Basic Research

Kidney Proximal Tubule Lipoapoptosis Is Regulated by Fatty Acid Transporter-2 (FATP2)

Shenaz Khan, Pablo D. Cabral, William P. Schilling, Zachary W. Schmidt, Asif N. Uddin, Amelia Gingras, Sethu M. Madhavan, Jeffrey L. Garvin and Jeffrey R. Schelling
JASN January 2018, 29 (1) 81-91; DOI: https://doi.org/10.1681/ASN.2017030314

Abstract

Albuminuria and tubular atrophy are among the highest risks for CKD progression to ESRD. A parsimonious mechanism involves leakage of albumin-bound nonesterified fatty acids (NEFAs) across the damaged glomerular filtration barrier and subsequent reabsorption by the downstream proximal tubule, causing lipoapoptosis. We sought to identify the apical proximal tubule transporter that mediates NEFA uptake and cytotoxicity. We observed transporter-mediated uptake of fluorescently labeled NEFA in cultured proximal tubule cells and microperfused rat proximal tubules, with greater uptake from the apical surface than from the basolateral surface. Protein and mRNA expression analyses revealed that kidney proximal tubules express transmembrane fatty acid transporter-2 (FATP2), encoded by Slc27a2, but not the other candidate transporters CD36 and free fatty acid receptor 1. Kidney FATP2 localized exclusively to proximal tubule epithelial cells along the apical but not the basolateral membrane. Treatment of mice with lipidated albumin to induce proteinuria caused a decrease in the proportion of tubular epithelial cells and an increase in the proportion of interstitial space in kidneys from wild-type but not Slc27a2−/−mice. Ex vivo microperfusion and in vitro experiments with NEFA-bound albumin at concentrations that mimic apical proximal tubule exposure during glomerular injury revealed significantly reduced NEFA uptake and palmitate-induced apoptosis in microperfused Slc27a2−/− proximal tubules and Slc27a2−/− or FATP2 shRNA-treated proximal tubule cell lines compared with wild-type or scrambled oligonucleotide–treated cells, respectively. We conclude that FATP2 is a major apical proximal tubule NEFA transporter that regulates lipoapoptosis and may be an amenable target for the prevention of CKD progression.

Over 26 million people in the United States are afflicted with CKD,1 and risks for CKD progression and mortality are albuminuria and decreased GFR.2 Although albuminuria reflects glomerular filtration barrier dysfunction, downstream tubular atrophy due to tubular epithelial cell apoptosis is superior to glomerular pathology as a predictor of CKD progression.3–6 Infusion of albumin in animal models7–9 or exposure of proximal tubule cells to albumin in vitro10–12 results in apoptosis. However, cytotoxicity is not observed with delipidated albumin administration,8,10,12,13 implying that albumin-bound fatty acids cause apoptosis. Mouse models of CKD show increased concentrations of NEFAs and NEFA metabolites in the kidney, which are hypothesized to cause renal function deterioration.12,14

Proposed mechanisms leading to proximal tubule lipid accumulation include increased uptake, increased synthesis, and diminished β-oxidation of NEFA,15–17 but the relative importance of each mechanism has not been determined. Increased NEFA uptake is plausible, because hyperlipidemia commonly coexists with CKD, resulting in enhanced extracellular NEFA supply for transport and intracellular storage by nonadipose tissues, such as in liver, skeletal muscle, and kidney.18 More importantly, in proteinuric renal diseases, such as diabetic nephropathy, the damaged glomerulus permits albumin-bound NEFA to be filtered and gain access to the previously unexposed proximal tubule luminal surface, where aberrant NEFA reabsorption could then occur.

Under normal circumstances, fatty acids are the preferred substrate for proximal tubule ATP generation.19,20 In plasma, esterified fatty acids circulate as triglycerides, whereas water-insoluble NEFAs are solubilized by complexing with albumin. NEFAs dissociate from albumin at the plasma membrane and are taken up by saturable, basolateral membrane transporters in proximal tubules.21–24 Under pathologic circumstances, the possibility of simultaneous apical and basolateral proximal tubule NEFA uptake combined with increased NEFA synthesis15,16 and diminished β-oxidation17 could lead to intracellular NEFA accumulation and tubular atrophy through a lipotoxicity mechanism.12,14

Our goal was to identify the apical membrane proximal tubule NEFA transporter, so that it could ultimately be exploited as a therapeutic target to prevent CKD progression. There are several candidate transporter classes, most notably the FATP family. FATP2 is exclusively expressed in kidney and liver, and it is the most abundant isoform in kidney.25 Using in vitro, ex vivo, and in vivo approaches, we show that FATP2 regulates proximal tubule apical NEFA transport and lipoapoptosis.

Results

NEFAs Are Absorbed by the Proximal Tubule

To screen for proximal tubule apical NEFA uptake, experiments were initially conducted in proximal tubule cell lines. Figure 1A shows rapid basolateral NEFA absorption in polarized LLC-PK1 cells, with time-dependent uptake. The rate and magnitude of NEFA uptake were considerably greater from the apical surface. Figure 1B shows concentration-dependent apical NEFA uptake.

Figure 1.
Figure 1.

NEFAs are absorbed by the proximal tubule in vitro. (A) LLC-PK1 cells were grown to confluence on permeable supports and then incubated with apical (AP) or basolateral (BL) BODIPY-labeled C16-NEFA (2.5 μM) complexed with albumin (0.2%) in buffer prewarmed to 37°C. BODIPY uptake was measured by fluorescence microscopy using a QBT kit (Molecular Devices; excitation/emission λ=488/510 nm; mean±SEM from 30 cells; n=3). (B) Concentration-dependent apical uptake of BODIPY-labeled NEFAs in LLC-PK1 maintained on permeable supports (mean±SEM from n=3).

