Iron transport in the kidney: implications for physiology and cadmium nephrotoxicity

Abstract

The kidney has recently emerged as an organ with a significant role in systemic iron (Fe) homeostasis. Substantial amounts of Fe are filtered by the kidney, which have to be reabsorbed to prevent Fe deficiency. Accordingly Fe transporters and receptors for protein-bound Fe are expressed in the nephron that may also function as entry pathways for toxic metals, such as cadmium (Cd), by way of “ionic and molecular mimicry”. Similarities, but also differences in handling of Cd by these transport routes offer rationales for the propensity of the kidney to develop Cd toxicity. This critical review provides a comprehensive update on Fe transport by the kidney and its relevance for physiology and Cd nephrotoxicity. Based on quantitative considerations, we have also estimated the in vivo relevance of the described transport pathways for physiology and toxicology. Under physiological conditions all segments of the kidney tubules are likely to utilize Fe for cellular Fe requiring processes for metabolic purposes and also to contribute to reabsorption of free and bound forms of Fe into the circulation. But Cd entering tubule cells disrupts metabolic pathways and is unable to exit. Furthermore, our quantitative analyses contest established models linking chronic Cd nephrotoxicity to proximal tubular uptake of metallothionein-bound Cd. Hence, Fe transport by the kidney may be beneficial by preventing losses from the body. But increased uptake of Fe or Cd that cannot exit tubule cells may lead to kidney injury, and Fe deficiency may facilitate renal Cd uptake.

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Frank Thévenod

Frank Thévenod received his MD from Goethe University, Frankfurt, Germany and his PhD in cellular physiology and biophysics from Case Western Reserve University (CWRU), Cleveland, OH. After postdoctoral training at the Max-Planck-Institute of Biophysics, Frankfurt, Germany and CWRU, he became assistant professor at Saarland University, Homburg, Germany and subsequently senior lecturer at the University of Manchester, UK. Since 2002 he holds the Chair of Physiology, Pathophysiology and Toxicology in the Faculty of Health at Witten/Herdecke University, Germany. His work focuses on renal physiology and toxicology of transition metals, epithelial transport, signal transduction, and molecular mechanisms of cellular metal homeostasis.

Natascha A. Wolff

Natascha A. Wolff received her diploma in biology from the University of Hannover, Germany, and her PhD from the Ruhr-University Bochum on work carried out at the Max-Planck-Institute of Molecular Physiology (formerly Systems Physiology), Dortmund, Germany. After postdoctoral studies at the NIEHS/NIH, NC, she became research associate at the Georg-August University Göttingen, Germany, where she received her Venia legendi (Habilitation) for Physiology. Since 2005, she is research associate at the University of Witten/Herdecke, Germany. Her work focuses on epithelial transport of organic ions and metals, as well as cellular metal homeostasis and nephrotoxicity by transition metals.

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1. Introduction

Until recently, the kidney was thought to play no role in systemic iron (Fe) homeostasis.¹ On the other hand, cadmium (Cd) has been known for decades to accumulate in the kidney (described in detail in the excellent review by G. F. Nordberg²). The major Fe compounds in biological systems are the redox pair ferrous (Fe²⁺) and ferric (Fe³⁺) iron whereas the major Cd compound is the divalent Cd ion (Cd²⁺). Due to their hydrophilicity Fe²⁺/Fe³⁺ and Cd²⁺ (and other metal ions) must cross cellular membranes via proteinous pathways, i.e. channels, transporters or receptors. In biological systems Fe²⁺/Fe³⁺ and Cd²⁺ are mostly found in bound form and are either complexed to small ligands, such as amino acids or peptides,³ or more or less specifically bound to proteins (e.g. ferritin, transferrin, metallothionein or albumin) whose affinity constants determine their residency as “free” or bound metal ions. As a non-essential metal in mammals, Cd²⁺ must exploit and compete for physiological entry pathways for essential metal ions, such as Fe²⁺, Cu²⁺, Zn²⁺ or Mn²⁺. To describe this process, the term “ionic and molecular mimicry” has been coined.⁴ In this context, molecular mimicry accounts for a condition in which a toxic metal ion forms a complex with an endogenous organic ligand (e.g. a peptide or a protein) and the resulting compound mimics the behavior of the endogenous ligand that binds to its receptor.

Interestingly, epidemiological studies had shown an inverse relationship between the size of the Fe stores and the Cd burden of the body (and kidneys),^5,6 thus hinting at a link between transport of Fe²⁺/Fe³⁺ and Cd²⁺ (reviewed in ref. 7). Mounting functional evidence for transport of Cd²⁺ by transporters and receptors for essential free and complexed metal ions in renal and other epithelia^8,9 was then superseded by the discovery that the first cloned Fe²⁺ transporter, the divalent metal transporter 1 (DMT1/Nramp2/DCT1/SLC11A2), is equally well permeated by Cd²⁺.^10,11 DMT1 (and other Fe²⁺/Cd²⁺ transporting proteins) is, however, highly expressed in the kidney.

This fact soon attracted the attention of Craig P. Smith from the University of Manchester (U.K.) who noticed the fact that the kidneys “…contain many if not all of the proteins that are central to iron balance, that in some cases are expressed in considerable amounts, implies that the kidney handles iron in some way that has demanded evolutionary conservation and therefore is likely to be of importance…”.¹² In a series of pioneering studies, his group measured Fe reabsorption by the rat kidney in vivo.¹³ From their data they estimated that under physiological conditions ∼0.4 mg Fe is filtered daily by rat kidneys, but only 0.7% is excreted in the urine and also appears to depend on the renal expression of DMT1.^14,15 Alterations of dietary Fe intake modulated renal DMT1 expression: iron restriction increased renal DMT1 whereas iron loading decreased renal DMT1 expression and DMT1 expression was inversely correlated with urinary Fe output.¹⁵ Therefore they concluded that long-term modulation of renal DMT1 expression may influence renal iron excretion rate. In addition, it soon became apparent that a certain proportion of filtered Fe is protein-bound and includes transferrin (Tf).^16,17 Meanwhile, additional renal Fe transport pathways have been identified and characterized that now allow a better understanding of the role of the kidney in Fe handling and physiological Fe homeostasis. Thus, the notion that the kidney is involved in transport and excretion of Fe and other metal ions¹² has gained recognition and has entered the fields of toxicology,¹⁸ iron biology^19,20 and nephrology.²¹

2. Systemic iron homeostasis

For detailed accounts of systemic Fe homeostasis, the reader is referred to excellent recent reviews.^1,22,23 Iron, with an amount of ∼2.5–4.5 g in adults, is the major transition metal in the body and is mostly localized in erythrocyte hemoglobin, amounting to roughly 60% of the human total body Fe. Yet, Fe is indispensable for other tissues as well, being an essential component of hundreds of proteins, including many enzymes. Thus, Fe is not only required for oxygen transport and storage with hemoglobin or myoglobin, but Fe-containing proteins are needed for a variety of additional functions, including, first and foremost, mitochondrial respiration (electron transport chain), but also metabolism and detoxification (cytochrome P450 enzymes), DNA synthesis (ribonucleotide reductase), antioxidant defense (catalase) and beneficial pro-oxidant functions, oxygen sensing (hypoxia-inducible factor (HIF) prolyl hydroxylases), and immune defense (myeloperoxidase).²⁴ Yet, Fe is also toxic due to the production of cell-damaging radicals through Fenton-type reactions.²⁵ Thus, body Fe homeostasis needs to be tightly regulated.

Mammalian Fe homeostasis is unusual in that it is mainly controlled at the level of intestinal Fe absorption. To date, there is no known regulated short-term mechanism of Fe excretion (but see Section 1 for an example of long-term modulation in the kidney; reviewed in ref. 12) and the small daily Fe loss of about 1 mg in healthy adult males is closely balanced by duodenal Fe uptake. The daily Fe loss mainly (∼80%) occurs via shedding of Fe-laden duodenal enterocytes, complemented by much smaller losses via other pathways, including the kidneys²⁶ (reviewed in ref. 27 and 28). The large fraction of Fe in erythrocyte hemoglobin is efficiently recycled, while excess Fe is stored in the liver (0.5–1 g) (reviewed in ref. 27 and 28). Under normal conditions, Fe loss is balanced via regulated intestinal absorption.²⁹ Two forms of dietary Fe are taken up into duodenal enterocytes via different mechanisms: while heme Fe uptake occurs via not yet clearly defined pathways (see ref. 30–32), non-heme Fe absorption is mediated by the proton-coupled divalent metal transporter 1 (DMT1/Nramp2/DCT1/SLC11A2)¹⁰ (reviewed in ref. 33) after reduction of dietary Fe³⁺ to Fe²⁺ by duodenal cytochrome B with ascorbate as an electron donor (reviewed in ref. 34). An apical intestinal Na⁺/H⁺ exchanger appears to be responsible for generating the proton gradient necessary for DMT1-mediated Fe²⁺ uptake.³⁵ Fe²⁺ is subsequently delivered to cytoplasmic ferritin for storage³⁶ by chaperones, including the poly(rC)-binding protein 1 (PCBP1),³⁷ or to the basolateral transporter ferroportin (FPN1/IREG1/MTP1/SLC40A1)^38–41 for efflux into the plasma. Efficient export requires the presence of members of a family of copper-containing ferroxidases, e.g. hephaestin and/or ceruloplasmin,^42–44 which convert effluxed Fe²⁺ to Fe³⁺ that mainly binds to the Fe-carrying serum protein Tf. Importantly, FPN1, to date the only known cellular Fe exporter,⁴⁵ is regulated by Fe loading⁴⁶ through homeostatically increased synthesis and release of the hepatic peptide hepcidin into the circulation that limits further absorption of dietary Fe and its release from stores (reviewed in ref. 47): hepcidin binds to FPN1, leading to its internalization and subsequent lysosomal degradation, hence preventing further Fe export into the plasma.⁴⁸

Free, unbound Fe is incompatible with either plasma Fe transport (it would precipitate) or with cytosolic Fe trafficking (it would damage the cellular environment).⁴⁹ Therefore, Fe must be complexed with appropriate ligands. The transport of Fe in plasma to its sites of use occurs predominantly as Tf-bound Fe⁵⁰ (TBI), and to a lesser extent associated with several other serum proteins, including ferritin, albumin, neutrophil gelatinase associated lipocalin (NGAL/24p3/lipocalin-2), and possibly lactoferrin and hepcidin. Collectively, these latter forms of serum Fe – with the exception of ferritin – are termed non-Tf-bound Fe (NTBI).^49,51,52 Under physiological conditions, NTBI is a minor entity within total serum Fe – although NTBI may become a relevant issue in patients with various pathological conditions in which Tf saturation is significantly elevated (reviewed in ref. 49). Ferritin, primarily an intracellular protein, is low in human serum under normal conditions, despite substantial inter-individual variations and substantial increases under Fe overload conditions⁵³ where it may be secreted through a non-classical lysosomal secretory pathway by macrophages and renal proximal tubule (PT) cells.⁵⁴ Additionally, serum Fe may exist in the form of holo-Tf bound to a soluble form of the Tf receptor.⁵⁵

3. Cellular iron homeostasis

One major mechanism for Fe assimilation by erythrocyte precursors and non-erythroid cells is the internalization of serum Tf-bound Fe³⁺.^56,57 Tf endocytosis is mediated by the ubiquitous Tf receptor 1 (TfR1)^58,59 (a TfR2 has been cloned, but its expression is limited to the liver and erythropoietic progenitors,⁶⁰ where it is thought to operate as an “Fe sensor”⁶¹). Endosomal acidification favors the release of iron from Tf, which itself remains bound to the receptor and is subsequently recycled to the cell surface, where the near neutral pH promotes dissociation of apo-Tf from the receptor and its release into the circulation.^62,63 Endosomal Fe³⁺ is quickly reduced to Fe²⁺ by an oxidoreductase activity, now known to be represented by “Steap” (sixtransmembrane epithelial antigen of the prostate) family proteins, namely Steap2 to Steap4,^64,65 (a reaction which may actually occur prior to dissociation from the Tf–TfR1 complex, especially since Fe³⁺ tightly binds to Tf, while Fe²⁺ does so only weakly⁶⁶). Subsequent endosomal efflux of Fe²⁺ is mediated by DMT1.^67,68 The transient receptor potential mucolipin 1 (TRPML1/ML1/MLN1/MCLN1) may function as another Fe²⁺ release channel in late endosomes and lysosomes.⁶⁹

The mechanisms of intracellular Fe trafficking to its sites of utilization are not well understood. In most cell types, it is agreed that Fe acquired during the Tf cycle is first released into the cytosol by entering a “labile cytosolic Fe pool” that is defined as a pool of chelatable and redox-active Fe²⁺ and represents a transition compartment for Fe sensing, metabolic utilization or storage.^70,71 A variety of low molecular weight compounds have been suggested as Fe chelators in this readily accessible Fe reservoir, including organic anions like citrate and phosphate, oligopeptides such as glutathione (GSH),⁷² membrane phospholipids, as well as “mammalian siderophores”, namely 2,5-dihydroxybenzoic acid (2,5-DHBA),⁷³ although an involvement of the latter has been recently challenged.⁷⁴ Further, the conserved cytosolic glutaredoxins Grx3 and Grx4 could also play an essential role in intracellular Fe sensing and trafficking, as their depletion in yeast leads to impaired Fe transport to mitochondria and defects in Fe-dependent pathways.⁷⁵

In contrast, in erythroid cells, kinetic and microscopy studies support a “kiss and run” hypothesis, which assumes the direct delivery of Tf-derived Fe to mitochondria through a transient contact with endosomes (reviewed in ref. 76): this concept was originally developed based on kinetic Fe release studies with ⁵⁹Fe–Tf in reticulocytes at 4 °C that contain very little chelatable cytosolic Fe, thus preventing Fe mobilization from other compartments.⁷⁷ Ponka and coworkers later observed direct, albeit transient inter-organellar contacts and a simultaneous increase in mitochondrial chelatable Fe at these sites by live confocal imaging.⁷⁸

The major Fe-utilizing cellular organelles are mitochondria that require Fe for the synthesis of heme and Fe–sulfur clusters.^79,80 Irrespective of whether Fe is delivered by cytosolic chaperones or direct endosome-mitochondria contacts, it has to cross two membranes to enter the mitochondrial matrix, where it is needed for synthetic processes. The outer mitochondrial membrane (OMM) has typically been assumed to be freely permeable to Fe due to the presence of “pores”⁸¹ represented by voltage-dependent anion channels (VDACs) that are regarded as the major permeability pathway of the OMM for small solutes.⁸² But this knowledge is based on in vitro studies that have been performed after reconstitution of VDAC in artificial membranes/planar lipid bilayers. Thus, in vivo the OMM may not be as freely permeable to inorganic cations as previously believed and VDAC function may be tightly regulated.^83,84 We have recently identified DMT1 in several tissues as a possible mechanism for Fe²⁺ transfer across the OMM using a variety of experimental approaches^85,86 (see Section 6.1.3), but additional pathways may also exist.

Entry of Fe into the mitochondrial matrix requires the SLC transporter mitoferrin-1 and -2 (also known as MFRN1/SLC25A37 and MFRN2/SLC25A28), which are found in the inner mitochondrial membrane (IMM).⁸⁷ Mitoferrin-1 is highly enriched in erythroid cells and is stabilized during differentiation whereas mitoferrin-2 is ubiquitously expressed and its half-life is not regulated.⁸⁸ The lack of functional mitoferrin-1 in the frascati zebrafish mutant is associated with severe defects in erythropoiesis, heme synthesis and Fe–sulfur clusters biogenesis.⁸⁷ Some studies have also suggested that the mitochondrial calcium uniporter in the IMM represents an additional route of Fe entry into the matrix (e.g.ref. 89).

Cells may eliminate excess intracellular Fe by secretion of Fe²⁺via FPN1 or by secretion of heme through the putative heme exporter FLVCR (feline leukemia virus, subgroup C, receptor).⁹⁰ Excess intracellular Fe may also be stored and detoxified in the cytosol by ferritin, which consists of 24 H (heavy) and L (light) subunits, encoded by two different genes.⁹¹ H-ferritin possesses ferroxidase activity, mediating conversion of ferrous Fe (Fe²⁺) to the ferric form (Fe³⁺), whereas L-ferritin chains provide a nucleation center. Ferritin assembles into a shell-like structure with a cavity of ∼80 nm that provides storage space for up to 4500 Fe³⁺ ions. Shuttling of Fe to ferritin appears to be mediated by the PCBP family chaperones^37,92 (see Section 2). Both the lysosomal and the proteasomal pathways of degradation seem to be recruited to mobilize Fe from ferritin, probably depending on the cell type and the cellular conditions (reviewed in detail in ref. 91). Mitochondria contain a nuclear-encoded ferritin isoform⁹³ whose expression is limited to a few organs, such as testis, neurons, heart and kidney, but not the liver or spleen.⁹⁴ Mitochondrial ferritin may cooperate with cytosolic ferritin in the maintenance of intracellular Fe balance or protect mitochondria from Fe-dependent oxidative damage and increased production of reactive oxygen species (ROS) in cells with high metabolic rate.⁹¹

4. Function of the kidney

Detailed state-of-the-art descriptions of the morphology of the kidneys, the structure of the nephrons, i.e. the functional units of the kidneys, and their functions are beyond the scope of this review and can be found in standard handbooks of renal physiology.^95–97 The aim of this very simple overview on kidney function is to introduce readers unfamiliar with renal physiology to basic principles that are required for a better understanding of the handling of Fe and Cd by the kidneys.

The plasma permanently equilibrates with the interstitial fluid of the extracellular space and – through the interstitial fluid – with the intracellular space. Together with the lungs and the intestine, the kidneys keep the body fluid homeostasis of mammalian organisms constant by selectively excreting metabolic wastes, excess solutes and water as well as xenobiotics from the body into the urine. Blood is constantly pumped through the kidneys where plasma fluid is filtered through a capillary network called the glomerulus. The driving force for ultrafiltration is generated by the effective filtration pressure in the capillaries, which is set by the glomerular blood pressure. To fulfill the excretory function of the kidneys, large quantities of plasma amounting to >60× its total body volume are filtered daily in the renal glomeruli, complemented by secretory pathways along the renal tubule epithelium. Filtered water and solutes still of use for the body are efficiently recycled to the circulation by obligatory and regulated reabsorptive processes in the tubular sections of the nephrons. By these means, about 180 l of primary filtrate is generated every day to produce about 1–3 l final urine. This indicates that about 99% of the primary urine is reabsorbed along the more than 2 million nephrons.

At the glomerulus, a three-layer anatomical barrier allows fluid and solutes <10 kDa and/or 18 Å to cross that barrier, but permeation decreases with increasing molecular mass (cutoff of ∼80 kDa), molecular size (<42 Å), and also depends on charge (cationic > neutral > anionic) (see however ref. 98 for a critical discussion). Hence, the primary urine also contains essential nutrients and electrolytes that need to be actively reabsorbed to avoid critical losses and ensuing deficiencies. On the other hand, some metabolic wastes are actively secreted by the kidney since their rate of production exceeds their rate of glomerular filtration. All these selective processes are carried out by the nephrons, epithelial tubular structures that consist of several interconnected segments with characteristic morphological and functional properties, the PT with its convoluted segments S1 and S2 (PCT) and straight segment S3, the loop of Henle (LOH), the distal tubule (DT) with its convoluted segment (DCT) and connecting tubule, and finally the collecting duct (CD) (see Fig. 1). Glomeruli, convoluted segments of the PT, DT with connecting tubule, and cortical CD are localized in the kidney cortex. Parts of the straight segment of the PT, parts of the thin limb of the LOH, the thick ascending limb of the LOH and outer medullary CD are in the outer medulla. The remaining segments (most of the thin limbs of the LOH as well as initial and terminal inner medullary CD) are found in the inner medulla of the kidney.

Fig. 1 Structure and function of the nephron. For further details, see Section 4.

