Metabolic Basis of Solute Transport - Brenner and Rector's The Kidney, 8th ed

Brenner and Rector's The Kidney, 8th ed.

CHAPTER 4. Metabolic Basis of Solute Transport

Bruce C. Kone

		ATP and Active Transport, 130
		Energy Consumption to Conduct Solute Transport, 131
		P-type ATPases, 132
		Vacuolar H⁺-ATPases, 134
		ATP-Binding-Cassette Transporters, 134
		Energy Production to Fuel Solute Transport, 135
		Mitochondrial ATP Production, 135
		Glycolysis, 139
		Tricarboxylic Acid Cycle, 139
		Ketone Body Metabolism, 139
		Fatty Acid Oxidation, 140
		Pentose-Phosphate Shunt, 140
		Gluconeogenesis and Its Role in Solute Transport, 140
		Coupling of Active Solute Transport and ATP Production, 141
		Whole Kidney Studies, 141
		Correlation Between Na⁺ Transport and QO₂ Along the Nephron, 141
		Coupling of Na⁺,K⁺-ATPase and Mitochondrial Oxidative Phosphorylation, 142
		ATP-sensitive Ion Channels and Coupling to Cation Transport, 144
		ATP (Purinergic) Receptors Modulating Active Na⁺ Transport, 145
		AMP-activated Protein Kinase Coupling Ion Transport and Metabolism, 145
		Functional Coupling of Glycolysis with Ion Pumps, 146
		Renal Substrate Preferences, 146
		Whole Kidney and Regional Profiles of Metabolism, 146
		Segmental Profiles of Nephron Metabolism, 146
		Solute Transport and Energy Availability During Hypoxic or Ischemic Conditions, 150
		Effects of L-Arginine Metabolism on Solute Transport and Cellular Energetics, 151
		L-Arginine Metabolism, 151
		Effects of NO on Renal Solute Transport, 151
		Effects of NO on Mitochondrial Respiration, 151
		Summary and Conclusions, 152

The normal kidney encounters extreme metabolic demands even under physiologic conditions. The reabsorption of 99% of the 180 L of glomerular ultrafiltrate each day expends considerable metabolic energy and requires commensurate levels of energy production. Accordingly, though contributing less than 1% of the total body mass, the kidney basally consumes 10% of the total body oxygen, a value surpassed only by the heart among parenchymal organs. The functional capacity of the kidney to perform active transport and biosynthetic processes is dependent on its energy supply. The energy needed to perform such work is chiefly derived from biologic oxidation (O₂-requiring) reactions that convert metabolic substrates into high-energy compounds such as ATP. These oxidative processes, in particular mitochondrial oxidative phosphorylation, account for roughly 95% of total renal ATP production, although nonoxidative ATP generation, principally glycolysis (lactate generation from glucose). generates a greater share of ATP to support active solute transport in certain nephron segments.

Through their direct or indirect influence on electrochemical gradients, membrane ionic conductances, membrane permeability, and ion pump activity, metabolic components exert considerable control over transepithelial solute transport. Given this level of importance to renal function, the relationship between renal solute transport and metabolism has been the subject of extensive study over the past century. This chapter reviews the elements that couple ATP synthesis to solute transport, the preferred metabolic substrates for these processes, and the integration of individual nephron segments in generating and consuming energy to conduct solute transport.

ATP AND ACTIVE TRANSPORT

Since membranes are generally impermeable to ions distributed across them, ion-motive pumps are used to interconvert chemical energy derived from the hydrolysis of ATP (ATP ➙ ADP+P_i) or other high-energy phosphate molecules into an electrochemical gradient to drive transport against a concentration gradient. This process is termed primary active transport ( Fig. 4-1 ). In the kidney, the principal mechanism for primary active transport is the Na⁺,K⁺-ATPase, an enzyme that serves to maintain the low concentration of Na⁺ and high concentration of K⁺ in the intracellular environment. Secondary active transport refers to transport that allows solutes to move along an electrochemical gradient, without chemical modification or direct consumption of energy (see Fig. 4-1 ). Thus, the energy stored in the steep Na⁺ gradient generated by the Na⁺,K⁺-ATPase can be used to direct Na⁺-coupled transport of sugars, amino acids, and a variety of other solutes along the nephron. H⁺-ATPases and Ca²⁺-ATPases in the plasma membranes of specific renal tubular epithelial cells can also generate ion gradients that can fuel Na⁺-independent secondary active transport. Indeed the H⁺-ATPase in the brush border membrane of the proximal tubule appears to be a significant energy-consuming process in certain species. Finally, the energy stored in the Na⁺gradient generated by the Na⁺,K⁺-ATPase can be indirectly used to drive the transport of other ions and organic molecules. As one example, peptide transport in the proximal tubule is driven by a H⁺ gradient across the brush border membrane. As another example, the Na⁺/H⁺ exchanger, a secondary active transport system located in the brush border membrane, couples the influx of Na⁺ into the cell with the efflux of H⁺ from the cell and is thus principally responsible for the existence of this H⁺ gradient (see Fig. 4-1 ). The driving force for the Na⁺/H⁺ exchanger, a transmembrane Na⁺ gradient, is in turn generated and maintained by the Na⁺,K⁺-ATPase, a primary active transport system in the basolateral membrane of these cells. In tertiary active transport, the H⁺ gradient generated by the operation of the Na⁺,K⁺-ATPase and Na⁺/H⁺ exchanger is used to drive the tertiary active transport of Cl^- across the brush border membrane via a Cl^-/HCO₃^- exchanger (see Fig. 4-1 ).



FIGURE 4-1 Active transport processes in renal epithelial cells. Models of three epithelial cells are shown to illustrate the different modes of active transport. Primary active transporters (in this example, the Na⁺,K⁺-ATPase) use the energy derived from ATP hydrolysis to power the transport of solutes across the plasma membrane against their electrochemical gradients. Secondary active transporters utilize the energy in the electrochemical gradient (in this example, Na⁺) generated by the primary active transport process to drive the influx of efflux of a coupled solute. Tertiary active transport links the transport of a solute (in this example, Cl^-) to the gradient (in this case, H⁺) created by the secondary active transport process.

ENERGY CONSUMPTION TO CONDUCT SOLUTE TRANSPORT

The intracellular ionic composition differs markedly from that of the extracellular fluid, and maintenance or restoration of this condition requires the input of energy. As previously emphasized, primary active transport processes make direct use of ATP, whereas secondary active transport makes use of the potential energy stored in transmembrane ion gradients. Various classes of ATP-driven solute pumps perform primary active transport. Many schemes exist to classify these pumps, but for simplicity, three broad classes important for renal transport are considered here ( Fig. 4-2 and Table 4-1 ). These energy-requiring pumps provide for all solute transport along the nephron either by conducting the transport themselves or by establishing electrochemical gradients that allow the solute transport by secondary or tertiary active transport processes. In general, the distribution and level of expression of specific ATPases along the nephron correlates with the demands for solute transport and the required metabolic machinery (e.g., mitochondria, enzymes for ATP synthetic pathways) to support the transport. Numerous studies have also established that the activities and level of expression of these active transporters in specific nephron segments respond to changes in overall ion balance, hormones, and autocoids.



FIGURE 4-2 Major classes of transport ATPases in the kidney. M=membrane-spanning domain; C=catalytic domain; MDR1=multi-drug resistance protein 1.

TABLE 4-1 -- Major Classes of Inorganic Ion-Translocating ATPases Active in Kidney and Their Structural and Functional Characteristics

Ion Pump	Class	Subunit Structure	Ionic Stoichiometry
Na⁺,K⁺-ATPase	P-type	αβ	3Na⁺:2K⁺:ATP
H⁺,K⁺-ATPase	P-type	αβ	H⁺:K⁺:ATP
Ca²⁺-ATPase	P-type	α	Ca²⁺: (H⁺):ATP
Cu²⁺-ATPase	P-type	α	Cu²⁺:ATP
H⁺-ATPase	V-type	F₁F₀ 12 subunits	H⁺:ATP

P-type ATPases are an evolutionarily conserved family of over 300 ATP hydrolysis-driven ion pumps that form an acylphosphate intermediate as part of the reaction mechanism and bear a seven amino acid signature motif, beginning with the aspartate to which the terminal phosphate of ATP is attached during the enzymatic cycle. These enzymes share four highly conserved protein domains ten transmembrane domains, as well as highly conserved phosphorylation and ATP-binding sites. The activity of most P-type ATPases is tightly controlled by extraregulatory domains or protein subunits. The Na⁺,K⁺-ATPase and the closely related gastric H⁺,K⁺-ATPase (H⁺,K⁺-ATPase α1 subunit) are the only members of the P-type ATPase family comprising more than one subunit. On the basis of sequence similarities, phylogenetic analyses, and substrate specificities, Axelsen and Palmgren^[1] classified this superfamily into 5 families and 11 subfamilies. The type Ia ATPases include the B subunit of the bacterial Kdp K⁺-ATPase. Type Ib ATPases transport heavy metals such as Cu²⁺, Cd²⁺, and Zn²⁺. Mutations in the ATP7A and ATP7B genes encoding Cu²⁺-ATPases are responsible for Menkes disease and Wilson's disease, respectively. The type II subfamily transports non-heavy metal cations and includes the sarcoplasmic-endoplasmic reticulum Ca²⁺-ATPases (SERCA; group IIa; three SERCA genes in humans); the secretory pathway Ca²⁺-ATPases (SPCA; two genes in humans), which transport both Ca²⁺ and Mn²⁺ in the Golgi lumen and therefore play an important role in the cytosolic and intra-Golgi Ca²⁺ and Mn²⁺ homeostasis; the plasma membrane Ca²⁺-ATPases (PMCA; group IIb, four genes in humans); and type IIc with the four isoforms of the Na⁺,K⁺-ATPase α-subunit and the gastric and “nongastric” H⁺,K⁺-ATPase a -subunits. The type III family is expressed only in plants and fungi and includes ATPases involved in the transport of Mg²⁺ and H⁺. Type IV subfamily members are exclusively expressed in eukaryotic cells and translocate phospholipids, rather than cations, from the outer and inner leaflet of membrane bilayers,^[2] and may play a role in vesicular (protein) trafficking in yeast. Finally, the recently identified type V subfamily, which is also exclusively expressed in eukaryotic cells, has been implicated in cellular Ca²⁺ homeostasis and endoplasmic reticulum function in yeast.

Vacuolar H⁺-ATPases (V-ATPases) are a family of multisubunit ATP-dependent H⁺ pumps responsible for acidification of intracellular organelles, including endosomes, lysomes, secretory vesicles, Golgi, and clathrin-coated vesicles, and of luminal or interstitial spaces. The V-ATPases are large multimeric complexes and differ from P-type ATPases in that they do not form a phosphoenzyme intermediate and are resistant to vanadate inhibition.

ATP-binding-cassette (ABC) transporters are a superfamily of proteins that couple ATP hydrolysis to the transport of a wide array of molecules across biologic membranes. Forty-nine ABC protein genes exist on human chromosomes. Eukaryotic ABC proteins were originally recognized as drug efflux pumps involved in the multidrug resistance of cancer cells, but it is now appreciated that they have multiple physiologic roles and their dysfunction can be often associated with human diseases. Collectively, they are known to transport a diverse array of drugs and toxins, conjugated organic anions comprising dietary and environmental carcinogens, pesticides, metals, metalloids, and lipid peroxidation products. They are recognized by their shared modular organization and by two sequence motifs that make up a nucleotide-binding fold. The functional protein generally contains two nuclear-binding folds and two transmembrane domains, typically consisting of six membrane-spanning α-helices.^[3] ABC proteins that confer drug resistance include (but are not limited to) P-glycoprotein (ABCB1), the multidrug resistance protein 1 (MDR1, gene symbol ABCC1), MDR2 (gene symbol ABCC2), and the breast cancer resistance protein (BCRP, gene symbol ABCG2). Proteins of the ABC transporter superfamily have been implicated in the outward transport, or “flopping,” of choline-containing phospholipids.^[2] Multidrug resistance protein (MDR) 3 (ABCB4) specifically translocates phosphatidylcholine from the inner to the outer leaflet of the bilayer when expressed in LLC-PK1 cells.^[4]MDR1 (ABCB1) protein, a broad-range xenobiotic transporter, translocates choline-containing lipids, including short-chain analogs of phosphatidylcholine, glucosylceramide, and platelet-activating factor. Studies of extracellular acidification rates in MDR1-transfected and null cells indicate that P-glycoprotein is tightly coupled to the metabolic state of the cell. The energy required for P-glycoprotein activation relative to the basal metabolic energy in glucose-deficient cells was double that of glucose-fed cells, suggesting cellular protection by P-glycoprotein even under conditions of starvation.^[5] Unlike P-glycoptorein and MDR1, ABCG2 is a half-transporter that must homodimerize to acquire transport activity. ABCG2 is found in a variety of stem cells and may protect them from exogenous and endogenous toxins. Its expression is upregulated under low-oxygen conditions, consistent with its high expression in tissues exposed to low-oxygen environments. ABCG2 interacts with heme and other porphyrins and protects cells and/or tissues from protoporphyrin accumulation under hypoxic conditions.^[6]

P-type ATPases

Na⁺,K⁺-ATPase.

The Na⁺,K⁺-ATPase is an oligomeric membrane protein that couples the hydrolysis of one ATP molecule to the translocation of three Na⁺ and two K⁺ ions against their electrochemical gradients to maintain or restore the normally high K⁺ and low Na⁺ concentrations inside mammalian cells. The Na⁺,K⁺-ATPase plays a central role in the regulation of the membrane potential, cell ion content, and cell volume. In renal tubular epithelial cells, this enzyme is distributed in the basolateral membrane, and it provides the principal driving force for net Na⁺ reabsorption and the secondary active transport of other ions and organic solutes. The specific activity of the purified Na⁺,K⁺-ATPase from renal medulla, approximately 10,000 ATP/min/enzyme molecule, is among the highest of any tissue.^{[7] [8]} Studies in the isolated perfused rat kidney suggest that the Na⁺,K⁺-ATPase directly accounts for about half of the total Na⁺ reabsorbed by the kidney.^[9]

Structurally, the minimal functional unit of the enzyme is a heterodimer of α and β subunits. The ≈100-kD α-subunit is responsible for ATP hydrolysis, cation transport, and ouabain binding, whereas the ≈40- to 60-kD (depending on the degree of glycosylation) β-subunit appears to play a role in the occlusion of K⁺, the modulation of the K⁺ and Na⁺ affinity of the enzyme, and directing the holoenzyme to the plasma membrane. The K_0.5 for ATP of the enzyme is between 5 and 400 μM,^{[7] [8]} so that the ATP concentration in most cells is saturating for the enzyme. Four α- and three β-subunit isoforms have been identified, and these exhibit different tissue distributions and produce Na⁺,K⁺-ATPase isozymes with different transport properties.^{[7] [8]}

The α1β1 enzyme is found in nearly every tissue and is the principal isozyme of the kidney. The renal expression of the other Na⁺,K⁺-ATPase isoforms has been debated.^{[10] [11] [12]} The α2 and α3 isoforms have been detected in renal cortex, medulla, and papilla and using RT-PCR,^[11] in situ hybridization,^[10] and differential [³H]ouabain titration analysis.^[12] Measurement of mRNA and protein levels together with [³H]ouabain titration data, however, indicates that the α2- and α3-isoforms constitute less than or equal to 0.1% of the α1β1 enzyme of the kidney.^[12] In contrast to the widespread expression of α1 and b1, the other α- and β-isoforms exhibit more restricted patterns of expression. The α2-isoform is expressed principally in adipocytes, skeletal and cardiac muscle, and brain. The α3-isoform is abundant in nervous tissues, whereas the α4-isoform is a testis-specific isoform.^{[7] [8]} The β2-isoform is present primarily in skeletal muscle, pineal gland, and neural tissues. Estimates of mRNA abundance by RNase protection assay showed that β2 constituted only 5% of the total b subunit mRNA.^[11] β3 is present in testis, retina, liver, and lung.^[13] The expression pattern of the Na⁺,K⁺-ATPase isoforms is under developmental and hormonal controls and may vary in pathologic states. However, no consistent data have emerged to indicate significant changes in the α1β1 enzyme predominance in the kidney under such transitions.

The distribution of the Na⁺,K⁺-ATPase α-subunit in the kidney has been extensively examined, using immunohistochemistry, Western blots, in situ hybridization, and RT-PCR analysis of mRNA levels in microdissected nephron segments. In the aggregate, these studies indicate that the highest levels of Na⁺,K⁺-ATPase expression are in the medullary thick ascending limb of Henle's loop (MTAL), cortical thick ascending limb of Henle's loop (CTAL), and the distal convoluted tubule (DCT). Lower levels are evident in the proximal convoluted tubule (PCT) and cortical collecting duct (CCD), and very low levels are expressed in glomeruli, descending and ascending thin limbs of Henle (DTL and ATL, respectively), outer medullary collecting duct (OMCD), and inner medullary collecting duct (IMCD). These data correlate with studies in rabbit, rat, and mouse that have examined the amount of Na⁺,K⁺-ATPase hydrolytic activity and specific binding of [³H]ouabain in isolated nephron segments, indicating that the highest activity is in the MTAL, CTAL, and DCT, intermediate activity is in the PCT and CCD, and very low activity is in the proximal straight tubule (PST), DTL, and ATL^{[14] [15]} ( Fig. 4-3 ). The distribution and relative abundance of the Na⁺,K⁺-ATPase along the nephron are generally comparable among these three species. Moreover, the differences in activity from different segments of the nephron appear to be the result of differences in pump number rather than differences in enzyme turnover rates or ATP dependence. El-Mernissi and Doucet^[15] found that Na⁺,K⁺-ATPase hydrolytic activity and specific binding of [³H]ouabain to its single site on the Na⁺,K⁺-ATPase α-subunit were similar along the nephron.



FIGURE 4-3 Relative levels of Na⁺,K⁺-ATPase activity measured in individual segments of the rat nephron. CCD, cortical collecting duct; CTAL, cortical thick ascending limb of Henle's loop; DCT, distal convoluted tubule; MCD, outer medullary collecting duct; MTAL, medullary thick ascending limb of Henle's loop; PCT, proximal convoluted tubule; PR, pars recta (proximal straight tubule); TAL, thin ascending limb of Henle's loop; TDL, thin descending limb of Henle's loop. (Data are normalized to that of the DCT and are redrawn from Katz AI, Doucet A, Morel F: Na⁺-K⁺-ATPase activity along the rabbit, rat, and mouse nephron. Am J Physiol 237:F114–F120, 1979.)

In addition, the Na⁺,K⁺-ATPase has been shown to be regulated in a tissue-specific and isoform-specific fashion by direct interaction with at least five of the seven members of the FXYD family, including FXYD1 (phospholemman), FXYD2 (the γ-subunit), FXYD3 (Mat-8), FXYD4 (corticosteroid hormone-induced factor [CHIF]), and FXYD7, all members of the FXYD family of type II transmembrane proteins.^[16] In addition, a dominant-negative mutation in FXYD2 has been linked to cases of autosomal dominant renal magnesium wasting.^[17] A signature FXYD motif in the N-terminus and conserved glycine and serine residues in the transmembrane domain characterizes the FXYD proteins. The γ-subunit is expressed principally in the kidney, and CHIF is expressed in the medullary collecting duct and in the epithelium of the distal colon.^[16] Two principal subtypes of the 8- to 14-kD hydrophobic polypeptide, termed “γ_a” and “γ_b,” have been characterized in rat kidney. Both isoforms are highly expressed with the Na⁺,K⁺-ATPase α1-subunit in the MTAL, whereas γ_a is specific for cells in the macula densa and principal cells of the CCD but is not expressed in the CTAL. In contrast, γ_b is present in the CTAL.^[16] It is becoming increasingly apparent that the γ-subunit is a component of the renal Na⁺,K⁺-ATPase and that this subunit may influence kinetic properties of the holoenzyme. The γ-subunit decreases the apparent Na⁺ affinity of the Na⁺,K⁺-ATPase, increases the apparent affinity for ATP, and increases the K⁺ antagonism of cytoplasmic Na⁺ activation.^[16]In contrast, CHIF increases the apparent Na⁺ affinity of the Na⁺,K⁺-ATPase.^[16] CHIF knockout mice exhibit twofold higher urine volumes and an increased glomerular filtration rate under K-loaded conditions. In addition, treatment of K⁺-loaded mice for 10 days with furosemide resulted in increased mortality in the knockout mice but not in the wild-type group. These findings are consistent with an effect of CHIF on the Na⁺,K⁺-ATPase of the OMCD and IMCD.^[18]

H⁺,K⁺-ATPases.

The H⁺,K⁺-ATPases are a family of integral membrane proteins closely related to the Na⁺,K⁺-ATPase. The H⁺,K⁺-ATPase in vertebrates exists as an a/b heterodimer and is encoded by at least two distinct genes. The H⁺,K⁺-ATPase α1-subunit (HKa1, also termed the “gastric” H⁺,K⁺-ATPase) was originally cloned from rat stomach,^[19] and cDNAs encoding the HKa1-subunit have since been cloned from a wide range of species, including humans. The H⁺,K⁺-ATPase α2-subunit (HKa2, also termed the “colonic” or “non-gastric” H⁺,K⁺-ATPase) was first cloned from rat colon,^[20] and an orthologous gene product ATP1AL1 has been characterized in humans.^[21] Alternative splicing gives rise to HKa2 N-terminal splice variants in rabbit^[22] and rat,^[23] but these are not encoded by the mouse or human genes.^[24] The members of the H⁺,K⁺-ATPase gene family appear to participate in the control of body K⁺ balance, renal HCO₃^- absorption, the enhanced ammonium secretion in the IMCD during chronic hypokalemia, and intracellular Na⁺ regulation in the macula densa.^[25] The two isoforms differ in their tissue distribution, response to chronic K⁺ depletion, and inhibitor sensitivity. The HKa1-subunit is expressed in stomach and the medullary collecting ducts,^[26] but not distal colon, and is inhibited by low concentrations of Sch-28080 but not by ouabain,^[27] and its expression in the collecting duct is not affected by chronic hypokalemia.^[26] In contrast, the HKa2-subunit is expressed in the medullary collecting ducts^[28] and distal colon, but not stomach,^[20] is Sch-28080–resistant and partially ouabain-sensitive,^[27] and its expression in the collecting duct is upregulated by chronic hypokalemia.^[28] From an energetics standpoint, both H⁺,K⁺-ATPase isoforms are believed to have identical stoichiometries for ATP.

