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Proc Natl Acad Sci U S A. Sep 29, 2009; 106(39): 16574–16579.
Published online Aug 31, 2009. doi:  10.1073/pnas.0902894106
PMCID: PMC2757848
From the Cover
Applied Biological Sciences

Imaging intracellular pH in a reef coral and symbiotic anemone

Abstract

The challenges corals and symbiotic cnidarians face from global environmental change brings new urgency to understanding fundamental elements of their physiology. Intracellular pH (pHi) influences almost all aspects of cellular physiology but has never been described in anthozoans or symbiotic cnidarians, despite its pivotal role in carbon concentration for photosynthesis and calcification. Using confocal microscopy and the pH sensitive probe carboxy SNARF-1, we mapped pHi in short-term light and dark-incubated cells of the reef coral Stylophora pistillata and the symbiotic anemone Anemonia viridis. In all cells isolated from both species, pHi was markedly lower than the surrounding seawater pH of 8.1. In cells that contained symbiotic algae, mean values of pHi were significantly higher in light treated cells than dark treated cells (7.41 ± 0.22 versus 7.13 ± 0.24 for S. pistillata; and 7.29 ± 0.15 versus 7.01 ± 0.27 for A. viridis). In contrast, there was no significant difference in pHi in light and dark treated cells without algal symbionts. Close inspection of the interface between host cytoplasm and algal symbionts revealed a distinct area of lower pH adjacent to the symbionts in both light and dark treated cells, possibly associated with the symbiosome membrane complex. These findings are significant developments for the elucidation of models of inorganic carbon transport for photosynthesis and calcification and also provide a cell imaging procedure for future investigations into how pHi and other fundamental intracellular parameters in corals respond to changes in the external environment such as reductions in seawater pH.

Keywords: cell biology, cnidaria, symbiosis, photosynthesis, climate change

Symbiotic cnidarians form the structural foundation of coral reefs, which rank among the most biodiverse and ecologically important ecosystems. Despite their environmental significance, key elements of coral physiology such as calcification and the symbiotic interactions between cnidarians and their intracellular algae (Symbiodinium) are poorly understood, largely due to knowledge gaps in fundamental aspects of cnidarian cell biology (13).

Intracellular pH (pHi) modulates virtually all aspects of cell metabolism, including membrane channel functioning, the structural and functional properties of enzymes, intracellular signaling, and ion availability (4, 5). As a consequence, pHi must be tightly controlled and changes in pHi are minimized by intracellular buffers and regulation by ion carriers on the cell membrane (6, 7). In symbiotic cnidarians such as corals, pHi is thought to be linked to processes of photosynthesis and calcification by influencing the flux of ions between host, symbiont, and the surrounding environment (6, 8). This particularly concerns the transport of dissolved organic carbon (DIC) between cell layers and the speciation of DIC between CO2, HCO3, and CO32− (911). Measurements of pHi in cnidarian cells are therefore an essential step for elucidating how DIC is transferred from the surrounding seawater to the respective sites of photosynthesis and calcification.

Understanding the links between pHi and processes of calcification and photosynthesis is becoming increasingly important given the challenges corals and symbiotic cnidarians face under global climate change (12). Oceanic absorption of atmospheric anthropogenic CO2 is decreasing both the availability of carbonate ions and seawater pH potentially compromising coral calcification (1315), while rising sea surface temperatures affect photosynthesis and cause dysfunction in the symbiotic relationship between corals and their intracellular algae (bleaching) (16, 17). Quantification of pHi is a key step toward determining the mechanistic basis behind the vulnerability of these physiological processes to environmental change.

Progress in the understanding of the physiology of symbiotic cnidarians like corals depends on the development of tools to examine their cell biology (2). In many other biological systems understanding of pHi regulation and other physiological parameters has improved significantly by the application of intracellular probes that emit fluorescence at 2 wavelengths which can be calibrated to the concentration of specific ions in the cell (e.g., H+) (1820). When coupled with confocal microscopy, this approach allows the mapping of dynamic physiological processes across cell structures (21, 22) and has enormous potential utility in physiological studies of symbiotic cnidarians, which harbor intracellular algae and thus require excellent spatial resolution to distinguish between animal and algal compartments.