In general, the PT is responsible for bulk reabsorption of the primary fluid that is filtrated into the lumen of that segment. About two-third of PT reabsorption occurs “paracellularly” at intercellular tight-junctions, through osmotically driven “solvent drag”. But amino acids, glucose, bicarbonate and several other essential molecules are also reabsorbed via luminal Na⁺-dependent transporters expressed in the luminal brush-border membranes (BBM) of PT cells and therefore require the energy of adenosine triphosphate (ATP) for activation of basolateral Na⁺/K⁺-ATPases to maintain these reabsorptive processes. The PT cells are also responsible for bulk reuptake of filtered proteins and peptides via a multi-ligand receptor complex expressed in the luminal BBM, megalin:cubilin:amnionless⁹⁹ (see Section 6.1.1), that also binds metalloproteins, such as Tf (an Fe binding protein) or metallothionein (MT) (a Cd²⁺ binding protein).¹⁰⁰ Finally, the PT is the major location for the secretion of xenobiotic and endogenous organic cations and anions. The LOH that follows the PT builds up the hyperosmotic interstitium surrounding the final segment of the nephron, the CD that is required for reabsorption of water to generate small volumes of concentrated urine (“antidiuresis”), thus preserving water for the body. Hyper-osmolarity of the kidney medulla is built up by several properties of the different segments of LOH, i.e. (1) active NaCl transport into the interstitium by the thick ascending limb of LOH; (2) high permeability of the descending LOH to water and low permeability of the ascending LOH; (3) increased permeability to urea in the medullary portions of LOH; and (4) magnification of the medullary hyper-osmolarity by the countercurrent flow within the descending and ascending limbs of LOH (“countercurrent multiplication”).^95–97 A hypo-osmotic fluid reaches the DT where divalent metal ions are reabsorbed, such as Ca²⁺ and Mg²⁺ (and Fe²⁺) (see Section 6.3) and the luminal fluid is further depleted by active NaCl reabsorption. In the CD the final composition of the urine is adjusted by “fine-tuning” through hormonal regulation of the CD cells. The paracellular permeability of the CD epithelial layer to ions and water is very low; therefore the final content of the urine in NaCl and water must be controlled by hormonal regulation of CD cells via aldosterone and antidiuretic hormone. This occurs through regulated and temporary incorporation of epithelial Na⁺ channels (ENaC) and aquaporin-2 water channels (AQP2), respectively, into the apical membrane of principal (light) cells of the CD. Additional regulated functions of the CD include acid–base balance (type A- and type B-intercalated cells) and K⁺ homeostasis. Apart from the CD, PT and DT also represent nephron segments where hormones (i.e. parathyroid hormone, calcitonin, calcitriol) control Ca²⁺ and PO₄³⁻ homeostasis. Fig. 1 summarizes the structure of the nephron with the major functions of the different nephron segments that are relevant to this review.

5. Plasma iron and renal glomerular filtration

Only recently has it been recognized that the kidney is also involved in systemic Fe homeostasis because certain Fe-containing complexes in plasma (e.g. Tf, NGAL/24p3/lipocalin-2, lactoferrin, albumin, hemoglobin, myoglobin and hepcidin) have the ability to cross the glomerular filter, even under physiological conditions^101,102 (reviewed in ref. 12 and 103). There is also a rising interest in the role of Fe in both acute kidney injury and chronic kidney disease.²¹ Renal Fe losses are minimal under physiological conditions²⁶ (reviewed in ref. 27). The lack of urinary Fe excretion has traditionally been attributed to binding of Fe (or, if erythrocytes are lysed within the blood, hemoglobin and free heme) to larger proteins that would ensure that little or no Fe is lost by glomerular filtration and entry into the urine because of the low protein permeability of the glomerular filter.^28,104 That would also include the large 24-subunit serum ferritin complex that is unlikely to reach the ultrafiltrate. But NGAL is present in plasma as monomers of 25 kDa and dimers of 45 kDa that should permeate the glomerular filter.¹⁰⁵ Similarly, the small molecule hepcidin (2–3 kDa) readily passes into the primary urine.⁴⁷ Moreover, early micropuncture studies in animals indicated significant glomerular filtration of high-molecular weight proteins (HMWP), such as albumin (reviewed in ref. 98 and 106). In accordance with these observations, patients with renal Fanconi syndrome, i.e. with compromised renal PT function, including protein reabsorption (reviewed in ref. 107), display increased urinary excretion of proteins up to 160 kDa,¹⁷ suggesting that substantial amounts of TBI, i.e. Fe bound to Tf (80 kDa), as well as NTBI (see Section 2), such as Fe bound to albumin (66.5 kDa) and lactoferrin (80 kDa), reach the primary filtrate and must be reabsorbed by the PT (see Section 6.1.1).

6. Iron transporters of the nephron

(See also Tables 1 and 2 for a summary).

Table 1 Localization and subcellular distribution of iron and cadmium transporting receptors, transporters and channels in the kidney

Receptor/transporter	Nephron localization	Subcellular localization	Species	Ref.	Comments
a PT: proximal tubule; LOH: loop of Henle; DT: distal tubule; CD: collecting duct. b NGAL: neutrophil gelatinase-associated lipocalin.
Megalin:cubilin:amnionless	PT^a	Apical; subapical	Human; mouse; rat; rabbit	108	Immuno-fluorescence/-histochemistry/-gold; co-labeling with segment-/organelle-specific marker
Transferrin receptor 1	PT; CD	Apical; subapical	Mouse	103	Immunofluorescence; poor resolution; specificity of antibody unclear
NGAL^b/24p3/lipocalin-2 receptor (SLC22A17)	DT; CD	Apical; subapical	Mouse; rat	182 and 214	Immunofluorescence/-histochemistry; co-labeling with segment-specific marker
DMT1 (SLC11A2)	PT	Intracellular (endosomes/lysosomes/mitochondria)	Mouse; rat	12, 14, 15, 85, 86, 137 and 140	Immuno-fluorescence/-histochemistry/-gold; co-labeling with segment-/organelle-specific marker
		Apical?	Mouse	138	Immunohistochemistry; poor resolution; collapsed tubules; no co-labeling with segment-specific marker
	LOH	Apical; intracellular	Rat	14	Immunofluorescence; co-labeling with segment-/cell-specific marker
	DT	Apical	Rat	14 and 15	Immunofluorescence; co-labeling with segment-/membrane-specific marker
		∅?	Mouse	138 and 140	Immunohistochemistry; poor resolution; collapsed tubules; no co-labeling with segment-specific marker; specificity of antibody unclear
	CD	Apical; intracellular; basolateral	Rat	14, 15 and 145	Immunofluorescence; co-labeling with segment-/cell-specific marker
ZIP8 (SLC39A8)	PT	Apical?; subapical	Mouse	153	Immunofluorescence/-histochemistry; co-labeling with membrane-specific marker, yet poor resolution (discussed in ref. 141)
ZIP14 (SLC39A14)	PT?	?	∅	∅	No staining of native tissue; staining in overexpressing cell lines only (discussed in ref. 141)
Ferroportin (FPN1/SLC40A1)	PT	Basolateral	Mouse; rat	123, 160 and 161	Immuno-fluorescence/-histochemistry/-gold; co-labeling with segment-/membrane-specific marker
		Apical/basolateral?	Mouse	140 and 162	Immunohistochemistry; poor resolution; no co-labeling with segment-specific marker; specificity of antibody unclear
	LOH	Basolateral	Mouse	123	Immunofluorescence; co-labeling with segment-/membrane-specific marker
		∅?	Mouse	140	Immunohistochemistry; poor resolution; no co-labeling with segment-specific marker; specificity of antibody unclear
	CD	Intracellular?	Mouse	140	Immunohistochemistry; poor resolution; no co-labeling with segment-specific marker; specificity of antibody unclear
		∅	Mouse	123	Immunofluorescence; co-labeling with segment-/membrane-specific marker
TRPV5 (ECaC1)	DT	Apical; subapical	Rat	164	Immunofluorescence; co-labeling with segment-specific markers
Cav3.1 (α1G)	DT; CD	Apical	Rat	175	Immunohistochemistry; co-labeling with segment-specific markers

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Table 2 Functional properties of iron and cadmium transporting receptors, transporters and channels of the kidney

Uptake pathway	Localization	Fe			Cd^2+
		Substrate/ligand	K D/K0.5 (nmol l^−1)	Ref.	Substrate/ligand	K D/K0.5 (nmol l^−1)	Ref.
a PT: proximal tubule; LOH: loop of Henle; DT: distal tubule; CD: collecting duct. b See Section 7.2 for detailed explanations. c NGAL: neutrophil gelatinase-associated lipocalin.
Megalin:cubilin:amnionless	PT^a	Transferrin	20	16	?^b	100 000	207
		NGAL	60	112	Cd^2+-metallothionein
Transferrin receptor 1	PT; CD	Transferrin	0.2–0.4	124 and 125	?	∅	∅
NGAL^c/24p3/lipocalin-2 receptor (SLC22A17)	DT; CD	Transferrin	100	182	Cd^2+-metallothionein	100	182
		NGAL	0.090	197
DMT1 (SLC11A2)	PT; LOH; DT; CD	Fe^2+	1000	11	Cd^2+	1000	11
ZIP8 (SLC39A8)	PT	Fe^2+	700	152	Cd^2+	620	148
ZIP14 (SLC39A14)	PT	Fe^2+	2300	151	Cd^2+	100–1100	149
Ferroportin (FPN1/SLC40A1)	PT; LOH; CD	Fe^2+	<100	41	∅	∅	41
TRPV5 (ECaC1)	DT	Fe^2+	<1000 (estimated)	169	Cd^2+	Micromolar (estimated)	297
Cav3.1 (α1G)	DT; CD	Fe^2+	Low micromolar (estimated)	180	Cd^2+	Low nanomolar (estimated)	295

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6.1. Iron transporters of the proximal tubule (PT)

6.1.1. Megalin:cubilin:amnionless. Megalin is a 600 kDa single transmembrane-domain receptor protein that belongs to the low-density lipoprotein receptor family. Megalin is responsible for the normal PT reabsorption of filtered plasma proteins, thus preventing the loss of these essential molecules into the urine.¹⁰⁸ The almost complete clearance of proteins from the ultrafiltrate by megalin-driven endocytosis is accomplished in cooperation with the 460 kDa glycosylated protein receptor cubilin.¹⁰⁹ The normal function of cubilin is also dependent on the 38 to 50 kDa, single transmembrane protein amnionless that is essential for the trafficking of cubilin to the apical plasma membrane.¹¹⁰ Megalin:cubilin:amnionless are expressed primarily in luminal plasma membranes of polarized absorptive epithelia.^99,108 Megalin binds a very wide range of ligands, including plasma transport proteins, peptides, hormones, etc. Known ligands of megalin normally filtered by the glomeruli include retinol-binding protein, transcobalamin-B12, insulin, α1- and β2-microglobulin, albumin, etc. (reviewed in ref. 108) (see Table 3). Although megalin and cubilin are structurally very different, some of the ligands are shared with cubilin, whereas others are specific for either megalin or cubilin (reviewed in ref. 108). The receptors are co-expressed in several tissues, where they interact to function: internalization of several cubilin ligands is strongly facilitated by megalin.¹⁰⁸ Megalin:cubilin:amnionless are highly expressed in the convoluted segments of the renal PT. Following binding to these receptors, ligands are internalized into coated vesicles and delivered to early and late endosomes. Whereas the receptors are recycled to the apical membrane, the ligands are transferred to late endosomes and lysosomes for protein degradation.¹⁰⁸

Table 3 Binding properties and estimated concentrations of ligands of megalin:cubilin in the glomerular filtrate of the kidney

Megalin:cubilin:amnionless
Ligand	K D (nmol l^−1)	Ref.	Concentration in plasma (μmol l^−1)	Ref.	Concentration in glomerular filtrate^a (nmol l^−1)
a Calculations are based on estimated glomerular sieving coefficients of plasma proteins.^17
Transferrin	20	16	35	17	2
NGAL [human]/siderocalin/24p3 [rodent]/lipocalin-2	60	112	7	116	650
Albumin	630	115	690	17	53
Metallothionein	100 000	207	0.0005–0.005	208 and 209	0.5–5
β2-Microglobulin	420	278	0.11	17	100
α1-Microglobulin	n.d.	108	1	17	92

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Reabsorption of filtered TBI in the renal PT has been mainly attributed to megalin-dependent cubilin-mediated endocytosis.¹⁶ Yet, cubilin-independent megalin-mediated uptake of Tf may also occur.¹¹¹ Based on its plasma concentration and calculated glomerular sieving coefficient (GSC; derived from studies in patients with renal Fanconi syndrome),¹⁷ the Tf concentration in the primary filtrate has been estimated to ∼2 nM, which would allow its PT reabsorption via cubilin because this receptor binds Tf with a K_D of ∼20 nM, as determined by surface plasmon resonance analysis.¹⁶ The same applies to other filtered Fe-binding proteins (see above) that are known substrates of cubilin and/or megalin, including NGAL,¹¹² albumin,^111,113 and hepcidin.¹¹⁴ Albumin requires cubilin for renal PT internalization, which is supported by experiments using cubilin-deficient mice.¹¹¹ The concentration of albumin in the glomerular filtrate has been calculated to ∼53 nM¹⁷ and the K_D of albumin to cubilin amounts to ∼0.63 μM.¹¹⁵ The plasma concentration of NGAL in healthy subjects amounts to ∼6.5 μM¹¹⁶ and should reach concentrations approximating ∼0.65 μM in the ultrafiltrate based on an estimated GSC of ∼0.1.¹⁷ Surface plasmon resonance analysis has demonstrated binding of apo-NGAL to megalin with a K_D of ∼60 nM,¹¹² which is about 10-fold lower than the estimated NGAL concentration in the primary filtrate (see Table 3). Although, both lactoferrin¹¹⁷ and hepcidin¹¹⁴ bind to megalin and are likely to be filtered by the glomerulus (the latter completely), their binding affinity to megalin has not be determined. Overall, significant amounts of both TBI and NTBI are filtered by the glomerulus and likely to be reabsorbed via megalin:cubilin:amnionless in the PT (but see also Section 6.1.2 for Tf reabsorption).

TBI and NTBI that has been reabsorbed by the PT can meet four possible and not mutually exclusive fates (see ref. 12 for a review): transcytosis; export back into the circulation via the Fe efflux transporter FPN1 aided by hephaestin (see Section 6.1.5); storage in ferritin (see Section 3); and utilization by PT cytosolic or mitochondrial Fe requiring processes (see Section 6.1.3). In vivo transcytosis of Fe transporting proteins has been recently reported by a number of groups. Thus, albumin transcytosis in the PT was inferred from intravital tracking of fluorescent albumin by two-photon microscopy¹¹⁸ as well as in a study showing the appearance of transgenic albumin specifically expressed in podocytes in the plasma where transcytosis was suggested to be mediated by a neonatal Fc (fragment crystallizable) receptor.¹¹⁹ However, the issue of whether transcytosis of intact albumin actually occurs in renal PT is highly controversial.^120,121 Whether bound Fe may be retained on albumin (and possibly other proteins) during such transcellular transfer, or may rather be released in some intracellular transit compartment, has, to our knowledge, not yet been investigated. Although transcytosis has also been reported for ferritin infused into the renal PT¹²² this process is unlikely to play a role in vivo due to the large size and therefore poor glomerular filtration of ferritin (see Section 5).

6.1.2. Transferrin receptor 1 (TfR1). Renal PT cells express TfR1 at the apical membrane, at least in some species (and possibly in CD of kidney medulla),^103,123 that could contribute to reabsorption of filtered TBI (see ref. 12 and 103 for reviews), considering the very high affinity of TfR1 to its natural ligand Tf (K_D ∼ 0.2–0.4 nM).^124,125 A recent study with cultured PT cells, to date published only in abstract form,¹²⁶ points to the possibility that, while megalin:cubilin mediated TBI reabsorption by the PT may predominate under Fe replete conditions, TfR1 becomes the principal receptor for Tf endocytosis under conditions of Fe restriction, due to differential regulation of these pathways by Fe.¹²⁶

6.1.3. DMT1 (SLC11A2). The first mammalian Fe transporter protein DMT1 (divalent metal transporter 1) was identified by expression cloning.¹⁰ DMT1 is a ferrous ion (Fe²⁺) transporter that is energized by the H⁺ electrochemical potential gradient while ferric ion (Fe³⁺) is excluded.¹⁰ However, H⁺ coupling may not always be necessary for Fe²⁺ transport.¹²⁷ The role of DMT1 as a Fe²⁺ transporter was confirmed with Belgrade rats (b/b) and mk mice that carry a DMT1 G185R mutation, which results in deficient Fe²⁺ uptake and microcytic anaemia.^68,128 DMT1 occurs in 4 major isoforms, which differ in their N- and C-termini. Isoforms 1A and 1B result from alternative 5′ promoter usage with the translation of isoform 1B actually starting at exon 2; alternative 3′ splicing yields two isoforms with different C-termini generated from transcripts containing (isoform I, +IRE) or lacking (isoform II, −IRE) an Fe-response element (IRE) in their 3′ untranslated region.^129,130 Functionally, the major human isoforms exhibit very similar characteristics when expressed in Xenopus laevis oocytes.¹³¹ Although Gunshin et al.¹⁰ demonstrated that, in addition to Fe²⁺, a broad range of transition metals (including Cd²⁺, Zn²⁺, Mn²⁺, Cu²⁺, Co²⁺, Ni²⁺ and Pb²⁺) could evoke inward currents in Xenopus oocytes expressing rat DMT1, subsequent studies using a combination of voltage clamp, radiotracer and fluorescence assays in Xenopus oocytes or transfected HEK293 cells have established that human DMT1 is only capable of efficiently transporting Cd²⁺, Fe²⁺ Co²⁺ and Mn²⁺.^11,132 Human DMT1 exhibited the highest affinity for Cd²⁺ and Fe²⁺ (K^M_0.5 ≈ 1 μM) (see Table 2), showed moderately high affinity for Co²⁺ and Mn²⁺ (K^M_0.5 in the range 3–4 μM), whereas human DMT1 reacted with Ni²⁺, VO²⁺, and Zn²⁺ at lower affinity (K^M_0.5 in the range 10–20 μM). At −70 mV and at pH 5.5, the selectivity of human DMT1 metal-ion substrates were ranked Cd²⁺ > Fe²⁺ > Co²⁺ and Mn²⁺ ≫ Zn²⁺, Ni²⁺, VO²⁺.¹¹ Whether DMT1 may also transport Cu²⁺ is still a matter of debate and may depend on the species and/or isoform investigated.^11,133

At the tissue level, DMT1 is ubiquitously expressed, most notably in the proximal duodenum, red blood cells, macrophages, but also in the kidneys and the brain.¹⁰ DMT1 is expressed in the plasma membrane, typically in enterocytes, where it mediates Tf-independent Fe²⁺ absorption into the organism.¹⁰ Alternatively, when DMT1 is located intracellularly, it is involved in the TfR1 pathway of Fe acquisition (as demonstrated in erythrocyte precursors or macrophages^67,134) (see Section 3). There, DMT1 is localized to intracellular endosomes and lysosomes that are formed during endocytosis of the Tf–TfR1 complex. Vacuolar-type ATPases acidify the endosomes and lysosomes which promotes dissociation of Fe³⁺. Fe³⁺ is reduced to Fe²⁺ by ferrireductase/oxidoreductase activity in the lumen of endosomes and lysosomes that is mediated by Steap proteins and that have also been found expressed at the mRNA level in epithelia, including the kidney.^64,65 This, in turn, activates DMT1 in the lysosomal membrane to co-transport the metal ion along with H⁺ into the cytosol.^67,134

The intracellular localization of DMT1 has also been demonstrated in epithelial cells,¹³⁵e.g. in immunolocalization studies of human 1B/+IRE and 1B/−IRE isoforms overexpressed in HEp-2 human larynx carcinoma cells.¹³⁶ In the kidney, the 1A/−IRE isoform was not detected by semiquantitative RT-PCR of total RNA from mouse kidney.¹³⁰ In contrast, we have detected all four DMT1 transcripts in RNA from rat renal cortex and a rat renal proximal tubule cell line, albeit with different abundance.¹³⁷ At the protein level, evidence could only be obtained for the presence of the +IRE isoforms in mouse kidney cortex that were expressed at the apical pole of PT cells.¹³⁸ In another study, murine +IRE and −IRE DMT1 isoforms were transfected in LLC-PK1 cells: the +IRE isoform was associated with a higher surface expression and slower rate of internalization, as opposed to the −IRE isoform, which was efficiently sorted to recycling endosomes upon internalization, whereas the +IRE isoform was not efficiently recycled and rather targeted to lysosomes.¹³⁹

Consistent with a major role of megalin:cubilin dependent endocytosis for Tf clearance from the ultrafiltrate, DMT1 has been detected in late endosomes and lysosomes of rat kidney PT cells by electron microscopy and also mainly co-localized with late endosomal and lysosomal markers in a renal PT cell line.¹³⁷ Furthermore, a marked increase of punctate intracellular DMT1 immunostaining was observed in rat renal PT upon Fe deprivation, whereas DMT1 was decreased when animals were fed an Fe enriched diet.¹⁵ Free Fe²⁺ has previously been postulated to be reabsorbed via DMT1 residing in the luminal membrane of mouse PT cells.¹³⁸ However, this localization contrasts with other reports indicating exclusive intracellular localization of DMT1, both in PT from rat^14,15,137 and mouse^12,140 (see Table 1 for an overview). Insufficient resolution of the immunohistochemical images in the study by Canonne-Hergaux and Gros¹³⁸ that could not distinguish between apical staining and staining of subapical vesicles has been proposed as a reason for this discrepancy.¹² Moreover, due to the high affinity of transferrin for Fe³⁺,¹⁴¹ a brush-border membrane DMT1 could only reabsorb Fe from NTBI as Fe³⁺ that would also require its reduction by a brush-border ferrireductase and that has not been described in the kidney so far (with the exception of anecdotal evidence for a ferrireductase activity of a prion protein expressed in the apical membrane of PT cells¹⁴²). Moreover, Wareing et al.¹³ have performed tracer microinjections of ⁵⁵FeCl₃ in the early PCT of rat kidney in vivo to determine the percentage of Fe reabsorption in the PT. Since urinary ⁵⁵Fe recovery was independent of the injection site (which varied between 1 and 6 mm from the glomerulus to the injection site) the authors concluded that free Fe is not reabsorbed across the surface convolutions of the PT. This further argues against a role for apical DMT1 (and other Fe transporters) in non-protein bound NTBI reabsorption by the PT.