Ca²⁺-ATPase.

The human plasma membrane Ca²⁺-ATPase (PMCA) isoforms are encoded by at least four separate genes. The diversity of these enzymes is further amplified by tissue-specific alternative splicing within regulatory sites, which generate multiple subtypes of each isoform. The functional enzyme is thought to consist an ≈115-kD monomer. The enzyme isolated from kidney is calmodulin-dependent and has a K_0.5 for Ca²⁺ of approximately 0.7 μM, which correlates with the value obtained for ATP-dependent Ca²⁺ uptake in basolateral membrane vesicles from kidney cortex.^[29]

Differences in the structure and localization of PMCA splice variants confer specific regulatory properties that may have consequences for proper cellular Ca²⁺ signaling. The isoforms are expressed in a tissue-dependent manner, with PMCA1 and PMCA4 widely expressed among tissues, whereas PMCA2 and PMCA3 are expressed predominantly in brain and skeletal muscle.^[30] PMCA in concert with the NCX1 Na⁺/Ca²⁺ exchanger regulates intracellular Ca²⁺ concentrations and mediates both basal and hormone-stimulated Ca²⁺ efflux by distal tubules. Among nephron segments, the DCT possesses the highest Ca²⁺-ATPase activity^[31] and exhibits the strongest immunocytochemical reactivity for PMCA protein expression.^[32] Magocsi and colleagues^[33] reported the presence of multiple PMCA isoform mRNAs detected by RT-PCR in the kidney. PMCA1 was found in cortex, outer medulla, and inner medulla, PMCA2 in cortex and outer medulla, and PMCA3 in outer medulla (the PMCA4 cDNA sequence was unknown at that time). RT-PCR of cDNA generated from microdissected rat tubules demonstrated PMCA2 expression exclusively in proximal tubules, CTALs, and distal tubules.^[33] In a subsequent RT-PCR analysis by another laboratory, mRNAs for PMCA1 and PMCA2 were discovered to be abundant in the glomerulus, PCT, DTL, DCT, and CCD. PMCA3 mRNA was identified in the DTL and CTAL, and PMCA4 was found throughout the nephron.^[34] The concordance of data obtained in human, rat, and mouse indicates that PMCA1 and PMCA4 are the principal transcripts and protein expressed in kidney,^[32] whereas PMCA2 mRNA constitutes less than 2% of the total PMCA mRNA in the kidney and PMCA3 could not be detected. Molecular studies in immortalized mouse DCT cells (mDCT) demonstrated that PMCA1 and PMCA4 are the isoforms in these cells.^[32] Transcripts of the SERCA3 isoforms, which like the other SERCA members is inhibited by thapsigargin but unlike the others does not appear to be regulated by phospholamban, have also been detected in kidney.^[35] However, the physiologic role of this pump in the kidney is unknown.

Cu²⁺-ATPases.

The Menkes protein ATP7A and the Wilson's disease protein ATP7B are monomeric proteins with eight predicted transmembrane domains that export Cu²⁺, and possibly other metals, from the cytoplasm to an intracellular organelle. The Menkes protein is essential for efficient dietary Cu²⁺ uptake in the small intestine but also in the delivery of Cu²⁺ to the brain across the blood-brain barrier and recovery of Cu²⁺ from the proximal tubules of the kidney. Patients with Menkes disease exhibit defective Cu²⁺ efflux. Under normal Cu²⁺ conditions, both ATP7A and ATP7B are located in the trans-Golgi network where they provide Cu²⁺ to secreted cuproenzymes.^[36] When intracellular Cu²⁺ levels rise, ATP7A traffics to the plasma membrane to export the excess Cu²⁺. Patients with Menkes disease and Wilson's disease both suffer Cu²⁺ accumulation in proximal tubules. In some Wilson's disease patients this Cu²⁺ accumulation causes tubular dysfunction, resulting in the increased urinary amino acid and Ca²⁺ excretion. In situ hybridization and immunolocalization studies have demonstrated that both Atp7a and Atp7b are expressed in glomeruli; however, Atp7b is also seen in the kidney medulla.^[37]

Vacuolar H⁺-ATPases

In the kidney, V-ATPases play an important role in H⁺ secretion in the proximal tubule and along the length of the distal tubule and collecting duct. Immunohistochemical, biochemical, and physiologic studies demonstrated that the V-ATPase is present in proximal tubules, TALs, DCT, and intercalated cells of the collecting duct.^[38] Several heritable diseases have been attributed to defects in genes that encode V-ATPase subunits, including renal tubular acidosis and osteo-petrosis.^[39] The renal V-ATPases can be inhibited by N,N′-dicyclohexyl carbodiimide (DCCD), the sulfhydryl reagent N-ethylmaleimide, and more specifically, by the macrolide antibiotic bafilomycin A. The H⁺:ATP stoichiometry of the V-ATPase has been suggested to be up to 3:1, and the K_m for ATP of the purified V-ATPases is roughly 150 μM. V-ATPases are composed of a cytoplasmic catalytic domain (V₁), responsible for ATP hydrolysis, and a transmembrane domain (V₀), responsible for H⁺ translocation. The V₁ domain consists of eight distinct subunits (A-H), with an aggregate molecular mass of ≈570 kD. The 260-kDa V₀ domain complex is composed of 5 subunits (subunits α-d).^[38] The V-ATPase in kidney is regulated at multiple levels from changes in gene transcription and protein synthesis, to alterations in membrane insertion and interaction with heterologous proteins. Monogenic defects in two subunits (ATP6V0A4, ATP6V1B1) of the V-ATPase have been observed in patients with distal renal tubular acidosis.^[38] As discussed later, recent work suggests that the V-ATPase may also be functionally coupled to ATP production from glycolysis.

ATP-Binding-Cassette Transporters

P-Glycoproteins.

P-glycoproteins are ABC transporters originally discovered as drug pumps in multidrug-resistant cancer cells, but have since been found in many normal tissues. The 170-kD multidrug resistance transporter (MDR) confers resistance by active, ATP-dependent extrusion of a broad range of drugs that do not share obvious structural characteristics. These include anticancer drugs, immunosuppressive agents such as cyclosporine and FK506, cardiac glycosides, and antibiotics.^[3] Humans have two known P-glycoprotein genes, ABCB1 and ABCB4 (formerly known as MDR1 and MDR3), whereas rodents have three genes, termed mdr1a, mdr1b, and mdr2. The human MDR1 and mouse mdr1a and mdr1b encode an ≈170-kD plasma membrane ATPase (K_m≈38 μM ATP) that transports a wide range of structurally unrelated drugs, steroids, and phospholipids, and thereby confers multidrug resistance. In contrast, MDR3 and its ortholog mouse mdr2 encode P-glycoproteins that are phosphatidylcholine translocases and that have limited ability to transport numerous drugs, although they may transport some drugs in cell culture systems.^[40] In the kidney, the MDR1 mRNA and protein are expressed in mesangial cells, proximal tubule, TAL, and collecting duct.^[41] In mesangial^[42] and proximal tubule cells, P-glycoprotein has been shown to transport xenobiotics. Human MDR2 was localized to the of proximal tubules by double and triple immunofluorescence microscopy.^[43]

Cystic Fibrosis Transmembrane Regulator.

Cystic fibrosis transmembrane regulator (CFTR), another member of the ABC transporter family, couples ATP signaling with ion transport. CFTR is regulated by phosphorylation of its regulatory R domain and ATP hydrolysis at two nucleotide-binding domains, but it is unique among the ABC transporter family in that it functions as a Cl^- channel. Mutations of this transporter lead to a defect of epithelial Cl^- secretion causing the disease cystic fibrosis. CFTR transcripts have been identified in all nephron segments, and the encoded protein participates in Cl^- secretion in the distal tubule and the principal cells of the CCD and IMCD.^[44] Although patients with cystic fibrosis do not manifest serious renal dysfunction, they do have impaired ability to concentrate and dilute the urine and to excrete certain drugs.^[44] In addition to Cl^- secretion, CFTR has also been shown to secrete ATP directly^[45] or to modulate other ATP release channels.^[46] In this manner, CFTR may regulate other ionic conductances, such as the epithelial Na⁺ channel (ENaC)^[47] and ATP-regulated K⁺ channels (Kir 1.1., also known as ROMK)^[48] in the collecting duct. The effect of CFTR on the renal K⁺ secretory channel is mediated by protein kinase A, which may provide a functional switch designating the distribution of open and ATP-inhibited K⁺ channels in apical membranes. Recent studies also revealed that overexpression of CFTR promotes cell volume recovery from swelling or a regulatory volume decrease, enhances both constitutive and volume-sensitive ATP release, and through purinergic receptors, facilitates autocrine control of cell volume.^[49] CFTR requires the hydrolysis of ATP for activity and has been shown to interact physically with AMP-activated protein kinase (AMPK), with activation of AMPK resulting in an inhibition of CFTR in epithelial cells colonic and pulmonary epithelial cells (detailed later).^[50]

ENERGY PRODUCTION TO FUEL SOLUTE TRANSPORT

The various metabolic pathways that support and regulate solute transport along the nephron are highly integrated and interdependent. The oxidation of carbohydrate, lipid, and protein is tightly coupled to the generation and utilization of energy. The tricarboxylic acid (TCA) cycle and b oxidation of fatty acids are tightly linked to mitochondrial electron transport via the supply and demand of nicotinamide and flavin nucleotides. Similarly, electron transport is tightly coupled to oxidative phosphorylation and the supply and demand for ADP and ATP. Given its high transport demands, the kidney favors the more efficient ATP generation of aerobic metabolism (36 ATP produced per glucose consumed) over anaerobic metabolism (e.g., 6 ATP per glucose consumed in anaerobic glycolysis). Studies using diverse experimental methods in a variety of species, including humans, have provided a model of the relative contributions and intrarenal localization of specific metabolic pathways that fuel renal solute transport under physiologic and pathophysiologic conditions. These methods have included studies in the intact organism, isolated perfused kidney, renal tissue slices, tubule suspensions, and isolated nephron segments. Techniques applied to these studies have included measurements of ¹⁴CO₂ production from ¹⁴C-labeled substrates, oxygen consumption (QO₂), ATP contents, and NADH fluorescence, ³¹P nuclear magnetic resonance (NMR) spectroscopy, blood oxygen level-dependent (BOLD)-MRI, and others.

Mitochondrial ATP Production

Oxidative Phosphorylation

In 1924, Otto Warburg characterized the O₂ transferring component of the “respiratory enzyme” and established the phenomenon of cellular respiration. A few years later, Lohmann, Fiske, and Subbarow isolated ATP from muscle extract, and in 1937, Kaclkar reported the link between cellular respiration and ATP synthesis. In 1961, Mitchell proposed the chemiosmotic theory, which states that the energy stored in an electrochemical gradient across the inner mitochondrial membrane could be coupled to ATP synthesis. Although this theory met with controversy until the mid-1970s, it has now gained widespread acceptance.

Oxidative phosphorylation occurs in the mitochondrial inner membrane and includes the oxidation of metabolic fuels by O₂ and the associated transduction of energy into ATP. The electron transport chain, or respiratory chain, is a system of mitochondrial enzymes and redox carrier molecules that transfer reducing equivalents (electrons), obtained from the oxidation of respiratory substrates, to O₂ ( Fig. 4-4 ). It comprises five enzyme complexes (complexes I-V), ubiquinone (or coenzyme Q), and cytochrome c. This set of enzymes consists of NADH:ubiquinone oxidoreductase (complex I), succinate:ubiquinone oxidoreductase (complex II), ubiquinone:cytochrome c oxidoreductase (complex III, cytochrome reductase, cytochrome bc1), cytochrome c oxidase (complex IV, cytochrome oxidase), and ATP synthase (complex V, F₁F₀-ATPase). Other membrane-bound enzymes, such as the energy linked transhydrogenase, fulfill ancillary roles. The crystal structures of the major complexes of the electron transport chain (except complex I) have been established, permitting detailed analyses of the mechanism of H⁺ pumping coupled to electron transport in the mitochondria.



FIGURE 4-4 Mitochondrial oxidative phosphorylation. The mitochondrial electron transport chain conducts the oxidation of NADH or FADH₂, generates an H⁺ gradient across the inner mitochondrial membrane to drive ATP synthesis, and consumes O₂. The proteins that make up the electron transport chain are integral membrane proteins of the inner mitochondrial membrane. Substrate-level dehydrogenase reactions within the mitochondrial space generate NADH, which contributes 2 electrons (e^-1) to complex I. These electrons are sequentially transferred to complexes III and IV, with O₂ as the final acceptor. Ubiquinone (UQ) and cytochrome c (cyt c) function as mobile carriers of electrons between complexes. The flow of electrons from higher to lower redox potentials generates energy that is used to extrude 10 to 12 H⁺ from the matrix space. The H⁺gradient across the inner mitochondrial membrane is used to drive ATP synthesis by the ATP synthase (F₁F₀-ATPase, complex V). An adenine nucleotide translocase, which exchanges ATP^4- for ADP^3-, and a PO₄^-/OH^- exchanger in the inner mitochondrial membrane function to deliver and extrude ADP, P_i, and ATP.

The first step in the oxidation process involves transfer of electrons from NADH to complex I. Alternatively, electrons can transfer from FADH₂ to complex II. Because the latter complex does not move H⁺, oxidation of FADH₂results in the movement of fewer H⁺ across the membrane and the production of less ATP. Hence, NAD-linked substrates give consistently higher ADP:O ratios (≈2.5) compared with FAD-linked substrates, such as succinate (ADP:O ratio ≈1.5). Variability of coupling may occur under some conditions but is generally not significant. The fractional values result from the coupling ratios of proton transport. An additional revision of ADP:O ratio may be required because a recent report of the structure of ATP synthase suggests that the H⁺:ATP ratio is 10:3, rather than 3, consistent with ADP:O ratios of 2.3 with NADH and 1.4 with succinate.^[51]

After entry into the respiratory chain, electrons are transferred sequentially to coenzyme Q, complex III), cytochrome c, complex IV, and finally to O₂, which is reduced to H₂O (see Fig. 4-4 ). The free energy released by the fall in redox potential of the passing electrons during electron transfer drives the translocation of H⁺ via complexes I, II, and IV from the mitochondrial matrix to the inner mitochondrial space. This process generates a H⁺ electrochemical potential gradient across the inner mitochondrial membrane of 200 to 230 mV,^[52] which is known as the proton-motive force. The H⁺ then reenters the matrix via the F₁F₀-ATPase of complex V (ATP synthase), driving ATP synthesis. The F₁F₀-ATPase complex (Mr ≈500,000) contains at least 12 distinct subunits, several of which are present in multiple copies. The catalytic F₁ head group, which contains three nucleotide binding sites, is connected by an oligomycin-sensitive stalk to a H⁺-conducting F₀ baseplate embedded in the mitochondrial inner membrane. Three H⁺ are thought to pass through the membrane for each molecule of ATP manufactured by the complex. An increase in Na⁺,K⁺-ATPase activity, therefore, decreases the intramitochondrial phosphorylation potential ([ATP]/[ADP][P_i]), which results in more rapid H⁺ entry via the F₁F₀-ATPase, a dissipation of the H⁺ gradient, and more rapid electron transfer along the electron transport chain to extrude H⁺ and increase QO₂.

The proton-motive force arises both from the change in membrane potential from the net movement of positive charge across the inner mitochondrial membrane and the pH gradient. Of these two components, the membrane potential contributes most of the energy stored in the gradient. The H⁺ gradient can also be dissipated by the presence of the uncoupling proteins UCP1, UCP2, UCP3, or by other transport processes for various ions and small molecules. Additional transporters influence the membrane potential of the inner mitochondrial membrane, and thereby the proton-motive force. The adenosine nucleotide translocator, which conducts the electrogenic 1:1 exchange of ADP^3-for ATP^4-, a phosphate transporter, which imports PO₄^- in exchange for OH^-, and a constitutive proton H⁺ leak reside in the inner mitochondrial membrane protein (see Fig. 4-4 ). The voltage-dependent anion channel localizes to the outer mitochondrial membrane. In addition to the proteins involved in the TCA cycle, the electron transport chain, and ATP synthesis, a number of other factors, including shuttles and transporters, are required for oxidative phosphorylation. Mitochondrial carrier proteins, integral membrane proteins that transport metabolites and cofactors across the inner membrane of mitochondria, are also required for ATP synthesis, the TCA cycle, fatty acid β-oxidation, and the malate shuttle.

Under certain stresses, such as free radical-mediated damage, mitochondria experience an irreversible autocatalytic collapse, with a loss of the normal membrane potential and a failure of ATP production, termed the mitochondrial permeability transition. The transition involves the integration of adenine nucleotide transporter subunits and other outer membrane proteins into a large pore, which allows free entrance of small ions to the mitochondrial interior. Atractyloside inactivates the ATP/ADP antiporter and favors pore formation, whereas bongkrekic acid and cyclosporine inhibit the process. Mitochondria that undergo the permeability transition release pro-apoptotic molecules.

Oxygen Consumption, Respiratory Control, and Coupled Respiration

Much of our understanding of mitochondrial respiration has come from studies of QO₂ in cells and isolated mitochondria. When normal mitochondria are incubated in an isotonic medium containing substrate and phosphate, ADP addition promotes a sudden increase in QO₂ as the ADP is converted into ATP. This active state of respiration, termed “state 3” respiration, distinguishes the maximal QO₂ that is coupled to ATP production. In permeabilized proximal tubules for example, ADP stimulates QO₂ by four- to fivefold over baseline.^[53] The subsequent slower rate of QO₂ after all the ADP has been phosphorylated to form ATP is referred to as state 4. The ratio of QO₂ in state 3 to that in state 4 is termed the respiratory control index, and it reflects the O₂ uptake (oxidation of NADH and/or FADH₂) by intact mitochondria and the simultaneous conversion of ADP and inorganic phosphate into ATP. Because the catalytic site for ATP synthesis by the F₁F₀-ATPase is in the mitochondrial matrix, respiratory control is likely related to the availability of ADP and the kinetics of its transport by the adenine nucleotide translocase, a hypothesis first proposed by Chance and Williams in the 1950s.^[54] Coupled respiration refers to O₂ uptake dependent on the presence of ADP and P_i. Respiratory control reflects the fact that the oxidation of NADH and FADH₂ is coupled to the H⁺ transport across the mitochondrial inner membrane. If the movement of H⁺ through the F₁F₀-ATPase to drive ATP synthesis does not dissipate the H⁺ gradient, the energy required for H⁺ translocation would exceed that derived from electron transfer and thus inhibit further electron transport. Competing hypotheses have been proposed to explain the factor(s) responsible for respiratory control. Lemasters and Sowers^[55] proposed that the kinetics of the adenine nucleotide translocase, an inner mitochondrial membrane protein that exchanges cytosolic ADP^3- for intramitochondrial ATP^[4], governs the rate of ATP synthesis. The adenine nucleotide translocase operates in parallel with a P_i/OH^- exchanger in the inner mitochondrial membrane that uses the H⁺ gradient to drive P_i entry (see Fig. 4-4 ). Inhibitors of the adenine nucleotide translocase, such as atractyloside, caused inhibition of ADP influx, respiration, and ATP synthesis.^[55] However, it is unclear whether [ADP] itself or the [ATP]/[ADP] ratio preferentially regulates the translocase.

The near-equilibrium hypothesis of Erecinska and Wilson^[56] proposed that respiration and ATP synthesis are mainly regulated by the phosphorylation potential and the NADH/NAD⁺ ratio. However, oxidative phosphorylation may not always be near equilibrium, and relative proximity to equilibrium does not necessarily exclude the contributions of the electron transport chain, H⁺ leak, F₁F₀-ATPase, or adenine nucleotide translocase to regulation of essential fluxes. In some instances, for example, respiration rate may correlate better with [ADP] than with phosphorylation potential, and may be relatively insensitive to mitochondrial NADH/NAD⁺ ratio. Although it is clear from these considerations that mitochondrial respiratory control is a complex process potentially involving ATP, ADP, P_i, the NAD redox state, cytochrome c, O₂, and other factors as signals, there is compelling evidence to support the concept that the dynamics of cytosolic ATP, ADP, and P_i participate in the coupling between mitochondrial respiration and active transport.

