Although fish respond effectively and restore blood pH during CO2 induced acid-base balance disturbance, recent studies have noted potential impacts of ocean acidification across a range of areas including neurosensory disruptions, increased otolith growth, altered mitochondrial function, and changes to metabolic rate1,2,3,4,5,6,7,8,9. A suggested unifying hypothesis is that the compensatory response induced during CO2 exposure to correct blood pH may have negative downstream consequences or induce tradeoffs1,4,10. This pH compensation is associated with a sustained increase of extra and intracellular concentration of HCO3 in response to the elevated partial pressure of CO2 (pCO2)11,12,13,14,15. Assuming the rate of adaptation does not keep pace with a rapidly acidifying ocean, fish in future oceans will require consistent and elevated levels of ion exchange to sustain elevated HCO3 and normal pH, a process that is anticipated to add to the cost of basic maintenance of homeostasis, most often quantified as standard metabolic rate (the minimum O2 consumption rate of a resting animal in the post-absorptive state16).

The gill, intestine, and kidney are the organs involved with acid-base balance and osmoregulation in marine fish13,17. Estimates vary widely, but metabolic cost of ion regulation has been proposed to range from 1–30% or 6–15% of whole-animal standard metabolic rate (reviewed in ref. 18,19). More targeted estimates of ion transport using specific isolated organs suggest that the gills and intestine account for ~4%20 and 5.6%21 of standard metabolic rate, respectively, which combined would account for about 10% of whole-animal standard metabolic rate. Interestingly, despite the need for HCO3 retention during a CO2-induced acidosis, toadfish experience ~34% increase in HCO3 ion loss through the intestine when exposed to near-future CO2 scenarios (1900 μatm CO2)22, a trend also apparent at higher CO2 levels23. Although the intestine is a metabolically active tissue central to water balance and survival in seawater, the functional consequence of this relatively high HCO3 loss during CO2 acclimation has not been examined. Elevated HCO3 secretion may also stimulate an increase in fish CaCO3 production and release to the marine environment, potentially altering the marine carbon cycle24. Since teleosts are the largest vertebrate group with a vital role in oceanic food webs, it is important to understand how impacts to the intestine during future ocean acidification scenarios could affect both fish and their surrounding environment. The first goal of the present study was to determine if the intestine of a marine teleost, an organ known to show phenotypic plasticity in other ion regulatory challenges25, would dynamically regulate intestinal function to reduce CO2-induced intestinal HCO3 loss in favor of whole-body HCO3 retention. We predicted that tissue from CO2-acclimated toadfish would show reduced bicarbonate secretion rates. After seeing a stimulation rather than a reduction of bicarbonate secretion rates, a second goal was to test the hypothesis that an increase in intestinal ion transport that occurs in response to elevated CO2 would be associated with an increased tissue metabolic demand. Experiments were conducted at 1900 μatm CO2, a level currently seen in certain coastal and upwelling zones26,27 and predicted globally in year 230028.


Bicarbonate secretion rates of isolated tissue

Contrary to expectations, anterior intestinal tissue from CO2 acclimated toadfish exhibited significantly increased HCO3 secretion rates (μmol cm−2 h−1) when compared to control tissue under identical conditions (Fig. 1). This result indicated prior acclimation to CO2 does not suppress but stimulates intestinal HCO3 transport by around 13%. In addition, within both the control and CO2 acclimated fish, HCO3 secretion rates using serosal salines mimicking plasma conditions at 1900 μatm CO2 were significantly higher than HCO3 secretion rates under control serosal salines (Two-way ANOVA, Fish treatment P < 0.009, Saline P < 0.001, Fish treatment × saline P < 0.490, Fig. 1). TEP and conductance remained stable during experiments and were in agreement with earlier reports (Supplementary Table 1)21,29.

