Acidified seawater impacts sea urchin larvae pH regulatory systems relevant for calcification

Abstract

Calcifying echinoid larvae respond to changes in seawater carbonate chemistry with reduced growth and developmental delay. To date, no information exists on how ocean acidification acts on pH homeostasis in echinoderm larvae. Understanding acid–base regulatory capacities is important because intracellular formation and maintenance of the calcium carbonate skeleton is dependent on pH homeostasis. Using H⁺-selective microelectrodes and the pH-sensitive fluorescent dye BCECF, we conducted in vivo measurements of extracellular and intracellular pH (pH_e and pH_i) in echinoderm larvae. We exposed pluteus larvae to a range of seawater CO₂ conditions and demonstrated that the extracellular compartment surrounding the calcifying primary mesenchyme cells (PMCs) conforms to the surrounding seawater with respect to pH during exposure to elevated seawater pCO₂. Using FITC dextran conjugates, we demonstrate that sea urchin larvae have a leaky integument. PMCs and spicules are therefore directly exposed to strong changes in pH_e whenever seawater pH changes. However, measurements of pH_i demonstrated that PMCs are able to fully compensate an induced intracellular acidosis. This was highly dependent on Na⁺ and HCO₃⁻, suggesting a bicarbonate buffer mechanism involving secondary active Na⁺-dependent membrane transport proteins. We suggest that, under ocean acidification, maintained pH_i enables calcification to proceed despite decreased pH_e. However, this probably causes enhanced costs. Increased costs for calcification or cellular homeostasis can be one of the main factors leading to modifications in energy partitioning, which then impacts growth and, ultimately, results in increased mortality of echinoid larvae during the pelagic life stage.

Sea urchin larvae have been shown to react with particular sensitivity to CO₂-induced reductions in seawater pH (1–4). When larvae are chronically exposed to elevated seawater pCO₂ of >0.1 kPa, e.g., as is predicted to occur during the next century in response to anthropogenic CO₂ emissions or through upwelling of low-pH deep water, this sensitivity is reflected in reduced growth and developmental rates (5, 6). Echinoderm larvae are considered to be especially vulnerable to seawater pH reduction and to the associated changes in calcium carbonate saturation state of seawater (Ω_Cal) because their internal skeleton is composed of high magnesium calcite, a highly soluble form of CaCO₃ (7, 8). However, long-term reductions in growth and development might just as well be evoked by other physiological mechanisms that are also sensitive to hypercapnia and the related acid–base disturbances. Recent studies conducted on several marine taxa including mollusks (9) and echinoderms (10) demonstrated increased metabolic rates in response to elevated seawater pCO₂. It was concluded that reductions in somatic growth and rate of development were caused by a shift of energy partitioning toward vital functions such as cellular acid–base homeostasis (9, 10). For example, sea urchin larvae exposed to acidified seawater (0.35 kPa CO₂, pH 7.25) maintained a fully calcified larval phenotype, albeit at a reduced rate of body growth (4). This indicates that biomineralization mechanisms remain functional despite CO₂-induced changes in the seawater carbonate system.

The calcification process in sea urchin larvae has been well investigated (e.g., refs. 7, 11–14). Primary mesenchyme cells (PMCs), located within the extracellular matrix of the primary body cavity, form a syncytium around the growing spicules of the pluteus larvae. This syncytial sheath covers the entire surface of the spicules and communicates with the extracellular environment of the primary body cavity (8, 15). The high magnesium calcitic spicules are formed through the production of a transient amorphous calcium carbonate (ACC) phase within vesicles in PMCs. ACC is subsequently released into the spicular cavity (7, 8, 12, 16). To fuel the calcification process, bicarbonate (HCO₃⁻) is absorbed from the seawater (40%) as well as generated from metabolic CO₂ (60%) (11). On the other hand, Ca²⁺ is exclusively obtained directly from the seawater (13). Although the general principle of calcification is well understood, mechanistic information concerning the nature of the transporters that facilitate Ca²⁺ or HCO₃⁻ uptake in PMCs is limited. Several pharmacological studies suggested that Ca²⁺ channels and transporters are key players in the provision of Ca²⁺ for spicule formation (17, 18). To remove protons generated during CaCO₃ precipitation, PMCs must possess efficient acid–base regulatory properties (19, 20). However, the mechanistic basis of acid–base regulation in PMCs has not been explored so far. Typically, a cell is required to secrete acid equivalents (e.g., H⁺) or to increase intracellular buffering capacity via the import of buffers (e.g., HCO₃⁻) to compensate for an intracellular acidosis. Acid–base equivalent transport is facilitated through ion transporters such as Na⁺/H⁺ exchangers (NHEs), Na⁺-dependent HCO₃⁻ transporters of the SLC4 transporter family, or primary active transporters (e.g., V-type H⁺-ATPases) (21, 22).

