ABSTRACT

The present study describes and validates a novel yet simple system for simultaneous in vivo measurements of rates of aquatic CO2 production (CO2) and oxygen consumption (O2), thus allowing the calculation of respiratory exchange ratios (RER). Diffusion of CO2 from the aquatic phase into a gas phase, across a hollow fibre membrane, enabled aquatic CO2 measurements with a high-precision infrared gas CO2 analyser. O2 was measured with a PO2 optode using a stop-flow approach. Injections of known amounts of CO2 into the apparatus yielded accurate and highly reproducible measurements of CO2 content (R2=0.997, P<0.001). The viability of in vivo measurements was demonstrated on aquatic dragonfly nymphs (Aeshnidae; wet mass 2.17 mg–1.46 g, n=15) and the apparatus produced precise CO2 (R2=0.967, P<0.001) and O2 (R2=0.957, P<0.001) measurements; average RER was 0.73±0.06. The described system is scalable, offering great potential for the study of a wide range of aquatic species, including fish.

INTRODUCTION

The development of high-precision non-dispersive infrared CO2 analysers has made available a powerful tool for the measurement of CO2 production rates (CO2) in air-breathers, but overlapping absorption spectra of CO2 and H2O preclude their use in aqueous media. However, it is possible to promote diffusion of CO2 from the water to a gas phase that can then be analysed with infrared analysers, a principle that has been used for discontinuous measurements of dissolved CO2 in aquaculture (Pfeiffer et al., 2011; Stiller et al., 2013), but relying entirely on the equilibration of water CO2 with an air-filled head space.

The present study describes a simple aquatic respirometer for simultaneous in vivo measurements of CO2 and oxygen consumption rate (O2) in small aquatic organisms. The novel aspect of this apparatus is the use of a hollow fibre membrane (HFM) as the interface between aquatic and gas phases, providing a large ratio of gas exchange surface area to system volume. Similar HFM designs are finding increasing application as oxygenators in human medicine (Nolan et al., 2011; Potkay, 2014; Strueber, 2015). The high precision, and thus suitability, of the apparatus for measuring small aquatic CO2 was validated both in vitro and in vivo using aeshnid dragonfly nymphs.

MATERIALS AND METHODS

Animal collection and husbandry

Aeshnid dragonfly nymphs (Aeshna spp. and Anax junius; wet mass 2.17 mg–1.46 g; n=15) were caught in the experimental ponds at the University of British Columbia (summer 2016), and were held in the laboratory for several weeks before experiments. Housing was in 1 l glass containers connected to a recirculation system supplied with dechlorinated Vancouver tap water at room temperature (22°C). Animals were fed various wild-caught aquatic insects, twice a week.

List of symbols and abbreviations
     
  • [CO2]

    concentration of CO2

  •  
  • f

    flow rate

  •  
  • HFM

    hollow fibre membrane

  •  
  • m

    mass

  •  
  • P

    pressure

  •  
  • PO2

    partial pressure of O2

  •  
  • RER

    respiratory exchange ratio

  •  
  • STP

    standard temperature and pressure

  •  
  • t

    time

  •  
  • V

    volume

  •  
  • CO2

    CO2 production rate

  •  
  • O2

    O2 consumption rate

  •  
  • αCO2

    solubility of CO2 in water

  •  
  • αO2

    solubility of O2 in water

Experimental setup

The central component of the apparatus for the measurement of CO2 and O2 was a HFM that created a high surface-area interface between counter-current flows of water and CO2-free air (Fig. 1). CO2 added to the water phase rapidly diffused down its partial pressure gradient, across the HFM and into the gas phase, where it was measured with a LI-7000 differential CO2 analyser (LiCor, Lincoln, NE, USA).

Fig. 1.

Schematic representation of the experimental apparatus for the measurement of rates of CO2 production (CO2) and O2 consumption (O2) in small aquatic organisms. Water was circulated in a closed loop indicated by the blue arrows, and CO2-free purge-gas is represented by the dashed arrows. The interface between water and air was a hollow fibre membrane (HFM) gas exchanger, in which water was circulated on the outside of a bundle of gas permeable fibres and CO2-free purge-gas was circulated through the lumen of the fibres in a counter-current fashion, similar to previous designs (Kaar et al., 2007; Arazawa et al., 2012). See Materials and methods, Experimental setup for specifications of system components.