To test NEFA uptake under native conditions, microdissected rat proximal tubules were perfused with fluorescently labeled NEFA using established methods.26 Figure 2, A and B depicts a proximal tubule immediately and then 10 minutes after luminal microperfusion with boron-dipyrromethene (BODIPY)-tagged NEFA, respectively. Figure 2C shows rapid time-dependent uptake and approximately fivefold greater NEFA internalization from the apical compared with the basolateral surface, consistent with Figure 1A in vitro data. Figures 1 and 2 collectively show that proximal tubule epithelial cells reabsorb NEFA from the apical surface and by a magnitude that exceeds basolateral uptake.

Figure 2.
Figure 2.

NEFA are absorbed from the proximal tubule apical surface. Fluorescence image of a rat proximal tubule (A) immediately and (B) 10 minutes after lumen perfusion with BODIPY-labeled NEFAs. (C) BODIPY-NEFA (2.5 μM; complexed with 0.2% albumin) uptake in a microdissected rat proximal tubule after luminal perfusion (left panel), addition of BODIPY-NEFA to the basolateral bath (center panel), and luminal perfusion of BODIPY-NEFA (500 nM) in Hepes (10-mM)–buffered saline (right panel). Data are representative of experiments from 12 tubules.

Proximal Tubule FATP2 Expression

FATPs (gene name Slc27) are a family of transmembrane spanning proteins, which contain six members with distinct tissue expression patterns. Of the FATP isoforms expressed in kidney, FATP1 and FATP4 are present in very small amounts25 (S. Khan et al., unpublished observations). FATP2 is expressed exclusively in kidney and liver and has previously been detected in total kidney and proximal tubule.25,27–32 Figure 3A shows FATP2 mRNA expression in wild-type mouse kidney, which is diminished in Slc27a2+/− and absent in Slc27a2−/− control kidneys. Figure 3, B–D reveals that FATP2 protein expression in kidney is restricted to proximal tubule. No FATP2 staining was detected in Slc27a2−/− kidneys (not shown). Figure 3E confirms FATP2 mRNA expression in Human Renal Proximal Tubule (HRPT) and HK-2 human proximal tubule cell lines. FATP2 transcripts were weakly detectable by RT-PCR in LLC-PK1 cells (Figure 3E), perhaps reflecting nucleotide sequence differences between human and porcine mRNA (porcine cDNA sequence is unknown).

Figure 3.
Figure 3.

FATP2 is expressed in proximal tubules. (A) Kidney FATP2 and GAPDH mRNA expression by RT-PCR. Immunohistochemical localization of (B) FATP2, (C) T. purpureus lectin staining, and (D) merged image in mouse kidney cortex. Gl, glomerulus. *Distal tubule. (E) FATP2 and GAPDH mRNA expression by RT-PCR in proximal tubule cell lines.

Because FATP2 may not be the sole FATP in the proximal tubule, screens for other plausible transporters were undertaken. CD36 has been reliably isolated in mouse kidney,33 but it was not detectable by immunohistochemistry in proximal tubules (Supplemental Figure 1)12,34 or by RT-PCR in HK-2 cells (not shown). The G protein–coupled GP40 and GP120 long-chain NEFA transporters have recently been deorphanized and termed FFA1 and FFA4, respectively; only FFA1 is expressed in kidney.35,36 Supplemental Figure 2 shows FFA1 expression in a glomerular epithelial cell (podocyte) pattern but not within tubules, consistent with the previously described role of podocyte FFA1-mediated NEFA uptake in the pathogenesis of the nephrotic syndrome.37 Unlike FATP2, CD36 and FFA1 are, therefore, not candidate transporters for regulation of proximal tubule NEFA reabsorption.

FATP2 Is Expressed in Proximal Tubule Apical Membranes

Like other members of the FATP family, FATP2 is predicted to be a membrane-associated transporter containing two transmembrane domains.38,39 To screen for FATP2 membrane expression, crude membrane fractions from cultured proximal tubules were probed by immunoblotting. Figure 4A reveals FATP2 expression in LLC-PK1 and HK-2 proximal tubule cell membranes. Immunocytochemistry experiments in postfixed HK-2 cells revealed FATP2 expression in a plasma membrane distribution (Figure 4B). Immunoprecipitation of biotin surface–labeled FATP2 also showed abundant expression in LLC-PK1, HK-2, and HRPT proximal tubule cell lines (Figure 4C), indicating that FATP2 is expressed on the plasma membrane.

Figure 4.
Figure 4.

FATP2 is expressed in proximal tubule membranes. (A) Lysates from crude membrane preparations were immunoblotted with anti-FATP2 antibodies (upper panel). Membranes were stripped and reprobed with anti–Na+/K+-ATPase antibodies as a loading control (lower panel). (B) HK-2 cells on coverslips were probed with anti-FATP2 antibodies and postfixed in paraformaldehyde (4%, 10 minutes at room temperature), and primary antibody detection was amplified with Alexa Fluor 568–conjugated goat anti-rabbit IgG. (C) Proximal tubule cell lines were surface labeled with biotin, and lysates were precipitated with streptavidin-coated beads, eluted, resolved by SDS-PAGE, and immunoblotted with anti-FATP2 antibodies (upper panel). Membranes were stripped and reprobed with anti-Glut5 antibodies as a loading control (lower panel).

