Pressure, O2, and CO2, in aquatic Closed Ecological Systems

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Advances in Space Research 51 (2013) 812–824 www.elsevier.com/locate/asr

Pressure, O2, and CO2, in aquatic Closed Ecological Systems Frieda B. Taub ⇑, Anna K. McLaskey School of Aquatic and Fishery Sciences, University of Washington, Seattle, WA 98195, USA Received 28 March 2012; received in revised form 22 August 2012; accepted 20 October 2012 Available online 29 October 2012

Abstract Pressure increased during net photosynthetic O2 production in the light and decreased during respiratory O2 uptake during the dark in aquatic Closed Ecological Systems (CESs) with small head gas volumes. Because most CO2 will be in the liquid phase as bicarbonate and carbonate anions, and CO2 is more soluble than O2, volumes of gaseous CO2 and gaseous O2 will not change in a compensatory manner, leading to the development of pressure. Pressure increases were greatest with nutrient rich medium with NaHCO3 as the carbon source. With more dilute media, pressure was greatest with NaHCO3, and less with cellulose or no-added carbon. Without adequate turbulence, pressure measurements lagged dissolved O2 concentrations by several hours and dark respiration would have been especially underestimated in our systems (250–1000 ml). With adequate turbulence (rotary shaker), pressure measurements and dissolved O2 concentrations generally agreed during lights on/off cycles, but O2 measurements provided more detail. At 20 °C, 29.9 times as much O2 will distribute into the gas phase as in the liquid, per unit volume, as a result of the limited solubility of O2 in water and according to Henry’s Law. Thus even a small head gas volume can contain more O2 than a larger volume of water. When both dissolved and gaseous O2 and CO2 are summed, the changes in Total O2 and CO2 are in relatively close agreement when NaHCO3 is the carbon source. These findings disprove an assumption made in some of Taub’s earlier research that aquatic CESs would remain at approximately atmospheric pressure because approximately equal molar quantities of O2 and CO2 would exchange during photosynthesis and respiration; this assumption neglected the distribution of O2 between water and gas phases. High pressures can occur when NaHCO3 is the carbon source in nutrient rich media and if head-gas volumes are small relative to the liquid volume; e.g., one “worse case” condition developed 800 mm Hg above atmospheric pressure and broke the glass container. Plastic screw cap closures are likely to leak at high pressures and should not be assumed to seal unless tested at appropriate pressures. Pressure can be reduced by having larger head-gas volumes and using less concentrated nutrient solutions. It is important that pressure changes be considered for both safety and closure, and if total O2 is used as the measure of net photosynthesis and respiration, the O2 in the gas phase must be added to the dissolved O2. Ó 2012 COSPAR. Published by Elsevier Ltd. All rights reserved. Keywords: Closed Ecological Systems; Algae; Daphnia; Photosynthesis; Respiration; Henry’s Law

1. Introduction Closed Ecological Systems (CESs), open to energy exchanges of light and heat but closed to material exchanges, allow us to study the two way feedbacks between biota and their local environment, in contrast to more traditional open systems in which the biota’s effects are mingled with atmospheric exchange. CESs are the only systems that allow us to study how a community of organ⇑ Corresponding author. Tel.: +1 206 484 1229; fax: +1 206 685 3275.

E-mail address: [email protected] (F.B. Taub).

isms, when restricted to their initial material recourses, will organize themselves and control their O2 and CO2 environment. Motivation for studying CESs has ranged from the potential of biologically controlled life support systems to basic research of how ecological communities function. The history of aquatic CESs is modest; early studies were summarized by Taub (1974) and some more recent work is reviewed in the discussion section. A goal of this laboratory has been to study the O2, CO2, and population dynamics in simple aquatic ecosystems consisting of a chemically defined medium, primary producers (algae), grazers (Daphnia) and associated microbes

0273-1177/$36.00 Ó 2012 COSPAR. Published by Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.asr.2012.10.016

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during light:dark cycles for periods of several weeks. Inherent in the goal was the measurement of properties that would represent the total ecosystem in the simplest way possible. Because of the size of the grazers (Daphnia magna, 0.7–3 mm), systems of 250–1000 ml have been used for our population studies. In earlier work with 259 ml aquatic CESs in our laboratory, the simplifying assumption was made that the CESs would remain at approximately atmospheric pressure because approximately equal molar quantities of O2 and CO2 would exchange during photosynthesis and respiration (Taub, 2009a,b, 2011). This assumption neglected to account for the limited solubility of O2 in water and ignored the O2 in the gas phase. As a result of this assumption, it was assumed that the seals were adequate to prevent leakage. However, when NaHCO3 was the carbon source, much more CO2 was calculated to have been removed than O2 produced; the differences were minor if cellulose or no-added carbon were tested. The lack of agreement between estimates of O2 and CO2 prompted a reevaluation of the assumption of pressure remaining near atmospheric pressure and leakage being unimportant. Although preliminary experiments suggested the CESs were not leaking, these tests have now been shown to have been inadequate. In contrast, Obenhuber (1986) and Obenhuber and Folsome (1988) working with marine microbial communities in 20 ml vials containing 15 ml liquid and 5 ml head gas space, used pressure changes as a surrogate for O2 changes. He stated, “The accumulation of reduced carbon in a closed system must produce gaseous oxygen due to the low solubility of oxygen in water. This increased partial pressure of oxygen will result in an increase in pressure within the container” (Obenhuber, 1986). He ignored the increased O2 dissolved in the water as being less than 5% of the total, and therefore treated it as negligible. He used the changes in pressure during light:dark cycles to estimate net photosynthesis, respiration, and biomass. His studies used NaHCO3 as a carbon source (2 mg/L or 23.8 mM), as did others using his pressure technique (Shaffer, 1991). Because his ecosystems’ pressure exceeded the sensitivity of his pressure monitors (1/3 atmosphere), he periodically opened his ecosystems and summed the total pressures obtained. He did not report lags between pressure and light–dark changes in his small units. His use of pressure to estimate O2 net production and respiration was ingenious, given that at that time stable O2 sensors (such as current optical O2 sensors) did not exist. The purpose of this paper is to test the assumption of gas pressure and to demonstrate considerations that must be taken into account when measuring O2 and/or CO2 in aquatic Closed Ecological Systems. Unlike the 20 ml systems studied by Obenhuber, the larger systems reported here (up to 1000 ml) had significant discrepancies between O2 and pressure unless turbulence was significant. Also, the implications of the Henry’s Law distribution of O2 in the gaseous phase and dissolved phase are evaluated; to assess

