Regulation of the energy sensor AMP-activated protein kinase in the kidney by di

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The Austin Research Institute and the Department of Nephrology, Austin Health, Department of Medicine, University of Melbourne, St. Vincent's Institute of Medical Research, St. Vincent's Hospital, and Commonwealth Scientific and Industrial Research Organization, Victoria, Australia2 d# Z- I, s" N( k% f/ V0 p

ABSTRACT! L4 W0 J: |( q! T
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The AMP-activated protein kinase (AMPK) is a key controller of cellular energy metabolism. We studied its expression and regulation by salt handling in the kidney. Immunoprecipitation and Western blots of protein lysates from whole rat kidney using subunit-specific antibodies showed that the 1-catalytic subunit is expressed in the kidney, associated with the 2- and either 1- or 2-subunits. Activated AMPK, detected by immunohistochemical staining for phospho-Thr172 AMPK (pThr172), was expressed on the apical surface of the cortical thick ascending limb of the loop of Henle, including the macula densa, and some parts of the distal convoluted tubule. Activated AMPK was also expressed on the basolateral surface of the cortical and medullary collecting ducts as well as some portions of the distal convoluted tubules. AMPK activity was increased by 25% in animals receiving a high-salt diet, and this was confirmed by Western blotting for pThr172. Low-salt diets were associated with reduced levels of the -subunit of AMPK, which was highly phosphorylated on Thr172. Surprisingly, both low- and high-salt media transiently activated AMPK in the macula densa cell line MMDD1, an effect due to changes in osmolality, rather than Na  or Cl– concentration. This study, therefore, demonstrates regulation of AMPK by both a high- and a low-salt intake in vivo and suggests a role for the kinase in the response to changes in osmolality within the kidney.
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sodium excretion; macula densa; tubuloglomerular feedback# y" B# i2 N: w7 B  k, L

RECENTLY, the AMP-activated protein kinase (AMPK) has been recognized as a key regulator of cellular energy utilization within cells and an important intermediate in control of appetite and fat metabolism (reviewed in Ref. 30). Although the actions of AMPK were initially defined as regulation of fatty acid and cholesterol synthesis pathways (9), it is now known that AMPK regulates many metabolic pathways, including glucose transport (1), fatty acid -oxidation (30), and glycolysis (28), as well as acting in the signaling pathways of leptin (36), isoproterenol (56), adiponectin (54), and resistin (3) to coordinate energy metabolism. Absence of AMPK signaling in the hypothalamus stimulates the sympathetic nervous system (51), and it is an important target for the antidiabetic drugs metformin and rosiglitazone (18). AMPK also regulates both endothelial nitric oxide synthase (eNOS) and neuronal NOS (nNOS) through phosphorylation (11, 12).

Structurally, the mammalian enzyme comprises a heterotrimer of one or other of the catalytic subunits 1 or 2, together with 1 or 2 and 1, 2, or 3 noncatalytic subunits (37). It is regulated through phosphorylation on Thr172 by an upstream kinase, recently identified as the tumor suppressor LKB-1, and intracellular levels of AMP (24, 46, 52). AMP binds to the CBS domains in the -subunit, whereas the -subunit possesses a glycogen-binding domain (27, 41).  _+ \7 ~1 h; n) q

Because the major energy-requiring process in the kidney is sodium transport (31), we hypothesized that AMPK is likely to be activated by salt loading and to phosphorylate molecules important in the response to changes in dietary salt. In this study, therefore, we determined the distribution of the active form of AMPK in normal rat kidney and demonstrated that salt intake regulates AMPK.

MATERIALS AND METHODS
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Animal models. The Animal Ethics Committee of Austin Health approved all animal procedures. Male Sprague-Dawley rats weighing 200–250 g, purchased from the Western Australian Research Facility, were fed a diet of high salt (8.95% NaCl; sodium toxicity diet, ICN, Costa Mesa, CA), low salt (0% NaCl; sodium-deficient diet, ICN), or standard rat chow (0.5% NaCl; Ridley Agriproducts, Pakenham, Victoria, Australia) for 2 wk. At the end of the experiment, the animals were anesthetized by an intraperitoneal injection of pentobarbital sodium (50 mg/kg). A midline incision was made, and the left kidney was freeze-clamped in situ to provide protein for biochemical analysis. For immunohistochemistry, kidneys were biopsied and immediately fixed by immersion in 4% paraformaldehyde. To confirm the immunohistochemical findings, perfusion-fixed kidneys from Sprague-Dawley rats, the generous gift of Dr. Wilkinson-Berka, Department of Anatomy, University of Melbourne, were also used.