To establish FATP2 localization, human kidney sections were probed with FATP2 and γ-glutamyl transferase-1 (a brush border enzyme) antibodies. Figure 5 reveals FATP2 and γ-glutamyl transferase-1 colocalization in an apical membrane pattern. To confirm FATP2 membrane domain localization, mouse kidney sections were colabeled with Glut5 (a luminal brush border protein) and Na+/K+-ATPase (to demarcate the basolateral membrane) antibodies. Supplemental Figure 3, A–D shows the diffuse FATP2 cytosolic pattern, although colocalization was also observed with Glut5, indicating that FATP2 is expressed on the proximal tubule apical membrane. In contrast, no FATP2 colocalization with Na+/K+-ATPase was noted (Supplemental Figure 3, E–G), indicating that FATP2 is not expressed in basolateral membranes and would, therefore, not mediate basolateral NEFA uptake (Figure 6C). Taken together, Figures 4 and 5 and Supplemental Figure 3 show that proximal tubule FATP2 is expressed most prominently within the apical plasma membrane.

Figure 5.
Figure 5.

FATP2 is expressed in proximal tubule apical membranes. Formalin-fixed paraffin sections from human kidneys were probed with (A) anti-FATP2 and Alexa Fluor 488 secondary antibodies or (B) anti–γ-glutamyl transferase-1 (GGT1) and Alexa Fluor 568–labeled secondary antibodies. (C) Nuclei were labeled with DAPI in the mounting medium. (D) Colocalization from merged images is depicted in yellow.

Figure 6.
Figure 6.

Proximal tubule FATP2-regulated NEFA uptake is time- and concentration-dependent. (A) BODIPY-NEFA (500 nM NEFA complexed with 0.2% albumin in Hepes [10-mM]–buffered saline) uptake after luminal perfusion in wild-type versus Slc27a2−/− tubules microdissected from 12-week-old mice. A representative tracing (from n=5) is shown. (B) BODIPY-NEFA (20–2500 nM NEFA in fivefold dilutions complexed with 0.2% albumin in Hepes [10-mM]–buffered saline) uptake after luminal perfusion in wild-type versus Slc27a2−/− tubules microdissected from 12-week-old mice (mean±SEM from n=8). (C) BODIPY-NEFA uptake from the basolateral bath. Representative tracings (from n=3) are shown for clarity due to overlapping error bars. (D) BODIPY-NEFA uptake in temperature-sensitive SV40 immortalized proximal tubule cell lines derived from wild-type and Slc27a2−/− mice (mean±SEM; n=4). (E) Concentration-dependent uptake of BODIPY-labeled NEFA in wild-type versus Slc27a2−/− mouse proximal tubule cell lines (mean from n=4).

Proximal Tubule FATP2-Regulated NEFA Uptake

FATP2-dependent NEFA uptake was initially evaluated in experiments with microdissected proximal tubules from wild-type and Slc27a2−/− mice, which were perfused with BODIPY-labeled NEFA. No difference was observed between male and female tubules, and therefore, results from both sexes are combined. Figure 6, A and B shows enhanced time- and concentration-dependent NEFA uptake from luminal perfusion of wild-type compared with Slc27a2−/− tubules. In contrast, basolateral NEFA was diminished compared with apical uptake, and the magnitude and kinetics of basolateral NEFA transport were identical in microperfused wild-type and Slc27a2−/− proximal tubules (Figure 6C), indicating that basolateral NEFA uptake does not require FATP2. NEFA uptake was also greater in proximal tubule cell lines derived from wild-type versus Slc27a2−/− mice (Figure 6, D and E), similar to results from tubules perfused ex vivo (Figure 6, A and B). These results show that a large proportion of apical (but not basolateral) proximal tubule NEFA uptake is mediated by FATP2.

FATP2 Mediates Tubulointerstitial Disease

To determine the pathophysiologic significance of proximal tubule FATP2, wild-type and Slc27a2−/− mice were treated with daily intraperitoneal injections of lipidated albumin to induce albuminuria (Figure 7A) and tubulointerstitial injury.8,40 Quantitative histomorphometric analysis revealed modest tubular atrophy and interstitial fibrosis, although tubular epithelial cell number and interstitial and tubular lumen areas were significantly different between wild-type and Slc27a2−/− mouse kidneys (Figure 7, B–D). Similar differences in tubular epithelial cell density were also observed in wild-type versus Slc27a2−/− mice after induction of albuminuria with daily LPS injections (Supplemental Figure 4).41,42 The pathophysiology of tubular atrophy is complex, but one proposed pathway is apoptosis, in the absence of compensatory hyperplasia.43 Figure 7E shows increased tubular epithelial cell apoptosis in lipidated albumin–injected mice, which was significantly attenuated in Slc27a2−/− compared with wild-type mice. Taken together, the in vivo data suggest that apical proximal tubule FATP2-mediated NEFA uptake contributes to tubular atrophy and interstitial fibrosis.

Figure 7.
Figure 7.

FATP2 mediates tubulointerstitial injury. Wild-type and Slc27a2−/− mice were treated with daily intraperitoneal lipidated albumin (or saline control) injections to induce albuminuria and tubulointerstitial disease as described in Concise Methods. After 3 weeks, mice were euthanized and evaluated for tubulointerstitial injury using previously described quantitative histomorphometry methods. (A) Coomassie Blue–stained gel showing albuminuria after the 3-week protocol was completed. Quantification of (B) tubular epithelial cells, (C) interstitium, (D) tubular lumen, and (E) apoptotic proximal tubular epithelial cells within the tubulointerstitial compartment. Data are expressed as mean±SEM (n=5). P values were generated using ANOVA with Bonferroni test for post hoc between-group comparisons. IP, intraperitoneal; NS, not significant (P>0.05).

