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

CHAPTER 6. Renal Handling of Organic Solutes

Orson W. Moe   Stephen H. Wright   Manuel Palacín

  

 

Glucose, 214

  

 

Physiology of Renal Glucose Transport, 214

  

 

Molecular Biology of Renal Glucose Transport Proteins, 216

  

 

Renal Glucose Transport in Diseases States, 217

  

 

Organic Cations, 218

  

 

Physiology of Renal Organic Cation Transport, 218

  

 

Molecular Biology of Renal Organic Cation Transport, 221

  

 

Clinical Diseases from Genetic Defects of Organic Cation Transporters, 225

  

 

Organic Anions, 225

  

 

Organic Anion Physiology, 225

  

 

Molecular Biology of Organic Anion Transporters, 227

  

 

Clinical Relevance of Organic Anion Transporters, 229

  

 

Amino Acids, 230

  

 

Physiology of Renal Amino Acid Transport, 230

  

 

Molecular Biology of Amino Acid Transport, 230

  

 

Inherited Aminoacidurias in Humans, 233

Archaic nephrons in lower life forms are largely secretory in nature. Kidneys in higher vertebrates handle solutes by the processes of filtration, reabsorption, and secretion. The handling of organic solutes involves all three of these means. The human kidney produces approximately 150 L to 170 L of low pro-tein cell-free ultrafiltrate per day. The renal tubules processes this large volume of fluid to conserve the essential nutrients (glucose, amino acids, Krebs cycle intermediates, vitamins), to eliminate potentially toxic substances (organic acids and bases), and to reduce the quantity of salt and water excreted in the final urine. The handling of organic solutes spans a wide range from a clearance that exceeds glomerular filtration rate (GFR) in the form of filtration followed by secretion (fractional excretion > 1) to filtration fol-lowed by complete reabsorption (fractional excretion ≈0), and everything in between (Fig. 6-1 ).

The kidney participates in homeostasis by adjusting the body content of specific solutes in the body as well as concentration of specific solutes in certain body fluid compartments; usually in the plasma. To achieve these regulatory functions, there must be sensing mechanisms for both the pool size and the concentration of the solute. Unlike inorganic solutes such as sodium or potassium, with organic solutes, the total pool is difficult to define as these solutes are constantly being synthesized and metabolized. For glucose, the maintenance of a discrete plasma concentration is clearly important. For amino acids and organic cation and anions, it is less clear whether plasma levels are as tightly regulated. The renal regulation of this latter group of organic solutes is more concerned with external balance and adjustment of urinary concentrations.

A filtration-reabsorption design is absolutely critical to maintain a high GFR, which is required for the complex metabolism and homeothermy of terrestrial mammals as tubular reabsorption salvages all the valuable solutes (e.g., sodium, bicarbonate, glucose) that would have otherwise be lost in the urine (see Fig. 6-1 ). In addition to allowance of high GFR, filtration-reabsorption com-mences by disposing everything and then selectively reclaims and retains substances the organism desires to keep in the appropriate amount. All that is not reclaimed is excreted. This mechanism economizes on genes and gene products required to identify and excrete the myriad of undesirable substances. In the filtration-secretion or secretion mode, the burden is on the kidney to recognize the substrates to be secreted. Therefore, in contrast to glucose transport (reabsorption), which is highly specific to certain hexose structures, organic anion and cation transport (secretion) can engage hundreds of structurally distinct substrates.

Unlike the handling of a lot of other solutes described in this textbook, the reabsorption and secretion of organic solutes are primarily performed by the proximal tubule with little or no contribution past the pars recta. This chapter summarizes the physiology, cell, and molecular biology of organic solute transport in the kidney, and highlights certain aspects of clinical relevance. Although only renal mechanisms will be covered in this chapter, it is important to note that homeostasis of organic solutes involves the concerted action of multiple organs.

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FIGURE 6-1  Secretory and reabsorptive modes of the mammalian nephron.

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GLUCOSE

Physiology of Renal Glucose Transport

Overview

Plasma glucose concentration is regulated at about 5 μM with balanced actions of glucose ingestion, glycogenolysis, and gluconeogenesis against glucose utilization and in some circumstances renal glucose excretion. Although transient increments and decrements of plasma glucose is tolerated in post-prandial and fasting states, neither hypoglycemia nor hyperglycemia is desirable for the organism. The robust metabolic rate of mammals mandates a high glomerular filtration rate (GFR) so the loss of glucose through the ultrafiltrate will be colossal if not reclaimed. Therefore the main physiologic task of the kidney is to retrieve as much glucose as possible so the normal urine is glucose-free. This was described by Cushny as early as 1917.[5]

Renal Glucose Handling

Plasma glucose is neither protein-bound nor complexed with macromolecules and is filtered freely at the glomerulus. Glucose reabsorption by the proximal tubule increases as the filtered load increases (Plasma [glucose] × GFR) until it reaches a threshold (TmGlucose) that represents the maximal reabsorptive capacity of the proximal tubule, then glycosuria ensues ( Fig. 6-2 ). This concept was first inducted by the classic studies of Shannon and is still quite valid today.[6] With normal GFR, the value of plasma glucose for glycosuria to occur is about 11 μM or 200 mg/dl. One can predict that glycosuria will occur at lower plasma glucose concentrations in physiologic states of hyperfiltration such as pregnancy or a unilateral kidney (e.g., nephrectomy, transplant allograft, etc.). In these circumstances, glycosuria may not indicate significant hyperglycemia. Conversely, in patients with renal insufficiency, it will take a plasma glucose concentration of more than 11 μM (200 mg/dl) for glycosuria to occur. The reduced filtered load for a given plasma glucose concentration is partially counterbalanced by lower TmGlucose (see Fig. 6-2 ) but glycosuria still occurs at a higher plasma glucose concentration. Some of the whole organism values for renal glucose handling in humans are summarized in Table 6-1 .[7]

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FIGURE 6-2  Urinary glucose excretion and tubular reabsorption as a function of filtered load. Tubular reabsorption increases linearly with filtered load as a part of glomerulotubular balance. When reabsorption reaches the tubular capacity (Tmglucose), glucose starts appearing in the urine. The plasma glucose concentration for the given GFR is the glycosuric threshold.

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TABLE 6-1   -- Renal Glucose Handing in Humans under Physiologic States

Excretion rate

2.7 μmoles/min (3.4 mmoles/day)

Urinary concentration

0.50 μM–0.65 μM

Reabsorptive capacity

1.85–2.17 mmoles/min

TmGlucose

(2664–3125 mmoles/day)

 

 

 

Microperfusion data for rabbit proximal tubules indicate that the maximal rate of glucose transport slows as one progresses from S1 to S3 ( Fig. 6-3 ).[1] However, the affinity for glucose rises, with a Michaelis constant (Km; concentration of substrate where half maximal rates of transport is attained) of approximately 2 μM in S1 to 0.4 μM in S3.[1] The net result of different Na+-glucose carrier kinetics along the length of the proximal tubule is that S1can reabsorb glucose with higher capacity but the S3 can decrease the tubule fluid glucose concentration to a much lower level. Theoretically, a single uniform segment cannot perform both high-capacity and high-gradient glucose absorption. Transport studies with brush border membrane vesicles and molecular cloning methods have now firmly established the existence of two Na+-glucose transport systems with kinetic characteristics consistent with earlier microperfusion findings.

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FIGURE 6-3  Relative magnitude of glucose transport characteristics in different segments of the proximal tubule. Jmax, maximal glucose transport rate; Km, affinity constant for glucose.  (Data from Barfuss DW, Schafer JA: Differences in active and passive glucose transport along the proximal nephron. Am J Physiol 241:F322, 1981.)

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When Na+ and glucose move as a net positive charge into the negatively charged cell interior, it partially depolarizes the cell interior. Consequently, when glucose is removed from the luminal solution, the PD becomes more negative (i.e., it hyperpolarizes). [8] [9] Using microelectrodes from the basolateral membrane to measure cell hyperpolarization, Biagi and coworkers found that elimination of Na+-glucose cotransport results in 14 mV of hyperpolarization in S1 and early S2 and about 4 mV in late S2.[10] Na+-glucose transport accounts for approximately 15% of the apical membrane current and for about half of the luminal negative PD in the early PCT.

Aronson and Sacktor first described Na+-dependent glucose transport in renal brush border vesicles in 1974. [11] [12] The two major Na+-glucose transporters are distinguished by their glucose transport capacity; their affinities for glucose, Na+, and phlorhizin; and their location within the kidney. In the outer cortex, where the S1 and S2 segments of the proximal tubule are located, there is predominantly a high-capacity, low-affinity glucose transport system.[13] [14] [15] The low-affinity system has a Km for glucose of approximately 6 μM. The transporter carries one Na+ per glucose with a Km for Na+ of 228 μM.[14] Phlorhizin binds and inhibits the transporter with a dissociation constant (Kd) of 1 μM to 2 μM.[14]

In the outer medulla, where S3 is located, there is a high-affinity system with a Km for glucose of approximately 0.3 μM, carrying two Na+ per glucose. [13] [14] [16] The coupling of two Na+ to one glucose allows the cotransporter to utilize the square of the electrochemical driving force of Na+ to energize glucose uptake. The S3 transporter has a K0.5 for Na+ of approximately 50 μM. Although the S3 transporter binds phlorhizin with a Kd of 1 μM to 2 μM, it inhibits glucose transport with a Kd of 50 μM. The S3 transporter has an affinity for D-galactose that is more than 10-fold higher than that of the S1 transporter.[13]

Molecular Biology of Renal Glucose Transport Proteins

Cell Model of Proximal Tubule Glucose Transport

Glucose reabsorption in the proximal tubule cell occurs in two steps: (1) carrier-mediated, Na+-glucose cotransport across the apical membrane, followed by (2) facilitated glucose transport and active Na+ extrusion across the basolateral membrane (see Fig. 6-2 ). Electroneutrality is maintained by either paracellular Cl- diffusion or Na+ back-diffusion, depending on the relative permeabilities of the intercellular tight junction to Na+ and Cl- ( Fig. 6-4 ). Two specific Na+-coupled carriers (sodium glucose cotransporter SGLT-1 and SGLT-2) have been identified in the proximal tubule cell apical membrane that bind Na+ and glucose in the tubule fluid. A third gene SGLT-3 has been cloned from a porcine kidney cell line and is transcribed in kidney.[17] SGLT3 has been studied in heterologously expressed system but its localization and functional role in the kidney is undetermined so the current paradigm still contains only SGLT-1 and -2 (see Fig. 6-4 ). The translocation of the Na+ and glucose across the apical cell membrane is driven by the electrochemical driving force for Na+ from tubule fluid to cell and is therefore termed “secondary active transport.” Exit of glucose across the basolateral membrane does not consume energy but is mediated by specific carriers belonging to the GLUT gene family (see Fig. 6-4 ). SGLT-1 and -2 belongs to a broader group of solute carriers called SLC5, which currently encompasses 11 members in the human genome of which 6 are Na+-glucose cotransporters.[18]

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FIGURE 6-4  Model of proximal tubule glucose absorption. The Na+-K+-ATPase lowers cell [Na+] and generates a negative interior voltage. This drives the uphill Na+-coupled glucose entry from the apical membrane via the SGLT transporters 1 and 2. Glucose leaves the basolateral membrane via the facilitative glucose transporters GLUT1 and GLUT down its electrochemical gradient.

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Transporter Proteins

Apical Entry

Molecular studies have confirmed with striking fidelity the physiologic data on glucose transport obtained in perfused tubules and membrane vesicles. It has been known since the 1960s that patients with the rare congenital disorder of glucose-galactose malabsorption have a partial defect in renal absorption of glucose, [19] [20] [21] [22] [23] [24] but patients with renal glycosuria have normal intestinal glucose transport.[20] This finding led to the conjecture that one of the two renal glucose transporters may also be found in the intestine. Hediger and co-workers cloned the intestinal Na+-glucose transporter and found expression in both intestine and kidney. [23] [24] Within the kidney, it was later shown to be expressed almost exclusively in the S3 segment of the proximal tubule.[25] Sequence comparison showed similarity to the proline transporter of Escherichia coli, the Na+-dependent neutral amino acid transporter, and the Na+-dependent myo-inositol transporter. This transporter, termed SGLT-1, has a Km for glucose of 0.4 μM, is inhibited by 5 μM to 10 μM of phlorhizin, and binds two Na+ with a Km for Na+ of 32 μM ( Table 6-2 ).[24] The high affinity allows SGLT-1 to reclaim even low concentrations glucose from the urinary lumen. The 2Na+: 1 glucose stoichiometry squares the electrochemical driving force of the lumen-to-cell Na+ gradient. These properties are virtually identical to those of the S3 glucose transport system determined from earlier microperfusion studies and transport studies in membrane vesicles.


TABLE 6-2   -- Na+-coupled Glucose Transporter Family

 

SGLT1

SGLT2

SGLT3

Gene name

SLC5A1

SLC5A2

SLC5A4

Human chromosome

22p13.1

16p11.2

22p12.1

OMIM

182380

233100

Genetic disease

Intestinal glucose galactose malabsorption

Familial renal glycosuria

Amino acids

664

672

659

Tissue distribution

Kidney, intestine

Kidney

Kidney, intestine, liver spleen

Renal expression

Proximal straight tubule

Proximal convoluted tubule

Unknown

Affinity glucose (K0.5, μM)

0.4

2

6

Hexose selectivity

Gluc = Gal

Gluc >> Gal

Gluc >> Gal

Affinity sodium (K0.5, μM)

32

100

1.5

Substrate stoichiometry

2Na+: glucose

Na+: glucose

2Na+: glucose

 

Gluc, Glucose; Gal, Galactose.

 

 

 

Hediger and colleagues cloned a second glucose transporter termed SGLT-2. [26] [27] This clone exhibits 59% homology to SGLT-1 and is expressed in kidney, but not intestine.[28] SGLT-2 confers phlorhizin-sensitive (1 μM to 5 μM) glucose transport with a Km for glucose of approximately 1.6 μM. One Na+ is bound per glucose with a Km for Na+ of 200 μM to 300 μM (see Table 6-2 ). In situ hybridization localized SGLT-2 to the cortex in S1 proximal tubule segments. SGLT-2 is most likely the previously described “low-affinity” glucose transporter.

SGLT-1 and SGLT-2 are responsible for bringing glucose into the proximal tubule cell via secondary active transport, but clearly a different system is needed to return this glucose from the cell to the blood. The transporter was found to be inhibited by phloretin and cytochalasin B, but not phlorhizin. [29] [30] Although stereospecific for D-glucose, it also transports 2-deoxy-D-glucose and 3-O-methyl-D-glucose, but not α-methyl-D-glucoside.[31] These characteristics are similar to those of proteins found in polarized intestinal and liver cells and to those of the insulin-sensitive D-glucose transporters in red blood cells, muscle cells, and adipocytes.[32] Another cDNA from the SGLT family was cloned from a pig kidney cell line and then subsequently human now termed SGLT-3. [33] [34] [35] SGLT-3 resembles SGLT-2 in terms of its low affinity for glucose and high specificity for glucose over other hexose substrates, but it functions more like SGLT-1 in terms of its tissue distribution and 2:1 Na+: glucose stoichiometry (see Table 6-2 ). [36] [37] [38] The SGLT-3 transcript is present in the kidney but in low levels[39]; nephron segmental distribution is not yet available. At present, SGLT-3 has been characterized in expression systems but its role in the kidney is unclear.

Basolateral Exit

The relationships between glucose transport in the proximal tubule basolateral membrane and glucose transport in other tissues has been clarified with the discovery of a large gene family termed the GLUT genes. There are now 17 known members of the GLUT gene family ( Table 6-3 ).[40] One classification based on sequence dendrograms has been proposed (see Table 6-3 ). A thorough discussion is beyond the scope of this book. Several excellent reviews are available. [40] [41] [42] At present, the two isoforms that are believed to be important for transepithelial glucose transport are GLUT1 and GLUT2 (see Fig. 6-4 ). The first member of the GLUT family to be discovered was GLUT-1, cloned via an antibody to the red blood cell glucose transporter. The carrier has a high affinity for glucose (1 μM to 2 μM) and is found at variable levels in virtually all nephron segments. [43] [44] Its expression may correlate with nutritive requirements of the cell,[45] and it is probably also the mechanism for glucose exit in S3.[46] GLUT-2 is a high-capacity, low-affinity (15 μM to 20 μM) basolateral transporter found in tissues with large glucose fluxes, such as intestine, liver, and pancreas, and the S1 segment of the PCT. [47] [48] GLUT-4 is the insulin-responsive glucose transporter found almost exclusively in fat and muscle. [49] [50] This transporter has also been found in glomeruli and renal microvessels.[51] The regulation of this and other glucose transporters in diabetes is discussed elsewhere. [40] [52] The role of GLUT-2 in renal glucose transport has been demonstrated by the presence of renal glycosuria in mice with GLUT-2 deletion[53] as well as in humans with GLUT-2 mutations who present interestingly with Fanconi syndrome, which is glycosuria with generalized proximal tubule dysfunction. [54] [55] Transcripts of some of the other GLUT transporters have been detected in the kidney but their roles are unclear.


TABLE 6-3   -- Facilitative Sugar Transporters

Protein

Gene

Glut Class

Renal Expression

GLUT1

SLC2A1

I

All nephron segments

Proximal tubule basolateral membrane S2

GLUT2

SLC2A2

I

Proximal tubule basolateral membrane S1

GLUT3

SLC2A3

I

Absent

GLUT4

SLC2A4

I

mRNA in situ in thick ascending limb

GLUT5

SLC2A5

II

mRNA in situ in proximal straight tubule

GLUT6

SLC2A6

III

Absent

GLUT7

SLC2A7

II

Unknown

GLUT8

SLC2A8

III

Absent

GLUT9

SLC2A9

II

mRNA present

GLUT10

SLC2A10

III

mRNA present

GLUT11

SLC2A11

II

Absent

GLUT12

SLC2A12

III

Unknown

HMIT

SLC2A13

III

Unknown

No gene product

SLC2A3P1

 

No gene product

SLC2A3P2

 

No gene product

SLC2A3P3

 

No gene product

SLC2AXP1

 

 

 

 

Renal Glucose Transport in Diseases States

Monogenic Defects of Glucose Transport

SGLT-1

The best characterized monogenic disease in the SGLT family is glucose-galactose malabsorption due to inactivating mutation of SGLT1 gene (OMIM 182380). [56] [57] [58] [59] [60] This rare autosomal recessive disease presents in infancy with an intestinal phenotype. The osmotic diarrhea resolves upon cessation of dietary glucose, galactose, and lactose; substrates of SGLT-1. The diarrhea returns when rechallenged with one of more these substrates. The diagnosis of the disease can be readily confirmed by oral administration of glucose or galactose (2 g/kg) followed by lactic acid determination in breath. Patients with inactivating mutations of SGLT-1 exhibit some degree of renal glycosuria. [19] [22] [61] In general the severity is very mild and reduction of tubular maximal absorptive capacity was not always demonstrable.[62] This is in keeping though with the low capacity late proximal tubule SGLT-1 transport system.