QO₂ measurements also provide important insights into the coupling of active Na⁺ transport and cellular respiration ( Table 4-2 ). As noted earlier, the state 3 rate of respiration provides an index of the maximal rate at which mitochondrial oxygen consumption is coupled to ATP production. Because tubule cells are impermeable to ADP, the tubules are first permeabilized by additions of low concentrations of digitonin, before addition of ADP. Assays of carbonylcyanide-m-chlorophenylhydrazone (CCCP)–uncoupled QO₂, provide similar information about mitochondrial respiratory capacity in the intact cell. The oligomycin-sensitive component of basal QO₂ represents that directly related to ATP synthesis. In proximal tubules, oligomycin inhibits at least 80% of basal QO₂.^[57] The ouabain-sensitive rate of respiration indicates that proportion of respiration devoted to the operation of the Na⁺,K⁺-ATPase and of secondary active transport coupled to the Na⁺,K⁺-ATPase. The value of ouabain-sensitive QO₂ varies among nephron segments ranging from 8% in the OMCD to 60% of total QO₂ in the PCT, according to the proportionate needs of active Na⁺ transport (see Fig. 4-8 ). The basal, ouabain-insensitive respiration reflects mitochondrial respiration devoted to Na⁺-independent processes, such as the operation of other primary and secondary active transport processes (e.g., H⁺-ATPase, Ca²⁺-ATPase), synthesis of DNA, RNA, protein, lipids, and glucose (gluconeogenesis), mitochondrial H⁺ leak, and substrate interconversions. For unclear reasons, the basal, ouabain-insensitive QO₂ is considerably higher in isolated renal cells and tubules (40%–90%), compared with measurements in whole kidney (<20%). Addition of cationophores, such as nystatin or monensin, allows the Na⁺ concentration to equilibrate across the membrane and provides maximal stimulation of the Na⁺,K⁺-ATPase without adversely affecting mitochondrial function. Thus, the effects of the maximal stimulation, with nystatin, and maximal inhibition, with ouabain, of Na⁺,K⁺-ATPase activity on QO₂ provide an assessment of the integrity of this ion pump and its coupling to Na⁺ entry and to mitochondrial respiration. Elegant studies in suspensions of cortical tubules by Mandel's laboratory, for example, found that nystatin stimulated QO₂ by nearly 60% above its spontaneous rate and to the respiratory capacity of the tubules as defined by the state 3 rate. In contrast, ouabain inhibited spontaneous QO₂ by about 50%.^[58] In these same studies, ATP contents were found to change little during these maneuvers: ATP declined by 15% during nystatin stimulation and increased by 6% during ouabain inhibition.^[59]

TABLE 4-2 -- Components of QO₂

QO₂ Parameter	Interpretation
Ouabain-insensitive	Basal rate, composed of Primary and secondary active transport not coupled to the Na⁺,K⁺-ATPase (e.g., H⁺-ATPase)
	Biosynthetic functions (lipids, glucose)
	Cell growth and repair
	Substrate interconversions and transformations
Ouabain-sensitive	Na⁺,K⁺-ATPase and secondary active transport coupled to the Na⁺,K⁺-ATPase
Nystatin-simulated	Maximal activation of the Na⁺,K⁺-ATPase; should be completely inhibited by ouabain
CCCP uncoupled	Maximal mitochondrial respiratory capacity
Oligomycin-sensitive	QO₂ coupled to ATP synthesis from mitochondrial oxidative phosphorylation

QO₂ can be used to dissect mechanisms that couple active Na⁺ transport and mitochondrial oxidative phosphorylation. The ouabain-insensitive QO₂ provides a measurement of basal QO₂ that is independent of Na⁺ transport. The ouabain-sensitive rate is related to active Na⁺ transport mediated by the Na⁺,K⁺-ATPase. The nystatin-stimulated QO₂ tests the integrity of the functions of and links between Na⁺ entry, Na⁺,K⁺-ATPase, and mitochondrial respiration. The carbonylcyanide-m-chlorophenylhydrazone (CCCP)-uncoupled QO₂ and the oligomycin-sensitive of QO₂ provide information regarding mitochondrial integrity and function.



FIGURE 4-8 Relative distributions of QO₂, mitochondrial density, ATP content, and Na⁺,K⁺-ATPase activity along the mammalian nephron. For each item, the results were normalized to the maximal value in the nephron. QO₂ data obtained from references 76, 157, 158, 171, 200–202 with the maximal value for total QO₂ in the MTAL (2000 ng/mg protein/h). The percentages indicated in the unshaded portion of the bars represent the ouabain-sensitive component of QO₂. Mitochondrial density data from reference 100 with a maximal value of 44% in the MAL. ATP content from reference 154 with a maximal value of 16.8 mmol/kg dry wt reported in DCT. Na⁺,K⁺-ATPase activities from reference 203 with the DCT value of 6679 pmol/mm tubule/h being maximal. CCD, cortical collecting duct; CTAL, cortical thick ascending limb of Henle's loop; GLOM, glomerulus; IMCD, inner medullary collecting duct; MTAL, medullary thick ascending limb of Henle's loop; OMCD, outer medullary collecting duct; S1, S1 segment of proximal tubule; S2, S2 segment of proximal tubule; S3, S3 segment of proximal tubule; TL, thin limbs of Henle's loop; (Adapted from Soltoff SP: ATP and the regulation of renal cell function. Annu Rev Physiol 48:9–31, 1986.)

Mitochondrial Substrate Entry

Many important metabolites show an asymmetrical distribution across the mitochondrial inner membrane. Therefore, normal operation of the respiratory chain requires highly specific transporters to control the movement of substrates across the membrane. These include electroneutral uptake mechanisms for phosphate, malate, succinate, 2-oxoglutarate, and citrate, as well as several exchangers for organic solutes ( Fig. 4-5 ). In general, these transporters exploit directly or indirectly the H⁺ gradient generated by the electron transport chain. Consequently, conditions that alter the mitochondrial pH gradient, such as intracellular acidosis or certain drugs, can change the concentrations of metabolites within the mitochondria and thereby influence oxidative metabolism. The transporter expression and specificity governs the ability of various tissues to conduct substrate oxidations (see Fig. 4-5 ). Although many of these transporters have been identified in the kidney, the intrarenal distribution of specific transporters has not yet been defined. Moreover, the role of these transporters has largely been studied in mitochondria from heart or liver, so that there is little information regarding the specific properties of these transporters in renal cells.



FIGURE 4-5 Major metabolite transporters in the mitochondrial inner membrane. These transporters permit the selective accumulation of organic solutes in the mitochondrial matrix that can be metabolized by the TCA cycle and other mitochondrial enzymes, as well as ADP and P_i needed for ATP synthesis. These pathways are driven directly or indirectly by the H⁺ gradient (matrix side alkaline), membrane potential (matrix side negative), and/or solute gradients.

Three of the most important metabolite carriers are the energy-driven aspartate/glutamate exchange (aspartate exchanges for glutamate plus a H⁺), the aspartate carrier, and the carnitine palmitoyltransferase (CPT) system. The aspartate/glutamate exchange plays an important role in maintaining the cytosolic compartment in a relatively “oxidizing” state (with a low NADH/NAD⁺ ratio), while keeping the mitochondrial space correspondingly reduced and suppressing the aerobic synthesis of lactic acid. The aspartate carrier plays a central role in the reoxidation of glycolytic NADH by the malate-aspartate cycle, involving malate dehydrogenase and glutamate oxaloacetate transaminase. This cycle overcomes the fact that the inner membrane is impermeable to NAD⁺ and NADH. The carnitine palmitoyltransferase (CPT) system (CPTI and CPTII) mediates the transport of long-chain fatty acids into the mitochondria for oxidation. CPTI exchanges carnitine for the CoA attached to long-chain fatty acids to create a fatty acid-carnitine conjugate. This conjugate is transported into the matrix by a transporter protein in the inner mitochondrial membrane. Once the fatty acid-carnitine conjugate is inside the matrix, CPTII exchanges CoA for carnitine to produce new fatty acid-CoA again, which is ready to enter fatty acid oxidation in the matrix. The liberated carnitine is exported to renew the cytoplasmic pool of carnitine and allow the transfer process to continue.

Inhibitors of Mitochondrial Function

Much of our knowledge of mitochondrial function has resulted from the use of specific inhibitors to probe discrete mechanisms. These compounds have been used to distinguish the electron transport system from the phosphorylation system and to establish the sequence of redox carriers along the respiratory chain ( Fig. 4-6 ). In addition, they have been used to probe the contributions of specific metabolic pathways to solute transport along the nephron. Several categories of mitochondrial inhibitors can be distinguished. Respiratory chain inhibitors (e.g., antimycin and rotenone) block mitochondrial respiration, even in the presence of either ADP or uncoupling agents. Rotenone inhibits complex I, and antimycin A inhibits cytochrome c. Phosphorylation inhibitors, such as oligomycin, block the burst of QO₂ and ATP manufacture after ADP addition, but only if the respiratory coupling is intact. Uncoupling agents (e.g., dinitrophenol, CCCP) inhibit the coupling between the respiratory chain and the phosphorylation system. By shuttling H⁺ across the inner mitochondrial membrane, uncoupling agents dissipate the H⁺ gradient, uncouple the processes of oxidation and phosphorylation, and block ATP synthesis. This results in a high rate of QO₂ in the absence of ADP until all O₂ is consumed. Transport inhibitors (e.g., atractyloside, bongkrekic acid, N-ethylmaleimide) prevent either ATP export or substrate import across the mitochondrial inner membrane. Ionophores (e.g., valinomycin, nigericin) render the inner membrane leaky to ions to which the membrane is usually impermeable.



FIGURE 4-6 Inhibitors of mitochondrial respiration. Representative inhibitors of mitochondrial oxidative phosphorylation and their sites of action in the inner mitochondrial membrane are shown. Glib, glibenclamide; Ruth red, ruthenium red.

Renal Transport Defects Associated with Mitochondrial Genetic Disorders

Genetic mutations in components of the electron transport chain or ATP synthase that affect renal solute transport have been described. In general, renal manifestations of these disorders tend to be more commonly observed in children than in adults. Most of these diseases affect other organ systems as well, particularly the central nervous system and muscle. Of the renal manifestations, the most common defect reported is a proximal tubulopathy resulting in a form of the Fanconi syndrome, but other patients developed tubulointerstitial nephritis, renal failure, or nephrotic syndrome.^[60] Because the Na⁺,K⁺-ATPase drives the secondary active transport of glucose, phosphate, and amino acids, insufficient ATP supply to this pump results in impaired reabsorption of these solutes. Renal biopsy often shows giant mitochondria and nonspecific abnormalities of the tubules, including tubular atrophy and casts.^[60]Diagnosis rests on establishing proper oxidoreduction status and activity of enzymatic complexes of the respiratory chain. For most patients—especially those presenting with tubulopathy—plasma lactate, pyruvate, βOH butyrate, acetoacetate, and their molar ratios were within the normal ranges. This could be ascribed to renal leakage, which contributes to lowering blood lactate. Thus, normal lactatemia does not rule out the hypothesis of a mitochondrial disorder in patients with renal involvement. However, urinary organic acids chromatography revealed high lactate and Krebs cycle intermediates, which are highly suggestive of mitochondrial disorder. Treatment is generally unsatisfactory. However, patients with complex I deficiency may be treated with riboflavin and ubidecarone (coenzyme Q10),^[61] and patients with complex III deficiency may benefit from menadione (vitamin K₃) or ubidecarone. Patients with secondary carnitine deficiency may benefit from carnitine supplementation.

Glycolysis

Glycolysis, the formation of lactate from glucose or glycogen, is accomplished by the actions of 11 cytoplasmic enzymes. In the kidney, glycolysis derives principally from glucose because glycogen stores are minimal. Glycolysis can occur under anaerobic or aerobic conditions, in which case mitochondria can continue the oxidation of glucose by converting lactate or pyruvate to CO₂. Glycolysis may be particularly important in the renal medulla and papilla where O₂ concentrations are low and aerobic metabolism is not effectively supported. The key regulatory enzymes—hexokinase, phosphofructokinase, and pyruvate kinase—are most active in the renal medulla. The distribution of these enzymes along the rodent and rabbit nephron has been examined^{[62] [63]} ( Fig. 4-7 ). In the rat, hexokinase and pyruvate kinase activities were relatively low in the proximal tubule but are at least 10-fold greater in the MTAL and about 8-fold greater in the DCT. In the rabbit, the activity of the hexokinase is lowest in the PCT and progressively increases in parallel with pyruvate kinase and phosphofructokinase along the distal nephron, including the MTAL, CTAL, DCT, and the entire collecting duct, with highest values in the connecting tubule. In addition, superficial nephrons exhibited lower activities of pyruvate kinase and phosphofructokinase compared with juxtamedullary nephrons.^[64] RT-PCR analysis of rat nephron segments demonstrated the expression of aldolase A and pyruvate kinase in the thin limb and collecting ducts of the medulla and in the distal tubules and glomeruli of the cortex.^[65]



FIGURE 4-7 Intrarenal distribution of enzymes involved in four major metabolic pathways. Assays of enzymatic activity for each of the indicated enzymes were performed on microdissected nephron segments. The data represent the percent of maximal activity for each enzyme. α-KGDH, α-ketoglutarate dehydrogenase; FACoA Ox, fatty acyl-CoA oxidase; FP, fructose bisphosphatase; GP, glucose-6-phosphatase; HK, hexokinase; ICDH, isocitrate dehydrogenase; OHACoA DH, 3-hydroxyacyl-CoA dehydrogenase; PEPCK, phosphoenolpyruvate carboxykinase; PFK, phosphofructokinase; PK, pyruvate kinase. (Data from references 62, 63, 66, 68, 169, 199.)

Tricarboxylic Acid Cycle

The enzymes for the TCA cycle are located in the matrix of the mitochondria. Acetyl coenzyme A is the major substrate entering the TCA cycle, and three products are generated: (1) H⁺, for oxidative phosphorylation, (2) CO₂ from the metabolism of carbohydrate, fat, and proteins, and (3) ATP, synthesized by a substrate phosphorylation step in the TCA cycle. The activity of the TCA cycle enzymes oxoglutarate dehydrogenase, citrate synthase, isocitrate dehydrogenase, and α-ketoglutarate dehydrogenase, and isocitrate dehydrogenase are particularly high in the CTAL, MTAL, and DCT, and very low in the TDL and the medullary collecting duct (see Fig. 4-7 ). Oxoglutarate dehydrogenase shows the lowest activities along the whole nephron and appears to catalyze the rate-limiting step of the TCA cycle.^[66]

Ketone Body Metabolism

The kidney can utilize ketone bodies as an energy source. In studies of cortical tissue slices, for example, acetoacetate met the majority of energy demands.^[67] Guder and colleagues^[68] described the distribution of two enzymes of ketone body metabolism, 3-oxoacid CoA-transferase and 3-hydroxybutyrate dehydrogenase in dissected segments of the mouse nephron. Both enzymes were found in all nephron segments, and their distribution generally paralleled the distribution of mitochondria along the nephron. The highest activities of the enzymes were observed in the TAL and DCT and decreased to only 20% of these values in the collecting duct. The proximal tubule exhibited heterogeneity, with uniform activity of 3-hydroxybutyrate dehydrogenase along its length, whereas the activity of 3-oxoacid CoA-transferase increased fivefold from PCT to PST (see Fig. 4-7 ).

Fatty Acid Oxidation

The oxidation of fatty acids is an important source of energy for ATP production in mitochondria through the entry of acetyl-CoA into the TCA cycle. Fatty acids can be oxidized in all nephron segments, with their rates of oxidation mirroring the mitochondrial mass of the nephron segments.^[69] Fatty acyl-CoA oxidase activity is greatest in the proximal tubule, increasing along its length, whereas there is little activity in more distal structures (see Fig. 4-7 ). The mitochondrial enzyme 3-hydroxyacyl-CoA dehydrogenase generally followed the distribution of mitochondrial density along the nephron, with highest activities in the PCT and DCT.^[66]

Pentose-Phosphate Shunt

The pentose-phosphate cycle yields the reducing energy needed for the biosynthesis of fatty acids, steroids, and nucleotides. It completely oxidizes glucose under anaerobic conditions. Glucose 6-phosphate dehydrogenase, the first enzyme in the pentose phosphate shunt, is found throughout the nephron, with highest catalytic activities in the proximal tubules and the DCT (see Fig. 4-7 ). Pentose-phosphate shunt activity has been demonstrated using radiolabeled glucose consumption in tissue slices and nephron segments.^[70] Based on these studies, this pathway is estimated to account for about 10% of total glucose utilization in the kidney.

GLUCONEOGENESIS AND ITS ROLE IN SOLUTE TRANSPORT

In 1937, Benoy and Elliott^[71] demonstrated the capacity of the kidney to perform gluconeogenesis, the de novo formation of glucose from noncarbohydrate precursors. Gram-for-gram, the kidney exhibits glucose synthetic rates several times higher than that observed in the liver.^[72] However, because arteriovenous difference measurements revealed that the kidneys of postabsorptive subjects show little or no net take-up or release of glucose, renal gluconeogenesis in humans under normal or pathologic conditions was believed to be of little consequence compared with that of the liver until the mid-1990s. Then, the combination of arteriovenous difference measurements with tracer techniques demonstrated that, in healthy postabsorptive humans, the renal glucose release approaches 20% of all glucose released in the circulation.^[73] The kidney has minimal glycogen stores, and its cells that are capable of storing glycogen lack glucose-6-phosphatase. Thus, renal glucose release occurs almost exclusively from gluconeogenesis. Renal glucose release is of the same order of magnitude as splanchnic glucose release during the postabsorptive period in humans, appears to be more sensitive to hormone action than hepatic glucose release, and may have a more important role during the adaptation to various physiologic and pathologic conditions. The kidney is now recognized as playing a key role in interorgan glucose metabolism, and particularly in the Cori cycle and glutamine-glucose cycle.

Compared with liver, renal gluconeogenesis has different substrate requirements and responds to different regulatory stimuli. The preferred gluconeogenic precursors in the kidney are lactate, glutamine, glycerol, α-ketoglutarate, and citrate. Of these, lactate is the predominant precursor for both renal and systemic gluconeogenesis in humans. Renal gluconeogenesis from lactate was shown to be 3.5-, 2.5-, and 9.6-fold greater than that from glycerol, glutamine, and alanine, respectively. These four substrates accounted for ≈90% of renal gluconeogenesis in humans.^[73] In vitro studies of microdissected S1, S2, and S3 human proximal tubules incubated with physiologic concentrations of glutamine or lactate revealed that the three successive segments have roughly the same capacity to synthesize glucose from glutamine, whereas the S2 and S3 segments synthesize more glucose from lactate than the S1 segment.^[74]Studies in multiple species indicate that it is difficult to extrapolate results concerning the metabolic heterogeneity of the nephron obtained in a given animal species to humans. In humans, renal gluconeogenesis is altered under physiologic and pathophysiologic states, including fasting, hypoglycemia, and diabetes mellitus. cAMP was a potent agonist of gluconeogenesis from lactate and glutamine by human proximal tubules, whereas adrenaline and noradrenaline were ineffective, suggesting that the stimulation of renal gluconeogenesis observed in vivo with adrenaline infusion may result from an indirect action on the renal proximal tubule.^[74]

Studies in animals and humans indicated that only the proximal tubule is capable of synthesizing glucose, and the only nephron segment that contains the key gluconeogenic enzymes glucose 6-phosphatase, fructose 1,6-diphosphatase, and phoshoenolpyruvate carboxykinase (PEPCK)^[69] (see Fig. 4-7 ). mRNA transcripts encoding fructose-1,6-bisphosphatase aldolase B and PEPCK have been detected in rat proximal cells.^[65] Aldolase B was found to be bound specifically to fructose 1,6-diphosphatase in these cells. In the absence of glutamine and lactate, glucose synthesis from endogenous substrates is negligible^[74] and consistent with the very low glycogen content of the proximal tubule in normal subjects. The proximal tubule is metabolically heterogeneous along its course, however, and species differences exist. Microdissected S1, S2, and S3 segments of the human proximal tubule synthesize glucose from lactate and glutamine at physiologic concentrations, with the S2 and S3 segments capable of more glucose from a physiologic concentration of lactate than the S1 segment. Similar heterogeneity was observed in the proximal tubule, in which the pars recta was shown to generate more glucose from lactate or pyruvate than the pars convoluta.^[75]

Studies in cortical tubule suspensions and the isolated perfused kidney revealed a reciprocal relationship between Na⁺ transport and gluconeogenesis. In some experiments, inhibition of Na⁺,K⁺-ATPase activity, and thereby, active Na⁺ reabsorption, stimulated gluconeogenesis in the kidney. Conversely, stimulation of Na⁺,K⁺-ATPase activity by nystatin^[76] or monensin inhibited gluconeogenesis in proximal tubules. Nagami and Lee^[77] demonstrated that increased Na⁺ transport in isolated proximal tubules was associated with a fall in glucose production and that inhibition of secondary active Na⁺ transport with amiloride increased glucose production. In agreement with this work, furosemide, ethacrynic acid, and chlorthiazide, all inhibitors of secondary active Na⁺ transport pathways, stimulated gluconeogenesis in kidney cortex slices.^[78] Because both gluconeogenesis and the Na⁺,K⁺-ATPase are ATP-consuming processes, it was proposed that they might compete for energy availability in the proximal tubule. However, ouabain inhibition of Na⁺,K⁺-ATPase does not always stimulate gluconeogenesis. In the isolated perfused kidneys of rat, for example, ouabain inhibited gluconeogenesis from lactate.^[79] In the aggregate, the studies suggested that under normal circumstances, the renal proximal tubule can meet the energetic demands of both gluconeogenesis and active Na⁺ transport and that these processes are subject to complex control.