Mitochondria heavily rely on Fe-dependent metabolism and are therefore intracellular targets for Fe trafficking, which is particularly relevant in the kidney PT and thick ascending limb of LOH where mitochondria provide ATP for active reabsorption and secretion of solutes. Recently, we have obtained evidence for expression of the four major DMT1 isoforms in the OMM in several cell lines and tissues from multiple origin, including the kidney PT, and proposed that mitochondrial DMT1 represents a possible entry pathway for Fe and other metal ions utilized by mitochondria.^85,86 We used a variety of methods, including (1) cryo-immunogold electron microscopy to detect DMT1 co-localization with the OMM protein VDAC1; (2) confocal immunofluorescence microscopy to visualize partial co-localization of DMT1 with the mitochondrial markers VDAC1 and Tom6 (translocase of outer membrane 6); (3) immunoblotting of OMM and IMM fractions to demonstrate co-purification with the OMM marker VDAC1, but not with the IMM marker adenine nucleotide translocase; (4) a split ubiquitin yeast-two hybrid screen where the mitochondrial protein cytochrome C oxidase subunit II (COXII) was identified as an interaction partner of DMT1; (5) co-immunoprecipitation of COXII with DMT1 from cell lysates.⁸⁵ Most importantly, preliminary studies indicate that mitochondria isolated from stably DMT1-transfected HEK293 cells exhibit substantially higher uptake of the known DMT1 substrate ⁵⁴Mn²⁺ when the cells had been pretreated with doxycycline to induce the DMT1 promoter.¹⁴³ Moreover, ⁵⁴Mn²⁺ uptake into mitochondria from induced cells was sensitive to a specific DMT1 inhibitor.¹⁴³ Taken together, these data suggest that DMT1 not only exports Fe²⁺ (and Mn²⁺) from endosomes and lysosomes, but also serves to import metal ions, including Mn²⁺ and Fe²⁺, for mitochondrial utilization in the kidney PT and other tissues and cells.

Homozygous Belgrade rats (b/b) have a G185R mutation of DMT1 that diminishes transport and results in significantly increased serum Fe levels due to the inability of the tissues to utilize Fe.^68,128 These animals have been investigated to estimate the role of renal DMT1 in reabsorption of Fe, however with conflicting results. Belgrade rats showed significantly reduced renal kidney ⁵⁹Fe³⁺ 2 hours after intravenous injection of Fe–Tf, compared to wild-type or heterozygous animals, suggesting that DMT1 is involved in Fe uptake by renal tissue.¹⁴⁴ In contrast, another study showed urinary iron excretion rates that were unchanged in b/b compared to heterozygous animals.¹⁴⁵ This study may cast doubts on a functional role of DMT1 in reabsorption of Fe in the kidney, but alternate path(s) for Fe reabsorption by renal cells may also compensate for the lack of DMT1 protein. Indeed, significantly increased urinary Ca²⁺ excretion was measured in Belgrade rats that did not show DMT1 dependence of urinary Fe excretion rates,¹⁴⁵ which may be explained by increased competition of Fe²⁺ with Ca²⁺ for renal reabsorption by Ca²⁺ channels in the DT of DMT1-deficient Belgrade rats (see Sections 6.3.1 and 6.3.2 for a further discussion). Another aspect needs also to be considered: a recent study with Belgrade rats has hinted to the fact that urinary Fe excretion increases with increasing age of the animals,¹⁴⁶ suggesting a subtle but cumulative impact of DMT1 (dys)function on Fe handling by the kidney. This study may provide another explanation for the negative results described previously where young animals had been used.¹⁴⁵ Consequently, we investigated renal Fe handling in >25 weeks old Belgrade rats and their heterozygous litter mates and measured ∼2-fold increased urinary Fe excretion (184 ± 40 versus 108 ± 9 μg l⁻¹ kg⁻¹ b.w.; n = 3) as well as ∼2-fold decreased kidney Fe concentrations (0.39 ± 0.11 versus 0.21 ± 0.03 mg g⁻¹ kidney tissue; n = 5) in Belgrade animals compared with heterozygous controls (F. Thévenod, A. R. Nair, W.-K. Lee & M. D. Garrick; unpublished), which is in agreement with the studies in Belgrade rats demonstrating the importance of DMT1 for renal Fe reabsorption.^144,146

6.1.4. ZIP8/ZIP14 (SLC39A8/SLC39A14). Two other candidate transporters for non-protein bound Fe have been described in the apical membrane of renal PT, namely the Zrt, Irt-related proteins 8 (ZIP8/SLC39A8) and 14 (ZIP14/SLC39A14).¹⁴⁷ Both carriers are believed to operate as HCO₃⁻ coupled divalent metal ion cotransporters,^148,149 and convincing experimental evidence has been provided that they are high-affinity Fe²⁺ transporters (for ZIP8 K^M_0.5 ≈ 0.7 μM, for ZIP14 K^M_0.5 ≈ 2.3 μM) when expressed in HEK-293H cells, Sf9 insect cells, or Xenopus laevis oocytes^150–152 (see Table 2). Yet, their subcellular localization in renal PT cells is not clear (see Table 1). Although Wang et al.¹⁵³ described the expression of ZIP8 in the BBM of PT cells by immunofluorescence staining of mouse renal cortical slices, the resolution of the images shown does not warrant this conclusion. Nor is it necessarily supported by the apical localization of the transporters in cell lines overexpressing ZIP8 and ZIP14. The pros and cons of plasma membrane ZIP8 and ZIP14 localization in cell lines and native tissues, such as kidney and liver, have been recently discussed.¹⁴⁷ Based on available evidence, the authors come to the conclusion that endogenous transporters may be more likely expressed in subapical endosomes and other intracellular organelles.¹⁴⁷ Clearly, more work is needed to define the subcellular localization of ZIP8 and ZIP14 in the PT.

6.1.5. Ferroportin (FPN1/SLC40A1). Ferroportin1 (FPN1), also known as iron-regulated transporter 1 (IREG1) or metal transporter protein 1 (MTP1), was independently cloned by three groups in 2000^38–40 (reviewed in ref. 45). To date, FPN1/SLC40A1 is the sole cellular Fe exporter described. Consistent with its assigned function, mammalian FPN1 was found expressed the basolateral pole of duodenal enterocytes and in splenic and hepatic macrophages by immunostaining.^38,39 Expression was also high in the basolateral membrane of the human placental syncytiotrophoblast, which is compatible with a role of FPN1 in Fe transfer to the fetal circulation.³⁹ Interestingly, despite the presence of a functional Fe-responsive element in its 5′ untranslated region,^38,40,154 which allows translational repression of FPN1 by Fe-response proteins under conditions of low cytosolic Fe (see ref. 155 for review), FPN1 was inversely regulated by Fe depletion in mouse duodenum and liver: whereas Fe deprivation resulted in the expected downregulation of FPN1 in liver, duodenal FPN1 was strongly upregulated.³⁸ This apparent paradox was later resolved when a FPN1 transcript lacking the Fe-responsive element and specifically expressed in mouse duodenum was discovered that escapes repression by Fe depletion.¹⁵⁶ Additionally, FPN1 is regulated at the post-translational level by the hepatic hormone hepcidin⁴⁸ (see also Section 2). Dietary Fe overload and inflammation increase, while increased erythropoietic drive/anemia decrease synthesis of hepcidin, which binds to FPN1 and leads to its internalization and subsequent lysosomal degradation, resulting in reduced dietary Fe absorption and Fe release from body Fe stores⁴⁸ (reviewed in ref. 47). FPN1-mediated Fe export into the circulation and subsequent binding to transferrin requires the presence of a ferrioxidase, namely hephaestin in duodenum¹⁵⁷ and ceruloplasmin in other cell types,¹⁵⁸ that convert released Fe²⁺ to Fe³⁺ (reviewed in ref. 159). Although FPN1 has been cloned for ∼15 years, relatively little is known about its functional characteristics: a human FPN1-enhanced green fluorescent protein fusion protein was expressed in Xenopus oocytes and was equally well permeated by microinjected ⁵⁵Fe²⁺ and ⁵⁷Co²⁺, and to some extent by ⁶⁵Zn²⁺.⁴¹ Notably, neither ¹⁰⁹Cd²⁺, ⁶⁴Cu²⁺ nor ⁵⁴Mn²⁺ were transported, even when applied at a concentration of 0.5 mM⁴¹ (see Table 2). FPN1-mediated efflux rate was found to be maximal at slightly alkaline pHo(utside) and abolished at pHo < 6.0, however, the mechanism of the pH effect on FPN1 transport is not understood.⁴¹

Using thoroughly characterized affinity-purified rabbit polyclonal antibodies against rat FPN1, we have previously reported that FPN1 is expressed in rat PT (S2 > S1 > S3) where it is mainly localized in the basolateral plasma membrane (and some intracellular vesicles), as evidenced by immunohistochemistry and immunogold electron microscopy at high magnification.¹⁶⁰ Interestingly, FPN1 was absent from glomeruli and DT. Iron loading resulted in increased surface expression of FPN1 in a rat renal PT cell line, as detected by immunofluorescence labeling of non-permeabilized cells as well as surface biotinylation experiments, but with no change in total cellular FPN1 expression, suggesting that FPN1 redistributes to the cell surface and that increased insertion of FPN1 into the plasma membrane may play a role in protecting PT cells from Fe overload.¹⁶⁰ The basolateral localization of FPN1 in PT was subsequently confirmed in hepcidin^{(−/−) 123} and heme oxygenase 1^(−/−) mice¹⁶¹ using commercial antibodies, but FPN1 expression was much weaker in control animals. In contrast to those studies, Veuthey et al. showed both apical and basolateral FPN1 distribution in the mouse PT,¹⁴⁰ and FPN1 was found only at the apex of PT cells in another mouse study.¹⁶² In addition to the poor resolution of the images shown in these mouse studies, the specificity and quality of the antibodies used is difficult to assess as they were either from commercial sources and/or poorly characterized (Dr B. Galy, European Molecular Biology Laboratory, Germany; personal communication) (information summarized in Table 1). There is also evidence to suggest that renal FPN1 expression is regulated by hepcidin: intraperitoneal hepcidin pre-injection (24 h) prevents FPN1 upregulation induced by ischemia-reperfusion injury, as demonstrated in whole membranes of mouse kidney.¹⁶³ Furthermore, intraperitoneal hepcidin injection in mice induces a rapid (1 h) degradation of FPN1 in kidney homogenates (Drs R. P. L. van Swelm & D. W. Swinkels, Department of Laboratory Medicine, RUMC, Nijmegen, The Netherlands, personal communication; manuscript submitted).

6.2. Iron transporters of the loop of Henle (LOH)

6.2.1. DMT1 (SLC11A2). Very few studies have investigated the role of the LOH in Fe transport. Wareing et al.¹³ performed two types of experiments: they used tracer microinjections of ⁵⁵FeCl₃ in early PCT or early DCT of rat kidney in vivo, determined the percentage of urinary ⁵⁵Fe recovery (18.5 ± 2.9% for PCT versus 46.1 ± 6.1% for DCT) and by interpolation of the data calculated that the LOH contributes to ∼40% of the total measured Fe transport. In addition, the authors microperfused LOHs in vivo with 7 μM ⁵⁵FeCl₃ by placing the perfusion pipette at the last accessible convolution of the PCT and collected the perfusate at the first accessible portion of the DCT, the LOHs being isolated from the rest of the tubule by injection of mineral oil blocks into the tubule lumen.¹³ By this approach they found that 52.7 ± 8.3% of perfused Fe was recovered from the DCT. This indicated that the LOH can reabsorb significant amounts of Fe. Furthermore, the same group found DMT1 expressed in the thick ascending limbs (TAL) of rat LOHs which exhibited punctate, DMT-1-specific immunoreactivity at the apical membrane and, more intensely, in the cytoplasm, and the intensity of staining increased progressively toward the DCT.¹⁴ This suggests that apical DMT1 in the TAL cells of the LOH may reabsorb Fe. Interestingly, Fe overload in hepcidin^(−/−) mice leads to increased Fe accumulation in TAL cells of the LOH as well as to increased basolateral FPN1 expression (see Section 6.2.2),¹²³ suggesting that TAL cells of the LOH express a pathway for apical Fe uptake that could represent DMT1.

6.2.2. Ferroportin (FPN1/SLC40A1). The Fe efflux transporter FPN1 has been detected basolaterally and intracellularly in the thick ascending limb of the LOH in hepcidin^(−/−) mice, but FPN1 expression was weak in the kidney cortex or medulla of control mice.¹²³ This study is in contrast to another study in mice in which no immunostaining of FPN1 was found in the LOH (the possible reasons for the discrepancies in both mouse studies have been discussed in Section 6.1.5 and Table 1).¹⁴⁰ In our own studies in the rat kidney, we mainly focused on the FPN1 distribution in PT,¹⁶⁰ but FPN1 was also expressed at low levels in the medulla, especially in the inner medulla (that includes the thin limbs of the LOH), as determined by immunoblotting.

6.3. Iron transporters of the distal tubule (DT)

6.3.1. TRPV5 Ca²⁺ channels. TRPV5 (epithelial Ca²⁺ channel 1 ECaC1) belongs to the vanilloid (V) family of the transient receptor channel (TRP) superfamily. In humans, TRPV5 is considered the renal isoform of that family. The human TRPV5 (hTRPV5) gene encodes 729 amino acids, along with a predicted molecular mass of around 83 kDa. In the kidney, TRPV5 is localized at the apical membrane of DCT and connecting tubules where it contributes to active Ca²⁺ reabsorption¹⁶⁴ (see Table 1). TRPV5 is a highly Ca²⁺-selective (permeability_Ca²⁺:permeability_Na⁺ > 100), strongly inward rectifying cation channel with a single channel conductance between 55 and 107 pS.^165,166 TRPV5 was shown to be permeable for Ca²⁺, Ba²⁺, Sr²⁺ and Mn²⁺ and inhibited by several di- and trivalent cations.^165,166 The expression of functional TRPV5 is regulated by several hormones such as parathyroid hormone, and 1.25 dihydroxy vitamin D at the transcriptional level^164,167 (reviewed in ref. 168). An orthologue to mammalian TRPV5 was cloned from the gill of pufferfish (Fugu rubripes) and characterized.¹⁶⁹ The F. rubripes ECaC (FrECaC) protein displays all structural features typical for mammalian ECaC. Functional expression of FrECaC in Madin-Darby canine kidney (MDCK) cells confirmed that the channel mediates Ca²⁺ influx, but FrECaC was also permeable to Fe²⁺ (and even better to Zn²⁺). Bulk Fe flux was measured with ascorbic acid to prevent oxidation of ⁵⁹Fe²⁺ to ⁵⁹Fe³⁺, and a modest increase of Fe influx was observed when flux buffer contained 0.24 μM ⁵⁹Fe²⁺ (without Ca²⁺).¹⁶⁹ Thus FrECaC (and possibly renal TRPV5) may serve as a pathway for Fe acquisition.

6.3.2. Ca_v3.1 Ca²⁺ channels. Ca_v3.1 is a T (transient opening)-type Ca²⁺ channel, also known as α_1G. T-type channels differ from the L(long lasting)-type Ca²⁺ channels due to their ability to be activated by more negative membrane potentials, their small single channel conductance, and their unresponsiveness to Ca²⁺ antagonist drugs.^170,171 As a member of the Ca_v3 subfamily of voltage-gated Ca²⁺ channels, T-type channels are important for the repetitive firing of action potentials in cells with rhythmic firing patterns such as cardiac muscle cells and neurons in the thalamus of the brain. Ca_v3.1 channels are widely expressed in excitable and non-excitable cells, including brain, ovary, placenta, heart, liver, bone, endocrine system and vascular smooth muscle.^170,172,173 Although Ca_v3.1 channels expression in the kidney has been primarily associated with the renal vasculature¹⁷⁴ one study revealed Ca_v3.1 expression in the DCT, in the connecting tubule and cortical collecting duct (CCD), and inner medullary collecting duct (IMCD) principal cells¹⁷⁵ (see Table 1). Using selective blockers, several reports have proposed that T-type Ca²⁺ channels are involved in steroid hormone-dependent luminal ⁴⁵Ca²⁺ uptake in isolated rabbit DCT.^176–178

Using a calcein-AM fluorescence assay to detect Fe in the cytosol under various Fe loading conditions, T-type calcium channels have been implicated in Fe²⁺ uptake by cardiomyocytes through the use of selective blockers.¹⁷⁹ In a more detailed study, Lopin et al.¹⁸⁰ examined the effects of extracellular Fe²⁺ on permeation and gating of Ca_v3.1 channels stably transfected in HEK293 cells, using whole-cell patch-clamp electrophysiology recording. In the absence of extracellular Ca²⁺, Fe²⁺ carried detectable, whole-cell, inward currents at millimolar concentrations (73 ± 7 pA at −60 mV with 10 mM extracellular Fe²⁺). With a two-site/three-barrier Eyring model for permeation of Ca_v3.1 channels,¹⁸¹ the authors estimated a transport rate for Fe²⁺ of ∼20 ions per s for each open channel at −60 mV, with 1 μM extracellular Fe²⁺ and in the presence of physiological Ca²⁺ concentrations (2 mM extracellular Ca²⁺). Reversal potentials indicated a Fe²⁺/Ca²⁺ permeability ratio of 0.06–0.18. Because Ca_v3.1 channels exhibit a significant “window current” at resting membrane voltage (open probability, ∼1%), the authors concluded that Ca_v3.1 channels represent a likely pathway for Fe²⁺ entry into cells at resting membrane potentials and possibly during the course of action potentials in excitable cells with clinically relevant concentrations of extracellular Fe^2+ 180 (see Table 2).

6.3.3. DMT1 (SLC11A2). The divalent metal transporter DMT1 (see Sections 6.1.3 and 6.2.1) has been found expressed in the luminal membrane of DCT. The most convincing localization study was performed by Ferguson et al.¹⁴ in rat kidney. Using an affinity-purified rabbit polyclonal antibody directed against a 21-amino acid region in the NH₂ terminus of rat DMT-1 that should recognize all known DMT-1 isoforms,¹³⁷ the authors showed extensive co-localization of DMT-1 and thiazide-sensitive Na⁺–Cl⁻ cotransporter (NCCT) in the apical membrane of DCTs. DMT-1 was absent in late DCT to early connecting segments. Furthermore, in another study expression of DMT1 in the apical membrane and subapical region of rat DCT showed an inverse correlation with the dietary Fe content.¹⁵ In support of a DCT localization of DMT1, in vivo tracer microinjection of ⁵⁵Fe into early rat DCT was associated with less than 50% urinary recovery, suggesting Fe reabsorption by late DCT segments and/or CD.¹³ In contrast, two other studies have failed to confirm expression of DMT1 in DCT of the mouse.^138,140 However, the immunohistochemical images used in those mouse studies showed poor resolution, and no attempt was made to identify the DMT1 labeled nephron segments with segment-specific markers.

6.3.4. Lipocalin-2 receptor (SLC22A17). The NGAL/24p3/lipocalin-2 receptor (Lip2-R) co-localizes with calbindin, a marker for late DCT and connecting tubule, in mouse and rat kidney.¹⁸² The majority of DCT cells demonstrated co-localization of the two proteins, Lip2-R apically and calbindin intracellularly. Other cells however, were Lip2-R-positive but calbindin-negative, suggesting that Lip2-R is expressed in both early and late DCT. Since Lip2-R is predominantly expressed in CD of rodent kidney it will be discussed in that section (see Section 6.4.2).