In the current study, we adapted a live cell imaging procedure that allowed us to make the first pHi measurements in symbiotic cnidarians and anthozoans. To do this, we observed endoderm cells from the reef coral Stylophora pistillata and the symbiotic anemone Anemonia viridis exposed to both light and dark conditions. pHi was determined by loading cells with the pH sensitive fluorescent probe, carboxyseminaphthorhodafluor-1 (SNARF-1), and analysis by confocal microscopy. Our in vivo cell imaging procedure was developed to allow spatial separation of cnidarian host and symbiont cell compartments and has a wide range of potential applications in the study of coral and cnidarian physiology.

Results

Observation of Cell Preparations.

Cell preparations isolated from A. viridis and S. pistillata included a variety of cell types including endoderm cells that contained 1 to 3 algae and cells without algal symbionts (Fig. 1).

Fig. 1.
Cnidarian endoderm cells isolated from A. viridis with and without symbiotic algae. (Row A–D) Different cell types: (A) endoderm cells without symbiotic algae. (B) endoderm cells containing one alga. C = animal cytoplasm A = intracellular symbiotic ...

Cells that contained 2 algae were the most suitable for measurements of pHi in algae-containing cells for 2 reasons. First, this cell type contained a larger area of host cytoplasm available for pHi measurements than single alga cells [in which the cytoplasm was closely stretched around the alga (Fig. 1, RowB)] and secondly, double cells were more numerous in cell preparations than cells that contained 3 algae.

In the case of A. viridis preparations, cells with no intracellular algae could be identified to be of endodermal origin because the isolation procedure allowed separation of this layer from other host tissues. The presence of these cells allowed comparisons of pHi to be made between endoderm cells with and without intracellular algae.

Analysis by TEM and staining with Hoecsht 33342 and rhodamine 123 identified the presence of nuclei and mitochondria in all cell types, which were generally located between the algal cells in host cells with 2 or more algae (Fig. 1, Column 2). The cytoplasm of host cells stained evenly with SNARF-1 with little or no exclusion or concentration of the dye within cell compartments such as nuclei or mitochondria (Fig. 1, Column 3), although compartmentalization of SNARF-1 began to occur in mitochondria and vacuoles after prolonged periods of greater than 90 min in perfusion chambers.

Intracellular pH in Coral and Anemone Cells.

pHi was measured by ratiometric analysis of SNARF-1 fluorescence at 585 and 640 nm within a region of interest (ROI) in host cytoplasm that did not overlap with algae in x, y, and z planes of each cell. Calibration curves of pHi with the SNARF-1 signal are given for S. pistillata and A. viridis as supporting information (SI) Fig. S1.

Analysis of vertically-stacked horizontal slices through the cell (z stack profiles) showed that pHi was stable (± pH 0.1) toward the center of most cells where SNARF-1 fluorescence was greatest (Fig. 2). Exceptions to this rule occurred with the presence of acidic vacuoles which had a pH of 6 or below the limits of our calibration.

Fig. 2.
Z (cell depth) profile of an A. viridis endodermal cell. (A) Representative Z stack slices with cell depth. White circle = ROI in the host cytoplasm. (B) Fluorescence intensity of SNARF-1 at 585 (circles) and 640 nm (triangles). (C) pHi with cell depth. ...

Using this approach, pHi was determined in endoderm cells of S. pistillata and A. viridis that had been incubated for 20 min under light or dark conditions (Table 1). In both species, pHi was significantly higher in light-incubated cells that contained algal symbionts than dark-incubated symbiont-containing cells (one way ANOVA, F5,114 = 8.135, P < 0.001). However, when light and dark-incubated symbiont free cells isolated from A. viridis were compared, no significant difference in pHi was found (Table 1). In both light and dark conditions pHi was not significantly different in coral relative to anemone cells.

Table 1.
Intracellular pH in S. pistillata and A. viridis endoderm cells treated with 20 min dark or light incubation following loading with the pH sensitive dye SNARF-1 for 30 min in the dark. Superscripted letters indicate homogeneous subsets as determined by ...

Manipulation of pHi by Perfusion with NH4Cl.