Renal Glycosuria

There is considerable controversy as to the inheritance pattern (autosomal dominant versus recessive), clinical classification of the reabsorptive defect (glucose threshold versus maximal absorptive capacity, versus both), and associated overlapping defects with aminoaciduria in this syndrome. [63] [64] Due to the lack of intestinal defect and the renal-restricted distribution of SGLT-2, the SGLT-2 gene has been repeatedly proposed as the candidate for renal glycosuria. To date, the strongest evidence that SGLT2 is the major transporter involved in the reabsorption of glucose from the glomerular filtrate comes from the analysis of one patient with autosomal recessive renal glycosuria with a homozygous nonsense mutation in exon 11 of SGLT2, and a heterozygous mutation at the same position in both parents and a younger brother.[65] In contrast, the linkage of the autosomal dominant form of renal glycosuria to the HLA complex on chromosome 6 are not supportive of the SGLT transporters being causative.[66] Based on circumstantial evidence, an autoimmune mechanism has been proposed for this disease.[67] It is possible that this entity represents a heterogeneous group of disorders.

Diseases of GLUTs

The first patient with Fanconi-Bickel syndrome[68] had hepatorenal glycogenosis and renal Fanconi syndrome.[69] This child presented at age 6 months with failure to thrive, polydipsia, and constipation followed later in childhood by osteopenia, short stature, hepatomegaly, and a proximal tubulopathy consisting of glycosuria, phosphaturia, aminoaciduria, proteinuria, and hyperuricemia. The liver was infiltrated with glycogen and fat. Disturbance of glucose homeostasis includes fasting hypoglycemia and ketosis and postprandially hyperglycemia. A mutation in the GLUT2 gene was demonstrated by Santer and co-workers.[70] Most patients with the Fanconi-Bickel syndrome are homozygous for the disease-related mutations consistent with an autosomal recessive pattern of inheritance. Some patients have been shown to be compound heterozygotes.[71] The mechanism by which GLUT2 mutation cause the proximal tubulopathy is unclear. It is conceivable that impaired basolateral exit of glucose in the proximal convoluted tubule can lead to glucose accumulation and glycotoxicity. GLUT2 gene deletion in rodents leads to glucose-insensitive islet cells but proximal tubulopathy was not described.[72] GLUT1 mutations presents with primarily a neurologic syndrome with no documented renal involvement. [68] [73] [74]

Pharmacologic Manipulation of SGLT

Antidiabetic therapy traditionally targets several broad levels: gut glucose absorption, insulin release, and insulin sensitivity. One additional strategy is providing a glucose sink to alleviate hyperglycemia and the ravages of glycotoxicity without actual direct manipulation of insulin secretion or sensitivity. If one decreases the capacity of proximal absorption, the same filtered load will lead to higher glycosuria resulting in lower plasma glucose concentration ( Fig. 6-5 ). In addition to providing a glucose sink, the proximal osmotic diuresis can potentially act via tubuloglomerular feedback and reduce GFR, especially in the setting of diabetic hyperfiltration. One advantage of this approach is the self-limiting effect. Increased filtered load from hyperglycemia in the presence of reduced proximal reabsorption increases glycosuria (see Fig. 6-5 ). Once hyperglycemia is corrected and filtered load is reduced, the renal glucose leak ceases even if the drug is still on board (see Fig. 6-5 ). This approach is receiving increasing attention[75] with new technical advances in high through-put screening.[76] A variety compounds with widely divergent structures has been shown to inhibit SGLT function. [77] [78] [79] [80] [81] [82] [83] Glycemic control with these agents has been shown in animal models. [84] [85] [86] The long-term consequence of escalated glycosylation of epithelial proteins exposed to the urinary lumen has not been examined. Because hyperglycemia fluctuates, so does osmotic diuresis. The staccato natriuresis may present a challenge in control of extracellular fluid volume.

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FIGURE 6-5  Effect of SGLT inhibition. Inhibition of proximal absorption leads to increased glucose excretion. Proximal osmotic diuresis activates tubuloglomerular (TG) feedback and reduces hyperfiltration. Right panel shows self-adjusting features of the renal glucose sink. As plasma glucose level falls, so does filtered load and glycosuria ceases even proximal absorption is still inhibited.

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ORGANIC CATIONS

Physiology of Renal Organic Cation Transport

Overview

The kidney is capable of clearing the plasma of a vast array of compounds that share little in common other than possessing a net positive charge at physiological pH. These “organic cations” (OCs) include a structurally diverse array of primary, secondary, tertiary, or quaternary amines, although compounds that have non-nitrogen cationic moieties (e.g., phosphoniums[87]) can also interact effectively with what is frequently referred to as the “classical organic cation secretory pathway”.[88] Studies employing the techniques of stop flow, micropuncture, and microperfusion identified the renal proximal tubule (RPT) as the principal site of renal OC secretion. [89] [90] [91] Although a number of endogenous OCs are actively secreted by the proximal tubule (e.g., N[1]-methylnicotinamide (NMN), choline, epinephrine, and dopamine; see Ref 90), an equally, if not more important function of this process is clearing the body of xenobiotic compounds, [18] [89] including a wide range of alkaloids and other positively charged, heterocyclic dietary constituents; cationic drugs of therapeutic or recreational use; or other cationic toxins of environmental origin (e.g., nicotine). Importantly, the secretory process is also a site of clinically significant interactions between OCs in humans. For example, therapeutic doses of cimetidine retard the renal elimination of procainamide [2] [92] and nicotine.[93]

The Cellular Basis of Renal Organic Cation Secretion

Renal OC secretion involves the concerted activity of a suite of distinct transport processes arranged in series (i.e., in the basolateral [peritubular] and apical [luminal] poles of RPT cells); or in parallel (i.e., within the same pole of RPT cells). In developing a model for the functional basis of this complexity, it is useful to consider the “Type I” and “Type II” classifications for different structural classes of organic cations developed to describe OC secretion in the liver.[94] In general, Type I OCs are comparatively small (generally <400 Da) monovalent compounds, such as tetraethylammonium (TEA), tributylmethylammonium, and procainamide ethobromide. Importantly, the majority of cationic drugs from a wide array of clinical classes, including antihistamines, skeletal muscle relaxants, antiarrhythmics, and β-adrenoceptor blocking agents, are adequately described as being Type I OCs. Type II OCs are usually bulkier (generally >500 Da) and frequently polyvalent, including d-tubocurarine, vecuronium, and hexafluorenium. Although the kidney plays a quantitatively significant role in the secretion of selected Type II OCs, the liver plays the predominant role in excretion (into the bile) of large hydrophobic cations (e.g., see Ref 95). In contrast, renal excretion is a predominant avenue for clearance of Type I OCs. Thus, although the processes associated with the renal handling of Type II OCs will be briefly described as currently understood, the renal transport of Type I OCs of substrates will be the central focus of this discussion. The reader is directed to recent reviews that consider the molecular biology and physiology of MDR1 (P-gp) in more depth. [96] [97] [98]

Basolateral Organic Cation Entry

Figure 6-6 shows a model for transcellular OC transport by RPT that is consistent with studies employing isolated renal plasma membranes and intact proximal tubules [89] [99] and supported by recent molecular data. The first step in transcellular OC secretion involves OC entry into RPT cells from the blood across the peritubular membrane. For Type I OCs this entry step involves either an electrogenic uniport (facilitated diffusion), driven by the inside-negative electrical potential difference (PD),[100] or an electroneutral antiport (exchange) of OCs [100] [101] (it is likely that these two mechanisms represent alternative modes of action of the same transporter(s)[102]). The PD across the basolateral membrane of RPT cells is in the order of 50 mV to 60 mV (inside negative [103] [104]), which is sufficient to account for an accumulation of OCs within proximal cells to levels approximately 10 times that in the blood. A hallmark of peritubular OC uptake is its broad selectivity, frequently termed “multispecificity”.[105] Studies by Ullrich and colleagues on the structural specificity of peritubular OC transport in microperfused rat proximal tubules in vivo indicated a clear correlation between an increase in substrate hydrophobicity and an increase in interaction with basolateral OC transporters, [105] [106] although it is also clear that steric factors influence this interaction. [107] [108] [109]

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FIGURE 6-6  Schematic model of the transport processes associated with the secretion of organic cations (OCs) by renal proximal tubule cells. Circles depict carrier mediated transport processes. Arrows indicate the direction of net substrate transport. Solid lines depict the principal pathways of OC transport; dotted lines indicate pathways that are believed to be of secondary importance; dashed line indicates diffusive movement. Na+-K+-ATPase; maintains the K+ gradient associated with the inside negative membrane potential and the inwardly directed Na+ gradient, both of which represent driving forces associated with active OC secretion. OCT1, OCT2, and OCT3; support electrogenic facilitated diffusion associated with basolateral uptake of Type I OCs (these processes are also believed to support electroneutral OC/OC exchange, as indicated by the outwardly directed arrows). MDR1; supports the ATP dependent, active luminal export of Type II OCs. NHE3; the Na+/H+ exchanger that plays a principal role in sustaining the inwardly directed hydrogen electrochemical gradient that, in turn, supports activity of transport processes mediated by physiologically characterized Type I OC/H+ exchanger, which includes MATE1 (the broadly specific process that accepts TEA as a prototypic substrate) and the narrowly specific process that accepts guanidine as a substrate, and OCTN1; supports electroneutral OC/H+ exchange but with selectivity properties that make it distinct from process 7. OCTN2 supports Na+-carnitine cotransport and the electrogenic flux of TEA and selected Type I substrates, as well as mediated exchange of TEA for Na+-carnitine. Finally, there is a physiologically characterized electrogenic choline reabsorption pathway.

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The molecular identity of the transport processes responsible for basolateral entry of Type I OCs is relatively clear. OCT1 (organic cation transporter 1; SLC22A1), OCT2 (SLC22A2), and OCT3 (SLC22A3) are electrogenic uniporters that are expressed in the basolateral membrane of renal proximal tubule cells (although their individual levels of ex-pression display marked species differences). Significantly, elimination of OCT1 and OCT2 activity (in OCT1/OCT2 null mice) completely eliminates active secretion of TEA.[3] The molecular biology of each of these processes is discussed in more detail in an upcoming section.

The process(es) responsible for basolateral entry of Type II OCs into RPT cells is(are) not clear. The bulky ring structures that characterize Type II OCs generally render them substantially more hydrophobic than Type I OCs and, in the liver, generally makes Type II OCs (e.g., rocuronium) substrates for one or more homologues of the organic anion transporting polypeptide (OATPs) family of transporters.[110] However, (in the rat) renal OATP expression is typically low, compared to the liver (the sole exception being Oatp5, the function and location of which in rat kidney is unknown, and for which no human ortholog has been identified[111]). It is likely that the marked hydrophobicity of most Type II OCs results in a substantial diffusive flux across the peritubular membrane that provides Type II OCs with a passive, electrically conductive avenue for entry into proximal cells.

Apical Organic Cation Exit

Exit of Type I OCs across the luminal membrane involves carrier-mediated antiport of OC for H+ (see Fig. 6-6 ), a process observed in brush border membrane vesicles (BBMV) isolated from human, rabbit, rat, dog, chicken, and snake kidneys.[89] Luminal OC efflux is the rate-limiting step in trans-tubular OC secretion.[112] It is unlikely that net OC secretion requires a transluminal H+-gradient. Indeed, in the early proximal tubule, where the pH of the tubular filtrate is on the order of 7.4 (i.e., the same as plasma), tubular secretion exceeds that of later segments[112] even though it is in these latter regions where an inwardly directed H+ gradient is most likely to develop.[113]Instead, it is the electrically silent nature of the exchanger (which involves the obligatory 1:1 exchange of monovalent cations[114]) that, even in the absence of an inwardly directed H+ gradient, will permit OCs to exit the electrically negative cytoplasm of RPT cells and develop a luminal concentration as large as (or larger than, if there is an inwardly directed H+ gradient) that in the cytoplasm. Net transepithelial secretion, therefore, is a consequence of combining luminal OC/H+ exchange with the electrically driven flux of OCs across the basolateral membrane. From an energetic perspective, OC/H+ antiport is the active step in the earlier outlined scenario because it depends on the displacement of H+ away from electrochemical equilibrium, a state maintained through the activity in the luminal membrane of the Na+/H+ exchanger [115] [116] and, to a lesser extent, a V-type H-ATPase (not shown in figure).[117] The basolateral Na,K-ATPase, ultimately, drives OC secretion by (1) maintaining the inside negative membrane potential that supports concentrative uptake of OCs across the basolateral membrane (the result of the developed K+ gradient); and (2) sustaining the inwardly directed Na gradient that drives the aforementioned luminal Na+/H+ exchange. Evidence on the structural specificity of luminal OC transport indicates that, as with the peritubular transport process, binding of substrate to the OC/H+ exchanger is profoundly influenced by substrate hydrophobicity and, to a lesser extent, the 3D structure of the substrate.[118] At least two distinct OC/H+ exchangers, distinguished by their substrate selectivities, have been described in renal cortical BBMV. One, which is regarded as being the principal avenue for luminal OC secretion, displays a very broad selectivity and accepts TEA as a substrate.[118] The second displays mechanistically similar characteristics to this former process, but displays a narrower selectivity and accepts guanidine as a substrate.[119]

Two members of the SLC22A family are suspected to play a role in mediating apical efflux of (at least) selected OC substrates. OCTN1 (organic cation transporter-novel 1; SLC22A4) supports mediated exchange of TEA and H+and, consequently, it has been suggested to contribute to luminal OC/H+ exchange activity.[120] However, the kinetics, selectivity, and tissue distribution of OCTN1 do not fit the physiological profile of the OC/H+ exchanger of renal BBMV,[18] making it unlikely that OCTN1 is a major contributor to renal secretion of Type I OCs. OCTN2 (SLC22A5) is unique in displaying both Na+ coupled transport of carnitine (and structurally related zwitterions[121]) and the electrogenic uniport of TEA and selected Type I OCs.[122] The potential significance of OCTN2 in mediating apical OC export is evident in the observation that a genetic defect in Octn2 in the jvs mouse is associated with a marked decrease in renal clearance of TEA.[3]

The recent cloning from human and mouse kidney of a member of the MOP (Multidrug/Oligosaccharidyl-lipid/Polysaccharide) superfamily of multidrug/H+ exchangers[123] offers the promise of identifying the principal elements in luminal OC/H+ exchange. The multidrug and toxicant extruder, MATE1, supports TEA/H+ exchange and in the human is expressed in the apical membrane of renal proximal tubules and in the canalicular membrane of hepatic cells[123] (i.e., locations known to contains OC/H+ exchange activity). Moreover, the kinetics and selectivity of the process [123] [124] are consistent with those of OC/H+ exchange characterized in isolated renal BBMV, further supporting the contention that MATE1 may comprise a quantitatively significant element in luminal OC secretion.

The apical export of Type II OCs is likely to involve the multidrug resistance transporter, MDR1 (ABCB1), which is expressed in the apical membrane of RPT cells and has been implicated in the apical efflux of Type II OCs (and other bulky hydrophobic substrates) in in vitro studies. [125] [126] [127] However, whereas the influence of MDR1 in biliary excretion of Type II OCs is evident (e.g., in studies employing Mdr1 knockout mice)[128]; the quantitative influence of MDR1 on renal secretion is less clear. For example, whereas biliary excretion of doxorubicin is markedly decreased in Mdr1 knockout mice, urinary clearance increases.[129] Similarly, elimination of Mdr1 activity in knockout mice is associated with marked changes in the distribution of Type II OCs across brain, intestinal, and hepatic barriers, whereas the renal phenotype in these animals is modest.[128]

Axial Distribution of Organic Cation Transport in the Renal Proximal Tubule

Renal secretion of TEA and procainamide by isolated perfused rabbit RPT shows a marked axial heterogeneity that differs from that of secretion of PAH [130] [131] with a profile of TEA secretion of S1 > S2 > S3,[112] and a profile of procainamide secretion of S1 = S2 > S3.[132] This axial distribution of secretory function is correlated (in rat and rabbit) with a marked difference in the distribution of distinct basolateral transporters, with OCT1 expression dominating in the early proximal tubule and OCT2 expression dominating in the mid and later portions of the proximal tubule. [133] [134] Despite these differences, the kinetics of basolateral TEA uptake, as determined in isolated, non-perfused tubules, is effectively the same in all three segments,[133] suggesting that the apical exit step for OCs is both rate limiting and the source of the axial heterogeneity observed for TEA secretion.[112] Consistent with this conclusion is the observation that the maximal rate of TEA/H+ exchange is significantly higher in rabbit renal BBMV isolated from outer cortex (enriched in membranes from S1/S2 segments) than from outer medulla (enriched in S3 segments[135]).

We can summarize the current, overall understanding of the cellular processes associated with secretion of organic cations as follows: Type I OCs enter RPT cells across the peritubular membrane via electrogenic facilitated diffusion (mediated by one or more OCT transporters) and leave cells across the luminal membrane via electroneutral exchange for H+ (possibly by means of one or more MATE and/or OCTN transporters). Type II OCs diffuse into proximal cells across the peritubular membrane and are exported into the tubule filtrate via the primary active MDR1 transporter. Importantly, considerable overlap appears to exist in the selectivity of these parallel transport pathways. [110] [136]

Organic Cation Reabsorption

Whereas secretion dominates the net flux of OCs transported by the proximal tubule, net reabsorption has been reported for a few cationic substrates, most notably choline. [89] [90] [137] [138] The apical membrane of renal proximal tubule cells expresses an electrogenic uniporter that accepts choline and structurally similar compounds with relatively high affinity. [139] [140] In contrast, the apical OC/H+ exchanger has a low affinity (but high capacity) for choline.[139] Consequently, choline is effectively reabsorbed when plasma concentration do not exceed the comparatively low, physiological concentrations (10 μM to 20 μM), and is secreted when concentrations are raised to levels >100 μM.[138]

Substrate Interactions and Renal Clearance of Organic Cations

The renal OC secretory process has sufficient transport capacity to extract >90% of many OCs, when present in low (clinically relevant) concentrations, during a single passage of blood through the kidney.[141] The presence of multiple OCs in the blood can result in competition between these compounds for one or more common elements in the OC secretory pathway, leading to decreased rates of elimination of one or more of these compounds with resultant elevation(s) in their blood levels (see Fig. 6-6 ). This has been shown to occur when the antiarrhythmic, procainamide, is administered with either cimetidine or ranitidine ( Fig. 6-7 ). [2] [92] [142] [143] The clinical impact of such interactions will depend on the therapeutic index of the drugs in question.