COUPLING OF ACTIVE SOLUTE TRANSPORT AND ATP PRODUCTION

Whole Kidney Studies

Given the high rates of active Na⁺ transport in the kidney and the high mitochondrial densities in many portions of the nephron, it is not surprising that there is a linear correlation between the rate of Na⁺ reabsorption (T_Na+) and the QO₂ over a wide range of Na⁺ transport rates. The relationship of T_Na+/QO₂ has been reported to be in the range of 26 to 30.^[80] Several investigators challenged the legitimacy of using T_Na+ as an accurate measure of active Na⁺reabsorption. For example, various provocative maneuvers, such as water deprivation or administration of mannitol,^[80] loop diuretics, or dopamine, resulted in a higher T_Na+/QO₂ coupling ratio than was predicted. In contrast, studies in tight epithelia such as frog skin or toad urinary bladder, revealed lower than predicted T_Na+/QO₂ coupling ratios, in the range of 16 to 20 and in closer agreement to the predicted value of 18.^[81] The greater T_Na+/QO₂coupling efficiency in the kidney compared with other systems was attributed to a significant (≈30%) proportion of passive Na⁺ transport. Studies subsequently designed to minimize passive transport obtained lower values for the coupling ratio, in the range of 12.5 to 20.^[82] Ostensen and colleagues^[83] used transport inhibitors to quantify the relative proportions of NaHCO₃ and NaCl transport that were actively and passively transported in the proximal and distal tubule of dogs. Under conditions of saturated distal NaCl reabsorption, acetazolamide, an inhibitor of carbonic anhydrase-dependent HCO₃^- and Na⁺ reabsorption in the proximal tubule, reduced HCO₃^- reabsorption, Na⁺reabsorption, and QO₂, whereas mannitol reduced NaCl reabsorption without significantly affecting NaHCO₃ reabsorption or QO₂. The investigators concluded that proximal NaHCO₃ reabsorption is the only important active Na⁺transport process that is sensitive to inhibition of carbonic anhydrase and that NaCl transport is passive in this segment. In addition, they concluded that the distal nephron exhibits a T_Na+/QO₂ of 18 when proximal Na⁺ reabsorption is blocked with acetazolamide and ouabain is added. These whole kidney studies, however, suffered from major limitations related to the numerous assumptions made and the inability to determine Na⁺ transport in specific nephron segments.

Correlation Between Na⁺ Transport and QO₂ Along the Nephron

The heterogeneity of oxidative metabolism and active Na⁺ transport along the nephron has been studied extensively in isolated cell or tubule suspensions from selected nephron segments using measurements of ouabain-sensitive QO₂, Na⁺,K⁺-ATPase activity, cellular ATP content, and mitochondrial density. In general, there is a strong correlation between active Na⁺ transport and these parameters along the nephron ( Fig. 4-8 ). Segments with high rates of active Na⁺ transport, such as the MTAL, exhibit high rates of ouabain-sensitive QO₂ and Na⁺,K⁺-ATPase activity, abundant cellular ATP, and many mitochondria (see Fig. 4-8 ). At the other end of the spectrum, the IMCD conducts lower amounts of active Na⁺ transport, has fewer mitochondria, and accordingly, has lower rates of ouabain-sensitive QO₂ and Na⁺,K⁺-ATPase activity.

In addition, the coupling efficiency of transepithelial Na⁺ transport and mitochondrial ATP generation differs considerably among nephron segments. In the proximal tubule, the T_Na+/QO₂ has been reported to be in the range of 24 to 30, indicating a Na⁺/ATP ratio of about 4 to 5 (assuming that six ATP are produced per O₂ consumed). Because the proximal tubule is a leaky epithelium, minor osmotic and electrochemical gradients are able to drive passive NaCl transport across the epithelium. Fromter and colleagues^[84] calculated that as much as 50% of net Na⁺ reabsorption in the proximal tubule is passively mediated, whereas Kiil and coworkers^[85] suggested that this number was 30%. Because ATP hydrolysis is not directly coupled to passive transepithelial Na⁺ transport, passive transport increases the T_Na+/QO₂ and Na⁺/ATP ratios. In addition, apical Na⁺ influx is coupled to H⁺ efflux by the Na⁺/H⁺ exchanger NHE3, a process that generates intracellular HCO₃^- ( Fig. 4-9 ). This HCO₃^- is then extruded across the basolateral membrane by an electrogenic Na⁺/HCO₃^- cotransporter kNCB1. The ionic stoichiometry of this transporter in the proximal tubule has been reported to be 3Na⁺:1HCO₃^-, although it may be 2Na⁺:1HCO₃^- in other cell types.^[86] Therefore, with these stoichiometries, one third of the Na⁺ transported into the cell by the apical Na⁺/H⁺ exchanger could be transported across the epithelium and extruded by the basolateral 3Na⁺:1HCO₃^- exchanger without the need for ATP hydrolysis. Thus, the Na⁺:ATP would be 4.5 (one third greater than the 3Na⁺:1ATP stoichiometry of the Na⁺, K⁺-ATPase), with a T_Na+/QO₂ ratio of 27:1. However, HCO₃^- absorption in the proximal tubule is also mediated by an apical H⁺-ATPase working in tandem with the apical Na⁺/H⁺ exchanger. The H⁺-ATPase has been estimated to mediate between 10% and 50% of the total apical H⁺ transport.^[57] Therefore, this transporter would contribute to HCO₃^- absorption at the expense of the Na⁺/H⁺ exchanger, which would reduce the Na⁺/ATP ratio toward 3:1, depending on the H⁺:ATP stoichiometry of the apical H⁺-ATPase. Thus, through the contributions of passive paracellular Na⁺ transport and of Na⁺-HCO₃^- coupled transport, the proximal tubule operates a highly efficient mechanism for transepithelial Na⁺ transport.



FIGURE 4-9 Cellular models characterizing the stoichiometric relationships between transepithelial sodium reabsorption (T_Na+) and QO₂ or ATP utilization (ATP) for the proximal tubule (PT), medullary thick ascending limb of Henle's loop (TAL), and cortical collecting duct (CCD) principal cell. See text for discussion.

The coupling efficiency of transepithelial Na⁺ transport and mitochondrial ATP generation in the MTAL is believed to be greater than that observed in other nephron segments, although this has not been directly established. Studies in model epithelia such as the shark rectal gland^[87] and tracheal epithelium^[88] suggest that the T_Na+/QO₂ of the MTAL is 36:1 and the Na⁺/ATP ratio is 6:1 (see Fig. 4-9 ). The high coupling efficiency in this epithelium is the result of the stoichiometry of the Na⁺/K⁺/2Cl^- transporter of the apical membrane, which allows the energy derived from the Na⁺ pump to drive the secretion of 2Cl^- molecules per Na⁺. The resulting Cl^- electrochemical gradient drives passive transepithelial Na⁺ transport via paracellular pathways. Like other high-resistance Na⁺-transporting epithelia, the CCD is thought to have a relatively low efficiency of Na⁺ to ATP coupling. Because the apical entry of Na⁺via the Na⁺ channels is not linked to the transport of other solutes, and the basolateral Na⁺,K⁺-ATPase represents the only significant pathway for Na⁺ exit, the Na⁺/ATP relationship is the same 3:1 ratio as that of the Na⁺,K⁺-ATPase (see Fig. 4-9 ).

Coupling of Na⁺,K⁺-ATPase and Mitochondrial Oxidative Phosphorylation

Na⁺/K⁺/O₂/ATP Stoichiometry

In the early 1960s, Whittam's laboratory^[89] provided experimental evidence for a functional coupling between active Na⁺ and K⁺ transport and respiration in renal cortical slices. In this classic model, ATP hydrolysis by the Na⁺,K⁺-ATPase produces ADP+P_i, which acts as a feedback signal to the mitochondrial for the regulation of ATP synthesis ( Fig. 4-10 ). In cells that rely exclusively on mitochondrial oxidative phosphorylation to support transport, activation of Na⁺ would be exactly coupled to coordinate increases in QO₂ and ATP generation. There is no direct experimental proof that adenine nucleotides are the coupling signal between Na⁺,K⁺-ATPase and mitochondrial oxidative phosphorylation, but there is compelling correlative evidence in the renal tubules. Balaban and colleagues^[59] made simultaneous measurements of the redox state of mitochondrial NAD fluorescence, cellular ATP and ADP concentrations, and QO₂ in renal cortical tubule suspensions (see Fig. 4-10 ). Ouabain caused a net reduction of NAD, a 30% increase in the ATP/ADP ratio, and a 54% inhibition of QO₂. Conversely, activation of Na⁺,K⁺-ATPase by addition of K⁺ to K⁺-depleted tubules promoted a 127% stimulation of QO₂, a large oxidation of NAD, and a 47% decrease in the cellular ATP/ADP ratio. These results suggested that the intracellular concentrations of ATP and ADP represented a cytoplasmic coupling signal in a feedback loop between ATP utilization by the Na⁺,K⁺-ATPase and ATP production by mitochondrial oxidative phosphorylation.



FIGURE 4-10 Whittam model for the coupling of ATP production by mitochondrial oxidative phosphorylation and ATP utilization by the Na⁺,K⁺-ATPase. ATP hydrolysis by the Na⁺,K⁺-ATPase produces ADP+P_i, which serve as a feedback regulator of mitochondrial ATP synthesis, and consequently, QO₂. This model suggests a direct relationship between Na⁺,K⁺-ATPase turnover, mitochondrial ATP synthesis, and QO₂ in cells that rely on mitochondrial ATP production. e^-1, electron; ETC, electron transport chain.

Additional studies examined the Na⁺/K⁺/O₂ stoichiometry in proximal tubule suspensions and their correlation with the known stoichiometries of the Na⁺,K⁺-ATPase and mitochondrial oxidative phosphorylation. Reintroducing K⁺ into a K⁺-depleted suspension of renal proximal tubules to activate the Na⁺,K⁺-ATPase resulted in an initial net K⁺ influx and simultaneous increase in QO₂.^{[53] [90]} In two studies, the K⁺ transport/QO₂ ratios were reported as 10.4 and 11.8,^{[53] [90]} near the expected theoretical value of 12 (two K⁺ transported per ATP hydrolyzed by the Na⁺,K⁺-ATPase and six ATP molecules produced per O₂ by mitochondrial oxidative phosphorylation). Extensions of these studies by different methods, including ²³Na NMR spectroscopy coupled with extracellular K⁺ electrodes^[91] and direct measurements of net Na⁺ and K⁺ fluxes by extracellular ion-sensitive electrodes,^[92] have largely concluded that the Na⁺/K⁺/O₂ in the intact proximal tubule is 18:12:1 as predicted.^[93]

ATP Dependence of Na⁺,K⁺-ATPase Activity

Given the coupling of Na⁺,K⁺-ATPase activity to ATP production, much attention has been directed at determining the ATP dependence of the Na⁺,K⁺-ATPase in the intact kidney, isolated tubules and cells under physiologic and provocative conditions. Different interpretations have arisen depending whether the studies were performed in intact or permeabilized tubules. In permeabilized proximal tubules, with Na⁺,K⁺-ATPase activity studied under V_maxconditions for Na⁺ and K⁺, Soltoff and Mandel^[94] reported that plots of Na⁺,K⁺-ATPase activity as a function of ATP concentration were hyperbolic, with K_0.5 of roughly 0.4 μM for ATP, in the range of values later obtained by others ( Fig. 4-11 ). In contrast, studies in the intact in vitro microperfused proximal tubule showed that a decrement in intracellular ATP content, achieved by pharmacologic inhibition of mitochondrial oxidative phosphorylation, resulted in coordinate reductions in Na⁺-dependent transport of fluid, phosphate, and glucose.^[95] Soltoff and Mandel^[94] extended these studies by examining the ATP dependence of Na⁺,K⁺-ATPase activity, indexed by the initial rate of K⁺ uptake in intact, K⁺-depleted proximal tubules, following graded reductions in intracellular ATP content achieved by serial reductions in the dose of rotenone. Because the Na⁺,K⁺-ATPase turnover is reduced in the K⁺-depleted tubules, the intracellular Na⁺ concentration is elevated and not limiting for subsequent activation of Na⁺,K⁺-ATPase turnover by reintroduction of K⁺. These investigators found that the relationship of Na⁺,K⁺-ATPase activity to ATP concentration was linear, even at ATP concentrations known to saturate the enzyme in in vitro enzymatic assays (see Fig. 4-11 ). The authors postulated that the differences in the data obtained in the permeabilized versus the intact proximal tubule might reflect differences in the local concentrations of ATP, ADP, and P_i in the vicinity for the Na⁺,K⁺-ATPase in the intact tubule compared with permeabilized preparations. In permeabilized tubules, respiration measured in the presence of excess extramitochondrial P_i is dependent on the extramitochondrial concentration of ADP, a relationship with saturable kinetics. The K_m for ADP has been reported to be 15 to 50 μM.^[54] Extrapolated to the intact proximal tubule, these values would saturate the ATP-ADP translocase of the mitochondria and maximally stimulate respiration. Most likely, differences in the measurements of the total cellular concentration of ADP in the cellular extract and the actual free ADP in the cytoplasm likely accounts for this discrepancy.



FIGURE 4-11 ATP dependence of the Na⁺,K⁺-ATPase in permeabilized (solid line) and intact (dashed line) rabbit proximal tubules. See text for discussion. (From Soltoff SP, Mandel LJ: Active ion transport in the renal proximal tubule. III. The ATP dependence of the Na⁺ pump. J Gen Physiol 84:643–662, 1984.)

Similarly, Amman and colleagues^[96] studying dog cortical tubules observed a fixed increment of ouabain-sensitive QO₂ upon stimulation of Na⁺,K⁺-ATPase activity at ATP concentrations ranging from 2 to 7 μM, and phosphorylation potential ([ATP]/[ADP][P_i]) ranging from 1.5 to 7.5 μM^-1. They concluded that the Na⁺/ATP stoichiometry of the Na⁺,K⁺-ATPase remains unmodified by intracellular ATP concentrations in these tubules even when the ATP content equals or exceeds the physiologic value. Taken together, these results suggest a complex coupling between the supply of ATP generated by mitochondrial oxidative phosphorylation and the energetic demands of the Na⁺,K⁺-ATPase and further suggest that the stoichiometry of the Na⁺,K⁺-ATPase is unaltered when ATP concentration or the phosphorylation potential is changed to values greater than or equal to normal. Furthermore, although neither ATP nor phosphorylation potential appears to be rate-limiting for the Na⁺,K⁺-ATPase in the proximal tubule under physiologic conditions, small reductions in these parameters lessen the activity of the Na⁺,K⁺-ATPase.

NAD Redox State

NADH fluorescence can be used to monitor the redox state of the cytoplasm. The ratio [NADH]/[NAD⁺] provides an index of the redox state within the cytoplasm. Inhibition of active transport causes reduction of NAD⁺, whereas increased transport work elicits oxidation of NAD⁺, both occurring as expected from mitochondrial transitions to a lesser or more active state, respectively ( Fig. 4-12 ). Another use of this method is the determination of the relative effectiveness of metabolic substrates to deliver reducing equivalents to the respiratory chain in a particular tissue. Combining QO₂ measurements with optical determinations of the redox level of the mitochondrial respiratory chain permits the distinction between energy utilization by primary transport processes, primary metabolic processes, or both (see Fig. 4-12 ). Balaban and coworkers^[97] applied this method coupled with measurements of QO₂ to study substrate utilization in suspensions of proximal tubules. Short-chain fatty acids increased NADH fluorescence and QO₂ to a greater extent than did carboxylic acids and amino acids. Because NADH fluorescence increased proportionally with QO₂, it was concluded that the substrates increase QO₂ by increasing the delivery of reducing equivalents to NAD and not by direct stimulation of ATP hydrolysis. In another example, glucose removal caused a decrease in QO₂ in proximal tubules in the presence or absence of butyrate, a readily oxidizable fatty acid. This result suggested that these QO₂ changes were related to the transport and not the metabolism of glucose.^[76] The nystatin-stimulated QO₂ was the same in the presence and absence of glucose, and NADH fluorescence showed that glucose addition to tubules suspended in glucose-free medium caused NAD oxidation. Thus, glucose increases respiration by stimulating Na⁺ entry followed by increased Na⁺ pump activity and its associated increase in mitochondrial respiration.



FIGURE 4-12 Mitochondrial responses to physiologic changes. QO₂, oxygen consumption. The indicator in the diamond represents the controlling factor for the mitochondrial response.

High-Resolution Nuclear Magnetic Resonance Spectroscopy

³¹P, ¹⁴N, and ¹H NMR spectroscopy are noninvasive means to measure in real-time renal metabolic pathways and the structure, dynamics, and interactions of biochemical compounds of the nephron in vivo and in vitro.^[98] An important strength of this method is that prior selection of metabolites or substrates is not required. The data obtained are determined principally by the nucleus used for detection (e.g., ³¹P, ¹⁴N, ¹³C, and ¹H). With ³¹P NMR, all manner of inorganic phosphate and high-energy phosphate compounds can be analyzed, including but not limited to ATP, ADP, P_i, and phosphocreatine. Studies of renal metabolism with ³¹P NMR measurements have been applied to the intact kidney in vivo and in vitro, extracts of kidney tissue, and isolated tubules. ³¹P spectra of kidney show strong signals for ATP, P_i, 2-3-diphosphoglycerate (from erythrocytes) but lack an appreciable phosphocreatine signal evident in the heart, brain, and other metabolically active tissues. In addition, ³¹P spectra show relatively little ADP or P_i in the intact kidney. Compared with enzymatic analysis, 100% of ATP, but only 25% of ADP and 27% of P_i, are visible to NMR in the intact kidney.^[99] These results suggest that most of the ADP and P_i is bound within the cell, likely unavailable for biochemical reactions, and invisible by NMR. Thus, of the estimated 0.5 to 1.0 μM ADP and 0.7 μM P_i in the cell, only about 25% of these compounds may be free in the cytoplasm. Consequently, the intracellular phosphorylation potential (i.e., [ATP]/[ADP][Pi]), as judged by ³¹P NMR, is some 10-fold greater than that assayed by biochemical techniques. Biochemical studies estimated a higher total of free cytosolic ADP concentration in the cytosol. Pfaller and colleagues^[100] isolated mitochondrial and cytosolic fractions from proximal tubular suspensions by the digitonin method and analyzed the amount of ATP for each compartment. In parallel experiments, they measured the absolute volumes of mitochondrial and extramitochondrial spaces in similar suspensions utilizing electron microscopic morphometry. When ATP content was related to the morphometrically determined absolute volumes, the ATP concentrations were calculated to be 4.33 mmol/L for the cytosolic and 2.62 mmol/L for the mitochondrial space. Thus, cytosolic and mitochondrial ATP represent 70% and 30% of the total cellular ATP, respectively, in these studies.

Saturation transfer NMR studies in the intact kidney revealed a ADP:O ratio of 2.45 and that the energy cost of Na⁺ transport, calculated from the theoretical Na:ATP of 3.0, exceeded the measured rate of ATP synthesis. Instead, Na:ATP for active transport in the perfused kidney was 12.^[99] ¹³C NMR spectroscopy using ¹³C-labelled substrates has also been used to examine substrate utilization and metabolic fluxes simultaneously through glycolysis, gluconeogenesis, and other cycles. For example, Kinne and coworkers^[101] used ¹³C NMR analysis of IMCD cell extracts to examine carbohydrate biochemistry with this nonradioactive isotope. The advent of magic-angle spinning (MAS) ¹H NMR technology has allowed the analysis of a wider range of biochemical processes in intact tissue samples, without the need for conventional acetonitrile extraction. However, this promising methodology has not been extensively applied to studies of renal transport and metabolism.

ATP-Sensitive Ion Channels and Coupling to Cation Transport

Several laboratories have identified ATP-sensitive K⁺ channels (K-ATP) in renal epithelial cells. These K⁺ channels are defined by their inhibition in response to increased cytosolic ATP concentrations. They couple the metabolic demands of the cell, including those imposed by the demands of the Na⁺,K⁺-ATPase, with the macroscopic K⁺ conductance of the plasma membrane. K-ATP channels are found in a variety of cell types, and their regulation by ATP, other nucleotides, and other molecular mediators varies depending on the roles these channels fulfill. Along the nephron, K-ATP channels are found in the proximal tubule, the TAL, and the CCD.^[102] Structurally, the K-ATP channels in the principal cell of the CCD are heteromultimers composed of an inwardly rectifying K⁺ channel and a sulphonylurea-binding subunit(s). Under basal conditions, these channels exhibit a high open probability and contribute to solute reabsorption and whole body K⁺ homeostasis. Elegant studies in the proximal tubule by Welling and coworkers^[103] found that the macroscopic K⁺ conductance of the basolateral membrane increased upon stimulation of Na⁺,K⁺-ATPase activity and that these responses were accompanied by a fall in intracellular ATP levels (see Fig. 4-9 ). Conversely, intracellular ATP loading uncoupled this response. It has been postulated that this pump-leak coupling via the nucleotide sensitivity of the K-ATP channel ensures that cell membrane potential, intracellular K⁺ activity, and cell volume are defended during physiologic variations in transepithelial ion transport.

The K-ATP channel plays an important role in K⁺ recycling in the TAL and in K⁺ secretion in the collecting duct (see Fig. 4-9 ). A cDNA encoding an inwardly rectifying K-ATP channel (ROMK1, Kir1.1a) was initially isolated from outer medulla of rat kidney by expression cloning.^[104] Since then, multiple alternative splice variants of ROMK have been characterized. In the rat, alternative splicing at the 5′-end gives rise to channel proteins differing at their amino-termini: ROMK2 (Kir1.1b) lacks the first 19 amino acids of ROMK1, and ROMK3 (Kir1.1c) contains a 7-amino acid extension.^[105] These splice variants are differentially expressed along the distal nephron, ROMK1 transcripts are expressed in the CCD and the OMCD, ROMK2 mRNA is highly expressed from the MTAL to the CCD and connecting tubule (CNT), and ROMK3 is expressed in the MTAL, macula densa, and DCT.^[106] ROMK protein has also been immunolocalized to the apical border of the TAL, macula densa, DCT, CNT, CCD principal cells, and OMCD principal cells.^[107] This localization is in agreement with the known distribution of low-conductance K⁺ channels identified in the apical membrane of the distal nephron segments.