Fig. 1.

Schematic representation of the experimental apparatus for the measurement of rates of CO2 production (CO2) and O2 consumption (O2) in small aquatic organisms. Water was circulated in a closed loop indicated by the blue arrows, and CO2-free purge-gas is represented by the dashed arrows. The interface between water and air was a hollow fibre membrane (HFM) gas exchanger, in which water was circulated on the outside of a bundle of gas permeable fibres and CO2-free purge-gas was circulated through the lumen of the fibres in a counter-current fashion, similar to previous designs (Kaar et al., 2007; Arazawa et al., 2012). See Materials and methods, Experimental setup for specifications of system components.

The HFM consisted of 34 lengths of Oxyplus polymethylpentene fibres (Membrana, Wuppertal, Germany; diameter=380 μm, length=100 mm, area=4059 mm2; see Fig. S5) arranged in parallel and fitted into a 1/8 inch acrylic tube, similar to previous designs (Kaar et al., 2007; Arazawa et al., 2012). The volume surrounding the fibres (0.6 ml) was perfused with water coming from the measuring chamber that was circulated in a closed loop by a peristaltic pump (Gilson MINIPULS 3, Middleton, WI, USA). The lumen of the fibres (0.4 ml volume) was perfused with CO2-free air, produced in a CDA4-CO2 purge-gas generator (Puregas, Broomfield, CO, USA) and passed through columns of Drierite (Hammond Drierite, Xenia, OH, USA) and soda lime (Spectrum Chemical, New Brunswick, NJ, USA). Air flow rates were set to either 20 or 50 ml min−1 (depending on the magnitude of the CO2 signal) with MC Standard Series mass-flow controllers (Alicat Scientific, Tucson, AZ, USA).

CO2-free air was water-saturated in a sparging column and passed through the HFM, where it took up CO2 from the water. Thereafter, the air was de-humidified in 25 cm of TT-070 Nafion tubing (Permapure, Lakewood, NJ, USA), housed in a chamber flushed with dry air at 600 ml min−1 (residual water content during analysis was <1 mmol mol−1). Air containing CO2 from the water was passed through channel B of the CO2 analyser and a separate stream of dry CO2-free purge-gas was passed through channel A. Channels A and B were subtracted from one another, providing an internal control for variability in purge-gas composition. The CO2 analyser was calibrated using analytical grade N2 and 1000 ppm CO2 in N2, and the calibration was confirmed periodically. Gas flow rates through the CO2 analyser were measured at the outlet with a DryCal Definer 220 positive displacement volumetric flow meter (MesaLabs, Butler, NJ, USA); these values were used for calculations.

The measuring chambers consisted of glass scintillation vials (1.9, 4.7 or 24.5 ml volume) fitted with ports for water inlets and outlets and a PO2 (partial pressure of O2) optode with MicroxTX3 meter (Loligo Systems, Viborg, Denmark); optodes were calibrated before each trial using a sodium sulfite solution and air-equilibrated dH2O. A PVC mesh was fitted inside the vials to provide a resting structure for the nymph, and mixing was with a magnetic stir bar that was covered by an aluminium grid to protect the animal (Fig. S5). Water flow rates replaced the vial volume every minute for the smaller vials and every 2 min for the largest vial (i.e. 1.9, 4.7, 12.3 ml min−1). Water pH was measured with a flow-through pH microelectrode (Microelectrodes Inc., Bedford, NH, USA) after the measuring chamber, which was calibrated before each run with BDH precision buffers (VWR, Radnor, PA, USA).

Experimental design

Before each trial, the system was flushed and filled with CO2-free, dechlorinated Vancouver tap water, which was generated in a glass vessel that was continuously sparged with CO2-free air from the purge-gas generator at 600 ml min−1. All experiments were carried out at room temperature (22°C).

In Series 1, the experimental setup was validated by injecting known amounts of CO2-saturated dH2O into the measuring chamber and recording the corresponding peaks in CO2 concentration ([CO2]) in the gas phase. CO2-saturated water was generated by sparging a column (height=0.35 m) of dH2O with 100% CO2. To ensure complete recovery of injected CO2 through the analyser, water in the system was acidified by addition of 10% (v/v) 0.01 mol l−1 HCl (average water pH was 3.6±0.1). After each set of four CO2 injections (5, 10, 15 and 20 μl), the water in the system was replaced, and six replicate runs were performed (n=6).