To determine the sensitivity of SNARF-1 fluorescence signal to induced pH changes, the pHi of algal-containing A. viridis endoderm cells were experimentally manipulated by short term perfusion of 20 mM NH4Cl. Perfusion with NH4Cl significantly increased pHi by approximately 0.4 pH units (one way ANOVA, F2, 16 = 10.90, P < 0.01) and returned to previous levels 10 min following removal of NH4Cl by washing with filtered seawater (FSW) (Fig. 3).

Fig. 3.
The impact of NH4Cl on pHi (mean ± SE) in endoderm cells in FSW isolated from A. viridis.* indicates significant difference to control as determined by Tukey's post hoc analysis, following one way ANOVA.

Spatial Patterns in pHi.

Analysis of Z stack images revealed that the principle source of pH heterogeneity within the cell was associated with the presence of symbiotic algal cells. In algae-containing endoderm cells isolated from both A. viridis and Stylophora pistillata the immediate area surrounding each alga was at lower pH than the rest of the host cytoplasm, at pH 6 or below the limits of our calibration using SNARF-1 (Fig. 4). The region of low pH was observed in both light and dark-incubated endoderm cells.

Fig. 4.
Ratiometric image of SNARF-1 emission (585 and 640 nm) in endoderm cell isolated from A. viridis. C = animal cytoplasm. A = Algal symbiont. Note: Red = algal autofluorescence and is not pH related.

Discussion

This is the first study to report intracellular pH (pHi) in anthozoans and symbiotic cnidarians and the first to use live cell confocal imaging in concert with a dual emission probe to monitor the physiology of coral and anemones. pHi has previously been determined in most model organisms that are routinely used for cell and developmental biology. However, data on pHi and many other physiological parameters are still lacking for cnidarians in general, despite growing interest in the cell biology of this group in both the fields of marine ecology and evolutionary-developmental biology (2, 23). Part of the reason for this lack of data lies in difficulties in obtaining viable cnidarian cells in continuous culture (2, 24, 25) and the limited number of studies that have undertaken imaging of physiological parameters in isolated living cells from corals and sea anemones (26, 27). Cell imaging is particularly challenging in symbiotic systems where the autofluorescence derived from the chlorophyll of intracellular algae can confound signals from many commercially available fluorescent dyes and the fact that algal cells occupy the majority of the cell volume leaving limited areas of animal cytoplasm to work with (28). The current study overcame these problems using confocal imaging to analyze each cell in 3D (Z-stack profile) and restrict pH determination to the regions of interest identified in host cytoplasm that did not overlap with algal symbionts. This approach allows the monitoring of intracellular parameters (in this case pHi) in both algal symbiont-containing and symbiont free cells. Overall, this work paves the way for future studies on symbiotic cnidaria utilizing the wide range of ratiometric fluorescent dyes designed to investigate numerous aspects of cell function including signal transduction, membrane potential, and the flux of metal and nonmetallic ions (29).

Values of pHi obtained for the reef coral S. pistillata (dark: 7.13 ± 0.24 and light: 7.41 ± 0.22) and the symbiotic sea anemone A. viridis (dark: 7.01 ± 0.27 and light: 7.29 ± 0.15) fall within the range typically observed for marine invertebrates, with low values having been observed in unfertilized sea urchin eggs (pH 6.7) (30) and higher values obtained in ascidian eggs (pH 7.6) (31). Two findings support the validity of our pHi measurements. First, perfusion experiments with NH4Cl cells displayed the classic pattern of alkalinization of pHi and return to previous pHi levels on removal of NH4Cl; this pattern has been observed before in more conventionally used cell types (21, 32). Secondly, the fact that light or dark treatment had no effect on pHi in A. viridis cells without algal symbionts excludes the possibility that differences observed in the SNARF-1 ratio between light and dark-symbiont-containing cells were caused by direct effects of light on the fluorescence behavior of the dye.

