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Acute leptin exposure reduces megalin expression and upregulates TGFβ1 in cultured renal proximal tubule cells
Molecular and Cellular Endocrinology 401 (2015) 25–34
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Molecular and Cellular Endocrinology j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / m c e
Acute leptin exposure reduces megalin expression and upregulates TGFβ1 in cultured renal proximal tubule cells Jessica F. Briffa a,b, Esther Grinfeld a, Michael L. Mathai a, Phillip Poronnik c, Andrew J. McAinch a, Deanne H. Hryciw a,b,* a
Centre for Chronic Disease Prevention and Management, College of Health and Biomedicine, Victoria University, St Albans, Vic. 3021, Australia Department of Physiology, The University of Melbourne, Parkville, Vic. 3010, Australia c School of Medical Sciences, The Bosch Institute, The University of Sydney, NSW 2006, Australia b
A R T I C L E
I N F O
Article history: Received 23 July 2014 Received in revised form 26 November 2014 Accepted 28 November 2014 Available online 2 December 2014 Keywords: Leptin Megalin Proximal tubule TGF-β1 AMPK
A B S T R A C T
Increased leptin concentrations observed in obesity can lead to proteinuria, suggesting that leptin may play a role in obesity-related kidney disease. Obesity reduces activation of AMP-activated protein kinase (AMPK) and increases transforming growth factor-β1 (TGF-β1) expression in the kidney, leading to albuminuria. Thus we investigated if elevated leptin altered AMPK and TGF-β1 signaling in proximal tubule cells (PTCs). In opossum kidney (OK) PTCs Western blot analysis demonstrated that leptin upregulates TGF-β1 secretion (0.50 μg/ml) and phosphorylated AMPKα (at 0.25, and 0.50 μg/ml), and downregulates megalin expression at all concentrations (0.05–0.50 μg/ml). Using the AMPK inhibitor, Compound C, leptin exposure regulated TGF-β1 expression and secretion in PTCs via an AMPK mediated pathway. In addition, elevated leptin exposure (0.50 μg/ml) reduced albumin handling in OK cells independently of megalin expression. This study demonstrates that leptin upregulates TGF-β1, reduces megalin, and reduces albumin handling in PTCs by an AMPK mediated pathway. © 2014 Elsevier Ireland Ltd. All rights reserved.
1. Introduction Obesity rates in Western cultures are rising rapidly, with comorbidities including chronic kidney disease (CKD) and end stage renal failure rising concurrently (Eknoyan, 2007). One of the earliest hallmark characteristics of CKD is the presence of albumin in the urine (albuminuria), which is typically the result of both a glomerular and proximal tubule dysfunction (Takeyama et al., 2011; Wohlfarth et al., 2003; Wolf et al., 2004). The hormone leptin is predominately produced by adipocytes (Vázquez-Vela et al., 2008), with plasma leptin concentrations reflecting adiposity (Garibotto et al., 1998), with higher leptin concentrations seen in obese individuals. Leptin binds to two transmembrane receptors, either the leptin receptor (ObRb) or the scavenger receptor megalin, in a cell specific manner. Megalin is responsible for the transportation of leptin in the hypothalamus, placenta, and renal tubules (Briffa et al., 2014; Hama et al., 2004), whereas ObR is responsible for leptin transport and signaling in most other tissues (Ahima, 2008). Circulating leptin is filtered from the blood by the glomerulus and is reabsorbed by cells of the proximal tubule by the scavenger
* Corresponding author. Department of Physiology, The University of Melbourne, Parkville, Vic. 3010, Australia. Tel.: +61 3 9035 9923; fax: +61 3 8344 5818. E-mail address: [email protected] (D.H. Hryciw). http://dx.doi.org/10.1016/j.mce.2014.11.024 0303-7207/© 2014 Elsevier Ireland Ltd. All rights reserved.
receptor megalin (Briffa et al., 2014; Hama et al., 2004), resulting in negligible leptin secretion in the urine (Meyer et al., 1997), even in obesity (Cumin et al., 1997; Lönnqvist et al., 1997). The plasma concentration of leptin in non-obese individuals is approximately 5.5 ng/ml, with 9.5% being transferred from the blood to the filtrate, with an estimated renal leptin clearance of 0.0595 μg/ml (Garibotto et al., 1998). The serum concentration of leptin in obese individuals is approximately 5 to 10 times higher than normal individuals (Garibotto et al., 1998). This value is disputed, with others determining that the maximum plasma leptin concentration observed in obesity is likely to be 200 ng/ml (Maffei et al., 1995). However, individuals with CKD demonstrate leptinemia at concentrations up to 490 ng/ml (Dagogo-Jack et al., 1998). Albuminuria is a hallmark characteristic of renal dysfunction, with studies demonstrating that exposure to elevated leptin can lead to a significant loss of albumin in the urine (Gunduz et al., 2005), suggesting that leptin may provide a link between CKD and obesity (Briffa et al., 2013). Albuminuria is typically the result of an increase in protein filtration through the glomerulus, caused by basement membrane thickening (Wohlfarth et al., 2003; Wolf et al., 2004), which is further compounded by an impairment in protein endocytosis by the proximal tubule cells (PTCs) which are unable to cope with the increased protein load (Takeyama et al., 2011). Importantly, exposure of the renal tubules to elevated concentrations of albumin induces the production of proinflammatory, profibrotic and vasoactive factors in vitro (Wolf et al., 2004). For example the
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profibrogenic cytokine transforming growth factor beta (TGF-β) is increased in the glomerular filtrate, which upregulates extracellular matrix production, ultimately resulting in basement membrane thickening (Wolf et al., 2004). Interestingly, exposure to elevated leptin has also been shown to increase TGF-β1 secretion in glomerular endothelial cells in vitro (Wolf et al., 1999). Exposure of renal cells to elevated protein and/or leptin also results in an accumulation of collagen in glomerular cells in vitro, which would result in tubulointerstitial fibrosis and basement membrane thickening in vivo (Wohlfarth et al., 2003). These data taken together suggest that leptin may play a role in obesity-related nephropathy by upregulating the expression of fibrotic mediators. Importantly, research by Gekle et al. (2003) has previously identified in opossum kidney (OK) cells, a well established model of the proximal tubule, exposure to 3 μg/ ml of TGF-β1 for 48 h significantly upregulates collagen secretion (types I and IV) and reduces albumin handling by decreasing megalin and cubilin protein expression. In addition to the fibrotic changes obesity causes in the kidney, obesity is also associated with tubular inflammation (Declèves et al., 2011), which further deteriorates kidney function. AMP-activated protein kinase (AMPK), a key regulatory of energy homeostasis, plays important roles in renal inflammation and fibrosis (Declèves et al., 2011; Sharma et al., 2008). Obesity results in a decrease in renal AMPK expression (Declèves et al., 2011), resulting in an upregulation of inflammatory mediators, as well as fibrotic changes to the kidneys (Declèves et al., 2011; Sharma et al., 2008). Specifically, research by Declèves et al. (2011) identified that one week of high fat feeding (60% fat) in six-week old C57BL/6 mice in vivo causes an increase in renal inflammatory markers that preceded albuminuria, with treatment of an AMPK activator preventing the increase in urinary expression of these mediators. Importantly, AMPK activation has been shown to decrease the fibrotic actions of TGF-β1, with an in vivo study identifying that AMPK activation, using Metformin, decreased the renal injury, adipokine expression, and macrophage infiltration that is associated with high fat feeding (60% fat) in six-week old C57BL/6 mice (Kim et al., 2013). In the kidney in vivo and in vitro (OK, LLC-PK1 and glomerular endothelial cells) leptin has been shown to upregulate fibrotic mediators and causes collagen deposition, which may compound renal function (Wohlfarth et al., 2003; Wolf et al., 1999), ultimately resulting in nephropathy. Therefore the overall aim of the current study was to identify if acute exposure to leptin will alter the expression of fibrotic mediators and, specifically, AMPK signaling in PTCs. We also aim to identify the effect leptin has on albumin handling in PTCs, and characterize changes in megalin expression. We hypothesize that exposure of PTCs to elevated leptin will upregulate fibrotic mediator expression and decrease AMPK expression, in addition to reducing albumin handling in PTCs.
2. Materials and methods 2.1. Cell culture We utilized OK cells for this study, as they are a widely used model for the renal proximal tubule as they express all proteins of the megalin signaling complex (Amsellem et al., 2010; Birn and Christensen, 2006; Hryciw et al., 2012). OK cells were maintained in Dulbecco’s Modified Eagle’s Medium and Ham’s F-12 (DMEM/ F12) media (Life Technologies; Victoria, Australia) supplemented with 10% fetal bovine serum (Life Technologies) with 1% penicillin/ streptomycin (Life Technologies), and incubated at 37 °C in 5% CO2. The cells were seeded at confluence and grown for two days in 25 ml flasks. Forty-eight hours prior to experimentation the cells were incubated in DMEM/F12 with 5 mM glucose medium lacking serum.
2.2. Animal care Experimental procedures were approved by the Howard Florey Institute Animal Ethics Committee (AEC 09-050). As used in our previous study (Jenkin et al., 2010), six-week old male Sprague Dawley rats (mean initial body weight approximately 178 g) were housed within individual cages in an environmentally controlled laboratory (ambient temperature 22–24 °C) with a 12 h light/dark cycle (7:00–19:00). Ad libitum access to food and water was maintained. At ten-weeks, rats were deeply anesthetized with sodium pentabarbitone (100 mg/kg; Virbac, Peakhurst, Australia) then euthanized via cardiac puncture. Kidney and white adipose tissues (WAT) were then removed and tissues were stored at −80 °C for subsequent Western blot analysis. 2.3. Immunoprecipitations OK cells were seeded onto 175 cm2 flasks and grown to confluence, and then serum starved for 48 h. The cells were then treated with either control (phosphate buffered saline: PBS) or human recombinant leptin (ProSpec; New Jersey, USA) for 15 min at 0.05 or 0.50 μg/ml. Protein was then isolated from OK cells using immunoprecipitation (IP) lysis buffer (10 mM Tris–HCl, 150 mM NaCl, 5% NP-40 with the pH adjusted to 7.5) supplemented with a Complete Mini Protease Inhibitor Cocktail (Roche; NSW, Australia). Immunoprecipitations (IPs) were then performed on 1 mg protein from each treatment. Cells were incubated overnight at 4 °C with no antibody, Normal Rabbit IgG (Merck Millipore; Victoria, Australia), Megalin antibody (ThermoFisher Scientific; Victoria, Australia), or Leptin antibody (Enzo Life Sciences; New York, USA) with endto-end rotation. The next day 50 μl pre-cleared Protein G Agarose (ThermoFisher Scientific) was added to each treatment and spun with end-to-end rotation at 4 °C for 5 h. The eppendorf tubes were then centrifuged and the supernatant was discarded, and the beads were washed with IP lysis buffer. Fifty microliters Laemmli Sample Buffer was then added to each treatment, and heated to 100 °C for 10 min. The treatments were then centrifuged and Western blots were performed on the supernatant. Equal aliquots (20 μl) of each treatment were separated on a 7.5% (Leptin IP) or 4–20% SDSPAGE gel (Megalin IP) (Bio-Rad; NSW, Australia) and transferred onto a nitrocellulose membrane. The membranes were then probed with either Leptin (Megalin IP) or Megalin (Leptin IP) antibodies to identify if leptin and megalin bind. 2.4. ‘Real-time’ polymerase chain reaction (PCR) OK PTCs were serum starved for 48 h (Briffa et al., 2014; Hryciw et al., 2006), then treated with human recombinant leptin (ProSpec) for 2 h at 0.05, 0.10, 0.25, and 0.50 μg/ml (Briffa et al., 2014). RNA was then extracted using TRIzol reagent according to the manufacturer’s instructions (Life Technologies) as described previously (Jenkin et al., 2010). The RNA was DNase treated, first strand cDNA was then generated from 0.5 μg of template RNA using the commercially available iScript™ cDNA synthesis kit (Bio-Rad) using random hexamers and oligo dTs as described previously (Jenkin et al., 2010). ‘Real-time’ PCR was conducted using MyiQ™ single color ‘realtime’ PCR detection system (Bio-Rad) with iQ™ SYBR Green Supermix (Bio-Rad) as the fluorescent agent. Forward and reverse oligonucleotide primers for megalin (forward 5′ TGC CCC ACC CGT TAT CCT A 3′ and reverse 5′ ACA GAC ATG GTT CTT ACA CTC AAA CAT 3′; Accession Number XM_007494924.1) and TGF-β1 (forward 5′ CCT GGA CAA CCA GTA CAG CA 3′ and reverse 5′ TTC CGG CCC ACA TAG TAG AC 3′; Accession Number XM_007491983.1) were designed using the OligoPerfect™ Suite (Life Technologies). Selective gene homology was confirmed using BLAST analysis (National Centre for Biotechnology Information; Maryland, USA). To compensate for
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variations in RNA input amounts and reverse transcriptase efficiency, mRNA abundance of the genes of interest were normalized to the housekeeping gene cyclophilin (forward 5′ CGG GTC ACT TTC GAG CTC TTT 3′ and reverse 5′ CTC AGA GCC CGG AAG TTT TCT 3′) (Jenkin et al., 2010). ‘Real-time’ PCR reactions were run for 50 cycles of 95 °C for 15 s and 60 °C for 60 s. Relative changes in mRNA abundance was quantified using the 2-ΔΔCT method as previously detailed (Jenkin et al., 2010) and reported in arbitrary units.
