Increased apical sodium-dependent glucose transporter abundance in the ctenidium of the giant clam Tridacna squamosa upon illumination

Tropical waters are poor in nutrients owing to a lack of overturn, and therefore referred to as ‘deserts’ of the sea. To compensate for nutrient shortage in tropical waters, some marine invertebrates, including giant clams and scleractinian corals, acquire and maintain symbiotic relationships with certain dinoflagellates (zooxanthellae) (Trench, 1987). Giant clams are common members of coral reefs throughout the tropical Indo-Pacific (Neo et al., 2017). They can harbor three genera of dinoflagellates of the family Symbiodiniaceae – Symbiodinium (formerly Symbiodinium clade A), Cladocopium (formerly Symbiodinium clade C) and Durusdinium (formerly Symbiodinium clade D) (Takabayashi et al., 2004; Hernawan, 2008; LaJeunesse et al., 2018) – extracellularly in a branched tubular system. These symbionts live inside the zooxanthellal tubules located mainly in the extensible and colorful outer mantle (Norton et al., 1992), where they conduct photosynthesis during insolation. They transfer >95% of photosynthates to the host, which can generally satisfy the host's energy requirements (Fitt, 1993; Griffiths and Klumpp, 1996). With that, giant clams can perform light-enhanced shell formation (calcification) and maintain a high growth rate in nutrient-deficient tropical waters where light is available (Lucas et al., 1989). As the photosynthesizing zooxanthellae require a supply of inorganic carbon, research in the past has focused on inorganic carbon uptake and metabolism in giant clams (Rees et al., 1993b; Baillie and Yellowlees, 1998; Leggat et al., 2002, 2005; Yellowlees et al., 2008). Nonetheless, giant clams can also obtain some nutrients through filter feeding (Fankboner and Reid, 1986), possible digestion of zooxanthellae in the digestive tract (Reid et al., 1984), and uptake of dissolved organic molecules from the external medium (Fitt, 1993).

More than a century ago, Putter (1909) suggested that marine organisms might be able to obtain nutrients by directly absorbing certain molecules from the ambient seawater. Since then, evidence has been gathered to substantiate the absorption of isotopically labelled sugars and amino acids against their concentration gradients through epidermal tissues of annelids, echinoderms and pogonophores (Stephens and Schinske, 1961; Ferguson, 1967; Ahearn and Gomme, 1975; Davis and Stephens, 1984; Manahan, 1989; Pajor et al., 1989). Péquignat (1973) provided categorical evidence to support the presence of a similar epidermal route of absorption for glucose and amino acids in the ctenidium (gill) and mantle of the filibranch bivalve Mytilus edulis. Subsequently, by measuring d-glucose transport in brush-border membrane vesicles derived from the ctenidium of M. edulis, the functional presence of a sodium-dependent glucose transporter (SGLT) in its labial palps has been confirmed (Pajor et al., 1989). Furthermore, in vitro incubation of the ctenidium of the oyster Crassostrea gigas in artificial seawater reveals that it can absorb d-glucose and d-galactose, but not 3-O-methyl-d-glucose, via an active carrier-mediated system (Bamford and Gingles, 1974). In the absence of exogenous Na⁺, glucose uptake occurs by simple diffusion, but an Na⁺-dependent carrier-mediated process sensitive to phlorizin [a potent inhibitor of sodium/glucose cotransporter 1 (SGLT1); Panayotova-Heiermann et al., 1996] inhibition is operating to absorb glucose in the ctenidium and the mantle of C. gigas. A kinetic analysis of glucose uptake confirms the presence of a saturable component at low substrate concentrations, and a diffusive component at high substrate concentrations. Subsequently, a sequence of SGLT-like has been obtained from the oyster C. gigas (Huvet et al., 2004) that has a high mRNA expression level in the ctenidium and the mantle edge but low expression in other tissues/organs (Hanquet et al., 2011).

SGLTs cotransport Na⁺ and glucose down the electrochemical potential gradient of Na⁺ but up against the concentration gradient of glucose across cell membranes. Six isoforms of the SGLT gene belonging to the SLC5 gene family have been identified in humans (Wright et al., 2011). All SGLTs have 14 transmembrane regions (TMs) in topology (Wright et al., 2011). In humans, SGLT1 is expressed mainly in the intestine and kidney, where it functions as a glucose/galactose transporter. In mouse intestine, SGLT1 is expressed in the brush-border membranes, and glucose absorption across these membranes disappears in SGLT1-deficient mouse, indicating that intestinal glucose absorption is mediated predominantly by SGLT1 (Gorboulev et al., 2012).

Until now, it was unknown whether giant clams express a homolog of SGLT1 (SGLT1-like) in their ctenidia, and whether they can absorb glucose from the ambient seawater. Nonetheless, giant clams are known to display light-dependent physiological properties, including light-enhanced shell formation (Sano et al., 2012; Ip et al., 2017a) and light-enhanced nitrogen uptake (Wilkerson and Trench, 1986; Chan et al., 2018). In addition, the gene and/or protein expression levels of some of their enzymes and transporters are also light dependent (Hiong et al., 2017a,b; Boo et al., 2017, 2018; Ip et al., 2015, 2017a,b, 2018; Koh et al., 2018; Chan et al., 2018; Chew et al., 2019). As such, it is highly probable that giant clams can increase the expression of SGLT1-like/SGLT1-like in the ctenidium and the rate of glucose uptake during light exposure.