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FIGURE 6-7  Effect of drug-drug competition at the level of renal organic cation secretion on mean plasma concentration-time profiles of procainamide 000321   and n—acetylprocainamide 000865   in six subjects with 000327   or without 000329   co-administration of cimetidine.  (From Somogyi A, McLean A, Heinzow B: Cimetidine-procainamide pharmacokinetic interaction in man: Evidence of competition for tubular secretion of basic drugs. Eur J Clin Pharmacol 25:339, 1983.)

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Molecular Biology of Renal Organic Cation Transport

The cloning in 1994 of OCT1[144] by Gründemann and Koepsell resulted in a rapid increase in understanding of the molecular and cellular basis of renal OC transport. As outlined earlier, strong evidence supports the conclusion that basolateral entry of Type I OCs into RPT cells occurs by a (species specific) combination of the activities of OCT1, OCT2, and OCT3; and that apical exit of Type I OCs includes a combination of the activities of MATE1, MATE2, OCTN1, and OCTN2. The OCTs and OCTNs are all found within the SLC22A family of solute carriers and share a common set of structural features that place them within the Major Facilitator Superfamily (MFS) of transport proteins,[145] whereas the MATEs are members of the Multidrug/Oligosaccharidyl-lipid/Polysaccharide (MOP) superfamily.[146] Renal secretion of Type II OCs involves MDR1 in the luminal membranes of proximal tubular cells, although the role its activity plays in clearance of these compounds from the body is currently the subject of speculation. Following is a discussion of the molecular characteristics of the earlier listed transport proteins.

Basolateral Organic Cation Transporters

Organic Cation Transporters

Basolateral OC transport is dominated by the combined ac-tivity of three members of the SLC22A family of transport proteins, OCT1 (SLC22A1), OCT2 (SLC22A2), and OCT3 (SLC22A3).[18] As MFS transporters, they share several structural characteristics including 12 transmembrane spanning helices (TMHs), cytoplasmic N- and C-termini, a long cytoplasmic loop between TMHs 6 and 7, and several conserved sequence motifs. [18] [147] Several additional features are unique to the OCT members of SLC22A, including a long (≈110 amino acid residues) extracellular loop between TMHs 1 and 2, as well as a distinguishing sequence motif.[148] The human orthologs of OCT1, OCT2, and OCT3 contain 554, 555, and 556 amino acid residues, respectively, and several consensus sites for PKC-, PKA-, PKG-, CKII-, and/or CaMII-mediated phosphorylation located within or near the long cytoplasmic loop between TMHs 6 and 7, or in the cytoplasmic C-terminal sequence. [147] [149] The long extracellular loop between TMHs 1 and 2 contains three N-linked glycosylation sites in all three homologs. Elimination of these sites is associated with both decreased trafficking of protein to the membrane and with changes in apparent affinity for substrate,[150] the latter observation suggesting that the configuration of the long extracellular loop influences the position of TMHs 1 and 2, which are elements of the hydrophilic “binding cleft” common to the OCTs and in which substrate is suspected to bind ( [4] [151]; and discussed later).

The human genes for OCT1, OCT2, and OCT3 have 11 coding exons.[152] Several alternatively spliced variants of OCT1 have been described. rOCT1A lacks putative TMHs 1 and 2 and the large extracellular loop that separates those two TMHs, yet supports mediated transport of TEA.[153] In the human, four alternatively spliced isoforms of OCT1 are present in human glioma cells,[154] a long (full-length) form and three shorter forms. Only the long form (hOCT1G/L554) supports transport when expressed in HEK293 cells.[154] Human kidney expresses at least one splice variant of OCT2. Designated hOCT2-A, it is characterized by the insertion of a 1169 bp sequence arising from the intron found between exons 7 and 8 of hOCT2[155] resulting in a truncated protein that is missing the last three putative TMHs (i.e., 10, 11, 12). Despite the absence of the last three TMHs, hOCT2-A retains the capacity to transport TEA and cimetidine, though guanidine transport is lost.

In the rat and rabbit, OCT1 expression appears to dominate basolateral OC entry in the early (S1 segment) of renal proximal tubule, whereas OCT2 expression appears to dominate the mid and late (S2 and S3) segments of RPT. [133] [134] In the human kidney it is likely that basolateral OC transport is dominated by activity of OCT2. OCT2 is heavily expressed in the human kidney, and the relative expression profile of mRNAs coding for OCT1, OCT2, and OCT3 in human renal cortex is 1:100:10.[156] However, the observation that, in the rabbit, OCT1 activity dominates OC transport in the early proximal tubule, despite the fact that OCT2 mRNA expression is >10 times larger,[133]suggests that it would be premature to conclude that OCT1 (and OCT3) have no influence on renal clearance of selected compounds by human kidney.

The relative role of OCTs expressed in the proximal tubule may also be influenced by their site of expression. In the rodent, as in the rabbit, OCT expression in the early proximal tubule is dominated by OCT1, whereas OCT2 expression is restricted to the later portions of the RPT. [3] [134] Jonker and colleagues[3] found that targeted elimination of OCT1 actually resulted in an increase in renal clearance of TEA (presumably reflecting the increase in plasma TEA levels that resulted from the elimination of OCT1-mediated hepatic clearance of TEA), and elimination of OCT2 had no effect on renal clearance of TEA ( Fig. 6-8 ). In other words, the level of functional expression of each transporter was sufficient to maintain fractional clearance of TEA at control levels in the absence of the other. Importantly, the elimination of both OCT1 and OCT2 completely eliminated active clearance of TEA (see Fig. 6-3 ),[3] indicating that (in the mouse) OCT3 plays no significant role in renal clearance of TEA. Indeed, mice in which OCT3 has been eliminated display no apparent renal phenotype[157] (although OCT3 may still play a role in the renal elimination of substrates for which it displays a particularly high affinity[133]). Thus, under normal conditions, transporters restricted to later portions of the RPT may see little or no substrate if that compound is effectively cleared by transporters located in “upstream” portions of the tubule. Transport capacity in later portions of the tubule may only come into play when the activity in the early RPT is saturated or inhibited, as may occur in the event of a drug-drug interaction.

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FIGURE 6-8  Renal clearance of TEA in wild-type and Oct1/2-/- mice. Renal clearance was calculated by dividing the amount of TEA excreted in the urine over 60 minutes by the plasma AUC(0–60). The estimated GFR was approximately 21 ml/h for both genotypes and is indicated with a dashed line.  (From Jonker JW, Wagenaar E, Van Eijl S, et al: Deficiency in the organic cation transporters 1 and 2 (Oct1/Oct2 [Slc22a1/Slc22a2]) in mice abolishes renal secretion of organic cations. Mol Cell Biol 23:7902, 2003.)

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All the OCTs share a common transport mechanism (i.e., electrogenic uniport). Transport is independent of extracellular Na+ and H+, with membrane potential providing the driving force for transport of cationic substrates. [102] [158] The transport of positively charged substrates is electrogenic, as shown directly in studies characterizing the saturable inward currents that result from exposing Xenopus oocytes injected with the cRNA for OCT1,[102] OCT2,[158] or OCT3[159] to increasing concentrations of substrate. Koepsell and colleagues[102] showed that membrane potential provides the driving force for OCT1-mediated TEA, NMN, and choline uptake, and that OCT1 can also support the electrogenic efflux of substrate in the presence of energetically favorable outwardly directed substrate gradients, as well as electroneutral OC/OC exchange.

Although the three OCTs display marked overlap in substrate selectivity, they are also distinguished by their selectivities for specific compounds. For example, OCT1 and OCT2 generally have a similar affinity for TEA (20 μM-200 μM)[133] whereas OCT3 has a very low affinity for TEA (≈2 μM[160]); Cimetidine has a much higher (50-fold) affinity for OCT2 than OCT1,[161] whereas tyramine has a higher affinity (20-fold) for OCT1 than OCT2[161]; and all three homologs display a similar, comparatively high affinity for MPP.[162] In general, the three homologs all support transport of a structurally diverse array of Type I OCs,[133] and interact with a limited number of neutral and even anionic substrates.[163] With respect to the latter observation that OCTs can interact with (selected) neutral or anionic substrates, Ullrich and colleagues observed “cross-over” interactions of a number of what they referred to as “bisubstrates” with both cation and anion transport pathways in rat kidney. [164] [165] Nevertheless, despite the (generally weak) interaction of neutral and anionic substrates with OCTs, the presence of a charged moiety clearly enhances interaction with these transporters, as shown in studies demonstrating more efficient interaction of the weak base, cimetidine (pKa = 6.9), with OCT2 when the substrate is protonated.[166] OCTs also typically interact with Type II OCs, though this interaction generally appears to be restricted to binding with modest or no translocation of substrate.[110]

It is likely that the kinetics of binding of the OCTs with many, if not most, substrates is asymmetric (i.e., differs when the interaction occurs at the extracellular versus cytoplasmic face of the membrane). Koepsell and colleagues,[167] using giant excised patches of Xenopus oocyte membrane, determined that the binding of corticosterone and tetrabutylammonium to the extracellular face of rat OCT2 is 20-fold lower versus 4-fold higher, respectively, than that measured for binding to the intracellular face of the transporter. This is not a surprising observation when considered in the light of information concerning the probable 3D structure of OCTs and the likelihood that binding regions will have (at least) modestly different 3D configurations when exposed to the inside versus the outside of cells. This issue is discussed in more detail in an upcoming section.

In addition to operating as electrogenic uniporters, the OCTs also support OC/OC exchange. [102] [168] [169] [170] [171] [172] Preloading Xenopus oocytes with unlabeled TEA, for example, stimulates the uptake of [3H]MPP by human, rabbit, mouse, and rat OCT1.[168] The symmetry of this type of trans-effect is apparent in observations of accelerated efflux of preloaded [3H]MPP from rOCT1-expressing oocytes in the presence of inwardly-directed gradients of unlabeled TEA[102] or MPP.[169] Human OCT1 also supports trans-stimulation of both influx and efflux (of TEA), but quantitative differences in the extent of these stimulated fluxes produced by some substrates (e.g., tributylmethylammonium) support the suggestion, as discussed earlier, of asymmetrical binding properties on the extracellular versus intracellular face of the transporter.[170]

Organic Cation Transporter Structure

The elucidation of the crystal structure of two MFS transporters, LacY,[173] and GlpT,[174] and the discovery that these two proteins share a marked structural homology (i.e., a common helical fold) despite having a low sequence homology (<15%), paved the way for efforts to use homology modeling as a means to develop structural models for other MFS transporters.[175] LacY and GlpT have served as “templates” for the modeling of OCT1[151] and OCT2,[4] respectively, and the resulting models share a number of common structural features (owing, in part, to shared structural features of the templates), including a large hydrophilic “cleft” formed by the juxtaposition of the N- and C-terminal halves of the proteins that is comprised of the amino acid residues of the “pore forming” helices: TMHs 1, 2, 4, 5, 7, 8, 10, and 11 ( Fig. 6-9 ). Significantly, amino acid residues that have been in-dependently shown in site-directed studies to influence substrate binding are found, in both models, at locations consistent with roles in stabilizing substrate-transporter interactions. In particular, an aspartate residue in TMH 11 that is conserved in all OCT homologues (i.e., D475 in hOCT2) and that markedly influences substrate binding in rOCT1[176] and in rOCT2,[177] is directed toward the hydrophilic pore at a position within the protein that coincides closely to the binding site identified in both GlpT[174] and in LacY[173] (see Fig. 6-9 ). Similarly, residues within TMHs 4 and 10 that influence substrate binding are also directed toward the pore region of OCT1[151] and OCT2,[4] including three residues in TMH10 that play a key role in defining the selectivity differences that distinguish OCT1 and OCT2. [4] [151] [178] The comparatively large extent of the pore or cleft region of OCTs (20 Å × 60 Å × 80 Å[151]), is consistent with the suggestion that the broad substrate selectivity these proteins reflects binding interactions over a large surface that contains several distinct sites or regions. [4] [178]

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FIGURE 6-9  Model of the 3D structure of the rabbit ortholog of OCT2, based on structural homology with the MFS transporter, GlpT. A, Side view of OCT2, with the cytoplasmic face directed downward. The helices (TMHs 1-6) comprising the N-terminal half of the protein are colored blue; the helices comprising the C-terminal half of the protein are colored cyan. The lighter colored helices (1, 2, 4, 5, 7, 8, 10, and 11) border the hydrophilic cleft region formed by the juxtaposition of the N- and C-terminal halves of the protein. The amino acid residues that comprise the postulated substrate-binding region within the cleft are rendered as sticks with a pink colored van der Waals surface. D475 is rendered as a space-filling reside in orange. Note: residues from the long extracellular loop (between TMHs 1 and 2) and the cytoplasmic loop (between TMHs 6 and 7) were eliminated to facilitate homology modeling with the GlpT template. B, An end-on view of the cleft and postulated binding region from the cytoplasmic aspect of the protein.  (From Zhang X, Shirahatti NV, Mahadevan D, et al: A conserved glutamate residue in transmembrane helix 10 influences substrate specificity of rabbit OCT2 (SLC22A2). J Biol Chem 280:34813, 2005.)

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The recent discovery of altered transport function of hOCT1 and hOCT2 that contain single nucleotide polymorphisms present in different ethnic populations [179] [180] [181] has underscored the importance of understanding structure-activity relationships for these processes. For example, 28 variable sites in the hOCT2 gene were discovered in a collection of 247 ethnically diverse DNA samples (White, African American, Asian American, Mexican American, and Pacific Islander). Eight of these polymorphisms caused non-synonymous amino acid changes, of which four were present in at least 1% of an ethnic population. These four displayed altered transporter function, including up to a threefold change in Kt values for MPP and TBA, changes that could result in differences in the pharmacokinetics of renal drug excretion between individuals expressing different variants of hOCT2. However, population-genetic analysis suggests that selection has acted against amino acid changes to hOCT2, which may reflect a necessary role of OCT2 in the renal elimination of endogenous amines or xenobiotics.[179]

Regulation of Organic Cation Transporter-Mediated Transport

Organic cation transporter activity responds to both short- and long-term regulation, although there appear to be significant species differences in the extent of such responses.[149] For example, when expressed heterologously, activation of protein kinase A acutely up-regulates rat OCT1-mediated transport,[182] but down-regulates human OCT1-mediated transport.[183] Of particular significance to the issue of short-term regulation of renal OC transport in humans is the observation that basolateral uptake of the fluorescent cation 4-[4-(dimethylamino)-styryl]-N-methylpyridinium into isolated single non-perfused proximal tubules from human kidney, is acutely down-regulated following activation of PKC,[184] and this presumably reflects acute regulation of OCT2 activity. In fact, hOCT2 (expressed heterologously) is acutely down-regulated following activation of PKA, PKC, Ca2+/CaM, or PI3-kinase.[185] The decrease in transport associated with acute activation of these kinases appears to reflect a decrease in the maximal rate of transport (i.e., Kt is not affected[186]), consistent with the hypothesis that acute downregulation of OCT2 activity reflects the rapid sequestration of transporters into an cytoplasmic vesicular compartment, a mechanism that has been shown to account for the acute down-regulation of the closely related OAT transporters.[187]

Sex steroids have been shown to regulate the long-term activity of OCT2. TEA uptake is greater in renal cortical slices of male rats than female rats, and this is correlated with a higher level of expression (mRNA and protein) of OCT2 in kidneys of male rats, with no sex-linked differences in either OCT1 or rOCT3.[188] Moreover, treatment of male and female rats with testosterone significantly increases OCT2 expression in the kidney,[189] via the androgen receptor mediated transcriptional pathway,[190] suggesting that testosterone plays a significant role in the transcriptional regulation of the OCT2 gene in rats. A similar profile is not, however, evident in all species. Although OCT2 mRNA expression is higher in kidneys from male rabbits than female rabbits, this difference does not extend to either protein expression or in rates of TEA transport in renal tubules isolated from male and female rabbit kidneys.[191] On the other hand, humans, like rats, exhibit a significant sex difference in the renal excretion of the OC substrate, amantadine (which is a substrate for OCT1, OCT2, and OCT3 [192] [193]). It is noteworthy that renal clearance involves transport across both basolateral and apical membranes. Thus, sex differences in renal clearance may involve apical transporters either in addition to or rather than basolateral transporters.

Apical Organic Cation Transporters

Although the physiological characteristics of apical OC transport have been studied extensively using isolated BBMV (see Refs 89, 133), the molecular identity of the processes that mediate the exit step in renal secretion of Type I OCs, particularly the identity of the “OC/H+ exchanger,” is still unclear. The OCTNs (i.e., OCTN1 and OCTN2) have been implicated and may well play a role in the mediating transport of at least some OCs. Perhaps more attractive, though still in a very early state of study, are the MATEs, which have a physiological profile consistent that of apical OC transport observed in isolated renal membranes and intact renal tubules.

OCTN

OCTN1 (SLC22A4) and OCTN2 (SLC22A5) are 551-557 amino acid peptides with ≈30% to 33% sequence identity with hOCT1. As with other members of the OCT family, the OCTNs have 12 putative TMHs and, consistent with the OCTs, 3 N-linked glycosylation sites and a number of consensus sites for phosphorylation mediated by PKA, PKC, and CKII. OCTN1, although widely expressed in human tissues, is weakly expressed in the kidney.[156]OCTN2, in contrast, is most heavily expressed in kidney, heart, placenta, skeletal muscle and pancreas.[194]

OCTN1 supports electroneutral transport of TEA, and flux of TEA across Xenopus oocyte membrane is trans-stimulated by oppositely oriented H+ gradients.[120] It is this latter observation that led to speculation that OCTN1 may play a role in the OC/H+ exchange observed in isolated renal BBMV preparations. However, TEA/H+ exchange has been noted in very few tissues, most notably in isolated membranes from kidney (apical BBMV[133]) and liver (canalicular BBMV[195]). OCTN1, however, is expressed in many tissues, including the placenta and intestine,[196] neither of which support TEA/H+ exchange. The comparatively low level of expression of OCTN1 in human kidney[156] also appears to be inconsistent with the observation that OC/H+ exchange is the dominant mechanism for OC flux across isolated human renal brush border membrane vesicles.[197] In addition, the kinetic/selectivity characteristics of OCTN1 are inconsistent with this process playing a major role in luminal OC transport. As noted earlier, a transporter with even modest levels of expression may play a quantitatively significant role in renal secretion of selected substrates, so OCTN1 could influence the secretion of some OCs. However, the paucity of data on the selectivity characteristics of OCTN1 makes it difficult to draw conclusions concerning its potential role in renal OC transport.

