PANDA: A simple and affordable chamber system for measuring the whole-plant net CO2flux

PANDA: A simple and affordable chamber system for measuring the whole-plant net CO2 flux | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results PANDA: A simple and affordable chamber system for measuring the whole-plant net CO 2 flux View ORCID Profile Philipp Schuler , View ORCID Profile Yuxin Li , Pierre Pittet , View ORCID Profile Patrick Favre , View ORCID Profile Mai-He Li , View ORCID Profile Yan-Li Zhang , View ORCID Profile Charlotte Grossiord doi: https://doi.org/10.1101/2025.06.02.657330 Philipp Schuler 1 Plant Ecology Research Laboratory PERL, School of Architecture , Civil and Environmental Engineering, EPFL, 1015 Lausanne, Switzerland 2 Community Ecology Unit, Swiss Federal Institute for Forest , Snow and Landscape WSL, 1015 Lausanne, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Philipp Schuler For correspondence: philipp.schuler{at}wsl.ch Yuxin Li 1 Plant Ecology Research Laboratory PERL, School of Architecture , Civil and Environmental Engineering, EPFL, 1015 Lausanne, Switzerland 3 Department of Ecology. College of Urban and Environmental Sciences, Peking University , 100871 Beijing, China Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Yuxin Li Pierre Pittet 4 Faculté des géosciences et de l’environnement, Université de Lausanne , 1015 Lausanne, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site Patrick Favre 1 Plant Ecology Research Laboratory PERL, School of Architecture , Civil and Environmental Engineering, EPFL, 1015 Lausanne, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Patrick Favre Mai-He Li 5 Forest Dynamics, Swiss Federal Institute for Forest , Snow and Landscape WSL, 8903 Birmensdorf, Switzerland 6 Key Laboratory of Geographical Processes and Ecological Security in Changbai Mountains, Ministry of Education, School of Geographical Sciences, Northeast Normal University , Changchun, Jilin, China 7 College of Life Science, Hebei University , Baoding, Hebei, China Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Mai-He Li Yan-Li Zhang 5 Forest Dynamics, Swiss Federal Institute for Forest , Snow and Landscape WSL, 8903 Birmensdorf, Switzerland 8 Department of Environmental Systems Science, Institute of Terrestrial Ecosystems , ETH Zürich, Zürich, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Yan-Li Zhang Charlotte Grossiord 1 Plant Ecology Research Laboratory PERL, School of Architecture , Civil and Environmental Engineering, EPFL, 1015 Lausanne, Switzerland 2 Community Ecology Unit, Swiss Federal Institute for Forest , Snow and Landscape WSL, 1015 Lausanne, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Charlotte Grossiord Abstract Full Text Info/History Metrics Preview PDF Summary The carbon (C) balance of plants is the sum of all source and sink processes. However, due to methodological limitations, most studies focus predominantly on measurements of leaf-level assimilation and respiration, with less attention given to these processes in heterotrophic organs or the whole-plant level. As a result, knowledge of the whole-plant net C balance is scarce, limiting our understanding of the dynamics between C source and sink activities. Therefore, we developed an easily reproducible chamber system for continuous measurements of whole-plant net CO 2 fluxes. We present the obtained dynamics of net CO 2 fluxes of several C 3 and CAM species, including germinating Quercus robur , over several days, as well as the whole-plant net CO 2 flux temperature response of Q. robur seedlings, identifying the temperature thresholds at which they shift from a net CO 2 sink to source. We show distinct diel patterns of net CO 2 fluxes in C 3 plants, likely driven by a dynamic diurnal up- and downregulation of sink activities in woody C 3 plants. These patterns appear temperature-driven, suggesting a dynamic response of plant’s sink and source activity to environmental drivers. Our results highlight the importance of whole-plant C balance measurements for understanding plant responses to environmental conditions. Introduction Determining net CO 2 fluxes at the whole-plant level comes with difficulties, since it integrates several parallel autotrophic and heterotrophic fluxes over the whole plant body, including leaves, branches, and roots. Most commonly, gas-exchange measurements are conducted at the leaf-level to quantify net assimilative CO 2 fixation (A net ) and respiratory (R) CO 2 release. In contrast, fewer studies address fluxes from other organs, such as branches, stems ( Han et al., 2017 ; Diao et al., 2020 ; Zhang et al., 2025 ), and roots (e.g., Atkin et al., 2000 ; Kuzyakov and Larionova, 2006 ). These organs, especially roots, act as major carbon (C) sinks and contribute significantly to the plant’s overall C balance by consuming up to more than 50% of all assimilates ( Lambers et al., 2002 ; Colombi et al., 2021 ), and it is known that the root- and stem-level metabolic responses to temperature can have a significant impact to whole plant level C relations ( Atkin et al., 2000 ; Wang and Hoch, 2022 ). Moreover, twig and stem CO 2 fixation can also significantly contribute to the whole plant C balance, reassimilatint 7 to 123% of respired CO 2 in non-succulent species ( Ávila et al., 2014 ). Still, there are methodological difficulties in measuring the net CO 2 flux of a particular piece of twig or stem, for example, stem photosynthesis can occur in plant species with and without stomata on these organs ( Ávila et al., 2014 ), making measurements difficult to interpret and hard to compare. Furthermore, CO 2 is being redistributed within the plant xylem by the sap stream ( Teskey et al., 2008 ; Grossiord et al., 2012 ; Trumbore et al., 2013 ), causing strong reduction of the measured CO 2 efflux of up to 40 % under high xylem sap velocity ( Gansert and Burgdorf, 