Here, we investigate how changes in seawater pCO₂ affect the direct environment of the calcifying PMCs and whether sea urchin larvae are able to regulate extracellular pH in response to both acute and chronic seawater acidification. Furthermore, we study acid–base regulatory features of the PMCs to provide information on the extent and mechanisms of intracellular pH regulatory abilities. This knowledge is crucial to understand the mechanisms underlying the control of pH homeostasis in PMCs and, thus, calcification during environmental hypercapnia. We hypothesize that a higher fraction of energy spent on acid–base compensatory processes of PMCs in response to CO₂-induced seawater acidification may be a critical reason for the reported reductions in growth and development of echinoid larvae.

Results and Discussion

Lack of pH Regulation Within the Primary Body Cavity.

H⁺-selective microelectrodes were adapted for extracellular pH (pH_e) measurements in marine invertebrate larvae. Surprisingly, pH_e in the body cavity followed seawater pH (pH_sw) without any measurable signs of active pH_e compensation. This observation was independent of acute or chronic exposure to elevated seawater pCO₂ (Fig. 1). Additionally, pH_e was also monitored noninvasively in larvae loaded with the pH-sensitive dye 2′,7′-bis-(carboxyethyl)-5(6)-carboxyfluorescein (BCECF) in an experiment to validate our electrode measurements. These measurements confirmed the lack of significant pH_e regulatory ability (Fig. S1). A bias on pH electrode measurements was excluded by performing nonselective voltage electrode measurements. No transepithelial potential difference was recorded between the bath solution and extracellular matrix (Fig. S2). The lack of any difference in pH_sw vs. pH_e is surprising because it would indicate very shallow pCO₂ diffusion gradients from the extracellular compartment to the seawater. Typically, pCO₂ values in extracellular fluids of heterotrophic marine metazoans are at least 0.1 kPa higher than those of the surrounding seawater, thereby enabling diffusive CO₂ excretion across a delimitating barrier, thus rendering pH_e more acidic than seawater (23). The pluteus ectoderm is very thin (∼1–2 µm) (Fig. 2E), comparable in thickness to respiratory epithelia of, e.g., teleost fish (23), thus enabling shallow diffusion gradients for pCO₂. However, the relatively high pH_e within the body cavity in our measurements could also be related to an ectoderm that is leaky for ions and small molecules. Using fluorescein-5-isothiocyanate (FITC) dextran conjugates of different molecular weights, we studied the permeability of the larval ectoderm. We exposed four-armed pluteus larvae to FITC dextran-containing seawater (Fig. 2) and found that free FITC (M_r = 0.4 kDa) equilibrated within 15 min with the extracellular matrix (ECM). Significant entry of FITC dextran conjugates of 4 kDa (15-, 60-min incubation) and 10 kDa (60-min incubation) and the absence of entry of 40-kDa particles within the 60-min incubation period (Fig. 2 A and B) indicated a comparatively high permeability of the pluteus ectoderm. Because no significant increases in FITC-derived fluorescence were measured in the cytosol of ectodermal or PMC cells (Fig. 2B), it appears that equilibration between ECM and seawater is mediated through paracellular pathways. It is likely that the pluteus ectoderm is characterized by a high permeability for ions as well. This assumption is supported by the rapid equilibration of pH_e with pH_sw in our experiments: a change in 0.3 pH units in the seawater medium typically resulted in stable pH_e within ∼20 s (Figs. S2 and S3).

Fig. 1.

(A) S. droebachiensis pluteus larvae attached to the holding pipette and the tip of the measuring electrode inside the primary body cavity (PBC). (B) Relationship between seawater and the PBC pH (NBS scale) during acute and chronic environmental hypercapnia. Bars represent ±SD; n = 4–5.

Fig. 2.