OCTN2 appears to be unique in that it supports both Na-dependent transport of the zwitterion, carnitine (and related compounds), and Na-independent, electrogenic facilitated diffusion of Type I OCs. Importantly, lesions in OCTN2 result in primary carnitine deficiency in humans [198] [199] and mice,[200] and the Na-dependent interaction of OCTN2 with carnitine and its role in supporting the reabsorption of this important metabolite is discussed extensively elsewhere. [201] [202] The Na-independent transport of TEA, MPP, and a wide range of other Type I OCs [122] [194] occurs electrogenically,[122] and operating in this mode OCTN2 would be expected to support OC reabsorption, rather than secretion. Significantly, however, jvs mice, which express an inoperative OCTN2 (and display primary carnitine deficiency), show a 50% reduction in TEA clearance and a 2.5-fold increase in the kidney-plasma ratio of TEA,[203] both of which implicate OCTN2 in the apical efflux step of TEA secretion. A possible, and intriguing, explanation for these results may be found in the observation that outwardly directed TEA gradients trans-stimulate OCTN2-mediated carnitine-Na cotransport.[203] Thus, OCTN2-mediated reabsorption of carnitine could serve as a driving force to support electroneutral luminal efflux of TEA (and selected Type I OCs); under normal physiological conditions, the comparatively high concentrations of Na+ and carnitine in the plasma (and filtrate) should maximize the operation of OCTN2 as a reabsorptive pathway for carnitine, rather than as an electrogenic pathway for Type I OC reabsorption, and in the presence of elevated cytoplasmic levels of a suitable Type I substrate (e.g., TEA), OCTN2 would mediate the secretory efflux of that molecule (e.g., Na-carnitine/TEA exchange) (see Fig. 6-6 ).

MATE

Two members of the MOP family of transport proteins, MATE1 and MATE2, were recently cloned in the human and mouse. [123] [124] A splice variant of MATE2 (MATE2-K) was also identified in human kidney.[204] The human ortholog of MATE1 is a 570 aa protein with 12 putative TMHs, no N-linked glycosylation sites (in extracellular loops), and 3 consensus PKG phosphorylation sites (and no PKC, PKA, CKII, or CaMII sites). In the human, MATE1 is expressed in the kidney and liver (and, less so in the heart), where it is found in the apical and canalicular membranes, respectively, of renal proximal tubules and hepatocytes.[123] Importantly, MATE1-mediated transport of TEA is electroneutral, pH sensitive, and markedly trans-stimulated by oppositely-oriented H+ gradients. TEA transport is cis-inhibited by Type I OCs, including cimetidine, quinidine, and MPP, only weakly inhibited by NMN and choline, and is refractory to guanidine, PAH and probenecid.[124] Thus, the (1) profile of expression, (2) energetic mechanism, and (3) selectivity characteristics of MATE1 are reasonably comparable to those of the apical OC/H+ exchanger as expressed in renal BBMV, making MATE1 a strong candidate for the molecular identity of this process. MATE2 is also expressed in the kidney, but not in the liver, and its physiological characteristics remain undetermined; its role in mediating renal OC transport is, therefore, unclear. However, its splice variant, MATE2-K, displays characteristics similar to those of MATE1, and its kidney-specific expression suggests that it, too, may play a role in the luminal export of OCs.

Multidrug Resistance

The multidrug resistance transporter (MDR1; ABCB1; also called the P-glycoprotein, or p-GP) was first characterized within the context of its role in the development of cross-resistance of cancer cells to a structurally diverse range of chemotherapeutic agents. The human ortholog of MDR1 is a protein of 1279 amino acids (141 kDa) and is composed of two homologous halves, each containing six TMDs and an ATP-binding domain, separated by a linker polypeptide. The normal expression of MDR1 in barrier epithelia, including the intestine, liver, and kidney, supports the conclusion that it plays a role in limiting absorption (in the intestine) and facilitating excretion (by the liver and kidney) of xenobiotic compounds. In the kidney, MDR1 is expressed in the apical membrane of proximal tubule cells in human[205] and mouse kidneys.[206] MDR1 is also expressed, albeit at apparently lower levels, in the mesangium, thick ascending limb of Henle's loop, and collecting tubule of the normal human kidney[207] MDR1 supports ATP-dependent export of a structurally diverse range of comparatively bulky, hydrophobic cationic substrates that, in general, fall within the Type II OC classification. These traditional MDR1 substrates include the vinca alkaloids (e.g., vinblastine, vincristine), cyclosporine, anthracyclines (e.g., daunorubicin, doxorubicin), and verapamil. In addition, MDR1 mediates the transport of a number of relatively hydrophobic compounds that are either uncharged or are neutral at physiological pH, including digoxin, colchicine, propafenone, and selected corticosteroids. Although substantial evidence supports the conclusion that MDR1 is involved in the luminal secretion of at least Type II OCs, its quantitative significance is not known.

Clinical Diseases from Genetic Defects of Organic Cation Transporters

Lesions in OCTN2 have been clearly linked to systemic carnitine deficiency, owing to the central role this trans-porter plays in reabsorption of carnitine from the tubular filtrate. [200] [208] [209] In addition, single nucleotide polymorphisms (SNPs) in the genes coding for OCTN1 and OCTN2 have been linked to increased incidence of inflammatory bowel diseases, including Crohn disease.[210] There is not, however, a clear disease phenotype associated with the failure of renal OC secretion, as shown by the normal phenotypes of mice in which OCT1, OCT2, and/or OCT3 have been eliminated. [3] [133] Instead, the focus has been on the influence of naturally occurring genetic variation in human populations for the OCTs and OCTNs and the influence such variation may have on the pharmacokinetics of drug elimination (see Ref 180, 211). For example, a study of six monozygotic twin pairs showed that genetic factors contribute substantially to the renal clearance of metformin, a drug that is a substrate of OCT2 and eliminated exclusively by the kidney. Indeed, the genetic component contributing to variation in the renal clearance of metformin, which undergoes transporter-mediated secretion, is suspected of being particularly high (>90%),[181] suggest that variation in the renal clearance of metformin has a strong genetic component, and that genetic variation in OCT2 may explain a large part of this pharmacokinetic variability. Common variants of OCT2, as well as genetic variants of OCT1, OCTN1, and OCTN2, may alter protein function and could cause inter-individual differences in the renal handling of organic cation drugs. Genetic variants of all these processes have been identified in human populations [179] [180] [181] [212] [213] [214] and studies in heterologous expressions systems have confirmed that common (typically, with occurrences in specific population groups of >1%) SNPs result in substantial changes in transporter activity. Further studies examining the pharmacokinetic phenotypes of individuals harboring genetic variants that change transport function should help to define the roles of each transporter in renal elimination.[211] Furthermore, such studies may help identify particular genetic variants that lead to susceptibility to drug toxicities resulting from drug-drug interactions.

ORGANIC ANIONS

Organic Anion Physiology

Organic anions represent an immensely broad group of solutes being transported by the kidney, which renders it barely justifiable to be discussed in a single section. An organic anion can be loosely defined as any organic compound that bears a net negative charge at the pH of the fluid in which the compound resides. These can be endogenous substances, or exogenously acquired toxins or drugs. The physiology can be poised for conservation with extremely low fractional excretion similar to glucose. Such is the case with metabolic intermediates like mono- and di-carboxylates ( Fig. 6-1 ; fractional excretion ≈ 0). On the other end, the system can gear itself for elimination utilizing combined glomerular filtration and secretion ( Fig. 6-1 ; fractional excretion >>1). In addition to the large range of fractional excretion, this group of transporters also has the broadest array of substrates that spans compounds with completely disparate chemical structures. Multi-specificity in substrate recognition is a prevalent feature within each gene family and across different families of organic anion transporters.

The precise analysis of the field of renal anion excretion was ushered by the seminal work of Marshall and co-workers who studied the elimination of dyes and arrived at the conclusion that mammalian renal tubules possess high capacity secretory function. [215] [216] This was followed by the classical studies of Smith and associates who described the tubular secretion of p-aminohippurate (PAH) and provided a marker for estimating renal blood flow (RPF) by PAH clearance for decades that followed.[217] Reabsorptive physiology was illustrated in the previous section on glucose. Figure 6-5 illustrates the secretory nature of the proximal tubule using PAH as a surrogate. In low plasma concentrations, PAH has a fractional excretion of >>1 and PAH clearance (CPAH) approaches renal plasma flow (RPF) as most of the PAH is removed from the plasma in a single pass. As plasma PAH increases, both filtered and secreted PAH increases and CPAH remains a good estimate of RPF. When the secretory maximal is reached and subsequently exceeded, the commensurate increment in excretion is contributed solely by increasing filtered load. At this stage, CPAH starts to gradually drift further below RPF toward the value of GFR ( Fig. 6-10 ).

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FIGURE 6-10  Illustration of filtration-secretion using PAH clearance (CPAH). PAH clearance = GFR + PAH secretion. For a given GFR, both secreted and filtered PAH increases with increasing plasma [PAH]. At this point, CPAH ap-proximates renal plasma flow (RPF). With increasing plasma [PAH], maximal secretion is reached and any further increase in CPAH is due to increasing filtered PAH. At high plasma [PAH], CPAH numerically drifts towards GFR.

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Classic studies using stop-flow, micropuncture, and microperfusion [218] [219] [220] in multiple species have demonstrated that organic anions are secreted in the proximal tubule. The study of Tune and co-workers definitely demonstrated uphill transport from peritubular fluid into urinary lumen.[220] As noted earlier, the secretory mode mandates broad substrate recognition and one simply cannot afford to devote one gene per compound that the organism wishes to excrete. Table 6-4 is an illustrative but incomplete inventory that demonstrates the extremely broad substrate spectra of organic anion handling by the kidney. It is impossible to fathom any structural similarities among these compounds. In addition, the number of substances transported far exceeds the number of proteins required to excrete these substance. This is not unlike proteins such as P-glycoprotein (ATP-binding cassette multidrug resistance protein) or the multi-ligand receptor megalin where the ability to engage with multiple compounds is intrinsic to their biologic function. [221] [222] The classical microperfusion study from Fritzch and co-workers proposed a minimal requirement of a hydropic region in the anion to be a substrate.[223] The protein structure that permits this broad range of substrate to be bound and transported is unknown but undoubtedly fascinating.


TABLE 6-4   -- Classes of Organic Anions Transported by the Proximal Tubule

Endogenous

Metabolic intermediates

α-ketoglutarate, succinate, citrate

Eicosanoids

PGE1, PGE2, PGD2, PGF2a, PGI2, TxB2

Cyclic nucleotides

cAMP, cGMP

Others

Urate, folate, bile acids, oxalate, 5-HIA, HVA

 

Metabolic Conjugates

Sulfate

Estrone sulfate, DHEAS,

Glucuronide

Estradiol glucuronide, salicylglucuronide

Acetyl

Acetylated sulphonamide

Glycine

PAH, o-hydroxyhippurate

Cysteine

CTFC, DCVC, N-acetyl-S-farnesyl-cysteine

 

Drugs

Antibiotics

β-lactam, cepham, tetracycline, sulphonamide

Antiviral

Acyclovir, amantadine, adefovir

Anti-inflammatory

Salicylates, indomethacin,

Diuretics

Loop diuretics, thiazides, acetazolamide

Antihypertensive

ACE inhibitors, ARB

Chemotherapeutic

Methotrexate, azathioprine, cyclophosphamide, 5-FU

Antiepileptic

Vaproate

Uricosuric

Probenicid

 

Environmental Toxins

Fungal products

Ochratoxin A & B, aflatoxin G1, patulin

Herbicides

2,4-dichlorophenoxyacetic acid

 

PG, prostaglandins; Tx, thromboxane; 5-HIA, 5-Hydroxyindoleacetate; HVA, homovanillic acid; DHEAS, dihydroxyepiandronesterone sulfate; PAH, p-amino hippurate; CTFC, S-(2-chloro-1,1,2-trifluoroethyl)-L-cysteine; DCVC, S-(1,2-dichlorovinyl)-L-cysteine; ACE inhibitors, angiotensin converting enzyme inhibitor; ARB, angiotensin receptor blockers; 5-FU, 5-fluorouracil.

 

 

 

Molecular Biology of Organic Anion Transporters

Several families of solute transporters can be included in this discussion. Three will be mentioned: the dicarboxylate-sulphate transporters (NaDC/NaS SLC13 family), the organic anion transporters (OAT SLC22 family), and the organic anion transporting polypeptides (OATP SLC21 family). A detail account is beyond the scope of this chapter. The reader is referred to several excellent recent reviews. [224] [225] [226] [227] [228] [229] [230] [231]

NaDC (SLC13A) Family

These transporters function to reclaim filtered solutes and are functionally directly opposite to the next group of secretory proteins. This family is related by similarities in primary sequences but the isoforms are quite distinct in their function. The nomenclature is still in a state of evolution and five genes are identified to date ( Table 6-5 ). [229] [230] NaS1 is a low-affinity sulfate transporter[232] expressed at the proximal tubule apical membrane (see Table 6-5 )[233] but does not take organic anions. NaS2 and NaCT are not expressed in the kidney (see Table 6-5 ). These proteins will not be discussed. NaDC1 and NaDC3 are the main transporters of interest in this discussion.


TABLE 6-5   -- Organic Anion Transporters

NaDC Family

Name

Gene Name

Human Chromosome

Renal Proximal Tubule Localization

Transport Mode/substrate (All Na+-dependent)

NaS1

SLC13A1

7q31-32

Apical

Sulphate, thiosulfate selenate

NaDC1

SLC13A2

17p11.1-q11.1

Apical

Succinate, citrate, α-ketoglutarate

NaDC3

SLC13A3

20q12-13.1

Basolateral

Succinate, citrate, α-ketoglutarate

NaS2

SLC13A4

7q33

Absent

Sulphate

NaCT

SLC13A5

12q12-13

Absent

Citrate, succinate, pyruvate

 

OAT Family

Name

Gene Name

Human Chromosome

Renal Proximal Tubule Localization

Transport Mode/substrate (Na+-independent)

OAT1

SLC22A6

11q12.3

Basolateral

OA dicarboxylate exchange

OAT2

SLC22A7

6q21.1-2

Basolateral

OA dicarboxylate exchange

OAT3

SLC22A8

11q12.3

Basolateral

OA dicarboxylate exchange

OAT4

SLC22A11

11q13.1

Apical

OA dicarboxylate exchange

URAT1

SLC22A12

11q13.1

Apical

Urate OA exchange

OAT5

Slc22a19

(murine)

 

 

OA, organic anion (broad substrate specificity).

 

OATP

Name

Gene Name

Human Chromosome

Renal Tubule Localization

Transport Mode/substrate (Na+-independent)

OATP4C1

SLCO4C1

5q21

PT: basolateral

Digoxin, ouabain, T3

OATP1A2

SLCO1A2

12p12

CCD: basolateral

Bile salts, estrogen conjugates PG's, T3, T4, antibiotics ouabain, ochratoxin A

OATP2A1

SLCO2A1

3q21

mRNA+

PG's

OATP2B1

SLCO2B1

11q13

mRNA+

Estrogen conjugates, antibiotics

OATP3A1

SLCO3A1

15q26

mRNA+

Estrogen conjugates, antibiotics

OATP4A1

SLCO4A1

20q13.1

mRNA+

Bile salts, estrogen conjugates, PG's, T3, T4, antibiotics

 

PT, proximal tubule; CCD, cortical collecting duct; T3, thyroid hormone; PGs, prostaglandin.

 

 

 

NaDC1

NaDC1 was first cloned by Pajor's group [234] [235] [236] [237] and subsequently by others. [238] [239] NaDC1 is found on apical membranes of both the renal proximal tubule and small intestine where it mediates absorption of tricarboxylic acid cycle intermediates from the glomerular filtrate or the intestinal lumen. The preferred substrates of NaDC1 are 4-carbon dicarboxylates such as succinate, fumarate, and α-ketoglutarate. Citrate exists mostly as a tricarboxylate at plasma pH, but in the proximal tubule lumen, because of apical H+ transport, citrate3- is titrated (citrate3-/citrate2- pK 5.7–6.0) and is taken up in protonated form as citrate2-. The Km for dicarboxylates ranges between 0.3 μM and 1 μM. Transport of one divalent anion substrate is coupled to three Na+ ions.

Once absorbed across the apical membrane, cytosolic citrate is either metabolized through ATP citrate lyase, which cleaves citrate to oxaloacetate and acetyl CoA, or transported into the mitochondria where it can be metabolized in the tricarboxylic acid cycle to neutral end products such as carbon dioxide ( Fig. 6-11 ). [240] [241] When a divalent organic anion is converted to neutral products, two H+ are consumed, which renders citrate2- an important urinary base.

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FIGURE 6-11  Proximal tubule citrate absorption and metabolism. The Na+-K+-ATPase generates the low cell [Na+]. As a secondary active transporter NaDC1 uses the electrochemical gradient to pick up filtered citrate, which metabolized in the cytoplasm or the mitochondria. Ambient and cytoplasmic pH increase citrate uptake and metabolism. (1) Acidification of urinary lumen titrates citrate to the divalent transported species; (2) NaDC1 is directly activated by pH and chronic low pH increases expression of NaDC1 (circled arrow); (3) Intracellular acidification increases the expression of ATP citrate lyase and aconitase (circled arrows).

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NaDC3

NaDC3 has a wider tissue distribution and much broader substrate specificity than NaDC1. NaDC3 is expressed on basolateral membranes in renal proximal tubule cells,[242] as well as liver, brain, and placenta. The basolateral location of NaDC3 was mapped to a motif in its amino-terminal cytoplasmic domain. [243] [244] [245] [246] The Km for succinate in NaDC3 is lower than NaDC1 (10 μM-100 μM).[246] Similarly, NADC3 displays a much higher affinity for α-ketoglutarate[247] than does NaDC1.[248] Like NaDC1, NaDC3 is sodium-coupled and electrogenic so it is very unlikely that NaDC3 will mediate citrate efflux from the proximal tubule into the peritubular space. It is more likely that the NaDC3 helps support the outwardly directed α-ketoglutarate gradient required for OAT transporters to perform organic anion exchange (see later). In fact, the activity of NaDC3 has been shown to support approximately 50% of the OAT-mediated uptake of the organic anion fluorescein across the basolateral membrane in isolated rabbit renal tubules[249] with half of this effect reflecting the accumulation of exogenous α-ketoglutarate from the blood, and the other half arising from “recycling” endogenous α-ketoglutarate that exited the cell in OAT-mediated exchange for the organic anion substrate.