Proximal Tubule Luminal FATP2-Mediated NEFA Uptake Is Cytotoxic

Accumulation of NEFAs and NEFA metabolites has been shown to be cytotoxic to proximal tubule epithelial cells (Supplemental Figure 5)8,10,12 and may contribute to tubular atrophy and CKD progression.12,14 To directly test whether FATP2 regulates lipotoxicity, BODIPY-tagged NEFA uptake was measured in FATP2 shRNA-silenced versus shRNA-scrambled sequence-treated HRPT human proximal tubule cell lines (Figure 8, A and B). Figure 8C depicts cells with oil red O–stained lipid droplets as an indirect measure of NEFA uptake and qualitatively confirms the efficacy of functional FATP2 knockdown with two of the three shRNA constructs. Oil red O quantification revealed reduced staining in FATP2 shRNA compared with scrambled nucleotide sequence control cells (Figure 8D), once again indicating that proximal tubule FATP2 mediates NEFA uptake. Figure 8, E and F shows that sustained NEFA exposure caused apoptosis, as shown by terminal deoxynucleotidyl transferase–mediated digoxigenin-deoxyuridine nick-end labeling (TUNEL) and caspase-2 activation, which was significantly reduced in FATP2 shRNA-expressing cells compared with scrambled nucleotide–treated cells. NEFA-induced apoptosis was also observed in proximal tubule cells lines derived from wild-type mice and significantly blunted in Slc27a2−/− mouse proximal tubule cells (Figure 8G). Taken together, these loss of function approaches show that FATP2 regulates lipoapoptosis.

Figure 8.
Figure 8.

FATP2 mediates proximal tubule lipoapoptosis. (A) HRPT cells were stably transfected with three FATP2 shRNA constructs or scrambled oligonucleotide (control). Lysates from a crude membrane fraction were probed by immunoblot for FATP2 expression. (B) Coomassie Blue–stained membrane as loading control. (C) Images of transfected HRPT cells exposed to apical 0.5% delipidated BSA complexed with palmitate (100 μM) for 24 hours. Fixed cells were stained with oil red O and counterstained with hematoxylin. (D) Oil red O was eluted with 100% propanol from shRNA- or scrambled oligonucleotide sequence (control)–transfected HRPT cells and quantitated by spectrophotometry (λ=492 nm; mean±SEM; n=3). FATP2 shRNA or scrambled sequence–treated HRPT proximal tubule cells were incubated with 0.2% albumin + palmitate (100 μM; 24 hours), and apoptosis was determined by (E) TUNEL or (F) immunoblot with rat antiactive (cleaved) caspase-2 IgG and α-tubulin loading control (n=3). (G) Proximal tubule cell lines derived from wild-type and Slc27a2−/− mice were incubated with 100 μM palmitate complexed with 0.2% fatty acid–free albumin (Palm) or albumin-only control (BSA) for 24 hours and then assayed for apoptosis by TUNEL (n=3). *P<0.05 compared with the BSA-treated group by t test; **P<0.05 compared with other groups by ANOVA.

Discussion

CKD associated with heavy albuminuria accounts for the vast majority of causes of ESRD. There has been significant progress in understanding the role of glomerular cells, particularly the glomerular epithelial cell (podocyte), in the pathophysiology of albuminuria, but it is often overlooked that tubular atrophy more accurately predicts CKD progression compared with glomerular pathology.3–6 Because each albumin molecule that leaks across the damaged glomerular filtration barrier has the capacity to bind up to seven NEFA molecules, it has been suggested that proximal tubule reabsorption and intracellular accumulation of NEFAs are cytotoxic and contribute to tubular atrophy pathogenesis.12,14 Using combined in vitro and ex vivo approaches, we show that proximal tubules rapidly reabsorb luminal NEFA from the apical surface and by a magnitude that is several fold greater than from the basolateral surface. Uptake was achieved by complexing NEFA with albumin at a concentration of 2 g/L, which corresponds to apical proximal tubule exposure of albumin under conditions that mimic glomerular filtration barrier dysfunction. Our data support the notion that aberrantly filtered NEFAs are efficiently reabsorbed by the proximal tubule and contribute to cytotoxicity by expediting lipoapoptosis.

A major advance in this report is the detailed characterization of the proximal tubule transporter, FATP2, which mediates apical NEFA uptake and lipoapoptosis. We confirmed that FATP2 is expressed in kidney and localizes to proximal tubule in an apical plasma membrane and cytoplasmic distribution. Subcellular identification was not pursued in detail in our studies, but the murine FATP2 cytosolic and plasma membrane immunohistochemical staining pattern is remarkably similar to the peroxisome and plasma membrane FATP2 expression pattern described in liver.31 Hepatic FATP2 exhibits peroxisomal very long–chain (>22-carbon chain length) acyl CoA synthetase activity and plasma membrane transport of long-chain (12–22 carbons) NEFAs, which ultimately undergo β-oxidation in mitochondria.32 The diffuse cytosolic and apical membrane pattern observed in Figures 3 and 4 suggests that both FATP2 functions may be operant in the proximal tubule. Using in vitro and ex vivo approaches, we also conclude that apical uptake of NEFA was largely mediated by membrane-localized FATP2.