2.5. Protein extraction and Western blotting To prepare the tissue lysates, 50 mg of kidney was lysed in 500 μl HNT buffer (20 mM HEPES, 120 mM NaCl, 5 mM EDTA, 0.6% Triton-X with the pH adjusted to 7.5) and 100 mg WAT was lysed in 100 μl lysis buffer (50 mM Tris–HCl, 120 mM NaCl, 1% NP-40 with the pH adjusted to 7.5) with protease and phosphatase inhibitors (Cell Signaling Technology; Massachusetts, USA). To clarify the tissue lysates, the kidney lysate was centrifuged at 10,000× g for 5 min at 4 °C and the WAT lysate was centrifuged at 12,000× g for 20 min at 4 °C with the centrifugation occurring three more times on the supernatant to remove the lipids. In addition, OK PTCs were serum starved for 48 h in minus media, then treated with a vehicle control (PBS) or human recombinant leptin for 2 h (0.05–0.50 μg/ml). Protein was then isolated from OK cells using IP lysis buffer supplemented with a Complete Mini Protease Inhibitor Cocktail and Halt Phosphatase Inhibitor Cocktail (ThermoFisher Scientific). Equal aliquots (50 μg) of the protein samples were separated on a 7.5% (megalin) or 4–20% SDS-PAGE gel and transferred onto a nitrocellulose membrane. Membranes were probed with AMPKα, phosphorylated AMPKα (p-AMPKα), AMPKβ and phosphorylated AMPKβ (p-AMPKβ) antibodies (Cell Signaling Technology), leptin antibody (Enzo Life Sciences), collagen type IV and TGF-β1 antibodies (Abcam; Massachusetts, USA) or Megalin antibody (ThermoFisher Scientific) were used following protocols described previously (Hryciw et al., 2006; Slattery et al., 2011). Densitometric analysis of the banding was performed using ImageJ software. In addition, cells were pre-treated with either 234.6 nM of the AMPK inhibitor Compound C (Merck Millipore), according to the manufacturer’s IC50 and previous research (Hao et al., 2010), or a DMSO control (Sigma-Aldrich; NSW, Australia) for 60 min. After the pretreatment the cells were then treated with human recombinant leptin (0.05–0.50 μg/ml) in the presence of 234.6 nM Compound C or DMSO and PBS (control treatment) for 2 h. Western blots were then performed as described above and probed for Megalin and collagen type IV.
2.6. TGF-β1 ELISA OK cells were seeded onto 6-well plates and serum starved for 48 h prior to experimentation. The cells were then treated with human recombinant leptin (0.05–0.50 μg/ml) for 2 h. Following treatment, the cell culture media was collected and centrifuged, and a protein assay was performed on the cell lysate. The cell culture media was treated with 1 M HCl for 15 min, and then neutralized with 1 M NaOH. Total TGF-β1 concentrations were determined using an enzyme linked immunoassay assay (ELISA) (Promega; NSW, Australia) as per the manufacturer’s instructions. The absorbance was read at 450 nm on the xMark Microplate Absorbance Spectrophotometer. Secreted TGF-β1 concentrations were standardized to the protein content of the cells, and are presented as concentrations normalized to total cellular protein. In addition cells were exposed to Compound C (as described in section 2.5), and a TGF-β1 ELISA was performed as described above.
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2.7. Texas Red (TR) and Dye Quenched (DQ) albumin uptake We used standard albumin uptake methods in OK cells, a cell line commonly used for the study of proximal tubule albumin uptake (Hryciw et al., 2005, 2006). OK cells were seeded onto 48-well plates and treated with human recombinant leptin (0.05–0.50 μg/ml) for 2 h. To determine albumin uptake, cells were exposed to 50 μg/ml Texas Red albumin (TR-albumin) (Life Technologies) and treated with their respective leptin concentrations (0.05–0.50 μg/ml) for 2 h. To determine albumin degradation, OK cells were exposed to 50 μg/ ml Dye Quenched albumin (DQ-albumin) (Life Technologies) and treated their respective leptin concentrations (0.05–0.50 μg/ml) for 2 h. Nonspecific binding was determined in cells exposed to TR- or DQ-albumin for 1 min. At the end of the uptake period, cells were washed in HEPES-Ringer buffer (122.5 mM NaCl, 5.4 mM KCl, 1.2 mM CaCl2, 0.8 mM MgCl2, 0.8 mM Na2HPO4, 0.2 mM NaH2PO4, 5.5 mM Glucose, 10 mM HEPES; pH 6 at 4 °C), then lysed in MOPS buffer (20 mM MOPS with 0.1% Triton X-100) for 30 min at 37 °C. The TRalbumin fluorescence was determined using the POLARstar Galaxy (Biodirect; Massachusetts, USA) at 580 nm excitation and 630 nm emission wavelengths. The DQ-albumin fluorescence was determined using the POLARstar Galaxy at 590 nm excitation and 620 nm emission wavelengths. TR-albumin and DQ-albumin uptake were standardized to total cellular protein, and the amount of fluorescence per microgram of cellular protein was adjusted for background. As AMPK plays a protective role in mitigating the fibrotic actions of TGF-β1 (Kim et al., 2013), we repeated the TR-albumin uptake assay using Compound C. OK cells were exposed to Compound C (as described in section 2.5); however, the cells were only treated with 0.50 μg/ml human recombinant leptin (the concentration that significantly reduced albumin uptake and endocytosis) and the TR-albumin uptake was then performed as described above. 2.8. Proteasomal activity assay To determine if leptin alters proteasomal function, we next performed a proteasomal activity assay using methods previously published (Hryciw et al., 2004). Briefly, OK cells were incubated with varying concentrations (0.05–0.50 μg/ml) of human recombinant leptin for 2 h. To determine proteasomal activity, 75 μM N-succinylLeu-Leu-Val-Tyr-7-amino-4-methylcoumarin (LLVY-AMC) (Enzo Life Sciences) was added to 250 μg of protein from each treatment and topped to a final volume of 200 μl. The reaction mixture was incubated at 30 °C for 60 min. The fluorescent intensities of each sample were measured on the POLARstar Galaxy at excitation and emission wavelengths of 360 nm and 460 nm respectively, and were adjusted for background. The proteasome inhibitor Z-Leu-Leu-Leual (MG132) (Sigma-Aldrich) was used as a negative control (Hryciw et al., 2004). 2.9. Statistical analysis Statistical analysis of all data was conducted using a one-way ANOVA with Tukey’s post-hoc test or a student’s t-test. All data are expressed as the mean ± the standard error of the mean (SEM), with a P value of less than 0.05 being considered statistically significant. 3. Results 3.1. Leptin binds to megalin in the proximal tubule Previous research in Sprague-Dawley rats has determined that megalin is the main receptor responsible for leptin uptake in the convoluted segment of the proximal tubule, with no leptin binding to ObR in the straight segment of the proximal tubule (Hama et al.,
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tubule. ‘Real-time’ PCR analysis identified that OK cells exposed to 0.50 μg/ml leptin significantly increased TGF-β1 mRNA expression (0.1211 ± 0.024 compared to control, n ≥ 3, P < 0.05; Fig. 3A). Interestingly, 0.50 μg/ml leptin mRNA expression was significantly different to all other leptin treatments (0.1211 ± 0.024 compared to 0.0486 ± 0.006, 0.0411 ± 0.004 and 0.0569 ± 0.009 for 0.05, 0.10 and 0.25 μg/ml leptin respectively, n ≥ 3, P < 0.05; Fig. 3A). Western blot analysis identified a significant decrease in mature TGF-β1 protein expression at the same leptin concentration (0.50 μg/ ml) (42.83 ± 3.1% compared to control, n = 3, P < 0.05; Fig. 3B and C) indicating that there may be an increase in leptin secretion at this concentration. ELISA analysis identified that leptin, at the same concentration (0.50 μg/ml), significantly increased TGF-β1 secretion from OK cells (0.2289 ± 0.049 compared to control respectively, n ≥ 4, P < 0.05; Fig. 3D). Thus, leptin increases the secretion of TGF-β1 from PTCs, which may lead to the tubulointerstitial injury observed in CKD. Research has previously shown that AMPK activation can regulate the fibrotic actions of TGF-β1 (Kim et al., 2013). To investigate this association, OK cells were pretreated with Compound C to inhibit AMPK, and then treated with leptin and Compound C. ELISA analysis identified that AMPK inhibition significantly decreased Fig. 1. Leptin binds to megalin in vitro. (A) Immunoprecipitations indicate that megalin binds to leptin in vitro, and that leptin binds to megalin in vitro (n = 2). (B) Control Western blots indicate that opossum kidney (OK) cells are capable of producing leptin (50 μg OK lysate). White adipose tissue (WAT) lysate (12.5 μg), kidney lysate (12.5 μg) and human recombinant leptin (2 μg) were used as positive controls.