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the total O2 changes, the gaseous and dissolved O2 should be summed. 2. Methods 2.1. Preliminary experiments The accumulation of pressure was tested by injecting hypodermic syringes (Multifit B-D Brand) into 259 ml bottles completely filled with medium (no head-gas volume) inoculated with algae, and laid on their sides to allow free movement of the syringe as gas accumulated. Subsequent tests were done in plastic “saline bags,” (EXCELÒ Container, B. Braun Medical Inc., Irvine, CA USA 92614-5895, empty and rinsed) which allowed pressure accumulation to expand the bag as gas bubbles formed, filled with 1000 ml of medium (B + 13.2 mM NaHCO3 and B/4 + 3.3 mM NaHCO3 as well as B, B/4, and B/16 with cellulose) inoculated with algae. 2.2. Differential pressure (Stationary) The measurement of pressure inside bottles was initially accomplished by drilling holes through the plastic and metal caps used and inserting a rubber serum stopper connected to a digital manometer (Model 8230 Mannix Testing & Measuring with range of 0–30 psi). These manometers measure differential pressure, i.e., deviation from atmospheric pressure. Atmospheric pressure was estimated from the University of Washington Department of Atmospheric Sciences web site: http://www.atmos.washington.edu/data/data.php?loc=local. Testing of caps to hold pressure was done by injecting air with a hypodermic syringe into the bottle through the rubber serum stopper and monitoring the decline in pressure over time. 2.3. Multi-probe experiments: pressure and O2 measurements with and without turbulence The multi-probe equipment was designed for a CES to screw on over the water sensors, allowing in situ measurements while maintaining a leak-proof seal at the pressures that these CESs develop. Because there is only one multiprobe, only one CES can be monitored at a time. There are no replicates so statistical analysis isn’t done. However, several separate experiments have been run on the multiprobe, each showing similar relationships. Initial testing of the equipment compared having the unit stationary, with a magnetic mixer inside (VWR Scientific Products, 375 Hot Plate and Stirrer with Teflon covered tissue culture stirrer magnet), and on a rotary shaker (Innova 2300 set at 110 rpm). Based on these tests our next six experiments using the multi-probe were all on the rotary shaker. B/4 with NaHCO3 as a carbon source, cellulose as a carbon source, and no-added carbon source was tested with the grazer Daphnia present and without Daphnia present.

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Detailed data presented here is B/4 with NaHCO3 as a carbon source with Daphnia and algae. 2.4. Organisms The organisms included three species of algae: Ankistrodesmus sp., Scenedesmus obliquus (previously known as Scenedesmus acutus), and Selenastrum capricornutum (also known as Pseudokirchneriella subcapitata and Raphidocelis subcapitata) all cultured in T82 medium (ASTM, 2012). Some CESs contained the grazer Daphnia magna, originally obtained from M.T. Brett, University of Washington, and cultured on B/4 media. All algal and Daphnia cultures contain microbes associated with them. Daphnia abundances were counted by eye and algal abundance was estimated from three in vivo fluorescence (Fm) measurements on PSM MarkII Plant Stress Meter (BioMonitor S.C.I. AB with a run time of 5 s, light level 4). 2.5. Media The B/4 medium has been previously published (Taub, 2009a) and B medium is four times as concentrated; these are modifications of the medium used in the Standardized Aquatic Microcosm protocol (ASTM, 2012) augmented with trace metals and vitamins that are required by Daphnia and not produced by the algae; complete media components are listed in Table 1. The B medium was 0.5 mM N, 0.04 mM P and with a carbon source of either 13.2 mM NaHCO3, 0.5 g/L cellulose, or no-added carbon. The B/4 was 0.125 mM N, 0.01 mM P, with a carbon source of 3.3 mM NaHCO3, 0.5 g/L cellulose, or no-added carbon. For B/16, C, N, and P were reduced by 1/4, but the other chemicals were maintained at B/4 so that osmolarity and conductivity were similar to B/4. The series of multiprobe experiments used B/4 with 3.3 NaHCO3, 0.5 g/L cellulose or no-added carbon. The ecosystems are inoculated with 10% by volume algal suspension, reducing the initial carbon concentration (e.g., 3.3 mM NaHCO3 to 3 mM and the alkalinity titrates at 3000 uM). At the time the algae are inoculated, most of the N and P are in the algal cells, and not in the liquid medium. Adding algal culture by volume insures chemical inputs are the same whether the N and P are in the spent medium or in the algal cells; as a consequence, the initial algal cell densities can vary slightly between experiments. 2.6. Incubation conditions CESs were incubated in a temperature controlled chamber maintained at 20° ± 1° C with 12:12 h light:dark cycles, unless stated otherwise. In a few cases, 6:18 or 18:6 h cycles were used. Light (fluorescent tubes) conditions were typical for incubators and ranged from 5.5 to 10.0 W/m2 from above and 1.5–2.64 W/m2 from the sides. Because bottles with opaque lids receive most of their light from the side,