OAT (SLC22A) Family

Two features of these transporters should once again be emphasized—their high capacity for substrate and tremendously diverse substrate selectivity (see Table 6-4 ). The importance of these proteins in rescuing the organism from succumbing to toxins cannot be over-emphasized. The uptake of substrates from the basolateral membrane of the proximal tubule is a thermodynamically uphill process utilizing tertiary active transport ( Fig. 6-12 ). The Na+ and voltage gradient generated by the Na+-K+-ATPase drives the accumulation of the dicarboxylate α-ketoglutarate in the proximal tubule via NaDC3, which in a tertiary fashion (thrice removed from ATP hydrolysis) energizes uptake of organic anions into the proximal tubule (see Fig. 6-12 ). Endogenously produced α-ketoglutarate from deamination and deamidation of glutamine (ammoniagenesis) may also participate in the exchange process. Some of the organic anions transported may be endogenous or relatively innocuous exogenous compounds but many of the substrates (see Table 6-4 ) are toxins. Although its function is in defending the body, the proximal tubule cells cannot afford a self-sacrificial approach as the end result can be destruction of the very mechanism that secretes these toxins. There exists detoxifying mechanism in the proximal tubule cell that protects the cell while the toxins are en route to the apical membrane to be disposed. The details of these mechanisms are still elusive but current data in isolated proximal tubules and cell culture models suggests compartmentalization that may serve to sequester the toxins from imparting their harmful effects.[250]

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FIGURE 6-12  NaDC (green), OAT (blue), and OATP (black) families of anionic transporters in the proximal tubule. OA-, organic anion; Ur-, urate. The intracellular transport and sequestration of organic anions are not understood.

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Basolateral Transporters

More than half a century after the seminal paper from Homer Smith's laboratory[251] that described PAH secretion into the urine, the “PAH transporter” was cloned by several laboratories almost contemporaneously. [252] [253] [254]The OAT members OAT1 and OAT3 are present in the basolateral membrane of the proximal tubule (see Fig. 6-12 , Table 6-5 ). OAT1-mediated uptake of PAH is stimulated by an outwardly directed gradient of dicarboxylates such as α-ketoglutarate, indicating that OAT1 is an organic anion-dicarboxylate exchanger.[255] The substrate selectivity of OAT1 is extremely broad with affinities for substrate that are comparable to that reported for the functional PAH transport system. OAT3 is localized in the basolateral membrane of the kidney and, like OAT1, has a broad extra-renal expression.[256] OAT3 also has a promiscuous substrate list comparable to that of OAT1.[231] The purpose for the OAT1/3 redundancy in the kidney is unclear. OAT2 was originally identified from the liver and its expression in the kidney appears to be weaker than OAT1 and OAT3.[257] It transports PAH, dicarboxylates, prostaglandins, salicylate, acetylsalicylate, and tetracycline.[231]

Apical Transporters

There is no overlap of polarized expression of OATs in the proximal tubule. OAT4 was cloned from the kidney and is expressed in the apical membrane of the proximal tubule.[258] When characterized in oocytes, it transports PAH, conjugated sex hormones, prostaglandins, and mycotoxins in an organic anion/dicarboxylate exchange mode and is capable of bi-directional movement of organic anions.[259] It is not known whether OAT4 represents an exceptional OAT-mediated luminal uptake, although it is hard to fathom from the list of candidate substrates why OAT will participate in absorption. The other apical transporter is URAT1, which is renal-specific in its expression.[260] The human URAT1 appears to be quite specific for urate transport.[260] As discussed later, the role of URAT1 as a urate transporter was proven at the whole organism levels from an experiment of nature in humans with renal hypouricemia.

OATP (SLCO) Family

This family of organic anion transporting polypeptides is expressed widely in the brain, choroid plexus, liver, heart, heart, intestine, kidney, placenta, and testis[261] and, like the OATs, they also have a wide spectrum of substrates.[262] The first member oatp1 was cloned from rat liver by Meier's group as a sodium-independent bile acid transporter.[263] Eleven human isoforms and even more rodent isoforms have been appended to the OATP family. [224] [264]In place of the older nomenclature of SLC21A, a new nomenclature has recently been assigned to the OATP family of solute transporters, [226] [231] which subdivides the OATP superfamily into multiple sub-families (reviewed in Ref 226 ). A comprehensive discussion of this complex classification is beyond the scope of this chapter. One noteworthy point is that there are considerable inter-species differences that engender difficulties in extrapolating rodent data to humans. Among human OATPs, only OATP4C1 is predominantly and definitively expressed in the kidney. The myriad of rodent isoforms that have not been confirmed in humans will not be discussed in this section (an excellent account can be found in two recent reviews). [226] [231] One important compound carried by OATP2C1 is the cardiac glycoside digoxin.[264] OATP4C1 is expressed exclusively in the basolateral membrane of proximal tubular cells and mediates the high-affinity transport of digoxin (Km: 7.8 μM) and ouabain (Km: 0.38 μM), as well as thyroid hormones such as triiodothyronine (Km: 5.9 μM). The apical pathway for digoxin has been presumed to be an ATP-dependent efflux pump such as P-glycoprotein.

Clinical Relevance of Organic Anion Transporters

NaDC1

The role of NaDC1 in physiology and pathophysiology has been well studied. Citrate has multiple functions in mammalian urine and the two most important ones are as a chelator for urinary calcium, and as a physiologic urinary base.[265] It is a tricarboxylic acid cycle intermediate, and the majority of citrate reabsorbed by the proximal tubule is oxidized to electroneutral end products so H+ is consumed in the process rendering citrate a major urinary base (see Fig. 6-11 ). Calcium associates in a one-to-one stoichiometry. The highest affinity and solubility is a monovalent anionic (Ca2+Citrate3-)- complex.[265]

The final urinary excretion of citrate is determined by reabsorption in the proximal tubule and the most important regulator of citrate reabsorption is proximal tubule cell pH. Acid loading increases citrate absorption by four mechanisms (see Fig. 6-11 ): (1) Low luminal pH titrates citrate3- to citrate2- which is the preferred transported species[266]; (2) NaDC1 is also gated by pH such that low pH acutely stimulates its activity[267]; (3) Intracellular acidosis increases expression of the NaDC1 transporter[268] and insertion of NaDC1 into the apical membrane; (4) Intracellular acidosis stimulates enzymes that metabolize citrate in the cytoplasm and mitochondria. [268] [270] This is a well concerted response and an appropriate response of the proximal tubule to cellular acidification is hypocitraturia. Although perfectly adaptive from an acid-base point of view, this response is detri-mental to prevention of calcium chelation. All condi-tions that lead to proximal tubular cellular acidification (e.g., distal renal tubular acidosis, high-protein diet, potas-sium deficiency) are clinical risk factors for calcareous nephrolithiasis. Hypocitraturia can cause kidney stones by itself or by acting with other risk factors such as hypercalciuria, and therapy with potassium citrate has been shown to reverse the biochemical defect and reduce stone recurrence.[271]

URAT1 and Hyperuricosuria

The model for renal handling of uric acid has been rather controversial with a popular but yet unproven paradigm of tandem filtration-reabsorption-secretion-reabsorption. Molecular identity and functional evidence of the proteins involved are just beginning to emerge. The current model suggest that apical urate absorption is mediated by URAT1, [260] [272] [273] [274] whereas apical secretion is mediated the ATP-binding cassette protein MRP4 [273] [274] and the galectin-9/uric acid transporter (UAT). [275] [276] [277]

A host of uricosuric substances such as probenecid, phenylbutazone, sulfinpyrazone, benzbromarone, and some non-steroidal anti-inflammatory agents inhibits URAT1 from the luminal side.[278] The angiotensin II receptor losartan, which lowers blood uric acid via its uricosuric actions[279] also inhibits URAT1.

Mutations of URAT1 (SLC22A12) cause idiopathic renal hypouricemia. [260] [280] This is a rare autosomal recessive disorder seen in Japanese and Iraqi Jews. The lack of functional URAT1 transporter leads to hypouricemia and hyperuricosuria resulting in crystalluria and kidney stones. Some patients can get exercise-induced acute renal failure from likely a combination of rhabdomyolysis and acute urate nephropathy.[281] Sequencing of SLC22A12 in Japanese cohorts with idiopathic renal hypouricemia revealed two patients who did not have missense mutations in this gene.[280] This suggests that non-coding sequences or additional loci related to urate transport or metabolism could be involved in renal hypouricemia.

AMINO ACIDS

Physiology of Renal Amino Acid Transport

Overview

The amino acid, cystine, was discovered in the urine of a patient suffering from urolithiasis in 1810.[282] We know now that the presence of this amino acid in the urine reflected the failure of this patient to reabsorb cystine properly. In fact, the filtered load of amino acids is comparatively large: with a total concentration of free amino acids in the plasma on the order of 2.5 μM,[283] the result is a daily filtered load at the glomerulus of some 400+ mmoles. Indeed, Cushny recognized in 1917[284] that potent reabsorptive mechanisms must be found in the tubular walls of the nephron to recover amino acids because almost none of the filtered load is actually lost in the urine.

As with the other substrates discussed in this chapter, the powerful techniques of stop flow, micropuncture, and microperfusion identified the renal proximal tubule (RPT) as the principal site of renal amino acid reabsorption.[283]However, although net transepithelial reabsorption typically predominates, there is also a physiologically important influx of many amino acids from the blood into renal cells across the basolateral membrane. The situation is further complicated by tubular amino acid metabolism. Renal glutamine breakdown, for example, plays a key role in acid-base balance by yielding NH3 for urinary acid excretion, and renal conversion of citrulline to arginine is the most important source of this dibasic amino acid in the whole body. [284] [285] Finally, unlike the other transport processes highlighted in this chapter that are generally restricted in their distribution to cells of the proximal tubule, all cells of the renal nephron express an array of distinct amino acid transporters that play important roles in supporting the metabolic needs of the cells. In addition, amino acid transporters distributed in cells of Henle's loop play critical roles in generating large medullary concentrations of amino acid that serve a protective role against the high ionic strength associated the urine concentrating mechanism. [286] [287] [288] [289] These latter process are, however, beyond the scope of the present discussion and the reader is directed to reviews that consider them in detail. [288] [289] The detailed discussion of the tubular and organ physiology of renal amino acid transport, as deduced from classical studies employing intact single renal tubules and perfused organs, is also beyond the scope of the present treatment, and the reader is directed to the discussion of these data by Silbernagl.[283] Here we focus our attention on the molecular and cellular physiology of the multiple amino acid transport processes of the proximal tubule.

Molecular Biology of Amino Acid Transport

Overview

The renal reabsorption of amino acids occurs mainly in the proximal convoluted tubule (S1-S2 segments),[283] and the absorption of these compounds occurs in the small intestine.[290] The plasma membrane of epithelial cells in these two locations has a similar set of amino acid transporters ( Fig. 6-13 ). Trans-epithelial flux of amino acids from the intestinal or renal tubular lumen to the intercellular space requires transport through apical and basolateral plasma membranes. Several amino acid transporters have been identified in the apical domain: (1) for neutral amino acids B0AT1 (system B0), ASCT2 (system ASC), SIT (system Imino), and PAT1 (also representing system Imino; reviewed in Ref 291); (2) for dibasic amino acids, the heterodimer complex rBAT/b0,+AT (system b0,+); and (3) for dicarboxylic amino acids, EAAC1 (system XAG-). Transporters localized in the basolateral domain of these cells are the heterodimers 4F2hc/y+LAT1 (system y+L) and 4F2hc/LAT2 (exchanger L for all neutral amino acids), and TAT1 (SLC16A10; aromatic amino acid (Trp) transporter). Several of these transporters present higher expression in the renal proximal convoluted (S1 and S2 segments) than in the straight tubule (S3 segment): rBAT/b0,+AT,[292] 4F2hc/y+LAT1,[293] and 4F2hc/LAT2, [293] [294] B0AT1, [295] [296] ASCT2,[297] and SIT. [298] [299] PAT1 is expressed in kidney, but its expression pattern along the nephron has not been studied.[300] Transporter ATB0,+ (SLC6A14; system B0,+: Na+ and Cl- dependent co-transporter for neutral and dibasic amino acids), which is not shown in Figure 6-13 , is expressed in distal ileum and colon but not in kidney, indicating a role for this transporter in the absorption of amino acids produced by bacterial metabolism.[301]

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FIGURE 6-13  Proximal tubule model for amino acid transporters involved in renal and intestinal reabsorption of amino acids. Transporters with a proven role in renal reabsorption or intestinal absorption of amino acids are colored, whereas those expressed in the plasma membrane of epithelial cells of the proximal convoluted tubule (or of the small intestine) but with no direct experimental evidence supporting their role in reabsorption, are shown in light blue. Amino acid fluxes in the reabsorption direction are in red. PEPT1 and PEPT2 are expressed in the small intestine and kidney, respectively.

000519

 

Neutral amino acids are mainly absorbed in the small intestine and reabsorbed in the proximal convoluted tubule by system B0. B0AT1 accounts for system B0 activity (electrogenic Na+ co-transport of neutral amino acids) (seeFig. 6-13 ). Two additional B0-like activities are expressed in the proximal straight tubule; the molecular identity of these transporters is unknown (reviewed in Ref 291). Mutations in B0AT1 cause Hartnup disease, characterized by wastage of all neutral amino acids in urine, with the exception of proline, hydroxyproline, glycine, and cystine.[302] This observation suggests that other transporters also mediate the reabsorption of proline. Indeed, renal iminoglycinuria, characterized by aminoaciduria of proline and glycine, also indicates that specific transporters contribute to the reabsorption of these amino acids. PAT1 and SIT are candidate transporters underlining the molecular bases of this disorder. PAT1 is a H+ co-transporter of proline, glycine and alanine[303] whereas SIT is Na+ co-transporter of proline and hydroxyproline (see Fig. 6-13 ). [298] [299] Although PAT1 is proton-dependent, sustained uptake in epithelial cells appears to be Na+-dependent because removal of H+ is coupled to the Na+ -gradient via the Na+/H+ exchanger.[300] Recently, Broer and co-workers[291] proposed a model for renal reabsorption of proline and glycine. PAT1, SIT, and B0AT1 together will reabsorb proline at the convoluted tubule with a capacity exceeding normal kidney load. In contrast, reabsorption of glycine will approach the capacity of two transporters (glycine is not a substrate for SIT): PAT1 and B0AT1. SIT would be the major player for intestinal reabsorption of proline in the small intestine. This model predicts that PAT1 mutations would result in iminoglycinuria: iminoglycinuria without intestinal phenotype may be caused by two mutated alleles in PAT1, whereas one mutated PAT1 allele would lead to isolated glycinuria. In contrast, iminoglycinuria with a defect in intestinal proline transport may be due to mutations in SIT. The possibility that a third gene is involved in renal iminoglycinuria cannot be ruled out. Indeed, the murine Slc6a18-knockout model presents with hyperglycinuria.[304] SLC6A18 codes for the orphan transporter XT2, which is expressed in the proximal straight tubule.[305]

System b0,+ mediates the influx of cystine and dibasic amino acids in exchange with neutral amino acids efflux (see Fig. 6-13 ). The high intracellular concentration of neutral amino acids drives the direction of this exchange. The membrane potential (negative inside) favors the influx of dibasic amino acids (i.e., with a net positive charge at neutral pH) and the intracellular reduction of cystine to cysteine favors the influx of cystine. As a result, patients with cystinuria presents with urinary hyperexcretion of cystine and dibasic amino acids but not other neutral amino acids. Interestingly, the mean and range (i.e., 5th-95th centile limits) of cystine, lysine, arginine, and ornithine in the urine of patients with mutations rBAT and in b0,+AT are almost identical (see patients AA with phenotype I and patients BB with non-I phenotype in Table 6-6 ). This result is expected because all b0,+AT heterodimerizes with rBAT in renal brush-border membranes, constituting the holotransporter b0,+.[292] Cystinuric patients may show almost no cystine reabsorption in kidney, whereas dibasic amino acid reabsorption in this organ remains intact (reviewed in Ref 292). This observation indicates that b0,+ is the main reabsorption system for cystine, but other transporters also participate in the reabsorption of dibasic amino acids. The molecular identity of these transporters is currently unknown.


TABLE 6-6   -- Urine Amino Acid Excretion in Patients Classified by Genotype and Clinical Type of Cystinuria

Genotype

Cystinuria Type

n

Urine Amino Acid Excreted (mmol/g creatinine)

 

 

 

Cystine

Lysine

Arginine

Ornithine

AA

I

34

1.66 [0.65–3.40]

6.58 [2.65–11.6]

3.14 [0.23–8.37]

1.74 [0.59–3.44]

AA

Mixed

3

0.78, 2.12, 5.56

3.31, 5.72, 11.4

1.23, 2.82, 7.03

0.72, 1.64, 1.92

AA(B)

Mixed

1

2.57

9.84

2.95

5.17

BB

I

1

2.69

2.28

1.11

0.30

BB

Non-I

37

1.62 [0.50–3.30]

6.51 [1.72–14.7]

3.45 [0.50–6.15]

2.20 [0.30–4.77]

B+

Non-I carriers

3

0.26[*], 0.44[*], 0.80

1.64[*], 2.45, 3.88

0.02[*], 0.12[*], 0.15[*]

0.04[*], 0.27[*], 0.29[*]

BB

Mixed

11

1.82 [0.43–3.18]

4.58 [1.57–8.72]

1.54 [0.21–3.51]

1.33 [0.47–2.45]

BB(A)

Mixed

1

0.43

3.27

0.489

0.603

Extracted from Font-Llitjos M, Jimenez-Vidal M, Bisceglia L, et al: New insights into cystinuria: 40 new mutations, genotype-phenotype correlation, and digenic inheritance causing partial phenotype. J Med Genet 42:58, 2005.

A, allele SLC3A1 mutated; B, allele SLC7A9 mutated; +, normal allele.

N, number of patients.

The mean of the amino acid levels for each group is indicated, with the exception of categories with less than 11 patients, where individual data points are shown. When applicable, the 5th and 95th percentile limits are in square brackets.

 

*

Excretion values below fifth percentile of homozygotes of cystinuria Type non-I (BB) in carriers of cystinuria Type non-I.

 

 

The intracellular concentration of neutral amino acids is a major determinant of the active uptake of cystine and dibasic amino acids via system b0,+. Apical (e.g., B0AT1) and basolateral (e.g., system A) co-transporters of Na+ and neutral amino acids should contribute to the high intracellular concentration of neutral amino acids (see Fig. 6-13 ). Moderate hyperexcretion of dibasic amino acids occurs in Hartnup disorder,[306] suggesting coordinated function between systems B0 and b0,+: a defective system B0 will reduce the intracellular concentration of neutral amino acids, which drives the influx of dibasic amino acids via system b0,+. In contrast, the impact of system A on renal reabsorption is unknown. The electrochemical gradient of Na+ drives the active transport of the Na+ co-transporters of neutral amino acids B0 and A. Thus, system b0,+ mediates active transport of cystine and dibasic amino acids with a tertiary active mechanism of transport.