Apical NEFA uptake capacity at equilibrium was significantly greater than basolateral uptake, suggesting that the apical-directed NEFA compartment size is different and larger than for basolateral uptake. Because fatty acids are a major energy source for the proximal tubule, we propose that, under normal circumstances, NEFA uptake across the basolateral membrane might be targeted to a subcellular compartment associated with metabolism, whereas damaged glomeruli filter albumin-bound NEFAs, which are transported by apical FATP2, and then may traffic to a compartment that ultimately initiates apoptosis. Thus, disparate cytotoxicity after apical versus basolateral NEFA uptake could be explained by differential NEFA partitioning. It is also noteworthy that apical NEFA uptake at equilibrium was significantly lower in Slc27a2−/− compared with wild-type proximal tubules, suggesting that metabolism and distribution to intracellular pools may differ after FATP2- and non–FATP2-mediated uptake. Plasma membrane FATPs are coupled to specific long-chain acyl CoA synthetases, which accelerate NEFA flux and shuttle fatty acids to binding proteins (FABPs) that then channel NEFAs to specific intracellular sites. We speculate that FATP/acyl CoA synthetase/FABP isoform combinations could also segregate NEFA to different compartments after transport across plasma membrane domains.44,45

We have previously shown that proximal tubule cells undergo apoptosis after exposure to NEFA (palmitate) under conditions that mimic apical albumin and NEFA concentrations in the context of glomerular filtration barrier injury.12 This concept of lipoapoptosis was invoked 35 years ago as a cause of tubular atrophy by Moorhead et al.,14 but the specific mechanisms that regulate tubular epithelial cell lipoapoptosis had not been previously described. At a molecular level, lipoapoptosis has been associated primarily with long-chain (C12–C22) NEFA accumulation, and cytotoxicity is proportional to acyl chain length and carbon bond saturation.46,47 In addition, the quantity of bound NEFA per albumin molecule may regulate toxicity as evidenced in clinical studies of minimal change disease, which does not progress due to absence of tubular atrophy and interstitial fibrosis. Despite massive albuminuria, the ultrafiltrate albumin was markedly NEFA deplete compared with serum in minimal change disease. In contrast, the ultrafiltrate NEFA content was relatively unchanged in other types of nephrotic syndrome,48 suggesting that the quantity and composition of NEFA bound to filtered albumin dictate whether albuminuric renal diseases progress or AKI develops after a relapse. With this caveat in mind, our in vivo and in vitro experiments were performed using albumin that was saturated with palmitate, the most toxic long-chain fatty acid. Slc27a2−/− mice developed less reduction in tubular epithelial cell number and interstitial fibrosis in the albumin overload model of inducible proteinuria, in which mice received daily intraperitoneal injections of palmitate-loaded albumin. Furthermore, Slc27a2−/− and shRNA-treated proximal tubule cell lines were almost completely protected from palmitate-induced apoptosis, suggesting that FATP2 plays a major pathophysiologic role in the apical uptake of filtered NEFAs that result from glomerular injury.

Proximal tubule lipotoxicity is likely to result from a combination of increased uptake, increased synthesis, and decreased catabolism of NEFA. The intracellular pathways are complex and may not be accessible to small molecule pharmacologic inhibitors, and therefore, they represent less rational therapeutic targets compared with apical FATP2. High-throughput screens of small molecule libraries for FATP2 inhibitors have been conducted to identify lead molecules for the treatment of hepatosteatosis.49,50 Some candidates have emerged, although enthusiasm has been dampened, because many of the compounds with the best inhibition profiles (e.g., anesthetics and tricyclic antidepressants) could cause nonspecific membrane disruption rather than specific inhibition of FATP2 and at high IC50 values (midmicromolar), which may be unattainable in blood. However, pharmacodynamics studies have not been conducted, and if these drugs are filtered and/or secreted by the kidneys, low doses could result in sufficiently high luminal concentrations to be effective inhibitors of proximal tubule FATP2 and lipoapoptosis-dependent tubular atrophy.

There are six FATP isoforms, all of which contain a signature Y/FIY/FTSGTTG AMP binding motif within a cytoplasmic loop, and they are predicted to contain two extracellular domains, which form an NEFA binding region after dimerization.25 As a group, FATPs are efficient transporters, with Km ranging from 0.3 to 20 μM,30,51,52 which is consistent with our observations for proximal tubule NEFA transport. However, the low- to midmicromolar concentrations used in our experiments represent total concentration, whereas circulating free NEFA concentration is in the low nanomolar range.53 There are two FATP2 splice variants: one with intrinsic acyl CoA synthetase activity (FATP2a), as previously mentioned, and one without acyl CoA synthetase activity (FATP2b); both splice variants are capable of transporting long-chain NEFA.32 Only FATP2a is detectable in kidney,32 which we confirmed by RT-PCR.

Apical NEFA uptake in Slc27a2−/− proximal tubules and cell lines was significantly diminished, although not eliminated, and basolateral NEFA was similar between wild-type and FATP2 knockout mice, implying the presence of other functional NEFA transporters in the proximal tubule. Considering the importance of NEFA β-oxidation to the proximal tubule metabolism, such redundancy is not surprising. One implication of this observation is that, if FATP2 is targeted as a therapy to reduce lipoapoptosis, the residual NEFA uptake by FATP2-independent mechanisms should be sufficient to support proximal tubule survival as evidenced by the viability of Slc27a2−/− cells (Figure 8) and previous observations that Slc27a2−/− mice exhibit no overt renal phenotype.54 We show that FFA1 and CD36 are expressed in kidney, but neither transporter localized to proximal tubule. CD36 has been implicated as a candidate kidney NEFA transporter in humans,33,34 but in mouse studies, it has reliably been shown only in kidney macrophages and inconsistently in proximal tubule.12,33,34,55 FABPpm is expressed in proximal tubule but localizes to mitochondria,56 and it is, therefore, not a likely candidate for NEFA uptake from the apical proximal tubule membrane. FATP1 and FATP4 have been detected in kidney, although at very low levels, and localization within the nephron has not been described. Nevertheless, these FATP isoforms could represent residual proximal tubule NEFA transporters.