top and side intensities are given. (Light intensity for the syringe experiment was 7.59 W/m2 from above and 1.94 W/m2 from the sides. For saline bags with NaHCO3 it was 5.49 W/m2 from above and 1.5 W/m2 from the sides. For saline bags with cellulose it was 9.86 W/m2 from above and 2.93 W/m2 from the sides. For the manometer experiments it was 5.49 W/m2 from above and 1.82 W/m2 from the sides. Light Intensity for multi-probe experiments was 7.02 W/m2 from above and 2.64 W/m2 from the sides.) 2.7. Glass bottles Glass bottles were Clear French Square neck finish 43400, Qorpak GLA-00832, nominal volume 240 ml, total volume 259 ml with either White Metal Caps with Plastisol Liner, neck finish 43-400, Container & Packaging Supply L018, or Black Ribbed Polypropylene caps, Qorpak CAP-00381 with Natural Teflon PTFE 0.010” discs, Qorpak CAP-00527. Larger CESs used Clear French Square neck finish 58-400, Qorpak GLA-00835, nominal volume 960 ml, total volume 1000 ml, with White Metal Caps with Plastisol Liner, Qorpak CAP-00454. 2.8. Multi-probe equipment The multi-probe was custom manufactured by Eureka Environmental Engineering (2113 Wells Branch Parkway, Suite 4400, Austin Texas 78728) and consisted of the Manta2 Multiprobe 4.0 in. body, fitted with a Temperature sensor, Central Cleaning Systems (Standard McVan Wiper), pH sensor Assembly, Reference pH Assembly, Low Range Depth Sensor (Pressure), M2 Optical HDO Sensor Assembly (O2), M2 Conductivity Sensor (Specific Conductance), M2 Chlorophyll Sensor (in vivo fluorescence), and a Custom Manta2 External Battery Cable (to connect to line power while logging and not running from computer). A clear plastic (acrylic) sampling chamber of l L volume was custom made to seal to the instrumentation by a screw fitting and “O” ring with a trace of grease (Fig. 1). The pH reference electrode had restricted flow to minimize KCl leakage into the chamber. The chlorophyll sensor was set at the least sensitive setting. (However, because much of the algae settled on the bottom, the chlorophyll was also measured weekly with a plant stress meter, reported here). The pressure sensor is sealed and does not respond to changes in atmospheric pressure. All sensors are calibrated at the beginning of an experimental run and at the end (to check for stability). The equipment was tested to be non-toxic to algae and Daphnia used in these experiments. To seal the sensors to the clear chamber, the sensor and chambers are screwed together tightly, and checked and retightened after an hour. As a result, the chambers start at a pressure slightly greater than atmospheric. Note that the sensors are not at exactly the same depth in the chamber (Fig. 1). Day 0 starts at the lights on (5 AM) of the next day.

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Table 1 Chemical composition of B and B/4 media. Values for vitamins and NaOH are corrected from (Taub, 2009a). Compound

Molecular weight

mMol per liter

Element

B medium mM

B/4 medium mM

mg element per liter B medium mg/L

B/4 medium mg/L

Medium B and B/4 for Closed Ecological Systems Carbon source (variable) NaHCO3 84 Or other carbon source Cellulose 0.5 g/L (C6H10O5)n subunit MW 162, 44% C

13.2

3.3

C

158.4

39.6

3.1

3.1

C

220.0

220.0

Major salts

mM

mM

mg/L

mg/L

0.5000 0.1000 0.0400 0.0320 1.0000 1.5000 0.0048 0.0800

0.1250 0.0250 0.0100 0.0080 0.2500 0.3750 0.0012 0.0200

7.0000 2.4300 1.2300 0.7400 40.000 34.500 0.2600 3.6800 2.2400

1.7500 0.6075 0.3075 0.1850 10.000 8.6250 0.0650 0.9200 0.5600

uM

uM

mg/L

mg/L

5.6000 7.1000 3.7500 0.1250 1.2500 0.1250 0.0250 0.0125

1.4000 1.7750 0.9375 0.0313 0.3125 0.0313 0.0063 0.0031

Fe EDTA B Zn Mn Mo Cu Co

0.3125 2.0730 0.0400 0.0075 0.0675 0.0120 0.0016 0.0008

0.0781 0.5183 0.0100 0.0019 0.0169 0.0030 0.0004 0.0002

mg/L

mg/L

Li Rb Sr Br I Se V

0.050000 0.050000 0.050000 0.013000 0.002500 0.001000 0.000500

0.012500 0.012500 0.012500 0.003250 0.000625 0.000250 0.000125

mg/L

mg/L

0.000500 0.000500 0.100000

0.000125 0.000125 0.025000

NaNO3 MgSO47H2O KH2PO4 NaOH CaCl22H2O NaCl Al2(SO4)318H2O Na2SiO39H2O

85.0 246.5 136.0 40.0 147.0 58.5 666.5 284.0

Trace metals FeSO47H2O EDTA H3BO3 ZnSO47H2O MnCl24H2O Na2MoO42H2O CuSO45H2O Co(NO3)26H2O

278.0 292.0 61.8 287.5 197.9 242.0 249.7 291.0

Additional trace metals for Daphnia

uM

uM

LiCl RbCl SrCl2 6H20 NaBr KI H2SeO3 Na3 VO4

7.3000 0.5800 0.5700 0.1600 0.0200 0.0130 0.0097

1.8250 0.1450 0.1425 0.0400 0.0050 0.0033 0.0024

uM

uM

3.69E04 2.05E03 2.96E01

9.22E05 5.12E04 7.41E02

42.4 120.9 266.6 102.9 166.0 129.0 183.8

Vitamins for Daphnia B12 (cyanocobalamin) Biotin (d-biotin) Thiamine (HCl)

1355.4 244.3 337.3

N Mg P Na Ca Na Al Na Si

B12 Biotin Thiamine

½AðaqÞ ½AðgÞ activity; fugacity; or concentration in aqueous phase ¼ activity; fugacity; or concentration in gaseous phase ð1Þ

2.9. O2 conversion of measured O2 measurements to total system O2

H¼

The O2 sensor measured the dissolved O2 in mg/L; these values were converted to lmoles/L by multiplying by 1000/ 32. The distribution between the liquid and gaseous phases at equilibrium is described by Henry’s Law, with the factors KH and H being commonly used (Stumm and Morgan, 1996). At 20° C there will be 29.9 times as much O2 in a liter of gas in equilibrium with a liter of water (see calculation below). Stumm and Morgan (1996) define the Henry’s law equilibrium constant H as:

In these systems, O2 is measured in the aqueous portion and O2 in the gaseous phase can be estimated by multiplying the dissolved O2 by 1/H (Eq. (2)). 1 ½AðgÞ ¼ H ½AðaqÞ

ð2Þ

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Drawn To Scale; length, not location of probes, is accurate.

