Branchial ammonia excretion in the Asian weatherloach Misgurnus anguillicaudatus

Comparative Biochemistry and Physiology, Part C 151 (2010) 40–50

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Comparative Biochemistry and Physiology, Part C j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / c b p c

Branchial ammonia excretion in the Asian weatherloach Misgurnus anguillicaudatus J. Moreira-Silva a,b, T.K.N. Tsui c, J. Coimbra a,b, M.M. Vijayan d, Y.K. Ip e, J.M. Wilson a,⁎ a

Laboratorio de Ecofisiologia, Centro Interdisciplinar de Investigação Marinha e Ambiental (CIIMAR), Rua dos Bragas 289, 4050-123, Porto, Portugal ICBAS–Instituto de Ciências Biomédicas de Abel Salazar da Universidade do Porto, Portugal c Department of Biology, McMaster University, Hamilton, Canada d Department of Biology, University of Waterloo, Waterloo, Canada e Department of Biological Sciences, National University of Singapore, Singapore b

a r t i c l e

i n f o

Article history: Received 2 June 2009 Received in revised form 29 July 2009 Accepted 11 August 2009 Available online 20 August 2009 Keywords: Ammonia Boundary layer acidification Membrane fluidity Na+/K+-ATPase Rhcg Slc42a3 H+-ATPase Weatherloach

a b s t r a c t The weatherloach, Misgurnus anguillicaudatus, is a freshwater, facultative air-breathing fish that lives in streams and rice paddy fields, where it may experience drought and/or high environmental ammonia (HEA) conditions. The aim of this study was to determine what roles branchial Na+/K+-ATPase, H+-ATPase, and Rhcg have in ammonia tolerance and how the weatherloach copes with ammonia loading conditions. The loach's high ammonia tolerance was confirmed as was evident from its high 96 h LC50 value and high tissue tolerance to ammonia. The weatherloach does not appear to make use of Na+/NH+ 4 -ATPase facilitated transport to excrete ammonia when exposed to HEA or to high environmental pH since no changes in activity were observed. Using immunofluorescence microscopy, distinct populations of vacuolar (V)-type H+-ATPase and Na+/K+-ATPase immunoreactive cells were identified in branchial epithelia, with apical and basolateral staining patterns, respectively. Rhesus C glycoprotein (Rhcg1), an ammonia transport protein, immunoreactivity was also found in a similar pattern as H+-ATPase. Rhcg1 (Slc42a3) mRNA expression also increased significantly during aerial exposure, although not significantly under ammonia loading conditions. The colocalization of H+-ATPase and Rhcg1 to the similar non-Na+/K+-ATPase immunoreactive cell type would support a role for H+-ATPase in ammonia excretion via Rhcg by NH+ 4 trapping. The importance of gill boundary layer acidification in net ammonia excretion was confirmed in this fish; however, it was not associated with an increase in H+-ATPase expression, since tissue activity and protein levels did not increase with high environmental pH and/or HEA. However the V-ATPase inhibitor, bafilomycin, did decrease net ammonia flux whereas other ion transport inhibitors (amiloride, SITS) had no effect. H+-ATPase inhibition also resulted in a consequent elevation in plasma ammonia levels and a decrease in the net acid flux. In gill, aerial exposure was also associated with a significant increase in membrane fluidity (or increase in permeability) which would presumably enhance NH3 permeation through the plasma membrane. Taken together, these results indicate the gill of the weatherloach is responsive to aerial conditions that would aid ammonia excretion. © 2009 Elsevier Inc. All rights reserved.

1. Introduction The weatherloach (Misgurnus anguillicaudatus) is a freshwater, facultative air-breathing fish (McMahon and Burggren, 1987) that inhabits streams and rice paddy fields, and is native to Siberia, Sakhalin, Korea, Japan, South China to Northern Vietnam (Kottelat and Freyhof, 2007). Like most other teleosts, their primary nitrogenous waste product is ammonia, which classifies them as being ammonotelic (Chew et al., 2001). This fish is subjected to drought conditions during summer, surviving weeks out of water, and also high environmental ammonia (HEA) during periods of agricultural fertilization (Ip et al., 2001). Both of these conditions will compromise ammonia excretion by reversing diffusion gradients for passive efflux (Wilkie, 2002; Ip et al., 2001).

⁎ Corresponding author. Tel.: +351 22 340 1809; fax: +351 22 339 0608. E-mail address: [email protected] (J.M. Wilson). 1532-0456/$ – see front matter © 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.cbpc.2009.08.006

The loach has a high tissue tolerance to ammonia (Chew et al., 2001) and is able to volatilize ammonia (Tsui et al., 2002). It was also shown that glutamine and urea are not used as detoxifying mechanisms (Chew et al., 2001; Tsui et al., 2002), but it is not known what role the gill may have in the remarkable ammonia tolerance of this species. The branchial Na+/K+-ATPase is important in ionoregulation providing the driving force for secondary active Cl− excretion in seawater fishes and Na+ uptake in freshwater fishes (Evans et al., 2005). It has also been implicated in ammonia excretion since similarities in the hydration radius of K+ and NH+ 4 allow substitution at transport sites. In the giant mudskipper (Periophthalmodon schlosseri), Na+/K+-ATPase has a role in active ammonia excretion (Randall et al., 1999). In the silver perch (Bidyanus bidyanus), Na+/K+-ATPase activity increased with exposure to ammonia indicating a role in ammonia excretion as well (Alam and Frankel, 2006). However, in the rainbow trout (Oncorhynchus mykiss) branchial Na+/K+-ATPase activity and mRNA expression were not modified by ammonia exposure (Salama et al., 1999; Nawata et al.,