6.4. Iron transporters of the collecting duct (CD)

6.4.1. DMT1 (SLC11A2). Wareing et al.¹³ attempted to identify the distal sites of renal Fe reabsorption by in vivo tracer microinjection of ⁵⁵FeCl₃ into the DCT segment of the rat nephron in vivo. Approximately 50% of the injected ⁵⁵Fe was recovered in the urine, suggesting that Fe is significantly reabsorbed by nephron segments distal to the DCT. This functional mapping of the distal nephron sites of Fe reabsorption is in good agreement with the immunofluorescence distribution of DMT1 in nephron segments in subsequent studies by the same group. Ferguson et al.^14,145 and Wareing et al.¹⁵ demonstrated strong DMT-1-specific immunofluorescence in the cortical and outer medullary CD. The signal gradually decreased in intensity from the cortex to the outer stripe and inner stripe of the medulla. The distribution of DMT1 in different cell types of the CD varied considerably: co-localization with the water channel aquaporin 2 showed that DMT-1 is present apically and intracellularly in principal cells of CD in the cortex and outer medulla. In superficial cortex, DMT-1 also co-localized with the vacuolar-type H⁺-ATPase at the apical membrane, thus indicating expression in A-intercalated cells, and also showed a bipolar distribution in some, but not all, B-intercalated cells. In the outer medullary region, DMT-1 was less intense at the apical membrane and more diffuse throughout the cytosol in intercalated cells. DMT-1 immunoreactivity decreased progressively along the length of the CD, and inner medullary CD ducts showed only faint DMT-1-specific staining.¹⁴ Another study confirmed DMT-1 in the renal medulla in mice,¹⁴⁰ but no information was provided on the cell types associated with DMT-1 in renal CD. The apical expression of DMT1 in type A intercalated cells of the cortical CD is interesting from a physiological point of view considering that Fe²⁺ transport by DMT1 is coupled to H⁺.¹⁰ Type A-intercalated cells could provide the pH gradient necessary to drive DMT1-mediated luminal Fe²⁺ uptake. Furthermore, the localization of DMT1 in type A-intercalated cells would be compatible with the recently postulated function of these CD cells as a critical barrier against infection¹⁸³ by depleting Fe from the urine that is necessary for bacterial growth. Hence, Fe²⁺ clearance from the lumen mediated by H⁺-driven Fe²⁺ uptake via DMT1 would represent another defense mechanism in addition to the suggested secretion of the Fe-bacterial siderophore sequestering peptide lipocalin-2/24p3/NGAL (neutrophil gelatinase-associated lipocalin) and urinary acidification as processes being responsible for bacteriostasis¹⁸³ (see Section 6.4.2).

6.4.2. Lipocalin-2 receptor (SLC22A17/Lip2-R). Neutrophil gelatinase-associated lipocalin (NGAL [human]/siderocalin/24p3 [rodent]/lipocalin-2 [human]) (Lip2) was discovered in neutrophils¹⁰⁵ and was also shown to be induced in intestinal epithelia by inflammation or cancer.¹⁸⁴ Lip2 binds Fe³⁺ through association with bacterial¹⁸⁵ and mammalian siderophores,^73,186 thereby affecting Fe homeostasis of target cells and their survival and proliferation. Hence, Lip2 may play a role as an Fe-sequestering protein in antibacterial innate immunity by decreasing susceptibility to bacterial infections,^185,187,188 and its interactions with bacterial siderophores have been very well characterized.¹⁸⁹ Lip2 may also deliver Fe to epithelia of the primordial kidney,¹⁹⁰ stimulate growth and differentiation, and promote repair and regeneration of damaged epithelia.¹⁹¹ Therefore, Lip2 is increasingly used as a sensitive biomarker of kidney damage in clinical settings.^192,193 During renal insults, e.g. acute kidney injury (AKI), Lip2 is thought to be secreted by the distal nephron (DCT and CD) and excreted into the urine¹⁹⁴ although earlier studies from the same laboratory had emphasized that Lip2 is secreted by the PT during AKI.^191,195 It has been postulated that Lip2 is secreted, possibly to limit injury and promote Fe-dependent regeneration of damaged epithelia,¹⁹¹ but how this happens is unclear. Despite a wealth of publications, the function of Lip2 in the kidney in health and disease as well as its mechanisms of secretion are still not well understood.

The mounting relevance of Lip2 in the medical field has increased the interest in identifying putative receptors of this ligand. Megalin, the epithelial multi-ligand receptor expressed in renal PT (see Section 6.1.1) binds Lip2 with high affinity.¹¹² In addition, a receptor for murine Lip2, Lip2-R, has also been cloned¹⁹⁶ whose mRNA encodes 520 amino acids (molecular mass ∼60 kDa and 11 or 12 transmembrane domains depending on the predicted topology) and whose affinity for Lip2 is ∼1000× higher (K_D ∼ 90 pM)¹⁹⁷ than that of megalin (K_D ∼ 60 nM).¹¹² According to the SLC (solute carrier) nomenclature system this receptor is also named SLC22A17 or BOCT (brain organic cation transporter).¹⁹⁸ However, classical substrates of organic cation transporters are not transported by SLC22A17 (ref. 199 and N. A. Wolff & F. Thévenod; unpublished). Interestingly, several short N- and C-terminal splicing variants (22 kDa and ∼30 kDa, respectively) of the Lip2-R have been described in humans and rodents, respectively,^196,200 but their function in health or disease is unknown. Although Lip2-R protein is expressed in the kidney¹⁹⁶ its localization and functions in that organ were unknown until recently. Using two affinity-purified polyclonal rabbit antibodies directed against the N- and C-terminal domains of Lip2-R, we showed apical expression of Lip2-R in rodent kidney DCT (where it co-localized with calbindin, Lip2-R being expressed apically and calbindin intracellularly) and CD (mainly inner medullary CD), but not in PT (where it was found weakly expressed intracellularly). In DCT, some cells were Lip2-R-positive but calbindin-negative, suggesting that Lip2-R is expressed in both early and late DCT (see Table 1). Lip2-R was also found expressed in respective mouse cell lines (mDCT209; mIMCD3, mCCDcl1), but not in PT cell lines (WKPT-0293 Cl.2) (ref. 182 and unpublished). We also confirmed the expression of several immunoreactive protein bands in purified plasma membranes by immunoblotting (MM ∼35 kDa, ∼45 kDa, ∼60 kDa and ∼130 kDa), thus confirming the presence of “short” and “long” forms of the protein that may represent splicing variants or dimers of the receptor, respectively (ref. 182 and unpublished). Chinese hamster ovary (CHO) overexpressing Lip2-R or mDCT209 cells expressing Lip2-R endogenously internalized submicromolar concentrations of fluorescence-labelled Tf, albumin, or MT and their uptake was blocked by 500 pM Lip2,¹⁸² which confirms that the uptake of these proteins is mediated by the Lip2-R. Using microscale thermophoresis, a powerful technique to quantify biomolecular interactions,²⁰¹ we showed that MT binds to Lip2-R with a K_D of ∼100 nM.¹⁸² Hence, Lip2-R seems to bind proteins filtered by the kidney, including Tf and MT, with high affinity and may contribute to receptor-mediated endocytosis of these proteins as well as of Lip2 in the distal nephron (see Table 2).

Is the uptake of metalloproteins, such as Tf, Lip2 or MT, by Lip2-R physiologically and pathophysiologically relevant when bulk protein reabsorption is thought to take place in the PT? Experimental evidence has demonstrated that physiologically a small but significant proportion of filtered proteins is reabsorbed by the distal segments of the nephron.^202–205 Although megalin:cubilin:amnionless is a high-capacity receptor complex for endocytotic reabsorption of filtered proteins,²⁰⁶ some proteins/metalloproteins may bypass reabsorption in the PT, either as the consequence of their low affinity to megalin and low concentration in the ultrafiltrate (e.g. MT with a K_D of ∼5–100 μM²⁰⁷ but a plasma concentration of ∼0.5–5 nM^208,209) (in this context see Table 3) or due to limited reabsorptive capacity of the system (e.g. following glomerular or PT damage and ensuing proteinuria).^17,210,211 A high-affinity protein receptor in the distal nephron such as Lip2-R could contribute to exhaustive protein/metalloprotein reabsorption and deplete the final urine from protein-bound Fe (and other metals) under physiological conditions, or limit losses associated with renal diseases, including various forms of inherited or acquired Fanconi syndrome.¹⁷ Indeed, two in vivo studies have demonstrated Fe uptake into the distal nephron of nephrotic rats²¹² or following glomerular damage induced by acute Fe overload.²¹³ Interestingly, Fe deposits were found in lysosomes of DT by electron microscopy²¹² and kidney medullary tubule cells by histochemistry.²¹³ Furthermore, in hepcidin^(−/−) mice, a model of the Fe overload disease hemochromatosis, Fe deposits were also found in the distal nephron.¹²³ Hence, increased uptake of proteins/metalloproteins by Lip2-R in the distal nephron could initiate or enhance kidney injury. Along these lines, a recent in vivo study has implicated the Lip2-R in the CD in contributing to initiation and/or aggravation of renal inflammation and fibrosis in response to proteinuria.²¹⁴

Correnti et al.⁷⁴ have recently questioned a role of Lip2 in cellular Fe metabolism based on their observation that gentisic acid (a putative mammalian siderophore) could not form a stable ternary complex with Lip2 and Fe and on their inability to demonstrate any physical interaction between Lip2 and N-(NTD) or C-terminal domains (CTD) of mouse Lip2-R by surface plasmon resonance analyses. However, using the 105 residue NTD of human Lip2-R and analysis of its interaction by microscale thermophoresis, isothermal titration calorimetry and nuclear magnetic resonance, we could demonstrate binding of human Lip2 to its cellular receptor NTD (A.-I. Cabedo Martinez et al.; submitted). Although the affinity we measured between human Lip2-R-NTD and human Lip2, i.e. ∼7 μM for apo-Lip2 and ∼20 μM for holo-Lip2 (Lip2 bound to the bacterial siderophore enterobactin) suggests that the N-terminus alone cannot account for the internalization of Lip2 by Lip2-R and that other parts of the receptor must contribute to the interaction, our results are in contradiction with the conclusions of Correnti et al.⁷⁴ We suspect that their failure to observe any direct interaction between Lip2 and mouse Lip2-R results from (1) their inability to control the state of their recombinant Lip2 (apo- or holo-) and (2) a lack of proper formation of the disulfide bridges of their mouse Lip2-R-NTD preparation, as the formation of aberrant disulfides would probably lead to forms of mouse Lip2-R-NTD that are unable to bind to Lip2 (A.-I. Cabedo Martinez et al.; submitted). Either or both of these points could explain their inability to observe an interaction between Lip2 and mouse Lip2-R-NTD. Overall, our data suggest that Lip2-R represents a high-affinity multiligand receptor for apical endocytosis of proteins and/or metalloproteins (such as Tf or Cd²⁺–MT) in renal epithelia. Increased endocytosis subsequent to glomerular and/or PT damage may promote renal epithelial damage by death, inflammation and fibrosis.

6.4.3. Ferroportin (FPN1/SLC40A1). Strong FPN1 (see Sections 6.1.5 and 6.2.2) immunostaining has been detected in inner medullary CD of mice but FPN1 expression decreased in anemic mice.¹⁴⁰ Specific immunostaining was found intracellularly. Outer medulla showed intracellular staining as well. In contrast, no medullary FPN1 staining was detected in another study in mice¹²³ and FPN1 expression was weak in inner medulla of rat kidney when measured by immunoblotting in our own studies.¹⁶⁰ The discrepancies observed in these mouse studies have been discussed in Section 6.1.5 and Table 1.

7. Cadmium toxicity

7.1. General considerations and link to iron transport

Pollution by cadmium (Cd) is rising worldwide because of intensified industrial activities that have increased its availability and because Cd cannot be degraded further.^7,215 Chronic exposure to low Cd concentrations is a significant health hazard for ∼10% of the world population that increases morbidity and mortality.²¹⁶ Indeed, Cd damages multiple organs in humans and other mammalian organisms by causing nephrotoxicity, osteoporosis, neurotoxicity, genotoxicity, teratogenicity, or endocrine and reproductive defects.²¹⁷

In mammalian organisms, Cd is a toxic-only element with no known role in physiological processes: as a non-essential metal ion Cd²⁺ competes with essential metal ions in cells where it disrupts cellular functions and leads to disease. Although Cd²⁺ is not capable of catalyzing Fenton chemistry in biological systems, it may initiate free radical chain reactions by depleting endogenous redox scavengers, inhibiting anti-oxidative enzymes, blocking the mitochondrial electron transport chain, and/or displacing redox active metals, such as Fe²⁺ or Cu²⁺ from their carrier proteins²¹⁸ and thereby trigger cell death by apoptosis (reviewed in ref. 219). Cd²⁺ can also substitute for Ca²⁺ in cellular signaling or for Zn²⁺ in many enzymes and transcription factors which may account for some of the biological effects of Cd²⁺.^219,220 In order for toxicity to occur Cd²⁺ must first enter cells by utilizing transport pathways for essential metals, such as Fe²⁺, Zn²⁺, Cu²⁺, Ca²⁺ or Mn²⁺, that are present in biological systems mostly as complexes with small organic molecules or as metalloproteins. These metal ion compounds are hydrophilic and must permeate lipophilic cellular membranes through intrinsic proteinous pathways. Hence, free or small complexed metal ions may be transported via ion channels or carrier proteins whereas metalloproteins are taken up by receptor-mediated endocytosis (RME). Cd²⁺ has similar physico-chemical properties as essential metal ions (for a detailed account see ref. 18 and references therein) and Cd²⁺ complexes are analogous to endogenous biological molecules, therefore this attribute has been termed “ionic and molecular mimicry”.^4,221 Hence, transport (and toxicity) of Cd²⁺ can only occur if cells possess pertinent transport pathways for essential metals or biological molecules. A number of pathways has been suggested to allow Cd²⁺ entry in excitable and non-excitable cells⁹ and the most likely candidates have been recently reviewed.^18,215

Chronic exposure to Cd²⁺ involves very low concentrations of Cd²⁺ that originate from environmental pollution and mainly results from dietary sources and cigarette smoking. Hence Cd²⁺ enters the body primarily through the lungs and the gastrointestinal (GI) tract: the absorption of Cd²⁺ from the lungs is much more effective than that from the gut; however, Cd²⁺ absorption from the GI tract is the main route of Cd²⁺ exposure in humans.²¹⁵ Following absorption in the lungs and/or intestine, Cd²⁺ in the blood at first largely binds to albumin and other thiol-containing HMWP and low molecular weight proteins (LMWP) in the plasma, including MT, as well as to blood cells. But Cd²⁺ tends to concentrate in blood cells (mainly erythrocytes) and <10% remains in the plasma.²²² Since intravenously injected MT-bound Cd²⁺ in mice is quickly cleared from the plasma by the kidneys²²³ this protein fraction in the circulation – that is assumed to originate from Cd²⁺ stored in liver cells as Cd²⁺–MT and is released from damaged cells (see below) – has been thought to be of great importance for the transport of Cd²⁺ to the kidney during long-term exposure^224–226 (although the plasma Cd²⁺–MT concentrations following experimental injections exceeded physiological MT concentrations by >2000-fold;^208,209 see Section 7.2 for a critical discussion). The blood level of Cd²⁺ largely reflects recent Cd²⁺ exposure with a half-life of 75–128 days.²²⁷ It ranges between 0.03 and 0.5 μg l⁻¹ (∼0.3–5 nM) depending on the preparation method and the populations studied (reviewed in ref. 228) and its concentration in plasma will be at least ten-fold lower.²²²

Cd²⁺ reaching the plasma is thought to be initially transported to the liver where intracellular Cd²⁺ induces the synthesis of the endogenous detoxicant MT, which binds, sequesters and detoxifies Cd²⁺ because its affinity to Cd²⁺ is very high with a K_D of ∼10⁻¹⁴ M (reviewed in ref. 229). Yet, a small proportion of liver (Cd²⁺–)MT is assumed to be slowly released into blood plasma as the hepatocytes in which Cd²⁺ is sequestered die off, either through normal turnover or as a result of Cd²⁺ injury.^224,230,231 Several studies have demonstrated that following long-term exposure to Cd²⁺ and even at long time intervals after a single exposure, the level of Cd²⁺ is initially highest in the liver and then gradually increases in the kidneys.^232,233 The strongest evidence for the concept that the major source of renal Cd²⁺ during chronic Cd²⁺ exposure is derived from hepatic Cd²⁺, which is transported in the form of Cd²⁺–MT in blood plasma, was derived from studies with transplanted livers of Cd²⁺-exposed rats to normal rats.²³⁴ Cd²⁺ and MT in the liver of recipient rats decreased over time after surgery whereas renal Cd²⁺ and MT levels increased and most of the Cd²⁺ in the kidney was bound to MT.²³⁴ Although none of these data proved that redistribution of Cd²⁺ from the liver to the kidney is mediated by circulating Cd²⁺–MT, this hypothesis still prevails in the literature (see Section 7.2 for a critical discussion).

Once absorbed Cd²⁺ is stored in various organs, including the kidneys and liver, with a half-life of several decades.^2,7,215 This happens because Cd²⁺ induces the expression of detoxifying molecules that form a complex with the metal ion and thereby alleviate its toxic effects. But this apparently beneficial effect is a two edged-sword because these seemingly harmless Cd²⁺ complexes represent an endogenous source of high concentrations of potentially toxic Cd²⁺. The major detoxifying tool of the cell for Cd²⁺ complexation is MT. MTs are low-molecular weight (MM ranging from 3.5–14 kDa), cysteine-rich metal-binding proteins that have the capacity to bind both physiological Zn²⁺ ions and toxic Cd²⁺ ions through the thiol group of their cysteine residues that represent nearly 30% of its amino acidic residues.^100,235,236

7.2. Cadmium handling by the kidney

As a consequence of its storage in tissues Cd²⁺ is very poorly excreted, mainly in urine and feces. With low, or even moderate, levels of exposure, little or no Cd²⁺ is excreted in the urine,²³⁷ which indicates that Cd²⁺ is reabsorbed and stored by the kidney. In humans, the amount of Cd²⁺ excreted daily in urine represents only about 0.005–0.015% of the total body burden²³⁷ and amounts to 0.05–0.2 μg l⁻¹ (reviewed in ref. 228). Most of the Cd²⁺ in urine is bound to MT^238,239 and it is assumed that urinary Cd²⁺ and MT stem from filtered Cd²⁺–MT and normal turnover and shedding of epithelial cells, or – perhaps – from exosomes derived from Cd²⁺–MT containing tubule epithelia. This supposition is based on chronic studies in several mammalian species showing that urinary excretion of Cd²⁺ increases slowly for a considerable time as a reflection of the level of Cd²⁺ exposure and the body burden of the toxic metal ion, which correlates with an increase of Cd²⁺ in the renal cortex (reviewed in ref. 240). But when the concentration of Cd²⁺ in the renal epithelial cells reaches a threshold value of ∼150–200 μg g⁻¹ wet weight Cd²⁺ disrupts tubular reabsorptive processes and the excretion of Cd²⁺ and MT begins to increase more strongly in a linear manner, which is associated with the onset of polyuria and proteinuria^241,242 (reviewed in ref. 240). When kidney dysfunction aggravates and a sharp increase in excretion of Cd²⁺ and MT occurs, a decrease in renal and liver Cd²⁺ concentrations is also observed.^243,244 Hence, the early, slow linear phases of Cd²⁺ and MT excretion likely mirror the level of chronic Cd²⁺ exposure whereas the later sharp increases in excretion reflect Cd²⁺-induced tubular injury.

In the previous paragraphs it has been emphasized that chronic exposure to low environmental or dietary Cd²⁺ concentrations results in accumulation of the metal ion in the kidney with a biological half-life of ∼20 years or more^2,7,215 where it may cause damage, fibrosis or failure,^245,246 or – with Cd²⁺ being a class 1 human carcinogen – cancer.²⁴⁷ In contrast, acute or subchronic Cd²⁺ nephrotoxicity is associated with a general transport defect of the PT that mimics the de Toni-Debré-Fanconi-syndrome^248,249 with proteinuria, aminoaciduria, glucosuria and phosphaturia (for review, see ref. 250).

For several decades the following scenario has prevailed to account for acute or chronic Cd²⁺ toxicity in the kidney: it has been presumed that Cd²⁺ in the circulation is filtered by the glomerulus because of the small molecular mass of most circulating Cd²⁺ forms: in the plasma, Cd²⁺ is thought to be loosely associated with molecules, such as LMWP – e.g. β-2 microglobulin, α-1 microglobulin, retinol-binding protein, insulin or parathyroid hormone – with amino acids or the sulfhydryl compounds GSH or cysteine, or tightly bound to specific metal-binding proteins such as the LMWP MT.²²⁹ Several HMWP, e.g. albumin, bind Cd²⁺ with low affinity,²⁵¹ also show some degree of glomerular filtration¹⁷ and may therefore carry Cd²⁺ into the ultrafiltrate. Furthermore, the Fe-binding protein Tf that is filtered by the glomerulus and ferritin (see Section 5) may also bind Cd²⁺ in plasma.^252–254 The PT largely contributes to the reabsorption of Cd²⁺ because as the first segment of the nephron it is responsible for bulk reabsorption of primary urine, which mainly takes place by solvent drag via paracellular routes (see Section 4). But PT cells also possess apical transporters (as proposed for ZIP8 and ZIP14 transporters that may carry both Fe²⁺ and Cd²⁺;^148,149 however see Section 6.1.4 and ref. 147 for a note of caution), amino acid transporters, metabolizing brush-border enzymes (such as γ-glutamyl transpeptidase that degrades GSH), and the receptor for protein endocytosis megalin:cubilin:amnionless¹⁰⁸ that may mediate apical uptake of Cd²⁺ ions and Cd²⁺ complexes (see Section 6.1.1, and ref. 9 and 18 for reviews). There is also evidence that Cd²⁺ is taken up at the basolateral surface of PT cells^255,256 and it has recently been shown to take place via the organic cation transporter 2 (OCT2).^257,258 Although it is mechanistically remarkable that an organic cation transporter is able to carry a divalent metal ion as a substrate, a K_m value of ∼54 μM for Cd^2+ 258 suggests that the in vivo toxicological relevance of this transporter is questionable.