In Series 2, the experimental setup was validated for the measurement of CO2 (and O2) in dragonfly nymphs. The system was flushed and filled with CO2-free water and a baseline measurement was performed without an animal (phase i). Thereafter, a nymph was introduced into the measuring chamber, which was then covered to minimise disturbance. Once stable readings were obtained (phase ii) the peristaltic pump was stopped, and the decline in water PO2 was recorded for the calculation of O2 (phase iii). When PO2 had decreased to 100 mmHg, water flow was re-started and the accumulated CO2 was washed out of the system, resulting in a peak in [CO2] measured in the gas phase (phase iv). After this washout period, a second set of animal recordings was performed (phase v). Immediately after the trial, the nymph was blotted dry and weighed on an XPE205 electronic balance (Mettler Toledo, Columbus, OH, USA).

Calculations and data analysis

All parameters were recorded continuously with a PowerLab data acquisition unit (ADInstruments, Dunedin, New Zealand) and raw data were analysed with Labchart v8.1.5.

In Series 1, the content of CO2 in the injected water was calculated as:
formula
(1)

where αCO2 (mmol l−1 mm Hg−1) is the solubility of CO2 in water (Boutilier et al., 1984); P is total gas pressure, calculated from atmospheric pressure (average 762±2 mmHg), the height of the sparging column (0.35 m) and subtracting the water vapour pressure (Dean, 1999); and V is the injected volume of CO2-saturated water.

The content of CO2 measured with the CO2 analyser was calculated as:
formula
(2)

where [CO2] is the concentration of CO2 (ppm), f is the gas flow rate (ml min−1; STP) and V is the volume of CO2 gas (22.4 l mol−1; STP). The integral under the [CO2] curve was calculated from the time of injection (t1) until [CO2] returned to baseline (t2) (see Fig. 2B).

Fig. 2.

Series 1, validation of the experimental apparatus. (A) Measured CO2 content (μmol) relative to injected CO2 content (Series 1). Known volumes of CO2-saturated water (5, 10, 15 or 20 μl) were injected into the measuring chamber of the apparatus, and CO2 content was calculated based on the physical solubility of CO2 in water under the tested conditions (see Fig. 1 for details on the apparatus). Measurements of CO2 content were carried out with an infrared CO2 analyser in the gas phase, after diffusion of CO2 across a hollow fibre membrane. All data are means±s.e.m. (n=6) and the linear regression was according to: CO2 measured=0.018±0.005+0.972±0.011×(CO2 injected). (B) Representative trace of CO2 concentration ([CO2], ppm) measured with an infrared CO2 analyser. In each trial, four volumes of CO2-saturated water were injected into the measuring chamber and injection time points are indicated by vertical dashed lines. Data were analysed by integrating under the [CO2] curve (green shading) and subtracting the average baseline reading (horizontal dashed line).

Fig. 2.

Series 1, validation of the experimental apparatus. (A) Measured CO2 content (μmol) relative to injected CO2 content (Series 1). Known volumes of CO2-saturated water (5, 10, 15 or 20 μl) were injected into the measuring chamber of the apparatus, and CO2 content was calculated based on the physical solubility of CO2 in water under the tested conditions (see Fig. 1 for details on the apparatus). Measurements of CO2 content were carried out with an infrared CO2 analyser in the gas phase, after diffusion of CO2 across a hollow fibre membrane. All data are means±s.e.m. (n=6) and the linear regression was according to: CO2 measured=0.018±0.005+0.972±0.011×(CO2 injected). (B) Representative trace of CO2 concentration ([CO2], ppm) measured with an infrared CO2 analyser. In each trial, four volumes of CO2-saturated water were injected into the measuring chamber and injection time points are indicated by vertical dashed lines. Data were analysed by integrating under the [CO2] curve (green shading) and subtracting the average baseline reading (horizontal dashed line).

In Series 2, continuous CO2 was calculated from the animal recording of [CO2] before (phase ii) and after the stop-flow period (phase v) and subtracting the baseline reading without the animal (phase i). CO2 during stop-flow was calculated as described in Eqn 2. The integral was taken from the time point at which the peristaltic pump was re-started (t1) until [CO2] returned to baseline (t2), which was calculated as the average [CO2] during continuous CO2 measurements corrected for drift during stop-flow; see Fig. S1B. The obtained CO2 content was then divided by the length of the stop-flow period (min).