2004). Further, we have previously shown that OK cells lack ObR expression (Briffa et al., 2014), demonstrating that leptin interacts with megalin in OK cells. To confirm an interaction, we first investigated if leptin and megalin bind via immunoprecipitation. Leptin and megalin IPs identified that multiple leptin molecules bind in OK cells (n = 2; Fig. 1A) at 0.05 and 0.50 μg/ml. Interestingly OK control IPs (not treated with leptin) were also positive for megalin and leptin binding, suggesting that OK PTCs may be capable of producing leptin. To investigate if OK cells express leptin we next performed a Western blot on OK lysate, kidney lysate, WAT lysate (both of which are known to express leptin) and human recombinant leptin (as a positive control) (ProSpec). Western blot analysis identified that OK cells do express leptin protein; however, it appears to have a higher molecular weight than rodent and human leptin (Fig. 1B), suggesting there is a species variation in leptin’s molecular weight. 3.2. Short term exposure of OK cells to leptin alters AMPK expression We investigated if leptin treatment alters the expression of AMPK in OK cells. Western blot analysis identified that leptin upregulates p-AMPKα at 0.25 and 0.50 μg/ml (149.16 ± 13.5% and 198.31 ± 10.7% compared to control respectively, n ≥ 3, P < 0.05; Fig. 2A and B). Interestingly, 0.50 μg/ml leptin was also significantly increased compared to all other leptin treatments (198.31 ± 10.7% compared to 118.34 ± 5.6%, 126.7 ± 20.83% and 149.16 ± 13.5% for 0.05, 0.10, and 0.25 respectively, n ≥ 3, P < 0.05; Fig. 2A and B). In contrast leptin did not alter p-AMPKβ expression (Fig. 2A and B). 3.3. Short term exposure of OK cells to leptin alters TGF-β1 expression and secretion by an AMPK mediated pathway As previous research has well established that exposure to elevated leptin upregulates TGF-β production and secretion in the glomerulus (Wolf et al., 1999), we next investigated the effect leptin has on TGF-β1 protein production and secretion in the proximal
Fig. 2. Leptin alters the expression of AMPK in OK cells. (A) In opossum kidney (OK) cells exposed to leptin for 2 h, phosphorylated AMPKα (p-AMPKα) protein was increased at 0.25 and 0.50 μg/ml. OK cells treated with 0.50 μg/ml leptin significantly altered p-AMPKα compared to all other leptin treatments. (B) Densitometric analysis confirmed the significant increase of p-AMPKα at 0.25 and 0.50 μg/ml (n ≥ 3, P < 0.05 vs. control) and a significant increase in p-AMPKα at 0.50 μg/ml compared to 0.05, 0.10 and 0.25 μg/ml leptin (n ≥ 3, P < 0.05 vs. 0.05, 0.10 and 0.25 μg/ml leptin). Densitometry analysis also identified no changes in p-AMPKβ expression in response to leptin. Significant differences between treatments are indicated by letters that differ (a, b, c); for example “a” is different from “b”, but not different from “ab”.
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Fig. 3. Leptin increases TGF-β1 mRNA transcription and TGF-β1 secretion in OK cells by an AMPK mediated pathway. (A) ‘Real-time’ PCR analysis identified that opossum kidney (OK) cells exposed to 0.50 μg/ml of leptin for 2 h significantly increased TGF-β1 mRNA transcription (n ≥ 3, P < 0.05 vs. control). Additionally, 0.50 μg/ml leptin mRNA expression of TGF-β1 was significantly greater than all other leptin concentrations (n ≥ 3, P < 0.05 vs. 0.05, 0.10 and 0.25 μg/ml leptin). (B) Western blot analysis identified a significant decrease in mature TGF-β1 protein in OK cells treated with 0.50 μg/ml leptin. (C) Densitometry analysis confirmed the significant decrease in mature TGF-β1 protein at 0.50 μg/ml leptin (n = 3, P < 0.05 vs. control). (D) ELISA analysis identified that OK cells exposed to leptin for 2 h significantly increased TGF-β1 secretion at 0.50 μg/ ml (n ≥ 4, P < 0.05 vs. control). (E) Pretreatment with Compound C resulted in a significant decrease in TGF-β1 being secreted from OK PTCs at 0.10, 0.25, and 0.50 μg/ml leptin (n ≥ 5, P < 0.05 vs. control), as well as compared to physiological (0.05 μg/ml) leptin (n ≥ 4, P < 0.05 vs. 0.05 μg/ml leptin). Significant differences between treatments are indicated by letters that differ (a, b); for example “a” is different from “b”, but not different from “ab”.
TGF-β1 secretion from OK cells at 0.10, 0.25, and 0.50 μg/ml leptin (0.0256 ± 0.004, 0.0298 ± 0.007 and 0.0222 ± 0.007 compared to control respectively, n ≥ 5, P < 0.05 vs. control; Fig. 3E). Importantly, these concentrations were significantly different compared to physiological (0.05 μg/ml) leptin concentrations (0.0256 ± 0.004, 0.0298 ± 0.007 and 0.0222 ± 0.007 compared to 0.0779 ± 0.012 for 0.05 μg/ml leptin, n ≥ 5, P < 0.05 vs. 0.05 μg/ml leptin; Fig. 3E), indicating that if not for the increased p-AMPKα expression leptin would not cause the increase in TGF-β1 secretion. These findings suggest, in the renal proximal tubule, that leptin increases AMPK activation, which positively regulates TGF-β1 expression. 3.4. Short term exposure of OK cells to elevated concentrations of leptin does not alter collagen type IV expression As TGF-β1 is known to upregulate collagen type IV production (Gekle et al., 2003; Grande et al., 2002), we next examined if leptin upregulates collagen type IV protein expression in cells of the proximal tubule. OK
cells exposed to leptin did not change collagen IV production (Fig. 4A and B). This finding is not surprising as the cells were only treated with leptin acutely (for 2 h), with previous research showing that prolonged exposure (48 h) of OK cells to TGF-β1 (3 μg/ml) upregulates Collagen IV protein expression (Gekle et al., 2003). As research has shown that AMPK attenuates the fibrotic changes brought on by TGF-β1 (Lee et al., 2013), we also investigated the effect AMPK inhibition has on collagen type IV expression. Western blot analysis identified that AMPK inhibition did not alter collagen type IV protein expression (compared to control) (n ≥ 3; Fig. 4C and D). 3.5. Short term exposure of OK cells to leptin decreases megalin protein expression by an AMPK mediated pathway Elevated TGF-β1 concentrations have been previously shown to decrease megalin protein expression in OK cells (Gekle et al., 2003). As leptin upregulates TGF-β1 secretion, we next examined the effect leptin has on megalin mRNA and protein expression. ‘Real-time’ PCR
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Fig. 4. Leptin does not alter collagen type IV production in OK cells. (A) In opossum kidney (OK) cells exposed to leptin for 2 h, collagen type IV protein expression did not change. (B) Densitometric analysis confirmed no change in collagen IV protein expression. (C) Western blot analysis identified that AMPK inhibition did not alter collagen type IV protein production. (D) Densitometry analysis confirmed that AMPK inhibition did not alter collagen type IV protein production.
Fig. 5. Leptin decreases megalin expression in OK cells by an AMPK mediated pathway. (A) In opossum kidney (OK) cells exposed to leptin for 2 h, there was no change in megalin mRNA transcription. (B) OK cells exposed to leptin for 2 h significantly decreased megalin expression across all leptin treatments (0.05–0.50 μg/ml). (C) Densitometric analysis confirmed the significant reduction in megalin protein across all leptin treatments (n = 4, P < 0.05 vs. control). (D) Western blot analysis identified that AMPK inhibition with Compound C restores megalin protein expression to values similar to control. (E) Densitometry analysis confirmed no change in megalin protein expression compared to control (n ≥ 5). (F) In OK cells, exposure to leptin for 2 h does not alter proteasomal function. The proteasome inhibitor Z-Leu-Leu-Leu-al (MG132) was used as a negative control, and significantly decreased proteasome function (n = 6, P < 0.05 vs. control). Significant differences between treatments are indicated by letters that differ (a, b); for example “a” is different from “b”, but not different from “ab”.
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Fig. 6. OK cells exposed to elevated leptin reduces albumin uptake and degradation by an AMPK mediated pathway. (A) Confluent monolayers of opossum kidney (OK) cells were incubated with 50 μg/ml of Texas Red albumin (TR-albumin) and leptin for 2 h. At 0.50 μg/ml of leptin, albumin uptake was significantly reduced compared to control (n = 4, P < 0.05 vs. control). Additionally, OK cells treated with 0.50 μg/ml leptin significantly reduced albumin uptake compared to all other leptin treatments (n = 4, P < 0.05 vs. 0.05, 0.10 and0.25 μg/ml leptin). (B) Confluent monolayers of OK cells were incubated with 50 μg/ml of Dye Quenched albumin (DQ-albumin) and leptin for 2 h. There was a significant decrease in albumin degradation, as measured by the DQ-albumin assay, in cells treated at 0.50 μg/ml of leptin (n = 4, P < 0.05 vs. control) which coincides with the reduction in albumin endocytosis. Additionally, OK cells treated with 0.50 μg/ml leptin significantly reduced albumin degradation compared to all other leptin treatments (n = 4, P < 0.05 vs. 0.05, 0.10 and0.25 μg/ml leptin). (C) Confluent monolayers of OK cells were treated with either Compound C or DMSO and 0.50 μg/ml leptin, with 50 μg/ml of TR-albumin for 2 h. AMPK inhibition resulted in a significant increase in albumin endocytosis in OK cells compared to cells treated with DMSO and leptin (n = 4, P < 0.05 vs. control). Significant differences between treatments are indicated by letters that differ (a, b); for example “a” is different from “b”, but not different from “ab”.
analysis identified that acute leptin exposure does not alter the expression of megalin mRNA (n ≥ 5; Fig. 5A). However, Western blot analysis identified that leptin significantly decreases megalin protein expression across all concentrations (55.7 ± 2.26%, 68.09 ± 7.25%, 64.59 ± 8.17% and 58.65 ± 9.16% compared to control respectively, n = 4, P < 0.05 vs. control; Figs. 4C and 5B). We demonstrated above that AMPK inhibition significantly decreases TGF-β1 secretion from OK cells compared to control (Fig. 3E), and as research has previously shown that TGF-β1 reduces megalin expression (Gekle et al., 2003), we next investigated the effect AMPK inhibition has on megalin protein expression in response to leptin. Western blot analysis identified that AMPK inhibition restored megalin protein expression to values similar to control (n ≥ 5; Fig. 5D and E). To date, the entire genome of the opossum has not yet been characterized, and as such the exact molecular weight of megalin in opossums is not known. However, the molecular size of our megalin product is similar to previous studies, which have demonstrated that opossum megalin is greater than 250 kDa (Labeau et al., 2001; Zou et al., 2004). 3.6. Short term exposure of OK cells to leptin does not alter proteasomal function As elevated albumin alters proteasomal activity in tubular cells (Hryciw et al., 2004), we also investigated if leptin alters the proteasomal pathway in OK cells. Proteasomal analysis identified that leptin (across all concentrations) does not alter the proteasomal activity in PTCs (n ≥ 6; Fig. 5F). The proteasome inhibitor MG132 was used as a negative control.