More recently, a stretch-activated nonselective cation channel that is inhibited by intracellular ATP (K_i=0.48 μM) has been discovered in the basolateral membrane of the proximal tubule. The channel allows permeation of Ca²⁺ and other cations, and its activity in cell-attached patches is completely inhibited by the venom of the common Chilean tarantula, Grammosola spatulata. The investigators postulated that the channel may be involved in cell volume regulation, intracellular Ca²⁺ homeostasis, and may have increased importance during states of ATP depletion.^[108] Finally, a small-conductance K⁺ channel, with properties similar to those of the K-ATP channel from the plasma membrane, has been characterized in the inner membrane of rat liver and beef heart mitochondria and designated the mitochondrial ATP-regulated K⁺ channel.^[109] The mitochondrial K-ATP channel is inhibited not only by ATP but also, like the plasma membrane K-ATP channel, by sulfonylureas. This channel, which is also expressed in kidney mitochondria, may play an important role in mitochondrial ATP production during physiologic and hypoxic states.^[110] K⁺ transport by this channel results in increased respiration and decreased in the inner mitochondrial membrane potential. Furthermore, diazoxide-triggered activation of ATP-sensitive K⁺ uptake results in decreased ATP hydrolysis through the reverse activity of the F₁F₀-ATPase when respiration is inhibited. Thus, this pathway may have a role in the prevention of mitochondrial ATP hydrolysis.^[111]

ATP (Purinergic) Receptors Modulating Active Na⁺ Transport

In addition to its role as a metabolic fuel, ATP and its metabolites play important roles in autocrine and/or paracrine processes through activation of P2 purinergic receptors in the kidney and most other tissues. Many of these local regulatory functions modulate solute transport in the kidney. P2 receptors are specific membrane-bound receptors sensitive to ATP and uridine triphosphate. Two major subtypes of P2 receptors—ionotropic P2X and metabotropic P2Y receptors—have been characterized. P2X receptors are ligand-gated channels, whereas the P2Y receptors are linked by G proteins to second-messenger systems. RT-PCR studies revealed more intense expression of P2Y(6) receptor mRNA in the proximal tubule and the TAL, less intense expression in the thin descending limb and the CCD and OMCD, and no detectable expression in either the thin ascending limb or the IMCD.^[112] Functional studies revealed luminal P2Y receptors in principal cells of mouse CCD but not in rabbit CCD.^[113] Patch-clamp studies demonstrated that extracellular ATP inhibits the small-conductance K⁺ channel on the apical membrane of the mouse CCD.^[114] Luminal ATP and UTP, presumably acting through P2Y2 receptors, inhibit amiloride-sensitive short-circuit current in perfused mouse CCD.^[115] Activation of P2 receptors also attenuates the inhibitory effect of PTH on Na⁺-dependent phosphate uptake,^[116] Ca²⁺ and Na⁺ absorption, as well as K⁺ secretion. ATP release from epithelial cells onto their luminal aspect, where ecto-nucleotidases promote their metabolism, results in adenosine generation, which may elicit further effects on ion transport, often opposite those of ATP. Moreover, ATP and adenosine may be important autocrine/paracrine mediators of cellular protection and regeneration after ischemic cell damage.^[117] ATP stimulation of purinergic P2Y receptors hydrolyzes anionic phospholipids of the inner leaflet of the plasma membrane, such as phosphatidylinositol-bisphosphates (PIP2), resulting in inhibition of ENaC activity.^[118] Recent studies have also demonstrated that aldosterone promotes ATP release from the basolateral side of target kidney cells, which then acts via a purinergic mechanism to produce contraction of small groups of neighboring epithelial cells and increased transepithelial electrical conductance in a process that involves phosphatidylinositol 3-kinase. It has been hypothesized that this lateral con-taction redistributes cell volume resulting in apical swelling, which, in turn, disrupts the interaction of ENaC with the F-actin cytoskeleton, opening the channel for increased sodium transport.^[119]

Purinergic P2Y(2) receptors are also expressed in mesangial cells and play an important role in the coupling of macular densa signaling to the mesangial cell-afferent arteriolar complex and thus the tubuloglomerular feedback response. Macula densa cells serve as biosensors, detecting changes in luminal NaCl concentration and relaying signals to the mesangial cell-afferent arteriolar complex. Patch-clamp and ATP bioassay studies of macula densa cells revealed a NaCl concentration-sensitive, ATP-permeable large-conductance (380 pS) anion channel that mediated release of ATP (up to 10 μM) across the basolateral membrane. Moreover, ATP released from macula densa cells via this maxi-anion channel acted at mesangial cell purinergic P2Y(2) receptors to provoke increased in intracellular Ca²⁺ concentration.^[120] In vivo studies demonstrated that ATP release occurs over the same range of NaCl concentrations known to effect the tubuloglomerular feedback response and, like the tubuloglomerular feedback response, was increased by dietary salt restriction, suggesting that macula densa ATP release is a signaling component of the tubuloglomerular feedback pathway.^[121]

AMP-activated Protein Kinase Coupling Ion Transport and Metabolism

AMP-activated protein kinase (AMPK) is a heterotrimeric serine/threonine kinase that functions as a sensor of intracellular energy stores, increasing its activity during conditions of metabolic stress as a result of an elevated intracellular AMP/ATP ratio. AMPK responds to alterations in cellular energy stores by adjusting both ATP-consuming and ATP-generating pathways. AMPK activation has been shown to inhibit fatty acid, triglyceride, and sterol synthesis and to stimulate glucose uptake, glycolysis, and fatty acid oxidation in various cell types.^[122] Recent studies indicate that as a metabolic sensor in cells, AMPK may be important in tuning transepithelial transport by CFTR and ENaC to cellular metabolism.^[123]

CFTR is unique among ion channels in requiring ATP hydrolysis for its gating, suggesting the coupling of CFTR activity with cellular energetics. Recent studies by Hallows and associates^{[50] [124] [125]} have shown that AMPK colocalizes with, binds to, and phosphorylates CFTR in colonic and lung epithelial cells and that AMPK phosphorylation of CFTR in vitro inhibits cAMP-activated CFTR conductance in Xenopus oocytes and T84 cells. AMPK has also been shown to inhibit ENaC activity in microinjected Xenopus oocytes expressing mouse ENaC and cultured cortical collecting duct mpkCCD(c14) cells. In contrast to CFTR, the AMPK-ENaC interaction effect appears to be indirect, because AMPK did not bind ENaC by in vivo pull-down assays, nor did it phosphorylate ENaC in vitro.^[126] Moreover, the AMPK-dependent ENaC inhibition appears to be mediated through a decrease in the number of active channels at the plasma membrane rather than a change in open probability of the channel.^[126] AMPK inhibition of ENaC may limit excessive Na⁺ influx under conditions of metabolic stress, thereby limiting Na⁺,K⁺-ATPase turnover and ATP utilization by this pump. It is intriguing to speculate that AMPK may serve to functionally couple the activities of CFTR and ENaC to coordinate transepithelial NaCl transport during states of metabolic stress.

Functional Coupling of Glycolysis with Ion Pumps

Several studies in different renal cell types have suggested a functional coupling of a primary active transport process with ATP generated locally from glycolysis. Paul and coworkers^[127] demonstrated a link between increases in activity of the Na⁺,K⁺-ATPase, consumption of ATP, and stimulation of aerobic glycolysis in vascular smooth muscle and suggested that carbohydrate metabolism is compartmentalized in these cells.^[128] Such coupling was also reported in the renal cell lines A6 and MDCK.^[129] Similarly, functional coupling of glycolytic ATP production to the energy requirements of the Ca²⁺-ATPase of the sarcoplasmic reticulum of skeletal and cardiac muscle^[130] and to the renal V-ATPase^[131] has been reported. Lu and colleagues^[131] established that the glycolytic enzyme aldolase directly binds the V-ATPase E subunit. The two proteins colocalized in the brush border of the proximal tubule, but not in the intercalated cells of the collecting duct. Expression studies in yeast indicated that the interaction with aldolase was necessary for proper association of the V₁ and V₀ domains of the V-ATPase. Further studies demonstrated that aldolase interacts with three different subunits of V-ATPase on distinct interfaces of the glycolytic enzyme and that this interaction increases dramatically in the presence of glucose, suggesting that aldolase may act as a glucose sensor for V-ATPase regulation.^[131] The effects of glucose on V-ATPase-dependent acidification of intracellular compartments and V-ATPase assembly and trafficking are mediated through a phosphatidylinositol 3-kinase-dependent pathway in renal epithelial cells.^[132] The authors postulated that the specific binding of aldolase to the V-ATPase provides a means for the generation of local ATP pools by glycolysis while at the same time regulating the local pH by excreting the H⁺ generated by glycolysis (glycolysis generates two H⁺ for every glucose molecule oxidized and one H⁺ for every ATP molecule produced).^[131] In addition to aldolase, phosphofructokinase-1, the rate-limiting step in glycolysis, has been shown to interact with the C-terminus of the α4 subunit of the V-ATPase and to colocalize in α-intercalated cells, providing another link between the pump and glycolysis.^[133]

RENAL SUBSTRATE PREFERENCES

Whole Kidney and Regional Profiles of Metabolism

By the early 1900s, it was established that the kidney exhibited a high QO₂ compared with other organs and that this was related in large part to Na⁺ reabsorption. In 1928, Gyoergy and coworkers^[134] reported metabolic differences between cortex and medulla. Studies in renal tissue slices later provided evidence for regional differences in intrarenal metabolism and suggested that this distinction may reflect the varying metabolic demands of different nephron segments. In vivo studies measuring carbon-labeled substrate consumption demonstrated the capacity of the kidney to oxidize numerous metabolic substrates. Of these fuels, lactate, glutamine, glucose, free fatty acids, citrate, and ketone bodies are most readily utilized.^{[135] [136]} Cellular respiration provides the majority of ATP needed for energy-dependent transport processes. A smaller proportion of ATP arises from glycolysis and substrate-level oxidation.^[137] These studies, in conjunction with studies in the isolated perfused rat kidney, provided considerable information about the integration of renal metabolism and ion transport. For example, glucose metabolism was found to be intimately involved in several overall transport processes in the isolated perfused kidney, including K⁺ secretion,^[138] HCO₃^- transport, H⁺ secretion,^[139] and active Na⁺ transport,^[137] However, because of the complex distribution of metabolic enzymes along the nephron, disparate metabolic needs and environments of various nephron segments, dicarboxylation reactions unrelated to oxidative metabolism, and methodologic factors, the relative contributions of individual substrates to overall renal metabolism have been difficult to assay.

Studies in renal tissue slices provided important early observations about renal metabolism. The cortex was shown to exhibit a high rate of aerobic metabolism, a low respiratory quotient (CO₂ generated per O₂ consumed), which is indicative of fatty acid oxidation,^[140] gluconeogenesis,^[71] and low glycogen content. In contrast, the medulla is characterized by a high rate of anaerobic or aerobic glycolysis, a respiratory quotient of unity,^[141] which is indicative of glucose oxidation,^[140] glucose consumption resulting in lactate accumulation in the medulla, and a high glycogen content.^[142] However, these studies were limited by the fact that the tissue preparation, with its collapsed tubules and prominent diffusion barriers to O₂ and solutes, did not reflect renal metabolism that occurs in the actively transporting nephron in vivo. Moreover, tissue slices did not allow investigators to attribute specific metabolic pathways to distinct nephron segments. With the development in the 1970s of microprocedures to analyze substrate utilization and enzymatic activities in single microdissected segments of rodents and rabbits, the heterogeneity of renal metabolism among nephron structures was appreciated.

Segmental Profiles of Nephron Metabolism

Substrate Preferences and Dominant Metabolic Pathways

Proximal Tubule.

Proximal tubules have relatively little glycolytic capacity and are thus dependent on aerobic mitochondrial metabolism for ATP synthesis. The intact proximal tubule normally conducts transport work at 50% to 60% of its maximal respiratory capacity, so that substantial reserve capacity is available under physiologic conditions to meet increased ATP utilization. This segment has the ability to oxidize a variety of metabolic substrates, and its prefer-ences for a given substrate are dictated by substrate availability and physiologic state. Experiments using QO₂, ¹⁴CO₂ production from radiolabeled substrates and ATP content in cortical tubule suspensions and microdissected proximal tubule segments revealed that lactate, glutamine, fatty acids, ketone bodies, and triglycerides are avidly oxidized, whereas glucose is poorly oxidized by the proximal tubule.^{[97] [143] [144] [145] [146]} In the rat proximal tubule, the rank order of preferred substrates is ketone bodies>fatty acids>lactate>glutamine ( Fig. 4-13 ). However, some variability in this hierarchy exists among species. The proximal tubule has also significant endogenous fuels, likely neutral lipids, that support about half of the respiratory energy in the absence of exogenous substrates.



FIGURE 4-13 Substrate preferences along the nephron. Summary of preferred substrates to fuel active transport in nephron segments as gleaned primarily from studies using QO₂, ion fluxes, ¹⁴CO₂ generation from ¹⁴C-labeled substrates, ATP contents, and NADH fluorescence. β-OHB, β-hydroxybutyrate.

Uchida and Endou^[146] analyzed the ability of exogenous substrates to support ATP content in individual mouse proximal tubules. ATP production by glucose alone was minimal in the early proximal tubule (S1) but was significant in the late proximal tubule (S3). Based on the support of ATP content, these investigators concluded that glutamine and lactate were the preferred substrates in proximal tubules. Similarly, studies in rabbit cortical tubule suspensions enriched for PST and PCT showed differences in the metabolic responses to glucose. In glucose-containing buffer, PST segments were able to maintain QO₂ and ATP contents at levels significantly higher than PCT segments.^[147]These differential responses between PST and PCT were glucose-dependent and suggested that the PCT cannot utilize glucose to support oxidative metabolism, whereas PST segments can oxidatively metabolize this substrate. Because these differences in glucose utilization do not correlate with the distribution of glycolytic enzyme activities, differential metabolic regulation of these enzymes may determine the ability of each segment to utilize glucose. Glucose cotransported with Na⁺ stimulated ouabain-sensitive QO₂ and NADH oxidation, indicating that it preferentially stimulated ATP utilization.^[148] In contrast, lactate maintains normal ATP content, stimulates QO₂, and increases NADH content in both the PCT and the PST.^{[97] [146]} Substrate-starved proximal tubules demonstrate an increased QO₂ and/or NADH fluorescence when provided TCA cycle intermediates, such as succinate, citrate, and malate, as well as glutamine and acetate.^{[97] [146]} β-hydroxybutyrate and glutamine supported ATP content in all proximal tubule segments.^[146]

Of the various substrates studied, the short-chain fatty acids butyrate, valerate, and heptanoate most dramatically stimulated QO₂ and NADH fluorescence.^[97] Harris and coworkers^[90] demonstrated that butyrate supported, to a greater extent than lactate, glucose, or alanine, the high rates of mitochondrial oxidative phosphorylation needed to sustain maximal rates of Na⁺,K⁺-ATPase activity, provoked by nystatin addition. In another study, addition of fatty acids (butyrate or valerate) or TCA intermediates (succinate or malate) to proximal tubules incubated in lactate, alanine, and glucose resulted in enhanced Na⁺-dependent phosphate transport in the absence of net fluid flux or ATP content.^[149] Butyrate was also shown to enhance the capacity of isolated, nonperfused PSTs to regulate volume under hypo-osmotic conditions by promoting NaCl transport.^[150] Gullans and coworkers^{[76] [144]} showed that succinate stimulated gluconeogenesis, hyperpolarized the plasma membrane potential, and promoted intracellular K⁺ accumulation without altering Na⁺,K⁺-ATPase activity. Thus, the proximal tubule responds to a variety of substrates to support ion transport but has varied metabolic capabilities and substrate preferences along its length.

Thin Limbs of Henle.

Relatively few studies have investigated the metabolic profile of the thin ascending limb of Henle or TDL. These segments have few mitochondria and limited oxidative metabolism. In studies of microdissected TDLs from short- and long-looped nephrons, ATP was depleted when the tubules were incubated in the absence of exogenous substrate at 37°C,^[151] indicating limited endogenous fuel stores. In the presence of exogenous substrates, however, TDLs from long-loop nephrons exhibited two to three times greater ATP contents per mm tubule length than did TDLs from short-loop nephrons. Glucose and pyruvate were the preferred substrates to sustain cellular ATP in TDLs from both short- and long-loop nephrons. ATP production from glutamine, β-hydroxybutyrate, and lactate was significant in TDLs from long-loop nephrons, whereas in TDLs from short-loop nephrons, glutamine was the preferred substrate, and β-hydroxybutyrate and lactate provided minimal metabolic support.^[151] The tubules exhibited a tight coupling of ATP production and active Na⁺ transport. When active Na⁺ transport was stimulated by the ionophore monensin, ATP levels were depleted, and conversely, ouabain inhibition of Na⁺ transport resulted in increased ATP levels.^[151] There have been no published studies profiling the metabolic pathways of the thin ascending limbs of Henle.

CTAL.

¹⁴C-labeled substrate studies demonstrated that glucose and lactate both efficiently generated ¹⁴CO₂ in the CTAL.^[70] Glutamine, glutamate, malate, 2-oxoglutarate, and palmitate were also oxidized, but to a lesser degree.^[152] The ATP content of CTALs was maintained in the presence of glucose, β-hydroxybutyrate, or lactate, but glutamine was ineffective.^[146] The isolated perfused CTAL of the rabbit nephron utilizes glucose and/or pyruvate, acetate, β-hydroxybutyrate, acetoacetate, and butyrate to energize active Na⁺ transport as measured by short-circuit current.^[153] Glutamine, glutamate, citrate, 2-oxoglutarate, and succinate were less effective in supporting maximal rates of transepithelial Na⁺ transport. Substrate removal resulted in a substantial decrease in I_sc over 10 minutes, indicating limited endogenous energy stores. Lactate was produced from glucose under aerobic conditions, and lactate synthesis was greatly enhanced when antimycin A was used to inhibit mitochondrial oxidative phosphorylation.^[154] The CTAL exhibits a tight coupling of active Na⁺ transport and QO₂. Ouabain inhibits about 40% to 50% of the QO₂ in CTALs in the presence of glucose, lactate, and alanine.

Inhibition of active Na⁺ transport with ouabain or furosemide abrogated ¹⁴CO₂ production with ¹⁴C-labeled lactate as substrate, indicating a tight coupling of active Na⁺ transport and lactate oxidation.^[152] In addition, some studies indicate that enzymes active in fatty acid oxidation are active in the CTAL and that CO₂ can be formed from palmitate. Other work, however, demonstrated that proprionate, caprylate, and oleate did not support active Na⁺transport in this segment, suggesting an inability of the CTAL to metabolize odd-chain or long-chain fatty acids.^[155] In the aggregate, these studies indicate that the preferred exogenous substrates to fuel active Na⁺ transport in the CTAL are glucose, lactate, pyruvate, ketone bodies, and fatty acids.

MTAL.

The TAL exhibits the highest rates of active Na⁺ transport among nephron segments, and possesses abundant mitochondria^[156] and a high QO₂ to meet these energy demands.^{[157] [158]} In fact, the QO₂ in the MTAL is nearly 50% greater than that of the proximal tubule.^[159] The MTAL has substantial endogenous energy reserves. In the absence of exogenous substrates, QO₂ is 85% of that achieved in the presence of substrate.^[158] Studies by Eveloff and coworkers,^[157] using pharmacologic inhibitors to block glycolysis, fatty acid oxidation, and amino acid transferase, showed that the endogenous fuels included glycogen, fatty acids, and amino acids. These endogenous fuels were inadequate to support fully nystatin-stimulated Na⁺,K⁺-ATPase activity, but addition of exogenous glucose stimulated QO₂ by nearly 20% and sustained higher rates of nystatin-stimulated QO₂. In the presence of glucose, inhibition of fatty acid oxidation had less inhibitory effect on QO₂. The short-chain fatty acid butyrate as well as acetoacetate and acetate but not lactate, further augmented QO₂ in the presence of glucose. These results suggested that the tubules were substrate-limited in the absence of fatty acids or ketone bodies. Glucose, acetate, malate, and succinate fully supported QO₂ in the MTAL.^[157] Inhibition of salt transport by furosemide or ouabain markedly decreased glucose oxidation.^[160]

Substrate oxidation assessed by measuring ¹⁴CO₂ production from tracer amounts of ¹⁴C-labeled substrates in microdissected MTALs showed that glucose, 2-oxoglutarate, palmitate, lactate, glutamate, and glutamine were utilized as fuels, whereas the TCA cycle intermediates succinate, citrate, and malate were not significantly oxidized.^[70] Leucine, a branched-chain amino acid, is also used as metabolic fuel by this nephron segment, although to a fivefold lower extent than glucose.^[160] In a separate study, glucose, β-hydroxybutyrate, and lactate supported normal ATP content, whereas glutamine was only partially effective in restoring ATP content.