Therefore, the present study was undertaken to clone and sequence SGLT1-like from the ctenidium of the fluted giant clam, Tridacna squamosa. Sequence similarity analysis was conducted to verify that the SGLT1-like of T. squamosa was derived from the host. Molecular characterization of the deduced SGLT1-like amino acid sequence was performed to elucidate its roles in the co-transport of Na⁺ and glucose. Furthermore, the gene expression of SGLT1-like in various organs and tissues of T. squamosa was examined. The hypothesis tested was that SGLT1-like was expressed predominantly in the ctenidium, where glucose uptake could occur. Based on the deduced amino acid sequences, a custom-made anti-SGLT1-like polyclonal antibody was produced commercially, and immunofluorescence microscopy was performed to test the proposition that SGLT1-like was localized in the apical membrane of the epithelial cells of the ctenidial filament and water channels, where it could engage in active glucose absorption. In addition, quantitative real-time PCR (qPCR) and western blotting were performed to examine whether the gene and/or protein expression of SGLT1-like/SGLT1-like in the ctenidium of T. squamosa could be upregulated by light. To verify that T. squamosa could indeed perform light-enhanced glucose absorption, the rates of glucose absorption in T. squamosa exposed to darkness (control) or to light were determined. Efforts were also made to elucidate the possible relationship between glucose and urea absorption in T. squamosa, as SGLT1 is known to have the capacity to transport urea (Panayotova-Heiermann and Wright, 2001; Bankir and Yang, 2012). As a crucial part of the coral reef, giant clams are sensitive to adverse changes in their environment, particularly in connection with global warming and ocean acidification. Results obtained from this study may shed light on the mechanisms of glucose (and urea) uptake in giant clams, which may offer insights into ways to enhance their growth and survivorship in this rapidly changing climate.

Adult Tridacna squamosa Lamarck 1819 (mass=500±180 g, N=42) were procured from Xanh Tuoi Tropical Fish (Ho Chi Minh City, Vietnam). The giant clams were maintained in tanks as described by Ip et al. (2015) but with slight modifications. The water temperature was maintained at 26±1°C, the salinity was 30–32 and the pH ranged between 8.1 and 8.3. The carbonate hardness was 143–179 ppm and the calcium concentration was 280–400 ppm. Each tank was illuminated from the top by two sets of Aquazonic T5 lighting systems (Yi Hu Fish Farm Trading, Singapore), and each system consisted of two sun and two actinic blue fluorescence tubes (39 W each). Using a Skye SKP 200 display meter connected with a SKP 215 PAR Quantum sensor (Skye Instruments, UK), the light intensity at the level of the giant clams was determined as ∼100 μmol m⁻² s⁻¹. Institutional approval was not necessary for research on giant clams (National University of Singapore Institutional Animal Care and Use Committee).

At the end of the 12 h dark period of the 12 h:12 h light:dark regime, one batch of giant clam (N=5; control) was killed and sampled. The other 15 individuals were separated into three batches (N=5 for every time point) and exposed to 3, 6 or 12 h of light before being killed for tissue sampling. They were anesthetized in 0.2% phenoxyethanol, and forced open to cut the adductor muscles. Then, samples of outer mantle, inner mantle, ctenidium, adductor muscle, foot muscle, hepatopancreas and kidney were excised. The tissue samples were dabbed dry and then freeze-clamped with aluminum tongs in liquid nitrogen. All samples were stored at −80°C prior to processing. Tissue samples for immunofluorescence microscopy were harvested separately from another batch of individuals that had been exposed to either darkness or light for 12 h (N=4 for each condition) followed with anesthetization in 0.2% phenoxyethanol.

Extraction of the total RNA from the ctenidium of T. squamosa was achieved using TRI Reagent^® (Sigma-Aldrich, St Louis, MO, USA). Further purification of extracted total RNA was accomplished using the RNeasy Plus Mini Kit (Qiagen, Hilden, Germany), and the concentration of total RNA was verified using a Shimadzu BioSpec-nano spectrophotometer (Shimadzu, Tokyo, Japan). Electrophoresis was used to verify total RNA integrity. The RevertAid™ first-strand cDNA synthesis kit (Thermo Fisher Scientific, Waltham, MA, USA) was used to reverse transcribe the purified total RNA into cDNA.

