrument accuracy and analyze fracture evolution with a fractal geometry method. This paper provides all steps to perform a uniaxial compression experiment with a briquette sample in different CO2 pressures with the gas-solid coupling test system. A briquette, cold-pressed by raw coal and sodium humate cement, is loaded in high-pressure CO2, and its surface is monitored in real-time using a camera. However, the similarity between the briquette and the raw coal still needs improvement, and a flammable gas such as methane (CH4) cannot be injected for the test. The results show that CO2 sorption leads to peak strength and elastic modulus reduction of the briquette, and the fracture evolution of the briquette in a failure state indicates fractal characteristics. The strength, elastic modulus, and fractal dimension are all correlated with CO2 pressure but not with a linear correlation. The visualized and constant-volume gas-solid coupling test system can serve as a platform for experimental research about rock mechanics considering the multifield coupling effect.

A Uniaxial Compression Experiment with CO2-Bearing Coal Using a Visualized and Constant-Volume Gas-Solid Coupling Test System

This protocol demonstrates how to prepare a briquette sample and conduct a uniaxial compression experiment with a briquette in different CO2 pressures using a visualized and constant-volume gas-solid coupling test system. It also aims to investigate changes in terms of coal&#8217;s physical and mechanical properties induced by CO2 adsorption.

Injecting carbon dioxide (CO2) into a deep coal seam is of great significance for reducing the concentration of greenhouse gases in the atmosphere and ...

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

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

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

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