Antibodies. As previously described (10, 19, 47), polyclonal antibodies were raised to nonconserved regions of the AMPK isoforms 1 (373–390, CARHTLDELNPQKSKHQG) and 2 (490–516, CSAAGLHRPRSSVDSSTAENHSLSG). Anti-AMPK 1-antibodies were raised against the peptide CSSPPGPYHQEPYISKPE-NH2 (181–197); this antibody is now recognized to detect both 1 and 2, which can be distinguished by their apparent molecular mass. AMPK 1-antibodies were raised against the peptide CQALVLTGGEKKP-COOH (319–331), whereas AMPK 2-antibodies were raised against the peptide CLTPAGAKQKETETE-COOH (555–569) and 3 against the peptide AESTGLEATFPKTTPLC (59–75). All antibodies were antigen affinity purified and tested for reactivity by ELISA with the immunizing peptide and for specificity by Western blotting using purified AMPK. A further antibody was raised against the activated 1-subunit with the phosphopeptide AMPK 1 KDGEFLRpTSCGSPNY (165–179). The antibody was purified from sera using the corresponding phosphopeptide affinity column after preclearing with a dephosphospho affinity column specific for the dephospho version of the immunising peptide (47). Affinity-purified rabbit polyclonal antibodies were produced against the phosphorylation site phospho-nNOS-Ser1412 as previously described (11). A monoclonal mouse antibody against nNOS (N31020 [GenBank] ) was purchased from BD Transduction Laboratories (Lexington, KY).! j/ [7 x* v. k9 p( }) ?
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Immunohistochemistry. Tissue was immersion fixed in 4% paraformaldehyde (BDH, Poole, UK), processed, and embedded in paraffin, as previously described (29). Briefly, 4-μm-thick paraffin sections were incubated overnight with the anti-pThr172 phosphopeptide antibody (5 μg/ml) at 4°C. Rabbit IgG was detected using the rabbit peroxidase-antiperoxidase (PAP) technique, comprising a goat anti-rabbit Ig (Dako, Carpinteria, CA) diluted at 1/100, followed by rabbit PAP (Dako) diluted 1/100. Peroxidase labeling was revealed using the liquid DAB substrate-chromagen system (Dako). Sections were counterstained with hematoxylin.0 K; i( t! n7 o4 t+ o7 \$ m

Localization of staining using tubular markers. To determine the location of tubular staining of activated AMPK, double staining confocal microscopy was performed with FITC-conjugated tubular markers and the AMPK anti-p-Thr172 phosphopeptide antibody (5 μg/ml). Paraffin sections were dewaxed and then quenched with 150 mM glycine for 20 min. Sections were permeabilized with 0.3% Triton X-100/0.025% CHAPS/PBS and nonspecific staining was blocked by the addition of 5% BSA. Staining was performed overnight and rabbit IgG was detected using anti-rabbit antibody conjugated to Alexa 594. To study the expression of activated AMPK in distal tubules and collecting ducts (CD), FITC-conjugated peanut agglutinin (Sigma, St. Louis, MO) was incubated overnight (final concentration 5 μg/ml). To stain proximal tubules, FITC-conjugated Phaseolus vulgaris erythroagglutinin (Sigma; PHA-E) was used (final concentration 5 μg/ml). Sections were then studied by confocal laser-scanning microscopy (Leica Microsystems, Heidelberg, Germany).

To examine activated AMPK expression in the thick ascending limb of the loop of Henle (TAL), sequential sections were stained by immunohistochemistry using a goat polyclonal antibody against Tamm-Horsfall protein (ICN; final concentration) (39). To determine the expression of activated AMPK in the macula densa (MD), sequential sections were stained by immunohistochemistry, using a monoclonal antibody against nNOS (N31020 [GenBank] ). For this purpose, kidneys were immersion fixed in zinc fixative.
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Immunoprecipitation and immunoblotting. Lysates of rat kidney, heart, liver, and skeletal muscle were prepared by homogenization into 4 ml of homogenization buffer (20 mM Tris﹞HCl, pH 8.0, 2 mM MgCl2, 200 mM sucrose, 1% Triton X-100, 50 mM NaF, and protease inhibitors 2 mM PMSF, 1 μM leupeptin, 0.2 μM aprotonin, 1 mM benzamidine, and 1 mM AEBSF) and centrifugation (15,000 g, 5 min). The resultant supernatant was further clarified by high-speed centrifugation (75,000 g, 30 min). The supernatant protein concentration was determined as described by Bradford using a commercial protein assay (Bio-Rad, Munich, Germany).
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Immunoprecipitations of AMPK were performed by incubating 20 mg of kidney lysate or 1 mg of cell lysate for 1 h with 5 μg antibody directed against the 1-, 2-, 1-, or 2-subunits of AMPK or an unrelated antigen. Protein A-Sepharose was used to immunoprecipitate immune complexes. Samples were separated using 12% SDS-polyacrylamide gel electrophoresis, transferred to PVDF membranes (Immobilon-P, Millipore, Bedford, MA), and immunoblotted with AMPK-specific antibodies. Partially purified AMPK from rat liver was used as a positive control. Immunoreactive proteins were detected by enhanced chemiluminescence (ECL) with protein A-horseradish peroxidase and the SuperSignal chemiluminescent system (Pierce). If the membrane was to be reprobed with another primary antibody, bound antibody was stripped with 1x Re-blot Stripping Solution (Chemicon, Temecula, CA) for 15 min. Quantitation of results was by densitometry (Scion Image for Windows, Scion, Frederick, MD). nNOS was immunoprecipitated with 2' 5' ADP Sepharose4B (Amersham Pharmacia Biotech AB, Uppsala, Sweden) from MMDD1 lysates, as previously described (11, 20).' Z) k/ b; [  ?/ O* \. T$ O
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AMPK activity assay. AMPK activity was measured by the extent to which the sample phosphorylated the AMPK consensus sequence of the ADR-1 peptide (LKKLTLRPSFSAQ-amide), as previously described (15, 35). Briefly, AMPK was immunoprecipitated and incubated at 30°C with 100 μM ADR-1 peptide in reaction buffer (50 mM HEPES, pH 7.5, 10 mM MgCl2, 5% glycerol, 1 mM DTT, 0.05% Triton X-100) with 250 μM [-32P]ATP (500 cpm/pmol) for 8 min. After the reaction, 25 μl of reaction mixture were spotted onto the phosphocellulose P81 paper and extensively washed with phosphoric acid. The radioactivity on the filter paper was measured by scintillation counting, and AMPK activity was expressed as picomoles of 32P incorporation into the peptide per minute per microgram of protein.7 d" \# x: b  F, O