Figure 1: Bicarbonate secretion rates of isolated anterior intestine from control and CO2 acclimated toadfish.
figure 1

Effect of blood-side saline composition and acclimation exposure on HCO3 secretion rates (means ± s.e.m.) of isolated anterior intestinal tissue obtained from gulf toadfish acclimated to control (~440 μatm CO2, n = 9) or ~1900 μatm CO2 (n = 10) for 2–4 weeks. Tissues from either control or 1900 μatm CO2 acclimated fish mounted in this dual chambered Ussing/pH-stat system were bathed on either side by salines that mimicked in vivo ionic composition. Each tissue received two blood-side (serosal) saline treatments, control and CO2 saline, representative of HCO3 and pCO2 previously measured in toadfish blood following control and 1900 μatm CO2 exposure (Supplementary Table 3). Two-way ANOVA; Acclimation exposure: P < 0.004, Saline: P < 0.001, Acclimation exposure × saline: P < 0.475. *Significant acclimation effect, Significant saline effect.

Oxygen consumption rates of isolated tissue

Under both saline compositions, tissue from 1900 μatm CO2 acclimated toadfish showed a significant 8% increase in oxygen consumption rate compared to tissue from control acclimated fish. In contrast to HCO3 secretion rates, oxygen consumption rates of isolated tissue from both control and 1900 μatm CO2 exposed fish showed no effect of saline composition (Two-way ANOVA, Fish treatment P < 0.033, Saline P < 0.769, Fish treatment × saline P value < 0.509, Fig. 2). A similar relationship was observed when data was corrected for body mass (Supplementary Fig. 1).

Figure 2: Oxygen consumption of isolated anterior tissue from control and CO2 acclimated toadfish.
figure 2

Effect of blood-side saline composition and acclimation exposure on oxygen consumption rates (means ± s.e.m.) of isolated anterior intestinal tissue taken from toadfish acclimated to control (~440 μatm CO2; n = 12) or ~1900 μatm CO2 (n = 8) for 2–4 weeks. Tissues from either control or 1900 μatm CO2 acclimated fish mounted in this dual-chambered epithelial respirometer were bathed on either side by salines designed to mimic in vivo ionic composition. Each tissue received two blood-side (serosal) saline treatments, control saline and CO2 saline, that were representative of HCO3 and previously measured in toadfish blood following acclimation at control and 1900 μatm CO2 (Supplementary Table 3). Two-way ANOVA; Acclimation exposure: P < 0.033, Saline: P < 0.769, Acclimation exposure × saline: P < 0.509. *Significant acclimation effect.


These results indicate that the marine fish intestine has a higher metabolic demand at 1900 μatm CO2 than at present-day ambient conditions (~400 μatm CO2), that is likely attributed to an increase in intestinal HCO3 loss from the body. This elevated CO2 level is predicted for year 2300 and is currently seen in upwelling coastal areas26. Similar to other fish experiencing elevated CO2 in a marine environment, gulf toadfish have been shown to defend blood pH following exposure to 1900 μatm CO2 by sustaining elevated levels of HCO3 in the face of higher pCO2 in the blood11. However, this compensation was associated with an increased intestinal HCO3 loss that was presumed to be associated with an activation of existing ionoregulatory transport pathways11,22. These pathways involve the movement of plasma HCO3 into the intestinal lumen in exchange for Cl and are critical for maintaining water balance30. However, from a whole-animal acid-base balance perspective, HCO3 loss during CO2 compensation counteracts the need to retain HCO3 and defend pH, suggesting that fish faced with longer term acclimation to CO2 must dynamically downregulate pathways involved in intestinal HCO3 secretion. Contrary to our initial hypothesis, comparison of HCO3 secretion rates of intestinal tissue from CO2 and control acclimated fish under identical saline conditions revealed that CO2 exposure leads to a stimulation, rather than a downregulation of HCO3 transport pathways.

We propose that the 13% increase in HCO3 loss from CO2 acclimated fish reflects an increased energetic demand, and thus increased CO2 production, in intestinal tissue. This suggestion is supported by an 8% increase in O2 consumption in tissue from CO2-acclimated fish. A detailed mechanistic explanation of this proposed response can be seen in Fig. 3. Sustained elevated plasma HCO3 following CO2 compensation leads to an increase in HCO3 movement from the blood into the intestinal cell that is paired with the movement of Na+ through basolateral NBC1, a Na+-HCO3 co-transporter31. The sustained Na+ influx must be compensated for by Na+ extrusion through the Na+-K+ ATPase (NKA). Increased NKA activity leads to an increased ATP and thus O2 demand. Meeting this demand results in increased endogenous CO2 production available for hydration via intracellular carbonic anhydrase (CAc) to form additional HCO3. Secretion of this excess HCO3 through apical SLC26a6 anion exchange likely accounts for the stimulation of HCO3 loss during CO2 exposure. Hydration of endogenous CO2 also results in formation of protons that must be eliminated from the intestinal cell and could put additional demand on the gill. This proton extrusion by the intestine may occur via Na+-dependent or independent pathways30, both of which are energy demanding and could contribute to the observed elevation of O2 consumption.