The partial SGLT1-like sequence was isolated using primers (given 5′ to 3′; forward: GGWTGGGTGTTTGTMCCTGT; reverse: GGWTGCACSAAYATYGCMTATC; where W, M, S and Y represent degenerate bases) designed from the homologous regions of Crassostrea gigas sodium glucose cotransporter (AY551098.1), Octopus bimaculoides sodium/glucose cotransporter 4-like (XM_014916561.1), Homo sapiens solute carrier family 5 member 1 (NM_000343.3; SGLT1) and Acropora digitifera sodium/glucose cotransporter 1-like (XM_015901071.1). The PCR reaction was conducted using DreamTaq™ polymerase (Thermo Fisher Scientific) in a 9902 Veriti 96-well thermal cycler (Applied Biosystems, Carlsbad, CA, USA). Following the methods previously described (Hiong et al., 2017a,b) with minor modifications, PCR and cloning experiments were used to isolate and analyze the gene. For the PCR experiments, initial denaturation was at 95°C held for 3 min, followed by 40 cycles of denaturation, annealing and extension at 95°C for 30 s, 55°C for 30 s and 72°C for 1.5 min. A final extension for 10 min was held at 72°C. An absence of isoforms was observed from the analysis of multiple clones of SGLT1-like fragments. 5′ and 3′ RACE (SMARTer™ RACE cDNA amplification kit, Clontech Laboratories, Mountain View, CA, USA) PCR was performed with specific primers (forward: GCCAGGTACAGGTTCAAGTGTACAGACT; reverse: ATCCAATGCCAAAGCGGGTACAATTCTG) to amplify and isolate the full coding sequence of SGLT1-like. The BigDye Terminator v3.1 Cycle Sequencing Kit (Thermo Fisher Scientific) with ethanol/sodium acetate precipitation was used to prepare the samples for gene sequencing. Sequencing was performed using a 3130XL Genetic Analyzer (Thermo Fisher Scientific). The sequence was subsequently analyzed and assembled using BioEdit version 7.2.5. The SGLT1-like sequence has been deposited into GenBank with accession number MF073182.

The SGLT1-like amino acid sequence was translated from the SGLT1-like nucleotide sequence using the ExPASy Proteomic server (http://web.expasy.org/translate/). To confirm the identity of SGLT1-like from T. squamosa, the deduced amino acid sequence was aligned with selected SGLT and SGLT1-like sequences from various species of mollusk and other animals to generate a sequence similarity table (see Table 1 for details). The TMs of the deduced amino acid sequence were predicted using the TMpred server of the ExPASy portal (https://embnet.vital-it.ch/software/TMPRED_form.html).

The mRNA expression of SGLT1-like in the outer mantle, inner mantle, ctenidium, adductor muscle, foot muscle, kidney and hepatopancreas of T. squamosa was investigated qualitatively by PCR using SGLT1-like specific primers (forward: AAATCTATCCAATGCCAAAGCGG; reverse: TTCTATCCCACAAATCCACTGACC). DreamTaq™ polymerase (Thermo Fisher Scientific) was used for each PCR reaction, with each reaction having a total volume of 10 µl. For the thermal cycling, initial denaturation was accomplished through holding 95°C for 3 min, followed by 35 cycles of 95°C (30 s) for denaturation, 55°C (30 s) for annealing and 72°C (1 min) for extension, and one final extension at 72°C (10 min). Electrophoresis in 1% agarose gel was used to separate the PCR products.

Absolute quantification through qPCR was performed using a StepOnePlus™ Real-Time PCR System (Thermo Fisher Scientific). cDNA (4 µg) was synthesized from RNA using random hexamer primers and the RevertAid™ first-strand cDNA synthesis kit. The mRNA expression level of SGLT1-like in the ctenidium was determined through the use of specific qPCR primers (forward: TCTTACACAGCCGATTGACGA; reverse: ATCCGAACACTGAGATGTCCT). The amplification efficiency for SGLT1-like was 86.6%. The absolute quantification of transcripts was calculated using the plasmid standard curve expressed as copy number per nanogram total RNA, with methodology following previous publications (Hiong et al., 2017a,b).

A custom-made rabbit polyclonal anti-SGLT1-like antibody was developed by a commercial firm (GenScript, Piscataway, NJ, USA) against residues 243–256 (PPDNSMNLIRSYDD) of SGLT1-like of T. squamosa, and used for immunofluorescence microscopy and immunoblotting. α-Tubulin was chosen as the reference protein for western blotting, and the anti-α-tubulin antibody (12G10) was procured from the Developmental Studies Hybridoma Bank (Department of Biological Sciences, University of Iowa, Iowa City, IA, USA)

The immuno-labeling of ctenidial samples of T. squamosa with the anti-SGLT1-like antibody (1.35 µg ml⁻¹) was performed following the methods of Hiong et al. (2017b) with Alexa Fluor 488, and goat anti-rabbit (Invitrogen, 2.5 µg ml⁻¹) secondary labeling in green. Nuclei were counterstained using 4′6′-diamidino-2-phenylindole (DAPI) (Sigma-Aldrich; 50 ng ml⁻¹). Slides were mounted in ProLong Gold Antifade Mountant (Life Technologies, USA) and cured in the dark at room temperature prior to storing at 4°C for further image acquisition. Image acquisition was conducted with a fluorescence microscope (Olympus BX60 equipped with a DP73 CCD digital camera) with the appropriate fluorescent filter sets (Ip et al., 2018). A DIC slider (U-DICT, Olympus) was used to produce differential interference contrast (DIC) images of the tissue structures such as ctenidial filaments and water channels. Micrographs were collected using CellSens Imaging Software (Olympus) under optimal exposure settings of 200–400 ms. Images were further processed with Adobe Photoshop CC (Adobe Systems, San Jose, CA, USA) with adjusted brightness and contrast.