Determination of epithelial permeability in pluteus larvae in vivo using FITC dextran conjugates of varying molecular weights and confocal microscopy. (A) Transmission image of pluteus larva; (B) confocal image of larva in a similar position as in A exposed to 4-kDa FITC dextran in seawater for 60 min; the green color in the extracellular matrix (ECM) indicates equilibration of FITC dextran between surrounding seawater (SW) and ECM; (C) confocal image of larva exposed to 40 kDa for 60 min; the dark color indicates lack of equilibration of FITC dextran between ECM and SW. (D) Equilibration of FITC dextrans of varying molecular weights between SW and ECM following 15- and 60-min incubation. Equilibration is represented as the ratio of FITC fluorescence within the ECM and the SW surrounding the larva. Low values indicate a low permeability. (E) Corresponding confocal (FM1-43-stained) and transmission images of the pluteus larva outer epithelium. Bars represent ±SD; n = 9.

Sea urchin larvae potentially sacrifice a certain degree of control over the abiotic conditions in the extracellular space to create favorable conditions for calcification (see discussion below). Vital stains using the dye FM1-43 indicate that the mesenchymal cells (PMCs, secondary mesenchyme cells) that populate the ECM are connected via a multitude of filopodia to each other and to cells of the ectoderm and the digestive tract (Fig. 3 and Fig. S4) (24, 25). These filopodial connections, which have been suggested to primarily aid in cell–cell signaling (e.g., refs. 26, 27), might also fulfill roles related to nutrient distribution that are classically mediated by the extracellular fluid in most adult invertebrates. This suggests that sea urchin larvae shift nutrient distribution to cellular syncytia to use the extracellular space more efficiently for the demands of ionic exchange processes relevant for calcification.

Fig. 3.

In vivo confocal images of pluteus larvae using the vital dye FM1-43 that stains membranes. Primary mesenchyme cell (PMC) syncytia and their sheaths, as well as filopodia connecting them with each other and epithelial cells are visible, as are vesicles within cells and filopodial connections. A is a close-up confocal image of the region depicted in the Inset in the transmission image (B). C depicts a similar region from a different larva as shown in A and B, but rotated by 90° clockwise.

Chronically Elevated Seawater pCO₂ Directly Affects the Sites of Calcification.

The present work demonstrates that, even under chronically decreased seawater pH, the primary body cavity conforms in pH with the environment. Under control conditions, a comparatively high pH_e (>8.1) in comparison with other marine invertebrates (28) might be beneficial for maintenance of the endoskeleton. Because the larval spicules are surrounded by sheaths of the PMC syncytium that do not fully isolate them from the extracellular medium (8, 15), high pH_e will enable a comparatively high CaCO₃ saturation state in the direct vicinity of the spicules. On the other hand, uncompensated pH_e during chronic acid challenge might lead to an increased energy demand for the PMCs to prevent dissolution of formed spicules within their sheaths when Ω decreases below 1. For example, Clark et al. (29) observed corrosion of larval spicules in Evechinus chloroticus and Pseudechinus huttoni pluteus larvae during exposure to pH 7.7 and 7.6, respectively. However, it has been demonstrated that sea urchin larvae are capable of maintaining calcification rates under acidified conditions when calcification rates are normalized to growth rate (4). Thus, pH conformity of the primary body cavity with the surrounding seawater suggests that PMCs themselves must have the ability to control intracellular pH, as well as pH in the microenvironment surrounding the spicules to at least partially protect the spicules from dissolution.

pH_i Regulatory Abilities of PMCs.