Like other LMWP, Cd²⁺–MT/MT is thought to be reabsorbed from primary urine into PT cells of the kidneys by megalin:cubilin:amnionless receptor-mediated endocytosis^{206,259–261} (see Table 3). Studies with cultured PT cells have provided evidence that Cd²⁺–MT/MT is trafficked to acidic late endosomes and lysosomes^262,263 where MT may be degraded by lysosomal proteases whereas Cd²⁺ may exit the endosomal/lysosomal compartment by DMT1-mediated efflux into the cytosol.^137,264 This may cause acute PT toxicity in cases where PT cells would have to handle high concentrations of endocytosed Cd²⁺–MT.²⁶⁴ However, if the Cd²⁺–MT stress is low PT cells may adapt by inducing the upregulation of detoxifying proteins, including MT²⁶⁵ that inactivate and complex Cd²⁺ released from lysosomes into the cytosol for long-term storage.²⁶⁶ Cd²⁺ accumulation in the PT (and storage as Cd²⁺–MT) may be likely further promoted by the absence of an efflux pathway for cytosolic Cd²⁺ into the extracellular fluid or blood plasma because FPN1 that is expressed at the basolateral cell side of PT cells¹⁶⁰ does not transport Cd²⁺ (as opposed to Fe²⁺)⁴¹ (see also Section 6.1.5).

The concept that endocytosis of filtered Cd²⁺–MT by megalin:cubilin:amnionless is mainly responsible for accumulation of Cd²⁺ in the PT was based on studies demonstrating redistribution of hepatic Cd²⁺ to the kidney that was supposed to be Cd²⁺–MT,^232–234 on in vivo animal studies with intravenously injected Cd²⁺–MT^{223,267–272} as well as on microinjections of Cd²⁺–MT in isolated PT.²⁷³ It was confirmed and elaborated in cell culture studies^260–263 (reviewed in ref. 274). However, all in vivo and cell culture studies applied Cd²⁺–MT at micromolar concentrations. Meanwhile surface plasmon resonance analyses, cell culture and in vivo studies have established that the binding affinity of megalin for MT is ∼100 μM,^207,260,272 which is compatible with the observations from the in vivo animal studies. But considering that plasma concentrations of MT are in the range of ∼0.5–5 nM^208,209 in humans (and healthy laboratory animals), the concept that filtered (Cd²⁺)–MT is also taken up by PT via megalin:cubilin:amnionless-dependent endocytosis under physiological conditions²⁷⁵ is unfounded, and thus the current models of Cd²⁺ accumulation (and chronic toxicity) in the PT should be revised (nevertheless Cd²⁺–MT may still be useful as a model compound to study RME of Cd²⁺–protein complexes in cell culture or in vivo as the high binding affinity of MT to Cd²⁺ precludes dissociation of toxic Cd²⁺ ions). It is more likely that other LMWP (e.g. α1- or β2-microglobulin) and albumin, which also bind Cd²⁺ and reach submicromolar concentrations in the ultrafiltrate,^276,277 are more relevant ligands (e.g. β2-microglobulin binds to megalin with a K_D of ∼0.42 μM²⁷⁸) (see also Table 3) that are endocytosed by megalin:cubilin:amnionless to induce Cd²⁺ accumulation and eventually PT toxicity. It could be argued that these proteins exhibit relatively low affinities to Cd²⁺ compared to MT (reliable K_D values of ∼10⁻⁶ M for Cd²⁺ and other divalent metal ions are only available for albumin and β2-microglobulin^251,279), indicating that at steady-state maximally 1% of these proteins in the circulation will form complexes with blood Cd²⁺ (with a concentration of 0.3–5 nM²²⁸) whereas MT in the circulation will be Cd²⁺-saturated (based on equivalent low nM concentrations of MT and Cd²⁺ and a K_D of ∼10⁻¹⁴ M²²⁹). Yet the relatively high concentration of microglobulins and albumin in the primary filtrate and their high binding affinity to megalin (see Table 3) combined with the multiplicative effect of their continuous glomerular filtration makes them more prone to accumulate in the PT and contribute to chronic renal PT toxicity than Cd²⁺–MT whose concentration in the primary filtrate is at least 10⁵-times lower^208,209 than its K_D for megalin binding.²⁰⁷

Because only 0.02–03% of filtered proteins, including MT (based on measured values for plasma and urinary MT 99.7% of filtered MT must be reabsorbed by the kidney^208,209), are excreted with the urine²⁸⁰ (reviewed in ref. 281) additional uptake pathways for proteins and protein–Cd²⁺ complexes must exist in the distal nephron (see below).

Cd²⁺ may not be only toxic to PT cells, but also to glomeruli and the distal nephron.²⁸² Glomerular damage with a decreased GFR has been observed in occupationally exposed workers²⁸³ and in environmentally exposed populations where it may occur at similar Cd²⁺ dose levels as the tubular damage.^246,284 But overall, the pathogenesis of the glomerular lesion in Cd²⁺ nephropathy is not well understood.²⁸⁵ Downstream segments of the nephron, both in the cortex and medulla, also exhibit a high permeability to Fe²⁺ and other metal ions (see Sections 6.2–6.4) and could hence contribute to uptake of Cd²⁺. This would be particularly the case when proximal segments of the nephron are defective or “overwhelmed” as a consequence of increased filtration (e.g. due to glomerular damage). Under those circumstances, later segments should become more relevant for uptake. As an example, a chronic in vivo study in ducks demonstrated substantial damage to glomerular podocytes following exposure to a combination of lead, methylmercury and cadmium that was associated with enhancement of degenerative changes in PT and CD; in contrast exposure to cadmium alone showed no podocyte damage and tubular damage was restricted to PT whereas the CD was not affected.²⁸⁶ Sporadic evidence for chronic Cd²⁺ toxicity of the distal portions of the nephron induced by Cd²⁺ exposure has also been obtained, both in experimental animals^287,288 and in Cd²⁺-exposed workers,²⁸⁹ but the mechanisms of distal nephron damage remain unclear.

In agreement with studies demonstrating in vivo⁵⁵Fe transport in the rat LOH¹³ that may be mediated by apical DMT1¹⁴ (see Section 6.2.1), Barbier et al.²⁹⁰ performed ¹⁰⁹Cd²⁺ tracer microinjections into the late PCT and early DCT of rat kidney and ¹⁰⁹Cd²⁺ reabsorption in the LOH was obtained by calculating the difference between ¹⁰⁹Cd²⁺ recovery after early DCT and late PCT. The authors obtained 46.8% unidirectional ¹⁰⁹Cd²⁺ fluxes that were reduced to 25.4% in the presence of 100 μM Fe²⁺, suggesting that DMT1 in the LOH is involved in ¹⁰⁹Cd²⁺ uptake (and competes with Fe²⁺ for reabsorption).²⁹⁰ This report is unique for its exhaustive characterization of Cd²⁺ transport by the nephron, but unfortunately no additional studies have been published to confirm its conclusions. Despite variable and partly questionable results of FPN1 expression and localization in the rodent LOH^123,140,160 (see Section 6.2.2), even if FPN1 were expressed in the LOH Cd²⁺ would remain trapped within the cells of the LOH because it is not transported by FPN1⁴¹ (see Section 6.1.5).

Given the large number of Ca²⁺ channels expressed throughout the body, the importance of Ca²⁺ signaling, and the large number of ions a channel can transport (∼10⁵ ions per s), even slight permeability of a Ca²⁺ channel to Cd²⁺ might lead to significant Cd²⁺ entry. Indeed, several Ca²⁺ channels that are expressed in the apical membrane of DCT^164,175 are known to transport Cd²⁺. T-type Ca²⁺ channels are blocked by Cd²⁺,^291,292 but their role in Cd²⁺ transport had not been investigated until recently. Ca_v3.1, also known as α_1G, is a T type Ca²⁺ channel and is expressed in the DT (see Section 6.3.2). Ca_v3.1 channels may be suitable for Cd²⁺ transport, because they have a well-defined and substantial window current at negative membrane potentials at which the driving force for divalent cation entry is high²⁹³ and they are ∼2-fold less selective for Ca²⁺ than are L-type Ca²⁺ channels,¹⁷⁰ which suggests that Cd²⁺ may have an increased chance of permeating the channel in the presence of competing Ca²⁺. Furthermore, development of resistance to Cd²⁺ in cell culture has been linked to down-regulation of Ca_v3.1, which suggested the involvement of this channel in Cd²⁺ toxicity.²⁹⁴ Consequently, Lopin et al.²⁹⁵ examined the effects of extracellular Cd²⁺ on permeation and gating of Ca_v3.1 channels stably transfected in HEK293 cells, by using whole-cell recording. In the absence of other permeant ions (Ca²⁺ and Na⁺ were replaced by N-methyl-D-glucamine), Cd²⁺ carried sizable inward currents through Cav3.1 channels (210 ± 20 pA at −60 mV with 2 mM Cd²⁺). Incubation with radiolabeled ¹⁰⁹Cd²⁺ confirmed uptake of Cd²⁺ into cells with Ca_v3.1 channels. With a two-site/three-barrier Eyring model for permeation of Ca_v3.1 channels,¹⁸¹ a transport rate for Cd²⁺ of ∼1 ion per s was estimated for each open channel at −60 mV, with 3–10 nM extracellular Cd²⁺ and in the presence of 2 mM extracellular Ca²⁺. On the basis of the Goldman–Hodgkin–Katz theory,²⁹⁶ a Cd²⁺/Ca²⁺ permeability ratio of 0.66 was calculated, with Cd²⁺ being only slightly less permeable than Ca²⁺.²⁹⁵ Blood Cd²⁺ concentrations range between 0.3 and 5 nM (reviewed in ref. 228). Following glomerular filtration, the concentration of the “free” ionic form of Cd²⁺ in the primary urine of the nephron may increase up to 15-fold in the lumen of the DT (see below). In addition, luminal ionic Cd²⁺ may be further increased by its release from small peptides that are degraded by brush-border enzymes (such as γ-glutamyl transpeptidase that degrades GSH). Hence, in view of the significant “window current” at negative voltages and the high permeability of Ca_v3.1 channels for Cd²⁺ at low nanomolar concentrations (in the presence of physiological Ca²⁺ concentrations), these channels are a likely candidate pathway for Cd²⁺ entry into cells expressing Ca_v3.1 channels, including the kidney DT. Thus, Ca_v3.1 channels could significantly contribute to the in vivo renal toxicity of Cd²⁺ (see Table 2). In another study, the human TRPV5 (ECaC1) of the vanilloid family of the transient receptor channel (TRP) superfamily was transiently expressed in the plasma membrane of human embryonic kidney (HEK293) cells²⁹⁷ (see also Section 6.3.1). Cd²⁺ (and less well Zn²⁺) permeated hTRPV5 in ion imaging experiments using Fura-2 or Newport Green DCF (dichlorofluorescein) with an EC₅₀ of ∼10 respective ∼100 μM Cd²⁺ depending on the absence or presence of 1 mM Ca²⁺ in the extracellular medium. The results were further confirmed using whole-cell patch clamp technique. Transient overexpression of hTRPV5 sensitized cells to Cd²⁺ toxicity. Hence, although micromolar concentrations of Cd²⁺ appeared to be required for permeation the results suggest that TRPV5 may also play a role in Cd²⁺ uptake by the DCT, especially under low Ca²⁺ dietary conditions, when these channels are maximally upregulated. Functional studies in vivo support these cell culture studies. Using both, ¹⁰⁹Cd²⁺ and ⁴⁵Ca²⁺ tracer microinjections into the early and late DCT of rat kidney and recovery in the urine, Barbier et al. showed about 20–25% unidirectional reabsorption of either of the tracers in the DCT that were almost completely abolished in the presence of 100 μM Fe²⁺ or 20 μM Cd²⁺, respectively (which, of course, seems too high from a viewpoint of physiological relevance).²⁹⁰ This suggests that Cd²⁺ is taken up by Fe²⁺ and/or Ca²⁺ transporters in the DCT, possibly DMT1 that is expressed in the luminal membrane of this nephron segment^14,15 (see Section 6.3.3), but also TRPV5/Ca_v3.1 Ca²⁺ channels (see above). However, the experimental design of this tracer microinjection study could not exclude that DMT1 expressed in the CD may also mediate ¹⁰⁹Cd²⁺ reabsorption²⁹⁰ (see below). Finally, we have previously shown apical expression of the Lip2-R in rodent kidney DT (see Sections 6.3.4 and 6.4.2) and cultured mDCT209 cells expressing Lip2-R at their surface internalized submicromolar concentrations of fluorescence-labelled MT that was blocked by 500 pM of the endogenous ligand Lip2.¹⁸² And Cd²⁺–MT caused cell death in mDCT209 cells that could be rescued by 500 pM Lip2.¹⁸² Hence, it is possible that Lip2-R contributes to receptor-mediated endocytosis of toxic Cd²⁺–MT and other Cd²⁺–protein complexes in the DT.

Cd²⁺ reabsorption by terminal nephron segments, i.e. CD, has been investigated by Barbier et al.²⁹⁰ using ¹⁰⁹Cd²⁺ and ⁴⁵Ca²⁺ tracer microinjections. Unidirectional ⁴⁵Ca²⁺ fluxes in the terminal segments of the nephron were not affected by 20 μM Cd²⁺, which suggests that Cd²⁺ permeating Ca²⁺ channels are less likely expressed in the CD. In contrast, they showed that 50–100 μM Fe²⁺, Co²⁺ and Zn²⁺ increased ¹⁰⁹Cd recovery in the urine after microinjection in the early DCT. This suggests involvement of DMT1 in nephron segments downstream of the early DCT, i.e. late DCT and CD (see Section 6.4.1). Mouse IMCD3 cells are sensitive to CdCl₂ (LC₅₀ ∼ 40 μM), indicating uptake of Cd²⁺ by these CD cells²⁹⁸ although the toxic concentrations of Cd²⁺ in these cells were much higher than the K_m of DMT1 for Cd²⁺ transport of ∼1 μM.^11,132 The likely Fe transport pathways of the distal nephron segments (LOH, DT, CD) that compete with Cd²⁺ for uptake are summarized in Table 2. Although Lip2-R is expressed in CD (see Section 6.4.2) and mediates uptake and toxicity of Cd²⁺–MT in various cultured cells,^182,299 its role in Cd²⁺–MT transport and cell damage in the CD has not been investigated so far.

A difficulty in the attempt to estimate the role of distal nephron segments (LOH, DT, CD) in Cd²⁺ uptake – and given the binding affinity of putative transport pathways for Cd²⁺ – is the inability to determine accurately the actual concentrations of ionic and complexed forms of Cd²⁺ in nephron segments downstream of the PT. Nevertheless, an approximation can be obtained from the ratio of inulin concentration in the tubule fluid over plasma (TF/P). Inulin is an indicator of the GFR, i.e. a molecule that is only filtered by the glomerulus and neither reabsorbed nor secreted by the nephron. The TF/P ratio of inulin therefore reflects fluid reabsorption by the nephron.³⁰⁰ The TF/P ratio of inulin increases from 1 to 3 at about 2/3 of the PT length. It reaches a value of 7 at the beginning of the DT and increases up to ∼15 towards its end to reach a final value of 10–200 in the final urine depending on the diuresis condition (water- and anti-diuresis, respectively). In other words, the concentration of non-reabsorbed solutes increases by a factor of 3 along the PT, varies between 7 and 15 along the DT and can increase up to 200-fold in the CD. Consequently, the concentrations of Cd²⁺ and MT/Cd²⁺–MT may increase up to 10–15-fold in the DT and up to 200-fold in the CD, suggesting that these nephron segments may be more relevant segments of the kidney cortex for Cd²⁺ and MT/Cd²⁺–MT uptake and accumulation under conditions of chronic low Cd²⁺ exposure than previously thought. DMT1 and the Lip2-R expressed in DT (cortex) and CD segments (cortex and medulla) are more likely to efficiently reabsorb Cd²⁺ and Cd²⁺–MT, respectively because of their high affinity to these Cd²⁺ compounds (K_m of DMT1 for Cd²⁺ ∼ 1 μM;^11,132K_D of Lip2-R for MT ∼ 120 nM¹⁸²). The renal medulla also accumulates significant amounts of both Cd²⁺ and (Cd²⁺–)MT in humans and concentrations of both Cd²⁺ compounds can reach ∼50% of the levels found in the cortex.^301,302 Indeed, MT has been detected by immunohistochemistry in distal segments of the nephron using cortical and medullary sections of rodent and human kidney^303–305 (although no co-localization with nephron segment-specific markers was performed) and its expression was increased by exposure to Cd²⁺.^304,305 But why then is Cd²⁺ nephrotoxicity less apparent in the DT and kidney medulla? The relative resistance of the DT and kidney medulla to Cd²⁺ toxicity may result from their lower sensitivity to oxidative stress,^306,307 their increased potential for adaptive responses, and stress-induced factors (e.g. hypoxia-inducible factor-1α (HIF-1α), hepcidin, neutrophil gelatinase-associated lipocalin (NGAL) to name a few),^21,308,309 and their metabolic profile (largely anaerobic glycolysis due to a low partial pressure for O₂ in the medullary segments).³¹⁰ All these issues are known to account for the resistance of the DT and kidney medulla to acute tubular damage (e.g. involving mitochondria)³¹¹ and necrosis (outer medulla (straight segment of PT, medullary thick ascending limb of LOH) > cortex (PT, DT) ≫ inner medulla) elicited by various inducers of AKI (reviewed in ref. 312).

8. Determinants of the fate of the kidney exposed to iron or cadmium

Irrespective of the nature of the Cd²⁺ compound that is taken up by tubule cells, its impact on cell viability differs from the effect of Fe. Both metal ions appear to be partly taken up as protein–metal complexes via RME. Moreover, both metal ions may accumulate intracellularly and be detoxified by binding to high-affinity chaperone proteins: for instance, the intracellular Fe storage protein ferritin is induced by overload of the PT with Fe in vivo.^{54,123,313,314} Similarly, following Cd²⁺ exposure in vivo PT cells upregulate the scavenger protein MT for Cd²⁺ storage.^234,235 Both proteins even share some degree of overlapping specificity for Cd²⁺ and Fe²⁺: apart from binding Fe with high affinity⁹¹ ferritin also binds Cd²⁺.^253,254,315 Conversely, although MT binds Cd²⁺ with very high affinity³¹⁶ it has the ability to form complexes with Fe²⁺ as well.³¹⁷ Yet, it is surprising that acute or chronic Fe overload generally does not cause manifest renal damage^213,314 whereas nephrotoxicity is not an unusual sequel of acute or chronic Cd²⁺ exposure.^245,246,250 Transport (TfR1, megalin:cubilin:amnionless, Lip2-R, DMT1, Ca_v3.1, etc.) (compare Sections 6 and 7 and Table 2) and detoxification/storage mechanisms (MT, ferritin) are shared by Fe²⁺ and Cd²⁺. Hence, the differential toxicity of Fe²⁺ and Cd²⁺ in PT cells could be attributed to disparate expression and/or upregulation of the intracellular metal scavenger proteins ferritin and MT,^318–320 which are both regulated by antioxidant response elements in their gene promoter region (reviewed in ref. 91 and 235). In addition, the differences in binding characteristics of these chaperone proteins for Fe and Cd²⁺ could be accountable: both, the binding capacity (maximally ∼4500 for ferritin⁹¹versus 7 metal ion binding sites for MT³²¹) and/or in the binding affinities of ferritin or MT to Fe and Cd^2+ 317,322 are at variance. However, currently there is no stringent evidence for the relative contribution of both chaperone proteins in determining the extent of Fe and Cd²⁺ toxicity in PT cells, therefore further work is needed to clarify these issues.

In contrast, obvious differences concern the cellular utilization or non-utilization (i.e. toxicity) of both metal ions, and the efflux pathway for Fe²⁺ and Cd²⁺. Fe²⁺ enters mitochondria possibly via DMT1 in the OMM⁸⁵ and mitoferrins in the IMM⁸⁷ for synthesis of heme and Fe–sulfur clusters.^79,80 In contrast, after crossing the OMM (possibly via DMT1 in the OMM⁸⁵) and entering the mitochondrial matrix through the mitochondrial Ca²⁺ uniporter in the IMM³²³ Cd²⁺ disrupts mitochondrial function,^324,325 which leads to increased formation of ROS and death through apoptosis and/or necrosis (reviewed in ref. 219 and 326). Another principle difference is the inability of FPN1 at the basolateral side of PT cells to transport Cd²⁺ into the circulation and thereby to clear it from the cell, which is in contrast to FPN1 handling of Fe²⁺.⁴¹ Hence, these differences may underlie – or at least contribute to – the mode of damage likely developing after acute exposure to high concentrations of Cd²⁺ (necrosis) or to nephrotoxicity induced by chronic accumulation of low concentrations of Cd²⁺ (apoptosis, cancer development).