The extraction of protein from ctenidial samples was performed following the methods of Hiong et al. (2017b). Twenty micrograms of ctenidial proteins was subjected to 8% SDS-PAGE and then blotted onto polyvinylidene difluoride membrane. The membranes were blocked with Pierce Fast blocking buffer for 30 min at 25°C, followed by incubation with anti-SGLT1-like (1.25 μg ml⁻¹) or anti-α-tubulin (12G10, 0.1 μg ml⁻¹) antibodies for 1 h at 25°C. Subsequently, the membranes were incubated in horseradish peroxidase-conjugated secondary antibodies (Santa Cruz Biotechnology; 1:10,000) for 1 h at 25°C. Blots were washed with TBST (0.05% Tween 20 in Tris-buffered saline: 20 mmol l⁻¹ Tris HCl; 500 mmol l⁻¹ NaCl, pH 7.6) three times and developed with the ECL system (Thermo Fisher Scientific). Scanning and quantification of the protein bands were performed as described by Ip et al. (2017a,b). The quantitation of the SGLT1-like protein abundance was normalized with that of α-tubulin. Prior to immunoblotting, immunizing peptide (Genscript) was incubated with anti-SGLT1-like antibody for 1 h to further determine the specificity of the custom-made antibody.

Experiments were designed to demonstrate glucose absorption in T. squamosa by monitoring the reduction in glucose concentration in the external medium, which represented the sum of glucose influx and efflux. It was not feasible to determine increases in glucose concentration in the hemolymph related to glucose absorption, as the hemolymph has high and fluctuating concentrations of glucose (500–600 mmol l⁻¹) owing to the donation of photosynthates, including glucose, from zooxanthellae (Deane and O'Brien, 1980; Rees et al., 1993a). Uptake of radiolabeled glucose was not adopted in the present study, because it would reflect at best the unidirectional influx of exogenous glucose without information on the possible glucose efflux.

Sixteen individuals of T. squamosa were randomly selected at the end of a 12 h:12 h light:dark regime, and transferred separately to clear plastic tanks (21.5×11.5×12.5 cm, length×width×height) in complete darkness. Each plastic tank contained eight volumes (8×mass of clam) of aerated artificial seawater, and the individual clam was allowed to acclimatize therein for 2 h in darkness. Then, concentrated glucose solution was added to the artificial seawater to make a final concentration of 50 µmol l⁻¹. Although natural seawater has relatively low glucose concentrations (∼0.23 μmol l⁻¹; Mopper et al., 1980), preliminary results indicated that 50 µmol l⁻¹ of exogenous glucose was needed for the giant clam to sustain a relatively linear uptake of glucose for 6 h. Approximately 2 min was required for the glucose concentration to become homogeneous; hence, 2 min after the addition of glucose was taken as time 0.

To examine the effects of light on glucose uptake by T. squamosa, five giant clams were exposed to light at 100–105 µmol m⁻² s⁻¹, and another five giant clams were kept as controls in darkness. In order to examine whether urea would interfere with glucose absorption, three individuals were exposed to light for 6 h in seawater containing 50 µmol l⁻¹ glucose, while another three individuals were exposed to light for 6 h in seawater containing 50 µmol l⁻¹ glucose+50 µmol l⁻¹ urea. The experiment lasted only 6 h because preliminary results obtained indicated that the glucose concentration would decrease by ∼50% over a 6-h period. Waters containing glucose or glucose+urea, but without giant clams, were regarded as blanks. Water samples were collected at 0, 2, 4 or 6 h, and kept at −20°C. The glucose assay, following the methods of Bergmeyer et al. (1974), was performed within 7 days of the uptake experiment. Briefly, the water samples were pre-incubated in 1.5 ml of reaction mixture containing 250 mmol l⁻¹ triethanolamine (pH 7.5), 2.5 mmol l⁻¹ MgSO₄, 0.8 mmol l⁻¹ NADP, 10 mmol l⁻¹ ATP and 2.1 i.u. glucose-6-phosphate dehydrogenase. After incubation at 25°C for 5 min, the absorbance was determined at 340 nm using a Shimadzu UV160 UV-VIS spectrophotometer and the reaction was initiated by the addition of 2.8 i.u. hexokinase. The absorbance was recorded after 10 min and the change in absorbance was used for calculation. Freshly prepared glucose solution was used as a standard for comparison. Percentage changes were used to express the decreases in the glucose concentration in the external medium, because of minor variations in the initial glucose concentrations at hour 0. The rate of glucose absorption was expressed as µmol g^–1 h^–1.