Compared with the proximal tubule, TAL segments exhibit a greater capacity for anaerobic metabolism to support cellular functions but still require mitochondrial respiration for maintenance of active Na⁺ transport. In in vitro microperfusion studies of isolated rabbit CTALs, removal of substrates led to a rapid decline in transepithelial Na⁺ transport, measured as short-circuit current. Addition of glucose from the basolateral side sustained short-circuit current, indicating that the CTAL of rabbit utilizes glucose to energize Na⁺ reabsorption. However, when mitochondrial oxidative phosphorylation was poisoned with cyanide, glucose was only minimally superior to lactate in supporting transport. Studies in both rat^[154] and mice^[146] CTALs demonstrated that inhibition of mitochondrial oxidative phosphorylation resulted in activation of glycolysis, but this was insufficient to maintain normal rates of NaCl transport. The MTAL exhibits a greater capacity for anaerobic glycolysis but, like the CTAL, requires ATP production from mitochondrial oxidative phosphorylation to maintain active Na⁺ transport. In the rat MTAL, lactate generation from glucose is greatly enhanced after chemical inhibition of cellular respiration,^[154] but insufficient to support normal ATP levels.^[146] Chamberlin and Mandel^[158] tested the effects of anoxia on Na⁺,K⁺-ATPase activity, ATP content, extracellular K⁺ release, and QO₂ in suspensions of rabbit MTALs. Under oxygenated conditions, the tubules exhibited efficient coupling between oxidative metabolism (six ATP/O₂) and Na⁺,K⁺-ATPase (two K⁺/ATP). When anoxic, the tubules released K⁺, indicating substrate limitation of Na⁺,K⁺-ATPase activity. However, this rate was accelerated with complete Na⁺,K⁺-ATPase blockade by ouabain, indicating a reserve of Na⁺,K⁺-ATPase activity even in anoxia. Anaerobic metabolism maintained 73% of cellular ATP during 10 min of anoxia, and iodoacetate, an inhibitor of glycolysis, produced a 57% decline in ATP levels and a 33% decline in K⁺ content during anoxia. Thus, glycolysis contributes significant energy during anoxia but is insufficient to maintain the high rates of active transport conducted by this segment.

Distal Convoluted Tubule.

The metabolic profile and substrate preferences of the DCT are not as clearly described as for other nephron segments. Vinay and coworkers^[161] established that this segment is glycolytic. The DCT contains abundant mitochondria, Na⁺, K⁺-ATPase activity,^[162] high ATP levels (as compared with the proximal tubule),^[163] and contains enzymatic activities that would support utilization of glucose, fatty acids, and ketone bodies.^{[68] [135]} In isolated DCTs from the mouse provided glucose, lactate, β-hydroxybutyrate, or L-glutamine as single substrates, lactate and β-hydroxybutyrate were preferred substrates for ATP maintenance. ATP contents were supported at somewhat lower levels by glucose alone, and glutamine did not increase ATP levels over basal conditions.^[146] Bagnasco and coworkers^[154] found that microdissected DCTs produced lactate with glucose as the only substrate and that inhibition of respiration with antimycin A increased lactate production by 98%. A single report of CO₂ production from ¹⁴C-labeled substrates demonstrated glucose oxidation in isolated DCT,^[164] but no data are published concerning QO₂ in this segment.

Cortical Collecting Duct.

Oxidative metabolism provides the majority of the support of cellular ATP content and active Na⁺ transport in this nephron segment. In measurements of metabolic CO₂ production from ¹⁴C-labelled lactate or glucose in microdissected nonperfused tubules, ouabain decreased by more than 50% the CO₂ production by CCD, indicating tight coupling of oxidative metabolism to active Na⁺ transport. Similarly, blockade of Na⁺ entry steps with amiloride reduced the rate of CO₂ production to an extent almost similar to that obtained with ouabain.^[165] Substrate deprivation for 30 min at 37°C produced no change in ATP content of the isolated rat CCD, indicating significant endogenous fuels.^[166] ATP production was greater from glucose than from lactate, β-hydroxybutyrate, or glutamine.^[146] Likewise, in the isolated perfused rabbit CCD, endogenous substrates supported a small component of Na⁺transport.^[167] Inhibition of mitochondrial respiration in the isolated rat CCD with antimycin A caused a significant decrement in cell ATP level within 5 min. Rabbit nonperfused CCDs subjected to hyperosmotic challenge undergo a regulatory volume increase in the presence of extracellular Na⁺ that is supported by butyrate,^[168] suggesting a role for fatty acids in this process, despite the relative low activity of enzymes for fatty acid oxidation present in the rat CCD.^[169]

Nonaka and Stokes^[170] examined the role of metabolism in the support of CCD ion transport using transepithelial electrical measurements and concurrent determination of lumen-to-bath Na⁺ flux. Glucose provided the better support of Na⁺ transport than lactate, pyruvate, glutamine, glutamate, alanine, and several short-chain fatty acids. With glucose absent, near-maximal support of Na⁺ transport was provided by lactate, butyrate hexanoate, or acetate. Hering-Smith and Hamm^[167] assayed lumen-to-bath ²²Na⁺ flux and HCO₃^- in microperfused rabbit CCDs before and after metabolic substrate changes or application of metabolic inhibitors. Both Na⁺ reabsorption, predominantly a principal cell function in the CCD, and HCO₃^- secretion, predominantly an intercalated cell process, were inhibited by antimycin A but were not significantly affected by inhibitors of glycolysis or the hexose-monophosphate shunt pathway.^[167] Basolateral perfusion of glucose and acetate best supported Na⁺ reabsorption, whereas either glucose or acetate fully maintained HCO₃^- secretion. In addition, luminal glucose to some degree supported HCO₃^-secretion, but not Na⁺ transport. The investigators concluded that principal cells and intercalated cells differ not only in their morphology and function but also in their metabolic support of transport.^[167]

OMCD.

The OMCD is a major site of H⁺ secretion along the nephron and is largely responsible for the final acidification of the urine. The OMCD appears to have appreciable endogenous fuels, presumably glycogen, as evidenced by the fact that substrate deprivation for 30 min at 37°C resulted in no change in ATP content of the isolated, nonperfused rat OMCD.^[166] Likewise, HCO₃^- secretion in the isolated perfused rabbit OMCD was fully supported by endogenous substrates.^[167] The segment exhibits has considerable reliance on oxidative metabolism. Addition of cyanide to inhibit mitochondrial oxidative phosphorylation depleted greater than 95% of the ATP content of isolated rabbit OMCD cells in the absence of glucose fuels.^[166] Anaerobic glycolysis can also contribute significantly to cellular energetics. Inhibition of oxidative phosphorylation with antimycin A resulted in an ≈350% increase in lactate production from the isolated rat OMCD.^[154] The OMCD has relatively low rates of active Na⁺ transport and, in the rabbit, exhibits a relatively low level of ouabain-sensitive QO₂. Active Na⁺ transport appears to be tightly coupled to oxidative metabolism. Ouabain decreased by more than 50% the CO₂ production from radiolabeled glucose in isolated, nonperfused rat OMCD, indicating that oxidative metabolism was substantially coupled to active Na⁺transport.^[165]

IMCD.

A major function of the IMCD is the final absorption of about 5% of filtered Na⁺. Studies in isolated rat IMCD cells showed that ouabain-sensitive QO₂ is only 25% to 35%,^[171] and accordingly, ATP turnover is relatively low.^[172]Cellular energetics are largely dependent upon glucose availability under aerobic or anaerobic conditions,^[172] although these cells appear to have considerable endogenous substrates. Stokes and colleagues^[171] found that, in the absence of any exogenous substrate, respiration and ATP contents were near-normal, but lactate production was markedly decreased. Based on ¹³C-NMR analysis of IMCD cell suspensions, enzyme assays on cell homogenates, and enzymatic determination of metabolites and cofactors, the major pathways of glucose metabolism in the IMCD are aerobic and anaerobic glycolysis, the pentose-phosphate shunt, and gluconeogenesis, although the latter two pathways represent minor ones for glucose metabolism in this tissue.^[101] Studies of suspensions of dog IMCDs incubated under aerobic and anaerobic conditions demonstrated that glucose is the preferred substrate for this segment, even if lactate can be oxidized under aerobic conditions.^[172] Glycogen consumption also occurs and to a greater extent during anoxia.^[172]

Under aerobic conditions, the net oxidation of glucose to CO₂ contributes significantly to the cellular energetics. In studies of isolated rat IMCDs, lactate production was at least three times greater than other distal nephron segments.^[154] In another study, aerobic glycolysis accounted for more than 20% of the ATP production in the IMCD.^[171] Cohen postulated that that the high rate of aerobic glycolysis in the presence of an adequate O₂ supply stems from the small mass of mitochondria in relation to the amount of work done by the papillary tissue. The limited ATP synthesis from mitochondrial oxidative phosphorylation shifts both the phosphorylation state ([ATP]/[ADP][P_i]) and the cytoplasmic redox state ([NAD⁺]/[NADH]) of the IMCD cells to a more reduced state, enhancing glycolytic rates and enabling these cells with few mitochondria to sustain substantial active transport in a low O₂ environment.

Given the low density of mitochondria in the IMCD and the low Po₂ it encounters, the IMCD relies to a greater degree on anaerobic glycolysis to sustain normal ion transport rates. Stokes and colleagues^[171] reported that glycolysis increased by 56% and was able to maintain the cellular ATP level at 65% of control values when mitochondrial oxidative phosphorylation was inhibited with rotenone. Similarly, Bagnasco and colleagues^[154] found that antimycin A treatment resulted in a 28% increase in lactate production from glucose. In studies that examined the metabolic determinants of K⁺ transport in the rabbit IMCD, glucose as sole substrate augmented basal QO₂ and cell K⁺ content by about 12% each, whereas iodoacetic acid, an inhibitor of glycolysis, or rotenone, an inhibitor of mitochondrial oxidative phosphorylation, promoted a release of cell K⁺, indicative of substrate limitation of the Na⁺,K⁺-ATPase.^[173]Similarly, in dog IMCD, anoxia resulted in a shift to glycolytic production of ATP, but both the apparent ATP turnover and the activity of the Na⁺, K⁺-ATPase were reduced.^[172] Collectively, these data indicate that both glycolysis and oxidative phosphorylation are required to maintain optimal cellular K⁺ gradients and ATP levels in the IMCD. In addition, Kinne and coworkers^[101] posit that substrate recycling helps to conserve carbohydrate. Based on IMCD cell isolation studies, they propose that sorbitol, taken up by neighboring interstitial cells, is converted into fructose and then recycled to the collecting duct cells. This cycle might represent a beneficial adaptation to low O₂ tension, low substrate supply, and extreme changes in extracellular osmolality in this region.

In summary, a variety of methodologies, both to examine specific metabolic pathways and to define intrarenal heterogeneity of metabolism, have been applied to the study of renal metabolism along the nephron. Differences are evident among cell types and are in large part dictated by the local environment and the requirements of the specific cell type to perform active Na⁺ transport. Though differences in mechanisms for substrate uptake differ among nephron cell populations, substrate availability is not generally rate-limiting under physiologic conditions in the kidney. To date, work has principally focused on the proximal tubule and the IMCD, so much is to be learned about the metabolism in other segments of the nephron. Because most studies have sought to isolate single metabolic pathways and used single substrates to analyze substrate preferences in substrate-starved cells, little is known about true preferences when multiple substrates are present and what factors may govern such preferences. There also appears to be considerable interspecies differences in metabolic profile among nephron segments, and very little is known about nephron heterogeneity of metabolism in humans.

SOLUTE TRANSPORT AND ENERGY AVAILABILITY DURING HYPOXIC OR ISCHEMIC CONDITIONS

Several investigators have explored the ability of isolated nephron segments supplied selected substrates to maintain active transport and ATP levels in the face of environmental or chemical inhibitors of mitochondrial oxidative phosphorylation. These studies sought to determine whether endogenous fuel reserves were sufficient to maintain active transport and ATP content, whether ATP-generation derived from anaerobic glycolysis was enhanced, and whether ATP from glycolytic metabolism could support cellular functions when mitochondrial oxidative respiration was limited. In the rabbit proximal tubule, disparate results have been obtained depending on the experimental model. Glycolysis, measured as lactate production from glucose, was not appreciably evident under aerobic conditions or when mitochondrial respiration was inhibited with antimycin A.^[154] In contrast, Dickman and Mandel,^[174]using different means to inhibit oxidative, namely hypoxia (1% O₂) or inhibition of the respiratory chain with rotenone, showed that suspensions of rabbit proximal tubules can generate lactate and ATP through anaerobic glycolysis to maintain 90% of basal ATP levels. In addition, a differential susceptibility of proximal tubule segments to hypoxia, related at least in part to differences in glycolytic capacity, was discovered: The PCT, with its more limited glycolytic capacity, was more vulnerable to hypoxia than the PST.^[175] Finally, in ischemia, the reduced ATP/ADP ratio would be predicted to increase the open probability of the K-ATP channels independently from pump activity, leading to detrimental imbalance of pump and K⁺ leak.

Weinberg and coworkers^{[176] [177]} conducted a comprehensive analysis of proximal tubule metabolism following hypoxia/reoxygenation. During hypoxia/reoxygenation, the cells developed severe energy deficits, respiratory inhibition, and diminished mitochondrial membrane potential. The decreased respiration persists for substantial periods of time before onset of the mitochondrial permeability transition and/or loss of cytochrome c. Interestingly, there is a high level of resistance to development of complex I dysfunction during hypoxia-reoxygenation in these cells, implicating events upstream of complex I to be important for the energetic deficit.^[177] The function of both the F₁F₀-ATPase and the adenine nucleotide translocase are largely intact, and uncoupling appears to play the principal role in the mitochondrial dysfunction.^[176] Provision of supplements, as substrates for anaerobic ATP generation, during either hypoxia or only during reoxygenation abrogated these abnormalities. Provision of the citric acid cycle metabolites α-ketoglutarate plus malate during either hypoxia or reoxygenation promotes mitochondrial anaerobic metabolism to increase ATP production by substrate-level phosphorylation and energization by anaerobic respiration in electron transport complexes I and II and provide succinate to bypass the complex I block when aerobic metabolism resumes. Accumulation of nonesterified fatty acids appears to underlie the energetic failure of reoxygenated proximal tubules. Moreover, lowering levels of nonesterified fatty acids is a major contributor to the benefit from supplementation with α-ketoglutarate and malate.^[176]

Compared with the proximal tubule, TAL segments exhibit a greater capacity for anaerobic metabolism to support cellular functions but still require mitochondrial respiration for maintenance of active Na⁺ transport. In in vitro microperfusion studies of isolated rabbit CTALs, removal of substrates led to a rapid decline in transepithelial Na⁺ transport, measured as I_sc. Addition of glucose from the basolateral side sustained the I_sc, indicating that the CTAL of rabbit utilizes glucose to energize salt reabsorption. However, when mitochondrial oxidative phosphorylation was poisoned with cyanide, glucose was only minimally superior to lactate in supporting transport.^[153] In studies in both rat^[154] and mice^[146] CTALs, inhibition of mitochondrial oxidative phosphorylation resulted in activation of glycolysis, but this was insufficient to maintain normal rates of NaCl transport. The MTAL exhibits a greater capacity for anaerobic glycolysis but, like the CTAL, requires ATP production from mitochondrial oxidative phosphorylation to maintain active Na⁺ transport. In the rat MTAL, lactate generation from glucose is greatly enhanced after chemical inhibition of cellular respiration^[154] but insufficient to support normal ATP levels.^[146] Chamberlin and Mandel^[158] tested the effects of anoxia on Na⁺,K⁺-ATPase activity, ATP content, extracellular K⁺ release, and QO₂ in suspensions of rabbit MTALs. Under oxygenated conditions, the tubules exhibited efficient coupling between oxidative metabolism (six ATP/O₂) and Na⁺,K⁺-ATPase (two K⁺/ATP). When anoxic, the tubules released K⁺, indicating insufficient Na⁺,K⁺-ATPase activity. However, this rate was accelerated with complete Na⁺,K⁺-ATPase blockade by ouabain, indicating a reserve of Na⁺,K⁺-ATPase activity even in anoxia. Anaerobic metabolism maintained 73% of cellular ATP during 10 min of anoxia, and iodoacetate, an inhibitor of glycolysis, produced a 57% decline in ATP levels and a 33% decline in potassium content during anoxia. Thus, glycolysis contributes significant energy during anoxia but is insufficient to maintain the high rates of active transport conducted by this segment.

Given the fact that requirement the MTAL in vitro requires mitochondrial respiration to maintain normal ATP contents and transport rates, it is somewhat surprising that the MTAL in vivo sustains high active transport rates despite the low O₂ tension of this region. The countercurrent flow of the vasa recta, coupled with the high rates of active solute transport and gradient generation by the MTAL, results in a steep corticomedullary gradient of oxygen, ranging from a Po₂ of 50 mm Hg in the cortex to 10 to 20 mm Hg in the medulla.^[178] The tenuous balance of oxygen supply and demand in the relatively hypoxic renal medulla places this nephron segment at high risk for hypoxic cellular dysfunction and injury. Prasad and coworkers^[179] used the noninvasive technique blood oxygenation level-dependent (BOLD) MRI, a method that exploits deoxygenated blood as an endogenous source of contrast, to demonstrate the dynamics of intrarenal oxygenation in humans. Furosemide, which inhibits active Na⁺ reabsorption and QO₂ in the MTAL, increased medullary Po₂ in healthy young adults, whereas acetazolamide, which principally inhibits solute reabsorption in the proximal tubules, had no effect on medullary Po₂.^[179] Indeed, Brezis and coworkers^[180] suggested that decreased transport workload in the MTAL may help to spare it from injury. In the isolated rat kidney perfused with hypoxic solutions, furosemide-treated kidneys exhibited 86% lower fraction of MTALs showing severe damage compared with vehicle-treated kidneys.

Given the low density of mitochondria in the IMCD and the low Po₂ it encounters, the IMCD relies to a greater degree on anaerobic glycolysis to sustain normal ion transport rates. In IMCD cell suspension from rabbit, glucose augmented basal QO₂ and cell K⁺ content by about 12% each, and iodoacetic acid, an inhibitor of glycolysis, promoted a release of cell K⁺. However, inhibition of mitochondrial oxidative phosphorylation with rotenone demonstrated that glycolysis alone could not maintain cell K⁺ content.^[173] In dog IMCD, anoxia resulted in a shift to glycolytic production of ATP, but both the apparent ATP turnover and the activity of the Na⁺, K⁺-ATPase were reduced.^[172] Thus, both glycolysis and oxidative phosphorylation are required to maintain optimal cellular K⁺ gradients and ATP levels in the IMCD. In addition, Kinne and coworkers^[101] postulated that substrate recycling helps to conserve carbohydrate. Based on IMCD cell isolation studies, they propose that sorbitol, taken up by neighboring interstitial cells, is converted into fructose and then recycled to the collecting duct cells. This cycle might represent a beneficial adaptation to low oxygen tension, low substrate supply, and extreme changes in extracellular osmolality in this region.

EFFECTS OF L-ARGININE METABOLISM ON SOLUTE TRANSPORT AND CELLULAR ENERGETICS

L-arginine is a semi-essential amino acid that is metabolized to several molecules that influence renal function, including NO, L-proline, or polyamines. L-Arginine is the only known substrate for all NO synthase isoforms and, as such, helps to regulate the potent effects of NO on solute transport and metabolism in the kidney.

L-Arginine Metabolism

The uptake, recycling, and degradation of L-arginine modulate NO production. The amino acid is transported into renal tubular epithelial cells by the cationic amino acid transporter (CAT) family of system y(+), proteins. In some cell types, CAT-2 activity is coordinately induced with NO synthase activity to support higher substrate demands of the enzyme. CAT-1 mRNA is expressed predominantly in the collecting ducts and vasa recta in the inner medulla, where L-arginine uptake by this transporter is important in the production of NO and maintenance of blood flow in the renal medulla.^[181] In addition to the importance of L-arginine uptake for NO generation, L-citrulline, which is formed as a byproduct of the NO synthase reaction, can be recycled to L-arginine by consecutive actions of argininosuccinate synthetase and argininosuccinate lyase. Immunolocalization studies in mouse kidney determined that both enzymes are expressed predominantly in the proximal tubule.^[182] In agreement with this work, studies examining the fate of radiolabeled L-citrulline documented substantial generation of radiolabeled L-arginine in the proximal tubule, predominantly in the proximal convoluted tubule.^[183]

In addition to its metabolism via NO synthases, L-arginine can be metabolized by arginase to ornithine and then via ornithine decarboxylase to growth stimulatory polyamines, and by arginine decarboxylase to agmatine following the action of arginine decarboxylase ( Fig. 4-14 ).^[184] These latter pathways effectively decrease intracellular arginine available for NO production. There is likely competition for substrate among the various arginine metabolic pathways, and the products of these enzymes appear to have a regulatory role on the various pathways.^[184] Immunolocalization studies have shown that the arginase II isoenzyme is strongly expressed in the outer stripe of the outer medulla, presumably in the proximal straight tubules, and in a subpopulation of the proximal convoluted tubules in the cortex.^[185] Similarly, within the mouse and rat nephron, ornithine decarboxylase activity is found exclusively in the proximal tubule, primarily in the proximal convoluted tubule.^[186] Filtered ornithine reabsorbed along the PCT may be a major source of ornithine for ornithine decarboxylase.



FIGURE 4-14 Pathways of L-arginine metabolism. ADC, arginine decarboxylase; BH₄, tetrahydrobiopterin, a necessary cofactor for NOS activity; NO, nitric oxide; NOS, NO synthase.