3.7. Short term exposure of OK cells to leptin alters PTC handling of albumin by an AMPK mediated pathway As TGF-β1 reduces albumin handling in PTCs (Gekle et al., 2003), we next investigated if leptin alters albumin handling in the proximal tubule. In OK cells exposed to 50 μg/ml of TR-albumin, in the presence of leptin, there was a significant reduction in albumin uptake in cells exposed to 0.50 μg/ml leptin (71.37 ± 5.2% compared to control, n = 4, P < 0.05 vs. control; Fig. 6A). Importantly, 0.50 μg/ml leptin reduced albumin endocytosis compared to all other leptin treatments (71.37 ± 5.2% compared to 92.1 ± 6.00%, 102.15 ± 3.50% and 87.13 ± 2.55% for 0.05, 0.10 and 0.25 μg/ml leptin respectively, n = 4; P < 0.05 vs. 0.05, 0.10 and 0.25 μg/ml leptin; Fig. 6A). Not surprisingly, in OK cells exposed to 50 μg/ml of DQalbumin, in the presence of leptin, there was also a significant reduction in albumin degradation in cells exposed to 0.50 μg/ml leptin (68.5 ± 2.72% compared to control, n = 4, P < 0.05 vs. control; Fig. 6B). Additionally a reduction in albumin degradation was observed at 0.50 μg/ml leptin compared to all other leptin treatments (68.5 ± 2.72% compared to 90.75 ± 4.87%, 85.00 ± 4.65% and 86.75 ± 3.97% for 0.05, 0.10 and 0.25 μg/ml leptin respectively, n = 4; P < 0.05 vs. 0.05, 0.10 and 0.25 μg/ml leptin; Fig. 6B). These data indicate that at elevated concentrations of leptin, albumin uptake and degradation are altered in the proximal tubule which is independent of changes in megalin receptor expression (Fig. 5B and C). To identify the role AMPK plays in regulating albumin endocytosis we repeated this experiment at 0.50 μg/ml leptin to identify if AMPK inhibition restores albumin endocytosis in response to leptin.
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TR-albumin uptake analysis identified that AMPK inhibition results in a significant increase in albumin uptake in OK cells compared to cells treated with a DMSO control and leptin (136 ± 10.88%, compared to control, n = 4, P < 0.05 vs. control; Fig. 6C). 4. Discussion This study describes the potentially significant role for the adipokine leptin in the renal dysfunction that is associated with obesity. Importantly, we have demonstrated that 1) the proximal tubule is capable of producing leptin, 2) leptin significantly upregulates TGF-β1 expression via an AMPK mediated pathway, 3) exposure to elevated leptin concentrations significantly reduces albumin handling of PTCs, and finally 4) the reduction in albumin handling that leptin causes is due to altered megalin function, not protein expression levels. We have previously shown that OK cells lack ObR expression (Briffa et al., 2014), which suggests that any alterations in signal transduction pathways are due to megalin-mediated signaling. However, there has been limited research investigating the ability of megalin to act as a signal transducer. Importantly, we have demonstrated that PTCs exposed to elevated leptin activates the mTOR signaling pathway (Briffa et al., 2014), TGFβ1 and AMPK (this study), which supports a role for megalin-mediated cell signaling in renal pathophysiology. Research by Zou et al. (2004) demonstrated that vitamin D binding protein (another megalin ligand) activates cell signaling pathways in OK cells in vitro through the cleavage of the intracellular carboxyl terminus of the receptor and activation of specific intracellular targets. Recent research in L2 rat yolk sac cells identified that megalin is trafficked to the endocytic recycling compartment for proteolytic processing and the modulation of cellular gene expression for the activation of signal transduction pathways (Shah et al., 2013). However, no studies thus far have demonstrated that albumin can activate megalin cell signaling pathways, and importantly the affinity for megalin binding to both endogenous and exogenous leptin has not been determined, as such, future studies are required to determine this association. In this study we chose to use several different concentrations of leptin (0.05–0.50 μg/ml) to investigate its effect on the renal proximal tubule. Despite the maximal concentration of leptin in the glomerular filtrate being ~47 ng/ml, assuming a normal glomerular barrier, this is likely to be an underestimate due to the following observations. Firstly, research in rabbit proximal tubules have identified that horseradish peroxidase (44 kDa) is transported from the basolateral membrane (Nielsen and Christensen, 1985), suggesting that if leptin is able to be transported from the basolateral membrane to the apical membrane in the renal proximal tubule, then the filtrate concentration of leptin may be significantly higher. More importantly, recent research has shown that kidney fibroblasts are capable of producing and secreting leptin in conditions reflecting renal fibrosis (Lin et al., 2011); however, if PTCs are capable of secreting leptin it is likely to be due to another mechanism as they are epithelial cells. Therefore, if PTCs are able to secrete leptin, especially under conditions of glomerular damage (Lin et al., 2011), then the concentration of leptin in the filtrate in these conditions is likely to be greater than our estimate of ~47 ng/ml. Importantly, we have shown that OK cells can produce leptin protein, however the molecular weight appears to be larger (closer to 20 kDa) compared to human and rodent leptin (~16 kDa). These differences in molecular weight are likely due to a species variation, as based on the predicted protein sequence of the opossum leptin protein (Accession Code – XP_007504322) the molecular weight has a predicted size of 22 kDa. Previous studies have shown that leptin upregulates fibrotic mediator expression. Specifically, leptin has been shown to upregulate TGF-β1 expression and collagen production both in vitro (glomerular endothelial cells) and in vivo (Wolf et al., 1999), which contributes to tubulointerstitial fibrosis. Research in leptin deficient mice (ob/ ob) that underwent unilateral urethral obstruction surgery has shown
a significant reduction in TGF-β mRNA and reduced fibrotic changes to the kidney compared to leptin-receptor deficient mice (db/db) and control animals, identifying that leptin is a co-activator of TGFβ1 (Kümpers et al., 2007). In support of this, glomerular endothelial cells treated with leptin (0.62–625 nM) for 48 h have a significant increase in TGF-β mRNA expression and secretion (Wolf et al., 1999). Additionally, leptin treatment (100 ng/ml) increased fibrotic mediator expression in glomerular mesenchymal cells (Cui et al., 2013). Specifically, TGF-β1 (6 h treatment), Collagen IV (24 h treatment), and Fibronectin (6 and 24 h treatment) are all increased with leptin treatment (Cui et al., 2013). Furthermore, leptin infusion in Wistar rats in vivo for 72 h or three weeks results in a significant increase in renal TGF-β mRNA at both time points, and an increase in collagen type IV mRNA at three weeks of leptin infusion (Wolf et al., 1999). Our data in the renal proximal tubule, with acute leptin treatment, support these findings, showing that leptin treatment significantly increases TGF-β1 secretion from OK cells; in addition to increasing TGF-β1 mRNA and decreasing TGF-β1 mature protein expression at significantly elevated concentrations (0.50 μg/ml). However, it is not surprising that our study did not find an increase in collagen type IV protein. Our study employed an acute leptin treatment. These findings are supported by the study by Wolf et al. (1999), which demonstrated that TGF-β1 expression increases prior to changes in collagen expression. Importantly, research has well established that TGF-β1 regulates collagen type IV production following 48 h of treatment (Gekle et al., 2003). In addition to leptin altering TGF-β1 expression, we also identified that leptin significantly upregulates p-AMPKα expression, with research well supporting a role for AMPK in inhibiting the fibrotic actions of TGF-β1 (Lee et al., 2013). Importantly, obesity is associated with reduced AMPK expression, a mediator that plays important roles in renal inflammation and fibrosis (Declèves et al., 2011; Sharma et al., 2008). In support of this, research by Declèves et al. (2011) demonstrated that C57BL/6 mice fed a high fat diet (60% fat) for one week in combination with AICAR, an AMPK activator, reduced renal hypertrophy and fibrotic mediator expression (H2O2 and monocyte chemotactic protein1). Surprisingly, we identified in conditions reflecting obesity (i.e. high leptin and low AMPK expression) in PTCs that there is a significant reduction in TGF-β1 secretion from OK cells at 0.10–0.50 μg/ml leptin. These differences may arise due to the receptor that leptin is signaling from, with leptin presumably signaling via ObRb in the glomerulus (Wolf et al., 1999), whereas in the proximal tubule leptin signals via megalin (Briffa et al., 2014; Hama et al., 2004). Collectively these findings suggest in renal PTCs, acute leptin treatment with elevated concentrations of leptin, activates AMPK, which upregulates TGF-β1 expression, which in turn may upregulate collagen type IV production with prolonged exposure, ultimately leading to fibrosis (Fig. 7). Research has previously identified that TGF-β1 reduces megalin receptor expression and albumin handling in the renal tubules (Gekle et al., 2003). Specifically, OK cells treated with TGF-β1 (0.3–10 ng/ml) for 48 h have a significant reduction in albumin endocytosis (Gekle et al., 2003). With additional research in OK cells identifying that 48 h exposure to 3 μg/ml TGF-β1 reduced albumin binding capacity at the apical membrane by reducing both megalin and cubilin expression, resulting in a reduction in albumin degradation (Gekle et al., 2003). In the current study we make the important observation that short-term exposure of OK cells to leptin significantly reduces megalin protein expression (0.05–0.50 μg/ml), although albumin endocytosis and degradation were only reduced at significantly elevated concentrations of leptin (0.50 μg/ ml), which is independent of changes in proteasomal function. This suggests that altered function of megalin, but not abundance of the protein, is involved in the renal pathophysiology associated with obesity. Leptin infusion in mice clearly results in albuminuria (Gunduz et al., 2005), with data from our study indicating that leptin directly reduces albumin handling in the proximal tubule. These findings are in contradiction with in vivo models, which clearly shows that leptin can
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References
Fig. 7. AMPKs role in activating fibrotic mediators. A proposed model of how AMPK is modulating fibrotic mediator expression by leptin in the renal proximal tubule. At elevated leptin concentrations, leptin activates AMPK which regulates TGF-β1 expression in renal proximal tubule cells, which may in turn upregulate collagen type IV production; ultimately resulting in fibrosis. Importantly, an upregulation of secreted TGF-β1 decreases megalin receptor expression and function, which impairs albumin uptake and degradation in the renal proximal tubule. Solid lines indicate observed changes seen in this study, with dashed lines indicating hypothesized changes (based on previous studies) due to a prolonged exposure of leptin.
attenuate albumin excretion in humans with lipodystrophy (Ebihara et al., 2007; Javor et al., 2004) and animals with diabetic nephropathy (Suganami et al., 2005), ultimately abolishing their albuminuria and proteinuria. However, under disease states, additional mediators may be affecting albumin handling in addition to leptin. Interestingly, AMPK inhibition restored megalin protein expression and resulted in a significant increase in albumin endocytosis, which supports the study by Gekle et al. (2003). As shown in Fig. 7 in the renal proximal tubule, leptin increasing TGF-β1 protein expression and secretion (via AMPK activation) decreases megalin receptor expression and function, which impairs albumin uptake and degradation in renal PTCs. Thus, leptin may reduce albumin uptake in pathophysiological conditions via fibrotic damage, which has been suggested in other nephropathy models before (Wohlfarth et al., 2003). In summary, this study is the first to identify that leptin upregulates TGF-β1 expression, and alters albumin processing in renal PTCs. Importantly this study provides novel data showing, in PTCs, that AMPK is mediating the expression of TGF-β1 which is altering the function of megalin, ultimately disrupting albumin handing. Thus altering leptins’ effects on AMPK within the PTC appears to warrant further investigation as a possible treatment target to reduce obesity associated renal damage. Acknowledgments This research was supported by the Helen McPherson Smith Trust (6662), through the Australian Government’s Collaborative Research Networks (CRN) program (AJM) and National Health and Medical Research Council grant (PP). JFB has an Australian Postgraduate Award (APA).