The pHi values obtained in the current study are markedly lower than the surrounding seawater pH of 8.1 revealing that coral and anemone cells maintain a strong gradient of pH between the outside medium and the cells. This finding and the observation that pHi was higher in light-incubated relative to dark-incubated symbiont-containing-cells have important implications when considering 2 processes that underpin coral reef ecosystems: photosynthesis and calcification. Cnidarian hosts supply DIC to their algal symbionts via a mechanism of DIC uptake and transport about which little is currently known (11, 33, 34). CO2 has been proposed as one species of DIC that enters hosts cells from the external environment (34, 35). Using the pHi measurements in the current study and a previous measurement of intracellular bicarbonate concentration [3.7 mM at minimum, (36)], it is possible to estimate the intracellular host CO2 concentration using the Henderson-Hasselbalch equation: pH = pK +log [HCO3]/[CO2] with pK = 6.1 (37). This would indicate that if CO2 were to enter host cells by diffusion, as proposed by previous authors (34, 35), external CO2 concentrations would have to exceed a minimum of ca. 460 μM to achieve diffusion into host cells.

Although the route by which DIC enters host cells is not clear, the majority of host intracellular DIC would be predicted to be HCO3 at the values of pHi determined in the present study. Previous authors have proposed that within endodermal cells that contain algae, the carbon requirements of algal photosynthesis are supplied by dehydration of host cell HCO3 to CO2 by carbonic anhydrase close to the symbiotic algae resulting in the production of OH (10). Previous observations that light-incubated endoderm cell layers excrete OH leading to alkalinization of the cnidarian coelenteron cavity (10, 38) are supporting evidence for this mechanism. As the production of OH within endoderm cells containing symbionts could be predicted to increase host cell pHi, the finding in the current study that pHi was significantly higher in light-incubated than dark-incubated symbiont-containing cells is important new evidence that also supports this proposed mechanism.

Close inspection of the interface by ratiometric imaging between the algae and host cell cytoplasm revealed that pH was low (<6) in the immediate area surrounding each alga in both dark and light-incubated cells of both species. One possibility is that this region is associated with the symbiosome membrane complex (39), as previous research indicates that the symbiosome in A. viridis could be as low as pH 5.7 (40), and symbiosomes in other symbiotic associations have also been found to be at low pH (41). Symbiosome pH is important to determine given the potential role of the symbiosome in the trafficking of compounds between host and symbiont, but the current data do not definitively show whether the low pH region is associated with the symbiosome membrane complex or the adjacent host cytoplasm. Future work using pH sensitive dyes with a lower pKa and recently developed symbiosome specific probes (42) have the potential to resolve this issue.

Turning to processes of calcification in corals, our findings must be taken into account when deciphering how ions are transported to and from the site of calcification. In particular, our results have implications for the speciation of carbon used for calcification. At the cnidarian pHi determined in the current study, the intracellular availability of CO32− relative to HCO3 would be even lower than predicted in previous studies (43), thus precluding the direct secretion of CO32− from cells at the site of calcification. To confirm this, pHi must be determined in calicoblastic cells. This research will be greatly facilitated by the recent development of protocols to maintain primary cultures of this cell type (25).

In conclusion, we report measurements of pHi in symbiotic cnidarian cells using a cell imaging procedure that paves the way for future investigations into the regulation of pHi and other physiological parameters in corals and other cnidarians. Given the importance of pHi in most elements of cell function, the observed differences in pHi between light and dark conditions may be a key regulator of cell physiology in symbiotic cnidarians. This possibility requires further research into pHi dynamics, building on the single time point measurements made in the current study, for a more compete understanding of the interactions between host pHi and photosynthesis. An additional research priority must also be to determine how changes in external parameters such as seawater acidification impact pHi in calicoblasts and other cell types. The information arising from investigations such as these will be essential for determining the mechanistic basis behind vulnerability of coral physiology to aspects of global environmental change and improving predictions about the future status of coral reefs.

Materials and Methods

Coral and Anemone Culture.

Colonies of S. pistillata and individuals of A. viridis were maintained at the Centre Scientifique de Monaco in aquaria supplied with flowing seawater from the Mediterranean Sea (exchange rate 2% h−1) at a salinity of 38.2 ppt, pH 8.1 ± 0.1 under an irradiance of 300 μmol photons m−2 s−1 (S. pistillata) or 100 μmol photons m−2 s−1 (A. viridis). Temperature was maintained at 25 ± 0.5 °C for S. pistillata and 21 ± 1 °C for A. viridis. Corals and anemones were fed 3 times a week with both frozen krill and live Artemia salina nauplii.

Preparation of Cells.