Cell culture. The murine MD cell line MMDD1 (55) was very kindly provided by Dr. J. Schnermann (National Institutes of Health, Bethesda, MD) and grown in D-MEM media supplemented with 10% FCS until confluence. Quiescent MMDD1 cells were then exposed to media of low-, normal-, or high-salt content with or without osmolality controls (Table 1) as previously described (13). Cells were incubated in lysis buffer for 5 min on ice and then centrifuged at 18,000 g for 5 min at 4°C and the resulting pellets were discarded. Protein concentration was then determined by Bradford assay using a commercial protein assay solution (Bio-Rad), and the lysates were used for immunopreciptiation.

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Statistics. Statistics were performed using Instat Version 3.05 (GraphPad Software, San Diego, CA). Data are presented as means ± SD. Unless stated, data were analyzed by ANOVA using Bonferroni's test of multiple comparisons. P values 9 S8 g6 |0 q  M/ S

RESULTS) n/ {# P& h6 c  P& e' D2 h

AMPK subunit expression in the kidney. The expression of AMPK subunits in the kidney was determined by immunoprecipitation experiments using antibodies against the 1-, 2-, 1-, and 2-subunits followed by Western blots using antibodies against all the known AMPK subunit isoforms (Fig. 1). The pattern of subunit expression in the kidney was found to be different to that in heart, liver, and skeletal muscle. In the kidney, the predominant isoform of the catalytic subunit was 1 (Fig. 1, A–D), whereas the 2-subunit was found to be expressed either as a faint band (Fig. 1B) or to be undetectable (Fig. 1, C and D). The 30-kDa 2-subunit was detected following immunoprecipitation with antibodies against 1-, 2-, 1-, and 2-subunits (Fig. 1). In contrast to the other tissues examined, the 36-kDa 1-subunit was generally not detected in kidney lysates (Fig. 1), although some other blots did demonstrate a faint 36-kDa 1 band consistent with a low level of expression (data not shown). The 1- and 2-subunits were found to coprecipitate with both -subunits (Fig. 1, A and B). This was confirmed by Western blots following immunoprecipitation with antibodies against the 1- and 2-subunits (Fig. 1, C and D). The skeletal muscle-specific 3-subunit was not detected following immunoprecipitation with antibodies against the 1-, 2-, and 3-subunits (data not shown). The 2-subunit has been described to exist as a long (63 kDa) and a short form (37 kDa), with the long form appearing predominant in the kidney at a mRNA level (14, 34). Surprisingly, we were unable to detect the 63-kDa long form of 2 by Western blot. However, a 37-kDa species consistent with the short form of 2 was detected in the kidney and other rat tissues. In fact, a double band close to 37 kDa was observed in the 2 blots. Whether this represents some form of posttranslational modification or an artifact caused by proteolysis is unclear.' U: z* m+ ?! F7 T; k
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Distribution of activated AMPK in the kidney. To determine the location of activated AMPK, a polyclonal antibody against AMPK phosphorylated at the Thr172 site (pThr172) in the activation loop of the 1-subunits and 2-subunits was used. Immunohistochemistry revealed labeling restricted to the apical or basolateral surface of a population of tubules (Fig. 2). In the cortex, there was apical staining of approximately half of the TAL (Fig. 2, A and F), demonstrated by staining of sequential sections with anti-Tamm-Horsfall antibody (not shown). The MD, identified by staining of sequential sections with nNOS antibody, also expressed activated AMPK (Fig. 2, E and G). Cells of the proximal convoluted tubule, identified by immunofluorescence microscopy using FITC-conjugated PHA, were negative (Fig. 3C). There was weak apical staining in some cells of the distal convoluted tubule (Fig. 3B), identified using the FITC-conjugated lectin PNA and immunofluorescence microscopy. CD in both the cortex and medulla showed basolateral staining (Fig. 2, B and D), although this was variable and sometimes weak. These tubules were identified using FITC-conjugated PNA (Fig. 3, A, D, and E). As a control for the specificity of staining, the antibody was preincubated with the immunizing phosphopeptide or the dephosphopeptide, both coupled to SulfoLink resin. Preincubation with the specific phosphopeptide removed all staining, whereas the dephoshopeptide had no effect (data not shown). Immunohistochemical staining of perfusion-fixed sections was identical to the immersion-fixed sections (data not shown).+ }1 L- D7 t5 _3 v( o
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Regulation of AMPK activity by salt in vivo. Because tubular salt reabsorption is a major determinant of energy consumption in the kidney, we investigated the effect of dietary salt concentration on AMPK activity. Rats were fed a low-, normal-, or high-salt diet and AMPK activity was determined by activity assay and detection of p-Thr172. There was a 25% increase in AMPK activity in rats fed a high-salt diet (P