In conclusion, intracellular accumulation of NEFA and NEFA metabolites leads to tubular atrophy and CKD progression. Normal proximal tubule metabolism is dependent on basolateral NEFA uptake, which is regulated by an FATP2-independent mechanism, whereas apical NEFA uptake is largely mediated by FATP2. We speculate that apical FATP2-directed NEFA uptake in conjunction with increased NEFA synthesis and decreased NEFA metabolism represent mechanisms of pathologic proximal tubule NEFA accumulation in proteinuric renal disease, such as diabetic nephropathy. Using in vivo, ex vivo, and in vitro techniques, we show that FATP2 is a major apical proximal tubule NEFA transporter, which regulates cytotoxicity. Because of this newly established role as a mediator of lipoapoptosis, we propose that FATP2 deserves attention as a potentially attractive target for the design of treatments for tubular atrophy and CKD progression.

Concise Methods

Animals

Sprague–Dawley rats weighing 100–150 g were purchased from Charles River Breeding Laboratories (Wilmington, MA). Wild-type and 129S-Slc27a2tm1Kds/J (Slc27a2−/−) mice were purchased from Jackson Laboratories (Bar Harbor, ME). All protocols and procedures were approved by the Institutional Animal Care and Use Committee of Case Western Reserve University in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals.

Human Samples

Formalin-fixed, paraffin-embedded normal human kidney sections were obtained from Imgenex Corp. (San Diego, CA). All studies involving human tissues were performed with approval of the institutional review board of the MetroHealth System, Case Western Reserve University.

Cell Lines

LLC-PK1 and HK-2 cell lines were purchased from ATCC (Manassas, VA); HRPT cells were a gift from Lorraine Racusen (Johns Hopkins University). LLC-PK1 and HRPT cells were maintained in DMEM-F12 (Invitrogen Life Technologies, Carlsbad, CA) plus 10% FBS (HyClone) and 1% penicillin/streptomycin-fungizone (Sigma-Aldrich, St. Louis, MO) as described previously.12 HK-2 cells were cultured in Keratinocyte-SFM supplemented with 5 ng/ml EGF and 40 μg/ml bovine pituitary extract (Gibco/Thermo Fisher Scientific, Waltham, MA).

Microperfusion Experiments

Microdissected proximal tubules were perfused with fluorescently labeled NEFA using established methods.26 Animals were anesthetized with ketamine (100 mg/kg body wt intraperitoneally) and xylazine (20 mg/kg body wt intraperitoneally). The abdominal cavity was opened, and the left kidney was superfused twice with ice cold 150 mM sodium chloride; then, it was removed and placed in physiologic saline at 4°C. Coronal slices were cut, and proximal tubule segments were isolated from the kidney cortex using microforceps under a stereomicroscope at 4°C to 10°C. S2 segment tubules ranging from 0.7 to 1.0 mm were transferred to a temperature-regulated chamber and perfused using concentric glass pipettes at 37°C. Tubules were perfused with BODIPY-conjugated 4,4-difuoro-5,7-dimethyl-4-bora-3a,4a-diaza-s-indacene-3-hexadecanoic acid (final concentration 2.5 μM; Invitrogen Life Technologies) complexed with 0.2% fatty acid–free albumin in buffer containing 130 mM NaCl, 2.5 mM NaH2PO4, 4 mM KCl, 1.2 mM MgSO4, 6 mM l-alanine, 1 mM trisodium citrate, 5.5 mM glucose, 2 mM calcium dilactate, and 10 mM Hepes, pH 7.4 at 37°C. The fluorescence detection system was mounted on a Nikon Diaphot inverted microscope (Nikon, Tokyo, Japan). The intracellular BODIPY dye was excited at 490 nm, and emitted fluorescence was measured using a 510-nm dichroic mirror. Images were recorded using a 40× immersion oil objective and a Coolsnap HQ digital camera (Photometrics, Tucson, AZ), and fluorescence measurements were recorded using Metafluor version 7 imaging software (Universal Imaging, Downingtown, PA).

NEFA Uptake in Proximal Tubule Cell Lines

Uptake studies were performed with minor modifications to published methods.57 Cells were cultured to confluence on permeable supports or 12-mm coverslips, which were mounted in a temperature-controlled perfusion chamber of a Leica DMIRE2 inverted microscope stage. Solutions containing BODIPY-conjugated dodecanoic acid (QBT fatty acid uptake assay; Molecular Devices, Sunnyvale, CA; final concentrations of 4 nM, 20 nM, 100 nM, 500 nM, and 2.5 μM) complexed with 0.2% essential fatty acid–free albumin were perfused into the recording chamber. Excitation λ=490-nm pulses were delivered and emission λ=510-nm fluorescence was recorded at 20-second intervals until steady-state uptake was achieved. Epifluorescence was recorded using a SPOT-RT camera (Diagnostic Instruments, Sterling Heights, MI), and images were acquired and analyzed using SimplePCI imaging software (Compix Inc., Cranberry Township, PA). The uptake rate was defined as the maximum slope within the first 10 seconds, which was determined using SigmaPlot 2000 software (SPSS, Chicago, IL).

RT-PCR

Total RNA was extracted from cell lines or mouse kidney cortex using the RNeasy Mini kit (Qiagen, Hilden, Germany) in accordance with the protocol described by the manufacturer. RNA concentrations were determined using the NanoDrop 2000 Spectrophotometer (Thermo Fisher Scientific). Reverse transcription was performed using SuperScript IV VILO Master-mix (Invitrogen Life Technologies) according to the manufacturer’s instructions. Each amplification reaction was conducted in 25-μl volume using Choice Taq DNA Polymerase (Denville Scientific Inc., Holliston, MA) according to the recommended protocol and PCR cycling conditions. PCR products underwent 1% agarose gel electrophoresis, and bands were identified by ethidium bromide staining and photographed. Human FATP2 primer sequences were 5′-GGAGATACATTCCGGTGGAA-3′ (forward) and 5′-TGATCTCAATGGTGTCCTGT-3′ (reverse), yielding a 257-bp amplicon. Human CD36 primer sequences were 5′-GGAACAGAGGCTGACAACTT-3′ (forward) and 5′-TCGCAGTGACTTTCCCAATAG-3′ (reverse), yielding a 347-bp amplicon. Mouse GAPDH primer sequences were 5′-CTGCCATTTGCAGTGGCAAAGTGG-3′ (forward) and 5′-TTGTCATGGATGACCTTGGCCAGG-3′ (reverse); human GAPDH primer sequences were 5′-GTCTTCACCACCATGGAGAAG-3′ (forward) and 5′-GCTTCACCACCTTCTTGATGTCATC-3′ (reverse).