2005 ). Moreover, diel patterns of stem CO 2 efflux differs between phylogenetic groups such as monocots, cycads, as well as angiosperm and gymnosperm trees ( Marler and Lindström, 2020 ), making an accurate comparable temporal tracking and quantification of tissue-specific measurements to the whole plant level challenging. Measuring whole-plant net CO 2 fluxes remains challenging due to the technical difficulties of enclosing entire plants, accurately capturing integrated gas exchange over time, and simultaneously excluding soil respiration, therefore, only few studies have been able to measure whole-plant net CO 2 fluxes ( Hand, 1973 ; Dutton et al., 1988 ; Zhao et al., 2013 ; Brauner et al., 2014 , 2018 ; Patono et al., 2023 ). This methodological gap limits our ability to fully understand how entire plants respond to environmental stressors under realistic conditions. One study that successfully measured whole plant net CO 2 fluxes demonstrated that the carbon balance of Thuja occidentalis L. becomes negative - reflecting respiratory losses exceed photosynthetic gains - at temperatures above 35°C and mild drought ( Zhao et al., 2013 ). This study highlights the high potential of such measurements for understanding plant responses to climatic stress and for predicting future shifts in the global C cycle. Recent systems developed for tracking the net CO 2 flux in larger plants ( Patono et al., 2023 ), while valuable, are often large, technically complex, and relatively difficult to reproduce, making them impractical for routine use in standard growth facilities or climate chambers. Smaller-scale systems suitable for controlled environments frequently measure only the aboveground biomass ( Kölling et al., 2015 ; Salvatori et al., 2021 ), which strongly bias estimates of whole plant net CO 2 balance - especially in larger plants such as trees, where root biomass can be comparable to that of the canopy ( Perry, 1982 ; Harris, 1992 ). However, existing small systems to capture whole plant net CO 2 flux are designed to measure small plants such as Arabidopsis thaliana (L.) Heynh.( Brauner et al., 2014 , 2018 ), and therefore are not suitable for larger plant species such as woody plants. Nevertheless, whole-plant C exchange measurements offer a key advantage in addressing current hot topics in plant science by capturing integrated responses that are missed by regular tissue-level measurements or ecosystem-scale approaches like eddy covariance, which are showing discrepancies in their evaluation of the whole-plant C dynamics ( Campioli et al., 2016 ). For instance, it is crucial to isolate how the C balance of whole plants is affected by the ongoing exacerbation of heatwaves ( Meehl and Tebaldi, 2004 ; Perkins-Kirkpatrick and Lewis, 2020 ), as these events can differentially impact photosynthesis, respiration, as well as C allocation across tissues ( Birami et al., 2018 ; Werner et al., 2020 ). For instance, stem photosynthesis can buffer against the sharp decline in leaf photosynthesis during heatwaves ( Ávila-Lovera et al., 2024 ) - effects that are only fully captured when measured at the whole-plant scale. Furthermore, direct measurements of whole plant net CO 2 fluxes could have the potential for a faster selection of heat-resistant genotypes in plant breeding, both in agriculture and forestry. Finally, Earth System Models are used to simulate the future pace and extent of CO₂ increases and climate change ( Prinn, 2013 ), integrating multiple subcomponents such as plant functional types, vegetation dynamics, C fluxes, as well as their response to climate on high temporal resolution. Therefore, an accurate knowledge of whole plant net CO 2 fluxes of different functional types responds to changes in temperature, how they change during plant development, and on a diel course, is essential for accurate predictions of the future trajectories of the global atmospheric CO 2 concentration. To address these gaps, we developed an easy-to-replicate and low-cost chamber system ( P lant A ssimilatio ND yn A mics (PANDA) that can be attached to any gas exchange system capable of measuring and regulating CO 2 concentrations, such as the WALZ GFS-3000 (Heinz Walz GmbH, 91090 Effeltrich, Germany) or the Li-6800 system (LI-COR Environmental, Lincoln, NE 68504, United States). To validate this system while simultaneously addressing a key uncertainty in whole-plant carbon fluxes, namely how daytime, growth and temperature influence the whole plant net CO₂ balance, we designed four experiments: To assess acclimation effects to the system, we first continuously measured the whole plant net CO 2 balance of three C 3 ( Brachychiton acerifolius (A.Cunn. ex G.Don) F.Muell., Trachycarpus fortunei (Hook.) H.Wendl., Coffea arabica L.) and three CAM species ( Aloe arborescens Mill., Opuntia Mill., Phalaenopsis Blume), the latter which exhibit distinct and well-studied diel changes in their net CO 2 flux ( Osmond et al., 1979 ), over two to three diel cycles. This experiment allowed us to test whether the measurements of a plant remains constant for several days of normal day/night cycles. In a second experiment, we monitored the whole-plant net CO 2 flux during several days of constantly light or dark conditions to investigate whether the system can be used to identify circadian cycles of the net CO 2 balance independent of light availability. To provide first ideas for potential plant physiological investigations with the system, we monitored the net CO 2 balance of germinating Quercus robur L. acorns over one month. Finally, we subjected the Q. robur seedlings twice to increasing temperatures of about 27, 34, and 43°C. Once after the first seedling growth fully finished, and once after the plants produced a second flush. With this, we could investigate potential changes of the response due to changes in ratios of autotrophic