Values are given as means±s.e.m. Student's t-test for independent samples was applied to compare the differences between two means. As for multiple means, the homogeneity of variance among the means was analyzed using Levene's test. Subsequently, one-way ANOVA was used to evaluate the difference between the means of data sets, followed by Dunnett’s T3 post hoc test or Tukey’s post hoc test, where appropriate. Statistical analysis of data was performed using SPSS Statistics version 19 (IBM Corporation, Armonk, NY, USA) with the significance level set at P<0.05.

The full cDNA coding sequence of SGLT1-like consisting of 1946 bp was obtained from the ctenidium of T. squamosa (GenBank accession no.: MF073182). The deduced SGLT1-like of T. squamosa comprised 649 amino acids with an estimated molecular mass of 72.4 kDa. It had the highest sequence similarity with the SGLTs of molluscs (84.3–86.0%; Table 1), followed by those of brachiopods, echinoderms, hemichordates and chordates (70.5–80.0%), but it had low sequence similarity with the SGLTs of plants (30.7–34.6%). A comparison with the SGLT sequence of the green alga Auxenochlorella protothecoides indicated only 34.6% similarity (Table 1). Hence, it can be confirmed that the SGLT1-like obtained from T. squamosa was derived from the host clam and not the zooxanthellae.

An alignment with SGLT1 sequences from other organisms (Crassostrea gigas, Homo sapiens and Vibrio parahaemolyticus) obtained from GenBank revealed that SGLT1-like of T. squamosa contained five glucose-binding residues (D12, E85, D187, K306 and Q440) (Fig. 1). Moreover, it also contained five residues (N61, Y275, W276, S377 and S378) known to be involved in the binding of Na⁺ and sugar in human SGLT1-like. Particularly, the conserved residues F436 and Q440 are known to contribute to urea permeability (Fig. 1). The backbone carbonyls of the conserved residues A59 and I62 also contributed to the formation of the sugar-binding site, whereas residues W274, Y275 and W276 (indicated in the open black box in Fig. 1) formed an aromatic triad contributing to the transport of sugar molecules. There were 14 predicted TMs in the SGLT1-like of T. squamosa. A domain analysis using PROSITE (Sigrist et al., 2002) denoted that SGLT1-like of T. squamosa displayed the profile of Sodium/Solute Symporter 3 (SSS3) family transporters.

The SGLT1-like gene was strongly expressed in the ctenidium but weakly expressed in the outer mantle, foot muscle and kidney of T. squamosa kept in darkness for 12 h (Fig. 2).

Immunofluorescence microscopy revealed that SGLT1-like was localized in the apical membrane of epithelial cells of the filaments (Fig. 3) and some epithelial cells of the water channels (Fig. 4) in the ctenidium of T. squamosa. An apparently stronger apical anti-SGLT1-like immunofluorescence was observed in the ctenidial filaments of giant clams exposed to 12 h of light than that of the control kept in 12 h of darkness, with reproducible results being obtained from four individual clams for each condition (Figs 3 and 4).

The SGLT1-like transcript level in the ctenidium of T. squamosa exposed to light for 3, 6 or 12 h of light remained unchanged as compared with the control kept in darkness for 12 h (Fig. 5). By contrast, there was a progressive, albeit insignificant, increase in the protein abundance of SGLT1-like in the ctenidium of clams exposed to light for 3 or 6 h, and by the 12th hour of light exposure, the protein abundance of SGLT1-like increased significantly (by ∼10-fold) as compared with the control kept in darkness (Fig. 6).

Without giant clams, the glucose concentration in seawater remained unchanged during the 6-h period. However, the glucose concentration in the seawater containing T. squamosa decreased almost linearly during 6 h of exposure to darkness (control) or light (N=5; Fig. 7A). The change in ambient glucose concentration was consistently greater in light than in darkness, with significant differences between them at hours 4 and 6. At hour 4, the calculated rate of glucose absorption in individuals exposed to light was significantly higher than that in individuals exposed to darkness (N=5; Fig. 7B).

When T. squamosa were exposed to glucose plus urea (both at 50 µmol l⁻¹), the decrease in ambient glucose concentration during exposure to light for 6 h was constantly and significantly lower than that of giant clams exposed to 50 µmol l⁻¹ glucose only (N=3; Fig. 8A). At hour 6, the calculated rate of glucose absorption in giant clams exposed to glucose plus urea was significantly lower than that exposed to glucose without urea (N=3; Fig. 8B).

There are two families of glucose transporters: SGLTs and facilitative glucose transporters (GLUTs). GLUTs transport glucose via facilitated diffusion, but SGLTs are secondary active glucose transporters that tap into the electrochemical gradient of Na⁺ generated by Na⁺/K⁺-ATPase across the plasma membrane. SGLT1 was first cloned by Hediger et al. (1987) from rabbit intestine, and soon after that the human SGLT1 analogue was discovered (Hediger et al., 1989). To date, 11 SGLT genes have been identified in the human genome and at least six of them are known to be expressed as integral proteins in the plasma membrane (Wright and Turk, 2004). Homologs of some of these SGLTs have been identified in invertebrates, including shrimp, lobsters, horseshoe crabs, insects, snails, mussels and oysters (see Martínez-Quintana and Yepiz-Plascencia, 2012 for a review). In the present study, we have successfully cloned the complete coding sequence of SGLT1-like from the ctenidium of T. squamosa.