Apical PEPT1 (SLC15A1) and PEPT2 (SLC15A1) are expressed in the small intestine and in kidney, respectively.[307] These transporters co-transport H+ with di- and tripeptides. The physiological role of PEPT2 in kidney is largely unknown.[308] The contribution of PEPT1 to the assimilation of amino acids has not been properly evaluated in mammals or humans, but it is assumed that absorption of di- and tri-peptides accounts for a significant proportion of the intestinal absorption of amino acids.[307] A deeper study of the phenotype of the Slc15A2-knockout mouse[308] and generation and study of the PEPT1 model may answer these questions. Meanwhile, the role of PEPT1 in amino acid nutrition is supported by observations of the lack of pathology associated with amino acid malabsorption in cystinuria and in many patients with Hartnup disorder. Patients with cystinuria do not show pathology, with the exception of cystine urolithiasis. It is believed that absorption of di- and tri-peptides via PEPT1 compensate for the defective absorption of cystine and dibasic amino acids via system b0,+. Similarly, phenotype severity in Hartnup disorder is reduced in well-nourished patients.

The heterodimer 4F2hc/y+LAT1 has a basolateral location and accounts for system y+L activity (the electroneutral efflux of dibasic amino acids in exchange with neutral amino acids plus sodium) (see Fig. 6-13 ). Mutations in y+LAT1 cause lysinuric protein intolerance (LPI), which is characterized by hyperdibasic aminoaciduria and malabsorption of dibasic amino acids. On the one hand, wastage of lysine in urine in LPI and cystinuria are similar, whereas that of arginine and ornithine are less severe in LPI than in cystinuria (see Table 6-6 ). Regarding the renal reabsorption of dibasic amino acids, these findings indicate that: (1) lysine appears to be a preferred substrate for basolateral efflux via 4F2hc/y+LAT1, and (2) other basolateral transporters mediate efflux of arginine and ornithine. The molecular identity of these transporters is unknown. On the other hand, LPI produces a larger depletion of the three dibasic amino acids in plasma than cystinuria (see Table 6-6 ). All these observations suggest that malabsorption of dibasic amino acids is more severe in LPI than in cystinuria. Two reasons may account for this: (1) the contribution of the apical peptide transporter PEPT1 cannot compensate for the basolateral defect associated with LPI (see Fig. 6-13 ); and (2) 4F2hc/y+LAT1 is probably the main basolateral system for intestinal absorption of dibasic amino acids.

The basolateral 4F2hc-LAT2 heterodimer is an exchanger with broad specificity for small and large neutral amino acids with characteristics of system L (see Fig. 6-13 ).[294] This transporter may be involved in intestinal absorption and renal reabsorption of neutral amino acids. Indeed, LAT2 knockdown experiments in the polarized opossum kidney cell line OK, derived from proximal tubule epithelial cells, demonstrated that LAT2 participates in the transepithelial flux of cystine, and the basolateral efflux of cysteine and influx of alanine, serine, and threonine.[309] To our knowledge, no inherited human disease has yet been related to LAT2 mutations. Therefore, a final demonstration of the role of LAT2 in reabsorption requires the generation of LAT2-knockout mouse models.

The model proposed in Figure 6-13 for renal reabsorption of amino acids requires a basolateral efflux system for neutral amino acids. A defective amino acid transport system for this efflux would increase the intracellular concentration of these compounds, resulting in their hyperexcretion in urine and intestinal malabsorption. Candidate transporters for this function may be found within transporter families SLC16 and SLC43. Amino acid transporters in these families mediate facilitated diffusion and may therefore mediate the efflux of neutral amino acids from the high intracellular concentration to the interstitial space. T-type amino acid transporter 1 (TAT1; SLC16A10) transports aromatic amino acids in a Na+- and H+-independent manner. [310] [311] TAT1 is expressed in human kidney and small intestine with a basolateral location and can function as a net efflux pathway for aromatic amino acids.[312] Thus, TAT1 may supply parallel exchangers (systems y+L and L) with recycling uptake substrates that could drive the efflux of other amino acids. The SLC16 family (also named MCT for monocarboxylate transporters) holds members that transport monocarboxylates and also thyroid hormones. Several members within this family are orphan transporters.[313] Knockout murine models for TAT1, and their related orphan transporters expressed in kidney cortex and small intestine, may help to identify the basolateral transporters involved in reabsorption of neutral amino acids. LAT3[314] and LAT4[315] within family SLC43 mediate the facilitated diffusion of neutral amino acids with characteristics of system L. Neither of these two transporters is expressed in epithelial cells of the renal proximal convoluted tubule or the small intestine. Interestingly, the SLC43 family has a third member with no identified transport function (EEG1[316]). Functional and tissue-expression studies are required to ascertain the role of EEG1 in the reabsorption of amino acids.

The bulk (>90%) of filtered acidic amino acids is reabsorbed within segment S1 (i.e., the first part of the proximal convoluted tubule). [317] [318] Two apical acidic transport systems have been described in the proximal tubule: one of high capacity and low affinity and the other of low capacity and high affinity.[319] The Na+/K+-dependent acidic amino acid transporter EAAC1 (also named EAAT3), which localized to chromosome 9p24[320] (system XAG-) is expressed mainly in the brush-border membranes of segments S2 and S3 of the nephron (see Fig. 6-13 ).[321] The transport character-istics of SLC1A1 correspond to the high-affinity system.[322] The Slc1a1-knockout mouse develops dicarboxylic aminoaciduria,[323] demonstrating the role of this transporter in renal reabsorption of dibarboxylic amino acids. Mutational analysis of SLC1A1 in patients with dicarboxylic aminoaciduria has not been performed. The apical low-affinity transport system for acidic amino acids in kidney has been characterized in brush-border membrane preparations,[324] but its molecular entity remains elusive. At renal basolateral plasma membranes, a high-affinity Na+/K+-dependent transport system for acidic amino acids has been reported,[325] but its molecular structure has not been identified. GLT1 (i.e., the glial high-affinity glutamate transporter, [326] [327] also named EAAT2; SLC1A2) may be responsible for this activity. GLT1 mRNA is expressed in rat kidney cortex and porcine small intestine [328] [329] but the expression of GLT1 protein has not been studied in kidney or intestine. Slc1a2-knockout mice show lethal spontaneous epileptic seizures[330] but the renal phenotype in these mice has not been examined.

Inherited Aminoacidurias in Humans

Overview

Primary inherited aminoacidurias (PIA) are caused by defective amino acid transport, which affect renal reabsorption of these compounds and may also affect intestinal absorption as well. Several PIA have been described ( Table 6-7). Inherited disorders of renal tubule like the renal Fanconi syndrome (MIM: 134600), which is a generalized dysfunction of the proximal tubule that results in wasting of phosphate, glucose, amino acid and bicarbonate, or cystinosis (MIM: 219800; 219900), affecting lysosomal efflux of cystine are not discussed in this chapter. Neither are the inherited defects of amino acid metabolism resulting in aminoaciduria (e.g., homocystinuria, MIM 236200) described in this chapter.


TABLE 6-7   -- Primary Inherited Aminoacidurias

 

Prevalence

Inheritance

Gene

Chromosome

Mutations

Transport System

Cystinuria[*]

1:7000

AR/ADIP

SLC3A1

2p16.3

112

 

 

 

 

SLC7A9

19q13.1

73

b0,+

Isolated cystinuria

Very rare

AR?

?

?

?

?

Lysine Protein Intolerance

∼200 cases

AR

SLC7A7

14q11

26

y+L

Hyperdibasic aminoaciduria Type 1

Very rare

AD

?

?

?

?

Isolated lysinuria

Very rare

AR?

?

?

?

?

Hartnup disorder

1:26000

AR

SLC6A19

5p15

10

B0

Renal familial iminoglycinuria

1 : 15000

AR

?

?

?

Imino(?)[†]

Dicarboxylic amino aciduria

Very rare

AR?

SLC1A1 (?)

9p24

KO null[‡]

XAG-

 

AR, autosomal recessive; ADIP, autosomal dominant with incomplete penetrance; AD, autosomal dominant; AR?, familial studies in the very few cases described for these diseases suggest an autosomal recessive mode of inheritance.

 

*

Three phenotypes of cystinuria, depending on the obligate heterozygotes, are considered: Type I (with AR inheritance), Type non-I (ADIP inheritance), and Mixed Type (combination of both).

The amino acids hyperexcreted in patients with renal familial iminoglycinuria (glycine and proline) suggest defects in Imino system.

Slc1a1-null knockout mice present dicarboxylic aminoaciduria, pointing to this gene as a candidate for the human disease.

 

Plasma membrane transport of dibasic amino acids (i.e., basic amino acids) is abnormal in four inherited diseases:

  

1.   

Cystinuria (MIM 220100; 600918), in which patients present hyperexcretion of cystine and dibasic amino acids (first described by Sir Archibald Garrod in 1908).[331] There is phenotypic variability in obligate heterozygotes (i.e., silent or hyperexcretors of amino acids).[332]

  

2.   

Lysinuric protein intolerance (LPI) (also named hyperdibasic aminoaciduria type 2, or familial protein intolerance; MIM 222700) (first described in Finland).[333]

  

3.   

Autosomal dominant hyperdibasic aminoaciduria type I (MIM 222690).[334]

  

4.   

Isolated lysinuria described in one Japanese patient.[335]

Cystinuria and LPI are caused by defective amino acid transporter systems b0,+ and y+L respectively. These two transporters belong to the family of heteromeric amino acid transporters (HAT). [336] [337] Mutations in the two subunits of system b0,+ (rBAT and b0,+AT) causes cystinuria. [338] [339] whereas mutations in one of the two subunits of system y+L (y+LAT1), but not in the other subunit (4F2hc), produce LPI. [340] [341] At the molecular level, the relationship between LPI, and the very rare autosomal dominant hyperdibasic aminoaciduria type I, and isolated lysinuria is unknown.

Plasma membrane transport of zwitterionic amino acids (i.e., neutral amino acids at physiological pH) is defective in three inherited diseases:

  

1.   

Hartnup disorder (MIM 234500), in which patients present hyperexcretion of neutral amino acids (first described in two siblings of the Hartnup family).[342]

  

2.   

Renal familial iminoglycinuria (MIM 242600) is an autosomal recessive benign disorder in which individuals present hyperexcretion of proline and glycine (first described in the sixties). [343] [344] There is phenotypic complexity in this disorder [345] [346]; (1) renal iminoglycinuria with defective intestinal absorption and normal heterozygotes; (2) renal iminoglycinuria without intestinal phenotype and normal heterozygotes; and (3) renal iminoglycinuria without intestinal phenotype and isolated glycinuria in heterozygotes.

  

3.   

Isolated cystinuria (MIM 238200), in which patients present hyperexcretion of cystine but not dibasic amino acids.[347]

Hartnup disorder is due to a defective amino acid transport system B0 (also named neutral brush border) caused by mutations in B0AT1 (SLC6A19). [296] [348] The relationship between isolated cystinuria and cystinuria at the molecular level is unknown. The molecular basis of iminoglycinuria is unknown but candidate genes are[291]SLC36A1 (coding for transporter PAT1),[300] SLC6A20 (coding for transporter SIT [also called IMINO]), [298] [299] and SLC6A18 (coding for the orphan transporter XT2).

Plasma membrane transport of dicarboxylic amino acids is defective in Dicarboxylic aminoaciduria (MIM 222730). [349] [350] The molecular basis of this disease is unknown but the glutamate transporter EAAC1 (SLC1A1[351]) is an obvious candidate because the murine knockout of Slc1a1 presents dicarboxylic aminoaciduria.[323]

Defects Associated with Heteromeric Amino Acid Transporters

Heteromeric amino acid transporters (HATs) are composed of a heavy subunit and a light subunit (see Table 6-2 ). [337] [352] [353] These are unique features among mammalian plasma membrane amino acid transporters. Two homologous heavy subunits from the SLC3 family have been cloned, rBAT (i.e., related to b0,+ amino acid transport) and 4F2hc (i.e., heavy chain of the surface antigen 4F2hc, also named CD98 or fusion regulatory protein 1 [FRP1]).[354] Ten light subunits (SLC7 family members from SLC7A5 to SLC7A14) have been identified. Six of these are partners of 4F2hc (LAT1, LAT2, y+LAT1, y+LAT2, asc1, and xCT); one forms a heterodimer with rBAT (b0,+AT); two (asc2 and AGT-1) appear to interact with as yet unknown heavy subunits [355] [356]; and the last one (arpAT) may interact with rBAT, 4F2hc, or an unidentified heavy subunit.[357] Two light subunits are not present in humans: asc2 is not found in the genome sequence and arpAT is heavily inactivated in this genome.[357] Members SLC7A1-4 of family SLC7 correspond to system y+ isoforms (i.e., cationic amino acid transporters; CATs) and related proteins, which on average show <25% amino acid identity with the light subunits of HATs.

The general features of HATs are as follows [352] [353]:

  

1.   

The heavy subunits (molecular mass of ≈90 and ≈80 kDa for rBAT and 4F2hc, respectively) are type II membrane N-glycoproteins with a single transmembrane domain, an intracellular N-terminus, and an extracellular C-terminus significantly homologous to insect and bacterial glucosidases ( Fig. 6-14 ). X-ray diffraction of the extracellular domain of human 4F2hc revealed a three-dimensional structure similar to that of bacterial glucosidases (a triose phosphate isomerase (TIM) barrel [(ab)8] and eight antiparallel β-strands); Fig. 6-14 (unpublished results).

  

2.   

The light subunits (≈50 kDa) are highly hydrophobic and not glycosylated. This results in anomalously high mobility in SDS-PAGE (35 kDa-40 kDa). Cysteine-scanning mutagenesis studies of xCT, as a model for the light subunits of HATs, support a 12-transmembrane-domain topology, with the N- and C- terminals located inside the cell and with a reentrant-like structure in the intracellular loop IL2-3 (see Fig. 6-14 ).[358]

  

3.   

The light and the corresponding heavy subunit are linked by a disulfide bridge (see Fig. 6-14 ). For this reason, HATs are also named glycoprotein-associated amino acid transporters. The intervening cysteine residues are located in the putative extracellular loop EL3-4 of the light subunit and a few residues away from the transmembrane domain of the heavy subunit (see Fig. 6-14 ).

  

4.   

The light subunit cannot reach the plasma membrane unless it interacts with the heavy subunit.

  

5.   

The light subunit confers specific amino acid transport activity to the heteromeric complex (LAT1 and LAT2 for system L isoforms, y+LAT1 and y+LAT2 for system y+L isoforms, asc1 and asc2 for system asc isoforms, xCT for system xc-, b0,+AT for system b0,+, AGT-1 for a system serving aspartate and glutamate transport, and arpAT for a transport system with aromatic amino acids as preferred substrates). Moreover, reconstitution in liposomes showed that the light subunit b0,+AT is fully functional in the absence of the heavy subunit rBAT.[382]

  

6.   

The light subunit b0,+AT stabilizes the heavy subunit rBAT. No data are available as to whether this also holds for 4F2hc and associated light subunits.

  

7.   

HAT are, with the exception of system asc isoforms, tightly coupled amino acid antiporters.[359]

 

000336

000519

FIGURE 6-14  Schematic representation of a heteromeric amino acid transporter. The heavy subunit (blue) and the light subunit (brown) are linked by a disulfide bridge with conserved cysteine residues (cysteine 158 for the human xCT and cysteine 109 for human 4F2hc). The heavy subunits (4F2hc or rBAT) are type II membrane glycoproteins with an intracellular NH2 terminus, a single transmembrane domain, and a bulky COOH terminal domain (≈50 kDa without glycosylation; i.e., similar to the size of the light subunits). This part of the protein shows homology with bacterial glycosidases (a schematic representation of the TIM barrel and the all-β domain is shown). The light subunits are polytopic proteins with 12 transmembrane domains, with the NH2 and COOH terminals located intracellularly and with a reentrant loop-like structure in the intracellular loop IL2-3 (on the basis of studies on xCT[358]).

000519

 

The transport characteristics of two HAT-associated transport systems are relevant to cystinuria and LPI (see Table 6-6 ): system b0,+ [due to the rBAT (SLC3A1) and b0,+AT (SLC7A9) heterodimer] is a tertiary active mechanism of renal reabsorption and intestinal absorption of dibasic amino acid and cystine in the apical plasma membrane. It mediates the electrogenic exchange of dibasic amino acids (influx) for neutral amino acids (efflux) (see Fig. 6-13 ). System y+L (4F2hc/y+LAT1 heterodimer) mediates the electroneutral exchange of dibasic amino acids (efflux) for neutral amino acids plus sodium (influx). [361] [362] [363] This transport system allows the efflux of dibasic amino acids against the membrane potential at the basolateral domain of epithelial cells (see Fig. 6-13 ). The 4F2hc/y+LAT2 heterodimer also mediates system y+L in many other cell types (see Ref 353 for review).

Cystinuria: Overview

Cystinuria is an autosomal inherited disorder, characterized by impaired transport of cystine and dibasic amino acids in the proximal renal tubule and the gastrointestinal tract. The overall prevalence of the disease is 1 in 7000 neonates, ranging from 1 in 2500 neonates in Libyan Jews to 1 in 100,000 among Swedes. Patients present with normal to low-normal levels in blood ( Table 6-8 ), hyperexcretion in urine, and intestinal malabsorption of these amino acids. High cystine concentration in the urinary tract most often causes the formation of recurring cystine stones as a result of the low solubility of this amino acid. This is the only symptom associated with the disease. Therefore, treatment attempts to increase cystine solubility in urine (increased hydration, urine alkalinization, and formation of soluble cystine adducts with thiol drugs). Cystinuria is not accompanied by malnutrition, suggesting that intestinal malabsorption is not severe. The transport defect occurs in the apical plasma membrane of renal and intestinal epithelial cells. Absorption of di- and tripeptides via PEPT1 (SLC15A1) may prevent malnutrition in cystinuria (see Fig. 6-13 ).[307]


TABLE 6-8   -- Plasma and Urine Amino Acids in Lysine Protein Intolerance and Cystinuria

Plasma Amino Acids (μM)

Amino Acid

Range in Normal Children

Patients with LPI

Controls Cystinuria (mean ± SD)

Patients with Cystinuria (mean ± SD)

Mean

Range

Lysine

71–151

70

32–179

171 ± 26

121 ± 30

Arginine

23–86

27

12–58

82 ± 16

46 ± 12

Ornithine

27–86

21

2–83

58 ± 11

36 ± 11

Cystine

48–140

80

57–105

79 ± 12

43 ± 12

Glutamine

57–467

5583

3644–7161

n.d.

n.d.

Alanine

173–305

772

417–1017

n.d.

n.d.