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Contents lists available at ScienceDirect
Molecular and Cellular Endocrinology j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / m c e
Acute leptin exposure reduces megalin expression and upregulates TGFβ1 in cultured renal proximal tubule cells Jessica F. Briffa a,b, Esther Grinfeld a, Michael L. Mathai a, Phillip Poronnik c, Andrew J. McAinch a, Deanne H. Hryciw a,b,* a
Centre for Chronic Disease Prevention and Management, College of Health and Biomedicine, Victoria University, St Albans, Vic. 3021, Australia Department of Physiology, The University of Melbourne, Parkville, Vic. 3010, Australia c School of Medical Sciences, The Bosch Institute, The University of Sydney, NSW 2006, Australia b
A R T I C L E
I N F O
Article history: Received 23 July 2014 Received in revised form 26 November 2014 Accepted 28 November 2014 Available online 2 December 2014 Keywords: Leptin Megalin Proximal tubule TGF-β1 AMPK
A B S T R A C T
Increased leptin concentrations observed in obesity can lead to proteinuria, suggesting that leptin may play a role in obesity-related kidney disease. Obesity reduces activation of AMP-activated protein kinase (AMPK) and increases transforming growth factor-β1 (TGF-β1) expression in the kidney, leading to albuminuria. Thus we investigated if elevated leptin altered AMPK and TGF-β1 signaling in proximal tubule cells (PTCs). In opossum kidney (OK) PTCs Western blot analysis demonstrated that leptin upregulates TGF-β1 secretion (0.50 μg/ml) and phosphorylated AMPKα (at 0.25, and 0.50 μg/ml), and downregulates megalin expression at all concentrations (0.05–0.50 μg/ml). Using the AMPK inhibitor, Compound C, leptin exposure regulated TGF-β1 expression and secretion in PTCs via an AMPK mediated pathway. In addition, elevated leptin exposure (0.50 μg/ml) reduced albumin handling in OK cells independently of megalin expression. This study demonstrates that leptin upregulates TGF-β1, reduces megalin, and reduces albumin handling in PTCs by an AMPK mediated pathway. © 2014 Elsevier Ireland Ltd. All rights reserved.
1. Introduction Obesity rates in Western cultures are rising rapidly, with comorbidities including chronic kidney disease (CKD) and end stage renal failure rising concurrently (Eknoyan, 2007). One of the earliest hallmark characteristics of CKD is the presence of albumin in the urine (albuminuria), which is typically the result of both a glomerular and proximal tubule dysfunction (Takeyama et al., 2011; Wohlfarth et al., 2003; Wolf et al., 2004). The hormone leptin is predominately produced by adipocytes (Vázquez-Vela et al., 2008), with plasma leptin concentrations reflecting adiposity (Garibotto et al., 1998), with higher leptin concentrations seen in obese individuals. Leptin binds to two transmembrane receptors, either the leptin receptor (ObRb) or the scavenger receptor megalin, in a cell specific manner. Megalin is responsible for the transportation of leptin in the hypothalamus, placenta, and renal tubules (Briffa et al., 2014; Hama et al., 2004), whereas ObR is responsible for leptin transport and signaling in most other tissues (Ahima, 2008). Circulating leptin is filtered from the blood by the glomerulus and is reabsorbed by cells of the proximal tubule by the scavenger
* Corresponding author. Department of Physiology, The University of Melbourne, Parkville, Vic. 3010, Australia. Tel.: +61 3 9035 9923; fax: +61 3 8344 5818. E-mail address: [email protected] (D.H. Hryciw). http://dx.doi.org/10.1016/j.mce.2014.11.024 0303-7207/© 2014 Elsevier Ireland Ltd. All rights reserved.
receptor megalin (Briffa et al., 2014; Hama et al., 2004), resulting in negligible leptin secretion in the urine (Meyer et al., 1997), even in obesity (Cumin et al., 1997; Lönnqvist et al., 1997). The plasma concentration of leptin in non-obese individuals is approximately 5.5 ng/ml, with 9.5% being transferred from the blood to the filtrate, with an estimated renal leptin clearance of 0.0595 μg/ml (Garibotto et al., 1998). The serum concentration of leptin in obese individuals is approximately 5 to 10 times higher than normal individuals (Garibotto et al., 1998). This value is disputed, with others determining that the maximum plasma leptin concentration observed in obesity is likely to be 200 ng/ml (Maffei et al., 1995). However, individuals with CKD demonstrate leptinemia at concentrations up to 490 ng/ml (Dagogo-Jack et al., 1998). Albuminuria is a hallmark characteristic of renal dysfunction, with studies demonstrating that exposure to elevated leptin can lead to a significant loss of albumin in the urine (Gunduz et al., 2005), suggesting that leptin may provide a link between CKD and obesity (Briffa et al., 2013). Albuminuria is typically the result of an increase in protein filtration through the glomerulus, caused by basement membrane thickening (Wohlfarth et al., 2003; Wolf et al., 2004), which is further compounded by an impairment in protein endocytosis by the proximal tubule cells (PTCs) which are unable to cope with the increased protein load (Takeyama et al., 2011). Importantly, exposure of the renal tubules to elevated concentrations of albumin induces the production of proinflammatory, profibrotic and vasoactive factors in vitro (Wolf et al., 2004). For example the
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profibrogenic cytokine transforming growth factor beta (TGF-β) is increased in the glomerular filtrate, which upregulates extracellular matrix production, ultimately resulting in basement membrane thickening (Wolf et al., 2004). Interestingly, exposure to elevated leptin has also been shown to increase TGF-β1 secretion in glomerular endothelial cells in vitro (Wolf et al., 1999). Exposure of renal cells to elevated protein and/or leptin also results in an accumulation of collagen in glomerular cells in vitro, which would result in tubulointerstitial fibrosis and basement membrane thickening in vivo (Wohlfarth et al., 2003). These data taken together suggest that leptin may play a role in obesity-related nephropathy by upregulating the expression of fibrotic mediators. Importantly, research by Gekle et al. (2003) has previously identified in opossum kidney (OK) cells, a well established model of the proximal tubule, exposure to 3 μg/ ml of TGF-β1 for 48 h significantly upregulates collagen secretion (types I and IV) and reduces albumin handling by decreasing megalin and cubilin protein expression. In addition to the fibrotic changes obesity causes in the kidney, obesity is also associated with tubular inflammation (Declèves et al., 2011), which further deteriorates kidney function. AMP-activated protein kinase (AMPK), a key regulatory of energy homeostasis, plays important roles in renal inflammation and fibrosis (Declèves et al., 2011; Sharma et al., 2008). Obesity results in a decrease in renal AMPK expression (Declèves et al., 2011), resulting in an upregulation of inflammatory mediators, as well as fibrotic changes to the kidneys (Declèves et al., 2011; Sharma et al., 2008). Specifically, research by Declèves et al. (2011) identified that one week of high fat feeding (60% fat) in six-week old C57BL/6 mice in vivo causes an increase in renal inflammatory markers that preceded albuminuria, with treatment of an AMPK activator preventing the increase in urinary expression of these mediators. Importantly, AMPK activation has been shown to decrease the fibrotic actions of TGF-β1, with an in vivo study identifying that AMPK activation, using Metformin, decreased the renal injury, adipokine expression, and macrophage infiltration that is associated with high fat feeding (60% fat) in six-week old C57BL/6 mice (Kim et al., 2013). In the kidney in vivo and in vitro (OK, LLC-PK1 and glomerular endothelial cells) leptin has been shown to upregulate fibrotic mediators and causes collagen deposition, which may compound renal function (Wohlfarth et al., 2003; Wolf et al., 1999), ultimately resulting in nephropathy. Therefore the overall aim of the current study was to identify if acute exposure to leptin will alter the expression of fibrotic mediators and, specifically, AMPK signaling in PTCs. We also aim to identify the effect leptin has on albumin handling in PTCs, and characterize changes in megalin expression. We hypothesize that exposure of PTCs to elevated leptin will upregulate fibrotic mediator expression and decrease AMPK expression, in addition to reducing albumin handling in PTCs.
2. Materials and methods 2.1. Cell culture We utilized OK cells for this study, as they are a widely used model for the renal proximal tubule as they express all proteins of the megalin signaling complex (Amsellem et al., 2010; Birn and Christensen, 2006; Hryciw et al., 2012). OK cells were maintained in Dulbecco’s Modified Eagle’s Medium and Ham’s F-12 (DMEM/ F12) media (Life Technologies; Victoria, Australia) supplemented with 10% fetal bovine serum (Life Technologies) with 1% penicillin/ streptomycin (Life Technologies), and incubated at 37 °C in 5% CO2. The cells were seeded at confluence and grown for two days in 25 ml flasks. Forty-eight hours prior to experimentation the cells were incubated in DMEM/F12 with 5 mM glucose medium lacking serum.
2.2. Animal care Experimental procedures were approved by the Howard Florey Institute Animal Ethics Committee (AEC 09-050). As used in our previous study (Jenkin et al., 2010), six-week old male Sprague Dawley rats (mean initial body weight approximately 178 g) were housed within individual cages in an environmentally controlled laboratory (ambient temperature 22–24 °C) with a 12 h light/dark cycle (7:00–19:00). Ad libitum access to food and water was maintained. At ten-weeks, rats were deeply anesthetized with sodium pentabarbitone (100 mg/kg; Virbac, Peakhurst, Australia) then euthanized via cardiac puncture. Kidney and white adipose tissues (WAT) were then removed and tissues were stored at −80 °C for subsequent Western blot analysis. 2.3. Immunoprecipitations OK cells were seeded onto 175 cm2 flasks and grown to confluence, and then serum starved for 48 h. The cells were then treated with either control (phosphate buffered saline: PBS) or human recombinant leptin (ProSpec; New Jersey, USA) for 15 min at 0.05 or 0.50 μg/ml. Protein was then isolated from OK cells using immunoprecipitation (IP) lysis buffer (10 mM Tris–HCl, 150 mM NaCl, 5% NP-40 with the pH adjusted to 7.5) supplemented with a Complete Mini Protease Inhibitor Cocktail (Roche; NSW, Australia). Immunoprecipitations (IPs) were then performed on 1 mg protein from each treatment. Cells were incubated overnight at 4 °C with no antibody, Normal Rabbit IgG (Merck Millipore; Victoria, Australia), Megalin antibody (ThermoFisher Scientific; Victoria, Australia), or Leptin antibody (Enzo Life Sciences; New York, USA) with endto-end rotation. The next day 50 μl pre-cleared Protein G Agarose (ThermoFisher Scientific) was added to each treatment and spun with end-to-end rotation at 4 °C for 5 h. The eppendorf tubes were then centrifuged and the supernatant was discarded, and the beads were washed with IP lysis buffer. Fifty microliters Laemmli Sample Buffer was then added to each treatment, and heated to 100 °C for 10 min. The treatments were then centrifuged and Western blots were performed on the supernatant. Equal aliquots (20 μl) of each treatment were separated on a 7.5% (Leptin IP) or 4–20% SDSPAGE gel (Megalin IP) (Bio-Rad; NSW, Australia) and transferred onto a nitrocellulose membrane. The membranes were then probed with either Leptin (Megalin IP) or Megalin (Leptin IP) antibodies to identify if leptin and megalin bind. 2.4. ‘Real-time’ polymerase chain reaction (PCR) OK PTCs were serum starved for 48 h (Briffa et al., 2014; Hryciw et al., 2006), then treated with human recombinant leptin (ProSpec) for 2 h at 0.05, 0.10, 0.25, and 0.50 μg/ml (Briffa et al., 2014). RNA was then extracted using TRIzol reagent according to the manufacturer’s instructions (Life Technologies) as described previously (Jenkin et al., 2010). The RNA was DNase treated, first strand cDNA was then generated from 0.5 μg of template RNA using the commercially available iScript™ cDNA synthesis kit (Bio-Rad) using random hexamers and oligo dTs as described previously (Jenkin et al., 2010). ‘Real-time’ PCR was conducted using MyiQ™ single color ‘realtime’ PCR detection system (Bio-Rad) with iQ™ SYBR Green Supermix (Bio-Rad) as the fluorescent agent. Forward and reverse oligonucleotide primers for megalin (forward 5′ TGC CCC ACC CGT TAT CCT A 3′ and reverse 5′ ACA GAC ATG GTT CTT ACA CTC AAA CAT 3′; Accession Number XM_007494924.1) and TGF-β1 (forward 5′ CCT GGA CAA CCA GTA CAG CA 3′ and reverse 5′ TTC CGG CCC ACA TAG TAG AC 3′; Accession Number XM_007491983.1) were designed using the OligoPerfect™ Suite (Life Technologies). Selective gene homology was confirmed using BLAST analysis (National Centre for Biotechnology Information; Maryland, USA). To compensate for
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variations in RNA input amounts and reverse transcriptase efficiency, mRNA abundance of the genes of interest were normalized to the housekeeping gene cyclophilin (forward 5′ CGG GTC ACT TTC GAG CTC TTT 3′ and reverse 5′ CTC AGA GCC CGG AAG TTT TCT 3′) (Jenkin et al., 2010). ‘Real-time’ PCR reactions were run for 50 cycles of 95 °C for 15 s and 60 °C for 60 s. Relative changes in mRNA abundance was quantified using the 2-ΔΔCT method as previously detailed (Jenkin et al., 2010) and reported in arbitrary units.