Figure 3: Proposed impacts of 1900 μatm CO2 on intestinal transport physiology in a marine teleost.
figure 3

Compensation for a CO2-induced acidosis increases HCO3 and pCO2 in extracellular fluids11. (A) Elevated serosal HCO3 during CO2 exposure stimulates transport via Na+:HCO3 co-transporter, NBC1, leading to both an increase influx of HCO3 and Na+ across the basolateral membrane. (B) Sustained influx of Na+ via NBC1 likely leads to a demand for increased Na+ extrusion via the energy-demanding Na+ K+ ATPase (~). (C) The metabolic demand and increased O2 consumption associated with handling an increased Na+ influx would generate additional endogenous CO2, increasing substrate for intracellular CAc hydration. (D) Intracellular HCO3 generated via this process likely accounts for the observed stimulation of bicarbonate secretion via SLC26a6 (a6) in isolated tissue from fish acclimated to 1900 μatm CO2 compared to control fish under serosal salines with identical bicarbonate concentrations. Protons generated in via carbonic anhydrase in step C would likely be extruded from the cell via NHE1 and add to the increase in intracellular Na+. For a more detailed overview of marine fish intestinal transport processes see review30.

It is likely that the 8% increase in intestinal tissue O2 consumption would not be observable by measurements of whole-animal metabolic rate. Difficulties in picking up small differences in organismal metabolic rate may explain some of the varying results from studies examining the effects of ocean acidification on fish standard metabolic rate1. In addition, there appears to be considerable intra- and inter-species variation in the response to elevated CO2 and there is inherent difficulty in comparing measurements using different methodologies32,33. As demonstrated in the present study and by others5,34,35,36,37, increased resolution and mechanistic insight into energetic tradeoffs and/or apparent consequences of ocean acidification may be obtained by integrating techniques and methods that examine multiple levels of organization. Altered mitochondrial capacity5, shifts in energetic budgets34,38,39, increased expression/activity of gill and intestinal ionoregulatory genes and proteins15, increased ventilation40, and increased protein turnover38 have all been noted during ocean acidification-relevant CO2 exposures with little impact to whole-animal measurements. The importance of integrating multiple techniques is apparent in a recent study on CO2 exposure (1,200 and 2,200 μatm CO2) in Atlantic cod exhibiting increased intestinal NKA mRNA and protein concentration, while exhibiting no change in NKA protein activity41. While the present study of isolated intestinal tissue precluded normal hormonal cascades or feedback mechanisms, it offered the advantage of careful control of blood-side (serosal) saline conditions and made it possible to identify mechanistic differences in tissue function that were impossible to observe in previous in vivo work22. One other caveat to note is that air, rather than custom CO2/O2 gas mixtures were used during respirometry experiments. Earlier work on this preparation revealed that tissue metabolic rate is not limited by O2 above 75% air saturation21. Thus, tissue was not O2-limited in the present study since saturation remained above 80%.

The marine fish intestine is fine-tuned to changes in plasma HCO3 to aid in water uptake and to handle the alkaline tide associated with digesting a meal21,31. Although the functional consequence of stimulated intestinal HCO3 loss remains to be fully elucidated, the increase in O2 consumption leads us to conclude that this CO2-induced response is potentially maladaptive but persists for protection of osmoregulatory and digestive functions. Support for this conclusion comes from a recent study, on Atlantic cod, demonstrating that high levels of CO2 (9200 μatm) lengthen the time needed to digest a meal42. Interestingly, reduced digestive efficiency has also been noted in the sea urchin, albeit through a different mechanism, reduced stomach pH. These urchins exhibited a behavioural adjustment, to counteract this reduced efficiency. Fish may also adjust feeding behaviour in such instances, but broad behavioural impairments across various species with CO2 exposure4,8 may impact such processes and should be further investigated. An increased metabolic demand from ion transport processes, as seen in the present study, may detract from energy available for digestive functions in the intestine, slowing the process of digestion. Calculations using estimates of toadfish standard metabolic rate43 suggest that the energetic cost of CO2 acclimation in the intestine would account ~0.5% of whole animal O2 consumption21. Albeit small, any factor that promotes an energy reallocation or increases standard metabolic rate could exacerbate already narrow metabolic constraints44 or possibly interact with projected temperature elevations to increase overall impact36,39,41,45.