To test whether PMCs themselves are able to control pH_i—necessarily linked to calcification and skeletogenesis—we measured their pH_i regulatory capacity using the fluorescent pH-sensitive dye BCECF (Fig. 4 and Fig. S5). Using pulses of NH₃/NH₄⁺ solution, we demonstrated pH_i regulatory abilities in PMCs (Fig. 4A). Under control conditions (pH_sw 8.1), PMCs of five larvae were characterized by a control ratio of 1.41 ± 0.06, which corresponds to a pH_i of 6.9 as determined by nigericine calibration (Fig. 4B). pH_i values for sea urchin eggs a few minutes to hours postfertilization were reported to range between 6.8 and 7.3 depending on the method and species used (30, 31). The initial alkalization of fertilized sea urchin eggs has been hypothesized to be one of several signals inducing protein synthesis and cell functionality (32, 33). pH_i values for marine invertebrates were reported to range from 6.9 to 7.4 depending on species and methods used (34–38). The 20 mM NH₃/NH₄⁺ solution induced an intracellular alkalinization leading to an increase of pH_i by ∼0.2 ratio units corresponding to a ΔpH of 0.7. After washout of the NH₃/NH₄⁺ solution, pH_i rapidly dropped by ∼0.7 pH units below the control value, followed by a pH compensation reaction to control levels within 15 min (Fig. 4). Buffering capacity (β) (mmol⋅L⁻¹⋅pH unit⁻¹, slykes) of PMCs was calculated according to Graber et al. (39) by dividing the amount of acid load by the measured change of pH_i produced by this load. β values obtained for PMCs under control conditions were 20.83 ± 7.78 slykes (n = 9). This buffering capacity is comparable with that of other invertebrates, which were typically reported to be in the range of 16–40 slykes (37, 38, 40–42). Rates of pH_i compensation are about five to seven times slower than recovery times reported for strong ion regulatory cells of rat epithelia [2-3 min in colon or Henle’s loop cells (22, 43–45)] but comparable with the recovery rate of crayfish neurons exposed to a similar protocol [∼20 min (46)], indicating a significant acid–base regulatory capacity in echinoid PMCs.

Fig. 4.

Ratiometric fluorimetry in PMCs using the pH-sensitive dye BCECF-AM. (A) Schematic illustration of a recording trace including ratio images (Top: the dashed lines represent the orientation of spicules). Data were obtained from the control period (control), after addition and removal of NH₃/NH₄⁺ (alkalosis and acidification; ammonium pulse), and during pH_i recovery. (B) Calibration curve of BCECF-AM in PMCs fitted by a function that flattens toward more acidic or alkaline conditions allowing the translation of ratios to pH values.

Na⁺-Dependent pH_i Regulation in PMC.

The pH_i regulatory machinery has been widely characterized for various animal cell types and is essential for the maintenance of enzyme functionality and cellular processes (for a review, see ] 22). In general, cells maintain pH homeostasis by the import/export of protons and anionic buffers (e.g., HCO₃⁻) (47). Our current data clearly indicate that the compensation reaction after NH₃/NH₄⁺-induced acidosis in PMCs is highly dependent on Na⁺ and HCO₃⁻ transport (Fig. 5 and Table S1, including baseline pH, pH values after NH₃/NH₄⁺ pulse, and recovery rate). Compared with control conditions, the pH_i recovery rate was reduced by 68% in the presence of 5 mM Na⁺ and by 74% in a HCO₃⁻-free solution (Fig. 5 A and C and Table S1). This suggests the involvement of Na⁺-dependent HCO₃⁻ import mechanisms during pH recovery, which are an indirect sink of ATP. Transporters of the SLC4 family, including Na⁺/HCO₃⁻ cotransporters and Na⁺-dependent Cl⁻/HCO₃⁻ exchangers were widely identified as key players in this regulatory process (21, 48–50). The sea urchin PMC transcriptome contains a large number of genes coding for ion transporters including Na⁺/K⁺-ATPase (NKA), Na⁺/HCO₃⁻ cotransporters (NBC), H⁺-ATPases (HA), and NHEs (51). This indicates that PMCs possess the necessary molecular machinery to regulate pH_i by means of proton secretion and HCO₃⁻ accumulation. However, besides the importance of Na⁺-dependent HCO₃⁻ import in PMCs, the present work also suggests that amiloride-sensitive proton extrusion mechanisms are probably not the major compensation pathway in response to an intracellular acidosis. Although a significant reduction in the extent of recovery in pH_i was observed in the presence of 100 µM amiloride (to 45% of control pH_i/ratio) (Fig. 5B), the slope of the recovery reaction was not significantly different from that under control conditions (Fig. 5C). Because nonmammalian NHEs were demonstrated to be less sensitive to amiloride, with some NHEs being insensitive up to concentrations of 500 µM (52), this experiment cannot provide definitive answers to the role of NHE proteins in PMC pH regulation. However, because protons are generated during the calcification process within the PMCs (20, 53), the identification and characterization of other acid secretion mechanisms involving transporters such as the V-type H⁺-ATPase or H⁺/K⁺-ATPase is an important task in further investigations.

Fig. 5.

Acid–base regulatory abilities of PMCs in the presence of low (5 mM) Na⁺, without HCO₃⁻ and with 100 µM amiloride. (A) Original recordings of control and treatment traces representing average values of four to five single measurements. Presence of NH₃/NH₄⁺-induced alkalosis and the washout-induced acidosis under the three experimental conditions. (B) The recovery given in percentage of the initial value, before alkalosis in response to control and treatment conditions. (C) Recovery rate represented by the compensatory slope after acidosis given in percentage of the control (100%) value, in response to control and treatment conditions. Bars represent ±SD; n = 4–5.