2.5. Protein extraction and Western blotting To prepare the tissue lysates, 50 mg of kidney was lysed in 500 μl HNT buffer (20 mM HEPES, 120 mM NaCl, 5 mM EDTA, 0.6% Triton-X with the pH adjusted to 7.5) and 100 mg WAT was lysed in 100 μl lysis buffer (50 mM Tris–HCl, 120 mM NaCl, 1% NP-40 with the pH adjusted to 7.5) with protease and phosphatase inhibitors (Cell Signaling Technology; Massachusetts, USA). To clarify the tissue lysates, the kidney lysate was centrifuged at 10,000× g for 5 min at 4 °C and the WAT lysate was centrifuged at 12,000× g for 20 min at 4 °C with the centrifugation occurring three more times on the supernatant to remove the lipids. In addition, OK PTCs were serum starved for 48 h in minus media, then treated with a vehicle control (PBS) or human recombinant leptin for 2 h (0.05–0.50 μg/ml). Protein was then isolated from OK cells using IP lysis buffer supplemented with a Complete Mini Protease Inhibitor Cocktail and Halt Phosphatase Inhibitor Cocktail (ThermoFisher Scientific). Equal aliquots (50 μg) of the protein samples were separated on a 7.5% (megalin) or 4–20% SDS-PAGE gel and transferred onto a nitrocellulose membrane. Membranes were probed with AMPKα, phosphorylated AMPKα (p-AMPKα), AMPKβ and phosphorylated AMPKβ (p-AMPKβ) antibodies (Cell Signaling Technology), leptin antibody (Enzo Life Sciences), collagen type IV and TGF-β1 antibodies (Abcam; Massachusetts, USA) or Megalin antibody (ThermoFisher Scientific) were used following protocols described previously (Hryciw et al., 2006; Slattery et al., 2011). Densitometric analysis of the banding was performed using ImageJ software. In addition, cells were pre-treated with either 234.6 nM of the AMPK inhibitor Compound C (Merck Millipore), according to the manufacturer’s IC50 and previous research (Hao et al., 2010), or a DMSO control (Sigma-Aldrich; NSW, Australia) for 60 min. After the pretreatment the cells were then treated with human recombinant leptin (0.05–0.50 μg/ml) in the presence of 234.6 nM Compound C or DMSO and PBS (control treatment) for 2 h. Western blots were then performed as described above and probed for Megalin and collagen type IV.
2.6. TGF-β1 ELISA OK cells were seeded onto 6-well plates and serum starved for 48 h prior to experimentation. The cells were then treated with human recombinant leptin (0.05–0.50 μg/ml) for 2 h. Following treatment, the cell culture media was collected and centrifuged, and a protein assay was performed on the cell lysate. The cell culture media was treated with 1 M HCl for 15 min, and then neutralized with 1 M NaOH. Total TGF-β1 concentrations were determined using an enzyme linked immunoassay assay (ELISA) (Promega; NSW, Australia) as per the manufacturer’s instructions. The absorbance was read at 450 nm on the xMark Microplate Absorbance Spectrophotometer. Secreted TGF-β1 concentrations were standardized to the protein content of the cells, and are presented as concentrations normalized to total cellular protein. In addition cells were exposed to Compound C (as described in section 2.5), and a TGF-β1 ELISA was performed as described above.
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2.7. Texas Red (TR) and Dye Quenched (DQ) albumin uptake We used standard albumin uptake methods in OK cells, a cell line commonly used for the study of proximal tubule albumin uptake (Hryciw et al., 2005, 2006). OK cells were seeded onto 48-well plates and treated with human recombinant leptin (0.05–0.50 μg/ml) for 2 h. To determine albumin uptake, cells were exposed to 50 μg/ml Texas Red albumin (TR-albumin) (Life Technologies) and treated with their respective leptin concentrations (0.05–0.50 μg/ml) for 2 h. To determine albumin degradation, OK cells were exposed to 50 μg/ ml Dye Quenched albumin (DQ-albumin) (Life Technologies) and treated their respective leptin concentrations (0.05–0.50 μg/ml) for 2 h. Nonspecific binding was determined in cells exposed to TR- or DQ-albumin for 1 min. At the end of the uptake period, cells were washed in HEPES-Ringer buffer (122.5 mM NaCl, 5.4 mM KCl, 1.2 mM CaCl2, 0.8 mM MgCl2, 0.8 mM Na2HPO4, 0.2 mM NaH2PO4, 5.5 mM Glucose, 10 mM HEPES; pH 6 at 4 °C), then lysed in MOPS buffer (20 mM MOPS with 0.1% Triton X-100) for 30 min at 37 °C. The TRalbumin fluorescence was determined using the POLARstar Galaxy (Biodirect; Massachusetts, USA) at 580 nm excitation and 630 nm emission wavelengths. The DQ-albumin fluorescence was determined using the POLARstar Galaxy at 590 nm excitation and 620 nm emission wavelengths. TR-albumin and DQ-albumin uptake were standardized to total cellular protein, and the amount of fluorescence per microgram of cellular protein was adjusted for background. As AMPK plays a protective role in mitigating the fibrotic actions of TGF-β1 (Kim et al., 2013), we repeated the TR-albumin uptake assay using Compound C. OK cells were exposed to Compound C (as described in section 2.5); however, the cells were only treated with 0.50 μg/ml human recombinant leptin (the concentration that significantly reduced albumin uptake and endocytosis) and the TR-albumin uptake was then performed as described above. 2.8. Proteasomal activity assay To determine if leptin alters proteasomal function, we next performed a proteasomal activity assay using methods previously published (Hryciw et al., 2004). Briefly, OK cells were incubated with varying concentrations (0.05–0.50 μg/ml) of human recombinant leptin for 2 h. To determine proteasomal activity, 75 μM N-succinylLeu-Leu-Val-Tyr-7-amino-4-methylcoumarin (LLVY-AMC) (Enzo Life Sciences) was added to 250 μg of protein from each treatment and topped to a final volume of 200 μl. The reaction mixture was incubated at 30 °C for 60 min. The fluorescent intensities of each sample were measured on the POLARstar Galaxy at excitation and emission wavelengths of 360 nm and 460 nm respectively, and were adjusted for background. The proteasome inhibitor Z-Leu-Leu-Leual (MG132) (Sigma-Aldrich) was used as a negative control (Hryciw et al., 2004). 2.9. Statistical analysis Statistical analysis of all data was conducted using a one-way ANOVA with Tukey’s post-hoc test or a student’s t-test. All data are expressed as the mean ± the standard error of the mean (SEM), with a P value of less than 0.05 being considered statistically significant. 3. Results 3.1. Leptin binds to megalin in the proximal tubule Previous research in Sprague-Dawley rats has determined that megalin is the main receptor responsible for leptin uptake in the convoluted segment of the proximal tubule, with no leptin binding to ObR in the straight segment of the proximal tubule (Hama et al.,
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tubule. ‘Real-time’ PCR analysis identified that OK cells exposed to 0.50 μg/ml leptin significantly increased TGF-β1 mRNA expression (0.1211 ± 0.024 compared to control, n ≥ 3, P < 0.05; Fig. 3A). Interestingly, 0.50 μg/ml leptin mRNA expression was significantly different to all other leptin treatments (0.1211 ± 0.024 compared to 0.0486 ± 0.006, 0.0411 ± 0.004 and 0.0569 ± 0.009 for 0.05, 0.10 and 0.25 μg/ml leptin respectively, n ≥ 3, P < 0.05; Fig. 3A). Western blot analysis identified a significant decrease in mature TGF-β1 protein expression at the same leptin concentration (0.50 μg/ ml) (42.83 ± 3.1% compared to control, n = 3, P < 0.05; Fig. 3B and C) indicating that there may be an increase in leptin secretion at this concentration. ELISA analysis identified that leptin, at the same concentration (0.50 μg/ml), significantly increased TGF-β1 secretion from OK cells (0.2289 ± 0.049 compared to control respectively, n ≥ 4, P < 0.05; Fig. 3D). Thus, leptin increases the secretion of TGF-β1 from PTCs, which may lead to the tubulointerstitial injury observed in CKD. Research has previously shown that AMPK activation can regulate the fibrotic actions of TGF-β1 (Kim et al., 2013). To investigate this association, OK cells were pretreated with Compound C to inhibit AMPK, and then treated with leptin and Compound C. ELISA analysis identified that AMPK inhibition significantly decreased Fig. 1. Leptin binds to megalin in vitro. (A) Immunoprecipitations indicate that megalin binds to leptin in vitro, and that leptin binds to megalin in vitro (n = 2). (B) Control Western blots indicate that opossum kidney (OK) cells are capable of producing leptin (50 μg OK lysate). White adipose tissue (WAT) lysate (12.5 μg), kidney lysate (12.5 μg) and human recombinant leptin (2 μg) were used as positive controls.