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2007) and in the juvenile European eel (Anguilla anguilla), there was actually a negative correlation between Na+/K+-ATPase activity and environmental ammonia levels (Moreira-Silva et al., 2009). Thus the role of Na+/K+-ATPase in fish ammonia tolerance is far from clear. The branchial vacuolar type proton ATPase (H+-ATPase) is generally preferentially expressed in the gills of freshwater versus seawater fishes and it has been linked to the uptake of Na+ and Cl− as well as acid–base regulation (Evans et al., 2005). There is also evidence for a role of the H+-ATPase in ammonia excretion (Nawata et al., 2007; Shih et al., 2008). The interrelationship of these latter two effects (acid–base regulation and ammonia excretion) likely has to do with a role for the H+-ATPase in boundary layer acidification. The Rhesus glycoproteins (Rhag, Rhbg, Rhcg) belong to the ammonia transporter/methylammonium permease/Rhesus glycoprotein (Amt/Mep/Rh) superfamily and are thought to transport ammonia (Marini et al., 1997). They were first detected in fish by Kitano and Saitou (2000) and have since been shown to be expressed in a number of teleosts (Takifugu rubripes — gill: Nakada et al., 2007a; Danio rerio — yolk sac, gill, kidney: Nakada et al., 2007b, — skin: Shih et al., 2008; Kryptolebias marmoratus — brain, eye, gill, gonad, gut, kidney, liver, skeletal muscle and skin: Hung et al., 2007; O. mykiss — blood, heart, brain, eye, gill, gonad, gut, kidney, liver, skeletal muscle and skin: Nawata et al., 2007 and gill cell culture: Tsui et al., 2009). Human RhBG and RhCG are expressed in diverse tissues, while RhAG is limited to red blood cells (Huang and Liu, 2001; Liu et al., 2000). Morpholino gene knockdown studies with zebrafish have shown that the Rhcg1 has a role in ammonia excretion (Shih et al., 2008). The mechanism of ammonia transport by Rh glycoproteins is still unclear (see Planelles, 2007), while some authors suggest an electrogenic NH+ 4 + movement (Nakhoul et al., 2006), or an electroneutral NH+ 4 /H mediated exchange (Ludewig, 2004), and others a direct NH3 transport associated with NH+ 4 transport (Bakouh et al., 2004). Independently of the mechanism used, it is agreed that Rh glycoproteins mediate ammonia transport. In aqueous solution ammonia exists as either unionized ammonia (NH3) or ammonium ion (NH+ 4 ). The equilibrium reaction can be + written as: NH+ 4 ↔ NH3 + H . Ammonia speciation will vary markedly with pH (USEPA, 1999; Ip et al., 2001). The separate fractions of each species (fNH3 and fNH4+) can be calculated by the following expressions: fNH3 = 1 / (1 + 10pK–pH), fNH4+ = 1 / (1+ 10pH–pK), and the sum of these two fractions is equal to 1, fNH3 +fNH4+ = 1. The dissociation constant definition is pK = 0.09018 + 2729.92 / (273.2 + T), where T is the temperature in degrees Celsius, and pH = −log10[H+] (Emerson et al., 1975). Temperature, ionic strength and pressure will also affect ammonia speciation by affecting ammonia pK, although to a lesser extent than pH (Ip et al., 2001). NH+ 4 is charged and larger than NH3, and because of this, ammonium ion is thought to be much less permeable through biomembranes than unionized ammonia (Knepper et al., 1989; McDonald et al., 1989). Thus increases in pH result in an increase of fNH3 and consequently a large increase in ammonia toxicity, due to increased uptake (USEPA, 1999). Indeed, in freshwater at 25 °C and pH 7 the fNH3 corresponds to 0.58% of total ammonia, and an increase in pH to 8 or 9, will increase this fraction to 5.49% (9.5 fold) and to 36.76% (64 fold), respectively. The gas permeability of membranes is dependent on composition and can be correlated with membrane fluidity (Lande et al., 1995). Ammonia is hypothesized to permeate membranes though the kinks in unsaturated fatty acids. In this study five experiments were conducted (acute ammonia exposure, environmental pH variation, boundary layer pH manipulation, in vivo pharmacological inhibition, and air versus ammonia exposure) with the aim of determining what roles branchial Na+/K+ATPase, H+-ATPase, and Rhcg have in ammonia tolerance and how the weatherloach copes with ammonia loading conditions. Immunofluorescence microscopy was used to study the distribution of these transporters in the gill, while semi-quantitative PCR was utilized to determine changes in Rhcg gene expression. Membrane fluidity

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measurements were also made in order to determine if modulation of membrane permeability plays a role in ammonia tolerance. For comparative purposes, the ammonia LC50 value was also determined for the weatherloach. 2. Materials and methods 2.1. Animals Adult weatherloaches were purchased from the main wet market in Yuen Long Hong Kong and stabilized at City University of Hong Kong where some of the studies were preformed. Some fish were transported to CIMAR (Centro Interdisciplinar de Investigação Marinha e Ambiental) by air freight for further work. Fish were transported with minimal water. Upon arrival fish were maintained in 100 L glass aquaria containing dechlorinated Porto city tap water (Na+ 0.5 mM, hardness 50 mg/l CaCO3, pH 8) for at least 1 week and maintained on a commercial diet (Granured, Sera Gmb, Germany). Animals were fasted for 48 h prior to experimentation. 2.2. Experiments 2.2.1. Acute ammonia exposure A 96 h LC50 experiment was carried out in order to determine a sublethal dose of ammonia for use with subsequent experiments and for the collection of tissue. A static-renewal test was conducted. Every 24 h half of the test solution was replace in the test aquaria. Fish [3.10 ± 1.09 g, mean ± SD (standard deviation)] were divided into five groups: control, 1, 5, 10 and 25 mM NH4Cl (n = 12 in each group). The water was buffered with Tris–HCl to a final concentration of 10 mM to stabilize the pH at ~8. During the experiment the pH was maintained at 7.89 ± 0.03 and temperature was kept at 26.5 ± 0.8 °C (Table 1). Water temperature and pH were measured throughout the experiment to determine NH3 and NH+ 4 fractions. Fish were not fed during the course of the experiment. The NH4Cl concentration lethal to 50% of the test fish (LC50) was determined using the Trimmed Spearman–Karber method. The number of moribund fish, determined as fish not responding to prodding, was recorded every 24 h for LC50 calculations, and moribund animals were removed. At the end of the experiment (96 h) the remaining fish were euthanized by an overdose of neutralized tricane methanesulphonate [1:5000 (w/v), Aquapharm UK]. Fish were measured to the nearest millimetre and weighed to the nearest milligram. The caudal peduncle was severed and exuded blood was sampled in heparinized capillary tubes, centrifuged, plasma collected, and immediately frozen in liquid nitrogen. Gill tissue was excised and placed in SEI buffer (150 mM sucrose, 10 mM Na2EDTA, 50 mM imidazole, pH adjusted to 7.3), and immediately frozen in liquid nitrogen. A piece of epaxial white muscle was excised and immediately frozen in liquid nitrogen. All samples were stored at −80 °C, until analysed. Gills of control fish were also fixed in 3% paraformaldehyde (PFA) in phosphate buffered saline (PBS, 137 mM NaCl, 2.7 mM KCl, 7.8 mM Na2HPO4, 1.5 mM KH2PO4, pH 7.4), for 12–24 h at room temperature (rt).

Table 1 Conditions of the LC50 test, conducted at pH 7.89 and 26.5 °C (TAN — total ammonia nitrogen). TAN

pH

NH+ 4 (µM)

NH3 (µM)

NH3 (%)

0 1 5 10

7.89 7.89 7.89 7.89

–

–

953 4,766 9,532

47 234 468

– 4.68 4.68 4.68

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2.2.2. Environmental pH variation The effect of environmental pH on ammonia tolerance was studied, since pH affects ammonia speciation. In aqueous solution ammonia exists as two species, NH3 and NH+ 4 , an increase in pH will increase the NH3 fraction. The aim of this experiment was to address the question of the relative toxicity of NH3 and NH+ 4 . Three pH levels were tested (pH 7, 8 and 9) in both control and ammonia conditions. Fish (2.80 ± 1.06 g, mean± SD) were divided into six groups: Ctrl pH 7, Ctrl pH 8, Ctrl pH 9 and HEA at pH 7, pH 8, and pH 9 (n = 4). For ammonia exposure a sublethal level was used, 6.3 mM NH4Cl (2 mM less than the LC50 value determined at pH 7.89 in the acute ammonia exposure experiment, corresponding to 375 μM NH3). Water temperature was maintained at 25.3 ± 0.8 °C (Table 2), and fish were not fed during the entire course of experiment. After 96 h exposure, survivors were euthanized and the same sampling procedure of the previous experiment was used. 2.2.3. Boundary layer pH manipulation (HEPES buffering) The importance of gill boundary layer acidification and NH+ 4 trapping to ammonia excretion was assessed by buffering the pH of the water with 10 mM HEPES buffer (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) at different water pHs (7 and 8). This buffer will eliminate the boundary layer pH difference with the bulk water. Fish (7.01 ± 1.82 g, mean ± SD) were divided into two groups: pH 7 and pH 8 (n = 6 in each group). An initial 3 h control flux with dechlorinated tapwater at pH 7 (adjusted with HCl) or pH 8 without HEPES was followed by 3 h and 24 h experimental flux periods with HEPES buffered water. Water samples were taken at 0, 1, 2, 3 and/or 24 h for measurement of total ammonia and titratable acidity. 2.2.4. In vivo pharmacological inhibition of ammonia flux The role of some ion transport proteins in ammonia excretion was studied using a pharmacological approach. The V-type H+-ATPase inhibitor bafilomycin A1 (0.1 μM; LC Laboratories), sodium transport inhibitor amiloride (0.1 mM; Sigma Aldrich), or chloride transport inhibitor disulfonic stilbene (SITS 0.1 mM; Sigma) were used. Ammonia flux rates were measured over a 3 h period as described in the previous experimental series. The bafilomycin experiment was repeated and titratable acidity also measured. Water and plasma samples were collected as described earlier for ammonia measurements. 2.2.5. Air versus ammonia exposure In order to determine if membrane permeability and Rhcg (Slc42a3) expression are involved in ammonia excretion in weatherloach gill, groups of animals were divided into three groups and acclimated under control conditions (dechlorinated tap water), high ammonia (30 mM NH4Cl at pH 7.2 and 21 °C in dechlorinated tap water, corresponding to 200 μM NH3), or aerial exposure (1 mL of dechlorinated tap water) for 7 days similar to conditions described by Tsui et al. (2002). At the end of the experiment the fish were euthanized as described previously and gill tissue excised and immediately frozen in liquid nitrogen, and stored at −80 °C.