to heterotrophic tissues between the curves after the first and second flush. Materials and Methods Chamber design and use The P lant A ssimilatio ND yn A mics (PANDA) chamber ( Fig. 1 ) consists of (Fig.1b, c) a 30 × 30 × 1 cm bottom plate (1) that features a circular recess (2; 24.5–25.5 cm diameter) accommodating air inlet/outlet ports (AI, AO) from and to the gas analyzing system (we used a GFS-3000 Gas-Exchange System, Heinz Walz GmbH, Germany), a water inlet (WI) for watering, where a watering hose is connected on both sides of the plate, and a cable gland (CG). Air and water inlet ports were connected with silicone tubes on both sides of the plate. Metal feet (F) measuring 8 cm are fixed at each corner. The internal fans and sensor connect to and are controlled by a Raspberry Pi 5 (Raspberry Pi Foundation, England) positioned outside the chamber, with live data displayed on an attached screen. Inside, two axial fans (V; Sunon MF60151V2-1000U-A99, Sunon Fans, Taiwan) and a temperature/humidity sensor (TH; Adafruit Sensirion SHT45 Precision Temperature Humidity Sensor, Adafruit, USA) are present. The chamber (a) is closed with an 80 cm tall, 25 cm diameter glass cylinder (C; Sandra Rich GMBH, Germany). To prevent condensation water from reaching the gas analyzer, a water trap (WT) had to be added to the air outlet tube outside the chamber, approximately 1m away from both the chamber and gas analyzer, with the water trap representing the lowest position of the whole system. It must be noted that the PANDA measuring chamber was not designed to regulate air temperature, relative humidity, or light but rather to serve an enclosed chamber to measure gas exchange fluxes. The temperature and light must be defined either by natural light and ambient temperature variation (e.g., by using the system outdoors) or actively being regulated with additional devices, such as heating units, LED light sources, or by placing the system inside a climate chamber. Due to the high evapotranspiration, the relative humidity (RH) raises rapidly (usually >80%), especially in well-irrigated plants, when connected to a standard gas analyzing system with a comparable low flow rate, such as the WALZ GFS-3000, which was used in this study. Hence, for researchers aiming to tightly control RH, we recommend either increasing the system’s flow rate, which must use a more powerful control unit than the here used WALZ GFS-3000, or by integrating a bypass desiccant system. Download figure Open in new tab Figure 1: The (a, b) PlantAssimilatioNDynAmics (PANDA) chamber consists of (c, d) a bottom plate (1; 30 x 30 x 1 cm) with a circular recess (2) with an inner diameter of 24.5 cm and an outer diameter of 25.5 cm milled into it. Inside the recess, an air inlet port (AI), an air outlet port (AO), a water inlet port (WI), and a cable gland (CG) are inserted. On each corner of the plate, 8 cm high metal feet (F) are attached. Inside the chamber (a), two fans (V; Sunon MF60151V2-1000U-A99 axial fan 12 V/DC) as well as a temperature and humidity sensor (TH; Adafruit Sensirion SHT45 Precision Temperature Humidity Sensor). The fans and the sensor are connected and controlled, as well as the data stored on a Raspberry Pi 5, which is located outside the chamber, the latter of which was attached to a screen for visually tracking chamber conditions (not shown in the scheme). To prevent condensation water from reaching the gas analyzer, a water trap (WT) is installed in the air outlet tubing, approximately 1m away from both the chamber and gas analyzer, with the water trap representing the lowest position of the whole system. The chamber is sealed by using an 80 cm high glass cylinder with a diameter of 25 cm (C; Glass vase CYLI clear cylindrical, Sandra Rich). Depending on the specific requirements of the planned experiments, one can add more access ports to the bottom plate of the chamber. For instance, we recommend adding an outlet port for condensation water, which can accumulate at the bottom of the chamber, especially during long-term measurements. Furthermore, the chamber size can also be adjusted by using different sizes of transparent (glass) cylinders or similar. The main limitation is the maximum flow rate of the analyzer unit, which limits the amount of CO 2 that can be added. Since we were working with two light sources on two sides of the chamber ( Fig. 1b ), we installed four panels covered in aluminium foil on each side of the chamber (only shown three of four in Fig. 1b ) to provide equal high light availability around the plants. Calculation of the net CO 2 flux The net CO 2 flux in µmol s -1 was calculated as: with CO 2abs being the CO 2 concentration delivered to the chamber, ca is the CO 2 concentration that is coming from the chamber, and k is the conversion factor to convert the measured CO 2 flux from ppm into µmol, which is calculated as follows: leading to a k of 0.0009 μmol CO 2 s −1 ppm −1 at a flow rate of 900 µmol s⁻¹. Due to the large chamber volume, we choose the maximum flow rate possible for the system. To convert the resulting µmol CO 2 s -1 into µmol glucose s -1 , one can divide µmol CO 2 s -1 by 6, representing the stoichiometric C ratio of 1 µmol CO₂ to 1 µmol glucose. Calculation of the cumulative CO 2 and C flux The cumulative CO 2 and C flux was calculated as: where F CO2, i is the net CO₂ flux at time point i (µmol CO₂ s⁻¹), and Δ ti is the time interval between consecutive measurements (seconds). The summation is done over all time intervals from the start to the end of the period of interest. If the time intervals are equal (e.g., measurements every minute), the formula simplifies to: Example Calculation: If you have flux measurements every minute (60 seconds) and the net CO₂ flux at three consecutive time points is 0.5, 0.7, and 0.6 µmol s⁻¹, the cumulative flux after 3 minutes would be: and to