2004). Further, we have previously shown that OK cells lack ObR expression (Briffa et al., 2014), demonstrating that leptin interacts with megalin in OK cells. To confirm an interaction, we first investigated if leptin and megalin bind via immunoprecipitation. Leptin and megalin IPs identified that multiple leptin molecules bind in OK cells (n = 2; Fig. 1A) at 0.05 and 0.50 μg/ml. Interestingly OK control IPs (not treated with leptin) were also positive for megalin and leptin binding, suggesting that OK PTCs may be capable of producing leptin. To investigate if OK cells express leptin we next performed a Western blot on OK lysate, kidney lysate, WAT lysate (both of which are known to express leptin) and human recombinant leptin (as a positive control) (ProSpec). Western blot analysis identified that OK cells do express leptin protein; however, it appears to have a higher molecular weight than rodent and human leptin (Fig. 1B), suggesting there is a species variation in leptin’s molecular weight. 3.2. Short term exposure of OK cells to leptin alters AMPK expression We investigated if leptin treatment alters the expression of AMPK in OK cells. Western blot analysis identified that leptin upregulates p-AMPKα at 0.25 and 0.50 μg/ml (149.16 ± 13.5% and 198.31 ± 10.7% compared to control respectively, n ≥ 3, P < 0.05; Fig. 2A and B). Interestingly, 0.50 μg/ml leptin was also significantly increased compared to all other leptin treatments (198.31 ± 10.7% compared to 118.34 ± 5.6%, 126.7 ± 20.83% and 149.16 ± 13.5% for 0.05, 0.10, and 0.25 respectively, n ≥ 3, P < 0.05; Fig. 2A and B). In contrast leptin did not alter p-AMPKβ expression (Fig. 2A and B). 3.3. Short term exposure of OK cells to leptin alters TGF-β1 expression and secretion by an AMPK mediated pathway As previous research has well established that exposure to elevated leptin upregulates TGF-β production and secretion in the glomerulus (Wolf et al., 1999), we next investigated the effect leptin has on TGF-β1 protein production and secretion in the proximal
Fig. 2. Leptin alters the expression of AMPK in OK cells. (A) In opossum kidney (OK) cells exposed to leptin for 2 h, phosphorylated AMPKα (p-AMPKα) protein was increased at 0.25 and 0.50 μg/ml. OK cells treated with 0.50 μg/ml leptin significantly altered p-AMPKα compared to all other leptin treatments. (B) Densitometric analysis confirmed the significant increase of p-AMPKα at 0.25 and 0.50 μg/ml (n ≥ 3, P < 0.05 vs. control) and a significant increase in p-AMPKα at 0.50 μg/ml compared to 0.05, 0.10 and 0.25 μg/ml leptin (n ≥ 3, P < 0.05 vs. 0.05, 0.10 and 0.25 μg/ml leptin). Densitometry analysis also identified no changes in p-AMPKβ expression in response to leptin. Significant differences between treatments are indicated by letters that differ (a, b, c); for example “a” is different from “b”, but not different from “ab”.
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Fig. 3. Leptin increases TGF-β1 mRNA transcription and TGF-β1 secretion in OK cells by an AMPK mediated pathway. (A) ‘Real-time’ PCR analysis identified that opossum kidney (OK) cells exposed to 0.50 μg/ml of leptin for 2 h significantly increased TGF-β1 mRNA transcription (n ≥ 3, P < 0.05 vs. control). Additionally, 0.50 μg/ml leptin mRNA expression of TGF-β1 was significantly greater than all other leptin concentrations (n ≥ 3, P < 0.05 vs. 0.05, 0.10 and 0.25 μg/ml leptin). (B) Western blot analysis identified a significant decrease in mature TGF-β1 protein in OK cells treated with 0.50 μg/ml leptin. (C) Densitometry analysis confirmed the significant decrease in mature TGF-β1 protein at 0.50 μg/ml leptin (n = 3, P < 0.05 vs. control). (D) ELISA analysis identified that OK cells exposed to leptin for 2 h significantly increased TGF-β1 secretion at 0.50 μg/ ml (n ≥ 4, P < 0.05 vs. control). (E) Pretreatment with Compound C resulted in a significant decrease in TGF-β1 being secreted from OK PTCs at 0.10, 0.25, and 0.50 μg/ml leptin (n ≥ 5, P < 0.05 vs. control), as well as compared to physiological (0.05 μg/ml) leptin (n ≥ 4, P < 0.05 vs. 0.05 μg/ml leptin). Significant differences between treatments are indicated by letters that differ (a, b); for example “a” is different from “b”, but not different from “ab”.
TGF-β1 secretion from OK cells at 0.10, 0.25, and 0.50 μg/ml leptin (0.0256 ± 0.004, 0.0298 ± 0.007 and 0.0222 ± 0.007 compared to control respectively, n ≥ 5, P < 0.05 vs. control; Fig. 3E). Importantly, these concentrations were significantly different compared to physiological (0.05 μg/ml) leptin concentrations (0.0256 ± 0.004, 0.0298 ± 0.007 and 0.0222 ± 0.007 compared to 0.0779 ± 0.012 for 0.05 μg/ml leptin, n ≥ 5, P < 0.05 vs. 0.05 μg/ml leptin; Fig. 3E), indicating that if not for the increased p-AMPKα expression leptin would not cause the increase in TGF-β1 secretion. These findings suggest, in the renal proximal tubule, that leptin increases AMPK activation, which positively regulates TGF-β1 expression. 3.4. Short term exposure of OK cells to elevated concentrations of leptin does not alter collagen type IV expression As TGF-β1 is known to upregulate collagen type IV production (Gekle et al., 2003; Grande et al., 2002), we next examined if leptin upregulates collagen type IV protein expression in cells of the proximal tubule. OK
cells exposed to leptin did not change collagen IV production (Fig. 4A and B). This finding is not surprising as the cells were only treated with leptin acutely (for 2 h), with previous research showing that prolonged exposure (48 h) of OK cells to TGF-β1 (3 μg/ml) upregulates Collagen IV protein expression (Gekle et al., 2003). As research has shown that AMPK attenuates the fibrotic changes brought on by TGF-β1 (Lee et al., 2013), we also investigated the effect AMPK inhibition has on collagen type IV expression. Western blot analysis identified that AMPK inhibition did not alter collagen type IV protein expression (compared to control) (n ≥ 3; Fig. 4C and D). 3.5. Short term exposure of OK cells to leptin decreases megalin protein expression by an AMPK mediated pathway Elevated TGF-β1 concentrations have been previously shown to decrease megalin protein expression in OK cells (Gekle et al., 2003). As leptin upregulates TGF-β1 secretion, we next examined the effect leptin has on megalin mRNA and protein expression. ‘Real-time’ PCR
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Fig. 4. Leptin does not alter collagen type IV production in OK cells. (A) In opossum kidney (OK) cells exposed to leptin for 2 h, collagen type IV protein expression did not change. (B) Densitometric analysis confirmed no change in collagen IV protein expression. (C) Western blot analysis identified that AMPK inhibition did not alter collagen type IV protein production. (D) Densitometry analysis confirmed that AMPK inhibition did not alter collagen type IV protein production.
Fig. 5. Leptin decreases megalin expression in OK cells by an AMPK mediated pathway. (A) In opossum kidney (OK) cells exposed to leptin for 2 h, there was no change in megalin mRNA transcription. (B) OK cells exposed to leptin for 2 h significantly decreased megalin expression across all leptin treatments (0.05–0.50 μg/ml). (C) Densitometric analysis confirmed the significant reduction in megalin protein across all leptin treatments (n = 4, P < 0.05 vs. control). (D) Western blot analysis identified that AMPK inhibition with Compound C restores megalin protein expression to values similar to control. (E) Densitometry analysis confirmed no change in megalin protein expression compared to control (n ≥ 5). (F) In OK cells, exposure to leptin for 2 h does not alter proteasomal function. The proteasome inhibitor Z-Leu-Leu-Leu-al (MG132) was used as a negative control, and significantly decreased proteasome function (n = 6, P < 0.05 vs. control). Significant differences between treatments are indicated by letters that differ (a, b); for example “a” is different from “b”, but not different from “ab”.
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Fig. 6. OK cells exposed to elevated leptin reduces albumin uptake and degradation by an AMPK mediated pathway. (A) Confluent monolayers of opossum kidney (OK) cells were incubated with 50 μg/ml of Texas Red albumin (TR-albumin) and leptin for 2 h. At 0.50 μg/ml of leptin, albumin uptake was significantly reduced compared to control (n = 4, P < 0.05 vs. control). Additionally, OK cells treated with 0.50 μg/ml leptin significantly reduced albumin uptake compared to all other leptin treatments (n = 4, P < 0.05 vs. 0.05, 0.10 and0.25 μg/ml leptin). (B) Confluent monolayers of OK cells were incubated with 50 μg/ml of Dye Quenched albumin (DQ-albumin) and leptin for 2 h. There was a significant decrease in albumin degradation, as measured by the DQ-albumin assay, in cells treated at 0.50 μg/ml of leptin (n = 4, P < 0.05 vs. control) which coincides with the reduction in albumin endocytosis. Additionally, OK cells treated with 0.50 μg/ml leptin significantly reduced albumin degradation compared to all other leptin treatments (n = 4, P < 0.05 vs. 0.05, 0.10 and0.25 μg/ml leptin). (C) Confluent monolayers of OK cells were treated with either Compound C or DMSO and 0.50 μg/ml leptin, with 50 μg/ml of TR-albumin for 2 h. AMPK inhibition resulted in a significant increase in albumin endocytosis in OK cells compared to cells treated with DMSO and leptin (n = 4, P < 0.05 vs. control). Significant differences between treatments are indicated by letters that differ (a, b); for example “a” is different from “b”, but not different from “ab”.
analysis identified that acute leptin exposure does not alter the expression of megalin mRNA (n ≥ 5; Fig. 5A). However, Western blot analysis identified that leptin significantly decreases megalin protein expression across all concentrations (55.7 ± 2.26%, 68.09 ± 7.25%, 64.59 ± 8.17% and 58.65 ± 9.16% compared to control respectively, n = 4, P < 0.05 vs. control; Figs. 4C and 5B). We demonstrated above that AMPK inhibition significantly decreases TGF-β1 secretion from OK cells compared to control (Fig. 3E), and as research has previously shown that TGF-β1 reduces megalin expression (Gekle et al., 2003), we next investigated the effect AMPK inhibition has on megalin protein expression in response to leptin. Western blot analysis identified that AMPK inhibition restored megalin protein expression to values similar to control (n ≥ 5; Fig. 5D and E). To date, the entire genome of the opossum has not yet been characterized, and as such the exact molecular weight of megalin in opossums is not known. However, the molecular size of our megalin product is similar to previous studies, which have demonstrated that opossum megalin is greater than 250 kDa (Labeau et al., 2001; Zou et al., 2004). 3.6. Short term exposure of OK cells to leptin does not alter proteasomal function As elevated albumin alters proteasomal activity in tubular cells (Hryciw et al., 2004), we also investigated if leptin alters the proteasomal pathway in OK cells. Proteasomal analysis identified that leptin (across all concentrations) does not alter the proteasomal activity in PTCs (n ≥ 6; Fig. 5F). The proteasome inhibitor MG132 was used as a negative control.