Regulation of AMPK activity by salt and osmolality in vitro. Incubation of cells in media of different composition has been used as a surrogate for changes in tubular fluid (13, 55). To determine the effect of salt concentrations on AMPK activity, therefore, the MD cell line MMDD1 (55) was incubated with low-, normal-, or high-salt media (Table 1) for periods up to 24 h. The AMPK subunit distribution of MMDD1 was first determined by immunoprecipitation and Western blot analysis and found to be similar but not identical to that of whole kidney. The major subunits present were 1, 1, 1, and 2 (Fig. 6), but there was no detectable 2, as in the whole kidney. The cells were incubated in low-, normal-, or high-salt media to determine the effect of salt on AMPK activation. There was a rapid (5 min) increase in AMPK activity in the low-salt medium, which peaked at 15 min (Fig. 7B), was maintained to 3 h, and returned to baseline by 24 h (Fig. 7D). This was confirmed by changes in p-Thr172, although there was a continued slight increase at 24 h when AMPK activity had returned to baseline (Fig. 7, A and C). The high-salt medium also showed an increase in AMPK activity, but not until 30 min (Fig. 7B) and had returned to baseline by 3 h (Fig. 7D). This was confirmed by increases in p-Thr172, with continued slight elevation at 3 h when AMPK activity had returned to baseline (Fig. 7, A and C). In the high-salt group, there was an apparent dissociation between p-Thr172 and AMPK activity at 24 h. This was inconsistent and associated with evidence of cellular toxicity.+ ?* M* y* F, i
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To determine whether the effect of salt concentration was mediated by changes in osmolality, osmotic controls were prepared using mannitol to restore the osmolality of the low-salt medium while the reduced salt concentration was maintained (Fig. 8, A and B) or to increase the osmolality of normal salt-containing medium to that of the high-salt medium (Fig. 9, A and B). These controls showed that restoration of normal osmolality prevented activation of AMPK associated with low-salt media, as determined by the presence of p-Thr172 (Fig. 8A) and AMPK activity assays (Fig. 8B). Moreover, increasing osmolality in the normal-salt medium to the level of the high-salt medium produced an equivalent increase in p-Thr172 and AMPK activity (Fig. 9, A and B). These data demonstrate that AMPK activation in MMDD1 cells is mediated by changes in osmolality and not salt concentration per se.

DISCUSSION  \6 z- s2 J1 o# w

To determine the AMPK subunits present in the kidney, lysates from kidney, heart, liver, and skeletal muscle were immunoprecipitated with antibodies against AMPK subunits and then immunoblotted. The kidney was unique when compared with liver, heart, and skeletal muscle, in the virtual absence of the 2-catalytic subunit and the 1-regulatory subunit. We next attempted to determine whether there was any region of the kidney that expressed activated AMPK. Immunohistochemical and immunofluorescence staining of normal rat kidneys with an antibody against pThr172 showed that the cortical TAL and MD strongly expressed activated AMPK on the apical surface. There was also some staining on the basolateral surface of the CD, although this was not present in all kidneys.
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Because we proposed a role for AMPK in regulation of salt reabsorption, rats were then placed on high-, normal-, or low-salt diets. AMPK activity was increased in the kidney by salt loading but not by salt restriction. More detailed study, however, showed that there was significantly reduced total 1-subunit expression in kidneys from rats receiving a low-salt diet. The AMPK present was highly phosphorylated on Thr172, although this did not achieve statistical significance.
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The unexpected ability of both high- and low-salt diets to modify AMPK activation was supported by studies performed in the MD cell line MMDD1. MMDD1 was used in these studies because the effect of salt concentration and tubular flows has been most well studied in the MD. In that location, the feedback signal for tubuloglomerular feedback (TGF) appears to be tubular Cl– concentration (43, 44). In MMDD1 cells, however, osmolality, whether high or low, was the stimulus for AMPK activation. Although this might argue against a role for AMPK, there is some evidence that osmolality can also mediate TGF (6, 7, 21).
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Although the data from the cell culture studies are supportive of the in vivo data, there are several caveats that must be made, beyond the usual reservations with cell culture as a model for events occurring in the whole animal. First, the time courses over which the observations were made clearly differ, with animals exposed to high-salt diets for 2 wk compared with only 24 h for cells in culture. In vivo, moreover, the tubules exposed to high salt might vary over time, as there was only about half of the TAL that showed detectable staining for p-Thr172 at any one time. Interestingly, the pattern of staining for p-Thr172 was the same in high-, normal-, and low-salt groups. Second, exposure of cultured cells to hypertonic or hypotonic stimuli is likely leading to changes in cell shape, which could stimulate AMPK. There is no evidence that tubular cells in vivo undergo similar changes in cell volume related to the more gradual changes in osmolality of the tubular fluid that they are likely to encounter.1 n. o7 t; P6 y( c1 B1 q
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High osmolality is a well-known activator of AMPK (25), but this is the first report of low osmolality causing AMPK activation. The ability of the two divergent stimuli, low and high osmolality, to transiently activate AMPK suggests the likelihood of different renal AMPK substrates, but their identity is currently unknown. Two other kinases are also regulated by dietary salt intake. Both erk1/2 and p38 MAPK are activated by increased dietary salt intake (57), but low-salt diets also activate p38 MAPK (13). The apparently paradoxical observation with AMPK, therefore, has a counterpart in regulation of p38 MAPK.
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The role of AMPK in the kidney is unknown, but the distribution of the activated kinase suggests several potential substrates. nNOS and eNOS are known substrates for AMPK (11, 12). nNOS is restricted in its immunohistochemical distribution to the MD (2, 50), although its mRNA is more widely distributed, especially in the medulla (53). Immunohistochemical staining for pThr172 AMPK demonstrated areas of codistributed in this study with nNOS in the MD. This may be relevant because AMPK is one of only two kinases known to phosphorylate nNOS (11, 26). NO produced by nNOS is important in resetting TGF and is probably important in excretion of a salt load (16, 42). If AMPK were to phosphorylate nNOS in vivo, then it could contribute to control of NO release from these cells, together with other stimuli such as intracellular Ca, which is reported to increase during salt loading (4, 40). Another potential role is in regulation of ion channels or sodium cotransporters, and preliminary data from laboratories including our own have suggested that both ENaC (22) and the Na-K-2Cl (NKCC) cotransporter could be regulated by AMPK (17).
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If AMPK has a role in regulation of MD function, it would provide another level of control of TGF by adenine nucleotides such as ATP and AMP. Much interest has centered recently on the regulation of TGF by ATP and the adenine nucleotide derivative adenosine. Adenosine receptor (A1AR) knockout mice demonstrate a complete absence of TGF responses to saturating increases in MD NaCl (8, 48) and adenosine receptor antagonists also inhibit TGF (49). The source of adenosine in the region of the MD is unclear; initially, it was considered that adenosine was generated in MD cells by dephosphorylation of ATP when a rise in transcellular NaCl transport increased cellular energy demand (38). Recent data suggest that cells of the MD release ATP from their basolateral surface in response to high-salt loads (5, 32). Extracellular ATP might then be converted to adenosine by ecto-5'-nucleotidase. AMPK is the most important of a family of kinases and other molecules that are regulated, in contrast, by changes in the intracellular level of the adenine nucleotides ATP and AMP (30, 33, 45).