Considering the competition between Fe²⁺ and Cd²⁺ for transport at renal entry pathways (“ionic and molecular mimicry”),^4,221 it should also be deduced that Fe deficiency may not only augment body and kidney Cd²⁺ burden⁵ by increased gastrointestinal absorption of Cd^2+ 327 but also facilitate renal Cd²⁺ reabsorption and thereby elicit a higher likelihood of renal tubule damage.

9. Outlook

Despite a wealth of novel data suggesting a contribution of the kidney to systemic Fe homeostasis and good evidence for uptake and reabsorption of Fe by specific transporters in various nephron segments as well as their involvement in Cd²⁺ uptake and nephrotoxicity, there is still not enough in vivo data available. Presently, a detailed characterization of Fe transporters has been performed in cell lines and heterologous expression systems and these studies have unambiguously demonstrated which transporters are Fe²⁺ and Cd²⁺ selective and which are not (see Table 2). In contrast, only one study has described in vivo transport of Fe by the nephron and this report is already 15 years old.¹³ Similarly, only one study has investigated the role of different nephron segments in uptake of Cd²⁺ and other divalent metal ions in vivo.²⁹⁰ These studies – as exhaustive and thorough as they are – would need to be confirmed and extended, in particular by using nephron specific transporter knockout models. Indeed, this approach has been successfully used to clarify the role of megalin:cubilin:amnionless in reabsorption of proteins by the PT,^115,278,328 including reabsorption of the Fe-binding protein Tf.¹⁶ Unfortunately, no study has attempted to investigate systemic Fe homeostasis in nephron specific megalin or megalin:cubilin deficient animals or whether the PT of these animals is protected against Cd²⁺ nephrotoxicity. Studies are underway that aim to investigate the role of renal Lip2-R in the uptake and toxicity of metalloproteins, including Cd²⁺–MT and Tf, in nephron specific lip2-R knockout mice, (F. Thévenod & S. de Seigneux; in preparation).

Another weakness of this area of research is the inconsistent characterization of the renal localization of renal Fe transporters that has resulted in contradictory results (see Table 1). These conflicting data have even inspired some authors to indiscriminately adopt models of renal Fe transport that are not compatible with renal epithelial physiology and membrane transport. A critical analysis of the relevant experimental reports identifies four principle methodological problems that may also be linked: the tissue sections used for immunostaining were often of poor quality (e.g. tubule lumina were collapsed); authors failed to properly identify nephron segments, e.g. by co-localization studies with nephron specific markers; the images showed poor resolution; and last not least, antibodies used were often of doubtful origin or their specificity had not been proven (see Table 1). This lack of methodological rigor (obviously immunostaining data were not important enough and therefore did not seem to concern the authors) has, at least in part, weakened this field of research. Once more, the use of renal Fe transporter knockout models will hopefully shed light on these confusing data.

Nevertheless, this review has clearly demonstrated that the kidney plays a previously unsuspected role in systemic iron balance and that renal Fe transporters are crucial for the accumulation of Cd²⁺ in the kidney and the development of nephrotoxicity. Future studies, as suggested above, should be able to verify the significance of Fe transporters described in this review and possibly identify additional relevant uptake pathways for renal Fe transport. Last not least, we think that the contribution of circulating Cd²⁺–MT (originating from the liver or not) to chronic Cd²⁺ accumulation in the PT and its toxicity may not be as important as previously suggested and that other hypotheses should be envisaged and experimentally tested.

Abbreviations

2,5-DHBA	2,5-Dihydroxybenzoic acid
AKI	Acute kidney injury
AQP2	Aquaporin 2
ATP	Adenosine triphosphate
BBM	Brush-border membrane
BOCT	Brain organic cation transporter (Lip2-R/Lipocalin-2 receptor)
calcein-AM	Calcein acetoxymethyl (AM) ester
CaV3.1	Calcium channel, voltage-dependent (T-type, α1G subunit)
CCD	Cortical collecting duct
CD	Collecting duct
CHO	Chinese hamster ovary (cell line)
COXII	Cytochrome C oxidase subunit II
CTD	Carboxy-terminal domain
DCF	Dichlorofluorescein
DCT	Distal convoluted tubule
DMT1/Nramp2/DCT1/SLC11A2	Proton-coupled divalent metal transporter 1
DT	Distal tubule
EC50	Half maximal effective concentration
ECaC	Epithelial calcium channel
ENaC	Epithelial sodium channel
Fc	Fragment crystallizable (region of an antibody)
FLVCR1	Feline leukemia virus, subgroup C, receptor (heme exporter)
FPN1/IREG1/MTP1/SLC40A1	Ferroportin
GFR	Glomerular filtration rate
GI	Gastrointestinal
Grx3_4	Glutaredoxin 3_4
GSC	Glomerular sieving coefficient
GSH	Glutathione
H-_L-ferritin	Heavy-_light ferritin subunit
HEK	Human embryonic kidney (cell line)
HIF-1α	Hypoxia-inducible factor-1α
HMWP	High-molecular weight protein
IMM	Inner mitochondrial membrane
IRE	Iron response element
K ^M0.5/Km	Substrate/metal concentration at which velocity is half-maximal
K D	Equilibrium dissociation constant
LC50	Half maximal lethal concentration
Lip2	Lipocalin-2 (NGAL/24p3)
Lip2-R	Lipocalin-2 receptor
LMWP	Low-molecular weight proteins
LOH	Loop of Henle
MDCK	Madin-Darby canine kidney (cell line)
MFRN1/SLC25A37	Mitoferrin-1
MFRN2/SLC25A28	Mitoferrin-2
MM	Molecular mass
MT	Metallothionein
NCCT	NaCl cotransporter
NGAL/24p3	Neutrophil gelatinase-associated lipocalin
NTBI	Non-transferrin-bound iron
NTD	Amino-terminal domain
OCT2	Organic cation transporter 2
OMCD	Outer medullary collecting duct
OMM	Outer mitochondrial membrane
pA	Picoampere
PCBP1	Poly(rC)-binding protein 1
PCT	Proximal convoluted tubule
pS	Picosiemens
PT	Proximal tubule
RME	Receptor-mediated endocytosis
ROS	Reactive oxygen species
RT-PCR	Reverse transcription polymerase chain reaction
S1_2_3	Segment 1_2_3 (PT)
SLC	Solute carrier
Steap	Sixtransmembrane epithelial antigen of the prostate (oxidoreductase)
T_L-type	Transient opening_long lasting-type (calcium channel)
TAL	Thick ascending limb
TBI	Transferrin-bound iron
Tf	Transferrin
TF/P	Tubule fluid over plasma
TfR1_2	Transferrin receptor 1_2
TOM6	Translocase of outer membrane 6
TRPML1/ML1/MLN1/MCLN1	Transient receptor potential mucolipin 1
TRPV	Vanilloid (V) family of the transient receptor potential channel (TRP) superfamily
VDAC	Voltage-dependent anion channel/porin
ZIP8_14/SLC39A8_14	Zrt, Irt-related proteins 8 (ZIP8/SLC39A8) and 14

Acknowledgements

Financial support was obtained by the Deutsche Forschungsgemeinschaft TH345 and the Center for Biomedical Training and Research ZBAF from the University of Witten/Herdecke. We thank our long-time collaborators Drs Craig P. Smith (University of Manchester, U.K.), Robert A. Fenton (Aarhus University, Denmark) and Michael D. Garrick (SUNY, Buffalo, U.S.A.) for valuable discussions and support.