SGLT1-like of T. squamosa comprised 14 predicted TMs, corresponding to the 14 TMs characterized in SGLT1 of H. sapiens (Turk et al., 1996). SGLT1 is known to have an extracellular N-terminus (Turk et al., 1996), which contains a highly conserved aspartate residue that is involved in sugar translocation (Turk et al., 2000). This aspartate residue (D12) is conserved in SGLT1-like of T. squamosa (Fig. 1). The hydrophobic C-terminus of SGLT1 is involved in the formation of a transmembrane helix (Turk et al., 1996), which is also reflected in SGLT1-like of T. squamosa (TM14; Fig. 1). Furthermore, SGLT1-like of T. squamosa contains the five conserved residues (D12, E85, D187, K306 and Q440) involved in binding of sugar molecules (Loo et al., 2013). It also comprises residues (N61, Y275, W276, S377 and S378) that bind to Na⁺ and sugar (Loo et al., 2013), as well as residues W274, Y275 and W276 (indicated by an open black box; Fig. 1) that form an aromatic triad crucial for both sugar and Na⁺ binding (Loo et al., 2013).

As SGLT1 can also transport urea (Leung et al., 2000), which may follow the path of sugar transport (Zeuthen et al., 2016), the sugar-binding residues in SGLT1-like of T. squamosa may also bind with urea (Panayotova-Heiermann and Wright, 2001; Wright et al., 2011). Moreover, residues F436 and Q440, which are conserved in SGLT1-like of T. squamosa, have been shown to contribute to urea permeability (Zeuthen et al., 2016). Hence, SGLT1-like of T. squamosa can probably transport not only glucose, but also urea, with Na⁺ being used as a motive force to drive their active uptake.

Many mollusks, including bivalves, cephalopods and aquatic gastropods, possess a pair of ctenidia inside their mantle cavities. A ctenidium is primarily a respiratory organ in water, but can also participate in filter feeding, ionoregulation and acid–base balance. In T. squamosa, the ctenidia are whitish in color, and each ctenidium consists of two demibranches (dorsal and ventral). The ctenidium is comb-shaped, and has a central part from which many filaments protrude and line up in a row to increase the surface area for various physiological functions (Norton and Jones, 1992). The surface area is further increased by numerous water channels found below the filaments inside the ctenidium.

The ctenidium of T. squamosa is known to express transporters and enzymes (Ip et al., 2015) related to nitrogen transport and assimilation (DUR3-like, Chan et al., 2018; Ammonia Transporter 1, Boo et al., 2018; Glutamine Synthetase, Hiong et al., 2017a), inorganic carbon absorption (Dual Domain Carbonic Anhydrase; Koh et al., 2018) and proton excretion (Na⁺/H⁺ Exchanger 3-like, Hiong et al., 2017b; Vacuolar-type H⁺-ATPase subunit A, Ip et al., 2018). In the present study, we demonstrated that SGLT1-like was expressed predominantly in the ctenidium of T. squamosa. Importantly, SGLT1-like had an apical localization in epithelial cells covering the ctenidial filament and lining the tertiary water channels. These results denote that SGLT1-like is positioned to transport Na⁺ together with glucose (or urea) from the ambient seawater into the ctenidial epithelial cells, and suggest the ctenidium as the site of active glucose uptake.

It has been reported that the protein expression level of SGLT1 in the intestinal brush border of mammals can be upregulated by glucose (Wood et al., 2000), and protein kinases (PKA and PKC) regulate the transport of SGLT1 by rapidly inserting it into the plasma membrane (Turk and Wright, 1997). Similarly, SGLT1-like of oyster (C. gigas) is regulated according to trophic conditions, including food abundance and quality (Hanquet et al., 2011). Nevertheless, this is the first report on the upregulation of the protein abundance of SGLT1-like in the ctenidium of T. squamosa in response to light. Our results indicate that SGLT1-like was regulated principally through translation, as light exposure did not have a significant effect on its transcript level. This differs from Dual Domain Carbonic Anhydrase (Koh et al., 2018), Glutamine Synthetase (Hiong et al., 2017a) and Na⁺/H⁺ Exchanger 3-like (Hiong et al., 2017b), of which both the transcript level and the protein abundance increase significantly in the ctenidium of T. squamosa during 12 h of light exposure. Overall, these results corroborate the proposition that T. squamosa could perform light-enhanced glucose/urea absorption.