finally convert the cumulative C balance into gravimetric units, we divide the resulting μmol (CO 2 ) by 6 to convert it into μmol (Glucose), and multiply this by the molar mass of glucose: Carbon-free substrate for plant growth The substrate used for the net CO 2 flux measurements is pure C-free perlite (Pflanzen Perlit, grain size 2-6mm, Otto Hauenstein Samen AG, Switzerland). The advantage of pure C-free perlite is that soil respiration is excluded from the measurement, as there is no organic matter or microbial C source to contribute to CO 2 emissions. Furthermore, since perlite possesses high porosity, it ensures rapid CO₂ exchange between the substrate and the chamber air space. Another advantage of using this pure white perlite is that it is easy to see contaminations such as algae or other microorganisms growing at its surface, something of great importance, especially during long-term measurements. As perlite lacks nutrients, they need to be added externally when plants are cultivated for a longer period of time, however, this also ensures that specific nutrient concentrations can also be precisely adjusted to the experimental needs. The high pore volume is providing well-aerated conditions at the root level but a relatively limited water-holding capacity ( Jackson, 1974 ) and therefore requires careful watering, which may not reflect the soil conditions certain plants typically experience in natural environments. Hence, root development and function might differ from those in more structured or biologically active soils, potentially influencing whole-plant physiological responses and C dynamics. Thus, it might be beneficial to grow plants in a more natural soil prior to placing them in the PANDA chamber, using C-free perlite only during the experimental stage. However, an optical check of the root development of the tested oak seedlings (Fig. S1) revealed no abnormalities, indicating normal root growth under the tested conditions. Plant cultivation prior to measurements and plant positioning in the chamber Plants are grown in perlite for at least 1 month to ensure the roots are acclimated to the porous growing material and the experimental light conditions. Plants were fertilized once per week with liquid inorganic fertilizer (Blumen-Dünger 1l (N-P-K 4-3-3), terrasan Schweiz GmbH, 8034 Zürich, Switzerland). The room temperature was 22-24°C, and the RH was ∼50-60%. Plants were grown under a 100W LED plant light (grow-led-pflanzenlampen.ch, Senn Services, Switzerland) with a 12-hour day/night cycle. To ensure that no organic C from root exudates or shedded root cells was contaminating the substrate, two days before the start of the experiment, the plants were carefully removed from the pots, the old perlite was carefully washed away from the roots, and the plants were repotted in new perlite. Therefore, any contribution of soil respiration to the measurements could be excluded. On the day the measurements started, plants were positioned with a horticultural saucer in the center of the bottom plate. The inner end of the watering hose was fixed below the pot, and the outer end of the watering hose was closed with a hemostatic clamp. After that, waterproof multi-purpose sealant (TEROSON RB IX, Henkel AG, Germany) was added by forming a 1 cm high and 2 cm wide wall on top of the circular recess, the glass cylinder was carefully positioned on the sealant, and the cylinder was tightly connected by applying gentle pressure on the top surface by weight. On two sides of the glass cylinder at a distance of 15cm, a 100W LED plant light (grow-led-pflanzenlampen.ch, Senn Services, Switzerland) was installed, running as well in a 12-hour day/night cycle. The photosynthetic active radiation, measured with the LI-6800 in the center of the bottom plate, was 1250 µmol m -2 s -1 when measured in the direction of either plant light. The temperature inside the chamber increased from an average of 22.1°C at night to an average of 31.8°C during the day (Fig. S2) due to the heat generated by the light. Due to the high evapotranspiration rate, the relative humidity (RH) was above 90%, so the vapor pressure deficit (VPD) remained below 1 kPa (Fig. S2). This temperature and RH regime was the same for all experiments in the next subsection, except for the temperature response curves of experiment 4. Continuous measurement of the whole plant net CO 2 flux The measurements were done by running an auto-program with the GFS-3000 system, which regulates the CO 2 concentration inside the chamber (ca ppm) at 420 ppm with a total flowrate of 900 µmol s -1 . Inflowing (CO 2abs ), outflowing (ca ppm), as well as the delta between the two (dCO 2 MP), were recorded every 2 minutes. Plants were watered by filling the horticultural saucer using a squeeze bottle via the water inlet port. To evaluate the performance of the system, we performed four experiments: Experiment 1: Diurnal variation in whole-plant CO 2 exchange in C 3 and CAM plants To test the sensitivity of the system and whether the results of the daily measurements are comparable between different days of measuring due to acclimation effects, we tracked the net CO₂ flux of single plants of three C 3 ( Brachychiton acerifolius (A.Cunn. ex G.Don) F.Muell., Trachycarpus fortunei (Hook.) H.Wendl., Coffea arabica L.) and three CAM species ( Aloe arborescens Mill., Opuntia Mill., Phalaenopsis Blume) over two to three diel cycles. Woody C 3 plants were included to investigate whether diel patterns could be observed in their net CO 2 flux, as has been observed in C 3 forbs (Resco De Dios et al., 2016 ), and CAM species, which are well known for their four diel oscillatory cycles of CAM photosynthesis ( Osmond et al., 1979 ), were included to evaluate the ability of the PANDA chamber to identify diel pattern. To compare the whole-plant net CO 2 flux with leaf-level net assimilation (A net at PAR = 1500, RH = 60%, T air = 30°C), leaf gas exchange was tracked with a Li-6800 over several hours of three leaves per plant. Experiment 2: Endogenous cycles in C 3 plants Additionally, we measured the net CO 2 flux of the C 3 plants under three to four days of continuous light or dark conditions to investigate potential endogenous rhythms ( Lüttge and Hertel, 2009 ) in their source- and sink dynamics. Indeed, many plants exhibit cyclical patterns of CO₂ uptake and release that follow a ∼24-hour rhythm. This suggests that photosynthesis and respiration are partially regulated by internal clocks, not just light availability (Resco De Dios et al., 2016 ). Understanding the extent of endogenous rhythms in CO 2 fluxes is important for accurate interpretation of gas exchange data, for the design of experiments, and for understanding and modeling plant C balance and growth. Experiment 3: Net CO 2 flux of establishing seedlings prior and during drought stress To evaluate whether the system can accurately track the C balance of developing plants over a longer period, we continuously tracked of the net CO 2 balance of germinating Quercus robur L. acorns. For this, we carefully peeled 15 slightly germinating (with initial root growth) acorns, weighed their initial weight, and planted 12 of them into fresh perlite in the same pot, and determined the initial dry mass of the remaining 3 acorns by drying them in the oven at 60°C for 7 days. The planted acorns and the pot have been placed inside the chamber, and the net CO 2 flux was continuously tracked over a whole month. Groups of three seedlings were carefully harvested after 9, 13, and 30 days to track the change in dry biomass of leaves, shoots, and roots after drying them in the oven at 60°C for 7 days. Finally, to study the impact of a mild drought on the net CO 2 flux of the seedlings, the plants were subjected to a mild drought by stopping the irrigation during the last four days of the germination experiments. Experiment 4: Shifts in whole-plant C balance with rising temperatures After the seedlings of experiment 3 finished their first growing cycle and had fully mature leaves, the three remaining plants were carefully taken out of the pot and the spent perlite was washed away. The plants were repotted in new perlite, again all plants in a single pot. Subsequently, the first of two temperature response experiments of whole plant net CO 2 flux was conducted (Round 1). For this, the plants were subjected to increasing target air temperatures of ∼27, ∼34, and ∼43°C over three days (i.e., changed temperature each day). The actual temperature values are displayed in Fig. S3. To decrease the air temperature in the system during the light hours of the first day, the temperature input of the light to the chamber surface was decreased by cooling the glass cylinder to about 27°C with two external desk fans. On the second day, the temperature input by the light was not altered, leading to a T air of about 34°C, while on the third day, T air was increased by carefully heating the system with an external heating vent (Trisa Heizlüfter Heat & Chill 2’000W, Trisa Electronics AG, Switzerland) to about 43°C. After this first temperature response experiment, the plants were flushed anew, and the plants grew inside the chamber until the leaves and shoots matured. While the new leaves and shoots were fully grown, the color of the new leaves was still slightly paler green compared to the ones of the first flush, indicating that their chlorophyll concentration did not reach the maximum. However, since the maximum CO 2 input into the system would have been surpassed, we continued with the second temperature response of the same temperature range (round 2) to compare the resulting response curves of the two plant sizes. Leaf area and biomass were determined by harvesting three seedlings after both rounds of temperature responses. Leaves were scanned (CanoScan LiDE 300, Canon, Tokyo, Japan), and the total leaf area was analyzed with ImageJ ( Schneider et al., 2012 ). The dry weight was measured (NewlassicMF, MS105DU, Mettler Toledo, Switzerland) after drying them in the oven at 60°C for 7 days. Results and Discussion Experiment 1: Diurnal variation in whole-plant CO 2 exchange in C 3 and CAM plants The net CO 2 flux measured with the PANDA chamber ( Fig. 2 ) showed robust tracking of the diel CO 2 fluxes over several days, which was reproducible between the different days, both in C 3 ( Fig. 2a ) and CAM ( Fig. 2b ) plants, thereby demonstrating the reliability and functionality of the system for capturing consistent whole-plant gas exchange dynamics. Download figure Open in new tab Figure 2: Diel dynamics of the net CO 2 flux of (a) the three measured C 3 and (b) CAM species over three to four days (n = 1 per species). Nighttime hours are shown in grey. The bold lines show the running means (10 measurements before and 10 measurements after each point, with one measurement every 2 minutes), and the fine lines show the corresponding measurements. All C 3 plants showed little variability in their CO 2 fluxes during the nighttime ( Fig. 2a ). However, their positive daytime fluxes were (i.e., indicating higher C uptake than release) highest at the beginning of the day, and showed species-specific trends, usually having the lowest net CO 2 balance just before the light turned off. Interestingly, the net CO 2 flux of the two dicotyledons B. acerifolius and C. arabica spiked, the former also with the overall most positive net CO 2 flux, at the beginning of the day, followed by a slow and steady increase after they reached a plateau, before declining steadily for the remainder of the day. However, the midday peak was less consistent between the two days of measurement in B. acerifolius . The smaller spike in C. arabica compared to B. acerifolius could be explained by its