Temperature

Distance from base plate to end of each probe:

Depth

Conductivity pH

Temp

Depth

Condt

Reference

pH

pH

Chlorophyll

Oxygen

oxygen: 7.5 cm chlorophyll a 7.5 cm pH Reference: 6 cm pH 5.25 cm Temperature 3.5 cm Depth 3.75 cm Conductivity 3.75 cm

Chlorophyll

pH Reference Oxygen

Liquid volume: 850 ml Head-gas volume: 90 ml

Fig. 1. Multi-probe instrumentation. The vertical diagram indicates the liquid volume (850 ml), head-gas volume 90 ml) and the length of the sensors. The circular diagram shows the placement of the sensors. The equipment has been tested in stationary position, with a magnetic mixer, and on a rotary shaker. The whole unit is 48 cm tall with 10.5 cm diameter; clear cylinder is 20 cm tall.

The vapor pressure of water at 20 °C = 17.535 mm Hg (CRC Handbook of Chemistry and Physics, 2011). To calculate the% O2 in wet air:

0:008496 mol O2 =L  32 g=mol O2

760  17:535  0:2092 ¼ 0:20437ðfrac:O2 in wet airÞ 760

At 20 °C, there are 0.009092 g O2/L water (Wetzel and Likens, 2000). Ratio of gaseous/aqueous:

ð3Þ

At 20 °C and 760 mm Hg, 1 mol gas = 24.055 L so 1 L gas = 0.04157 mol. To calculate the moles of O2 in 1 L of wet air:

¼ 0:271875 g O2 =lgas

0:271875 g O2 =L gas ¼ 29:903 0:009092 g O2 =L aqueous

ð5Þ

ð6Þ

0:04157 mol=L  0:20437 ðfrac: O2 in wet airÞ ¼ 0:008496 mol O2 in L wet air Converting to grams O2/liter gas:

ð4Þ

The Yaws’ Handbook of Properties for Environmental and Green Engineering (Yaws, 2008) gives the Henry’s Law constant for O2 in water at 293 K as 39890 atm/mol

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frac, which when converted to a ratio of concentrations (http://www.epa.gov/athens/learn2model/part-two/onsite/ henryslaw.html) is 29.9. The Henry’s Law Constant represents the distribution of oxygen per unit volume and must be applied to each specific system. For example, our multi-probe systems are 850 ml liquid and 90 ml head gas space, therefore total O2 = (0.85 L  dissolved O2 lmol/L) + (0.09 L  29.9  dissolved O2 lmol/L) or, factoring out dissolved O2 lmol/L, Total O2 = dissolved O2 lmol/L (0.85 L + (0.09 L  29.9) = dissolved O2  3.541.

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between O2 concentration and pressure in single replicate multi-probe experiments were run using Excel 2007 Data Analysis Toolpak. The regressions between O2 concentration and pressure are given for three different experiments: B/4 with NaHCO3 as a carbon source, B/4 with cellulose, and B/4 with no-additional carbon source, each with Daphnia. 3. Results 3.1. Critical re-testing of atmospheric pressure in aquatic CESs

2.10. CO2 calculations Measured pH values were converted to carbonate species concentrations using the CO2_sys model for Excel (Pierrot et al., 2006). This model uses two known CO2 parameters, in our case pH and total alkalinity, titrated using the Gran function according to Wetzel and Likens (2000). The “freshwater” option was used, which supplies parameters for distilled water, whereas the freshwater media used here are more concentrated than distilled water but much less concentrated than sea water (the other option). The measured pH and calculated TCO2, CO2, HCO3, CO32, and OH for the NaHCO3 experiment with Daphnia are shown in the results as P lmol/L. (In some formats TCO2 would be expressed as CO2). The gas–liquid distribution is defined by Henry’s Law Constant, which at 293 K is 1423 atm/mol frac (Yaws, 2008) which when converted to a ratio of concentrations air:water of 1.07 (http://www.epa.gov/athens/learn2model/ part-two/onsite/henryslaw.html). Note that only dissolved CO2 (not HCO3 nor CO2 3 ) will enter into equilibrium with the gas phase. At high pH only a small proportion of the Dissolved Inorganic Carbon (DIC) is dissolved CO2. If pH is high, very little of the DIC is CO2, most is HCO3 and CO32. Also note that the Henry’s Law constant (air:water) for CO2 is much smaller than that of O2. The Henry’s Law Constant represents the distribution of CO2 per unit volume and must be calculated for the specific gas volume, in our case 0.09 L. Since only the dissolved CO2 portion of the DIC will be in equilibrium with the air, 0.09  1.07  CO2 = lmol will be in the gas phase. However, the full TCO2 in the liquid must be considered in calculating the change in TCO2 during photosynthesis and respiration, 0.85 L DIC. The Total CO2 of the CES = (0.09  1.07  dissolved CO2) + (0.850  TCO2 in the liquid).

Although earlier testing in our laboratory appeared to support our original assumption that CESs remained near atmospheric pressure, three series of more rigorous tests demonstrated their shortcomings. (1) Hypodermic syringes connected via needles injected into the head-gas space or liquid of aquatic CESs in a vertical (upright) position had not increased in volume; this had been interpreted as pressure not having increased. However, when 259 ml bottles were completely filled with medium (no head-gas volume), and the needles and 10 ml syringes were tested in a horizontal position, the barrels of the syringes did extend (10–3.4 ml), and the horizontal position of the bottle allowed us to observe the development of gas bubbles against the top surface. In a vertical position, gas bubble formation was hidden by the black plastic cap, and the weight of the syringe barrel and friction between the barrel and outer wall prevented the extension of the syringe. (2) Flooding the incubator with N2 gas did not result in measurable changes of O2 within 259 ml aquatic CESs, and this had been interpreted as indication that the seals prevented exchange between the aquatic CESs contents and the incubator atmosphere. However, this test was preformed with a N- and P- free solution and in the absence of algae, and the aquatic CESs were at atmospheric pressure during the one day test. (3) Plastic “saline bags” of B and B/4 with NaHCO3 as the carbon source developed 32.2 and 9 ml of gas, respectively, by day 12, and 52 and 11 ml by day 21. When B, B/4, and B/16 with cellulose as a carbon source were inoculated with algae, they developed gas bubbles more slowly; only 1–2 ml on day 35, but by day 126 had developed 17, 9.3 and 6.3 ml respectively. As a result of these preliminary studies, the assumption that the aquatic CESs remained at approximately atmospheric pressure was rejected, and it was decided to measure pressure changes.