Table 2 Conditions of the pH experiment, conducted at pH 7, 8 and 9, at 25.3 °C (TAN — total ammonia nitrogen). TAN

pH

NH+ 4 (mM)

NH3 (mM)

NH3 (%)

0 0 0 6.3 6.3 6.3

7.15 7.95 8.65 7.11 7.96 9.03

– – – 6.25 5.98 3.88

– – – 0.05 0.32 2.42

– – – 0.75 5.08 38.38

2.3. Analysis 2.3.1. Ammonia measurements Ammonia content in plasma and muscle were measured using an enzymatic microplate technique, modified from Bergmeyer and Beutler (1983). Water ammonia concentrations were measured using a modified quantitative microplate technique from Verdouw et al. (1978). For details see Moreira-Silva et al. (2009). 2.3.2. Net ammonia and acid fluxes Total ammonia was measured in 0, 1, 2, 3 and 24 h samples for the calculation of net ammonia flux (JAmm) (µmol ammonia kg− 1 h− 1). Titratable acidity flux rate (JTA) (µEq ·H+ mL− 1) was calculated through titratable alkalinity measurements (see McDonald and Wood, 1981) in 3 and 24 h water samples and the net acid flux calculated as JAcid = JAmm + JTA (µEq kg− 1 h− 1), signs considered. 2.3.3. ATPase activity Activities of Na+/K+-ATPase and V-type H+-ATPase were measured using a modified microplate technique from McCormick (1993) with ouabain and bafilomcyin A1, respectively, as specific inhibitors as described in Moreira-Silva et al. (2009). ATPase activities are expressed as µmol ADP h− 1 mg protein− 1. 2.3.4. Immunofluorescence (IF) microscopy Gills were fixed in PFA (3% paraformaldehyde) in PBS, paraffin embedded, sectioned and processed for immunofluorescence microscopy as described by Wilson et al. (2007). Sections were incubated with mouse monoclonal (clone α5) anti-Na+/K+-ATPase antibody (1:100) and affinity purified rabbit polyclonal anti-H+-ATPase (B2/ BvA1) antibody (1:500) or rabbit polyclonal RhCG1 (Ab740) antibody (1:200), both diluted in 1% BSA/TPBS (0.05% Tween-20/phosphate buffered saline, pH 7.4)/0.05% sodium azide, overnight at 4 °C in a moist chamber. Negative control incubations were performed simultaneously under the same conditions, using isotyped hybridoma culture supernatant (clone J3), and either preimmune rabbit serum or antibody pre-absorbed with excess peptide (pre-absorbed overnight at 4 °C on an orbital shaker) equivalently diluted as the primary antibodies. Secondary incubations were performed with goat antimouse Alexa Fluor 488 and goat anti-rabbit Alexa Fluor 594 conjugated secondary antibodies (Invitrogen S.A., Barcelona, Spain) diluted 1:400 in TPBS, for 1 h at 37 °C. Nuclei were counterstained with 4',6-Diamidino-2-phenylindole (DAPI) and coverslips were mounted with a glycerol based mounting media (10% Mowiol/40% glycerol/2.5% 1,4-Diazabicyclo[2.2.2]octane (DABCO) in 0.1 M Tris, pH 8.5) and observed on an epifluorescence microscope (Leica Microsystems DM6000 B, Germany). Images of fluorescent staining were captured with a Leica DFX340 camera, along with the corresponding differential interference contrast (DIC) image. Plates were assembled using Adobe Photoshop CS3 software, and images enhanced while maintaining the integrity of the data. No quantitative analysis of the images was performed. 2.3.5. SDS-PAGE and immunoblotting (IB) Remaining sample supernatants from the ATPase activity assay were used for sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoblotting. Supernatants were diluted to 0.5 µg/µL in Laemmli's buffer as described by Wilson et al. (2007), with equal amounts of proteins loaded per lane (15 µg/lane). Proteins were separated on 10% resolving and 4% stacking SDS-polyacrylamide gels by electrophoresis under denaturing conditions (running buffer — 25 mM Tris, 191 mM glycine, 0.1% SDS), using a vertical apparatus Mini-Protean III System (Bio-Rad, Hercules, CA, USA). After size separation, proteins were transferred to polyvinylidene difluoride (PVDF) Hybond ECL membranes (GE Healthcare, Carnaxide, Portugal), using a Trans-Blot SD Semi-Dry Transfer Cell (Bio-Rad). Resulting