Effects of NO on Renal Solute Transport

In the proximal tubule, NO has been reported to both stimulate and inhibit net fluid and HCO₃^- flux, but only inhibitory effects of NO have been found on the Na⁺/H⁺ exchanger and Na⁺,K⁺-ATPase activity in this segment.^[187] In the MTAL, NO inhibits net Cl^- and HCO₃^- absorption, effects in part mediated by a direct inhibitory action of NO on the Na⁺-K⁺-2Cl^- cotransporter and the Na⁺/H⁺ exchanger.^[187] In contrast, NO stimulates the activity of apical K⁺ channels in this segment.^[188] In the collecting duct, NO inhibits Na⁺ absorption and vasopressin-stimulated osmotic water permeability.^[187] In addition, Lu and coworkers^[189] demonstrated that NO inhibits apical Na⁺channels in the CCD and linked this mechanism to the inhibition of the basolateral small-conductance K⁺ channel. NO has also been reported to inhibit the H⁺-ATPase of intercalated cells of the collecting duct.^[190] In the dog, nonselective NO synthase inhibitors renal increase QO₂ and T_Na+/QO₂. Rats treated with nonselective NO synthase inhibitors exhibited increased T_Na+/QO₂, and these inhibitors also increased QO₂ in proximal tubules in vitro at presumed lower levels of vectorial NaCl transport, suggesting that this effect was not mediated by influences on sodium transport alone. Thus, nonselective NO synthase inhibition increases the oxygen costs of kidney function via angiotensin II-independent mechanisms.^[191]

Effects of NO on Mitochondrial Respiration

Experiments on isolated mitochondria and intact cells have shown that NO plays important roles in regulating mitochondrial QO₂, membrane potential, ATP production, and free radical generation.^[192] Several groups demonstrated that NO potently and reversibly inhibits cytochrome oxidase through interactions with complex IV and S-nitrosation of complex I^[193] with very rapid binding and dissociation kinetics and reduces the affinity of the enzyme for O₂.^[194] In addition, mounting evidence indicates that an NO synthase, recently identified as the full-length neuronal NO synthase isoform with unique post-translational modifications, is expressed in mitochondria and produces NO under physiologic conditions.^[195] Transcripts for this mitochondrial NO synthase were detected in kidney.^[195] Although mitochondria cannot release physiologically relevant levels of NO, they produce biologically active nitrates through arginine-independent mechanisms, which raises the possibility that modulating mitochondrial functions can alter nitrate metabolism. In the aggregate, these findings suggested that NO might serve as a physiologic regulator of cellular respiration. Studies in renal tubules and isolated mitochondria indicated that NO could potently and reversibly inhibit respiration at nanomolar concentrations. There were no differences in sensitivity to NO-mediated inhibition between outer medullary and cortical tubules. The result suggested that, because of its low Po₂, the renal outer medulla may be more vulnerable to hypoxia, not simply because of the low Po₂ as such, but more likely because of the competition between NO and O₂ to control respiration.^[196]

In addition to limiting cellular respiration, the inhibition of cytochrome oxidase by NO also shifts the electron transport chain to a more reduced state, favoring the formation of superoxide anions (O₂^-) at the level of complexes I and III of the electron transport chain. The O₂^- can then be converted by superoxide dismutase into hydrogen peroxide or, depending on intracellular redox conditions, react with NO to form peroxynitrite (ONOO^-).^[197] Both reactive species can produce alteration of solute transport pathways, cell damage, and apoptosis. The attendant depletion of the glutathione pool and enhanced production of ONOO^- in the mitochondria promotes the induction of the permeability transition pore, which collapses the membrane potential and leads to mitochondrial swelling, rupture of the outer mitochondrial membrane, release of pro-apoptotic factors, and apoptosis.^[198]

SUMMARY AND CONCLUSIONS

The heterogeneity of renal transport systems to maintain cellular homeostasis is coupled in a dynamic and interactive manner to a diverse, but highly integrated system of metabolic pathways that effect or influence energy production. A family of ion-translocating ATPases mediates the primary active transport of Na⁺, K⁺, Ca²⁺, and perhaps Cl^-, drives the secondary active transport of other ions and solutes, and consumes the majority of the renal energy supply. To address these demands, kidney epithelial cells have a high capacity for mitochondrial oxidative phosphorylation to generate ATP. Studies in whole kidney, isolated tubules, and renal cells have demonstrated that the energy demands of active Na⁺ transport are coupled to the availability and synthesis of ATP, often indexed by QO₂. The coupling factors between these processes appear to be the cytosolic concentrations of ATP, ADP, and P_i. These adenine nucleotides also influence ATP-sensitive K⁺ channels and link the activity of the Na⁺ pump with the K⁺ leak in at least some nephron segments. There is considerable variability along the nephron among ion transport demand, specific ion transport pathways, metabolic substrate preferences to fuel ion transport, and biochemical pathways and mitochondrial density used to generate ATP and reducing equivalents. In addition to its role as a metabolic fuel, ATP and its metabolites serve as autocrine and/or paracrine regulators of solute transport via activation of P2 purinergic receptors in the kidney. Similarly, the generation of NO from L-arginine affects both specific ion transport mechanisms and mitochondrial respiration. Finally, a wealth of data indicate that the ion transport mechanisms and metabolic processes that support and regulate them along the nephron are subject to complex control by changes in local and systemic environmental conditions (e.g., diet, hormones, drugs, dysfunction of other organs). The extent to which the responses of ion transport and metabolism remain coordinated is largely predictive of successful adaptation to the environmental challenge.

References

1. Axelsen KB, Palmgren MG: Evolution of substrate specificities in the P-type ATPase superfamily. J Mol Evol 1998; 46:84-101.

2. Paulusma CC, Oude Elferink RP: The type 4 subfamily of P-type ATPases, putative aminophospholipid translocases with a role in human disease. Biochim Biophys Acta 2005; 1741:11-24.

3. Gottesman MM, Fojo T, Bates SE: Multidrug resistance in cancer: Role of ATP-dependent transporters. Nat Rev Cancer 2002; 2:48-58.

4. van Helvoort A, Smith AJ, Sprong H, et al: MDR1 P-glycoprotein is a lipid translocase of broad specificity, while MDR3 P-glycoprotein specifically translocates phosphatidylcholine. Cell 1996; 87:507-517.

5. Gatlik-Landwojtowicz E, Aanismaa P, Seelig A: The rate of P-glycoprotein activation depends on the metabolic state of the cell. Biochemistry 2004; 43:14840-14851.

6. Krishnamurthy P, Schuetz JD: Role of ABCG2/BCRP in biology and medicine. Annu Rev Pharmacol Toxicol 2006; 46:381-410.

7. Horisberger JD: Recent insights into the structure and mechanism of the sodium pump. Physiology (Bethesda) 2004; 19:377-387.

8. Jorgensen PL, Hakansson KO, Karlish SJ: Structure and mechanism of Na,K-ATPase: Functional sites and their interactions. Annu Rev Physiol 2003; 65:817-849.

9. Ross B, Leaf A, Silva P, Epstein FH: Na-K-ATPase in sodium transport by the perfused rat kidney. Am J Physiol 1974; 226:624-629.

10. Ahn KY, Madsen KM, Tisher CC, Kone BC: Differential expression and cellular distribution of mRNAs encoding a- and b-isoforms of Na+-K+-ATPase in rat kidney. Am J Physiol 1993; 265:F792-F801.

11. Clapp WL, Bowman P, Shaw GS, et al: Segmental localization of mRNAs encoding Na+-K+-ATPase a- and b-subunit isoforms in rat kidney using RT-PCR. Kidney Int 1994; 46:627-638.

12. Lucking K, Nielsen JM, Pedersen PA, Jorgensen PL: Na-K-ATPase isoform (a3, α2, α1) abundance in rat kidney estimated by competitive RT-PCR and ouabain binding. Am J Physiol 1996; 271:F253-F260.

13. Arystarkhova E, Sweadner KJ: Tissue-specific expression of the Na,K-ATPase b3 subunit. The presence of b3 in lung and liver addresses the problem of the missing subunit. J Biol Chem 1997; 272:22405-22408.

14. Katz AI: Distribution and function of classes of ATPases along the nephron. Kidney Int 1986; 29:21-31.

15. El Mernissi G, Doucet A: Quantitation of [3H]ouabain binding and turnover of Na-K-ATPase along the rabbit nephron. Am J Physiol 1984; 247:F158-F167.

16. Geering K: FXYD proteins: New regulators of Na-K-ATPase. Am J Physiol Renal Physiol 2006; 290:F241-F250.

17. Meij IC, Koenderink JB, van Bokhoven H, et al: Dominant isolated renal magnesium loss is caused by misrouting of the Na+,K+-ATPase g-subunit. Nat Genet 2000; 26:265-266.

18. Aizman R, Asher C, Fuzesi M, et al: Generation and phenotypic analysis of CHIF knockout mice. Am J Physiol Renal Physiol 2002; 283:F277-F569.

19. Shull GE, Lingrel JB: Molecular cloning of the rat stomach (H++K+)-ATPase. J Biol Chem 1986; 261:16788-16791.

20. Crowson MS, Shull GE: Isolation and characterization of a cDNA encoding the putative distal colon H+,K+-ATPase. Similarity of deduced amino acid sequence to gastric H+,K+-ATPase and Na+,K+-ATPase and mRNA expression in distal colon, kidney, and uterus. J Biol Chem 1992; 267:13740-13748.

21. Grishin AV, Sverdlov VE, Kostina MB, Modyanov NN: Cloning and characterization of the entire cDNA encoded by ATP1AL1—A member of the human Na,K/H,K-ATPase gene family. FEBS Lett 1994; 349:144-150.

22. Campbell WG, Weiner ID, Wingo CS, Cain BD: H-K-ATPase in the RCCT-28A rabbit cortical collecting duct cell line. Am J Physiol 1999; 276:F237-F245.

23. Kone BC, Higham SC: A novel N-terminal splice variant of the rat H+-K+-ATPase α2 subunit. Cloning, functional expression, and renal adaptive response to chronic hypokalemia. J Biol Chem 1998; 273:2543-2552.

24. Zhang W, Kuncewicz T, Higham SC, Kone BC: Structure, promoter analysis, and chromosomal localization of the murine H+/K+-ATPase α2 subunit gene. J Am Soc Nephrol 2001; 12:2554-2564.

25. Kone BC: Renal H,K-ATPase: Structure, function and regulation. Miner Electrolyte Metab 1996; 22:349-365.

26. Ahn KY, Turner PB, Madsen KM, Kone BC: Effects of chronic hypokalemia on renal expression of the “gastric” H+-K+-ATPase α-subunit gene. Am J Physiol 1996; 270:F557-F566.

27. Codina J, Kone BC, Delmas-Mata JT, DuBose Jr TD: Functional expression of the colonic H+,K+-ATPase α-subunit. Pharmacologic properties and assembly with X+,K+-ATPase b-subunits. J Biol Chem 1996; 271:29759-29763.

28. Ahn KY, Park KY, Kim KK, Kone BC: Chronic hypokalemia enhances expression of the H+-K+-ATPase α2-subunit gene in renal medulla. Am J Physiol 1996; 271:F314-F321.

29. Ramachandran C, Chan M, Brunette MG: Characterization of ATP-dependent Ca2+ transport in the basolateral membrane vesicles from proximal and distal tubules of the rabbit kidney. Biochem Cell Biol 1991; 69:109-114.

30. Strehler EE, Zacharias DA: Role of alternative splicing in generating isoform diversity among plasma membrane calcium pumps. Physiol Rev 2001; 81:21-50.

31. Doucet A, Katz AI: High-affinity Ca-Mg-ATPase along the rabbit nephron. Am J Physiol 1982; 242:F346-F352.

32. Magyar CE, White KE, Rojas R, et al: Plasma membrane Ca2+-ATPase and NCX1 Na+/Ca2+ exchanger expression in distal convoluted tubule cells. Am J Physiol Renal Physiol 2002; 283:F29-F40.

33. Magosci M, Yamaki M, Penniston JT, Dousa TP: Localization of mRNAs coding for isozymes of plasma membrane Ca2+-ATPase pump in rat kidney. Am J Physiol 1992; 263:F7-F14.

34. Caride AJ, Chini EN, Homma S, et al: mRNA encoding four isoforms of the plasma membrane calcium pump and their variants in rat kidney and nephron segments. J Lab Clin Med 1998; 132:149-156.

35. Martin V, Bredoux R, Corvazier E, et al: Three novel sarco/endoplasmic reticulum Ca2+-ATPase (SERCA) 3 isoforms. Expression, regulation, and function of the membranes of the SERCA3 family. J Biol Chem 2002; 277:24442-24452.

36. Greenough M, Pase L, Voskoboinik I, et al: Signals regulating trafficking of Menkes (MNK; ATP7A) copper-translocating P-type ATPase in polarized MDCK cells. Am J Physiol Cell Physiol 2004; 287:C1463-C1471.

37. Moore SD, Cox DW: Expression in mouse kidney of membrane copper transporters ATP7a and ATP7b. Nephron 2002; 92:629-634.

38. Wagner CA, Finberg KE, Breton S, et al: Renal vacuolar H+-ATPase. Physiol Rev 2004; 84:1263-1314.

39. Borthwick KJ, Karet FE: Inherited disorders of the H+-ATPase. Curr Opin Nephrol Hypertens 2002; 11:563-568.

40. Smith AJ, van Helvoort A, van Meer G, et al: MDR3 P-glycoprotein, a phosphatidylcholine translocase, transports several cytotoxic drugs and directly interacts with drugs as judged by interference with nucleotide trapping. J Biol Chem 2000; 275:23530-23539.

41. Ernest S, Rajaraman S, Megyesi J, Bello-Reuss EN: Expression of MDR1 (multidrug resistance) gene and its protein in normal human kidney. Nephron 1997; 77:284-289.

42. Bello-Reuss E, Ernest S: Expression and function of P-glycoprotein in human mesangial cells. Am J Physiol 1994; 267:C1351-C1358.

43. Schaub TP, Kartenbeck J, Konig J, et al: Expression of the MRP2 gene-encoded conjugate export pump in human kidney proximal tubules and in renal cell carcinoma. J Am Soc Nephrol 1999; 10:1159-1169.

44. Stanton BA: Cystic fibrosis transmembrane conductance regulator (CFTR) and renal function. Wien Klin Wochenschr 1997; 109:457-464.

45. Cantiello HF: Electrodiffusional ATP movement through CFTR and other ABC transporters. Pflugers Arch 2001; 443(suppl 1):S22-S27.

46. Sugita M, Yue Y, Foskett JK: CFTR Cl- channel and CFTR-associated ATP channel: Distinct pores regulated by common gates. Embo J 1998; 17:898-908.

47. Ismailov II, Awayda MS, Jovov B, et al: Regulation of epithelial sodium channels by the cystic fibrosis transmembrane conductance regulator. J Biol Chem 1996; 271:4725-4732.

48. Ruknudin A, Schulze DH, Sullivan SK, et al: Novel subunit composition of a renal epithelial KATP channel. J Biol Chem 1998; 273:14165-14171.

49. Braunstein GM, Roman RM, Clancy JP, et al: Cystic fibrosis transmembrane conductance regulator facilitates ATP release by stimulating a separate ATP release channel for autocrine control of cell volume regulation. J Biol Chem 2001; 276:6621-6630.

50. Hallows KR, Raghuram V, Kemp BE, et al: Inhibition of cystic fibrosis transmembrane conductance regulator by novel interaction with the metabolic sensor AMP-activated protein kinase. J Clin Invest 2000; 105:1711-1721.

51. Hinkle PC: P/O ratios of mitochondrial oxidative phosphorylation. Biochim Biophys Acta 2005; 1706:1-11.

52. Mitchell P, Moyle J: Estimation of membrane potential and pH difference across the cristae membrane of rat liver mitochondria. Eur J Biochem 1969; 7:471-484.

53. Harris SI, Balaban RS, Mandel LJ: Oxygen consumption and cellular ion transport: Evidence for adenosine triphosphate to O2 ratio near 6 in intact cell. Science 1980; 208:1148-1150.

54. Jacobus WE, Moreadith RW, Vandegaer KM: Mitochondrial respiratory control. Evidence against the regulation of respiration by extramitochondrial phosphorylation potentials or by [ATP]/[ADP] ratios. J Biol Chem 1982; 257:2397-2402.

55. Lemasters JJ, Sowers AE: Phosphate dependence and atractyloside inhibition of mitochondrial oxidative phosphorylation. The ADP-ATP carrier is rate-limiting. J Biol Chem 1979; 254:1248-1251.

56. Erecinska M, Wilson DF: Regulation of cellular energy metabolism. J Membr Biol 1982; 70:1-14.

57. Noel J, Vinay P, Tejedor A, et al: Metabolic cost of bafilomycin-sensitive H+ pump in intact dog, rabbit, and hamster proximal tubules. Am J Physiol 1993; 264:F655-F661.

58. Harris SI, Balaban RS, Barrett L, Mandel LJ: Mitochondrial respiratory capacity and Na+- and K+-dependent adenosine triphosphatase-mediated ion transport in the intact renal cell. J Biol Chem 1981; 256:10319-10328.

59. Balaban RS, Mandel LJ, Soltoff SP, Storey JM: Coupling of active ion transport and aerobic respiratory rate in isolated renal tubules. Proc Natl Acad Sci U S A 1980; 77:447-451.

60. Niaudet P: Mitochondrial disorders and the kidney. Arch Dis Child 1998; 78:387-390.

61. Rotig A, Appelkvist EL, Geromel V, et al: Quinone-responsive multiple respiratory-chain dysfunction due to widespread coenzyme Q10 deficiency. Lancet 2000; 356:391-395.

62. Guder WG, Schmidt U: The localization of gluconeogenesis in rat nephron. Determination of phosphoenolpyruvate carboxykinase in microdissected tubules. Hoppe Seylers Z Physiol Chem 1974; 355:273-278.

63. Burch HB, Narins RG, Chu C, et al: Distribution along the rat nephron of three enzymes of gluconeogenesis in acidosis and starvation. Am J Physiol 1978; 235:F246-F253.

64. Vandewalle A, Wirthensohn G, Heidrich HG, Guder WG: Distribution of hexokinase and phosphoenolpyruvate carboxykinase along the rabbit nephron. Am J Physiol 1981; 240:F492-F500.

65. Yanez AJ, Ludwig HC, Bertinat R, et al: Different involvement for aldolase isoenzymes in kidney glucose metabolism: Aldolase B but not aldolase A colocalizes and forms a complex with FBPase. J Cell Physiol 2005; 202:743-753.

66. Le Hir M, Dubach UC: Activities of enzymes of the tricarboxylic acid cycle in segments of the rat nephron. Pflugers Arch 1982; 395:239-243.

67. Weidemann MJ, Krebs HA: The fuel of respiration of rat kidney cortex. Biochem J 1969; 112:149-166.

68. Guder WG, Purschel S, Wirthensohn G: Renal ketone body metabolism. Distribution of 3-oxoacid CoA-transferase and 3-hydroxybutyrate dehydrogenase along the mouse nephron. Hoppe Seylers Z Physiol Chem 1983; 364:1727-1737.

69. Guder WG, Ross BD: Enzyme distribution along the nephron. Kidney Int 1984; 26:101-111.

70. Klein KI, Wang MS, Torikai S, et al: Substrate oxidation by defined single nephron segments of rat kidney. Int J Biochem 1980; 12:53-54.

71. Benoy MP, Elliot KAC: The metabolism of lactic and pyruvic acids in normal and tubule tissue. V. Synthesis of carbohydrate. Biochem J 1937; 31:1268-1275.

72. Krebs HA: Renal gluconeogenesis. Adv Enzyme Regul 1963; 1:385-400.

73. Meyer C, Stumvoll M, Dostou J, et al: Renal substrate exchange and gluconeogenesis in normal postabsorptive humans. Am J Physiol Endocrinol Metab 2002; 282:E428-E434.

74. Conjard A, Martin M, Guitton J, et al: Gluconeogenesis from glutamine and lactate in the isolated human renal proximal tubule: Longitudinal heterogeneity and lack of response to adrenaline. Biochem J 2001; 360:371-377.

75. Maleque A, Endou H, Koseki C, Sakai F: Nephron heterogeneity: Gluconeogenesis from pyruvate in rabbit nephron. FEBS Lett 1980; 116:154-156.

76. Gullans SR, Brazy PC, Dennis VW, Mandel LJ: Interactions between gluconeogenesis and sodium transport in rabbit proximal tubule. Am J Physiol 1984; 246:F859-F869.

77. Nagami GT, Lee P: Effect of luminal perfusion on glucose production by isolated proximal tubules. Am J Physiol 1989; 256:F120-F127.

78. Fulgraff G, Nunemann H, Sudhoff DE: Effects of the diuretics furosemide, ethacrynic acid, and chlorothiazide on gluconeogenesis from various substrates in rat kidney cortex slices. Naunyn Schmiedebergs Arch Pharmacol 1972; 273:86-98.

79. Silva P, Hallac R, Spokes K, Epstein FH: Relationship among gluconeogenesis, QO2, and Na+ transport in the perfused rat kidney. Am J Physiol 1982; 242:F508-F513.

80. Kiil F, Aukland K, Refsum HE: Renal sodium transport and oxygen consumption. Am J Physiol 1961; 201:511-526.

81. Sen AK, Post RL: Stoichiometry and localization of adenosine triphosphate-dependent sodium and potassium transport in the erythrocyte. J Biol Chem 1964; 239:345-352.

82. Mathisen O, Monclair T, Kiil F: Oxygen requirement of bicarbonate-dependent sodium reabsorption in the dog kidney. Am J Physiol 1980; 238:F175-F180.

83. Ostensen J, Stokke ES, Hartmann A, et al: Low oxygen cost of carbonic anhydrase-dependent sodium reabsorption in the dog kidney. Acta Physiol Scand 1989; 137:189-198.

84. Fromter E, Rumrich G, Ullrich KJ: Phenomenologic description of Na+, Cl-, and HCO3- absorption from proximal tubules of the rat kidney. Pflugers Arch 1973; 343:189-220.

85. Kiil F, Sejersted OM, Steen PA: Energetics and specificity of transcellular NaCl transport in the dog kidney. Int J Biochem 1980; 12:245-250.

86. Gross E, Hawkins K, Abuladze N, et al: The stoichiometry of the electrogenic sodium bicarbonate cotransporter NBC1 is cell-type dependent. J Physiol 2001; 531:597-603.

87. Silva P, Myers MA: Stoichiometry of sodium chloride transport by rectal gland of Squalus acanthias.. Am J Physiol 1986; 250:F516-F519.