If ATP is lost through a specific release mechanism on the basolateral surface of cells when tubular NaCl and flows are high, then intracellular levels may fall, so disturbing the AMP/ATP ratio. In addition, salt reabsorption is the major energy-consuming process in the kidney (31), generally through the action of a basolateral Na-K-ATPase. In the MD, there is very little Na-K-ATPase and most of the sodium extrusion from the cell appears to be due to an apical H-K-ATPase that also transports Na (reviewed in Ref. 4). The net effect, however, might be similar in that high-salt loads would lead to greater activity of the Na-H/K-ATPase and greater production of AMP at the expense of ATP. AMPK activation, therefore, could occur when tubular flows are high through release of ATP into the extracellular environment and generation of AMP by the apical Na-H/K-ATPase. This proposition would be strengthened by direct measurements of AMP levels within cells of the MD during salt loading.
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Hallows et al. (23) previously suggested that AMPK could link the metabolic state of a cell and its handling of ion transport. This study supports that prediction, showing that AMPK is activated in the kidney by salt intake. Activation of AMPK in cells of the MD also suggests a potential role for intracellular adenine nucleotides in control of TGF. This complements emerging evidence for the role of extracellular ATP and adenosine in TGF and is an exciting possibility for future study.! I4 e. g0 n/ |, ^9 o

GRANTS8 H: K) z3 _6 N) L

This work was supported by National Health and Medical Research Council of Australia (NHMRC) grants to D. A. Power and B. E. Kemp. S. Fraser is a Dora Lush Postgraduate Scholar of the NHMRC, P. Mount is a Medical Postgraduate Scholar of the NHMRC, V. Levidiotis is a Clinical Research Fellow of the NHMRC, D. Stapleton is a R. D. Wright Fellow of the NHMRC, and B. Kemp is an ARC Federation Fellow. This work has been reported in abstract form: (Fraser SA, Mount PF, Stapleton D, Kemp BE, and Power DA. J Am Soc Nephrol 13: 490A, 2002)./ j  B* \) X5 T# y8 s
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FOOTNOTES

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.5 R! j+ i: d+ t
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REFERENCES
* G. P& z' T3 V, L' J  j
Andersson U, Filipsson K, Abbott CR, Woods A, Smith K, Bloom SR, Carling D, and Small CJ. AMP-activated protein kinase plays a role in the control of food intake. J Biol Chem 279: 12005–12008, 2004.

Bachmann S, Bosse HM, and Mundel P. Topography of nitric oxide synthesis by localizing constitutive NO synthases in mammalian kidney. Am J Physiol Renal Fluid Electrolyte Physiol 268: F885–F898, 1995.

Banerjee RR, Rangwala SM, Shapiro JS, Rich AS, Rhoades B, Qi Y, Wang J, Rajala MW, Pocai A, Scherer PE, Steppan CM, Ahima RS, Obici S, Rossetti L, and Lazar MA. Regulation of fasted blood glucose by resistin. Science 303: 1195–1198, 2004.  B1 ?" ]5 W5 M
0 R9 |; K0 Z7 \2 }- N" e
Bell PD, Lapointe JY, and Peti-Peterdi J. Macula densa cell signaling. Annu Rev Physiol 65: 481–500, 2003.. a  G, B6 ^: t- L& W* h

Bell PD, Lapointe JY, Sabirov R, Hayashi S, Peti-Peterdi J, Manabe K, Kovacs G, and Okada Y. Macula densa cell signaling involves ATP release through a maxi anion channel. Proc Natl Acad Sci USA 100: 4322–4327, 2003.: N/ P0 s% M9 q# [- l5 L% O
  }) J( X; s* q) K$ A
Bell PD, McLean CB, and Navar LG. Dissociation of tubuloglomerular feedback responses from distal tubular chloride concentration in the rat. Am J Physiol Renal Fluid Electrolyte Physiol 240: F111–F119, 1981.
% m5 p' |2 R* e# _2 l
Bell PD, Navar LG, Ploth DW, and McLean CB. Tubuloglomerular feedback responses during perfusion with nonelectrolyte solutions in the rat. Kidney Int 18: 460–471, 1980.