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blots were incubated with the primary antibodies, diluted in 1% BSA/ TTBS/0.05% azide (Na+/K+-ATPase α5, 1:500; H+-ATPase B2/BvA1, 1:1000; tubulin 12G10, 1:200), overnight at room temperature. Secondary incubations were performed with either goat anti-mouse antibody (for mouse monoclonal primary antibodies) or goat antirabbit antibody (for rabbit primary antibodies) horse radish peroxidase (HRP) conjugates, diluted (1:25,000) in TTBS. Between incubation steps blots were rinsed in TTBS, (20 mM Tris, 500 mM NaCl, and 0.05% Tween 20). Proteins were detected by enhanced chemiluminescent ECL (ECL Western Blotting Detection, GE Healthcare), using X-ray film (ECL Hyperfilm, GE Healthcare). The film was scanned (Agfa Duoscan T1200, Mortsel, Belgium), area and average intensity was determined for semi-quantification using an image analysis software program (SigmaScan Pro 5.0, SPSS, Chicago, IL, USA). The results are expressed as the relative difference from the control group of the ratio of either α5 (α subunit of Na+/K+-ATPase) or B2/BvA1 (H+-ATPase B subunit) to 12G10 (α tubulin) protein expression. 2.3.5.1. Antibodies. The Na+/K+-ATPase α5 antibody, a mouse monoclonal antibody raised against α-subunit of chicken NKA (Fambrough, D.M., Developmental Studies Hybridoma Bank), was used. This is a pan specific antibody for Na+/K+-ATPase a subunit and is not isoform specific (Takeyasu et al., 1988) is routinely used to detect gill Na+/K+-ATPase mitochondrion rich cells (for review see Wilson and Laurent, 2002). The affinity purified H+-ATPase B2/BvA1 antibody, a rabbit polyclonal antibody raised against a synthetic peptide of B-subunit of the eel H+-ATPase (Davids Biotechnologie GmbH), was used (Wilson et al., 2007). This antibody crossreacts with H+-ATPase B subunit in a number of fishes (A. anguilla Wilson et al., 2007; Petromyzon marinus Reis-Santos et al., 2008; Acipenser transmontanus Baker et al., 2009). The Rhcg1 rabbit polyclonal antibody (Ab740) raised against a recombinant zebrafish Rhcg1 COOH terminus (amino acid residues 425–488) protein (Nakada et al., 2007b) was used. This antibody with the preimmune serum was the generous gift of S. Hirose (Tokyo Institute of Technology, Japan). The tubulin 12G10 antibody, a mouse monoclonal antibody raised against Tetrahymena thermophila and Tetrahymena pyriformis αtubulin (Frankel, J. and Nelsen, E.M., Developmental Studies Hybridoma Bank) was used (Thazhath et al., 2002) as a loading control for Immunoblotting, since the α-tubulin expression with different environmental ammonia levels did not change. The α5, J3 and 12G10 antibodies were obtained as culture supernatant from the Developmental Studies Hybridoma Bank, University of Iowa, IA, USA, under contract N01-HD-7-3263 from NICHD. 2.3.6. Semi-quantitative PCR to assess Rhcg expression 2.3.6.1. Total RNA isolation and cDNA synthesis. Total RNA was isolated and cDNA prepared as described by Gonçalves et al. (2007). In brief, pieces of frozen gill tissue were ground under liquid nitrogen with a mortar and pestle and total RNA was isolated using the acid guanidinium thiocyanate-phenol-chloroform method (Chomczynski and Sacchi, 1987) using phase-lock gel tubes (Eppendorf AG, Hamburg, Germany). RNA samples were resuspended in DEPC (diethylpyrocarbonate) treated water, their concentration and purity

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was measured spectrophotometrically, at 260 and 280 nm (Jenway 6405 UV/VIS Spectrophotometer, Dunmow, England). Integrity of RNA samples was assessed by electrophoresis (1.2% Agarose with formaldehyde gel) (Mini-Sub Cell GT Cell, Bio-Rad). The cDNA was synthesized from 1 μg of total RNA in the presence of oligo-dT primer and M-MuLV Reverse transcriptase (BIORON GmbH, Ludwigshafen, Germany). Twenty µl reactions were carried out in a MJ MINI Personal Thermal Cycler (Bio-Rad). 2.3.6.2. Gene isolation. Primers were designed from conserved regions of Slc42a3 (D. rerio, Fugu rubripes, Oncorhynchus mykiss, Tetraodon nigroviridis; fwd 5'-GCACACTGTTCCTGTGGATG-3'; rev 5'-CAGCAGGATCTCCCCAGA3'). β-actin was used as a “house-keeping” gene and the primers were originally designed for Sparus aurata (fwd 5'-GGCCGCGACCTACAGACTAC3'; rev 5'-ACCGAGGAAGGATGGCTGGAA-3'; Santos et al., 1997). Expected sizes of the amplification products were 585 and 250 bp, respectively. PCR was performed with 1 μL of sample cDNA, 2 mM MgCl2 (for both Rhcg and β-actin) and 1 unit of DFS-Taq DNA polymerase (BIORON) in a 25 µl reaction volume. Single bands were retrieved from 1% agarose gels in TBE (Tris–borate–EDTA) buffer (GFX column, GE Healthcare) and directly sequenced (StabVida, Oeiras, Portugal). The PCR profiles for the semi-quantitative analysis were as follows. β-actin: a first cycle of 2 min at 94 °C, was followed by 30 s at 94 °C, 30 s at 60 °C, 45 s at 72 °C for 30 cycles and 72 °C for 5 min. Slc42a3: a first cycle of 2 min at 94 °C, was followed by 30 s at 94 °C, 30 s at 52 °C, 30 s at 72 °C for 35 cycles and 72 °C for 5 min. Trial runs were performed to determine the optimal cycle conditions for semi-quantitative analysis (Marone et al., 2001). Reactions were carried out in the MJ MINI Personal Thermal Cycler (Bio-Rad). 2.3.6.3. Gel image acquisition and quantitative analysis. PCR products were loaded onto 2% agarose gels in TBE buffer run at 80 V. A 100 bp DNA ladder (BIORON) was run on every gel to confirm expected size of the amplification product. Gels were stained with ethidium bromide and images acquired with a Kodak EDAS 290 image acquisition system. Band area and average intensity was determined for semi-quantification using an image analysis software program (SigmaScan Pro 5.0, SPSS). The results are calculated as the ratio of Rhcg (Slc42a3) with β-actin mRNA in order to normalize expression and presented as a fold change from the control group. It should be noted that although semi-quantitative analysis using RT-PCR has some limitations, in the present study we performed a careful validation of the technique for Rhcg expression analysis (following recommendations from the review of Freeman et al., 1999). 2.3.7. Gill membrane fluidity Gill plasma membranes were extracted and purified using the method of Daveloose et al. (1993), based on density gradient centrifugation. In brief, samples were pooled into three groups (each group having 7, 7 and 8 samples) and homogenized with a Dounce homogenizer (pestle A) on ice. To separate the membranes on a sucrose step gradient, a Beckman Coulter Optima™ Max with a MLS-50 Swinging-Bucket Rotor (Beckman Coulter, Fullerton, CA, USA) ultracentrifuge was used. To determine membrane enrichment, total protein was determined by the BCA method (Smith et al., 1985) and Na+/K+ATPase and lactate dehydrogenase enzymatic assays were performed.

Table 3 LC50 (lethal concentration to 50% of the test fish) values and 95% confidence intervals (in parentheses) to NH4Cl in weatherloach, at 26.5 °C and pH 7.89 (TAN — total ammonia nitrogen, N — nitrogen). 24 h TAN mM NH3 µM mg N/L TAN mg NH3-N/L

11.99 561 168 7.86

48 h (9.62–14.95) (450–700) (135–209) (6.30–9.80)

9.56 447 134 6.27

72 h (7.64–11.97) (357–560) (107–168) (5.01–7.85)

9.66 452 135 6.33

96 h (7.70–12.11) (360–567) (108–170) (5.05–7.94)

8.31 389 116 5.45

(6.90–10.00) (323–468) (97–140) (4.52–6.55)

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(control, ammonia and air) and compared by ANCOVA (R software, 2008). 3. Results 3.1. LC50 determination and mortalities The concentration of NH4Cl that was lethal to 50% of the weatherloach at 26.5 ± 0.8 °C and pH 7.89 ± 0.03, within 24, 48, 72 and 96 h was estimated as 11.99, 9.56, 9.66 and 8.31 mM, respectively (Table 3). An overlap of the 95% confidence limits within all the LC50 determinations was observed. When expressed as NH3, the 24, 48, 72 and 96 h LC50 values were 561, 447, 452 and 389 μM, respectively. During the acute ammonia exposure, 100% mortality occurred in the group exposed to 25 mM NH4Cl. In the pH experiments the ammonia group 6.3 mM NH4Cl pH 9 was terminated since all specimens died Fig. 1. Effect of different ammonia exposure levels (control, 1 mM, 5 mM and 10 mM NH4Cl) on the concentration of ammonia in the muscle. Values are mean±SE (n = 8– 11, with the exception of 10 mM group n = 2), bars with like characters are not significantly different (P > 0.05). Abbreviation: TAN, total ammonia nitrogen.