Conclusion.

Gene expression analysis of sea urchin larvae reared under acidified conditions revealed an increased expression of NKA α-subunit, the key enzyme that provides the electrochemical gradient for most secondary active transport processes (4, 54, 55). This up-regulation of NKA is accompanied by an increase in metabolic rate, indicating a higher energy demand during environmental hypercapnia (10, 54). Because up to 77% of larval metabolism in sea urchin pluteus larvae is required to fuel the energetic demands of NKA under control conditions (56), it is likely that additional acid–base stress will significantly impact the organism’s energy budget. We suggest that a higher fraction of energy is spent on PMC acid–base regulation because these have to maintain full cell functionality in a more acidified extracellular medium. Thus, calcification during environmental hypercapnia could impact energy allocation. Because vesicular ACC precipitation occurs intracellularly in PMCs (8), the processes of pH_i regulation and calcification are intrinsically linked: membrane transporters that are involved in ACC precipitation depend on H⁺ and HCO₃⁻ concentrations in the cytosol, which in turn depend on the action of the pH_i regulatory machinery (summarized in Fig. 6). In this way, our data support the concept that maintenance of calcification rates in sea urchin larvae under elevated seawater pCO₂ is primarily an energetic, rather than a physicochemical problem. This helps to explain the observed phenomenon of an altered energy allocation under acidified conditions. Decreases in larval developmental rates and delays in metamorphosis will likely result in a higher mortality during the planktonic life phase (57–59). Furthermore, because the energy reserves required by the juvenile sea urchin to support the first weeks after metamorphosis are accumulated by the larvae (60), energetic compromises faced by larval stages may translate into juvenile fitness problems and thus will define the long-term fate of these keystone species in future marine habitats.

Fig. 6.

Schematic model summarizing the interplay of calcification, pH regulation, and energetic costs in sea urchin larvae during environmental acidification. The primary body cavity (PBC) pH conforms to the surrounding seawater pH (pH_sw). A decrease in pH_e directly affects the calcifying primary mesenchyme cell (PMC) syncytia challenging the intracellular pH (pH_i) regulatory machinery due to decreased proton gradients. The vesicular precipitation of amorphous calcium carbonate (ACC) within PMCs is intrinsically linked to pH_i regulation. pH homeostasis is maintained by ion transporters, which either directly or indirectly depend on the consumption of energy. Increased energetic costs due to decreased proton gradients lead to a shift in the larvae’s energy budget, which decreases scope for growth and may also translate into juvenile impaired fitness. ST, stomach; putative transporters are in gray.

Materials and Methods

Sea Urchin Larvae Culture and CO₂ Perturbation Experiment.

All animal experiments were performed according to the German law for animal welfare and were approved by the animal welfare officer of the Christian Albrechts University, Kiel. Adult Strongylocentrotus droebachiensis were collected in Winter 2010 and 2011 in the Kattegat (Dröbak, Norway) by divers. Spawning was induced by injection of 2 mL of 0.5 mM KCl into the coelomic cavity. For each experiment, eggs of one to two females were collected in separate 1-L beakers, washed, and fertilized by adding dry sperm (20 µL) of two males. Fertilization was followed by monitoring the fertilization-induced elevation of the oocyte membrane under a stereomicroscope (fertilization rates, >95%). Zygotes were allowed to divide once before they were pooled, concentrated in 25 mL, and split into 2-L (three replicates per pCO₂ treatment, Erlenmeyer flasks) culture vessels, which were preequilibrated at three different pCO₂, control (∼0.05 kPa, pH 8.1), intermediate (0.12 kPa, pH 7.7), and high pCO₂ (∼0.24 kPa, pH 7.4) by a central automatic gas mixing facility (Linde Gas, HTK Hamburg, Germany). Water pH_NBS, salinity, and temperature were monitored daily in the setup during the entire incubation period. Seawater samples (0.5 L) for determination of total dissolved inorganic carbon, C_T, were collected weekly. C_T was determined using an AIRICA analyzer (Marianda). C_T was corrected using certified reference material provided by Andrew Dickson (Scripps Institution of Oceanography, La Jolla, CA) (61). Seawater carbonate system speciation was calculated from pH_NBS and C_T with the open-source program CO2SYS (62) using the dissociation constants by Mehrbach et al. (63) as refitted by Dickson and Millero (64). Water parameters measured during the experimental period are summarized in Table S2.