2.11. Statistics

3.2. Differential pressure changes with B and B/4 medium; no turbulence

The lack of equipment permitting simultaneous measurements of the response variables in more than one experimental unit prevented the establishment of experiments with true replicates and thus the variability of the described phenomena could not be estimated. Regressions

The measurement of head-gas pressure by a manometer connected to the head-gas space in glass bottle CESs provided increasing pressures, but they were not tightly related to the light–dark cycle (Fig. 2). Pressures increased during the 12 h lighted phase but continued to increase for some

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The plastic screw lined caps without holes, and with Teflon liners (used for non-instrumented CES) leaked at pressures exceeding 25–50 mm Hg. Metal caps with plastisol liners appear to hold pressure up to 200 mm Hg for several days. These latter two values are minimum estimates because holes had to be made in the caps to attach the manometers, and these holes may have contributed to leakage. Still, it is unsafe to assume caps hold pressures unless tested for multiple days to weeks and at the maximal pressures likely to develop.

300

250 B/4 with NaHCO3

Pressure (mmHg)

200

B with NaHCO3

150

100

50

3.4. Multi-probe: simultaneous measurements of pressure and O2 (effects of turbulence)

0 0.0

1.0

2.0

3.0

4.0

-50

Days Sealed

Fig. 2. Pressure changes during 12:12 h L:D cycles in 247 liquid + 12 ml head-gas volumes in stationary aquatic CESs with either B + 13.2 mM NaHCO3 or B/4 + 3.3 mM NaHCO3. The dark bars represent dark periods. Note that there were lags before the pressure started to increase during the lighted periods and pressures continued to increase during the beginning of the dark period, and showed little if any decrease (dark respiration). These experiments were not run simultaneously.

hours after darkness, and eventually approached being constant, but rarely decreased during the dark phase. At the beginning of the 12 h lighted period the pressure began to increase slowly after a brief lag, and then increased at a faster rate. The more concentrated medium (B with 13.2 mM NaHCO3) increased pressure faster than the less concentrated medium (B/4 with 3.3 mM NaHCO3), but the pattern of increasing pressure well into the dark phase was similar. It was difficult to estimate rates of photosynthesis and respiration from these patterns; had we done so, we would have slightly underestimated net photosynthesis and grossly underestimated dark respiration. To further test the ability of CESs to develop pressure, highly concentrated medium (B + 13.2 mM NaHCO3) was inoculated with the algal mixture in a 500 ml bottle with 12 ml of head-gas volume and 488 ml of medium, pressure reached 1100 mm Hg above atmospheric pressure. In a 1 L bottle with 12 ml of head-gas space and 988 ml of medium, the glass broke at 800 mm Hg above atmospheric pressure. 3.3. Closure studies of caps Given the recognition that CESs can develop substantial pressures, especially with NaHCO3 as the carbon source, the various caps and seals were tested to determine leakage. In earlier equipment (Data Acquisition Device), plastic screw cap lids with Teflon liners were used in the noninstrumented units, and rubber serum stoppers holding O2 and pH sensors through holes in the caps were used in instrumented units. The caps with holes for sensors (instrumented) leaked at 10 mm Hg pressure, but these stoppers had been in use more than 5 years, and were worn.

The multi-probe equipment was the first to provide simultaneous measurements of pressure, O2 concentration, pH and other attributes. The act of closure increases the internal pressure 40–60 mm Hg above atmospheric. In stationary cultures (using B/4 + 3.3 mM NaHCO3 and algae, no Daphnia) O2 increased each lighted period and continued to increase each dark period (Fig. 3A). With a magnetic mixer, the O2 changes still corresponded to the light:dark cycle and the pressure readings showed more relationship to the light cycles, but there were lags so that pressure continued to increase at the start of the dark period, and there were only small decreases during the dark phase (Fig. 3B). In overnight experiments, the magnetic mixer killed Daphnia, so it would not have been suitable for experiments with these grazers. The following rotary shaker experiment included Daphnia. With the multi-probe on the rotary shaker (Fig. 3C) the O2 and pressure changes were in much greater agreement but the O2 trace shows more detail than the pressure (e.g., the O2 data suggest net photosynthesis and dark respiration are at faster rates immediately after the light change, whereas this does not show up in the pressure measurements). Further evidence of the importance of turbulence (agitation) is shown in the change of data when the rotary shaker was accidently left off for slightly more than a 24 h period (Fig. 3D); O2 continued to conform to the light:dark cycle with some irregularities, but the pressure showed only modest changes, not conforming to the light:dark cycles. (Note that during this time period, we were varying the light:dark cycles from 12:12 to 6:18 h.) Regression analysis showed the relationship between O2 concentration (mg/L) and pressure (mm Hg) was greater on the rotary shaker (Days 0–10, R2 = 0.997, n = 2821), than with a magnetic mixer (Days 8–18, R2 = 0.775, n = 2870) or stationary (Days 0– 10, R2 = 0.865, n = 2853). Separate, single replicate experiments were run using B/ 4 with NaHCO3 as a carbon source, cellulose, and noadded carbon source, all with Daphnia, on the rotary shaker. They all showed similar O2 and pressure regression relationships. With NaHCO3 as a carbon source, over the course of the 60 day experiment, R 2 = 0.997, n = 16,601. With cellulose over 76 days, R 2 = 0.961, n = 20,125 and no-added carbon over 34 days R2 = 0.932, n = 9674.

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250

25

200

20

200

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In the first few uses of the acrylic chamber with algae only, the plastic surface became covered with tiny bubbles, presumably of O2. With use, and multiple washings, these bubbles did not form. Had we seen bubbles in our earlier studies, we would have suspected super-saturation of O2, but apparently bubbles only form if the surface has high surface tension, and this decreases with use and washing.

inorganic carbon) and HCO3 decreased, CO2 3 increased, and OH started to increase (Fig. 3C). After day 14, when grazers were abundant the pattern was reversed, TCO2, and HCO3 increased while CO2 and OH decreased. 3 Note that free CO2 was always very low in concentration. When the total O2 produced (liquid + gas phase) and the CO2 consumed are compared (Fig. 4D) the agreement is good, but not exact.