Brown R, Ollerstam A, Johansson B, Skott O, Gebre-Medhin S, Fredholm B, and Persson AE. Abolished tubuloglomerular feedback and increased plasma renin in adenosine A1 receptor-deficient mice. Am J Physiol Regul Integr Comp Physiol 281: R1362–R1367, 2001.
# v6 f/ n+ g, g2 ^# @
Carling D, Zammit VA, and Hardie DG. A common bicyclic protein kinase cascade inactivates the regulatory enzymes of fatty acid and cholesterol biosynthesis. FEBS Lett 223: 217–222, 1987.
5 C7 e+ L0 m' S6 x1 r7 ?4 s, U# m
Chen Z, Heierhorst J, Mann RJ, Mitchelhill KI, Michell BJ, Witters LA, Lynch GS, Kemp BE, and Stapleton D. Expression of the AMP-activated protein kinase 1 and 2 subunits in skeletal muscle. FEBS Lett 460: 343–348, 1999., n- ]6 M1 q  O! ?: D( f
3 m  x+ M$ X* _. N6 z4 y
Chen ZP, McConell GK, Michell BJ, Snow RJ, Canny BJ, and Kemp BE. AMPK signaling in contracting human skeletal muscle: acetyl-CoA carboxylase and NO synthase phosphorylation. Am J Physiol Endocrinol Metab 279: E1202–E1206, 2000.

Chen ZP, Mitchelhill KI, Michell BJ, Stapleton D, Rodriguez-Crespo I, Witters LA, Power DA, Ortiz de Montellano PR, and Kemp BE. AMP-activated protein kinase phosphorylation of endothelial NO synthase. FEBS Lett 443: 285–289, 1999." n( d# ]) C3 Q4 M

Cheng HF, Wang JL, Zhang MZ, McKanna JA, and Harris RC. Role of p38 in the regulation of renal cortical cyclooxygenase-2 expression by extracellular chloride. J Clin Invest 106: 681–688, 2000.' r. {3 w! x/ `7 Z: u( K$ L

Cheung PC, Salt IP, Davies SP, Hardie DG, and Carling D. Characterization of AMP-activated protein kinase -subunit isoforms and their role in AMP binding. Biochem J 346: 659–669, 2000.
' D) U. j5 Q+ r4 \+ N- s
Davies SP, Carling D, and Hardie DG. Tissue distribution of the AMP-activated protein kinase, and lack of activation by cyclic-AMP-dependent protein kinase, studied using a specific and sensitive peptide assay. Eur J Biochem 186: 123–128, 1989.% d! S3 u# e0 T7 T" r" U

Deng A, Hammes JS, and Thomson SC. Hemodynamics of early tubuloglomerular feedback resetting during reduced proximal reabsorption. Kidney Int 62: 2136–2143, 2002.2 Y, V& X" |0 B( m' O

Fraser SA, Mount PF, Hill R, Stapleton D, Kemp BE, and Power DA. Inhibition of the Na-K-2Cl cotransporter by novel interaction with the metabolic sensor AMP-activated protein kinase. J Am Soc Nephrol 14: 545A, 2003.

Fryer LG, Parbu-Patel A, and Carling D. The anti-diabetic drugs rosiglitazone and metformin stimulate AMP-activated protein kinase through distinct signaling pathways. J Biol Chem 277: 25226–25232, 2002.# ~1 S$ E7 C- |0 u
3 ~7 G8 W3 ]* y
Gao G, Fernandez CS, Stapleton D, Auster AS, Widmer J, Dyck JR, Kemp BE, and Witters LA. Noncatalytic - and -subunit isoforms of the 5'-AMP-activated protein kinase. J Biol Chem 271: 8675–8681, 1996.9 O7 m# Q% U( `" u

Gerber NC and Ortiz de Montellano PR. Neuronal nitric oxide synthase. J Biol Chem 270: 17791–17796, 1995.0 L& g& `+ ]8 D% f# v
7 u! t3 u% }) J
Goldwasser P and Fotino S. Acute renal failure following massive mannitol infusion. Appropriate response of tubuloglomerular feedback Arch Intern Med 144: 2214–2216, 1984.
! t! ~) b2 g0 k  r3 C7 J  v3 ^
Hallows KR, Grieser HJ, Bruns JR, Carattino MD, Kleyman TR, Johnson JP, and Edinger RS. AMP-activated kinase modulates epithelial Na  channel in oocytes and polarized mouse CCD cells. J Am Soc Nephrol 14: 81A, 2003.

Hallows KR, Raghuram V, Kemp BE, Witters LA, and Foskett JK. Inhibition of cystic fibrosis transmembrane conductance regulator by novel interaction with the metabolic sensor AMP-activated protein kinase. J Clin Invest 105: 1711–1721, 2000.

Hawley SA, Boudeau J, Reid JL, Mustard KJ, Udd L, Makela TP, Alessi DR, and Hardie DG. Complexes between the LKB1 tumor suppressor, STRAD/ and MO25/ are upstream kinases in the AMP-activated protein kinase cascade. J Biol Chem 2: 28, 2003.