relatively low photosynthetic temperature optimum of 24°C ( Nunes et al., 1968 ; Kumar and Tieszen, 1980 ; DaMatta et al., 2018 ), consistent with its origin in the cooler montane rainforests of Ethiopia ( Kufa and Burkhardt, 2011 ). In contrast, B. acerifolius , native to tropical lowland forests (“CSIRO - Australian Tropical Rainforest Plants,” 2020) is better adapted to temperatures above 30°C. T. fortunei , a monocot palm, also showed a peak in net CO 2 flux at the beginning of the day, followed by a steady decline throughout the daylight hours. This trend may be partly explained by a continuous increase in meristem temperature, which is known to increase the growth rate in monocots ( Ben-Haj-Salah and Tardieu, 1995 ) and contribute to a steady increase in growth respiration. At the leaf-level, the net assimilation rate in the C 3 species (Fig. S3) were less consistent under conditions of 30°C and 60% RH over several hours, suggesting that the daytime flux variability may result from complex interactions between C source and sink activities and the environment, such as an increased sink activity due to slowly increasing soil and shoot temperatures ( Wang et al., 2006 ) and an increased availability of fresh assimilates during daytime hours increasing respiration ( Lai et al., 2016 ; Brauner et al., 2018 ). At night, all three species exhibited peak respiration at the start of the dark period, likely caused by residual heat and substrate availability following light-off ( Frantz, 2004 ). Thereafter, whole plant net CO 2 flux remained consistently negative and stable, indicating that plant C dynamics are more variable during the day than at night, which is in contrast to earlier studies in forbs where distinct patterns during the nighttime were found ( Gessler et al., 2017 ). This might either indicate that forbs regulate their respiration differently during the night, or that the nighttime C fluxes are coordinated on the whole-plant level. Measurements of the CAM plants ( Fig. 2b ) clearly captured all four classical phases of CAM photosynthesis, as described by ( Osmond, 1978 ). In phase I, during the night, stomata open to allow CO₂ uptake. In phase II, as the light switches on, stomata begin to close to prevent excessive water loss. However, during this transitional phase, both PEP carboxylase and Rubisco are active, allowing simultaneous CO₂ fixation via the CAM and C 3 pathways. This dual activity results in small, visible spike in net CO 2 flux at the beginning of the day (not observed in Opuntia ). In phase III, with stomata fully closed during most of the day, water loss is minimized, thus only respiratory (i.e., negative) CO 2 fluxes of heterotrophic tissues such as roots can be measured. Finally, in phase IV, in the late afternoon and absence of water stress, stomata reopen briefly, allowing additional CO₂ uptake via the C 3 pathway. This is reflected in the observed in net CO 2 flux late in the day. Following the transition to darkness, a sharp decline in net CO 2 flux can be observed, marking the metabolic shift from C 3 back to CAM pathway. Together, these results highlight distinct diurnal CO₂ exchange patterns in C 3 and CAM plants, emphasizing the PANDA chamber’s utility in resolving species- and pathway-specific gas exchange dynamics over time, such as cryptic or low level CAM photosynthesis ( Winter and Holtum, 2015 ; Surridge, 2019 ; Winter et al., 2019 ). Experiment 2: Circadian cycles in C 3 plants Species-specific responses emerged when the C 3 plants were exposed to extended periods of constant light or darkness ( Fig. 3 ). In B. acerifolius ( Fig. 3a ), net CO 2 flux stabilized over the three days of constant light, with the highest variability occurring on the first day. Under constant darkness, its net CO 2 flux steadily declined, becoming increasingly negative over time. In contrast, T. fortunei ( Fig. 3b ) displayed pronounced fluctuations in net CO 2 flux throughout the light period, with multiple sharp increases and decreases. This pattern might indicate an inherent cyclic growth rhythm in this monocot palm species, which becomes disrupted under prolonged light exposure. In C. arabica ( Fig. 3c ), the initially positive net CO 2 flux during constant light gradually declined over the three days, exhibiting a characteristic wavelike or cyclic pattern. This decline could reflect either a gradual reduction in photosynthetic activity due to sustained exposure to high light and temperatures above 30°C, or a progressive increase in sink demand driven by continuous assimilate supply. The latter interpretation suggests a relatively stable source-sink coordination, potentially governed by recurring episodes of elevated sink activity. Under constant darkness, net CO 2 flux in both B. acerifolius and C. arabica became progressively less negative, possibly indicating a depletion of non-structural carbohydrates ( Weber et al., 2018 ) and a corresponding decline in sink activity ( Sevanto et al., 2014 ; Collins et al., 2021 ). In contrast, T. fortunei showed no such trend, likely due to the mobilization of substantial non-structural carbohydrate reserves inside its stem ( Tomlinson, 1990 ). However, the absence of clear patterns in respiratory nighttime CO 2 flux is, as experiment 1, in contrast to earlier studies in forbs, where distinct patterns during the nighttime were found ( Gessler et al., 2017 ). Download figure Open in new tab Figure 3: The net CO 2 flux of the three tested C 3 species (n = 1 per species) when exposed to three to four consecutive days of constant light and darkness. The lighter grey marks indicate the hypothetical nighttime hours, while the larger, darker grey area on the right indicates the actual time in darkness. The red lines show the running means (10 measurements before and 10 measurements after each point, with one measurement every 2 minutes), and