3.5. Multi-probe; mixed (shaker) study of algae, Daphnia, and microbes Algae increased for the first two weeks, until Daphnia populations increased and their grazing reduced the algal population (Fig. 4A). As long as the rotary shaker was on, the pressure and O2 measurements agreed closely (Fig. 4B); the only exception was during a period when the rotary shaker was accidently left off (data also shown in Fig. 3D). The pH started at above eight, and increased during the first two weeks while photosynthesis dominated the processes; during this time, TCO2 (total dissolved

3.6. Effects of carbon sources on pressure (multi-probe measurements with rotary shaker) With B/4 medium, NaHCO3 caused the highest pressures to develop (>250 mm Hg above atmospheric pressure), while the addition of cellulose as the carbon source or no-added carbon source caused lower pressures to develop (slightly exceeding 100 mm Hg above atmospheric pressure, including the 50 mm Hg from closure of the ecosystems; Fig. 5). These experiments were with algae

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and Daphnia, and represent the balance between primary production and grazing.

In aquatic CESs there are two mechanisms that contribute to changes in O2 having greater influence than changes in CO2 on head-gas volume and pressure. First, of the DIC, only the dissolved CO2, not the HCO3 nor CO2 3 , will come into equilibrium with the gas phase. At high pH, there is very little dissolved CO2, most of the DIC is HCO3 and CO32; in the multi-probe experiment shown here, Fig. 3C, pH was always >8.3, and CO2 was very small. In some other experiments where pH was lower at times, CO2 was more significant and its contribution to the gas phase is important. Second, the distribution between the gas:water phases, as expressed by Henry’s Law, is smaller for CO2, 1.07 as compared to 29.9 for O2 . The results clearly show that the aquatic CESs did not remain at  atmospheric pressure if NaHCO3 at 13.2 or 3.3 mM was used as the carbon source, and if sufficient N, P, and other elements were present for algal growth (media B and B/4). Pressure can be expected to increase

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significantly because most of the O2 produced by photosynthesis will be in the gas phase, 29.9 times as much as compared to an equal volume of water. The O2 produced under these conditions (B or B/4 medium with NaHCO3) in earlier publications (Taub, 2009a,b, 2011) would have been much underestimated because (1) the high pressures would have been likely to cause leakage with the serum caps holding the O2 and pH electrodes (2) head-gas would have been lost when the caps with electrodes were removed weekly to check and re-calibrate the Clark-type O2 electrodes, and (3) the O2 in the head-gas volume was ignored in the mistaken belief that its 12 ml was trivial compared to the 247 ml of culture. This would be an adequate explanation for the disagreement between the O2 produced and the CO2 taken up in those studies. Other data for those studies appear valid (e.g., in vivo fluorescence, Daphnia counts, pH, DIC dynamics). With cellulose or no-added carbon source, pressure increases were much less, but could have still compromised the seals. So published data using these conditions are closer to being correct, but leakage cannot be excluded. Also, since most of the O2 will be in the head-gas phase, it should be included in calculations; this was not done in the 2009 and 2011 studies (Taub). The development of stable dissolved O2 sensors (optical sensors) have made possible the monitoring of O2, pH, pressure, temperature, conductivity, and in vivo fluorescence without the need to check O2 sensors weekly. This instrumentation is expensive, and has precluded our using replicates for these measurements. We have used replicates in non-instrumented bottles with metal caps (to be reported later). The use of pressure as a surrogate for O2 production and uptake must be used with caution. It was apparently successful with 20 ml vials in the work of Obenhuber (1986), Obenhuber and Folsome, (1988), because his results (Fig. 3 in his paper) did not appear to show an increase after lights were off. However, mixing was necessary in our 250 and 1 L volumes. In the absence of adequate mixing (e.g., stationary or magnetic mixer) the multiday trend is similar between O2 production and pressure, but within the light:dark diurnal cycle, the O2 production could be slightly underestimated, and the dark respiration would be extremely underestimated. One mechanism might be that the production and use of O2 occurs largely near the bottom of the liquid, but the pressure changes (also measured in the liquid) do not become measureable until the O2 is transferred to the gas phase. Without adequate turbulence, the transfer of excess O2 from liquid to air during net photosynthesis requires several hours and the reverse transfer from the air to the liquid during dark respiration is even slower. The differences between the Total O2 produced and the Total CO2 removed (Fig. 4D) can be caused by a number of processes: (1) the biomass formed may be rich in lipids which have a lower O2 content per C than carbohydrates or protein (2) the O2 measurements may not be exact because all the solubility units assume distilled water, and