Branch tips of S. pistillata were removed using surgical bone cutters and the tissue removed by gentle brushing with a soft bristle toothbrush in 0.45 μm FSW. For A. viridis, cells were prepared by scraping endoderm cells from longitudinally sectioned tentacles and suspending in FSW (44). Suspensions of anemone or coral cells were then filtered through a 40 μm mesh made up to a volume of 45 ml and centrifuged at 350 g, for 4 min, at 20 °C. For confocal microscopy, the resulting pellets were resuspended in 2 ml of FSW to obtain a density of approximately 106 cells ml−1 and loaded into perfusion chambers that were constructed from silane slides with glass coverslips mounted on thin runners of scotch tape sealed to the slide on 2 sides by silicone grease (45). In the following procedures exposure of cells to fluorescence dyes and buffer solutions was achieved by perfusion, allowing the elimination of further centrifugation steps which reduce the yield of viable endoderm cells. Viability staining using a Live/Dead Viability Kits (Invitrogen) confirmed cells remained viable in perfusion chambers for at least 4 h in FSW.

Dye Loading.

Loading of SNARF-1 into cnidarian cells was achieved using the cell permeant acetoxymethyl ester acetate of SNARF-1 (SNARF-1 AM). Solutions were prepared from 20 mM stock in DMSO with working concentrations at 10 μM SNARF-1 a.m., 0.01% Pluronic F-127, and 0.1% DMSO in FSW for 30 min dark incubation at 21–24 °C. Concentrations fell within the range used in previous investigations (18). Inspection of cells treated with 5 μM digitonin in FSW to permeablize the cell membrane confirmed that any unhydrolyzed SNARF-1 a.m. did not accumulate with cell compartments.

Host Cell Nuclei and Mitochondria Were Labeled by Incubation in 10 μg/ml Hoechst 33342 and 5 μM Rhodamine 123 in FSW for 10 min. After dye loading, cells were washed with 2 ml FSW. All dyes used for microscopy were purchased from Invitrogen. Other chemicals were purchased from Sigma-Aldrich.

Confocal Microscopy.

Confocal microscopy was conducted using a Leica SP5 confocal microscope (Leica) equipped with UV and visible laser lines. Cells loaded with SNARF-1 a.m. were excited at 543 nm at a laser power at 25%, scan speed 400htz, and pinhole set at 1.51 units using a X100-fold oil immersion lens. Each cell was analyzed by capturing 15 frames in the x/y plane, throughout the depth (z plane) of the cell (z-stack profile). Prior optimization of settings ensured no loss of the SNARF-1 signal by photobleaching during the 12 sec required for acquisition of each z-stack. SNARF-1 fluorescence emission was captured in 2 channels at 585 and 640 ± 10 nm whilst simultaneously monitoring in transmission. As SNARF-1 a.m. has been shown previously to be responsive to temperature (46); temperature was maintained between 22 and 24 °C during observation of cells.

In cells containing algal symbionts, the use of 543 nm as the excitation wavelength minimized chlorophyll autofluorescence, as 543 nm lies outside of absorption spectra of chlorophyll a and in a low region of absorption of the Peridinin-Chlorophyll-Protein complex (47).

Cell nuclei stained with Hoechst 33342 were visualized using UV excitation with image capture at 450 nm ± 10 nm. Mitochondria stained with Rhodamine 123 were visualized by excitation at 496 nm and captured at 530 nm ± 10 nm and in transmission. When imaging host nuclei and mitochondria, autofluoresence of intracellular algae was captured in a separate channel at 670 ± 10 nm and used to identify the position of algal symbionts within the cell.

Light and Dark Incubation of Cells.

After dark loading with SNARF-1 and perfusion with 2 ml FSW (described above), cell preparations were incubated for a further 20 min in the dark or under illumination from a fiber optic light source (Bioblock Scientific, SFBN Mechanik GmbH). The duration of light incubation was chosen based on previous research showing that light-dependant increases in coelenteron pH reached a maximum after 20 min light exposure in tentacles of A. viridis (38). During light incubation, A. viridis or S. pistillata cells were exposed to levels of photosynthetically active radiation (PAR) that matched the conditions under which the anemones and corals were cultured (100 and 300 μmol photons m−2 s−1, respectively). During confocal analysis, light-incubated cells continued to receive light while locating and moving between cells in the preparation, but the light source was turned off during the 12 sec required for each z-stack profile acquisition. All pH measurements were performed within 20 min following light or dark illumination, with no loss of SNARF-1 signal in either 585 and 640 nm channels.