Larval culturing and monitoring was essentially conducted as described in previous studies (10, 54).

Selective Ion Electrode Technique.

Selective ion electrode measurements were essentially performed after Kuehl and Revsbech (65) to measure H⁺ concentrations in the extracellular gel matrix (ECM) of sea urchin larvae. Glass capillary tubes (borosilicate; inner diameter, 1.2 mm; outer diameter, 1.5 mm; Hilgenberg) were pulled on a DMZ-Universal puller (Zeitz Instruments) into micropipettes with tip diameters of 2–3 µm. These were then baked at 200 °C for 4 h and vapor-silanized with dimethyl chlorosilane (Sigma-Aldrich) overnight. The micropipettes were front-loaded with a 200-µm column of liquid ion exchanger mixture (H⁺ ionophore III; Sigma-Aldrich) diluted in 2-nitrophenyl ether at a concentration of 10.5 mg⋅mL⁻¹. Additionally, micropipettes were again front-loaded with a 100-µm column of the ionophore mixture containing a polyvinylchloride/tetrahydrofuran (330 mg⋅mL⁻¹) solution with a ratio of 1:3 to seal the opening of the electrode tip. The micropipette was backfilled with a 4-cm column of pH electrolyte solution (300 mM KCl, 50 mM NaPO₄, pH 7) to create an ion selective microelectrode (probe). To calibrate the ion-selective probe, the Nernstian property of each microelectrode was measured by placing the microelectrode in a series of artificial seawater (ASW) solutions (pH 6, 7, 8, and 9) with reference to an Ag/AgCl electrode. By plotting the voltage output of the probe against log [H⁺] values, a linear regression yielded a Nernstian slope of 49.8 ± 2.3 mV (n = 12) for 1 pH unit. With this method, we were able to resolve a minimum difference of 0.1 pH units (corresponding to a change in pCO₂ of ∼0.02–0.04 kPa). Electrode response times in the perfusion solution (0.79 ± 0.21 s⋅nM [H⁺]⁻¹) and within the primary body cavity (1.75 ± 0.42 s⋅nM [H⁺]⁻¹) were calculated by the time needed to reach 95–100% of the fully stabilized value after a pH change (usually 0.3 or 0.4 pH units between pH 8.1, 7.7, and 7.4) in the perfusion bath.

Measurement of Extracellular H⁺ Concentrations.

The electrode measurements were performed at 10 °C on an inverse microscope (Axiovert 135; Zeiss) equipped with a temperature-controlled perfusion system, allowing the quick exchange of different pH solutions inside the perfusion chamber. The pH of the bath solutions was adjusted by equilibrating ASW with pure CO₂ to the desired pH. The larva was placed inside the perfusion bath and kept in position via a holding pipette (30- to 40-µm tip diameter) to which a slight vacuum was applied. The ion-selective probe was mounted on a remote-controlled micromanipulator (Phytron) and was introduced into the ECM from the base of the arms at the oral side of the larvae (10–20 µm inside the ECM) to record the ionic activities (Fig. 1A). The epidermis and basal lamina formed a seal around the entrance point of the electrode, preventing fluid exchange between seawater and the primary body cavity. pH recordings were performed on pluteus larvae [8–10 d postfertilization (dpf)] reared under control conditions, which were exposed to acute changes in seawater pH ranging from 6.9 to 8.2. Furthermore, the same measurements were also performed on pluteus larvae (8–10 dpf) from the CO₂ perturbation experiment to address the effects of chronically elevated seawater pCO₂ on extracellular pH homeostasis.

Visualization of Mesenchyme Cell Filopodial Network.

Four-armed pluteus larvae (96 h) were incubated for 15 min in control (pH 8.1) seawater with 20 µM FM1-43 at 10 °C. Following incubation, larvae were placed on microscopy slides suspended in the incubation solution and covered with glass coverslips supported by ∼100-µm-thick fibers as spacers. Larvae were then imaged with a Zeiss LSM510 using a Plan Neofluar 40×/1.3 objective and a 505-nm long-pass filter at an excitation wavelength of 488 nm. Images were collected within 20 min (n = 20) at room temperature (20 °C).