Membrane fluidity was measured using a fluorimetric method (fluorescence anisotropy) (Crockett and Hazel, 1995). In brief, fluorescence anisotropy of the probe 1,6-diphenyl-1,3,5-hexatriene (DPH) incubated with the membranes, was measured with a POLARstar Galaxy microplate fluorometer (BMG Laboratories, Germany), with excitation and emission monochrometers set at 360 nm and 430 nm respectively, in a temperature gradient (27, 29, 34, 37 and 39 °C). The anisotropy of the probe DPH gives an indication of lipid order with higher anisotropy corresponding to a more ordered membrane. A more ordered membrane is less permeable. 2.4. Statistics Results are presented as means ± S.E.M. (standard error of the mean) or 95% confidence interval. Comparison of LC50 values was made by examining overlap of confidence limits as recommended by APHA (1989). One-way analysis of variance followed by Dunn's or Student–Newman–Keuls post-tests and repeated measurements analysis of variance followed by SNK–Friedman post-test were used to compare differences between means when applicable. For two group comparisons, t-test or Mann–Whitney rank sum tests were performed. A Pearson Product Moment Correlation (PPMC) was carried out (SigmaStat 3.0, SPSS, Chicago, IL, USA). For membrane fluidity analysis, linear regressions were performed on each group

Fig. 2. Effect of different pH conditions (pH 7, 8 and 9 control; with and without 6.3 mM NH4Cl) on muscle ammonia concentrations. Values are mean±SE (n = 7–10), bars with like characters are not significantly different (P > 0.05). The × mark indicates the terminated ammonia group at pH 9. An asterisk indicates significant differences from the control group at the same pH (P < 0.05).

Fig. 3. Effects of HEPES buffering (10 mM) at pH 7 and 8 on ammonia (JAmm, net ammonia flux) and net acid flux (JAcid = JAmm +JTA, JTA being the titratable acidity flux rate in M. anguillicaudatus. (a) 3 h control flux, dechlorinated Porto tap water at pH 7 or 8. (b) 3 h and (c) 24 h experimental flux, water buffered at pH 7 and 8 with HEPES 10 mM. Values are mean±SE (n = 6) (P > 0.05), bars with like characters are not significantly different (P > 0.05), asterisk indicates significant differences in the same flux period between different pHs (P < 0.05).

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experimental flux. However, in the 24 h experimental flux JTA (P = 0.024) and JAcid (P = 0.017) were significantly higher at pH 8. Acute exposure to bafilomycin A1 significantly inhibited JAmm by over 60% whereas neither amiloride nor SITS had any effect on JAmm (Fig. 4). Bafilomycin exposure resulted in a tripling of plasma ammonia levels (P =0.007), and a significantly lower JAcid (635.65 ± 127.40 µEq kg− 1 h− 1) compared to control animals (1013.62 ± 219.19 µEq kg− 1 h− 1) (P=0.022). 3.4. Gill Na+/K+-ATPase and V-type H+-ATPase activities and protein expression

−1

−1

Fig. 4. Effects of pharmacological inhibitors on net ammonia fluxes (μmol kg h ) in loaches over a 3 h flux period. Fish were exposed to the H+-ATPase inhibitor bafilomycin A1 (baf 0.1 μM), sodium transport inhibitor amiloride (AMIL 0.1 mM), or chloride transport inhibitor disulfonic stilbene (SITS 0.1 mM) or kept under control conditions (Ctrl). The inset shows plasma total ammonia levels in control and bafilomycin exposed fish.

within 4.5 h of starting of the experiment (× in Fig. 2). In the HEPES experiment no mortalities were observed.

In the acute ammonia exposure experiment no significant differences (P>0.05) were found in either Na+/K+-ATPase or H+-ATPase activities with ammonia exposure (Tables 4 and 5). Protein level expression, measured by immunoblotting, also indicated no significant differences in either Na+/K+-ATPase α subunit or H+-ATPase B subunit protein expression with ammonia exposure (Tables 4 and 5; Figs. 5 and 6). In the pH experiment branchial Na+/K+-ATPase and H+-ATPase activities did not change with pH or with pH together with ammonia exposure. There were also no changes reflected at the protein level as measured by immunoblotting (Tables 4 and 5; Figs. 5 and 6).

3.2. Muscle ammonia 3.5. Immunofluorescence In the acute ammonia exposure experiment there was a significant increase (P < 0.05) in the muscle ammonia content of fishes acclimated to 5 and 10 mM NH4Cl (Fig. 1). This shows that there was an increased building up of ammonia when environmental ammonia was elevated. Moreover there was a good positive correlation (r = 0.988, P = 0.0122 with PPMC) between environmental ammonia concentration and tissue ammonia accumulation. In the pH experiment there was a gradual but significant muscle ammonia content increase in the control groups (P < 0.05) as environmental pH increased (Fig. 2). Ammonia content in muscle of fish exposed to ammonia at pH 8 was significantly higher (P < 0.001) than in fish exposed to ammonia at pH 7. At pH 7 a higher ammonia accumulation in the muscle of fish exposed to ammonia (2.73 µmol g− 1) compared to control fish (0.54 µmol g− 1) was observed (P < 0.05). The largest ammonia accumulation occurred at pH 8 in ammonia exposed fish (16.33 µmol g− 1) when compared to controls (2.69 µmol g− 1) (P < 0.001).

In weatherloach gills strong Na+/K+-ATPase immunoreactivity (IR) was identified in discrete cells in both the filament and lamellar epithelia. Whole cell body staining indicative of localization to the cellular tubular system was typical. The shape of these Na+/K+ATPase-IR cells ranged from flasked shaped to cuboidal (Figs. 7a,c,e and 8a,c,e). The immunolocalization of H+-ATPase was apical in epithelial cells in a discontinuous pattern (Fig. 7a,b,e). The apical staining pattern was either diffuse in the subapical, supranuclear region or sharply along the apical boarder of the cell. These cells did not show Na+/K+ATPase immunoreactivity. Pre-absorption of the antibody with excess peptide eliminated the apical staining pattern. Rhcg1-like immunoreactivity was found apically in a similar pattern as H+-ATPase (Fig. 8a,b,e). However, the frequency of Rhcg1 immunoreactivity appeared greater. Incubation of sections with preimmune serum produced negligible staining.

3.3. Net ammonia and acid fluxes

3.6. Semi-quantitative PCR to assess Rhcg expression

Buffering the water by the addition of HEPES (see Fig. 3) at pH 7 did not inhibit net ammonia flux (JAmm) although at pH 8 a decrease in JAmm was observed when HEPES was added in both 3 h, and 24 h experiments. JAcid (net acid flux) and JTA (titratable acidity) did not change when comparing the 3 h Ctrl, 3 h Exp, and 24 h Exp fluxes, within pH 7 or pH 8 groups. In comparisons between the pH 7 with pH 8 groups, no differences were found in JAmm, JTA, or JAcid during the initial 3 h control flux or 3 h

The PCR experiments to isolate Rhcg (Slc42a3) resulted in single band. Direct sequencing followed by BLASTx (GenBank, version 2.2.19, Altschul et al., 1997) sequence analysis confirmed that, indeed, the retrieved band corresponds to the gene ortholog of Slc42a3-1 in M. anguillicaudatus. The partial Slc42a3-1 nucleotide sequence has been deposited in GenBank with the following accession number FJ982777. Further comparisons with other vertebrate sequences highlight a substantial degree of conservation with percentage amino acid