the grey lines show the measured values. These findings demonstrate the PANDA system’s capacity to resolve subtle, species-specific endogenous rhythms in CO₂ exchange, highlighting differences in source–sink coordination and carbohydrate storage strategies among C 3 plants under constant environmental conditions. Future work should investigate the circadian regulation of the whole-plant net CO 2 flux between different phylogenetic groups such as monocotyledons and dicotyledons, or angiosperms and gymnosperms, as well as different life forms, such as grasses, forbs, palms, and trees. Experiment 3: The net CO 2 flux over one month of germinating Quercus robur acorns Tracking acorn germination with the PANDA system ( Fig. 4 ) demonstrated its ability to monitor net CO 2 flux in growing plants continuously and accurately over more than a month. Mean daytime temperature and RH were 31.8°C and 96.6% (± 0.6°C and ± 2.6%, respectively), while nighttime values were 22.1°C and 100% (±0.4°C and ±0%). Net CO 2 flux (µmol s -1 acorn -1 ) ( Fig. 4a ) captured the emergence of diel patterns from germination on February 22 nd to the fully developed seedling on March 19 th , including the onset of a short drought treatment on March 21 st . Seedling development at selected time point is illustrated in Fig. 5 . Download figure Open in new tab Figure 4: The (a) net CO 2 flux in µmol s -1 acorn -1 per plant (12 replicates until March 3, 9 until March 7, and 6 until March 24) of germinating acorns of Quercus robur over one month, (b) the cumulative CO 2 balance in mmol acorn -1 during this period, and (c) the calculated resulting change in biomass based as cumulative C balance in mg per plant calculated by the measured CO 2 fluxes (red line) as well as the dry mass measured during the subsequent harvesting of three seedlings for the corresponding date (light blue boxplots). In (a), the light grey areas indicate the nighttime hours, and the yellow area indicates the days with reduced water access. The bold line in (a) shows the running means (10 measurements before and 10 measurements after each point, one measurement every 2 minutes), and in (b) and (c), the bold red line shows the actual values. In (c), the central horizontal line within each box represents the median of 3 harvested plants, while the box edges correspond to the 25 th and 75 th percentiles. Whiskers extend to data points within 1.5 × interquartile range. The black circle indicates the mean value for each harvest date. Download figure Open in new tab Figure 5: Time-lapse of the germinating Quercus robur acorns from the 25 th of February 2025 (3 days after sowing) until the 17th of April 2025, the day after the second round of temperature response curves. Early in the germination process, net CO 2 flux was consistently negative, indicating a fully heterotrophic metabolism (cf. Fig. 5 , Feb. 25). This also highlights the system’s sensitivity to below-ground respiration. Daytime fluxes were more negative than nighttime, likely due to ∼12°C higher daytime temperature. As root and shoot growth progressed, CO 2 flux remained negative – even after initial leaf emergence on March 3 rd , 5 th , and 7 th ( Fig. 5 ), likely due to high growth respiration. A sharp nighttime peak in negative flux developed at the onset of darkness, likely reflecting high tissue respiration at still high temperature, while a small positive peak appeared at lights-on, possibly due to cooler morning temperatures. Increasing photosynthesis led daytime net CO 2 flux turning positive on March 8 th . Cumulative CO 2 ( Fig. 4b ) and C ( Fig. 4c ) balances reached a turning point between March 8 th - and 10 th , after which it turned from negative to positive. After this, the strongly increasing photosynthetic activity ( Fig. 4a ) led to a progressively increasing positive net CO 2 flux during the daytime hours, while nighttime respiration remained did not increase at a similar rate. This led to a steep increase in cumulative CO 2 ( Fig. 4b ) and C ( Fig. 4c ) balance. By March 20 th , seedling growth finished and net CO 2 flux plateaued, a mild drought was induced. This did not impact the early morning peak of net CO 2 uptake ( Fig. 4a ) but decreased the net CO 2 uptake during the later hours of the day. This resulting midday depression of photosynthesis resembles field observations ( Muraoka et al., 2000 ; Koyama and Takemoto, 2014 ), though here it likely results from soil water limitation rather than photoinhibition. Finally, a comparison of cumulative C balance with dry biomass ( Fig. 4C ) showed strong agreement: the calculated C gain per seedling was 609 mg glucose equivalent, closely matching the 544 mg average of seedlings harvested on March 24. The slight offset likely reflects developmental variability ( Fig. 5 ) and sample size (i.e., only three per time point). Experiment 4: Shifts in whole-plant C balance with rising temperatures The net CO 2 flux responded strongly to increasing T air ( Figs. 6 , S4), with notable differences between the two experimental rounds. Between the two rounds, total leaf area increased by 595 cm 2 (from 873 cm 2 to 1468 cm 2 ), and leaf mass fraction (the percentage the leaf weight contributes to the total weight) rose from 15.7% to 34.4%. On the first day (average daytime T air of 27.6°C for both rounds), net CO 2 uptake was 4.4 times higher in the second round (1.15 µmol s -1 ) than in the first (0.26 µmol s -1 ). On the second day, daytime T air averaged 33.6°C in the first round and 31.8°C in the second. Correspondingly, the net CO 2 flux declined by 31% in the first round and 10% in the second ( Fig. 6b ). Despite this, flux on the second round (1.03 µmol s -1 ) remained 5.7 times higher than in the first (0.18 µmol s -1 ). On the third day, with average T air reaching 42.9°C and 42.3°C respectively, the net CO 2 flux dropped sharply in both rounds: to 14% of the original value in the first (0.04 µmol s -1 ) and to 30% in the second (0.35 µmol s -1 ), which was still 8.8 times higher than in the first. Download figure Open in new tab Figure 6: (a) The running means of the net CO 2 flux in µmol s -1 during the three days with increasing air temperature (T air ) of Quercus robur seedlings (all three seedlings combined) after their first (Round 1, blue, average daytime T air 27.6, 33.6, and 42.9°C) and second (Round 2, red, average daytime T air 27.6, 31.8, and 42.3°C) development stages finished, where negative values indicating a net CO 2 release whereas positive values indicate a net CO 2 uptake. Empty circles indicate the average net CO 2 flux during daytime, while triangles indicate average nighttime flux at room temperature (∼22°C). (b) The corresponding temperature response (blue = round 1, red = round 2, black = combined) of the mean daytime net CO 2 flux (straight line) and the diel-corrected net CO 2 flux (i.e., average daytime flux – average nighttime flux; dashed line) normalized to the mean of the first day with a 27.6°C, the quadratic fit for the mean of both rounds, as well as the temperature thresholds at which the net CO 2 flux turns negative (triangles for the diel-corrected, i.e. daytime – nighttime flux and circles for the daytime fluxes, number). The red and blue numbers indicate average values of the corresponding days and nights, respectively. Quadratic regressions models fitted to the normalized mean daytime net CO 2 flux ( Fig. 6b ) predicted the temperature at which net flux becomes zero to be 45.1°C (first round), 45.7°C (second round), and 45.6°C (combined data). When diel CO 2 flux was calculated by subtracting nighttime from daytime values ( Fig. 6b ), the flux turned neutral at 40.3°C, 43.9°C, and 42.1°C for the first, second, and combined rounds, respectively. However, these measurements were based on a single pseudo-replicate, and the temperature responses therefore overfitted. As such, these results should only be viewed as a proof of concept, illustrating the capabilities of this method with the PANDA system. Future experiments with appropriate replication are needed to robustly quantify how the net CO₂ flux responds to increasing temperature, and how this response is influenced by factors like plant size and leaf mass fraction. Conclusions We presented a replicable, low-cost chamber system – PANDA - that enables accurate, continuous, and reliable monitoring of whole-plant net CO₂ flux. This study demonstrates that the PANDA system provides a practical and flexible platform for studying plant C balance at the whole-plant level in a laboratory setting. Its application includes (1) monitoring diel dynamics of net CO 2 fluxes in both C 3 and CAM plants, (2) investigating circadian rhythms under prolonged exposure to light or dark conditions, (3) tracking the C balance during germination and early growth, and (4) assessing responses to environmental stressors such as drought and high temperature. PANDA’s modular design allows for easy adaptation to diverse experimental needs, such as scaling chamber size, integrating active temperature control, or situating the entire setup within a climate-controlled environment. We highlight the value of shifting from measurements isolated plant organs to whole-plant assessments, offering a more integrated perspective on plant C dynamics under changing environmental conditions. Given its simplicity, affordability, and reproducibility, the PANDA system holds strong potential to become a widely adopted tool for advancing our understanding of whole-plant responses to climate-related stresses, thereby contributing to improved predictions of plant behavior in future climates. Author contributions PS conceptualized the chamber and planned the experiments. PF and PS built the chamber system. PS, PP and YL conducted the experimental work. PS led the writing of the manuscript. YL, PP, PF, MHL, YLZ, and CG critically revised the manuscript. Conflict of interest All authors confirm that they declare having no conflict of interest. Funding PS, YL, PP, PF, and CG were supported by the Swiss National Science Foundation SNSF (310030_204697 and CRSK-3_220989) and the Sandoz Family Foundation. Data availability After publication, all raw data will be published on a publicly available respiratory ( www.envidat.ch ). 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Share PANDA: A simple and affordable chamber system for measuring the whole-plant net CO 2 flux Philipp Schuler , Yuxin Li , Pierre Pittet , Patrick Favre , Mai-He Li , Yan-Li Zhang , Charlotte Grossiord bioRxiv 2025.06.02.657330; doi: https://doi.org/10.1101/2025.06.02.657330 Share This Article: Copy Citation Tools PANDA: A simple and affordable chamber system for measuring the whole-plant net CO 2 flux Philipp Schuler , Yuxin Li , Pierre Pittet , Patrick Favre , Mai-He Li , Yan-Li Zhang , Charlotte Grossiord bioRxiv 2025.06.02.657330; doi: https://doi.org/10.1101/2025.06.02.657330 Citation Manager Formats BibTeX Bookends EasyBib EndNote (tagged) EndNote 8 (xml) Medlars Mendeley Papers RefWorks Tagged Ref Manager RIS Zotero Tweet Widget Facebook Like Google Plus One Subject Area Plant Biology Subject Areas All Articles Animal Behavior and Cognition (7624) Biochemistry (17651) Bioengineering (13871) Bioinformatics (41882) Biophysics (21424) Cancer Biology (18566) Cell Biology (25461) Clinical Trials (138) Developmental Biology (13365) Ecology (19867) Epidemiology (2067) Evolutionary Biology (24290) Genetics (15590) Genomics (22476) Immunology (17714) Microbiology (40331) Molecular Biology (17148) Neuroscience (88483) Paleontology (666) Pathology (2828) Pharmacology and Toxicology (4817) Physiology (7635) Plant Biology (15114) Scientific Communication and Education (2044) Synthetic Biology (4286) Systems Biology (9815) Zoology (2268)