The demand to secrete HCO3 in the intestine to maintain water balance is well-conserved across marine teleosts30. Increased intestinal HCO3 secretion with CO2 exposure as reported here has also been demonstrated at higher CO2 levels in the plainfin midshipmen (~50,000 μatm CO2)46, in the toadfish (5000–20,000 μatm CO2)23, and suggested from gene expression and/or protein assays in the Japanese ricefish (7000 μatm CO2)34 and the Atlantic cod (1,200 and 2,200 μatm CO2)41. These studies suggest that increased intestinal HCO3 secretion and metabolic demand during CO2 exposure could be a ubiquitous response to elevated CO2 throughout marine bony fishes. Finally, intestinal HCO3 secretion results in formation and excretion of CaCO3 by marine fish which amounts to at least 3–15% of the marine inorganic CaCO3 production24. Although a study on the toadfish reported unaltered CaCO3 excretion rates22 at 1900 μatm a more recent study on toadfish23 and an earlier study on midshipmen46 may suggest otherwise. Thus, increased intestinal HCO3 loss at elevated CO2 in other species may impact the magnitude of this globally important calcification process, a possibility worthy of further study. In this context, it is important to consider an alternative scenario in which the increase stimulation of HCO3 secretion during CO2 exposure serves to assist the animal in buffering ingested acidified seawater, making conditions more favorable for carbonate precipitation and continued water absorption. However, this cannot be the sole purpose of the increase HCO3 secretion, as the extra protons ingested in a given amount of 1900 μatm CO2 water is several orders of magnitude lower than the increase in HCO3 secretion due to acclimation. These calculations support the idea that HCO3 loss during CO2 exposure is not beneficial for the animal.

Surprisingly few studies have examined the impact of sustained global increases of carbon dioxide on acid-base and osmoregulatory processes in marine fish. While the presents study reports relatively small increases in bicarbonate secretion and oxygen consumption in the intestine, it is important to remember that these changes likely require reallocations or adjustments in an organism that may already be facing other downstream impacts of CO2. Although compensation for elevated CO2 in marine fish typically occurs within days11, it cannot be ruled out that longer acclimation periods, transgenerational effects, and/or adaptation may affect the dynamics of acid-base balance. Furthermore, interspecies differences associated with the cost of ionoregulatory demands are likely40.


Animal collection and care

Gulf toadfish (Opsanus beta) were obtained from local shrimpers as by-catch in Biscayne Bay and acclimated to flow-through, aerated, sand-filtered seawater from Bear Cut, FL (22–25 °C, 32–35 ppt) in the laboratory at the University of Miami for at least two weeks prior to experimentation. During this period, fish were fed squid twice weekly. Once introduced to experimental tanks, toadfish (HCO3 secretion mass range: 21.7–30.9 g, O2 consumption mass range: 28.1–48.6 g) were fed 5% of their body weight weekly. Food was withheld at least 4 days prior to experimentation, a time period previously demonstrated to ensure that confounding effects of specific dynamic action (SDA) would not be a factor for HCO3 secretion or oxygen consumption measurements21. Fish were sacrificed using a lethal dose of 0.2 g/L MS-222 buffered with 0.3 g/L NaHCO3. All general animal care and animal sacrifice protocols were carried out in accordance with relevant guidelines for experiments on teleost vertebrates provided by University of Miami IACUC (Institutional Animal Care and Use Committee). All experimental protocols were approved by University of Miami Institutional Animal Care and Use Committee. (IACUC 13-225). IACUC is accredited by the Association for Assessment and Accreditation of Laboratory Animal Care (AALAC). Toadfish were collected with the approval and in accordance with guidelines outlined by the Florida Fish and Wildlife Conservation Commission (Permit SAL-12-0729 SR).