Table 4 Branchial Na+/K+-ATPase activity (µmol ADP h− 1 mg protein− 1) and α subunit expression (as a ratio with α-tubulin and relative to the control group) in M. anguillicaudatus following acute ammonia exposure and environmental pH variation with or without ammonia exposure (6.3 mM TAN) (TAN — total ammonia nitrogen). Acute ammonia exposure

Ctrl

1 mM

5 mM

10 mM

Activity Protein expression

0.77 ± 0.19 1.00 ± 0.22

0.70 ± 0.04 0.69 ± 0.14

0.76 ± 0.09 0.88 ± 0.17

0.55 ± 0.19 0.90 ± 0.07

pH variation

Ctrl pH 7

Ctrl pH 8

Ctrl pH 9

Amm pH 7

Amm pH 8

Activity Protein expression

0.72 ± 0.10 1.00 ± 0.14

0.69 ± 0.07 1.02 ± 0.17

0.87 ± 0.09 0.94 ± 0.07

0.79 ± 0.07 1.00 ± 0.07

0.61 ± 0.07 1.39 ± 0.09

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Table 5 Branchial V-type H+-ATPase activity (µmol ADP h− 1 mg protein− 1) and B subunit expression (as a ratio with α-tubulin and relative to the control group) in M. anguillicaudatus following acute ammonia exposure and environmental pH variation with or without ammonia exposure (6.3 mM TAN). Acute ammonia exposure

Ctrl

1 mM

5 mM

10 mM

Activity Protein expression

0.22 ± 0.04 1.00 ± 0.19

0.10 ± 0.02 0.90 ± 0.07

0.17 ± 0.05 1.19 ± 0.07

0.25 ± 0.15 1.39 ± 0.60

pH variation

Ctrl pH 7

Ctrl pH 8

Ctrl pH 9

Amm pH 7

Amm pH 8

Activity Protein expression

0.10 ± 0.04 1.00 ± 0.14

0.06 ± 0.02 1.06 ± 0.23

0.11 ± 0.02 0.94 ± 0.07

0.18 ± 0.04 1.00 ± 0.09

0.14 ± 0.05 1.46 ± 0.17

identity ranging between 62% (Homo sapiens) with SLC42A3 and 79% (D. rerio) with Slc42a3-1 and 68% (D. rerio) with Slc42a3-2. A Rhesus C-glycoprotein (Slc42a3) phylogenetic tree (Fig. 9) formed two main clusters: one composed by mammalian, chicken and frog sequences and the other one composed by fishes. The fish cluster is further subdivided into two smaller clusters corresponding to isoform 1 and isoform 2. Weatherloach Slc42a3 falls into the isoform 1 fish cluster. Following acclimation to different conditions (Fig. 10), Rhcg (Slc42a3) mRNA levels were significantly higher (P < 0.05) in fish aerially exposed when compared to controls. No significant differences were found in the ammonia group compared to the control. 3.7. Membrane fluidity Anisotropy was temperature dependent and regressions of control, ammonia and aerial exposure were parallel. Gill fluorescence anisotropy was found to be lower, greater membrane fluidity, in the aerially exposed fish (intercept = 0.283417 − 0.045917, P = 0.0299) compared to control (intercept = 0.283417), while a higher fluorescence anisotropy, lower membrane fluidity, although non-significant was observed in ammonia exposed fish (intercept = 0.283417 + 0.016978, P = 0.3455). 4. Discussion The weatherloach had a very high ammonia tolerance as demonstrated by its acute 96 h LC50 to ammonia of 389 μM NH3, when compared to the range of 5–59 μM for salmonids or 19–176 μM NH3 for non-salmonid species (USEPA, 1999; these ranges were calculated from the species 96 h LC50 values presented in “Appendix 4. Acute values”. The species values were averaged from all tests on a given species). This value exceeds the most tolerant freshwater fish species referred to in the USEPA (1999) survey, which includes Ictalurus punctatus, Gambusia affinis, Notropis lutrensis that have 96 h LC50 values of 123, 156 and 176 μM NH3, respectively. However, the loach value is comparable to another tropical air-breathing species the African sharp-tooth catfish Clarias gariepinus that has a 96 h LC50 of 380 μM NH3 (Oellermann, 1995), but is still well short of the exceptionally tolerant swamp eel Monopterus albus [1092 µM NH3; 193 mM TAN (total ammonia nitrogen) at pH 7.0 Ip et al., 2004]. The

Fig. 5. Representative immunoblot showing branchial Na+/K+-ATPase α subunit (α5) and V-type H+-ATPase B subunit (B2) expression in M. anguillicaudatus following acute ammonia exposure. The α tubulin was used as a loading control.

weatherloach ranks amongst the very ammonia tolerant freshwater fishes. In the weatherloach H+-ATPase activity and expression were found in gill, and localized by immunofluorescence microscopy to a distinction population of cells found in both filament and lamellar epithelia. The H+-ATPase labelling was generally concentrated in the apical region of the cell. These immunoreactive (IR) cells did not have strong immunoreactivity for Na+/K+-ATPase which would indicate that they are not “chloride” type mitochondrion-rich cells (Wilson and Laurent, 2002). In the loach, Cobitis taenia, Pisam et al. (1990) characterized two types of mitochondrion-rich cells (MRC), light and dark based cytoplasmic density and differences in cell fine structure. However, on the basis of cell shape and location there is no clear correlation of these morphotypes and the Na+/K+-ATPase and H+ATPase IR cells described in this study In other fishes, H+-ATPase has been localized to both gill epithelial mitochondrion-rich cells and pavement cells (see Evans et al., 2005). Work by Lin and Randall (1995) and more recently by Nawata et al. (2007) indicates that H+-ATPase contributes to gill boundary layer acidification. This has also been elegantly demonstrated in zebrafish skin using a vibrating probe technique to measure H+ and NH+ 4 fluxes across individual cells (Shih et al., 2008). In addition, both H+-ATPase knock down and bafilomycin A1 were demonstrated to reduce ammonia efflux (Shih et al., 2008). In our study, bafilomycin also reduced ammonia excretion and acid flux but activity and protein expression of this transporter did not change in weatherloach exposed to higher ammonia levels. It is possible that the basal levels of activity of H+-ATPase produce enough H+s to trap ammonia as NH+ 4 and that an increase in expression was not required. Or alternatively a similar mechanism that exists in kidney A-type intercalated cells is employed. Specifically, the insertion of H+-ATPase vesicles into the apical membrane from an intracellular vesicle pool (see Brown and Breton, 1996). The very similar pattern of H+-ATPase and Rhcg1 immunostaining suggests that these two transporters are likely colocalized to the same cell type. This has been demonstrated in zebrafish using a combination of lectin histochemisty, in situ hybridization and immunocytochemistry (Nakada et al., 2007b, Shih et al., 2008). Furthermore, in the study by Shih et al. (2008) elimination of acidification by the H+-ATPase by

Fig. 6. Representative immunoblot showing branchial Na+/K+-ATPase α subunit (α5) and V-type H+-ATPase B subunit (B2) expression in M. anguillicaudatus following environmental pH variation with or without ammonia exposure (6.3 mM TAN). The α tubulin was used as a loading control.