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the freshwater medium is not distilled water (3) the estimates of DIC may be inexact because the CO2 model uses dissociation constants for distilled water (4) our estimates of alkalinity (used in the CO2 model) may be slightly off because our technique has rather high variability, and (5) we assume that alkalinity remains constant during the course of the experiment, whereas a few researchers have reported alkalinity to vary (Goldman and Brewer, 1980; Uusitalo, 1996). Experimenters must be aware of the likely increases in pressure and be certain that their apparatus can sustain closure; ignoring pressure increases the risk of leakage. For example, our use of concentrated B medium + 13.2 mM NaHCO3 with 250 ml of liquid medium and only 12 ml of head-gas volume (as in our earlier research) was unwise. To avoid excessive pressure and the likelihood of leakage, the head-gas volume should vastly exceed the O2 volume likely to be produced. We are now using 200 ml medium in 259 ml bottles or 800 ml medium in 1000 ml bottles, and are not using medium more concentrated than B/4. If leakage of the gas phase occurs, much more O2 will be lost than CO2 because of the limited solubility of O2 in water as expressed by the Henry’s Law constant of 29.9 (ratio of air:water at 20 °C). For CO2, only dissolved CO2, not the HCO3 and CO2 3 , will be in equilibrium with the gaseous phase, and the Henry’s Law constant of 1.07 (ratio of air:water at 20° C) means that a smaller proportion of the CO2 will be in the gas phase than O2. It is useful to estimate the pressures likely to occur. The increase in pressure will be approximately proportional to the O2 volume produced by net photosynthesis (minus the 3% that will remain in solution, which is being ignored for these calculations). The usual assumption is that 50% of the bicarbonate will be available for photosynthesis and the other half will accumulate as carbonate: 2 HCO3 M CO2 + CO32 + H2O. In the modeled distribution of the DIC (Fig. 4B) almost half of the TCO2, supplied as bicarbonate, disappears. In the following examples, we are assuming 50% of the bicarbonate to be available for photosynthesis and 1 mM of O2 to be produced for each mM bicarbonate involved in photosynthesis. For example, if B/4 + 3.3 mM NaHCO3 is used as medium in a container of total volume 259 ml, 200 ml of medium and 59 ml headgas volume, then 0.33 mmol of O2 could be produced (3.3 mM/2  200 ml/1000 ml = 0.33 mmol O2). To convert this additional gas to pressure, we use the Ideal Gas Equation (Chang, 2005) PV = nRT, or for our use, P = nRT/V, where P is pressure in atm (760 mm Hg/atm), volume = liters, n = moles of gas, T = Temperature in Kelvin (293.15 K for 20° C) and R, the gas constant is 0.082057 L  atm/K  mol. So the additional 0.33 mmol (0.00033 mol) of O2 introduced into the 59 ml (0.059 L) head-gas volume will cause the pressure to increase 102 mm Hg above atmospheric pressure, i.e. (0.00033 mol)  (0.082057 L  atm/K  mol)  (293.15 K)  (760 mm Hg/atm)/ (0.059 L) = 102 mm Hg pressure. Using similar reasoning, 800 ml of B/4 medium and 200 ml of head-gas volume

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could produce 1.32 mmol O2 (3.3 mM/2  800 ml/1000 = 1.32 mmol = 0.00132 mol of O2). This additional O2 can be expected to develop a pressure of 121 mm Hg above atmospheric pressure, i.e. (0.00132 mol)  (0.082057 L  atm/K  mol)  (293.15)  (760 mm Hg/atm)/(0.200 L) = 121 mm Hg pressure. By way of checking the pressures measured in the multi-probe apparatus of 850 ml of medium and 90 ml of head-gas space, 284 mm Hg above atmospheric pressure is calculated to potentially occur from photosynthesis (plus the initial 50 mm Hg pressure increase that is caused by sealing and tightening the apparatus). We have measured values approaching 300 mm Hg. So these calculations appear to be reasonable estimates. Pressure increase was much less when cellulose or noadded carbon was tested. Cellulose supports some algal growth and supports larger Daphnia populations which persist longer than NaHCO3. Even if no carbon is added to the ecosystems, some is present from the atmosphere at time of sealing, and the organisms bring in a carbon supply, some of which is respired, so some photosynthesis is possible and a few Daphnia are sustained. Pressure and leakage can be minimized by reducing net photosynthesis by reducing nutrients such as N, P and carbon, by avoiding NaHCO3, and by increasing head-gas volume. Intense grazing also reduces net photosynthesis and pressure, and returns carbon to the dissolved inorganic pool (Fig. 4).

study the carbon dynamics of a plant–soil–atmosphere subjected to temperature and CO2 changes (Lukac et al., 2011; Milcu et al., 2012a, b). Although Closed Ecological Systems have not had wide acceptance as a research tool, they provide thought provoking experiments in student research and have the potential to provide new information on how ecological communities develop within the constraints of limited resources. In open ecosystems, CO2 and O2, being invisible gases, diffuse in and out of containers and remain unmeasured in most studies. Student experiments have used non-instrumented aquatic CESs to test hypotheses of their own. Given the relative inexpensive nature of these experiments, they usually establish 24 or more units, with four or more treatments of five or six replicates each. The use of replicates allows students to observe biological variability and to compare treatments statistically. Students have tested the effects of different nutrient concentrations, container sizes, temperatures, organisms, pesticides and light conditions. Most students expect the animals to be dead within a day of sealing the experiments, and are delighted when animal populations develop for weeks to months. Hypoxic conditions have occurred only rarely and usually when Daphnia populations have become very dense and overgrazed the algae.

4.2. CESs as a research technique

Closed Ecological Systems could also be useful for studying the effects of volatile chemicals, especially those that are noxious, on aquatic communities. Given Henry’s Law, it is possible to calculate the distribution of the test chemical between gas and water, and algal abundance and Daphnia populations can be estimated without opening the containers. Degradation products, including volatile chemicals, could be assessed at the end of the experiment.

At this point, CESs have not become a common research tool and not all authors have explicitly stated the head-gas and liquid volumes. Eley and Myers (1964) tested a mouse-alga CES life support system which incorporated a variable volume control system (brass bellows) to maintain constant pressure; their experiments had to be terminated when the capacity of the variable volume system was exceeded. Maguire (1980) tested 12.5, 125, and 1250 ml subsamples (aliquots) of different communities, with head-gas volume being approximately equal to the liquid; more species survived longer in the larger systems. Folsome and Hanson (1986) developed ecosystems with bright red shrimp that were 50% saline water and 50% air; these provided the background for the commercially available EcospheresÒ which range from 10–25 cm diameter and are at least 30% head gas space (http:// www.eco-sphere.com/). Obenhuber, a student of Folsome’s, used 15 ml liquid and 5 ml head-gas to show long term photosynthesis and respiration in marine communities (1988), and his explanation of the O2 pressure relationships sparked the studies reported here. The Chinese space program published on plant-grazer CES (Hao et al., 2012; Wang et al., 2004, 2006, 2008). Biosphere II combined terrestrial and marine components but its volume was largely atmospheric and contained a “lung” to stabilize pressures, especially those caused by temperature differences (Marino and Odum, 1999). A terrestrial CES has been developed to

4.3. CESs as potential bioassays or toxicity tests

4.4. Experiments conducted in space CES experiments aboard spacecraft are expected to remain sealed and leakage at one atmosphere of pressure would be a problem. The problem would be exacerbated if it became necessary to store an experiment at reduced pressure. In which case, the differential between the internal and external pressure would be greater, and leakage more likely. 4.5. Energetic considerations If CO2 and O2 can be measured in aquatic CESs, the potential exists of converting these changes to energy (joules) using the same assumptions as used for individual organism bioenergetics. For example, Jobling (1994), gives carbohydrate, lipid, and protein (ammonia as a waste product) oxycalorific coefficients as 14.76, 13.72, and 13.36 J/mg O2 respectively (average 13.95 J/mg O2  32 g/mol = 446.2 kJ/mol O2). Hansen et al. (2004), reviewing calorimetry studies in which O2 and heat output

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are measured, suggests 455 ± 15 kJ/mol O2 as Thornton’s rule.