General experimental procedures

Toadfish were acclimated to either control or CO2 (~440 and ~1900 μatm CO2, respectively, Supplementary Table 2) for 2–4 weeks, a time period previously deemed sufficient to elicit a CO2 compensatory response in the toadfish11. Throughout the methods section, reference to CO2 refers to the 1900 μatm CO2 level, with respect to both treatment exposure and saline. Following acclimation, anterior intestinal tissue was dissected and immediately placed between two half-chambers where the tissue was bathed on either side by salines representative of in vivo conditions. Gut lumen-side (mucosal) salines were identical throughout all treatments, however, two different blood-side (serosal) salines (“control” or “CO2”) were used to mimic measured values at control or 1900 μatm CO2 (Supplementary Table 3)11. These Ussing chambers were combined with pH-stat titration to allow for measurement of HCO3 secretion rates from the blood-side to the gut-side saline29. In addition, the O2 consumption rate of isolated intestinal segments were measured using a custom-built epithelial respirometer21. Each mounted tissue was treated with both the control and the CO2 serosal saline, allowing for examination of the effect of prior CO2 acclimation along with the effect of blood-side saline conditions that mimicked in vivo measured blood chemistry. If dynamic regulation was occurring in tissue from fish exposed to CO2 to reduce HCO3 loss, reduced HCO3 secretion rates would be expected in tissue from CO2 acclimated fish.

Seawater manipulation

Stable CO2 levels were achieved using a Loligo pH-based negative feedback system (Loligo Systems, Tjele, Denmark) as previously described11,22. In this system, a standard curve was generated based on the relationship between known gas standards and seawater pH. Once a pH setpoint corresponding to 1900 μatm CO2 was calculated following calibration and measurement of ambient seawater, 100% CO2 was gassed directly into flow-through tanks via solenoid valves controlled by pH electrodes (Sentix H, wtw Germany) and a pCO2/pH DAQ-M digital relay instrument connected to CapCTRL software (Loligo Systems, Tjele, Denmark) to achieve the desired pCO2 setpoint. pCO2 levels typically stayed within 4–10 percent of setpoint values. Independent pH measurements were taken multiple times a week using a separate pH electrode (pHNBS, PHC3005, Radiometer, France). Target CO2 levels were confirmed with measurements of seawater TCO2 using a Corning 965 CO2 Analyzer (Corning 965, Corning Diagnostics, UK). TA (titratable alkalinity) and pCO2 levels were calculated from measured pHNBS and TCO2 measurements into CO2SYS software47. These calculations confirmed target pCO2 values for control (ambient-~439 μatm CO2) and 1900 (1878 μatm CO2) were reached. Water chemistry parameters including temperature and salinity are reported in Supplementary Table 2.

Electrophysiological measurements

In the Ussing chamber systems (Physiological Instruments, San Diego, CA, USA), current and voltage electrodes attached to an amplifier measured transepithelial potential (TEP, mV) differences between a baseline 0 μA current clamp and a 3 second pulse of 50 μA that was applied every 60s. Measurements were logged using Acknowledge software (v. 3.8.1, BIOPAC Systems). TEP values are reported with a luminal reference of 0 mV and conductance was calculated using Ohm’s law (Supplementary Table 1).

In vitro Ussing chamber/pH stat electrophysiological and bicarbonate secretion measurements

Simultaneous measurement of electrophysiological parameters and bicarbonate secretion of isolated tissue was achieved using Ussing chamber systems (Physiological Instruments, San Diego, CA, USA) combined with automated pH-stat titration (TIM854/856 Titralab and Titramaster software v.5.1.0, Radiometer, Copenhagen, Denmark)29. Following acclimation in either control or CO2 treatment tanks (1900 μatm CO2, 2–4 weeks; Supplementary Table 2), anterior intestine segments were mounted on tissue holders designed to expose 0.71 cm2 of tissue to two half-chambers (1.6 mL) in the Ussing chamber system (P2400, Physiological Instruments). In this setup, isolated tissue was bathed in pre-gassed serosal (blood-side) or mucosal (gut-side) salines continuously mixed by air-lift gassing and held at 25 °C using a recirculating water bath. Mucosal saline composition remained unaltered throughout all experiments (Supplementary Table 3) and was gassed with 100% O2. “Control” and “CO2” serosal salines were designed to mimic in vivo HCO3 and pCO2 levels previously measured in the plasma of toadfish during exposure to control and 1900 μatm CO2 (3.3 and 6.3 mM HCO3, 0.225 and 0.462% CO211, respectively; Supplementary Table 3).