 

Amino Acids in Urine

Amino Acid

Range in Controls (mmol/g creatinine)

Patients with LPI (mmol/1.73 m2/24 h)

Patients with Cystinuria Type B (mmol/g creatinine)

Mean

Range[*]

Mean

Range

Mean

Range[*]

Lysine

0.18

0.04–0.50

4.13

1.02–7.00

6.51

1.72–14.7

Arginine

0.02

0.00–0.05

0.36

0.08–0.69

3.45

0.50–6.15

Ornithine

0.03

0.01–0.07

0.11

0.09–0.13

2.20

0.30–4.77

Cystine

0.05

0.02–0.11

0.12

0.06–0.21

1.62

0.50–3.30

 

Plasma amino acids are expressed in μM. Data for normal children, and patients with LPI (n = 20) are from (Simell, 2001). Plasma glutamine data also include asparagine concentration. Plasma amino acids from controls (n = 12) and patients with cystinuria (n = 8) are from Morin CL, Thompson MW, Jackson SH, et al: Biochemical and genetic studies in cystinuria: Observations on double heterozygotes of genotype I-II. J Clin Invest 50:1961, 1971. In patients with LPI, urinary excretion is expressed in mmol/1.73 m2/24 h (n = 4) (Simell, 2001), whereas in controls (n = 83) (Dello SL, Pras E, Pontesilli C, et al: Comparison between SLC3A1 and SLC7A9 cystinuria patients and carriers: A need for a new classification. J Am Soc Nephrol 13:2547, 2002) and patients with cystinuria type B (i.e., due to mutations in SLC7A9) (n = 37) (Font-Llitjos M, Jimenez-Vidal M, Bisceglia L, et al: New insights into cystinuria: 40 new mutations, genotype-phenotype correlation, and digenic inheritance causing partial phenotype. J Med Genet 42:58, 2005.) it is expressed in mmol/g creatinine.

SD, standard deviation. n.d., not determined.

 

*

Fifth-95th centile range.

 

Traditionally, three types of cystinuria have been recognized in humans: type I, type II, and type III.[364] This classification correlates poorly with molecular findings, and it has recently been revised to type I (MIM 220100) and non-type I (MIM 600918) cystinuria (with the latter corresponding to old types II and III). These two are distinguished on the basis of the cystine and dibasic aminoaciduria of the obligate heterozygotes[332]: type I heterozygotes are silent, whereas non-type I heterozygotes present a variable degree of urinary hyperexcretion of cystine and dibasic amino acids that is higher in type II than in type III. This indicates that type I cystinuria is transmitted as an autosomal recessive trait, whereas non-type I is transmitted dominantly, with incomplete penetrance. Not surprisingly, urolithiasis has been reported in a minority of non-type I heterozygotes. Thus, in the cohort of patients of the International Cystinuria Consortium (ICC) three type non-I heterozygotes with cystine urolithiasis have been identified from 164 cystinuria probands ( Table 6-9 ).[365] Patients with a mixed type, inheriting type I and non-type I alleles from either parent, have also been described.[366] Data on the relative proportion of the two types in specific populations are scarce. In 97 well-characterized families of the ICC cohort of patients, mainly from Italy, Spain, and Israel, 38%, 47%, and 14% transmitted type I, non-type I, and mixed cystinuria, respectively (see Table 6-9 ).[365] This cohort is not a registry, and therefore it may not be representative of the whole population within these countries.[367]


TABLE 6-9   -- Cystinuria Type and Genetic Frequencies of the Probands from the International Cystinuria Consortium

Cystinuria Phenotype

Genotype

I

Non-I

Non-I Carriers[*]

Mixed

Untyped

Total Probands (%)

 

AA

29

 

 

2

25

56 (34.1)

 

AA(B)

 

 

 

1

 

1 (0.6)

 

BB

1

34

 

7

23

65 (39.6)

 

B+

 

 

3

 

 

3 (1.8)

 

BB(A)

 

 

 

1

 

1 (0.6)

126 (76.8)

A?

5

 

 

1

5

11 (6.7)

 

B?

2

7

 

2

11

22 (13.3)

33 (20.1)

??

 

2

 

 

3

5 (3.0)

5 (3.0)

Total probands (%)

37 (22.6)

43 (26.2)

3 (1.8)

14 (8.5)

67 (40.9)

164 (100)

 

Total alleles

74

89

3

28

134

 

 

Explained alleles (%)

67 (90.5)

78 (87.6)

3 (100)

25 (89.3)

112 (83.6)

 

 

Extracted from Font-Llitjos M, Jimenez-Vidal M, Bisceglia L, et al: New insights into cystinuria: 40 new mutations, genotype-phenotype correlation, and digenic inheritance causing partial phenotype. J Med Genet 42:58, 2005.

A, allele SLC3A1 mutated; B, allele SLC7A9 mutated; +, normal allele; ?, unknown allele.

A total of 164 probands have been studied. In 126 probands (76.8%) the alleles causing the disease have been identified. In 33 probands (20.1%) one of the two mutated alleles has been identified. In five probands (3.0%) no mutated alleles have been identified.

 

*

Heterozygote probands with cystine lithiasis. For patients AA(B) and BB(A), two alleles causing the disease and two explained alleles in each case have been taken into account in the calculations.

 

 

The characteristics of rBAT expression in the plasma membrane of the epithelial cells of kidney proximal tubule and small intestine, and induction of system b0,+ in oocytes (for review see Ref 352) pointed to this gene as a candidate for cystinuria. In 1994 it was demonstrated that mutations in SLC3A1 cause type I cystinuria.[338] Since then 112 distinct rBAT mutations have been described, including nonsense, missense, splice-site, and frameshift mutations, as well as large deletions and chromosome rearrangements (mutations are listed in Ref 365; for more recently described mutations, see Refs 368-370). Cystinuria resembling type I caused by mutations in canine Slc3a1 has been reported in Newfoundland dogs.[371] Similarly, Pebbels mice (homozygous for the rBAT mutation D140G) develop type I cystinuria with urolithiasis.[372]

Most of the cystinuria-associated rBAT missense mutations occur in the ectodomain.[365] Unfortunately, amino acid sequence identity of the ectodomains of rBAT and 4F2hc is very low. This precludes the generation of a structural model of the ectodomain of rBAT using the crystal structure of that of 4F2hc. Therefore, at present, functional analysis is the easiest study available for missense rBAT mutations. The most common SLC3A1 mutation, M467T, showed a trafficking defect, with the protein reaching the plasma membrane inefficiently.[373] A trafficking defect, also suggested for other SLC3A1 mutations, [359] [373] [374] is consistent with the proposed role of rBAT as an ancillary subunit of b0,+AT. SLC3A1 mutations may also affect transport properties of the holotransporter b0,+: the cystinuria-specific mutation R365W, in addition to temperature-sensitive protein stability and trafficking defect, shows a defect in the efflux of arginine but not in its influx.[359] This observation indicates two pathways for the transport unit of system b0,+, one for influx and the other for efflux. This scenario is consistent with two additional sets of results: (1) the intestinal system b0,+ of the chicken has a sequential mechanism of exchange, compatible with the formation of a ternary complex (i.e., the transporter bound to its intracellular and extracellular amino acid substrates)[375]; and (2) the analog amino isobutyrate (AIB) induces an unequal exchange with other substrates through the rBAT-induced system b0,+ in oocytes (i.e., using the endogenous b0,+AT subunit). The oligomeric structure of system b0,+ (i.e., rBAT-b0,+AT heteromer) is unknown. Functional coordination of two rBAT-b0,+AT heterodimers in a heterotetrameric structure would explain these results. If this is not the case, the transport defect associated with mutation R365W would suggest that a single b0,+AT subunit contains two translocation pathways.

The gene causing non-type I cystinuria was assigned to 19q12-13.1 by linkage analysis. [376] [377] [378] SLC7A9 was a positional and functional candidate gene for non-type I cystinuria (i.e., appropriate chromosomal location, rBAT-associated amino acid transport activity (system b0,+), and proper tissue ex-pression in kidney and small intestine (reviewed in Ref 379). In 1999 the non-type I cystinuria gene was identified as SLC7A9.[339] The protein product encoded by SLC7A9 was termed b0,+AT for b0,+ amino acid transporter. Seventy-three SLC7A9 mutations causing cystinuria have been described (mutations are listed in Ref 365; for more recently described mutations see Refs 368-370). Mutation G105R is the most frequent SLC7A9 mutation in the ICC cohort of patients (≈27% of the SLC7A9 alleles identified). Similarly to the human disease, the Slc7a9-knockout mouse presents with non-type I cystinuria with urolithiasis.[380] Several cystinuria-specific SLC7A9 missense mutations have been reported to lead to a defect in transport function.[381] Reconstitution in proteoliposomes showed that A182T-mutated b0,+AT is active, whereas mutation A354T renders the transporter inactive.[382]

Very recently, the ICC performed an exhaustive mutational analysis of 164 probands[365]: ≈87% of the independent alleles were identified. The coverage of identified alleles was similar in all cystinuria types (see Table 6-9 ). The unidentified alleles (≈13%) may be due to mutations in intronic or promoter regions, to SLC3A1 or SLC7A9 polymorphisms in combination with cystinuria-specific mutations in the other allele,[383] or to unidentified genes. These three possibilities have not been confirmed or ruled out. Of particular interest is the possibility of a third cystinuria gene. In this regard, Goodyer's group proposed SLC7A10 as a candidate.[384] This gene is located near the cystinuria gene SLC7A9 on chromosome 19q13.1 and codes for asc1, a 4F2hc-associated renal light subunit with substrate specificity for cysteine and other small neutral amino acids. Moreover, these authors identified the missense mutation E112D associated with cystinuria. In contrast, recent studies have ruled out this hypothesis[359]: (1) cystinuria-specific mutations are not found in patients with alleles not explained by mutations in the two cystinuria genes; (2) the conservative mutation E112D does not affect transport of 4F2hc-asc1; and (3) asc1 mRNA is expressed in the distal tubule where renal reabsorption of amino acids is not relevant but where asc1 may have a role in osmoregulation. The possibility of a third cystinuria gene cannot be discarded, but it would be relegated to a very small proportion of patients (only 3% of the probands of the ICC show no mutation in either of the two cystinuria genes) (see Table 6-9).

Genotype/Phenotype Correlations in Cystinuria

Initial data suggested a close correlation between the phenotype and the mutated gene (mutations in SLC3A1 resulted in type I, and mutations in SLC7A9 resulted in non-type I). [385] [386] In contrast to this simple view, recent data show a more complex scenario. On the one hand, all SLC3A1 mutations in well-characterized families cause type I cystinuria, with the exception of mutation dupE5–E9, which shows the non-type I phenotype in four of six heterozygotes studied.[365] This mutation consists of a gene rearrangement c.(891+1524_1618-1600)dup, which results in the duplication of exons 5-9 and the corresponding in-frame duplication of amino acid residues E298–D539 of rBAT, as shown by RNA studies. [365] [387] Functional studies are required to explain the dominant negative effect of dup E5–E9 mutation on the rBAT/b0,+AT heteromeric complex. On the other hand, most of the heterozygotes carrying a SLC7A9 mutation have a phenotype of non-I (i.e., hyperexcretion of dibasic amino acids and cystine), but may also have a phenotype of I (i.e., silent heterozygotes). Approximately 14% of the SLC7A9 heterozygotes have phenotype I.[367] SLC7A9 mutations associated with phenotype I in some families are I44T, G63R, G105R, T123M, A126T, V170M (the Libyan Jewish mutation), A182T, G195R, Y232C, P261L, W69X, and c.614dupA. [365] [388]There is no clear explanation of why these mutations associate with phenotype I because some proteins show residual transport activity when expressed in heterologous expression systems whereas others do not. A182T is the most frequent SLC7A9 mutation associated with phenotype I (i.e., 6 of 11 A182T heterozygotes in the ICC cohort), and this mutation leads to a protein with 50% residual transport activity at the plasma membrane. Moreover, in the ICC cohort the 11 mixed cystinuria patients and the single patient with cystinuria type I, which all carry two mutations in SLC7A9 presented aminoaciduria in the lower range of non-type I patients with two mutations in this gene.[365]This suggests that, in addition to individual and population variability, mild SLC7A9 mutations may be more prone to associate with silent phenotype in heterozygotes (i.e., phenotype I).

The lack of a direct relationship between the mutated cystinuria gene and the type of cystinuria led the ICC to propose a parallel classification to describe cystinuria on the basis of the genotype of the patients (type A due to mutations in SLC3A1, type B due to mutations in SLC7A9, and type AB to define a possible digenic cystinuria).[367] Table 6-8 summarizes the double classification for 78 cystinuria probands by the ICC as follows: (1) most type I patients have two mutations in SLC3A1 (i.e., individuals AA); (2) all non-type I patients (including type non-type I heterozygotes with urolithiasis) have mutations in SLC7A9 (i.e., individuals BB and B+); (3) patients with mixed cystinuria carry mutations in SLC3A1 (2 probands AA) or in SLC7A9 (7 probands BB); and (4) 2 of 126 fully genotyped probands carry mutations in both genes.

To our knowledge, only four patients with mutations in both cystinuria genes have been described. [365] [389] There is no report of the urine phenotype of the Swedish patient AA(B). Two sisters AA(B) and one male BB(A) from 2 families of 126 fully genotyped families in the ICC cohort have been identified and classified as mixed cystinuria patients (i.e., each of the two mutated alleles in the same gene is associated with phenotype I or non-phenotype I in the obligate heterozygotes). The aminoaciduria levels of these patients and their double-heterozygote (i.e., AB) relatives indicate that digenic inheritance in cystinuria has only a partial effect on the phenotype, restricted to a variable impact on the aminoaciduria. Indeed, none of the individuals AB presented urolithiasis. Given that the frequencies of type A and B alleles are similar in this cohort, if digenic inheritance was the rule in cystinuria, we would expect a quarter of patients to be AA, a quarter to be BB, and half to be AB. This indicates that digenic inheritance affecting phenotype is an exception in cystinuria. However, the possibility that some combinations of mutations A and B produce enough cystine hyperexcretion to cause urolithiasis cannot be ruled out.

A working hypothesis on the biogenesis of the rBAT/b0,+AT heterodimer may explain the urine phenotypes and the ap-parent lack of full digenic inheritance in cystinuria: b0,+AT controls the amount of active holotransporter at the plasma membrane. Thus, the rBAT protein would be produced in excess in kidney, and therefore an rBAT mutation in heterozygosis in humans and in mice does not lead to hyperexcretion of amino acids (phenotype I). The only exception to this rule that has been identified to date is the human rBAT mutation dupE5–E9, thereby indicating a dominant effect for this mutation. b0,+AT controls the expression of the functional rBAT/b0,+AT heterodimeric complex: interaction with b0,+AT stabilizes rBAT, and the excess of rBAT is degraded, as shown in transfected cells. [382] [390] As a result, a half dose of b0,+AT (i.e., heterozygotes of severe human SLC7A9 mutations or of the Slc7a9-knockout mice) causes hyperexcretion of cystine and dibasic amino acids (i.e., phenotype non-I) as a result of a significant decrease in the expression of functional rBAT/b0,+AT (system b0,+). In this scenario, the lack of a full cystinuria phenotype because of digenic inheritance indicates that in double heterozygotes (AB), the mutated rBAT does not compromise the heterodimerization and trafficking to the plasma membrane of the half dose of wild-type b0,+AT with the half dose of wild-type rBAT. Thus individuals AB behave as heterozygotes B with a variable degree of aminoaciduria, which could be greater than that of single heterozygotes within the family, depending on the particular combination of mutations. Demonstration of this hypothesis requires an in-depth study of the effect of cystinuria-specific rBAT and b0,+AT mutations on the biogenesis of the heteromeric complex rBAT/b0,+AT both in cell culture studies and in vivo, using double heterozygote mice (Slc3a1 D140G/+Slc7a9 -/-).

Urolithiasis shows a clear gender and individual variability among cystinuria patients.[367] In the ICC cohort, the age of onset of lithiasis ranges from 2 to 40 years with a median of 12 and 15 years for males and females, respectively. Similarly, the number of total stone events (i.e., spontaneously emitted stones plus those surgically removed) is higher in males than females (0.42 and 0.21 events per year in males and females, respectively). Of the 224 patients studied, ten with full genetic confirmation of the disease and presenting aminoaciduria did not develop renal stones, and two of these patients were over 40 years of age. In contrast, clinical symptoms (i.e., urolithiasis and its consequences) are almost identically represented in the two cystinuria types when either the clinical or the genetic classification is considered. The differences in severity between the genders and marked differences between siblings sharing the same mutations[367] suggest that other lithogenic factors, genetic or environmental, contribute to the urolithiasis phenotype. Indeed, only about half of the Slc7a9-knockout mice, in a mixed genetic background, develop urolithiasis.[380] Moreover, lithiasic and non lithiasic Slc7a9-knockout mice hyperexcrete similar levels of cystine. Studies in Slc7a9-knockout mice with distinct genetic backgrounds may unravel the genetic factors, in addition to mutations in Slc7a9 and the cystine levels in urine, which contribute to urolithiasis.