References

1. T. Ganz, Physiol. Rev., 2013, 93, 1721–1741.
2. G. F. Nordberg, Toxicol. Appl. Pharmacol., 2009, 238, 192–200.
3. I. Sovago and K. Varnagy, Met. Ions Life Sci., 2013, 11, 275–302.
4. T. W. Clarkson, Annu. Rev. Pharmacol. Toxicol., 1993, 33, 545–571.
5. L. Jarup, M. Berglund, C. G. Elinder, G. Nordberg and M. Vahter, Scand. J. Work, Environ. Health, 1998, 24(Suppl 1), 1–51.
6. M. Berglund, A. Akesson, B. Nermell and M. Vahter, Environ. Health Perspect., 1994, 102, 1058–1066.
7. L. Jarup and A. Akesson, Toxicol. Appl. Pharmacol., 2009, 238, 201–208.
8. G. L. Diamond, in Molecular biology and toxicology of metals, ed. R. K. Zalups and J. Koropatnick, Taylor & Francis, London and New York, 2000, ch. 11, pp. 300–345.
9. R. K. Zalups and S. Ahmad, Toxicol. Appl. Pharmacol., 2003, 186, 163–188.
10. H. Gunshin, B. Mackenzie, U. V. Berger, Y. Gunshin, M. F. Romero, W. F. Boron, S. Nussberger, J. L. Gollan and M. A. Hediger, Nature, 1997, 388, 482–488.
11. A. C. Illing, A. Shawki, C. L. Cunningham and B. Mackenzie, J. Biol. Chem., 2012, 287, 30485–30496.
12. C. P. Smith and F. Thévenod, Biochim. Biophys. Acta, 2009, 1790, 724–730.
13. M. Wareing, C. J. Ferguson, R. Green, D. Riccardi and C. P. Smith, J. Physiol., 2000, 524(pt 2), 581–586.
14. C. J. Ferguson, M. Wareing, D. T. Ward, R. Green, C. P. Smith and D. Riccardi, Am. J. Physiol. Renal Physiol., 2001, 280, F803–F814.
15. M. Wareing, C. J. Ferguson, M. Delannoy, A. G. Cox, R. F. McMahon, R. Green, D. Riccardi and C. P. Smith, Am. J. Physiol. Renal Physiol., 2003, 285, F1050–F1059.
16. R. Kozyraki, J. Fyfe, P. J. Verroust, C. Jacobsen, A. Dautry-Varsat, J. Gburek, T. E. Willnow, E. I. Christensen and S. K. Moestrup, Proc. Natl. Acad. Sci. U. S. A., 2001, 98, 12491–12496.
17. A. G. Norden, M. Lapsley, P. J. Lee, C. D. Pusey, S. J. Scheinman, F. W. Tam, R. V. Thakker, R. J. Unwin and O. Wrong, Kidney Int., 2001, 60, 1885–1892.
18. F. Thévenod, BioMetals, 2010, 23, 857–875.
19. J. Wang and K. Pantopoulos, Biochem. J., 2011, 434, 365–381.
20. K. Gkouvatsos, G. Papanikolaou and K. Pantopoulos, Biochim. Biophys. Acta, 2012, 1820, 188–202.
21. A. M. Martines, R. Masereeuw, H. Tjalsma, J. G. Hoenderop, J. F. Wetzels and D. W. Swinkels, Nat. Rev. Nephrol., 2013, 9, 385–398.
22. I. De Domenico, D. McVey Ward and J. Kaplan, Nat. Rev. Mol. Cell Biol., 2008, 9, 72–81.
23. K. Pantopoulos, S. K. Porwal, A. Tartakoff and L. Devireddy, Biochemistry, 2012, 51, 5705–5724.
24. Iron in biochemistry and medicine, II., ed. A. Jacobs and M. Worwood, Academic Press, London, 1980, pp. 1–706.
25. J. Prousek, Pure Appl. Chem., 2007, 79, 2325–2338.
26. R. Green, R. Charlton, H. Seftel, T. Bothwell, F. Mayet, B. Adams, C. Finch and M. Layrisse, Am. J. Med., 1968, 45, 336–353.
27. M. Hynes, J. Clin. Pathol., 1948, 1, 57–67.
28. M. C. Linder, Nutrients, 2013, 5, 4022–4050.
29. C. Finch, Blood, 1994, 84, 1697–1702.
30. M. Shayeghi, G. O. Latunde-Dada, J. S. Oakhill, A. H. Laftah, K. Takeuchi, N. Halliday, Y. Khan, A. Warley, F. E. McCann, R. C. Hider, D. M. Frazer, G. J. Anderson, C. D. Vulpe, R. J. Simpson and A. T. McKie, Cell, 2005, 122, 789–801.
31. A. Qiu, M. Jansen, A. Sakaris, S. H. Min, S. Chattopadhyay, E. Tsai, C. Sandoval, R. Zhao, M. H. Akabas and I. D. Goldman, Cell, 2006, 127, 917–928.
32. A. Rajagopal, A. U. Rao, J. Amigo, M. Tian, S. K. Upadhyay, C. Hall, S. Uhm, M. K. Mathew, M. D. Fleming, B. H. Paw, M. Krause and I. Hamza, Nature, 2008, 453, 1127–1131.
33. A. Shawki, P. B. Knight, B. D. Maliken, E. J. Niespodzany and B. Mackenzie, Curr. Top. Membr., 2012, 70, 169–214.
34. D. J. Lane, D. H. Bae, A. M. Merlot, S. Sahni and D. R. Richardson, Nutrients, 2015, 7, 2274–2296.
35. B. Mackenzie, A. Shawki, R. Kim, S. R. Anthony, P. B. Knight, E. M. Bradford and G. E. Shull, FASEB J., 2011, 25, 238.231 (abstract).
36. L. Vanoaica, D. Darshan, L. Richman, K. Schumann and L. C. Kuhn, Cell Metab., 2010, 12, 273–282.
37. H. Shi, K. Z. Bencze, T. L. Stemmler and C. C. Philpott, Science, 2008, 320, 1207–1210.
38. S. Abboud and D. J. Haile, J. Biol. Chem., 2000, 275, 19906–19912.
39. A. Donovan, A. Brownlie, Y. Zhou, J. Shepard, S. J. Pratt, J. Moynihan, B. H. Paw, A. Drejer, B. Barut, A. Zapata, T. C. Law, C. Brugnara, S. E. Lux, G. S. Pinkus, J. L. Pinkus, P. D. Kingsley, J. Palis, M. D. Fleming, N. C. Andrews and L. I. Zon, Nature, 2000, 403, 776–781.
40. A. T. McKie, P. Marciani, A. Rolfs, K. Brennan, K. Wehr, D. Barrow, S. Miret, A. Bomford, T. J. Peters, F. Farzaneh, M. A. Hediger, M. W. Hentze and R. J. Simpson, Mol. Cell, 2000, 5, 299–309.
41. C. J. Mitchell, A. Shawki, T. Ganz, E. Nemeth and B. Mackenzie, Am. J. Physiol.: Cell Physiol., 2014, 306, C450–C459.
42. Z. L. Harris, A. P. Durley, T. K. Man and J. D. Gitlin, Proc. Natl. Acad. Sci. U. S. A., 1999, 96, 10812–10817.
43. C. D. Vulpe, Y. M. Kuo, T. L. Murphy, L. Cowley, C. Askwith, N. Libina, J. Gitschier and G. J. Anderson, Nat. Genet., 1999, 21, 195–199.
44. S. Cherukuri, R. Potla, J. Sarkar, S. Nurko, Z. L. Harris and P. L. Fox, Cell Metab., 2005, 2, 309–319.
45. N. Montalbetti, A. Simonin, G. Kovacs and M. A. Hediger, Mol. Aspects Med., 2013, 34, 270–287.
46. E. Ramos, L. Kautz, R. Rodriguez, M. Hansen, V. Gabayan, Y. Ginzburg, M. P. Roth, E. Nemeth and T. Ganz, Hepatology, 2011, 53, 1333–1341.
47. T. Ganz and E. Nemeth, Biochim. Biophys. Acta, 2012, 1823, 1434–1443.
48. E. Nemeth, M. S. Tuttle, J. Powelson, M. B. Vaughn, A. Donovan, D. M. Ward, T. Ganz and J. Kaplan, Science, 2004, 306, 2090–2093.
49. P. Brissot, M. Ropert, C. Le Lan and O. Loreal, Biochim. Biophys. Acta, 2012, 1820, 403–410.
50. B. J. Scott and A. R. Bradwell, Clin. Chem., 1983, 29, 629–633.
51. C. Hershko and T. E. Peto, Br. J. Haematol., 1987, 66, 149–151.
52. R. C. Hider, Eur. J. Clin. Invest., 2002, 32(suppl 1), 50–54.
53. C. A. Finch, V. Bellotti, S. Stray, D. A. Lipschitz, J. D. Cook, M. J. Pippard and H. A. Huebers, West. J. Med., 1986, 145, 657–663.
54. L. A. Cohen, L. Gutierrez, A. Weiss, Y. Leichtmann-Bardoogo, D. L. Zhang, D. R. Crooks, R. Sougrat, A. Morgenstern, B. Galy, M. W. Hentze, F. J. Lazaro, T. A. Rouault and E. G. Meyron-Holtz, Blood, 2010, 116, 1574–1584.
55. B. S. Skikne, Am. J. Hematol., 2008, 83, 872–875.
56. D. M. Frazer and G. J. Anderson, BioFactors, 2014, 40, 206–214.
57. A. N. Luck and A. B. Mason, Curr. Top. Membr., 2012, 69, 3–35.
58. P. Aisen, Int. J. Biochem. Cell Biol., 2004, 36, 2137–2143.
59. J. E. Levy, O. Jin, Y. Fujiwara, F. Kuo and N. C. Andrews, Nat. Genet., 1999, 21, 396–399.
60. H. Kawabata, R. Yang, T. Hirama, P. T. Vuong, S. Kawano, A. F. Gombart and H. P. Koeffler, J. Biol. Chem., 1999, 274, 20826–20832.
61. J. Chen and C. A. Enns, Biochim. Biophys. Acta, 2012, 1820, 256–263.
62. B. D. Grant and J. G. Donaldson, Nat. Rev. Mol. Cell Biol., 2009, 10, 597–608.
63. P. Ponka, C. Beaumont and D. R. Richardson, Semin. Hematol., 1998, 35, 35–54.
64. R. S. Ohgami, D. R. Campagna, E. L. Greer, B. Antiochos, A. McDonald, J. Chen, J. J. Sharp, Y. Fujiwara, J. E. Barker and M. D. Fleming, Nat. Genet., 2005, 37, 1264–1269.
65. R. S. Ohgami, D. R. Campagna, A. McDonald and M. D. Fleming, Blood, 2006, 108, 1388–1394.
66. S. Dhungana, C. H. Taboy, O. Zak, M. Larvie, A. L. Crumbliss and P. Aisen, Biochemistry, 2004, 43, 205–209.
67. S. Gruenheid, F. Canonne-Hergaux, S. Gauthier, D. J. Hackam, S. Grinstein and P. Gros, J. Exp. Med., 1999, 189, 831–841.
68. M. D. Fleming, M. A. Romano, M. A. Su, L. M. Garrick, M. D. Garrick and N. C. Andrews, Proc. Natl. Acad. Sci. U. S. A., 1998, 95, 1148–1153.
69. X. P. Dong, X. Cheng, E. Mills, M. Delling, F. Wang, T. Kurz and H. Xu, Nature, 2008, 455, 992–996.
70. W. Breuer, M. Shvartsman and Z. I. Cabantchik, Int. J. Biochem. Cell Biol., 2008, 40, 350–354.
71. F. Petrat, H. de Groot, R. Sustmann and U. Rauen, Biol. Chem., 2002, 383, 489–502.
72. R. C. Hider and X. L. Kong, BioMetals, 2011, 24, 1179–1187.
73. L. R. Devireddy, D. O. Hart, D. H. Goetz and M. R. Green, Cell, 2010, 141, 1006–1017.
74. C. Correnti, V. Richardson, A. K. Sia, A. D. Bandaranayake, M. Ruiz, Y. Suryo Rahmanto, Z. Kovacevic, M. C. Clifton, M. A. Holmes, B. K. Kaiser, J. Barasch, K. N. Raymond, D. R. Richardson and R. K. Strong, PLoS One, 2012, 7, e43696.
75. U. Muhlenhoff, S. Molik, J. R. Godoy, M. A. Uzarska, N. Richter, A. Seubert, Y. Zhang, J. Stubbe, F. Pierrel, E. Herrero, C. H. Lillig and R. Lill, Cell Metab., 2010, 12, 373–385.
76. D. R. Richardson, D. J. Lane, E. M. Becker, M. L. Huang, M. Whitnall, Y. Suryo Rahmanto, A. D. Sheftel and P. Ponka, Proc. Natl. Acad. Sci. U. S. A., 2010, 107, 10775–10782.
77. A. S. Zhang, A. D. Sheftel and P. Ponka, Blood, 2005, 105, 368–375.
78. A. D. Sheftel, A. S. Zhang, C. Brown, O. S. Shirihai and P. Ponka, Blood, 2007, 110, 125–132.
79. R. S. Ajioka, J. D. Phillips and J. P. Kushner, Biochim. Biophys. Acta, 2006, 1763, 723–736.
80. R. Lill, B. Hoffmann, S. Molik, A. J. Pierik, N. Rietzschel, O. Stehling, M. A. Uzarska, H. Webert, C. Wilbrecht and U. Muhlenhoff, Biochim. Biophys. Acta, 2012, 1823, 1491–1508.
81. W. Xu, T. Barrientos and N. C. Andrews, Cell Metab., 2013, 17, 319–328.
82. R. Benz, Biochim. Biophys. Acta, 1994, 1197, 167–196.
83. W. Tan and M. Colombini, Biochim. Biophys. Acta, 2007, 1768, 2510–2515.
84. M. Y. Liu and M. Colombini, J. Bioenerg. Biomembr., 1992, 24, 41–46.
85. N. A. Wolff, A. J. Ghio, L. M. Garrick, M. D. Garrick, L. Zhao, R. A. Fenton and F. Thévenod, FASEB J., 2014, 28, 2134–2145.
86. N. A. Wolff, L. M. Garrick, L. Zhao, M. D. Garrick and F. Thévenod, Channels, 2014, 8, 458–466.
87. G. C. Shaw, J. J. Cope, L. Li, K. Corson, C. Hersey, G. E. Ackermann, B. Gwynn, A. J. Lambert, R. A. Wingert, D. Traver, N. S. Trede, B. A. Barut, Y. Zhou, E. Minet, A. Donovan, A. Brownlie, R. Balzan, M. J. Weiss, L. L. Peters, J. Kaplan, L. I. Zon and B. H. Paw, Nature, 2006, 440, 96–100.
88. P. N. Paradkar, K. B. Zumbrennen, B. H. Paw, D. M. Ward and J. Kaplan, Mol. Cell. Biol., 2009, 29, 1007–1016.
89. J. Sripetchwandee, S. B. KenKnight, J. Sanit, S. Chattipakorn and N. Chattipakorn, Acta Physiol., 2014, 210, 330–341.
90. S. B. Keel, R. T. Doty, Z. Yang, J. G. Quigley, J. Chen, S. Knoblaugh, P. D. Kingsley, I. De Domenico, M. B. Vaughn, J. Kaplan, J. Palis and J. L. Abkowitz, Science, 2008, 319, 825–828.
91. D. Finazzi and P. Arosio, Arch. Toxicol., 2014, 88, 1787–1802.
92. S. Leidgens, K. Z. Bullough, H. Shi, F. Li, M. Shakoury-Elizeh, T. Yabe, P. Subramanian, E. Hsu, N. Natarajan, A. Nandal, T. L. Stemmler and C. C. Philpott, J. Biol. Chem., 2013, 288, 17791–17802.
93. S. Levi, B. Corsi, M. Bosisio, R. Invernizzi, A. Volz, D. Sanford, P. Arosio and J. Drysdale, J. Biol. Chem., 2001, 276, 24437–24440.
94. P. Santambrogio, G. Biasiotto, F. Sanvito, S. Olivieri, P. Arosio and S. Levi, J. Histochem. Cytochem., 2007, 55, 1129–1137.
95. Brenner & Rector's The Kidney, ed. M. W. Taal, G. M. Chertow, P. A. Marsden, K. Skorecki, A. S. L. Yu and B. M. Brenner, Saunders, Philadelphia, 9th edn, 2012, pp. 1–3064.
96. Seldin and Giebisch's The Kidney: Physiology & Pathophysiology, ed. R. J. Alpern, M. J. Caplan and O. W. Moe, Academic Press, London, 5th edn, 2013, pp. 1–3352.
97. Handbook of Physiology, ed. E. E. Windhager, Oxford University Press, Oxford, 1992, pp. 1–2544.
98. W. D. Comper, B. Haraldsson and W. M. Deen, J. Am. Soc. Nephrol., 2008, 19, 427–432.
99. E. I. Christensen and H. Birn, Nat. Rev. Mol. Cell Biol., 2002, 3, 256–266.
100. C. D. Klaassen, J. Liu and B. A. Diwan, Toxicol. Appl. Pharmacol., 2009, 238, 215–220.
101. M. J. Burne, T. M. Osicka and W. D. Comper, Kidney Int., 1999, 55, 261–270.
102. P. R. Cutillas, R. J. Chalkley, K. C. Hansen, R. Cramer, A. G. Norden, M. D. Waterfield, A. L. Burlingame and R. J. Unwin, Am. J. Physiol. Renal Physiol., 2004, 287, F353–F364.
103. D. Zhang, E. Meyron-Holtz and T. A. Rouault, J. Am. Soc. Nephrol., 2007, 18, 401–406.
104. P. Aisen, in Iron metabolism in health and disease, ed. J. H. Brock, J. W. Halliday, M. J. Pippard, L. W. Powell and W. B. Saunders, London, 1994, pp. 1–30.
105. L. Kjeldsen, A. H. Johnsen, H. Sengelov and N. Borregaard, J. Biol. Chem., 1993, 268, 10425–10432.
106. A. Tojo and S. Kinugasa, Internet J. Nephrol., 2012, 2012, 481520.
107. E. D. Klootwijk, M. Reichold, R. J. Unwin, R. Kleta, R. Warth and D. Bockenhauer, Nephrol., Dial., Transplant., 2015, 30, 1456–1460.
108. E. I. Christensen, H. Birn, T. Storm, K. Weyer and R. Nielsen, Physiology, 2012, 27, 223–236.
109. B. Seetharam, E. I. Christensen, S. K. Moestrup, T. G. Hammond and P. J. Verroust, J. Clin. Invest., 1997, 99, 2317–2322.
110. J. C. Fyfe, M. Madsen, P. Hojrup, E. I. Christensen, S. M. Tanner, A. de la Chapelle, Q. He and S. K. Moestrup, Blood, 2004, 103, 1573–1579.
111. S. Amsellem, J. Gburek, G. Hamard, R. Nielsen, T. E. Willnow, O. Devuyst, E. Nexo, P. J. Verroust, E. I. Christensen and R. Kozyraki, J. Am. Soc. Nephrol., 2010, 21, 1859–1867.
112. V. Hvidberg, C. Jacobsen, R. K. Strong, J. B. Cowland, S. K. Moestrup and N. Borregaard, FEBS Lett., 2005, 579, 773–777.
113. S. Cui, P. J. Verroust, S. K. Moestrup and E. I. Christensen, Am. J. Physiol., 1996, 271, F900–F907.
114. H. P. Peters, C. M. Laarakkers, P. Pickkers, R. Masereeuw, O. C. Boerman, A. Eek, E. A. Cornelissen, D. W. Swinkels and J. F. Wetzels, BMC Nephrol., 2013, 14, 70.
115. H. Birn, J. C. Fyfe, C. Jacobsen, F. Mounier, P. J. Verroust, H. Orskov, T. E. Willnow, S. K. Moestrup and E. I. Christensen, J. Clin. Invest., 2000, 105, 1353–1361.
116. N. E. Magnusson, M. Hornum, K. A. Jorgensen, J. M. Hansen, C. Bistrup, B. Feldt-Rasmussen and A. Flyvbjerg, BMC Nephrol., 2012, 13, 8.
117. R. A. Orlando, M. Exner, R. P. Czekay, H. Yamazaki, A. Saito, R. Ullrich, D. Kerjaschki and M. G. Farquhar, Proc. Natl. Acad. Sci. U. S. A., 1997, 94, 2368–2373.
118. R. M. Sandoval, M. C. Wagner, M. Patel, S. B. Campos-Bilderback, G. J. Rhodes, E. Wang, S. E. Wean, S. S. Clendenon and B. A. Molitoris, J. Am. Soc. Nephrol., 2012, 23, 447–457.
119. V. Tenten, S. Menzel, U. Kunter, E. M. Sicking, C. R. van Roeyen, S. K. Sanden, M. Kaldenbach, P. Boor, A. Fuss, S. Uhlig, R. Lanzmich, B. Willemsen, H. Dijkman, M. Grepl, K. Wild, W. Kriz, B. Smeets, J. Floege and M. J. Moeller, J. Am. Soc. Nephrol., 2013, 24, 1966–1980.
120. E. I. Christensen and H. Birn, Nat. Rev. Nephrol., 2013, 9, 700–702.
121. L. E. Dickson, M. C. Wagner, R. M. Sandoval and B. A. Molitoris, J. Am. Soc. Nephrol., 2014, 25, 443–453.
122. S. Cui, S. Nielsen, T. Bjerke and E. I. Christensen, Exp. Nephrol., 1993, 1, 309–318.
123. B. Moulouel, D. Houamel, C. Delaby, D. Tchernitchko, S. Vaulont, P. Letteron, O. Thibaudeau, H. Puy, L. Gouya, C. Beaumont and Z. Karim, Kidney Int., 2013, 84, 756–766.
124. H. Tsunoo and H. H. Sussman, J. Biol. Chem., 1983, 258, 4118–4122.
125. H. Fuchs and R. Gessner, Biochim. Biophys. Acta, 2002, 1570, 19–26.
126. M. Haley, M. Lowe and C. P. Smith, Proc. Physiol. Soc., 2012, 27, 24P (abstract C41).
127. B. Mackenzie, M. L. Ujwal, M. H. Chang, M. F. Romero and M. A. Hediger, Pfluegers Arch., 2006, 451, 544–558.
128. M. A. Su, C. C. Trenor, J. C. Fleming, M. D. Fleming and N. C. Andrews, Blood, 1998, 92, 2157–2163.
129. P. L. Lee, T. Gelbart, C. West, C. Halloran and E. Beutler, Blood Cells, Mol., Dis., 1998, 24, 199–215.
130. N. Hubert and M. W. Hentze, Proc. Natl. Acad. Sci. U. S. A., 2002, 99, 12345–12350.
131. B. Mackenzie, H. Takanaga, N. Hubert, A. Rolfs and M. A. Hediger, Biochem. J., 2007, 403, 59–69.
132. M. Okubo, K. Yamada, M. Hosoyamada, T. Shibasaki and H. Endou, Toxicol. Appl. Pharmacol., 2003, 187, 162–167.
133. M. Arredondo, M. J. Mendiburo, S. Flores, S. T. Singleton and M. D. Garrick, BioMetals, 2014, 27, 115–123.
134. N. Jabado, F. Canonne-Hergaux, S. Gruenheid, V. Picard and P. Gros, Blood, 2002, 100, 2617–2622.
135. M. Tabuchi, T. Yoshimori, K. Yamaguchi, T. Yoshida and F. Kishi, J. Biol. Chem., 2000, 275, 22220–22228.
136. M. Tabuchi, N. Tanaka, J. Nishida-Kitayama, H. Ohno and F. Kishi, Mol. Biol. Cell, 2002, 13, 4371–4387.
137. M. Abouhamed, J. Gburek, W. Liu, B. Torchalski, A. Wilhelm, N. A. Wolff, E. I. Christensen, F. Thévenod and C. P. Smith, Am. J. Physiol. Renal Physiol., 2006, 290, F1525–F1533.
138. F. Canonne-Hergaux and P. Gros, Kidney Int., 2002, 62, 147–156.
139. S. Lam-Yuk-Tseung and P. Gros, Biochemistry, 2006, 45, 2294–2301.
140. T. Veuthey, M. C. D'Anna and M. E. Roque, Am. J. Physiol. Renal Physiol., 2008, 295, F1213–F1221.
141. P. Aisen, A. Leibman and J. Zweier, J. Biol. Chem., 1978, 253, 1930–1937.
142. S. Haldar, A. Tripathi, J. Qian, A. Beserra, S. Suda, M. McElwee, J. Turner, U. Hopfer and N. Singh, J. Biol. Chem., 2015, 290, 5512–5522.
143. N. A. Wolff, M. D. Garrick, L. M. Garrick and F. Thévenod, Acta Physiol., 2015, 213(suppl 699), 83(P039).
144. A. C. Chua and E. H. Morgan, J. Comp. Physiol., B, 1997, 167, 361–369.
145. C. J. Ferguson, M. Wareing, M. Delannoy, R. Fenton, S. J. McLarnon, N. Ashton, A. G. Cox, R. F. McMahon, L. M. Garrick, R. Green, C. P. Smith and D. Riccardi, Kidney Int., 2003, 64, 1755–1764.
146. T. Veuthey, D. Hoffmann, V. S. Vaidya and M. Wessling-Resnick, Am. J. Physiol. Renal Physiol., 2014, 306, F333–F343.
147. S. Jenkitkasemwong, C. Y. Wang, B. Mackenzie and M. D. Knutson, BioMetals, 2012, 25, 643–655.
148. L. He, K. Girijashanker, T. P. Dalton, J. Reed, H. Li, M. Soleimani and D. W. Nebert, Mol. Pharmacol., 2006, 70, 171–180.
149. K. Girijashanker, L. He, M. Soleimani, J. M. Reed, H. Li, Z. Liu, B. Wang, T. P. Dalton and D. W. Nebert, Mol. Pharmacol., 2008, 73, 1413–1423.
150. J. P. Liuzzi, F. Aydemir, H. Nam, M. D. Knutson and R. J. Cousins, Proc. Natl. Acad. Sci. U. S. A., 2006, 103, 13612–13617.
151. J. J. Pinilla-Tenas, B. K. Sparkman, A. Shawki, A. C. Illing, C. J. Mitchell, N. Zhao, J. P. Liuzzi, R. J. Cousins, M. D. Knutson and B. Mackenzie, Am. J. Physiol.: Cell Physiol., 2011, 301, C862–C871.
152. C. Y. Wang, S. Jenkitkasemwong, S. Duarte, B. K. Sparkman, A. Shawki, B. Mackenzie and M. D. Knutson, J. Biol. Chem., 2012, 287, 34032–34043.
153. B. Wang, S. N. Schneider, N. Dragin, K. Girijashanker, T. P. Dalton, L. He, M. L. Miller, K. F. Stringer, M. Soleimani, D. D. Richardson and D. W. Nebert, Am. J. Physiol.: Cell Physiol., 2007, 292, C1523–C1535.
154. A. Lymboussaki, E. Pignatti, G. Montosi, C. Garuti, D. J. Haile and A. Pietrangelo, J. Hepatol., 2003, 39, 710–715.
155. N. Wilkinson and K. Pantopoulos, Front. Pharmacol., 2014, 5, 176.
156. D. L. Zhang, R. M. Hughes, H. Ollivierre-Wilson, M. C. Ghosh and T. A. Rouault, Cell Metab., 2009, 9, 461–473.
157. B. K. Fuqua, Y. Lu, D. Darshan, D. M. Frazer, S. J. Wilkins, N. Wolkow, A. G. Bell, J. Hsu, C. C. Yu, H. Chen, J. L. Dunaief, G. J. Anderson and C. D. Vulpe, PLoS One, 2014, 9, e98792.
158. I. De Domenico, D. M. Ward, M. C. di Patti, S. Y. Jeong, S. David, G. Musci and J. Kaplan, EMBO J., 2007, 26, 2823–2831.
159. D. M. Ward and J. Kaplan, Biochim. Biophys. Acta, 2012, 1823, 1426–1433.