A pH electrode (PHC4000-8, Radiometer, Denmark) and an acid-dispensing microburette were submerged into the mucosal half-chamber. The rate of acid titrant addition and the titrant concentration (0.005 N HCl) needed to hold the mucosal saline at a constant pH of 7.8 were used to calculate the epithelial HCO3 secretion rate. Electrophysiological measurements were taken simultaneously with bicarbonate secretion rates (Supplementary Table 1). Once tissue preparations were considered stable based upon steady bicarbonate secretion and electrophysiological parameters, a minimum 30-minute measurement interval was recorded prior to a saline change. Each isolated intestinal tissue received both control and CO2 serosal saline treatments. Although tissues have been demonstrated to be viable for at least 5 hours in prior studies using this species and setup29, the order of serosal salines applied were randomized. During a saline change, the first serosal saline was carefully removed with a syringe, the half chamber was rinsed, and replaced by the second serosal saline treatment. Measurements post-saline change were continued until stable levels were recorded for a minimum of 30 minutes.

In vitro oxygen consumption measurements

Following the same acclimation procedures outlined for bicarbonate secretion experiments, the oxygen consumption rate of isolated anterior intestine was measured using a custom-designed epithelial respirometer (Loligo Systems, Tjele, Denmark). Intestinal tissue was mounted so that 0.87 cm2 of tissue was exposed to two half-chambers (2.80 mL), mucosal saline on the gut side and one of two serosal saline treatments (described above) on the blood-side. All salines were pre-gassed with air (100% O2 saturation) rather than custom O2 mixes to make measurements of oxygen consumption rates comparable to whole-animal measurements. Saline HCO3 concentrations were kept the same in the control and CO2 serosal salines (Supplementary Table 3).

Salines in half chambers were continuously mixed by micromagnetic glass-coated Teflon stir bars (Loligo Systems), and a Teflon tissue mount ensured that the system was gas-tight21. Oxygen measurements were conducted using a fiber-optic cable secured to the outside wall of either glass half-chamber that illuminated a fiber-optic sensor spot glued to the inside wall of each respective side. Each cable was connected to a separate single-channel oxygen meter (Fibox 3) used in conjunction with Oxy-View software (PST3-V6.02; PreSens, Regensburg, Germany). Prior to daily experiments, calibrations were performed with salines pre-gassed with air for 100% air saturation and gassed with N2 conditions representing no oxygen saturation.

Intermittent-flow respirometry was performed to determine oxygen consumption rates of isolated tissue by flushing and replacing salines using a manual gravity-fed system. Flush cycles guaranteed complete saline replacement during open cycles and time intervals during closed measurements were limited to ~20 minutes, to ensure that air saturation of tissue did not drop below 80%, since values below 75% were previously shown to restrict this tissue21. Previous use of this respirometry system has shown negligible rates of gas back-flux with the atmosphere or across chambers and a constant O2 consumption rate at air saturation above 75%21. Tissue O2 consumption was estimated from the mucosal and serosal O2 depletion rates.

Statistical analysis

Two-way ANOVAs were used to analyze bicarbonate secretion (Fig. 1), oxygen consumption (Fig. 2, Supplementary Fig. 1), and electrophysiological measurements (Supplementary Table 1) with treatment exposure (acclimation) and saline exposure as independent variables. Significance for all tests was determined at P < 0.05 for all tests and all data are presented as means ± s.e.m. unless specifically noted otherwise.

Additional Information

How to cite this article: Heuer, R. M. and Grosell, M. Elevated CO2 increases energetic cost and ion movement in the marine fish intestine. Sci. Rep. 6, 34480; doi: 10.1038/srep34480 (2016).