O2 was determined from the slope of the PO2 curve during stop-flow, selecting a 10 min period immediately before re-starting the flow; see Fig. S1A. O2 was then calculated as:
formula
(3)

where –(δPO2t) (mmHg s−1) is the slope of the PO2 curve during stop-flow, Vc and Va (ml) are the volumes of the chamber and the animal (based on animal wet mass and assuming a density equal to water), respectively, and αO2 (μmol l−1 mmHg−1) is the solubility of O2 in water (Boutilier et al., 1984). A control stop-flow recording was performed without an animal and the obtained PO2 slope was subtracted from every animal recording (on average, control PO2 slope was 14±3% of animal respiration slopes). Respiratory exchange ratio (RER) was calculated as stop-flow CO2 over O2.

All data were analysed in RStudio v0.98.1049 (Rv3.3.1). Results of linear regression analyses are presented as adjusted R2 and P-values. Differences in regressions slopes and between O2 and CO2 measurements were compared by ANCOVA with animal mass as a covariate, and RER, pH and PO2 were compared by ANOVA (P<0.05). All data are means±s.e.m.

RESULTS AND DISCUSSION

In Series 1, injections of CO2-saturated water into the measuring chamber were used to demonstrate the precision and accuracy of the apparatus. After initiating a run, [CO2] readings quickly decreased to an average baseline of 2.6±0.1 ppm. Injections of CO2 were reflected in a peak in [CO2] in the gas phase (Fig. 2B) with a time lag of several seconds for the tested gas flow rates. The measured content of CO2 corresponded well with that injected (Fig. 2A; R2=0.997, P<0.001). Thus, injected CO2 quickly and completely diffused from the water to the gas phase across the HFM.

In Series 2, dragonfly nymphs spanning a wide range of wet masses were used to validate the apparatus for in vivo measurements of CO2 and O2. Few other studies have measured respiratory rates in aeschnid dragonfly nymphs and, to our knowledge, no other study has performed simultaneous measurements of CO2 and O2. Petitpren and Knight (1970) measured O2 in nymphs of Anax junius over a range of animal dry masses. Re-calculating their data (assuming that dry mass is 25% of animal wet mass; Dermott and Paterson, 1974) yielded O2 values (μmol min−1) that scaled with wet mass (m) according to:
formula
(4)
and these results were not significantly different from the data generated in the present study (P=0.652; Fig. S2):
formula
(5)

In addition, both scaling exponents for O2 are within the range that has been previously reported for invertebrates (Glazier, 2005). Therefore, O2 can be viewed as a reliable internal reference for the validation of the novel CO2 measurements.

Continuous measurements of CO2, before and after stop-flow, were not significantly different (P=0.273) and averaged over all animals 0.070±0.017 μmol min−1. Stop-flow CO2 was significantly lower than continuous CO2 (P=0.014) and was on average 0.051±0.011 μmol min−1 (Fig. S3). This discrepancy was driven entirely by the largest individuals measured (m>1100 mg, n=4), where CO2 was lower during stop-flow compared with continuous readings, and in a subset of data excluding these individuals there was no significant difference between continuous and stop-flow measurements of CO2 (P=0.176, n=11). This is an important result and indicates that there is no methodological discrepancy between the two measurements, and CO2 can be determined reliably using the continuous and stop-flow approaches.

The observed discrepancy between CO2 measurements in the largest individuals may be the result of: (1) CO2 loss to the environment during stop-flow; (2) formation of HCO3 during stop-flow and its retention during washout; or (3) a lower animal CO2 during stop-flow. Loss of CO2 from the system seems unlikely, as CO2 injected in Series 1 was recovered entirely across the HFM, using peak [CO2] values that were in line with those produced by the animal. Likewise, CO2 retained in the system as HCO3 should be reflected in a lower pH and a higher CO2 in the continuous animal recording after stop-flow (phase v), compared with the initial recording (phase ii), neither of which was observed (in the whole dataset or in the subset of largest individuals; Figs S3, S4). Thus it seems possible that CO2 in the largest individuals was lower during stop-flow periods because of a reduction in metabolic rate, perhaps related to the accumulation of metabolites in the water, decreasing pH and PO2.