Calibration of pH in Cnidarian Cells.

Measurements of intracellular pH were performed by ratiometric analysis of SNARF-1 fluorescence (48). The ratio of fluorescence intensity of SNARF-1 at 585/640 nm was calibrated in vivo in the range pH 6–8.5 by washing endoderm cells in calibration buffer adjusted to the desired pH by the addition of HCl or NaOH. The ionophore nigericin was added at 30 μM to equilibriate pHi with the surrounding media and eliminate pH gradients within the cell (18). Calibration buffers contained approximate cytosolic concentrations of major ions in cnidarian cells (49, 50) (60 mM Na+, 200 mM K+, 190 mM Cl, 25 mM Pipes (pH 6- 7.5) or Tricine (pH 8–8.5). Mannitol was added to adjust osmolarity to 1100 mosm l−1.

Calibration curves of pHi in S. pistillata and A. viridis endoderm cells are provided as Fig. S1.

Image Analysis.

After inspection of each cell in x, y and z planes, SNARF-1 fluorescence intensity at 585 and 640 ± 10 nm was recorded within an ROI selected in an area of host cytoplasm that did not overlap with algal cells. Examination of cells not loaded with SNARF-1 confirmed that algal autofluorescence was not detectable in ROIs in this location. R was calculated after subtracting background fluorescence recorded in a second ROI in the surrounding cell media.

The 585/640 nm fluorescence intensity ratio (R) was related to pH by the following equation:

equation image

where (F) is fluorescence intensity measured at 640 nm (λ2) and the subscripts A and B represent the limiting values at the acidic and basic end points of the titration respectively (i.e., pH 6 and 8.5).

Procedures used for calibration and image analysis were validated by checking the methods on a mammalian cell line (Murine C2C12 mesenchymal cells) in PBS buffer. pHi was determined to be in the range of pH 7–7.2 which fits the values determined previously for murine cells (e.g., 46).

Manipulation of Intracellular pH by NH4Cl.

Endoderm cells were suspended in FSW and dark loaded in perfusion chambers as described above and pHi determined (n = 8 cells). Cells were then perfused with FSW containing 20 mM NH4Cl and left for 10 min before pHi was determined again. NH4CL was then washed out by perfusion with 5 ml FSW, left for 10 min and pHi determined for a third time.

Transmission Electron Microscopy (TEM).

Fixation and observation of samples for TEM was carried out by methods described in ref. 51. Briefly, cell pellets were fixed at 4 °C with 4% glutaraldehyde in 0.085 M Sorensen phosphate buffer at pH 7.8 with 0.5 M sucrose. Samples were rinsed in Sorensen buffer, then postfixed for 1 h at ambient temperature with 1% osmium tetroxide in Sorensen phosphate buffer. Samples were dehydrated by transfer through a graded series of ethanol ending with a concentration of 100%. Following dehydration, samples were embedded in Epon resin. Ultrathin sections were cut with a diamond knife, transferred to copper grids coated with Formvar, stained with uranyl acetate and lead citrate, rinsed in distilled water, and observed with a CM12 Phillips TEM at 80 kV at the Centre Commun de Microscopie Appliquée at the University of Nice-Sophia Antipolis, Nice, France.

TEM examination of cells immediately after centrifugation, following loading with SNARF-1 a.m. and 90 min post SNARF-1 a.m. loading, confirmed there were no impacts of these preparation steps on cell ultrastructure.

Supplementary Material

Supporting Information:

Acknowledgments.

We thank Evelyn Houliston and Jonathan Erez for advice on microscopy, Sophie Pagnotta for assistance with TEM, and Mia Hoogenboom and 2 anonymous reviewers for comments on the manuscript. This research was funded by the government of the Principality of Monaco.

Footnotes

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

See Commentary on page 16541.

This article contains supporting information online at www.pnas.org/cgi/content/full/0902894106/DCSupplemental.

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