88. Welsh MJ: Energetics of chloride secretion in canine tracheal epithelium. Comparison of the metabolic cost of chloride transport with the metabolic cost of sodium transport. J Clin Invest 1984; 74:262-268.

89. Whittam R: Active cation transport as a pacemaker of respiration. Nature 1961; 191:603-604.

90. Harris SI, Patton L, Barrett L, Mandel LJ: (Na+,K+)-ATPase kinetics within the intact renal cell. The role of oxidative metabolism. J Biol Chem 1982; 257:6996-7002.

91. Avison MJ, Gullans SR, Ogino T, et al: Measurement of Na+-K+ coupling ratio of Na+-K+-ATPase in rabbit proximal tubules. Am J Physiol 1987; 253:C126-C136.

92. Brady HR, Kone BC, Gullans SR: Extracellular Na+ electrode for monitoring net Na+ flux in cell suspensions. Am J Physiol 1989; 256:C1105-C1110.

93. Blostein R, Harvey WJ: Na+, K+-pump stoichiometry and coupling in inside-out vesicles from red blood cell membranes. Methods Enzymol 1989; 173:377-380.

94. Soltoff SP, Mandel LJ: Active ion transport in the renal proximal tubule. III. The ATP dependence of the Na pump. J Gen Physiol 1984; 84:643-662.

95. Gullans SR, Brazy PC, Soltoff SP, et al: Metabolic inhibitors: Effects on metabolism and transport in the proximal tubule. Am J Physiol 1982; 243:F133-F140.

96. Ammann H, Noel J, Boulanger Y, Vinay P: Relationship between intracellular ATP and the sodium pump activity in dog renal tubules. Can J Physiol Pharmacol 1990; 68:57-67.

97. Balaban RS, Mandel LJ: Metabolic substrate utilization by rabbit proximal tubule. An NADH fluorescence study. Am J Physiol 1988; 254:F407-F416.

98. Foxall PJ, Nicholson JK: Nuclear magnetic resonance spectroscopy: A non-invasive probe of kidney metabolism and function. Exp Nephrol 1998; 6:409-414.

99. Freeman D, Bartlett S, Radda G, Ross B: Energetics of sodium transport in the kidney. Saturation transfer 31P-NMR. Biochim Biophys Acta 1983; 762:325-336.

100. Pfaller W, Guder WG, Gstraunthaler G, et al: Compartmentation of ATP within renal proximal tubular cells. Biochim Biophys Acta 1984; 805:152-157.

101. Kinne RK, Grunewald RW, Ruhfus B, Kinne-Saffran E: Biochemistry and physiology of carbohydrates in the renal collecting duct. J Exp Zool 1997; 279:436-442.

102. Quast U: ATP-sensitive K+ channels in the kidney. Naunyn Schmiedebergs Arch Pharmacol 1996; 354:213-225.

103. Tsuchiya K, Wang W, Giebisch G, Welling PA: ATP is a coupling modulator of parallel Na,K-ATPase-K-channel activity in the renal proximal tubule. Proc Natl Acad Sci U S A 1992; 89:6418-6422.

104. Ho K, Nichols CG, Lederer WJ, et al: Cloning and expression of an inwardly rectifying ATP-regulated potassium channel. Nature 1993; 362:31-38.

105. Boim MA, Ho K, Shuck ME, et al: ROMK inwardly rectifying ATP-sensitive K+ channel. II. Cloning and distribution of alternative forms. Am J Physiol 1995; 268:F1132-F1140.

106. Lee WS, Hebert SC: ROMK inwardly rectifying ATP-sensitive K+ channel. I. Expression in rat distal nephron segments. Am J Physiol 1995; 268:F1124-F1131.

107. Kohda Y, Ding W, Phan E, et al: Localization of the ROMK potassium channel to the apical membrane of distal nephron in rat kidney. Kidney Int 1998; 54:1214-1223.

108. Hurwitz CG, Hu VY, Segal AS: A mechanogated nonselective cation channel in proximal tubule that is ATP sensitive. Am J Physiol Renal Physiol 2002; 283:F93-F104.

109. Paucek P, Mironova G, Mahdi F, et al: Reconstitution and partial purification of the glibenclamide-sensitive, ATP-dependent K+ channel from rat liver and beef heart mitochondria. J Biol Chem 1992; 267:26062-26069.

110. Garlid KD, Paucek P: The mitochondrial potassium cycle. IUBMB Life 2001; 52:153-158.

111. Cancherini DV, Trabuco LG, Reboucas NA, Kowaltowski AJ: ATP-sensitive K+ channels in renal mitochondria. Am J Physiol Renal Physiol 2003; 285:F1291-F1296.

112. Bailey MA, Imbert-Teboul M, Turner C, et al: Evidence for basolateral P2Y(6) receptors along the rat proximal tubule: Functional and molecular characterization. J Am Soc Nephrol 2001; 12:1640-1647.

113. Deetjen P, Thomas J, Lehrmann H, et al: The luminal P2Y receptor in the isolated perfused mouse cortical collecting duct. J Am Soc Nephrol 2000; 11:1798-1806.

114. Lu M, MacGregor GG, Wang W, Giebisch G: Extracellular ATP inhibits the small-conductance K channel on the apical membrane of the cortical collecting duct from mouse kidney. J Gen Physiol 2000; 116:299-310.

115. Lehrmann H, Thomas J, Kim SJ, et al: Luminal P2Y2 receptor-mediated inhibition of Na+ absorption in isolated perfused mouse CCD. J Am Soc Nephrol 2002; 13:10-18.

116. Lederer ED, McLeish KR: P2 purinoceptor stimulation attenuates PTH inhibition of phosphate uptake by a G protein-dependent mechanism. Am J Physiol 1995; 269:F309-F316.

117. Leipziger J: Control of epithelial transport via luminal P2 receptors. Am J Physiol 2003; 284:F419-F432.

118. Ishikawa T, Jiang C, Stutts MJ, et al: Regulation of the epithelial Na+ channel by cytosolic ATP. J Biol Chem 2003; 278:38276-38286.

119. Gorelik J, Zhang Y, Sanchez D, et al: Aldosterone acts via an ATP autocrine/paracrine system: The Edelman ATP hypothesis revisited. Proc Natl Acad Sci U S A 2005; 102:15000-15005.

120. Bell PD, Lapointe JY, Sabirov R, et al: Macula densa cell signaling involves ATP release through a maxi anion channel. Proc Natl Acad Sci U S A 2003; 100:4322-4327.

121. Komlosi P, Peti-Peterdi J, Fuson AL, et al: Macula densa basolateral ATP release is regulated by luminal [NaCl] and dietary salt intake. Am J Physiol Renal Physiol 2004; 286:F1054-F1058.

122. Carling D: AMP-activated protein kinase: Balancing the scales. Biochimie 2005; 87:87-91.

123. Hallows KR: Emerging role of AMP-activated protein kinase in coupling mem-brane transport to cellular metabolism. Curr Opin Nephrol Hypertens 2005; 14:464-471.

124. Hallows KR, Kobinger GP, Wilson JM, et al: Physiological modulation of CFTR activity by AMP-activated protein kinase in polarized T84 cells. Am J Physiol Cell Physiol 2003; 284:C1297-C1308.

125. Hallows KR, McCane JE, Kemp BE, et al: Regulation of channel gating by AMP-activated protein kinase modulates cystic fibrosis transmembrane conductance regulator activity in lung submucosal cells. J Biol Chem 2003; 278:998-1004.

126. Carattino MD, Edinger RS, Grieser HJ, et al: Epithelial sodium channel inhibition by AMP-activated protein kinase in oocytes and polarized renal epithelial cells. J Biol Chem 2005; 280:17608-17616.

127. Paul RJ, Bauer M, Pease W: Vascular smooth muscle: Aerobic glycolysis linked to sodium and potassium transport processes. Science 1979; 206:1414-1416.

128. Lynch RM, Paul RJ: Compartmentation of glycolytic and glycogenolytic metabolism in vascular smooth muscle. Science 1983; 222:1344-1346.

129. Lynch RM, Balaban RS: Coupling of aerobic glycolysis and Na+-K+-ATPase in renal cell line MDCK. Am J Physiol 1987; 253:C269-C276.

130. Xu KY, Zweier JL, Becker LC: Functional coupling between glycolysis and sarcoplasmic reticulum Ca2+ transport. Circ Res 1995; 77:88-97.

131. Lu M, Sautin YY, Holliday LS, Gluck S: The glycolytic enzyme aldolase mediates assembly, expression, and activity of vacuolar H+-ATPase. J Biol Chem 2004; 279:8732-8739.

132. Sautin YY, Lu M, Gaugler A, et al: Phosphatidylinositol 3-kinase-mediated effects of glucose on vacuolar H+-ATPase assembly, translocation, and acidification of intracellular compartments in renal epithelial cells. Mol Cell Biol 2005; 25:575-589.

133. Su Y, Zhou A, Al-Lamki RS, Karet FE: The α-subunit of the V-type H+-ATPase interacts with phosphofructokinase-1 in humans. J Biol Chem 2003; 278:20013-20018.

134. Gyoergy P, Keller W, Brehme TH: Nierenstoffwechsel und Nierenentwicklung. Biochem Zeitschr 1928; 200:356-366.

135. Wirthensohn G, Guder WG: Renal substrate metabolism. Physiol Rev 1986; 66:469-497.

136. Guder WG, Wagner S, Wirthensohn G: Metabolic fuels along the nephron: Pathways and intracellular mechanisms of interaction. Kidney Int 1986; 29:41-45.

137. Ross BD, Epstein FH, Leaf A: Sodium reabsorption in the perfused rat kidney. Am J Physiol 1973; 225:1165-1171.

138. Silva P, Ross BD, Charney AN, et al: Potassium transport by the isolated perfused kidney. J Clin Invest 1975; 56:862-869.

139. Besarab A, Silva P, Ross B, Epstein FH: Bicarbonate and sodium reabsorption by the isolated perfused kidney. Am J Physiol 1975; 228:1525-1530.

140. Dickens F, Simer F: The respiratory quotient and the relationship of respiration to glycolysis. Biochem J 1930; 24:1301-1326.

141. Lee JB, Peter HM: Effect of oxygen tension on glucose metabolism in rabbit kidney cortex and medulla. Am J Physiol 1969; 217:1464-1471.

142. Cohen JJ: Is the function of the renal papilla coupled exclusively to an anaerobic pattern of metabolism?. Am J Physiol 1979; 236:F423-F433.

143. Gullans SR, Brazy PC, Mandel LJ, Dennis VW: Stimulation of phosphate transport in the proximal tubule by metabolic substrates. Am J Physiol 1984; 247:F582-F587.

144. Gullans SR, Kone BC, Avison MJ, Giebisch G: Succinate alters respiration, membrane potential, and intracellular K+ in proximal tubule. Am J Physiol 1988; 255:F1170-F1177.

145. Soltoff SP, Mandel LJ: Active ion transport in the renal proximal tubule. I. Transport and metabolic studies. J Gen Physiol 1984; 84:601-622.

146. Uchida S, Endou H: Substrate specificity to maintain cellular ATP along the mouse nephron. Am J Physiol 1988; 255:F977-F983.

147. Ruegg CE, Mandel LJ: Bulk isolation of renal PCT and PST. I. Glucose-dependent metabolic differences. Am J Physiol 1990; 259:F164-F175.

148. Gullans SR, Harris SI, Mandel LJ: Glucose-dependent respiration in suspensions of rabbit cortical tubules. J Membr Biol 1984; 78:257-262.

149. Mandel LJ: Use of noninvasive fluorometry and spectrophotometry to study epithelial metabolism and transport. Fed Proc 1982; 41:36-41.

150. Rome L, Grantham J, Savin V, et al: Proximal tubule volume regulation in hyperosmotic media: Intracellular K+, Na+, and Cl. Am J Physiol 1989; 257:C1093-C1100.

151. Jung KY, Endou H: Cellular adenosine triphosphate production and consumption in the descending thin limb of Henle's loop in the rat. Ren Physiol Biochem 1990; 13:248-258.

152. LeBouffant F, Hus-Citharel A, Morel F: vitro 14CO2 production by single pieces of cortical thick ascending limbs and its coupling to active salt transport. In: Morel F, ed. Biochemistry of Kidney Functions, Amsterdam: Elsevier Biomedical Press; 1982:363-370.INSERM Symposium 21

153. Wittner M, Weidtke C, Schlatter E, et al: Substrate utilization in the isolated perfused cortical thick ascending limb of rabbit nephron. Pflugers Arch 1984; 402:52-62.

154. Bagnasco S, Good D, Balaban R, Burg M: Lactate production in isolated segments of the rat nephron. Am J Physiol 1985; 248:F522-F526.

155. Guder WG, Wirthensohn G: Metabolism of isolated kidney tubules. Interactions between lactate, glutamine and oleate metabolism. Eur J Biochem 1979; 99:577-584.

156. Kone BC, Madsen KM, Tisher CC: Ultrastructure of the thick ascending limb of Henle in the rat kidney. Am J Anat 1984; 171:217-226.

157. Eveloff J, Bayerdorffer E, Silva P, Kinne R: Sodium-chloride transport in the thick ascending limb of Henle's loop. Oxygen consumption studies in isolated cells. Pflugers Arch 1981; 389:263-270.

158. Chamberlin ME, Mandel LJ: Na+-K+-ATPase activity in medullary thick ascending limb during short-term anoxia. Am J Physiol 1987; 252:F838-F843.

159. Mandel LJ: Primary active sodium transport, oxygen consumption, and ATP: Coupling and regulation. Kidney Int 1986; 29:3-9.

160. Trinh-Trang-Tan MM, Levillain O, Bankir L: Contribution of leucine to oxidative metabolism of the rat medullary thick ascending limb. Pflugers Arch 1988; 411:676-680.

161. Vinay P, Gougoux A, Lemieux G: Isolation of a pure suspension of rat proximal tubules. Am J Physiol 1981; 241:F403-F411.

162. Katz AI, Doucet A, Morel F: Na-K-ATPase activity along the rabbit, rat, and mouse nephron. Am J Physiol 1979; 237:F114-F120.

163. Kiebzak GM, Yusufi AN, Kusano E, et al: ATP and cAMP system in the in vitro response of microdissected cortical tubules to PTH. Am J Physiol 1985; 248:F152-F159.

164. Le Bouffant F, Hus-Citharel A, Morel F: Metabolic CO2 production by isolated single pieces of rat distal nephron segments. Pflugers Arch 1984; 401:346-353.

165. Hus-Citharel A, Morel F: Coupling of metabolic CO2 production to ion transport in isolated rat thick ascending limbs and collecting tubules. Pflugers Arch 1986; 407:421-427.

166. Torikai S: Dependency of microdissected nephron segments upon oxidative phosphorylation and exogenous substrates: A relationship between tubular anatomical location in the kidney and metabolic activity. Clin Sci (Lond) 1989; 77:287-295.

167. Hering-Smith KS, Hamm LL: Metabolic support of collecting duct transport. Kidney Int 1998; 53:408-415.

168. Natke Jr E: Cell volume regulation of rabbit cortical collecting tubule in anisotonic media. Am J Physiol 1990; 258:F1657-F1665.

169. Le Hir M, Dubach UC: Peroxisomal and mitochondrial beta-oxidation in the rat kidney: Distribution of fatty acyl-coenzyme A oxidase and 3-hydroxyacyl-coenzyme A dehydrogenase activities along the nephron. J Histochem Cytochem 1982; 30:441-444.

170. Nonaka T, Stokes JB: Metabolic support of Na+ transport by the rabbit CCD: Analysis of the use of equivalent current. Kidney Int 1994; 45:743-752.

171. Stokes JB, Grupp C, Kinne RK: Purification of rat papillary collecting duct cells: Functional and metabolic assessment. Am J Physiol 1987; 253:F251-F262.

172. Meury L, Noel J, Tejedor A, et al: Glucose metabolism in dog inner medullary collecting ducts. Ren Physiol Biochem 1994; 17:246-266.

173. Kone BC, Kikeri D, Zeidel ML, Gullans SR: Cellular pathways of potassium transport in renal inner medullary collecting duct. Am J Physiol 1989; 256:C823-C830.

174. Dickman KG, Mandel LJ: Differential effects of respiratory inhibitors on glycolysis in proximal tubules. Am J Physiol 1990; 258:F1608-F1615.

175. Ruegg CE, Mandel LJ: Bulk isolation of renal PCT and PST. II. Differential responses to anoxia or hypoxia. Am J Physiol 1990; 259:F176-F185.

176. Feldkamp T, Kribben A, Roeser NF, et al: Accumulation of nonesterified fatty acids causes the sustained energetic deficit in kidney proximal tubules after hypoxia-reoxygenation. Am J Physiol 2005; 288:F1092-F1102.

177. Feldkamp T, Kribben A, Roeser NF, et al: Preservation of complex I function during hypoxia-reoxygenation-induced mitochondrial injury in proximal tubules. Am J Physiol 2004; 286:F749-F759.

178. Leichtweiss HP, Lubbers DW, Weiss C, et al: The oxygen supply of the rat kidney: Measurements of intrarenal pO2. Pflugers Arch 1969; 309:328-349.

179. Prasad PV, Edelman RR, Epstein FH: Noninvasive evaluation of intrarenal oxygenation with BOLD-MRI. Circulation 1996; 94:3271-3275.

180. Brezis M, Rosen S, Silva P, Epstein FH: Transport activity modifies thick ascending limb damage in the isolated perfused kidney. Kidney Int 1984; 25:65-72.

181. Kakoki M, Kim HS, Arendshorst WJ, Mattson DL: L-Arginine uptake affects nitric oxide production and blood flow in the renal medulla. Am J Physiol Regul Integr Comp Physiol 2004; 287:R1478-R1485.

182. Morris Jr SM, Sweeney Jr WE, Kepka DM, et al: Localization of arginine biosynthetic enzymes in renal proximal tubules and abundance of mRNA during development. Pediatr Res 1991; 29:151-154.

183. Levillain O, Hus-Citharel A, Morel F, Bankir L: Localization of arginine synthesis along rat nephron. Am J Physiol 1990; 259:F916-F923.

184. Blantz RC, Satriano J, Gabbai F, Kelly C: Biological effects of arginine metabolites. Acta Physiol Scand 2000; 168:21-25.

185. Miyanaka K, Gotoh T, Nagasaki A, et al: Immunohistochemical localization of arginase II and other enzymes of arginine metabolism in rat kidney and liver. Histochem J 1998; 30:741-751.

186. Levillain O, Hus-Citharel A, Morel F, Bankir L: Arginine synthesis in mouse and rabbit nephron: Localization and functional significance. Am J Physiol 1993; 264:F1038-F1045.

187. Ortiz PA, Garvin JL: Role of nitric oxide in the regulation of nephron transport. Am J Physiol Renal Physiol 2002; 282:F777-F784.

188. Lu M, Wang X, Wang W: Nitric oxide increases the activity of the apical 70-pS K+ channel in TAL of rat kidney. Am J Physiol 1998; 274:F946-F950.

189. Lu M, Giebisch G, Wang W: Nitric oxide links the apical Na+ transport to the basolateral K+ conductance in the rat cortical collecting duct. J Gen Physiol 1997; 110:717-726.

190. Tojo A, Guzman NJ, Garg LC, et al: Nitric oxide inhibits bafilomycin-sensitive H+-ATPase activity in rat cortical collecting duct. Am J Physiol 1994; 267:F509-F515.

191. Deng A, Miracle CM, Suarez JM, et al: Oxygen consumption in the kidney: Effects of nitric oxide synthase isoforms and angiotensin II. Kidney Int 2005; 68:723-730.

192. Moncada S, Erusalimsky JD: Does nitric oxide modulate mitochondrial energy generation and apoptosis?. Nat Rev Mol Cell Biol 2002; 3:214-220.

193. Burwell LS, Nadtochiv SM, Tompkins AJ, et al: Direct evidence for S-nitrosation of mitochondrial complex I. Biochem J 2006; 15:627-634.

194. Loke KE, McConnell PI, Tuzman JM, et al: Endogenous endothelial nitric oxide synthase-derived nitric oxide is a physiological regulator of myocardial oxygen consumption. Circ Res 1999; 84:840-845.

195. Elfering SL, Sarkela TM, Giulivi C: Biochemistry of mitochondrial nitric-oxide synthase. J Biol Chem 2002; 277:38079-38086.

196. Garvin JL, Hong NJ: Nitric oxide inhibits sodium/hydrogen exchange activity in the thick ascending limb. Am J Physiol 1999; 277:F377-F382.

197. Packer MA, Porteous CM, Murphy MP: Superoxide production by mitochondria in the presence of nitric oxide forms peroxynitrite. Biochem Mol Biol Int 1996; 40:527-534.

198. Halestrap AP, Doran E, Gillespie JP, O'Toole A: Mitochondria and cell death. Biochem Soc Trans 2000; 28:170-177.

199. Schmid H, Scholz M, Mall A, et al: Carbohydrate metabolism in rat kidney: Heterogeneous distribution of glycolytic and gluconeogenic key enzymes. Curr Probl Clin Biochem 1977; 8:282-289.

200. Zeidel ML, Seifter JL, Lear S, et al: Atrial peptides inhibit oxygen consumption in kidney medullary collecting duct cells. Am J Physiol 1986; 251:F379-F383.

201. Grunewald RW, Kinne RK: Sugar transport in isolated rat kidney papillary collecting duct cells. Pflugers Arch 1988; 413:32-37.

202. Chamberlin ME, Mandel LJ: Substrate support of medullary thick ascending limb oxygen consumption. Am J Physiol 1986; 251:F758-F763.

203. Terada Y, Knepper MA: Na+-K+-ATPase activities in renal tubule segments of rat inner medulla. Am J Physiol 1989; 256:F218-F223.

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