Hayashi T, Hirshman MF, Fujii N, Habinowski SA, Witters LA, and Goodyear LJ. Metabolic stress and altered glucose transport: activation of AMP-activated protein kinase as a unifying coupling mechanism. Diabetes 49: 527–531, 2000.
5 m) @4 X$ {1 ^$ ^! i- j3 H
Hayashi Y, Nishio M, Naito Y, Yokokura H, Nimura Y, Hidaka H, and Watanabe Y. Regulation of neuronal nitric oxide synthase by calmodulin kinases. J Biol Chem 274: 20597–20602, 1999.
% B, I" r/ P4 L/ `
Hudson ER, Pan DA, James J, Lucocq JM, Hawley SA, Green KA, Baba O, Terashima T, and Hardie DG. A novel domain in AMP-activated protein kinase causes glycogen storage bodies similar to those seen in hereditary cardiac arrhythmias. Curr Biol 13: 861–866, 2003.* l$ Z) I! ?, s- m9 Y$ e
. u3 X% d3 o+ Q7 K
Hue L, Beauloye C, Marsin AS, Bertrand L, Horman S, and Rider MH. Insulin and ischemia stimulate glycolysis by acting on the same targets through different and opposing signaling pathways. J Mol Cell Cardiol 34: 1091–1097, 2002.

Kanellis J, Fraser S, Katerelos M, and Power DA. Vascular endothelial growth factor is a survival factor for renal tubular epithelial cells. Am J Physiol Renal Physiol 278: F905–F915, 2000.8 A/ F! a" y0 K3 I. f  A
3 l1 q8 c9 t2 f. R  V# a; ?
Kemp BE, Stapleton D, Campbell DJ, Chen ZP, Murthy S, Walter M, Gupta A, Adams JJ, Katsis F, van Denderen B, Jennings IG, Iseli T, Michell BJ, and Witters LA. AMP-activated protein kinase, super metabolic regulator. Biochem Soc Trans 31: 162–168, 2003.3 S- V) j- X& h2 Q

Kiil F, Aukland K, and Refsum HE. Renal sodium transport and oxygen consumption. Am J Physiol 201: 511–516, 1961.$ E  a- A  |! M: N. {; }6 U0 r
) Y% [: ?* i& i, `5 L( d9 k
Komlosi P, Peti-Peterdi J, Fuson AL, Fintha A, Rosivall L, and Bell PD. Macula densa basolateral ATP release is regulated by luminal [NaCl] and dietary salt intake. Am J Physiol Renal Physiol 286: F1054–F1058, 2004.
- @1 P  h  l+ f9 _
Lizcano JM, Goransson O, Toth R, Deak M, Morrice NA, Boudeau J, Hawley SA, Udd L, Makela TP, Hardie DG, and Alessi DR. LKB1 is a master kinase that activates 13 kinases of the AMPK subfamily, including MARK/PAR-1. EMBO J 23: 833–843, 2004.
9 U9 m5 @7 [. {+ C+ {
Mahlapuu M, Johansson C, Lindgren K, Hjalm G, Barnes BR, Krook A, Zierath JR, Andersson L, and Marklund S. Expression profiling of the -subunit isoforms of AMP-activated protein kinase suggests a major role for 3 in white skeletal muscle. Am J Physiol Endocrinol Metab 286: E194–E200, 2004.& e- ~7 }. B6 [0 I1 ~5 \; Q
; A$ S( t# M5 K. H, E( i, C: v
Michell BJ, Stapleton D, Mitchelhill KI, House CM, Katsis F, Witters LA, and Kemp BE. Isoform-specific purification and substrate specificity of the 5'-AMP-activated protein kinase. J Biol Chem 271: 28445–28450, 1996.
2 H$ D) j3 z* U9 s' r$ B9 n
Minokoshi Y, Kim YB, Peroni OD, Fryer LG, Muller C, Carling D, and Kahn BB. Leptin stimulates fatty-acid oxidation by activating AMP-activated protein kinase. Nature 415: 339–343, 2002.

Mitchelhill KI, Stapleton D, Gao G, House C, Michell B, Katsis F, Witters LA, and Kemp BE. Mammalian AMP-activated protein kinase shares structural and functional homology with the catalytic domain of yeast Snf1 protein kinase. J Biol Chem 269: 2361–2364, 1994.
3 y% {1 G; j% h4 U, }0 A$ @- U
Osswald H, Hermes HH, and Nabakowski G. Role of adenosine in signal transmission of tubuloglomerular feedback. Kidney Int 12: S136–S142, 1982.# A3 k9 T% L1 Z8 B: i

Paizis K, Kirkland G, Polihronis M, Katerelos M, Kanellis J, and Power DA. Heparin-binding epidermal growth factor-like growth factor in experimental models of membranous and minimal change nephropathy. Kidney Int 53: 1162–1171, 1998.* b$ Q- b9 o6 a
$ {+ {3 [0 ^" Q/ c# W2 V
Peti-Peterdi J and Bell PD. Cytosolic [Ca2 ] signaling pathway in macula densa cells. Am J Physiol Renal Physiol 277: F472–F476, 1999.
) u4 P7 e! ?6 \4 |
Polekhina G, Gupta A, Michell BJ, van Denderen B, Murthy S, Feil SC, Jennings IG, Campbell DJ, Witters LA, Parker MW, Kemp BE, and Stapleton D. AMPK  subunit targets metabolic stress sensing to glycogen. Curr Biol 13: 867–871, 2003.
2 @" O; _1 j1 ]3 r+ C: p# q( x: D
Schnermann J and Levine DZ. Paracrine factors in tubuloglomerular feedback: adenosine, ATP, and nitric oxide. Annu Rev Physiol 65: 501–529, 2003.
! n8 H& Z; h3 v: F+ \" J2 `3 p
Schnermann J, Ploth DW, and Hermle M. Activation of tubuloglomerular feedback by chloride transport. Pflügers Arch 362: 229–240, 1976.( H, w& L4 p9 ~7 e% Z" R