Epithelial Permeability Measurements with FITC Dextran.

To measure epithelial permeability, four-armed pluteus larvae (96 h) were incubated for 15 and 60 min in control (pH 8.1) seawater with 0.4 mM FITC dextran (free FITC, 0.4, 4, 10, 40 kDa; Sigma-Aldrich) at 10 °C. FITC dextran solutions were dialyzed overnight in ASW in MWCO-dialysis tubing (Roth) before experiments. Following incubation, larvae were imaged on a Zeiss LSM510 as described above. Images were collected within 10 min (n = 9 larvae per treatment) at room temperature (20 °C).

Bag Isolation Procedure and Dye (BCECF-Acetoxymethylester) Loading.

To measure PMC intracellular pH (pH_i), epidermal cells of 8–10 dpf larvae were removed according to Harkey and Whiteley (66). Using this protocol, both epithelial as well as PMCs can be maintained viable for hours (66). Larvae were incubated at 10 °C with a final concentration of 50 μmol L⁻¹ BCECF-acetoxymethylester (BCECF-AM) for 30 min to allow sufficient uptake and cleavage of the esterified dye by intracellular esterases. Thereafter, only larvae, which were firmly attached to the perfusion bath bottom, were used for measurement. The flow rate of the perfusion solutions was 2–3 mL⋅min⁻¹. With this dye-loading procedure, we achieved a signal-to-noise ratio for the emission signal of >10 throughout an experimental period of >2 h without appreciable loss of signal intensity.

Solutions.

ASW solutions were mixed according to Zeebe and Wolf-Gladrow (67). Osmolality (980 ± 10 mosm⋅kg⁻¹) and salinity (31 ± 1) were chosen to match the natural seawater used in the larval cultures. All ingredients of the experimental solutions are given in Table S3. Amiloride was added at a final concentration of 100 μM to the ASW. DMSO did not exceed a concentration of 0.1%.

Microfluorimetry.

Fluorescence was monitored with an imaging system (Visitron). The dye was alternatively excited at a rate of 0.2 Hz at 486 and 440 nm (±10-nm bandwidth) for 24 and 60 ms, respectively. Emission was recorded at 525 nm (±13-nm bandwidth), and the integrated ratio of the emission intensities at the two excitation wavelengths over the whole cell was calculated after subtraction of system immanent camera offset and background signal (MetaFluor Software 7.6.1; Molecular Devices). From each larva, the recordings of four to five PMCs were averaged and treated as one replicate (n = 1). Four to five control and treatment larvae were measured in an alternate mode and used for further analysis. Nigericin was used to calibrate pH_i of living cells as previously described by Suffrian et al. (68). PMCs of pluteus larvae were exposed to 10 μmol⋅L⁻¹ nigericin in the presence of 150 mmol⋅L⁻¹ K⁺ at pH 5.5, 6.5, 7, 7.5, and 8.5. This [K⁺]_i is in the same range reported for fertilized and unfertilized sea urchin eggs (69) and intestinal tissues of the sea urchin S. droebachiensis (70). The calibration curve (Fig. 4) allowed estimation of the relationship between detected emission ratio of BCECF and the respective pH_i. A change of 0.1 ratio units corresponded to a change of 0.35 pH units.

For all experiments, the bath was exchanged at a rate of 2–3 mL⋅min⁻¹ at 10 °C, and pH_i was continuously monitored. All larvae were exposed to ASW followed by the 20 mM NH₃/NH₄⁺ prepulse. Acidosis was induced by NH₃/NH₄⁺ washout using the respective solutions, 5 mM Na⁺,HCO₃⁻-free solutions and ASW plus 100 μM amiloride or control solutions (e.g., ASW or ASW plus DMSO), which were directly applied to the larvae via the perfusion system.

Traces of intracellular pH measurements were analyzed according to the parameters depicted in Fig. 3A. To determine the ability of PMCs to recover from an NH₃/NH₄⁺-induced acidosis, we calculated the ratio of the recovery value (in the stationary phase of the recovery phase) and the initial control value. Additionally, the slope of the compensation reaction was used to characterize the rate of recovery after an induced acidosis by determining the change in the ratio as a function of time.

Statistical Analysis.

Statistical differences between response times of ion-selective electrode measurements and pH regulatory abilities of PMCs were analyzed with Student t test with significance levels of P < 0.05 (*) and P < 0.01 (**).