Immunohistochemistry

Methods have been described previously in detail.58 Mouse kidney was frozen at −80°C. Samples were sectioned to 5-μm thickness by cryostat, fixed in paraformaldehyde (4%; 10 minutes at room temperature), rinsed in PBS, and then permeabilized with Triton X-100 (Sigma-Aldrich; 0.2% in PBS, 10 minutes at room temperature). Sections were blocked with rabbit or goat serum (5% in PBS, 1 hour at room temperature) or Mouse on Mouse reagent (Vector, Burlingame, CA). Primary antibodies were rabbit anti-FATP2 IgG (GeneTex, Irvine, CA; 1:50, 16 hours, 4°C), rabbit polyclonal FATP2 antisera31 (1:50, 16 hours, 4°C), rabbit anti-FFA1 IgG (Alamone, Jerusalem, Israel; 1:200, 16 hours, 4°C), mouse anti-CD36 IgA (BD Pharmingen, San Jose, CA; 1:200, 16 hours, 4°C), mouse anti–Na+/K+-ATPase IgG (Santa Cruz Biotechnology, Santa Cruz, CA; 1:100, 16 hours, 4°C), or goat anti-Glut5 IgG (Santa Cruz Biotechnology; 1:100, 16 hours, 4°C). Secondary Alexa Fluor 488 and 568 antibodies were used (Invitrogen Life Technologies; 1:200, 1 hour at room temperature). Proximal tubules were labeled with Texas red–conjugated Tetragonolobus purpureus lectin as previously described.5 Sections were mounted in Vectashield aqueous mounting media containing DAPI nuclear counterstain (Vector) and viewed by confocal microscopy (Leica, Wetzlar, Germany).

Immunoblot Analyses and Immunoprecipitation

Lysates were probed from whole cells or crude membrane fractions. Crude membranes were harvested by incubation with hypotonic buffer (1:10 dilution of PBS with protease inhibitor cocktail; Thermo Fisher Scientific). Lysed cells were homogenized, and large particles were removed by centrifugation at 500×g for 2 minutes. For immunoblots, the supernatant was centrifuged at 14,000×g for 30 minutes, and the resulting pellet was suspended in Laemmli buffer (125 mM Tris, pH 6.8, 2% SDS, and 5% glycerol), assayed for protein concentration by DC protein assay (Bio-Rad, Hercules, CA), and frozen at −80°C.

For biotinylation experiments, PBS-washed cells were incubated with sulfo-NHS-LC-Biotin (Thermo Fisher Scientific; 2 mM, 5 minutes, 4°C), washed again with PBS, and then incubated with glycine quenching buffer (0.1 M, 5 minutes, 4°C). Cells were lysed in 1 ml lysis buffer (25 mM Tris, pH 7.4, 50 mM NaCl, 25 mM NaF, 10% glycerol, and 1% Triton X-100) and centrifuged (14,000×g, 10 minutes, 4°C), and supernatants were assayed for protein concentration and stored at −80°C. Thawed samples of equal protein concentration were mixed with streptavidin agarose beads (Thermo Fisher Scientific; 20 μl bead volume) with gentle rocking for 3 hours at 4°C and then centrifuged (1000×g, 30 seconds, 4°C). The pelleted beads were then washed with lysis buffer at 4°C.

Whole-cell, crude membrane, and streptavidin-precipitated samples were lysed and denatured in boiling SDS-PAGE buffer (125 mM Tris, pH 6.8, 2% SDS, 5% glycerol, 1% β-mercaptoethanol, and 0.003% bromphenol blue) for 5 minutes. Immunoblot methods have been described previously.12 Briefly, samples (20 μg protein per lane) were resolved by SDS-PAGE and transferred to polyvinylidine difluoride membranes. Blots were blocked in 5% nonfat dried milk and probed with anti-FATP2 (GeneTex; 1:500, 16 hours, 4°C) or anticaspase-2 (Abcam; 10 μg/ml, 16 hours, 4°C) IgG and then HRP-conjugated IgG (1:10,000, 1 hour at room temperature). Band intensity was detected by enhanced chemiluminescence. Blots were exposed to stripping buffer (Thermo Fisher Scientific; 10 minutes at room temperature) and then reprobed with anti–α-tubulin (Santa Cruz Biotechnology; 1:3000, 1 hour at room temperature), anti–Na+/K+-ATPase (Santa Cruz Biotechnology; 1:1000, 1 hour at room temperature), or anti-Glut5 (Santa Cruz Biotechnology; 1:500, 1 hour at room temperature) IgG followed by HRP-conjugated IgG (1:10,000, 1 hour at room temperature).