Lysinuric Protein Intolerance: Overview

Lysinuric protein intolerance is a primary inherited aminoaciduria with an autosomal recessive mode of inheritance predominantly reported in Finland where the prevalence of the disorder is 1 in 60,000. Two other geographic locations with a relatively high prevalence are Southern Italy and Japan,[332] with the northern part of Iwate (Japan) registering a prevalence of 1 in 50,000.[391] The diagnosis of LPI is often difficult because of an unspecific clinical presentation. Therefore it is not surprising that LPI is mainly known in Finland, Italy, and Japan (≈200 patients described) where clinicians are accustomed to diagnose this disorder.[332]

In LPI there is massive urinary excretion of dibasic amino acids, especially lysine, and the intestinal absorption of these amino acids is poor; therefore the concentration of dibasic amino acids in plasma is low (see Table 6-8 ). [392] [393] Arginine and ornithine are intermediates of the urea cycle that provide the carbon skeleton to the cycle. Their reduced availability results in a functional deficiency of the urea cycle.[394] Other characteristics of the LPI phenotype are as follows. Protein malnutrition and deficiency of the essential amino acid lysine contribute to the patient's failure to thrive. Patients with LPI are usually asymptomatic while breast-feeding, and symptoms (e.g., vomiting, diarrhea, and hyperammonemic coma when force-fed high-protein food) appear only after weaning. After infancy, patients with LPI reject high-protein diets, and show a delay in bone growth and prominent osteoporosis, hepatosplenomegaly, muscle hypotonia, and sparse hair. Most patients have a normal mental development, but some may show moderate retardation. Low-protein diet and citrulline, a urea cycle intermediate, are used to correct the functional deficiency of intermediates of the urea cycle. The final height in treated patients is slightly subnormal or low normal. This treatment does not correct all symptoms such as poor growth, hepatosplenomegaly, delayed bone age, and osteoporosis, which are all probably due to the lysine deficiency. Recently, Simell's group reported recovery of plasma lysine by oral supplementation with the amino acid.[395]

About two thirds of patients with LPI have interstitial changes in chest radiographs, and some develop acute or chronic respiratory insufficiency[396] that can lead to fatal pulmonary alveolar proteinosis and to multiple organ dysfunction syndrome. Further symptoms suggesting that the immune system is affected are glomerulonephritis and erythroblastophagia. [397] [398]

System y+L transports dibasic amino acids with high affinity (Km in the micromolar range) in a sodium-independent way, but requires sodium to transport neutral amino acids with high affinity[399]: (1) in the absence of sodium, the transport of neutral amino acids through system y+L is of very low affinity; and (2) system y+L catalyzes the electroneutral efflux of cationic amino acids in exchange for neutral amino acids plus sodium, using the driving force of the sodium concentration gradient. In the early 1990s, two groups described the expression of a system y+-like transport activity in Xenopus oocytes after injection of 4F2hc cRNA. [400] [401] Two closely related proteins (y+LAT-1 and y+LAT-2) that induce y+L transport activity when expressed together with 4F2hc were identified by homology screening,[402] using the light subunit of HAT LAT-1. [403] [404] The transport characteristics of 4F2hc/y+LAT-1 have been studied in heterologous expression systems, where co-immunoprecipitation of these two proteins has been substantiated. [363] [402] [405] The transport activity elicited matches the characteristics of system y+L; electroneutral exchange of dibasic amino acids for neutral amino acids plus sodium with a 1:1:1 stoichiometry. y+LAT-1 is expressed in the basolateral plasma membrane in the epithelial cells of kidney tubules and polarized cellular models (see Fig. 6-13 ).[293]

The gene responsible for LPI was localized to 14q11.2 in Finnish and non-Finnish populations. [406] [407] The cloning of y+LAT-1, encoded by SLC7A7, revealed characteristics that made this gene an excellent candidate for LPI (i.e., appropriate chromosome location, co-expression of system y+L with 4F2hc, and proper expression in LPI affected tissues).

In 1999 two consortiums [340] [341] independently reported the first mutational analysis of SLC7A7 in patients with LPI. A single Finnish mutant allele (1181-2A>T) was found with an A>T transversion at position -2 of the acceptor splice site in intron 6 of SLC7A7. This inactivates the normal splice site acceptor and activates a cryptic acceptor 10 bp downstream with the result that 10 bp of the ORF are deleted and the reading frame is shifted. This mutation has been found in all Finnish LPI patients (i.e., “the Finnish mutation”).[408] These two seminal studies also identified LPI-specific SLC7A7 mutations in Spanish and Italian patients, and established that mutations in SLC7A7 cause LPI. The fact that system y+L activity is present in LPI erythrocytes or fibroblasts [409] [410] indicates the expression of a distinct y+L transporter isoform in these cells, most probably y+LAT-2. Additional studies showed the nonsense mutation W242X and the insertion 1625insATAC as the most prevalent mutations in the south of Italy[411] and the nonsense mutation R410X as the most prevalent in Japan. A total of 26 SLC7A7 mutations of any kind (large genomic rearrangements, missense and nonsense mutations, splicing mutations, insertions and deletions) has been described in 106 patients with LPI (>90% allele explained).[412] No LPI-associated mutations have been reported in SLC3A2, coding for the heavy subunit of y+LAT-1 (4F2hc). This strongly suggests that SLC7A7 is the only gene involved in the primary cause of LPI. It is believed that mutations in SLC3A2 would be deleterious. 4F2hc serves as the heavy subunit of six other HAT (see earlier). Therefore, a defect in 4F2hc will result in six defective amino acid transport activities expressed in many cell types and tissues. Indeed, the murine Slc3a2 knockout is lethal.[413]

Functional studies in oocytes and transfected cells showed that frameshift mutations (e.g., 1291delCTTT, 1548delC, and the Finnish mutation) produce a severe trafficking defect (e.g., the mutated proteins do not localize to the plasma membrane when co-expressed with 4F2hc). [408] [414] In contrast, the missense mutations G54V and L334R inactivate the transporter (e.g., the mutated proteins reach the plasma membrane when co-expressed with 4F2hc but no transport activity is elicited). [408] [414] Mutation E36del showed a dominant negative effect when expressed in Xenopus oocytes.[415] The molecular basis for this effect is not yet fully understood.

Pathophysiology of Lysinuric Protein Intolerance

Lysinuric protein intolerance is a multisystemic disease. Some of the symptoms of this disease, like the renal and intestinal phenotypes, are easily explained by a defect in the basolateral amino acid transport system y+L. Urea cycle malfunction is a characteristic of patients with LPI after weaning. Patients with LPI have a decreased tolerance for nitrogen and present with hyperammonemia after ingestion of even moderate amounts of protein. The malfunction of the urea cycle in LPI is less severe than that caused by defects in the enzymes of the cycle. y+LAT1 is not expressed in hepatoytes. [402] [405] It is believed that urea cycle malfunction is due to diminished availability of the intermediates of this cycle because of their low concentration in plasma (“intermediate functional deficiency hypothesis”). The mechanisms underlying the LPI-associated immune-related disorders (e.g., alveolar proteinosis, erythroblastophagia, and glomerulonephritis) are unknown. In addition, individual phenotypic variability precluded establishment of genotype/phenotype correlations. [408] [411] Thus, Finnish patients with LPI, all with the same Finnish mutation in homozygosis, show a wide range of phenotypic severity ranging from nearly normal growth with minimal protein intolerance to severe cases with hepatosplenomegaly, osteoporosis, alveolar proteinosis, and severe protein intolerance. In the following part of this section, the mechanisms that explain the renal and intestinal pathophysiology in LPI are discussed.

Table 6-9 compares plasma and urine levels for several amino acids in patients with LPI and cystinuria. Plasma concentrations of the dibasic amino acids (i.e., lysine, arginine, and ornithine) are usually subnormal (one third to one half of the normal values), but occasionally may fall within the normal range. Similarly, but to a lesser extent, plasma dibasic and cystine concentrations are lower in patients with cystinuria.[416] This observation indicates that the defects in renal reabsorption and intestinal absorption of dibasic amino acids may have a greater impact in LPI than in cystinuria, and therefore produce a larger depletion of these amino acids in plasma. In contrast to dibasic amino acids, the plasma concentrations of the neutral amino acids glutamine and alanine are increased in patients with LPI (see Table 6-9 ), and to a lesser extent serine, glycine, citrulline, and proline. The considerable increase in plasma glutamine and alanine in LPI is believed to be the result of the large amount of waste nitro-gen not incorporated into urea as a result of urea cycle malfunction.

In LPI, urinary excretion and renal clearance of lysine is massively increased, whereas that of arginine and ornithine is moderately augmented[417]: lysine excretion is 10-fold and 30-fold that of arginine and ornithine in LPI patients, respectively (see Table 6-8 ). In contrast, lysine excretion is only twofold to threefold higher than that of arginine and ornithine in patients with cystinuria. Renal reabsorption of lysine is comparable in LPI and cystinuria, whereas hyperexcretion of arginine and ornithine are lower in LPI than in cystinuria (see Table 6-8 ). These observations indicate that the LPI-defective transporter (y+LAT-1/4F2hc) may have a more pronounced role in the reabsorption of lysine than of the other dibasic amino acids. In contrast to cystinuria, where cystine excretion in urine is four times lower than that of lysine, in LPI there is only a slight increase of renal cystine excretion (see Table 6-8 ). This may be explained by the large tubular lysine load (i.e., caused by the reabsorption defect of lysine) that competes for absorption through the apical system b0,+, and shares uptake of cystine and dibasic amino acids in exchange with other neutral amino acids. The increased plasma concentration of serine, glycine, citrulline, proline, alanine, and glutamine in LPI explains hyperexcretion of these amino acids, and their renal clearance is within the normal range.

The defect in kidney and intestine in LPI is located in the basolateral membrane and thus affects the basolateral efflux of dibasic amino acids. [418] [419] An oral load with the dipeptide lysyl-glycine increased glycine plasma concentrations, but plasma lysine remained almost unchanged in patients with LPI, whereas both amino acids increased in plasma of control subjects or in patients with cystinuria. [420] [421] Figure 6-13 shows the present knowledge on the molecular bases of the intestinal absorption of dibasic amino acids. At the luminal membrane of the enterocyte, the transport of oligopeptides (not shared with amino acids) is mediated by PEPT1.[422] A major route for dibasic amino acids across the apical membrane is system b0,+ (i.e., the transporter defective in cystinuria). The absorbed peptides are hydrolyzed to release amino acids in the cytoplasm of the enterocyte [423] [424] [425] and are able to cross the basolateral membrane only as free amino acids. The lack of increased plasma lysine after the lysyl-glycine load, but normal increase in plasma glycine, shows that the basolateral efflux of the intracellularly delivered lysine is defective in LPI. In patients with cystinuria, the cleaved glycine and lysine cross the epithelial cell normally because the defect is apical (i.e., system b0,+) (see Fig. 6-13 ). The defect in the basolateral system y+L explains the renal and the intestinal phenotypes in LPI. The protein y+LAT-1 has a basolateral location in epithelial cells. System y+L (i.e., the 4F2hc/y+LAT-1 heteromeric complex) mediates the efflux of cationic amino acids by exchange with extracellular neutral amino acids and sodium (see Fig. 6-13 ). Thus, the loss of transport function of the LPI-associated y+LAT-1 mutations results in a defective basolateral efflux of dibasic amino acids in the intestinal absorptive and renal reabsorptive epithelial cells.

Hartnup Disorder

The original patients with Hartnup disorder presented cerebellar ataxia, tremor, nystagmus, pellagra-like photosensitive skin rash, and delayed intellectual development. Hartnup disorder affects the renal reabsorption and intestinal absorption of neutral amino acids with the exception of proline, hydroxyproline, glycine, and cystine. Pellagra-like symptoms (i.e., niacin deficiency) are frequent in patients with this disorder. Low tryptophan availability (i.e., defective renal and intestinal reabsorption of the amino acids) appears to be at the basis of the niacin deficiency: tryptophan and niacin deficiencies are thought to generate similar symptoms because this amino acid is a major source of NAD(P)H in humans. In this regard, pellagra-like symptoms respond to nicotinic acid supplementation.

The incidence of Hartnup disorder has been estimated at 1 in 26,000 in newborn screening programs.[306] The trait is transmitted in autosomal recessive fashion, but clinical manifestations are probably modulated by environmental and genetic factors.[426]

System B0 neutral amino acid transporter has been considered the defective transporter in Hartnup disorder. Large neutral amino acids are mainly absorbed in the small intestine and reabsorbed in the proximal convoluted tubule (i.e., S1-S2 segments) by the apical system B0 (reviewed in Ref 291). Functional studies in renal and intestinal brush-border membrane vesicles and derived cell models defined system B0 (B for broad and 0 for neutral charge[427]) as a transporter serving a broad spectrum of neutral amino acids. System B0 mediates co-transport of Na+ and neutral amino acids with 1:1 stoichiometry, where Na+ and amino acid affects each other's kinetic parameters (reviewed in Ref 291).

Broer's group demonstrated that mouse B0AT1 (previously the orphan XTR2-related transporter) when expressed in Xenopus oocytes induces Na+-dependent and Cl-independent transport of neutral amino acids with broad specificity, matching the characteristic of system B0. [295] [428] Apparent Km for neutral amino acids ranges from 1 μM to 10 μM with the following substrate specificity (one letter code for amino acids): M=L=I=V > Q=N=C=F=A > S=G=Y=T=H=P > W.[428] The human ortholog showed similar transport characterisics. [296] [348] Human B0AT1 mRNA is expressed mainly in kidney and small intestine, and to a lesser extent in colon, pancreas, and prostate.[296] [348] Mouse B0AT1 was localized to the brush-border membrane of the epithelial cells of the renal proximal convoluted tubule (S1-S2 segments) and of the small intestine with a gradient of expression from the crypts toward the tip of the microvilli. [295] [296] Human B0AT1 gene (SLC6A19) localized to chromosome 5p15.33,[348] and Hartnup disorder to chromosome 5p15 in Japanese families transmitting the disease.[429] Thus, SLC6A19 was an obvious functional and positional candidate gene for Hartnup disorder.

In 2004, two independent studies demonstrated that mutations in SLC6A19 are associated with Hartnup disorder and confirmed the recessive mode of inheritance. [296] [348] Patients from the Hartnup family were homozygotes for mutation IVS8+2T>G affecting the donor splice consensus sequence of exon 8.[296] In seven Australian pedigrees, six distinct mutations that cosegregated with the disorder were identified (three missense, one nonsense, and two splice site mutations), including one Australian family transmitting the Hartnup family mutation in one allele. In the Australian population, D173N and R240X mutations occur at a frequency of 1 in 140 and 1 in 1000 people, respectively. Four further mutations were identified in three Japanese families (one missense, one nonsense, and two small deletions causing frameshift).[296] In total, 10 Hartnup disorder-specific SLC6A19 mutations have been identified in 13 independent pedigrees. This implies that ≈73% of the independently studied alleles have been identified (19 of 26 alleles).

Recently the crystal structure of a prokaryotic homolog (LeuTAa from Aquifex aeolicus) of the SLC6 family has been reported.[430] This structure will be very useful to ascertain the molecular events underlining the defects associated with Hartnup disorder mutations. The four Hartnup disorder-specific SLC6A19 missense mutations (R57C, D173N, L242P, E501K) were checked for function in oocytes. [296] [348] These mutations showed no transport function, with the exception of the most common mutation D173N, which has residual transport activity (≈50%). Figure 6-15 shows the location of these mutations within the topology of SLC6 transporters. R57C destroys a saline bridge with residue Asp486. This bond helps to hold the position of TM1b, which interacts with the amino acid substrate and the two Na+ ions. Leu242 involves the first of two residues constituting the extracellular b1 sheet, and mutation L242P, most likely, disrupt this structure. Glu501 in TM10 interacts with one of two water molecules that holds the structure of the unwound residues between TM6a and TM6b, which interact with the amino acid substrate and one of the Na+ ions. Then, mutation E501K most probably affects the folding of this unwound region. Finally, mutation D173N is a conservative amino acid substitution affecting a residue not conserved among the SLC6 transporters in the extracellular α-helix EL2. Then, not surprisingly, this mutation retains significant transport activity.[348]

000352

000519

FIGURE 6-15  The predicted topology is based on the crystal structure of LeuTAa from bacteria Aquifex aeolicus.[409] There is a structural repeat, not based on amino acid sequence, in the first ten transmembrane (TM) helices of LeuTAa, relating TM1–TM5 (pink triangle) and TM6–TM10 (blue triangle) by a pseudo-twofold axis located in the plane of the membrane. LeuTAa is a bacterial homolog of Na+/Cl--dependent neurotransmitter transporters (family SLC6), to which the defective Hartnup disorder transporter (B0AT1) belongs. Amino acid sequence homology of B0AT1 and LeuTAa is ≈20% and covers the whole sequences (CLUSTAL alignment; data not shown). Major amino acid sequence differences between LeuTAa and the eukaryotic SLC6 transporters are located at the N- and C termini, between TM3 and TM4, and between extracellular α-helices EL4a and EL4b (these segments are longer in B0AT1 and in other SLC6 eukaryotic transporters). The positions of the substrate leucine and the two sodium ions are shown as a yellow triangle and two blue circles, respectively. Residues interacting with the substrate and ions are located within and surrounding the unwound regions between TM1a and TM1b, and TM6a and TM6b, as well as in TM3 and TM8. Hartnup disorder-specific missense B0AT1 mutations are indicated within the LeuTAa topology. Rectangle, α-helix. Arrow, β-sheet.  (Figure modified from Smith DW, Scriver CR, Simell O: Lysinuric protein intolerance mutation is not expressed in the plasma membrane of erythrocytes. Hum Genet 80:395, 1988.) Location of mutations within the topology of SLC6 transporters. R57C destroys a saline bridge with Asp486 holding the position of TM1b, which interacts with the amino acid substrate and the two Na+ ions. Leu242 involves the extracellular b1 sheet, and L242P, most probably, disrupt this structure. Glu501 in TM10 interacts with one of two water molecules that hold the structure of the unwound residues between TM6a and TM6b, which interact with the amino acid substrate and one of the Na+ ions. Mutation E501K most probably affects the folding of this unwound region. Finally, mutation D173N is a conservative amino acid substitution affecting a residue not conserved among the SLC6 transporters in the extracellular α-helix EL2.

000519

 

 

Genetic Heterogeneity and Phenotype Variability

Taken together, these results demonstrated that mutations in SLC6A19 cause Hartnup disorder. However, individuals that display Hartnup-like aminoaciduria without apparent mutations in SLC6A19 (in two American pedigrees[296]) have been reported; similar results have been described by the Australian Hartnup Consortium.[291] Indeed, genetic linkage of Hartnup disorder with the 5p15 region has been excluded in an American family.[296] This finding indicates that additional Hartnup disorder genes may be involved and remain to be identified. Five candidate neutral amino acid transporters have been excluded (genetic linkage exclusion and/or lack of co-segregating mutations) as causative genes of the disorder[348]SLC3A2 (4F2hc), SLC7A8 (LAT2), SLC1A5 (ASCT2 or ATB0), SLC6A18 (orphan Xtrp2), and SLCA20 (orphan XT3). A system B0-like activity in the proximal straight tubule (S3 segment), which has not yet been identified, is an obvious candidate for Hartnup disorder.[291]

Patients with Hartnup disorder display a wide phenotype range. This was described in the original report of the Hartnup family: of the four siblings with clear aminoaciduria, two presented severe clinical symptoms, one had mild symptoms, and one was asymptomatic. Symptoms most likely appear in individuals with subnormal plasma amino acid levels (reviewed in Ref 426). Intestinal absorption of peptides, via PEPT1, is thought to compensate for the lack of amino acid transport in Hartnup disorder (see Fig. 6-13 ).[307] This compensation has two consequences. On the one hand, in developed societies, characterized by high protein intake, most patients will remain asymptomatic. Only a limited number of patients will display symptoms (e.g., subnormal body weight, episodes of diarrhea, pellagra-like rash, etc.).[431] On the other hand, genetic factors may predispose individuals to a more severe deficiency in amino acid uptake. The phenotype of Hartnup disorder could be influenced by the amino acid transporters, which participate in the renal reabsorption and intestinal absorption of amino acids: other apical transporters for neutral amino acids (e.g., the B0-like activity in the proximal straight tubule) and basolateral transporters. Polymorphisms in these transporters may contribute to heterogeneity in the phenotype of Hartnup disorder.

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