Schnermann J, Wright FS, Davis JM, von Stackelberg W, and Grill G. Regulation of superficial nephron filtration rate by tubuloglomerular feedback. Pflügers Arch 318: 147–175, 1970.! X7 |3 [% s9 W) {0 p2 c0 w

Scott JW, Hawley SA, Green KA, Anis M, Stewart G, Scullion GA, Norman DG, and Hardie DG. CBS domains form energy-sensing modules whose binding of adenosine ligands is disrupted by disease mutations. J Clin Invest 113: 274–284, 2004.

Shaw RJ, Kosmatka M, Bardeesy N, Hurley RL, Witters LA, DePinho RA, and Cantley LC. The tumor suppressor LKB1 kinase directly activates AMP-activated kinase and regulates apoptosis in response to energy stress. Proc Natl Acad Sci USA 101: 3329–3335, 2004.

Stapleton D, Mitchelhill KI, Gao G, Widmer J, Michell BJ, Teh T, House CM, Fernandez CS, Cox T, Witters LA, and Kemp BE. Mammalian AMP-activated protein kinase subfamily. J Biol Chem 271: 611–614, 1996., O" g: {0 f* P" N

Sun D, Samuelson LC, Yang T, Huang Y, Paliege A, Saunders T, Briggs J, and Schnermann J. Mediation of tubuloglomerular feedback by adenosine: evidence from mice lacking adenosine 1 receptors. Proc Natl Acad Sci USA 98: 9983–9988, 2001.( l" Y3 t2 U& d& N! q/ p8 {) C

Thomson S, Bao D, Deng A, and Vallon V. Adenosine formed by 5'-nucleotidase mediates tubuloglomerular feedback. J Clin Invest 106: 289–298, 2000.
, _+ j# s% M! r! S$ L; Z
Tojo A, Gross SS, Zhang L, Tisher CC, Schmidt HH, Wilcox CS, and Madsen KM. Immunocytochemical localization of distinct isoforms of nitric oxide synthase in the juxtaglomerular apparatus of normal rat kidney. J Am Soc Nephrol 4: 1438–1447, 1994.

Viollet B, Andreelli F, Jorgensen SB, Perrin C, Geloen A, Flamez D, Mu J, Lenzner C, Baud O, Bennoun M, Gomas E, Nicolas G, Wojtaszewski JF, Kahn A, Carling D, Schuit FC, Birnbaum MJ, Richter EA, Burcelin R, and Vaulont S. The AMP-activated protein kinase 2 catalytic subunit controls whole body insulin sensitivity. J Clin Invest 111: 91–98, 2003.& E: t4 A- ?5 f7 _2 D/ F

Woods A, Johnstone SR, Dickerson K, Leiper FC, Fryer LG, Neumann D, Schlattner U, Wallimann T, Carlson M, and Carling D. LKB1 is the upstream kinase in the AMP-activated protein kinase cascade. Curr Biol 13: 2004–2008, 2003.; y2 \5 }3 U7 A5 m* }4 F0 v
' N  P  _& w4 W& j7 i
Wu F, Park F, Cowley AW Jr, and Mattson DL. Quantification of nitric oxide synthase activity in microdissected segments of the rat kidney. Am J Physiol Renal Physiol 276: F874–F881, 1999.

Yamauchi T, Kamon J, Minokoshi Y, Ito Y, Waki H, Uchida S, Yamashita S, Noda M, Kita S, Ueki K, Eto K, Akanuma Y, Froguel P, Foufelle F, Ferre P, Carling D, Kimura S, Nagai R, Kahn BB, and Kadowaki T. Adiponectin stimulates glucose utilization and fatty-acid oxidation by activating AMP-activated protein kinase. Nat Med 8: 1288–1295, 2002.  ]1 M' G2 c- M& M' `5 _

Yang T, Park JM, Arend L, Huang Y, Topaloglu R, Pasumarthy A, Praetorius H, Spring K, Briggs JP, and Schnermann J. Low chloride stimulation of prostaglandin E2 release and cyclooxygenase-2 expression in a mouse macula densa cell line. J Biol Chem 275: 37922–37929, 2000., s: o( s! q+ r/ a
& C4 V9 p: _& ]( u  p# W
Yin W, Mu J, and Birnbaum MJ. Role of AMP-activated protein kinase in cyclic AMP-dependent lipolysis in 3T3–L1 adipocytes. J Biol Chem 278: 43074–43080, 2003.6 T( O  [3 _  g5 i
5 ~3 _8 ]% S" _* _) l- u" K
Ying WZ and Sanders PW. Dietary salt intake activates MAP kinases in the rat kidney. FASEB J 16: 1683–1684, 2002.(Scott Fraser, Peter Mount)