Immunocytochemistry

Methods have previously been described in detail.59 Cells were maintained on sterile glass coverslips within six-well plates, blocked with 5% low-IgG BSA and 0.2% Triton X-100 (Sigma-Aldrich) for 30 minutes at room temperature, incubated with rabbit anti-FATP2 IgG (1:50, 2 hours at room temperature), and then fixed in paraformaldehyde (4%, 10 minutes at room temperature). Postfixed cells were incubated with Alexa Fluor 568 goat anti-rabbit IgG (1:200, 2 hours at room temperature). Coverslips were mounted in antifade, aqueous media containing DAPI (Vectashield; Vector) on standard microscope slides. Images were viewed using a Leica confocal microscope with appropriate fluorescence filters.

shRNA Transfection

To achieve FATP2 knockdown, HRPT cells were transfected with lentiviral shRNA vectors containing a puromycin selection cassette and targeting human SLC27A2 (Ref-Seq NM_003645.3) according to previously described methods.12 Oligonucleotides were purchased from the Mission shRNA Gene Set and cloned into pLKO.1-puro plasmid (Sigma-Aldrich), and lentiviral particles were produced in HEK 293T packaging cells using the ViraPower Expression System (Invitrogen Life Technologies). The viral supernatant was added to HRPT monolayers grown on 10-cm plates, and stably transfected cells were identified for further studies. Three constructs were screened by immunoblotting for silencing of FATP2 expression: ccggCCATACTTCTTCCAGGACATACTCGAGTATGTCCTGGAAGAAGTATGGtttttg (shRNA1), ccggGCTGATTACCTACCTAGTTATCTCGAGATAACTAGGTAGGTAATCAGCtttttg (shRNA2), and ccggCCTATGACTGAGGACATCTATCTCGAGATAGATGTCCTCAGTCATAGGtttttg (shRNA3).

In Vivo Induction of Proteinuria

Proteinuria and tubulointerstitial disease were induced by adapted protein overload8,40 and LPS41,42 protocols. Both protocols used 6- to 8-week-old wild-type and Slc27a2−/− mice on 129S genetic backgrounds. For the protein overload model, the right and left abdomen were alternately cleansed with isopropanol on consecutive days for intraperitoneal injections. Mice received daily (Monday through Friday) sterile injections of endotoxin-free BSA (A-9430; Sigma-Aldrich; 10 mg/g body wt) 5 d/wk for 3 weeks. For the LPS model, mice were injected with endotoxin-free LPS (L-3137; Sigma-Aldrich; 10 mg per mouse) for 3 consecutive days. All mice were euthanized the day after protocol completion. Urine was collected by bladder puncture, and kidneys were harvested for quantitative histomorphometry analyses.

Urine Protein Electrophoresis

To determine albumin excretion, urine samples (10 μl per lane + 2 μl 6× SDS-PAGE buffer) were boiled for 10 minutes, loaded on 10%–20% Tris-glycine gels, and electrophoresed at 150 V. Gels were then stained overnight in solution containing 0.1% Coomassie Brilliant Blue R-250, 50% methanol, and 70% glacial acetic acid. Gels were then destained in 50% methanol + 7% acetic acid solution, and digital images were obtained with a Canon Canoscan LiDE 120 scanner.

Quantitative Histomorphometry

Methods were conducted as previously described.12 Briefly, studies were conducted by two observers blinded to experimental conditions on images viewed at 40× magnification, which were overlaid with a 16×22 grid within Adobe Photoshop (San Jose, CA) or ImageJ to assess for tubular atrophy and interstitial fibrosis. Ten sections per kidney were randomly selected for analysis. Coincidence of intersecting grid lines with tubule (nucleus, cytoplasm, or brush border), tubule lumen, or Masson trichrome–stained interstitium was counted, whereas glomeruli and blood vessels were omitted from calculations. The total of the three compartments was defined as 1.0, and mean values between the two observers for the proportion of the total composed of tubule cells, tubule lumen, or interstitium were compared between experimental and control (saline-injected) mice.

Oil Red O Staining and Quantification

After incubation with palmitic acid (100 μM, 24 hours) complexed with 0.2% albumin, cells were fixed with paraformaldehyde (4%, 10 minutes at room temperature), rinsed with PBS, and stained with the Oil Red O (0.5%; Biovision; 5 minutes at room temperature) as previously described.12 Stained cells were rinsed with PBS and viewed with a Leica Dmi8 inverted light microscope. For quantification, Oil Red O was incubated for 30 minutes and washed with PBS and then 60% isopropanol. Oil Red O was eluted with isopropanol (100%, 10 minutes at room temperature) followed by eluate absorbance measurement at 492 nm with a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific).

TUNEL Assays

Proximal tubule epithelial cell lines were cultured on coverslips and grown to near confluence. Ten to 12 random fields per coverslip were assayed for apoptosis, which was assessed by TUNEL as previously described.59 Nuclei of all cells were counterstained with DAPI, and data are presented as percentage of apoptotic cells.

Statistical Analyses

Unless otherwise noted, all results are expressed as means±SEM. Two-tailed paired t tests were used for statistical analysis between two groups. Two-way ANOVA was used for comparisons between more than two groups. Statistical significance is defined as P≤0.05.

Disclosures

None.

Acknowledgments

Fatty acid transporter-2 antibodies were a gift from Dr. Andreas Stahl (University of California at Berkeley).

This work was supported by National Institutes of Health grants R01 HL128053 (to J.L.G.) and R01 DK067528 (to J.R.S.).

Footnotes

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Kidney Proximal Tubule Lipoapoptosis Is Regulated by Fatty Acid Transporter-2 (FATP2)
Shenaz Khan, Pablo D. Cabral, William P. Schilling, Zachary W. Schmidt, Asif N. Uddin, Amelia Gingras, Sethu M. Madhavan, Jeffrey L. Garvin, Jeffrey R. Schelling
JASN Jan 2018, 29 (1) 81-91; DOI: 10.1681/ASN.2017030314
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