3.7. Short term exposure of OK cells to leptin alters PTC handling of albumin by an AMPK mediated pathway As TGF-β1 reduces albumin handling in PTCs (Gekle et al., 2003), we next investigated if leptin alters albumin handling in the proximal tubule. In OK cells exposed to 50 μg/ml of TR-albumin, in the presence of leptin, there was a significant reduction in albumin uptake in cells exposed to 0.50 μg/ml leptin (71.37 ± 5.2% compared to control, n = 4, P < 0.05 vs. control; Fig. 6A). Importantly, 0.50 μg/ml leptin reduced albumin endocytosis compared to all other leptin treatments (71.37 ± 5.2% compared to 92.1 ± 6.00%, 102.15 ± 3.50% and 87.13 ± 2.55% for 0.05, 0.10 and 0.25 μg/ml leptin respectively, n = 4; P < 0.05 vs. 0.05, 0.10 and 0.25 μg/ml leptin; Fig. 6A). Not surprisingly, in OK cells exposed to 50 μg/ml of DQalbumin, in the presence of leptin, there was also a significant reduction in albumin degradation in cells exposed to 0.50 μg/ml leptin (68.5 ± 2.72% compared to control, n = 4, P < 0.05 vs. control; Fig. 6B). Additionally a reduction in albumin degradation was observed at 0.50 μg/ml leptin compared to all other leptin treatments (68.5 ± 2.72% compared to 90.75 ± 4.87%, 85.00 ± 4.65% and 86.75 ± 3.97% for 0.05, 0.10 and 0.25 μg/ml leptin respectively, n = 4; P < 0.05 vs. 0.05, 0.10 and 0.25 μg/ml leptin; Fig. 6B). These data indicate that at elevated concentrations of leptin, albumin uptake and degradation are altered in the proximal tubule which is independent of changes in megalin receptor expression (Fig. 5B and C). To identify the role AMPK plays in regulating albumin endocytosis we repeated this experiment at 0.50 μg/ml leptin to identify if AMPK inhibition restores albumin endocytosis in response to leptin.
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TR-albumin uptake analysis identified that AMPK inhibition results in a significant increase in albumin uptake in OK cells compared to cells treated with a DMSO control and leptin (136 ± 10.88%, compared to control, n = 4, P < 0.05 vs. control; Fig. 6C). 4. Discussion This study describes the potentially significant role for the adipokine leptin in the renal dysfunction that is associated with obesity. Importantly, we have demonstrated that 1) the proximal tubule is capable of producing leptin, 2) leptin significantly upregulates TGF-β1 expression via an AMPK mediated pathway, 3) exposure to elevated leptin concentrations significantly reduces albumin handling of PTCs, and finally 4) the reduction in albumin handling that leptin causes is due to altered megalin function, not protein expression levels. We have previously shown that OK cells lack ObR expression (Briffa et al., 2014), which suggests that any alterations in signal transduction pathways are due to megalin-mediated signaling. However, there has been limited research investigating the ability of megalin to act as a signal transducer. Importantly, we have demonstrated that PTCs exposed to elevated leptin activates the mTOR signaling pathway (Briffa et al., 2014), TGFβ1 and AMPK (this study), which supports a role for megalin-mediated cell signaling in renal pathophysiology. Research by Zou et al. (2004) demonstrated that vitamin D binding protein (another megalin ligand) activates cell signaling pathways in OK cells in vitro through the cleavage of the intracellular carboxyl terminus of the receptor and activation of specific intracellular targets. Recent research in L2 rat yolk sac cells identified that megalin is trafficked to the endocytic recycling compartment for proteolytic processing and the modulation of cellular gene expression for the activation of signal transduction pathways (Shah et al., 2013). However, no studies thus far have demonstrated that albumin can activate megalin cell signaling pathways, and importantly the affinity for megalin binding to both endogenous and exogenous leptin has not been determined, as such, future studies are required to determine this association. In this study we chose to use several different concentrations of leptin (0.05–0.50 μg/ml) to investigate its effect on the renal proximal tubule. Despite the maximal concentration of leptin in the glomerular filtrate being ~47 ng/ml, assuming a normal glomerular barrier, this is likely to be an underestimate due to the following observations. Firstly, research in rabbit proximal tubules have identified that horseradish peroxidase (44 kDa) is transported from the basolateral membrane (Nielsen and Christensen, 1985), suggesting that if leptin is able to be transported from the basolateral membrane to the apical membrane in the renal proximal tubule, then the filtrate concentration of leptin may be significantly higher. More importantly, recent research has shown that kidney fibroblasts are capable of producing and secreting leptin in conditions reflecting renal fibrosis (Lin et al., 2011); however, if PTCs are capable of secreting leptin it is likely to be due to another mechanism as they are epithelial cells. Therefore, if PTCs are able to secrete leptin, especially under conditions of glomerular damage (Lin et al., 2011), then the concentration of leptin in the filtrate in these conditions is likely to be greater than our estimate of ~47 ng/ml. Importantly, we have shown that OK cells can produce leptin protein, however the molecular weight appears to be larger (closer to 20 kDa) compared to human and rodent leptin (~16 kDa). These differences in molecular weight are likely due to a species variation, as based on the predicted protein sequence of the opossum leptin protein (Accession Code – XP_007504322) the molecular weight has a predicted size of 22 kDa. Previous studies have shown that leptin upregulates fibrotic mediator expression. Specifically, leptin has been shown to upregulate TGF-β1 expression and collagen production both in vitro (glomerular endothelial cells) and in vivo (Wolf et al., 1999), which contributes to tubulointerstitial fibrosis. Research in leptin deficient mice (ob/ ob) that underwent unilateral urethral obstruction surgery has shown
a significant reduction in TGF-β mRNA and reduced fibrotic changes to the kidney compared to leptin-receptor deficient mice (db/db) and control animals, identifying that leptin is a co-activator of TGFβ1 (Kümpers et al., 2007). In support of this, glomerular endothelial cells treated with leptin (0.62–625 nM) for 48 h have a significant increase in TGF-β mRNA expression and secretion (Wolf et al., 1999). Additionally, leptin treatment (100 ng/ml) increased fibrotic mediator expression in glomerular mesenchymal cells (Cui et al., 2013). Specifically, TGF-β1 (6 h treatment), Collagen IV (24 h treatment), and Fibronectin (6 and 24 h treatment) are all increased with leptin treatment (Cui et al., 2013). Furthermore, leptin infusion in Wistar rats in vivo for 72 h or three weeks results in a significant increase in renal TGF-β mRNA at both time points, and an increase in collagen type IV mRNA at three weeks of leptin infusion (Wolf et al., 1999). Our data in the renal proximal tubule, with acute leptin treatment, support these findings, showing that leptin treatment significantly increases TGF-β1 secretion from OK cells; in addition to increasing TGF-β1 mRNA and decreasing TGF-β1 mature protein expression at significantly elevated concentrations (0.50 μg/ml). However, it is not surprising that our study did not find an increase in collagen type IV protein. Our study employed an acute leptin treatment. These findings are supported by the study by Wolf et al. (1999), which demonstrated that TGF-β1 expression increases prior to changes in collagen expression. Importantly, research has well established that TGF-β1 regulates collagen type IV production following 48 h of treatment (Gekle et al., 2003). In addition to leptin altering TGF-β1 expression, we also identified that leptin significantly upregulates p-AMPKα expression, with research well supporting a role for AMPK in inhibiting the fibrotic actions of TGF-β1 (Lee et al., 2013). Importantly, obesity is associated with reduced AMPK expression, a mediator that plays important roles in renal inflammation and fibrosis (Declèves et al., 2011; Sharma et al., 2008). In support of this, research by Declèves et al. (2011) demonstrated that C57BL/6 mice fed a high fat diet (60% fat) for one week in combination with AICAR, an AMPK activator, reduced renal hypertrophy and fibrotic mediator expression (H2O2 and monocyte chemotactic protein1). Surprisingly, we identified in conditions reflecting obesity (i.e. high leptin and low AMPK expression) in PTCs that there is a significant reduction in TGF-β1 secretion from OK cells at 0.10–0.50 μg/ml leptin. These differences may arise due to the receptor that leptin is signaling from, with leptin presumably signaling via ObRb in the glomerulus (Wolf et al., 1999), whereas in the proximal tubule leptin signals via megalin (Briffa et al., 2014; Hama et al., 2004). Collectively these findings suggest in renal PTCs, acute leptin treatment with elevated concentrations of leptin, activates AMPK, which upregulates TGF-β1 expression, which in turn may upregulate collagen type IV production with prolonged exposure, ultimately leading to fibrosis (Fig. 7). Research has previously identified that TGF-β1 reduces megalin receptor expression and albumin handling in the renal tubules (Gekle et al., 2003). Specifically, OK cells treated with TGF-β1 (0.3–10 ng/ml) for 48 h have a significant reduction in albumin endocytosis (Gekle et al., 2003). With additional research in OK cells identifying that 48 h exposure to 3 μg/ml TGF-β1 reduced albumin binding capacity at the apical membrane by reducing both megalin and cubilin expression, resulting in a reduction in albumin degradation (Gekle et al., 2003). In the current study we make the important observation that short-term exposure of OK cells to leptin significantly reduces megalin protein expression (0.05–0.50 μg/ml), although albumin endocytosis and degradation were only reduced at significantly elevated concentrations of leptin (0.50 μg/ ml), which is independent of changes in proteasomal function. This suggests that altered function of megalin, but not abundance of the protein, is involved in the renal pathophysiology associated with obesity. Leptin infusion in mice clearly results in albuminuria (Gunduz et al., 2005), with data from our study indicating that leptin directly reduces albumin handling in the proximal tubule. These findings are in contradiction with in vivo models, which clearly shows that leptin can
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References
Fig. 7. AMPKs role in activating fibrotic mediators. A proposed model of how AMPK is modulating fibrotic mediator expression by leptin in the renal proximal tubule. At elevated leptin concentrations, leptin activates AMPK which regulates TGF-β1 expression in renal proximal tubule cells, which may in turn upregulate collagen type IV production; ultimately resulting in fibrosis. Importantly, an upregulation of secreted TGF-β1 decreases megalin receptor expression and function, which impairs albumin uptake and degradation in the renal proximal tubule. Solid lines indicate observed changes seen in this study, with dashed lines indicating hypothesized changes (based on previous studies) due to a prolonged exposure of leptin.
attenuate albumin excretion in humans with lipodystrophy (Ebihara et al., 2007; Javor et al., 2004) and animals with diabetic nephropathy (Suganami et al., 2005), ultimately abolishing their albuminuria and proteinuria. However, under disease states, additional mediators may be affecting albumin handling in addition to leptin. Interestingly, AMPK inhibition restored megalin protein expression and resulted in a significant increase in albumin endocytosis, which supports the study by Gekle et al. (2003). As shown in Fig. 7 in the renal proximal tubule, leptin increasing TGF-β1 protein expression and secretion (via AMPK activation) decreases megalin receptor expression and function, which impairs albumin uptake and degradation in renal PTCs. Thus, leptin may reduce albumin uptake in pathophysiological conditions via fibrotic damage, which has been suggested in other nephropathy models before (Wohlfarth et al., 2003). In summary, this study is the first to identify that leptin upregulates TGF-β1 expression, and alters albumin processing in renal PTCs. Importantly this study provides novel data showing, in PTCs, that AMPK is mediating the expression of TGF-β1 which is altering the function of megalin, ultimately disrupting albumin handing. Thus altering leptins’ effects on AMPK within the PTC appears to warrant further investigation as a possible treatment target to reduce obesity associated renal damage. Acknowledgments This research was supported by the Helen McPherson Smith Trust (6662), through the Australian Government’s Collaborative Research Networks (CRN) program (AJM) and National Health and Medical Research Council grant (PP). JFB has an Australian Postgraduate Award (APA).
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