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Fig. 7. (a) Localization of V-type H+-ATPase (B subunit; green) and Na+/K+-ATPase (a5; red) double immunofluorescent labelling of a sagittal section of a M. anguillicaudatus gill filament toward the afferent side. To provide structural information nuclei have been counterstained with DAPI (a,d,e; blue) and the DIC image overlaid. A higher magnification view of the separate (b) H+-ATPase, and (c) Na+/K+-ATPase immunoreactive cells as well as merger image are given. Arrowheads indicate Na+/K+-ATPase immunoreactive cells, while arrows and crossed arrows indicate sharp and diffuse H+-ATPase staining, respectively. Asterisks indicate erythrocytes that showed weak fluorescence. Scale bar = 50 μm (a) and 10 μm (b–e). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

knockdown or bafilomcyin reduced ammonia flux. Since the Rhcg likely functions as a bidirectional NH3 channel, boundary layer acidification is an important mechanism for maintaining an outward concentration gradient for NH3 necessary for excretion (Nawata et al., 2007; Shih et al., 2008). However, in some studies Rhcg1 has been found to colocalize to NKA-IR cells (T. rubripes; Nakada et al., 2007a) and thus the pattern of Rhcg1 expression can be species specific. In the loach, aerial exposure resulted in a significant increase of Rhcg1 mRNA expression and membrane fluidity also showed a significant increase. Both of these changes would increase gill

Fig. 8. (a) Double immunofluorescent localization of Rhcg1(green) and Na+/K+-ATPase (red) in a sagittal section of the same gill filament seen in Fig. 7. A region at higher magnification is shown with the colour channels separate [(b) Rhcg1, (c) Na+/K+ATPase, (d) DAPI] and (e) merged with the DIC overlay. Arrowheads indicate Na+/K+ATPase immunoreactive cells, while arrows and crossed arrows indicate strong sharp and weak diffuse apical Rhcg1 staining, respectively. Asterisks indicate erythrocytes that showed weak fluorescence. Scale bar = 50 μm (a) and 10 μm (b–e). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

ammonia permeability. Thus although the intestine has been implicated as an important site of ammonia excretion through volatilization in weatherloach (Tsui et al., 2002), the gill may still have a role to play during aerial exposure. The mangrove killifish K. marmoratus, is also very ammonia tolerant like the weatherloach, surviving aerial exposure and volatilizing ammonia although through the skin (Frick and Wright, 2002). In this fish, expression levels of Rhcg1 and 2 are high in gill (Hung et al., 2007) and exposure to high ammonia resulted in increases in Rhcg2 expression in both gill and skin, while Rhcg1 expression only increased in skin. Significant increases in both skin Rhcg1 and 2 expressions were found following aerial exposure; however, no results on gill were presented for comparison. In the loach no significant changes in Rhcg1 expression occurred in the ammonia exposed fish either. This might be related to the environmental ammonia level used or indicate that expression is regulated at another level. However, a consistent pattern appears to be emerging in which the branchial Rhcg1 is not responsive to ammonia exposure since studies with rainbow trout (Nawata et al., 2007; Tsui et al., 2009), and zebrafish (Nakada et al., 2007b) also found no differences in Rhcg1 expression with ammonia exposure. Instead, branchial Rhcg2 has been shown to be responsive to ammonia in both in vivo (Nawata et al., 2007) and in vitro (Tsui et al., 2009) studies of trout. Weatherloach exposed to water buffered with HEPES 10 mM at pH 8 showed a decrease in JAmm at 3 and 24 h, when compared to unbuffered control water. Buffering the water eliminates the gill surface boundary layer acidification since the buffer binds H+ arising from either CO2 hydration via carbonic anhydrase (CA) or direct H+ pumping or excretion. Buffering the water at pH 7 had no effect on JAmm. Other studies in rainbow trout support these results since JAmm was reduced when boundary layer acidification was eliminated with either Tris 0.4 mM at pH 8 (Wright et al., 1989), HEPES 5 mM at pH 8 (Wilson et al., 1994), or HEPES 5 mM at pH ranging from 7.7 to 8.2 (Salama et al., 1999). This reduction in JAmm in weatherloach exposed to buffered water confirms the importance of the gill boundary layer in ammonia excretion in this fish. In our study the JAmm, under control conditions at pH 8, was 342– 400 µmol kg− 1 h− 1. Chew et al. (2001) and Tsui et al. (2002) obtained JAmm in submerged weatherloach of around 18 µmol g− 1 day− 1, which corresponds to 750 µmol kg− 1 h− 1. This difference in net ammonia excretion rates might be related with differences in nutritional state and the shorter fasting period of 24 h used in the Singapore experiments compared to our experiments of 48 h (Fromm, 1963). Also the lower water pH (7.2) in Singapore would favour NH+ 4

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Fig. 10. (a) Rhcg mRNA expression in control (n = 3), ammonia (n = 4) and air exposed (n = 4) M. anguillicaudatus determined by semi-quantitative RT-PCR. Data are normalized to β-actin and expressed relative to the control group. The asterisk indicates significant difference from the control group (P < 0.05). (b) Representative ethidium bromide gel showing expression of Rhcg in control, ammonia and air exposure conditions.

Fig. 9. Rooted phylogenetic tree of Slc42a3 homolog sequences. Tree was constructed by the neighbor-joining method with 1000 bootstrap trials using ClustalX. Accession numbers are as follows: Tunicate (Ciona savignyi) Slc42a3 (AAY41908), African clawed frog (Xenopus laevis) Slc42a3 (AAH84943), Chicken (Gallus gallus) Slc42a3 (AAP49833), Pig (Sus scrofa) Slc42a3 (ABF69687), Rhesus monkey (Macaca mulatta) Slc42a3 (ABD72472), Chimpanzee (Pan troglodytes) Slc42a3 (AAX39717), Human (Homo sapiens) SLC42A3 (AAG02171), Rabbit (Oryctolagus cuniculus) Slc42a3 (AAK14653), House mouse (Mus musculus) Slc42a3 (AAF19373), Norway rat (Rattus norvegicus) Slc42a3 (NP_898876), Zebrafish (Danio rerio) Slc42a3-2 (BAF63791), Torafugu (Takifugu rubripes) Slc42a3-2 (AAM48579), Mangrove rivulus (Rivulus marmoratus) Slc42a3-2 (ABD83662), Stickleback (Gasterosteus aculeatus) Slc42a3 (ABF69690), Rainbow trout (Oncorhynchus mykiss) Slc42a3 (AAU89494), Mangrove rivulus Slc42a3-1 (ABN41463), Green pufferfish (Tetraodon nigroviridis) Slc42a3 (AAY41907), Torafugu Slc42a3-1 (AAM48578), Zebrafish Slc42a3-1 (AAM90586), and Weatherloach (Misgurnus anguillicaudatus) Slc42a3 (FJ982777).

trapping and thus ammonia excretion. Nevertheless, the values reported in the loach studies are all well within the range reported for freshwater teleosts (Wood, 1993). Most likely in the weatherloach, as in the majority of other freshwater teleost fishes (Wilkie, 2002), ammonia excretion occurs primarily according to a NH3 diffusion gradient not requiring active transport. In addition, levels of gill Na+/K+-ATPase activity are low as in other freshwater fishes, when compared to seawater fishes (Epstein et al., 1980; Karnaky, 1998; Moreira-Silva et al., 2009). These two factors make it improbable that Na+/NH+ 4 -ATPase is a significant mechanism in freshwater fish (Wilkie, 1997). Probably the weatherloach is not an exception and Na+/NH+ 4 -ATPase active transport is not used to excrete ammonia when exposed to HEA or to high environmental pH. Indeed, activity and protein level expression of branchial Na+/K+-ATPase did not change in weatherloach exposed to different ammonia levels or to different water pHs.