Fig. 3A depicts stop-flow CO2 data in direct comparison with the control measurements of O2 taken over the same stop-flow period; CO2 scaled with wet mass according to:

Fig. 3.

Series 2, validation of the experimental apparatus for in vivo measurements. (A) Rates of CO2 production (CO2, μmol min−1, red triangles) and O2 consumption (O2, blue circles) of aeschnid dragonfly nymphs (n=15) as a function of animal wet mass (g) on a double-log scale (Series 2). Data points represent individual measurements and power regression lines are indicated for both parameters, corresponding to the linear models: log10(CO2)=–1.124±0.038+0.672±0.033log10(m) (R2=0.967, P<0.001) and log10(O2)=–0.965±0.044+0.678±0.038log10(m) (R2=0.957, P<0.001). The slopes for both parameters were significantly different (ANCOVA, P=0.015). (B) Average respiratory exchange ratio (RER) calculated from the individual data in A, as CO2/O2. Data are means±s.e.m.

Fig. 3.

Series 2, validation of the experimental apparatus for in vivo measurements. (A) Rates of CO2 production (CO2, μmol min−1, red triangles) and O2 consumption (O2, blue circles) of aeschnid dragonfly nymphs (n=15) as a function of animal wet mass (g) on a double-log scale (Series 2). Data points represent individual measurements and power regression lines are indicated for both parameters, corresponding to the linear models: log10(CO2)=–1.124±0.038+0.672±0.033log10(m) (R2=0.967, P<0.001) and log10(O2)=–0.965±0.044+0.678±0.038log10(m) (R2=0.957, P<0.001). The slopes for both parameters were significantly different (ANCOVA, P=0.015). (B) Average respiratory exchange ratio (RER) calculated from the individual data in A, as CO2/O2. Data are means±s.e.m.

formula
(6)

These results lend further support to the validity of stop-flow CO2 measurements, which corresponded well with the previously validated O2 data, producing physiologically relevant RER (0.73±0.06; Fig. 3B), and RER in the subset of the largest individuals was not significantly different from that of all others (P=0.886). Thus it appears that the largest individuals had lower stop-flow CO2 as well as O2, which is in line with a decreased metabolic rate during stop-flow (but not with CO2 loss to the environment or the retention of HCO3). Clearly, the effects of changing stop-flow respirometry conditions on the respiratory rates of dragonfly nymphs deserve further attention, especially as these effects may be sensitive to animal life stage.

The range of animal sizes investigated here spans nearly the entire aquatic life stage of aeschnid dragonflies and a 700-fold increase in body mass. The two smallest individuals were hatched in the laboratory, and both were third instar nymphs when measured (2.17 and 2.40 mg wet mass). Even for these small animals, the CO2 signal after stop-flow (duration 125 and 71 min) was on average 17±3-fold larger than baseline readings. Therefore, it appears feasible to measure CO2 in animals that are yet another order of magnitude smaller. Finally, because the water phase is in constant contact with the CO2-free (but normoxic) purge-gas across the HFM, long-term experiments are possible without the animal depleting O2 in the system (Fig. S4). Likewise, gas tensions in the measuring chamber can be easily controlled by changing the composition of the purge-gas, presenting great potential for hypoxia studies. The described apparatus is scalable, and thus a wide range of aquatic species may be studied, from invertebrates to larval and juvenile vertebrates such as fish and amphibians.

Acknowledgements

Thanks are due to Mike Sackville and Alexander Cheng for their help with animal capture and care.

Footnotes

Author contributions

T.S.H. and P.G.D.M. conceived the study. T.S.H. built the apparatus, performed the experiments and wrote the manuscript. All authors participated in the analysis and interpretation of the data and revised the manuscript.

Funding

This study was supported by Natural Sciences and Engineering Research Council of Canada (NSERC) Accelerator [462242-2014 to P.M.; 446005-13 to C.J.B.] and Discovery Grants [2014-05794 to P.G.D.M.; 261924-13 to C.J.B.] and a Canada Foundation for Innovation (CFI) John R. Evans Leaders Fund grant [JELF 33979 to P.G.D.M.]

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Competing interests

The authors declare no competing or financial interests.

Supplementary information