The hemolymph of giant clams is known to have very high concentrations of glucose (500–600 mmol l⁻¹; Deane and O'Brien, 1980; Rees et al., 1993a) owing to the donation of photosynthates from zooxanthellae. Yet, T. squamosa could reduce the ambient glucose concentration (50 mmol l⁻¹) by 45% during 6 h of light exposure despite such a huge concentration gradient. Hence, it can be concluded that T. squamosa could actively absorb glucose form the environment. Notably, the observation on the rate of glucose uptake being higher in light than in darkness is novel. Although glucose uptake is known to occur in non-symbiotic mussels (Péquignat, 1973) and oysters (Bamford and Gingles, 1974), there is no indication of it being a light-dependent process. Scleractinian corals (e.g. Fungia) can also absorb glucose from the external medium (Stephens, 1960, 1962), but the possibility of it being enhanced by light has not been explored.

Natural seawater contains dissolved organic molecules that comprise a wide variety of carbohydrates, lipids, amino acids, as well as vitamins and hormones (Duursma, 1965). Specifically, glucose is the major monosaccharide in seawater (Ittekkot et al., 1981), derived largely from oligosaccharide hydrolysis. Phytoplankton produces and stores glucan oligosaccharides (Lewin, 1974), which can be released to the ambient seawater through exudation or cell lysis. The degradation of these glucan oligosaccharides in seawater produces monosaccharides including glucose. Generally, the concentration of glucose in open seawater ranges between 10⁻⁶ and 10⁻⁸ mol l⁻¹ (Vaccaro et al., 1968). Although the glucose concentration can be higher in waters around coral reefs owing to both algal and coral exudations that contain saccharides, proteins and lipids (Haas and Wild, 2010; Nelson et al., 2013), it is unlikely to reach the concentration of 0.05 mmol l⁻¹ adopted in this study. Therefore, it is possible that the rate of glucose uptake obtained for T. squamosa is non-physiological, although its ability to absorb glucose is affirmed. However, in effect, the absorption rate of glucose should be defined by the concentrations of glucose in the unstirred mucus layer covering the ctenidial epithelium, and not the glucose concentrations in the ambient seawater.

In scleractinian corals, the oral ectoderm, consisting of multiple specialized cells such as cnidocytes, mucocytes and epitheliomuscular cells (Fautin and Mariscal, 1991), is involved in the transport of Ca²⁺ into calicoblastic cells associated with calcification (Tambutté et al., 2007). During insolation, the Ca²⁺ concentration (14.5±0.7 mmol l⁻¹) in the 10–20 μm thick mucus layer coating the oral ectoderm is consistently higher than that (11.3±0.3 mmol l⁻¹) in the ambient seawater (Clode and Marshall, 2002). Hence, the existence of a mucus boundary between the seawater and oral ectoderm could facilitate the efficient absorption of Ca²⁺ into the oral ectodermal cells. Similarly, type 2 and type 3 mucin gland cells have been identified in the ctenidium of T. squamosa, and these cells can secrete mucus onto surfaces of the epithelial cells lining the filaments and water channels (Norton and Jones, 1992). If the microbiome of the ctenidial surface can perform oligosaccharide degradation, the glucose concentration in the mucus layer could be higher than that in the seawater, but the confirmation of this proposition awaits future study.

The assimilation of dissolved organic matter by sponge holobionts is known to facilitate the cycling of dissolved organic matters in benthic habitats (de Goeij et al., 2013). Microbes contribute to this assimilation process (Rix et al., 2016), accounting for ∼90% of the holobiont's total heterotrophic carbon uptake (Morganti et al., 2017; Hoer et al., 2018). In corals, the associated microbial communities are also important components of the coral holobionts (Leggat et al., 2007; Rosenberg et al., 2007), as they participate in nutrient cycling and influence coral health and diseases (Rosenberg and Loya, 2004; Rosenberg et al., 2007). Kimes et al. (2010) analyzed the microbiome associated with the scleractinian coral Montastraea faveolata and detected 134 ribulose-1,5-bisphosphate carboxylase/oxygenase genes (RuBisCO) of archaea and bacteria origins, indicating that these microbes could fix CO₂ and produce organic compounds through photosynthesis (Berg et al., 2002). Furthermore, Kimes et al. (2010) detected 825 microbial genes involved in polysaccharide degradation, including bacterial, archaeal and fungal cellulases (359 sequences), chitinases (206 sequences), mannanases (55 sequences) and polygalactases (63 sequences). Degradation of polysaccharide by these microbes could release glucose and other monosaccharides close to the surface of the host. In fact, the ability of microbes to metabolize nutrients, which can then be translocated to their host, is likely a driver in the establishment of coral-associated microbial assemblages. Assuming that the surface microbiome of giant clams can do the same, the concentration of glucose in the mucus layer covering the ctenidial epithelium could be higher than that in the ambient seawater, rendering an effective uptake of glucose though SGLT1-like.

However, the contribution of light-enhanced glucose absorption through ctenidial SGLT1-like to the overall metabolic needs of the host clam could be inconsequential when compared with the donation of carbohydrates, including glucose, from the photosynthesizing zooxanthellae (Rees et al., 1993a; Ishikura et al., 1999). In fact, it has been established that the photosynthates transferred from the symbionts to the host clam can satisfy ∼100% of the host's energy requirements (Fisher et al., 1985; Klumpp et al., 1992). Therefore, the physiological significance of active glucose absorption in symbiotic T. squamosa is obscure. Nonetheless, there could be three physiological reasons, related separately to the metabolic needs of the aposymbiotic larva, the ctenidium and the zooxanthellae.