The increased build up of ammonia observed in the weatherloach exposed to 5 and 10 mM NH4Cl (at pH 7.89 and 26.5 °C) and the higher ammonia accumulation in the muscle of ammonia exposed fish (6.3 mM NH4Cl, 25.3 °C) compared to control fish at pH 7 and pH 8, confirmed that high environmental ammonia impaired net ammonia efflux and favored ammonia entry. Other authors (see review by Wood, 1993) have also shown a build up of ammonia, represented by an increase in plasma ammonia levels. Assuming that NH3 permeability is higher than NH+ 4 (Knepper et al., 1989; McDonald et al., 1989) in the loach as in other freshwater fishes, ammonia excretion will occur principally according to the transbranchial PNH3 gradient (see Wilkie, 2002). Therefore HEA will reverse this outward diffusion gradient and ammonia entry will occur (Wood, 1993). Indeed it has been shown in the present study that there was a good correlation between HEA and the ammonia accumulated in the muscle. Thus the loach does not seem exceptional in its ability to prevent the influx of environmental ammonia and therefore likely conforms to the common mechanism for aquatic ammonia excretion (passive NH3 diffusion). Other studies on weatherloach confirm the high tissue tolerance observed in the present study (Chew et al., 2001; Tsui et al., 2002). The weatherloach exposed to terrestrial conditions showed a build up of ammonia to around 15 µmol g− 1 after 48 h and 72 h (Chew et al., 2001). This value is similar to the one obtained in our study in fish exposed 10 mM NH4Cl (16.89 µmol g− 1). In the work of Tsui et al. (2002), the weatherloach were exposed to the subleathal dose of 30 mM NH4Cl at pH 7.2 (289 µM NH3) for 48 h and muscle ammonia increased to 18.9 µmol g− 1, which is similar to fish exposed to 10 mM at pH 7.89 (468 µM NH3) for 96 h (16.89 µmol g− 1), but markedly higher than fish exposed to a similar NH3 level of 234 µM NH3 (5 mM TAN at pH 7.89) with 7.70 µmol·g− 1 muscle ammonia. In control animals muscle ammonia levels were similar amongst these three studies (2.8–3.6 µmol g− 1 TAN). The gradual increase in muscle ammonia content as pH increases, in both control and ammonia exposed fish, is consistent with the observation that the increase in pH will increase the more permeant NH3 fraction of ammonia (Knepper et al., 1989; McDonald et al., 1989). High pH will also inhibit ammonia excretion which will lead to the retention of endogenously produced ammonia (Wilkie, 2002). In

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fact these results agree with other studies that have shown an increase in plasma ammonia in salmonids exposed to alkaline water pH (see Wilkie, 2002). Weatherloach exposed to ammonia (6.3 mM TAN) in alkaline water at pH 9.03 did not survive. Most likely they could not cope with the increase in the NH3 concentration to 2441 µM, which would have increased the ammonia body burden, and impaired ammonia efflux. In conclusion, the apical colocalization of Rhcg1 and H+-ATPase to non Na+/K+-ATPase immunoreactive cells in the gill and the increase in Rhcg mRNA abundance with air exposure suggest the involvement of this putative ammonia transporter in branchial ammonia excretion of this freshwater fish. Gill ammonia permeability is also increased with aerial exposure, by both the up-regulation of Rhcg and the increase in gill membrane fluidity. We found no correlation between branchial Na+/K+-ATPase expression and HEA to indicate a role in ammonia excretion. The weatherloach has high environmental and tissue ammonia tolerance and does not actively excrete ammonia. The H+-ATPase clearly has a role in ammonia excretion through boundary layer acidification, although without the need to increase branchial expression levels. Acknowledgements The present study was supported by the Foundation for Science and Technology (FCT) grant POCTI/BSE/47585/2002. J. Moreira-Silva was supported by a PhD fellowship (SFRH/BD/16760/2004). The authors thank Dr. Marta Ferreira and Hugo Santos. References Alam, M., Frankel, T.L., 2006. Gill ATPase activities of silver perch, Bidyanus bidyanus (Mitchell), and golden perch, Macquaria ambigua (Richardson): effects of environmental salt and ammonia. Aquaculture 251, 118–133. Altschul, S.F., Madden, T.L., Schaffer, A.A., Zhang, J., Zhang, Z., Miller, W., Lipman, D.J., 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25, 3389–3402. APHA, 1989. Standard Methods for the Examination of Water and Wastewater, 17th Edition. American Public Health Association, Washington DC. Baker, D.W., Matey, V., Huynh, K.T., Wilson, J.M., Morgan, J.D., Brauner, C.J., 2009. Complete intracellular pH protection during extracellular pH depression is associated with hypercarbia tolerance in white sturgeon, Acipenser transmontanus. Am. J. Physiol Regul. Integr. Comp Physiol 296, R1868–R1880. Bakouh, N., Benjelloun, F., Hulin, P., Brouillard, F., Edelman, A., Cherif-Zahar, B., Planelles, G., 2004. NH3 is involved in the NH+ 4 transport induced by the functional expression of the human Rh C glycoprotein. J. Biol. Chem. 279, 15975–15983. Bergmeyer, H.U., Beutler, H.O., 1983. Ammonia. In: Bergmeyer, H.U. (Ed.), Methods of Enzymatic Analysis. Verlag Chemie, Weinheim, pp. 454–461. Brown, D., Breton, B., 1996. Mitochondria-rich, proton-secreting epithelial cells. J. Exp. Biol. 199, 2345–2358. Chew, S.F., Jin, Y., Ip, Y.K., 2001. The loach Misgurnus anguillicaudatus reduces amino acid catabolism and accumulates alanine and glutamine during aerial exposure. Physiol. Biochem. Zool. 74, 226–237. Chomczynski, P., Sacchi, N., 1987. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem. 162, 156–159. Crockett, E.L., Hazel, J.R., 1995. Chloesterol levels explain inverse compensation of membrane order in brush border but not homeoviscous adaptation in basolateral membranes from the intestinal epithelia of rainbow trout. J. Exp. Biol. 198, 1105–1113. Daveloose, D., Vezin, H., Basse, F., Viret, J., 1993. Fluidity of chicken ventricular plasma membranes during development in-ovo and after birth: spin labelling and fluorescence studies. J. Mol. Cell. Cardiol. 25, 1439–1444. Emerson, K., Russo, R.C., Lund, R.E., Thurston, R.V., 1975. Aqueous ammonia equilibrium calculations: effect of pH and temperature. J. Fish. Res. Bd. Can. 32, 2379–2383. Epstein, F.H., Silva, P., Kormanik, G., 1980. Role of Na–K-ATPase in chloride cell function. Am. J. Physiol. 238, R246–R250. Evans, D.H., Piermarini, P.M., Choe, K.P., 2005. The multifunctional fish gill: dominant site of gas exchange, osmoregulation, acid–base regulation, and excretion of nitrogenous waste. Physiol. Rev. 85, 97–177. Freeman, W.M., Walker, S.J., Vrana, K.E., 1999. Quantitative RT-PCR: pitfalls and potential. Biotechniques 26, 112–115. Frick, N.T., Wright, P.A., 2002. Nitrogen metabolism and excretion in the mangrove killifish Rivulus marmoratus II. Significant ammonia volatilization in a teleost during air-exposure. J. Exp. Biol. 205, 91–100. Fromm, P.O., 1963. Studies on renal and extrarenal excretion in a freshwater teleost, Salmo gairdneri. Comp. Biochem. Physiol. 10, 121–128. Gonçalves, A.F., Castro, L.F., Pereira-Wilson, C., Coimbra, J., Wilson, J.M., 2007. Is there a compromise between nutrient uptake and gas exchange in the gut of Misgurnus

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