160. N. A. Wolff, W. Liu, R. A. Fenton, W. K. Lee, F. Thévenod and C. P. Smith, J. Cell. Mol. Med., 2011, 15, 209–219.
161. R. R. Starzynski, F. Canonne-Hergaux, M. Lenartowicz, W. Krzeptowski, A. Willemetz, A. Stys, J. Bierla, P. Pietrzak, T. Dziaman and P. Lipinski, Biochem. J., 2013, 449, 69–78.
162. A. Zarjou, S. Bolisetty, R. Joseph, A. Traylor, E. O. Apostolov, P. Arosio, J. Balla, J. Verlander, D. Darshan, L. C. Kuhn and A. Agarwal, J. Clin. Invest., 2013, 123, 4423–4434.
163. Y. Scindia, P. Dey, A. Thirunagari, H. Liping, D. L. Rosin, M. Floris, M. D. Okusa and S. Swaminathan, J. Am. Soc. Nephrol., 2015 DOI:10.1681/ASN.2014101037.
164. J. G. Hoenderop, D. Muller, A. W. Van Der Kemp, A. Hartog, M. Suzuki, K. Ishibashi, M. Imai, F. Sweep, P. H. Willems, C. H. Van Os and R. J. Bindels, J. Am. Soc. Nephrol., 2001, 12, 1342–1349.
165. R. Vennekens, J. G. Hoenderop, J. Prenen, M. Stuiver, P. H. Willems, G. Droogmans, B. Nilius and R. J. Bindels, J. Biol. Chem., 2000, 275, 3963–3969.
166. B. Nilius, R. Vennekens, J. Prenen, J. G. Hoenderop, R. J. Bindels and G. Droogmans, J. Physiol., 2000, 527(Pt 2), 239–248.
167. M. van Abel, J. G. Hoenderop, A. W. van der Kemp, M. M. Friedlaender, J. P. van Leeuwen and R. J. Bindels, Kidney Int., 2005, 68, 1708–1721.
168. T. de Groot, R. J. Bindels and J. G. Hoenderop, Kidney Int., 2008, 74, 1241–1246.
169. A. Qiu and C. Hogstrand, Gene, 2004, 342, 113–123.
170. E. Perez-Reyes, Physiol. Rev., 2003, 83, 117–161.
171. W. A. Catterall, Cold Spring Harbor Perspect. Biol., 2011, 3, a003947.
172. D. M. Rodman, K. Reese, J. Harral, B. Fouty, S. Wu, J. West, M. Hoedt-Miller, Y. Tada, K. X. Li, C. Cool, K. Fagan and L. Cribbs, Circ. Res., 2005, 96, 864–872.
173. A. M. Yunker and M. W. McEnery, J. Bioenerg. Biomembr., 2003, 35, 533–575.
174. P. B. Hansen, Am. J. Physiol.: Regul., Integr. Comp. Physiol., 2015, 308, R227–R237.
175. D. Andreasen, B. L. Jensen, P. B. Hansen, T. H. Kwon, S. Nielsen and O. Skott, Am. J. Physiol. Renal Physiol., 2000, 279, F997–F1005.
176. M. Leclerc, M. G. Brunette and D. Couchourel, Kidney Int., 2004, 66, 242–250.
177. M. G. Brunette, M. Leclerc, D. Couchourel, J. Mailloux and Y. Bourgeois, Can. J. Physiol. Pharmacol., 2004, 82, 30–37.
178. D. Couchourel, M. Leclerc, J. Filep and M. G. Brunette, Mol. Cell. Endocrinol., 2004, 222, 71–81.
179. S. Kumfu, S. Chattipakorn, S. Srichairatanakool, J. Settakorn, S. Fucharoen and N. Chattipakorn, Eur. J. Haematol., 2011, 86, 156–166.
180. K. V. Lopin, I. P. Gray, C. A. Obejero-Paz, F. Thévenod and S. W. Jones, Mol. Pharmacol., 2012, 82, 1194–1204.
181. K. V. Lopin, C. A. Obejero-Paz and S. W. Jones, J. Membr. Biol., 2010, 235, 131–143.
182. C. Langelueddecke, E. Roussa, R. A. Fenton, N. A. Wolff, W. K. Lee and F. Thévenod, J. Biol. Chem., 2012, 287, 159–169.
183. N. Paragas, R. Kulkarni, M. Werth, K. M. Schmidt-Ott, C. Forster, R. Deng, Q. Zhang, E. Singer, A. D. Klose, T. H. Shen, K. P. Francis, S. Ray, S. Vijayakumar, S. Seward, M. E. Bovino, K. Xu, Y. Takabe, F. E. Amaral, S. Mohan, R. Wax, K. Corbin, S. Sanna-Cherchi, K. Mori, L. Johnson, T. Nickolas, V. D'Agati, C. S. Lin, A. Qiu, Q. Al-Awqati, A. J. Ratner and J. Barasch, J. Clin. Invest., 2014, 124, 2963–2976.
184. B. S. Nielsen, N. Borregaard, J. R. Bundgaard, S. Timshel, M. Sehested and L. Kjeldsen, Gut, 1996, 38, 414–420.
185. D. H. Goetz, M. A. Holmes, N. Borregaard, M. E. Bluhm, K. N. Raymond and R. K. Strong, Mol. Cell, 2002, 10, 1033–1043.
186. G. Bao, M. Clifton, T. M. Hoette, K. Mori, S. X. Deng, A. Qiu, M. Viltard, D. Williams, N. Paragas, T. Leete, R. Kulkarni, X. Li, B. Lee, A. Kalandadze, A. J. Ratner, J. C. Pizarro, K. M. Schmidt-Ott, D. W. Landry, K. N. Raymond, R. K. Strong and J. Barasch, Nat. Chem. Biol., 2010, 6, 602–609.
187. T. H. Flo, K. D. Smith, S. Sato, D. J. Rodriguez, M. A. Holmes, R. K. Strong, S. Akira and A. Aderem, Nature, 2004, 432, 917–921.
188. T. Berger, A. Togawa, G. S. Duncan, A. J. Elia, A. You-Ten, A. Wakeham, H. E. Fong, C. C. Cheung and T. W. Mak, Proc. Natl. Acad. Sci. U. S. A., 2006, 103, 1834–1839.
189. R. J. Abergel, M. C. Clifton, J. C. Pizarro, J. A. Warner, D. K. Shuh, R. K. Strong and K. N. Raymond, J. Am. Chem. Soc., 2008, 130, 11524–11534.
190. J. Yang, K. Mori, J. Y. Li and J. Barasch, Am. J. Physiol. Renal Physiol., 2003, 285, F9–F18.
191. K. M. Schmidt-Ott, K. Mori, J. Y. Li, A. Kalandadze, D. J. Cohen, P. Devarajan and J. Barasch, J. Am. Soc. Nephrol., 2007, 18, 407–413.
192. A. Haase-Fielitz, M. Haase and P. Devarajan, Ann. Clin. Biochem., 2014, 51, 335–351.
193. W. F. t. Peacock, A. Maisel, J. Kim and C. Ronco, Postgrad. Med., 2013, 125, 82–93.
194. N. Paragas, A. Qiu, Q. Zhang, B. Samstein, S. X. Deng, K. M. Schmidt-Ott, M. Viltard, W. Yu, C. S. Forster, G. Gong, Y. Liu, R. Kulkarni, K. Mori, A. Kalandadze, A. J. Ratner, P. Devarajan, D. W. Landry, V. D'Agati, C. S. Lin and J. Barasch, Nat. Med., 2011, 17, 216–222.
195. J. Mishra, Q. Ma, A. Prada, M. Mitsnefes, K. Zahedi, J. Yang, J. Barasch and P. Devarajan, J. Am. Soc. Nephrol., 2003, 14, 2534–2543.
196. L. R. Devireddy, C. Gazin, X. Zhu and M. R. Green, Cell, 2005, 123, 1293–1305.
197. L. R. Devireddy, J. G. Teodoro, F. A. Richard and M. R. Green, Science, 2001, 293, 829–834.
198. H. Koepsell and H. Endou, Pfluegers Arch., 2004, 447, 666–676.
199. K. M. Bennett, J. Liu, C. Hoelting and J. Stoll, Mol. Cell. Biochem., 2011, 352, 143–154.
200. W. K. Fang, L. Y. Xu, X. F. Lu, L. D. Liao, W. J. Cai, Z. Y. Shen and E. M. Li, Biochem. J., 2007, 403, 297–303.
201. C. J. Wienken, P. Baaske, U. Rothbauer, D. Braun and S. Duhr, Nat. Commun., 2010, 1, 100.
202. A. Tojo and H. Endou, Am. J. Physiol., 1992, 263, F601–F606.
203. P. D. Leber and D. J. Marsh, Am. J. Physiol., 1970, 219, 358–363.
204. R. G. Galaske, J. B. Van Liew and L. G. Feld, Kidney Int., 1979, 16, 394–403.
205. K. M. Madsen, R. H. Harris and C. C. Tisher, Kidney Int., 1982, 21, 354–361.
206. E. I. Christensen, P. J. Verroust and R. Nielsen, Pfluegers Arch., 2009, 458, 1039–1048.
207. R. B. Klassen, K. Crenshaw, R. Kozyraki, P. J. Verroust, L. Tio, S. Atrian, P. L. Allen and T. G. Hammond, Am. J. Physiol. Renal Physiol., 2004, 287, F393–F403.
208. D. F. Akintola, B. Sampson and A. Fleck, J. Lab. Clin. Med., 1995, 126, 119–127.
209. H. Milnerowicz and A. Bizon, Acta Biochim. Pol., 2010, 57, 99–104.
210. R. Nielsen and E. I. Christensen, Pediatr. Nephrol., 2010, 25, 813–822.
211. C. Kastner, M. Pohl, M. Sendeski, G. Stange, C. A. Wagner, B. Jensen, A. Patzak, S. Bachmann and F. Theilig, Am. J. Physiol. Renal Physiol., 2009, 296, F902–F911.
212. A. C. Alfrey and W. S. Hammond, Kidney Int., 1990, 37, 1409–1413.
213. N. Ishizaka, K. Saito, E. Noiri, M. Sata, I. Mori, M. Ohno and R. Nagai, Nephron Physiol., 2004, 98, p107–p113.
214. E. Dizin, U. Hasler, S. Nlandu-Khodo, M. Fila, I. Roth, T. Ernandez, A. Doucet, P. Y. Martin, E. Feraille and S. de Seigneux, Am. J. Physiol. Renal Physiol., 2013, 305, F1053–F1063.
215. F. Thévenod and W. K. Lee, Met. Ions Life Sci., 2013, 11, 415–490.
216. J. M. Moulis, BioMetals, 2010, 23, 877–896.
217. T. S. Nawrot, J. A. Staessen, H. A. Roels, E. Munters, A. Cuypers, T. Richart, A. Ruttens, K. Smeets, H. Clijsters and J. Vangronsveld, BioMetals, 2010, 23, 769–782.
218. A. Cuypers, M. Plusquin, T. Remans, M. Jozefczak, E. Keunen, H. Gielen, K. Opdenakker, A. R. Nair, E. Munters, T. J. Artois, T. Nawrot, J. Vangronsveld and K. Smeets, BioMetals, 2010, 23, 927–940.
219. F. Thévenod and W. K. Lee, Arch. Toxicol., 2013, 87, 1743–1786.
220. W. Maret and J. M. Moulis, Met. Ions Life Sci., 2013, 11, 1–29.
221. C. C. Bridges and R. K. Zalups, Toxicol. Appl. Pharmacol., 2005, 204, 274–308.
222. R. Cornelis, B. Heinzow, R. F. Herber, J. M. Christensen, O. M. Poulsen, E. Sabbioni, D. M. Templeton, Y. Thomassen, M. Vahter and O. Vesterberg, J. Trace Elem. Med. Biol., 1996, 10, 103–127.
223. M. Nordberg and G. F. Nordberg, Environ. Health Perspect., 1975, 12, 103–108.
224. R. E. Dudley, L. M. Gammal and C. D. Klaassen, Toxicol. Appl. Pharmacol., 1985, 77, 414–426.
225. K. Nomiyama, H. Nomiyama and N. Kameda, Toxicology, 1998, 129, 157–168.
226. S. Thijssen, J. Maringwa, C. Faes, I. Lambrichts and E. Van Kerkhove, Toxicology, 2007, 229, 145–156.
227. L. Jarup, A. Rogenfelt, C. G. Elinder, K. Nogawa and T. Kjellstrom, Scand. J. Work, Environ. Health, 1983, 9, 327–331.
228. K. Klotz, W. Weistenhofer and H. Drexler, Met. Ions Life Sci., 2013, 11, 85–98.
229. E. Freisinger and M. Vasak, Met. Ions Life Sci., 2013, 11, 339–371.
230. H. M. Chan, Y. Tamura, M. G. Cherian and R. A. Goyer, Proc. Soc. Exp. Biol. Med., 1993, 202, 420–427.
231. K. Tanaka, Dev. Toxicol. Environ. Sci., 1982, 9, 237–249.
232. S. A. Gunn and T. C. Gould, Proc. Soc. Exp. Biol. Med., 1957, 96, 820–823.
233. G. F. Nordberg and K. Nishiyama, Arch. Environ. Health, 1972, 24, 209–214.
234. H. M. Chan, L. F. Zhu, R. Zhong, D. Grant, R. A. Goyer and M. G. Cherian, Toxicol. Appl. Pharmacol., 1993, 123, 89–96.
235. I. Sabolic, D. Breljak, M. Skarica and C. M. Herak-Kramberger, BioMetals, 2010, 23, 897–926.
236. W. Maret, J. Biol. Inorg. Chem., 2011, 16, 1079–1086.
237. T. Kjellstrom, Environ. Health Perspect., 1979, 28, 169–197.
238. H. Roels, R. Lauwerys, J. P. Buchet, A. Bernard, J. S. Garvey and H. J. Linton, Int. Arch. Occup. Environ. Health, 1983, 52, 159–166.
239. C. Tohyama, Z. A. Shaikh, K. Nogawa, E. Kobayashi and R. Honda, Toxicology, 1981, 20, 289–297.
240. W. C. Prozialeck and J. R. Edwards, BioMetals, 2010, 23, 793–809.
241. L. Jarup, Nephrol., Dial., Transplant., 2002, 17(suppl 2), 35–39.
242. H. Roels, A. Bernard, J. P. Buchet, A. Goret, R. Lauwerys, D. R. Chettle, T. C. Harvey and I. A. Haddad, Lancet, 1979, 1, 221.
243. WHO, Cadmium, World Health Organization, 1992.
244. Y. Suzuki, J. Toxicol. Environ. Health, 1980, 6, 469–482.
245. P. M. Ferraro, S. Costanzi, A. Naticchia, A. Sturniolo and G. Gambaro, BMC Public Health, 2010, 10, 304.
246. A. Navas-Acien, M. Tellez-Plaza, E. Guallar, P. Muntner, E. Silbergeld, B. Jaar and V. Weaver, Am. J. Epidemiol., 2009, 170, 1156–1164.
247. A. Hartwig, Met. Ions Life Sci., 2013, 11, 491–507.
248. A. Bernard, J. P. Buchet, H. Roels, P. Masson and R. Lauwerys, Eur. J. Clin. Invest., 1979, 9, 11–22.
249. K. Nogawa, A. Ishizaki, M. Fukushima, I. Shibata and N. Hagino, Environ. Res., 1975, 10, 280–307.
250. N. Johri, G. Jacquillet and R. Unwin, BioMetals, 2010, 23, 783–792.
251. W. Goumakos, J. P. Laussac and B. Sarkar, Biochem. Cell Biol., 1991, 69, 809–820.
252. W. R. Harris and L. J. Madsen, Biochemistry, 1988, 27, 284–288.
253. D. J. Price and J. G. Joshi, J. Biol. Chem., 1983, 258, 10873–10880.
254. T. Granier, G. Comberton, B. Gallois, B. L. d'Estaintot, A. Dautant, R. R. Crichton and G. Precigoux, Proteins, 1998, 31, 477–485.
255. W. C. Prozialeck and P. C. Lamar, Arch. Toxicol., 1993, 67, 113–119.
256. R. K. Zalups, Toxicol. Appl. Pharmacol., 2000, 164, 15–23.
257. S. Soodvilai, J. Nantavishit, C. Muanprasat and V. Chatsudthipong, Toxicol. Lett., 2011, 204, 38–42.
258. F. Thévenod, G. Ciarimboli, M. Leistner, N. A. Wolff, W. K. Lee, I. Schatz, T. Keller, R. Al-Monajjed, V. Gorboulev and H. Koepsell, Mol. Pharmacol., 2013, 10, 3045–3056.
259. C. D. Klaassen, J. Liu and S. Choudhuri, Annu. Rev. Pharmacol. Toxicol., 1999, 39, 267–294.
260. N. A. Wolff, M. Abouhamed, P. J. Verroust and F. Thévenod, J. Pharmacol. Exp. Ther., 2006, 318, 782–791.
261. N. A. Wolff, W. K. Lee, M. Abouhamed and F. Thévenod, Toxicol. Appl. Pharmacol., 2008, 230, 78–85.
262. C. Erfurt, E. Roussa and F. Thévenod, Am. J. Physiol.: Cell Physiol., 2003, 285, C1367–C1376.
263. N. A. Wolff, W. K. Lee and F. Thévenod, Toxicol. Lett., 2011, 203, 210–218.
264. M. Abouhamed, N. A. Wolff, W. K. Lee, C. P. Smith and F. Thévenod, Am. J. Physiol. Renal Physiol., 2007, 293, F705–F712.
265. R. D. Palmiter, Proc. Natl. Acad. Sci. U. S. A., 1994, 91, 1219–1223.
266. Y. Liu, J. Liu, S. M. Habeebu, M. P. Waalkes and C. D. Klaassen, Toxicol. Sci., 2000, 57, 167–176.
267. C. Dorian, V. H. Gattone II and C. D. Klaassen, Toxicol. Appl. Pharmacol., 1992, 114, 173–181.
268. C. Dorian, V. H. Gattone II and C. D. Klaassen, Toxicol. Appl. Pharmacol., 1992, 117, 242–248.
269. W. Tang and Z. A. Shaikh, J. Toxicol. Environ. Health, Part A, 2001, 63, 221–235.
270. M. Nordberg, Environ. Health Perspect., 1984, 54, 13–20.
271. J. Sudo, T. Hayashi, J. Terui, M. Soyama, M. Fukata and K. Kakuno, Eur. J. Pharmacol., 1994, 270, 229–235.
272. A. Onodera, M. Tani, T. Michigami, M. Yamagata, K. S. Min, K. Tanaka, T. Nakanishi, T. Kimura and N. Itoh, Toxicol. Lett., 2012, 212, 91–96.
273. E. Felley-Bosco and J. Diezi, Toxicol. Appl. Pharmacol., 1987, 91, 204–211.
274. F. Thévenod, Nephron Physiol., 2003, 93, 87–93.
275. A. Chaumont, M. Nickmilder, X. Dumont, T. Lundh, S. Skerfving and A. Bernard, Toxicol. Lett., 2012, 210, 345–352.
276. P. E. Evrin and L. Wibell, Scand. J. Clin. Lab. Invest., 1972, 29, 69–74.
277. K. Takagi, K. Kin, Y. Itoh, H. Enomoto and T. Kawai, J. Clin. Pathol., 1980, 33, 786–791.
278. J. R. Leheste, B. Rolinski, H. Vorum, J. Hilpert, A. Nykjaer, C. Jacobsen, P. Aucouturier, J. O. Moskaug, A. Otto, E. I. Christensen and T. E. Willnow, Am. J. Pathol., 1999, 155, 1361–1370.
279. C. M. Eakin, J. D. Knight, C. J. Morgan, M. A. Gelfand and A. D. Miranker, Biochemistry, 2002, 41, 10646–10656.
280. A. G. Norden, P. Sharratt, P. R. Cutillas, R. Cramer, S. C. Gardner and R. J. Unwin, Kidney Int., 2004, 66, 1994–2003.
281. H. Birn and E. I. Christensen, Kidney Int., 2006, 69, 440–449.
282. J. Liu, R. A. Goyer and M. P. Waalkes, in Casarett and Doull's Toxicology, ed. C. D. Klaassen, McGraw-Hill, New York, NY, 5th edn, 2008, pp. 931–979.
283. L. Jarup, B. Persson and C. G. Elinder, Occup. Environ. Med., 1995, 52, 818–822.
284. A. Akesson, T. Lundh, M. Vahter, P. Bjellerup, J. Lidfeldt, C. Nerbrand, G. Samsioe, U. Stromberg and S. Skerfving, Environ. Health Perspect., 2005, 113, 1627–1631.
285. W. Xiao, Y. Liu and D. M. Templeton, Toxicol. Appl. Pharmacol., 2009, 238, 315–326.
286. P. V. Rao, S. A. Jordan and M. K. Bhatnagar, J. Environ. Pathol., Toxicol. Oncol., 1989, 9, 19–44.
287. J. P. Girolami, J. L. Bascands, C. Pecher, G. Cabos, J. P. Moatti, J. F. Mercier, J. M. Haguenoer and Y. Manuel, Toxicology, 1989, 55, 117–129.
288. G. Oner, U. K. Senturk and N. Izgut-Uysal, Biol. Trace Elem. Res., 1995, 48, 111–117.
289. R. Lauwerys and A. Bernard, Toxicol. Lett., 1989, 46, 13–29.
290. O. Barbier, G. Jacquillet, M. Tauc, P. Poujeol and M. Cougnon, Am. J. Physiol. Renal Physiol., 2004, 287, F1067–F1075.
291. L. Lacinova, N. Klugbauer and F. Hofmann, Neuropharmacology, 2000, 39, 1254–1266.
292. D. Diaz, R. Bartolo, D. M. Delgadillo, F. Higueldo and J. C. Gomora, J. Membr. Biol., 2005, 207, 91–105.
293. J. R. Serrano, E. Perez-Reyes and S. W. Jones, J. Gen. Physiol., 1999, 114, 185–201.
294. E. M. Leslie, J. Liu, C. D. Klaassen and M. P. Waalkes, Mol. Pharmacol., 2006, 69, 629–639.
295. K. V. Lopin, F. Thévenod, J. C. Page and S. W. Jones, Mol. Pharmacol., 2012, 82, 1183–1193.
296. C. J. Frazier, E. G. George and S. W. Jones, Biophys. J., 2000, 78, 1872–1880.
297. G. Kovacs, N. Montalbetti, M. C. Franz, S. Graeter, A. Simonin and M. A. Hediger, Cell Calcium, 2013, 54, 276–286.
298. E. K. Park, S. K. Mak and B. D. Hammock, Interdiscip. Toxicol., 2013, 6, 157–158.
299. C. Langelueddecke, W. K. Lee and F. Thévenod, Toxicol. Lett., 2014, 226, 228–235.
300. K. J. Ullrich, in Kurzgefaßtes Lehrbuch der Physiologie, ed. W. D. Keidel, Georg Thieme Verlag, Stuttgart, 1975, ch. 10.
301. M. Torra, J. To-Figueras, M. Brunet, M. Rodamilans and J. Corbella, Bull. Environ. Contam. Toxicol., 1994, 53, 509–515.
302. M. Yoshida, H. Ohta, Y. Yamauchi, Y. Seki, M. Sagi, K. Yamazaki and Y. Sumi, Biol. Trace Elem. Res., 1998, 63, 167–175.
303. S. H. Garrett, M. A. Sens, J. H. Todd, S. Somji and D. A. Sens, Toxicol. Lett., 1999, 105, 207–214.
304. T. Nagamine, K. Nakazato, K. Suzuki, T. Kusakabe, T. Sakai, M. Oikawa, T. Satoh, T. Kamiya and K. Arakawa, Biol. Trace Elem. Res., 2007, 117, 115–126.
305. L. Wang, D. Chen, H. Wang and Z. Liu, Biol. Trace Elem. Res., 2009, 129, 190–199.
306. R. Campos, F. Maureira, A. Garrido and A. Valenzuela, Comp. Biochem. Physiol., Part B: Biochem. Mol. Biol., 1993, 105, 157–163.
307. B. Gonzalez-Flecha, P. Evelson, N. Sterin-Speziale and A. Boveris, Biochim. Biophys. Acta, 1993, 1157, 155–161.
308. A. P. Zou and A. W. Cowley, Jr., Acta Physiol. Scand., 2003, 179, 233–241.
309. R. A. Zager, A. C. Johnson and K. B. Frostad, Am. J. Physiol. Renal Physiol., 2014, 307, F856–F868.
310. A. A. McDonough and S. C. Thompson, in Brenner & Rector's The Kidney, ed. M. W. Taal, G. M. Chertow, P. A. Marsden, K. Skorecki, A. S. L. Yu and B. M. Brenner, Saunders, Philadelphia, 9th edn, 2012, ch. 4, vol. 1, pp. 138–157.
311. A. M. Hall, G. J. Rhodes, R. M. Sandoval, P. R. Corridon and B. A. Molitoris, Kidney Int., 2013, 83, 72–83.
312. A. Sharfuddin, S. D. Weisbord, P. M. Palevsky and B. A. Molitoris, in Brenner & Rector's The Kidney, ed. M. W. Taal, G. M. Chertow, P. A. Marsden, K. Skorecki, A. S. L. Yu and B. M. Brenner, Saunders, Philadelphia, 9th edn, 2012, ch. 30, vol. 1, pp. 1044–1099.
313. G. W. Richter, J. Exp. Med., 1957, 106, 203–218.
314. S. Fagoonee, J. Gburek, E. Hirsch, S. Marro, S. K. Moestrup, J. M. Laurberg, E. I. Christensen, L. Silengo, F. Altruda and E. Tolosano, Am. J. Pathol., 2005, 166, 973–983.
315. J. G. Wardeska, B. Viglione and N. D. Chasteen, J. Biol. Chem., 1986, 261, 6677–6683.
316. J. H. Kagi and B. L. Vallee, J. Biol. Chem., 1961, 236, 2435–2442.
317. X. Ding, E. Bill, M. Good, A. X. Trautwein and M. Vasak, Eur. J. Biochem., 1988, 171, 711–714.
318. J. P. Muller, M. Vedel, M. J. Monnot, N. Touzet and M. Wegnez, DNA Cell Biol., 1991, 10, 571–579.
319. D. M. Durnam and R. D. Palmiter, Mol. Cell. Biol., 1984, 4, 484–491.
320. J. C. Fleet, G. K. Andrews and C. C. McCormick, J. Nutr., 1990, 120, 1214–1222.
321. J. H. Kagi and A. Schaffer, Biochemistry, 1988, 27, 8509–8515.
322. D. Price and J. G. Joshi, Proc. Natl. Acad. Sci. U. S. A., 1982, 79, 3116–3119.
323. W. K. Lee, U. Bork, F. Gholamrezaei and F. Thévenod, Am. J. Physiol. Renal Physiol., 2005, 288, F27–F39.
324. Y. Wang, J. Fang, S. S. Leonard and K. M. Rao, Free Radical Biol. Med., 2004, 36, 1434–1443.
325. W. K. Lee, M. Spielmann, U. Bork and F. Thévenod, Am. J. Physiol.: Cell Physiol., 2005, 289, C656–C664.
326. W. K. Lee and F. Thévenod, Am. J. Physiol.: Cell Physiol., 2006, 291, C195–C202.
327. M. Kippler, W. Goessler, B. Nermell, E. C. Ekstrom, B. Lonnerdal, S. El Arifeen and M. Vahter, Environ. Res., 2009, 109, 914–921.
328. K. Weyer, T. Storm, J. Shan, S. Vainio, R. Kozyraki, P. J. Verroust, E. I. Christensen and R. Nielsen, Nephrol., Dial., Transplant., 2011, 26, 3446–3451.

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