Giant clam larvae are aposymbiotic, and they acquire symbiotic zooxanthellae only during the veliger stage through filter feeding (Fitt and Trench, 1981; Heslinga et al., 1984; Mies and Sumida, 2012). It has been reported that veliger larvae of oyster (C. gigas) and red abalone (Haliotis rufescens) absorb glucose and some other complex sugars from the external medium, which allows them to utilize a greater part of the dissolved organic material in the sea as a source of nutrition (Welborn and Manahan, 1990). Therefore, it is probable that the absorption of exogenous glucose through SGLT1-like in the rudimentary ctenidium contributes to the successful survival of giant clam larvae, which needs to be confirmed in future studies. Nevertheless, this could lead to the retention of SGLT1-like expression in the ctenidium of symbiotic T. squamosa.

The ctenidium of T. squamosa is metabolically active owing to its important roles in, for example, inorganic carbon absorption (Koh et al., 2018), excretion of excess H⁺ (Hiong et al., 2017b; Ip et al., 2018), and nitrogen transport and assimilation (Chan et al., 2018; Boo et al., 2018; Hiong et al., 2017a). It contains few zooxanthellae, and is located inside the mantle cavity with no direct exposure to sunlight. Although the ctenidium can undoubtedly receive some photosynthates from the outer mantle through the hemolymph, these two organs are separated far apart. Importantly, other tissues such as the inner mantle, which adjoins the outer mantle and conducts light-enhanced shell formation, would also compete for nutrients, especially during insolation. Therefore, the expression of SGLT1-like in the ctenidium could be advantageous to symbiotic T. squamosa, as the absorbed glucose can specifically fuel the metabolic needs of this organ. It is possible that the problem concerning low ambient glucose concentrations could be remedied by the activities of its surface microbiome.

It has been established that SGLT1 can transport urea in addition to glucose (Leung et al., 2000; Panayotova-Heiermann and Wright, 2001), with the coupling of urea transport to its cotransport cycle (Zeuthen et al., 2001). This could explain why urea interfered with light-enhanced glucose absorption in T. squamosa, and the rate of glucose uptake in individuals exposed to glucose+urea was lower than that in individuals exposed to glucose only. Although the host clam benefits from photosynthates donated by symbiotic zooxanthellae (Streamer et al., 1993), the symbionts require a supply of nutrients from the host (Furla et al., 2005). Specifically, the host clam would need to absorb exogenous nitrogen and supply it to the symbiotic zooxanthellae, which are nitrogen deficient. Dissolved inorganic nitrogen is present in seawater as ammonium, nitrite and nitrate, whereas dissolved organic nitrogen is represented mainly by urea and amino acids. In reef environments, urea concentrations range between 0.2 and 2.0 µmol N l⁻¹ (Wafar et al., 1986; Beauregard, 2004), but the urea concentration in the mucus covering the ctenidial epithelium of T. squamosa could be higher, attributable to the activities of its microbiome. Indeed, nitrogen cycling is often cited as a probable role filled by the microbial community of coral reefs (Rosenberg et al., 2007), and bacteria are known to produce and excrete urea (Pedersen et al., 1993).

Recently, it has been demonstrated that T. squamosa can perform light-enhanced urea absorption, and its ctenidium expresses a urea-active (energy-dependent) transporter, DUR3-like, of animal origin (Chan et al., 2018). Urea can be a good nitrogen source for the symbiotic zooxanthellae because it contains two nitrogen atoms and one carbon atom (H₂NCONH₂), and zooxanthellae possess urease, which can catabolize urea. The degradation of urea by urease in zooxanthellae releases NH₃ to support amino acid metabolism and CO₂ to sustain photosynthesis. Therefore, it would be beneficial for T. squamosa to possess multiple mechanisms in the ctenidium, including DUR3-like and SGLT1-like, to absorb urea from the ambient seawater. Notably, the competition between glucose and urea for transport by SGLT1-like would be defined by the transporter's kinetic properties towards these two substrates, as well as the effective concentrations of these two substrates in the vicinity of the transporter.

Tridacna squamosa expresses SGLT1-like in its ctenidia, and the protein abundance of SGLT1-like is upregulated by light exposure. It also increases the absorption of glucose from the ambient seawater in response to light. When taken together with information in the literature, it can be concluded that the ctenidium of T. squamosa is not simply a respiratory organ, as it also participates in various absorptive and excretory processes. Many of these processes are light dependent and they involve enzymes and transporters that can respond to light through transcriptional and/or translational changes. Such phenomena probably stem from the symbiotic association of the clam host with phototrophic zooxanthellae, as the light-dependent properties of these enzymes and transporters would allow the host to react to light in synchrony with the photosynthetic activity of its symbionts.