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With NaHCO3 as the carbon source, aquatic CESs do not remain near atmospheric pressure; pressure was less with cellulose or no-added carbon, but could still cause leakage.

Although it would be helpful to have replicated experiments, three lines of evidence support the relationship between O2 production and pressure: (A) The laws describing the distribution of oxygen between aqueous and gas phases i.e., Henry’s Law (Stumm and Morgan, 1996) (B) Obenhuber’s work using pressure to estimate oxygen (Obenhuber, 1986; Obenhuber and Folsome, 1988) and (C) the measured relationships between O2 measurements and pressure reported here (R2 values >0.9) in several singular experiments.

5.2. Oxygen production

Acknowledgments

Most of the O2 will be in the gas phase (29.9 times that in an equal volume of water), so it is critical to know the volume of the gas phase, and include its O2 in total calculations. The amount of O2 produced will be a function of the medium richness, the carbon source, and the volume of the liquid. The total pressure will be controlled by the O2 produced and relative head-gas to liquid volume.

Dr. Jonathon Cole for alerting me to the implications of low O2 solubility in water and the gas–water distribution. Dr. Oscar Monje, NASA and Dr. Donald Obenhuber for checking calculations and commenting on the manuscript. Three anonymous reviewers for suggesting improvements.

5.3. Pressure as an index of O2

CRC Handbook of Chemistry and Physics. 92 ed. CRC Press, 2011. ASTM. E1366 Standard practice for standardized aquatic microcosms: fresh water. Annual Book of Standards, vol. 11.06. ASTM International, W. Conshohocken, PA, 2012. Chang, R. Chemistry, 8th ed McGraw-Hill, New York, 2005. Ecosphere Associates, Inc. http://www.eco-sphere.com/. Eley, J.H., Myers, J. Study of a photosynthetic gas exchanger: a quantitative repetition of the priestly experiment. Tex. J. Sci. 16, 296–333, 1964. Folsome, C.E., Hanson, J.A. The emergence of materially-closed-system ecology, in: Polunin, N. (Ed.), Ecosystem Theory and Application. John Wiley & Sons Ltd., pp. 269–288, 1986. Goldman, J.C., Brewer, P.G. Effect of nitrogen source and growth rate on Phytoplankton-mediated changes in alkalinity. Limnol. Oceanogr. 25, 352–357, 1980. Hansen, L.D., Macfarlane, C., McKinnon, N., Smith, B.N., Criddle, R.S. Use of calorespirometric ratios, heat per CO2 and heat per O2, to quantify metabolic paths and energetics of growing cells. Thermochim. Acta 422, 55–61, 2004. Hao, Z., Li, Y., Cai, W., Wu, P., Liu, Y., Wang, G. Possible nutrient limiting factor in long term operation of closed aquatic ecosystem. Adv. Space Res. 49, 841–849, 2012. Jobling, M. Fish Bioenergetics, 1st ed Chapman & Hall, London, New York, 1994. Lukac, M., Milcu, A., Wildman, D., Anderson, R., Sloan, T., Ineson, P. Non-intrusive monitoring of atmospheric CO2 in analogue models of terrestrial carbon cycle. Methods Ecol. Evol. 2, 103–109, 2011. Maguire Jr., B. Some patterns in post-closure ecosystem dynamics (failure), in: Giesy, J.P. (Ed.), Microcosms in Ecological Research. Technical Information Center, U.S. Department of Energy, Springfield, Virginia, pp. 319–332, 1980. Marino, B.D.V., Odum, H.T. Biosphere 2: Research Past and Present. Elsevier Science B.V, Amsterdam, 1999. Milcu, A., Lukac, M., Ineson, P. The role of closed ecological systems in carbon cycle modelling. Clim. Change 112, 709–716, 2012a. Milcu, A., Lukac, M., Subke, J.A., et al. Biotic carbon feedbacks in a materially closed soil–vegetation–atmosphere system. Nat. Clim. Change 2, 281–284, 2012b. Obenhuber, D.C. The Persistence of Life Measured by Carbon Cycling in Closed Ecological Systems. Microbiology. University of Hawaii, Honolulu, HI, 1986. Obenhuber, D.C., Folsome, C.E. Carbon recycling in materially closed ecological life support systems. Biosystems 21, 165–173, 1988.

5. Conclusions 5.1. Pressure

Turbulence is recommended if pressure is to be used as a surrogate for O2. In l000 ml ecosystems, pressure changes lag O2 changes by hours, unless turbulence is adequate. The dark respiration would be significantly underestimated if mixing were inadequate. 5.4. Dissolved Inorganic Carbon (DIC) Calculations of DIC in earlier studies were probably acceptable since the high pH values (most above eight) would have caused most of the DIC to be in the ionic forms of bicarbonate and carbonate; very little CO2 (which would be in equilibrium between the liquid and gas) would be present. However, this assumption is not true during periods of intense grazing when pH can be less than seven. We now include the gaseous phase of CO2 in our TCO2 calculations. 5.5. Algae–Daphnia interactions The biological reactions between the algal growth and Daphnia populations seem much the same in systems with better seals as earlier studies. The Daphnia population lags the algal growth, but eventually overgrazes, and most of the late born animals starve. These systems rarely go anaerobic, in spite of the probable O2 leakage of the early instrumented CESs. 5.6. Simultaneous measurements of pressure and O2 The multi-probe with rotary mixing appears to provide a satisfactory method for studying O2 and CO2 dynamics, including the potential of using pressure as an approximate O2 surrogate, but O2 measurements are better, if the sensor can be afforded.

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