Obesity is associated with induction of the ER (endoplasmic reticulum)-stress response signalling and insulin resistance. PTP1B (protein tyrosine phosphatase 1B) is a major regulator of adiposity and insulin sensitivity. The aim of the present study was to investigate the role of L-PTP1B (liver-specific PTP1B) in chronically HFD (high-fat diet) and pharmacologically induced (tunicamycin and thapsigargin) ER-stress response signalling in vitro and in vivo. We assessed the effects of ER-stress response induction on hepatic PTP1B expression, and consequences of hepatic-PTP1B deficiency, in cells and mouse liver, on components of ER-stress response signalling. We found that PTP1B protein and mRNA expression levels were up-regulated in response to acute and/or chronic ER stress, in vitro and in vivo. Silencing PTP1B in hepatic cell lines or mouse liver (L-PTP1B−/−) protected against induction of pharmacologically induced and/or obesity-induced ER stress. The HFD-induced increase in CHOP (CCAAT/enhancer-binding protein homologous protein) and BIP (binding immunoglobulin protein) mRNA levels were partially inhibited, whereas ATF4 (activated transcription factor 4), GADD34 (growth-arrest and DNA-damage-inducible protein 34), GRP94 (glucose-regulated protein 94), ERDJ4 (ER-localized DnaJ homologue) mRNAs and ATF6 protein cleavage were completely suppressed in L-PTP1B−/− mice relative to control littermates. L-PTP1B−/− mice also had increased nuclear translocation of spliced XBP-1 (X box-binding protein-1) via increased p85α binding. We demonstrate that the ER-stress response and L-PTP1B expression are interlinked in obesity- and pharmacologically induced ER stress and this may be one of the mechanisms behind improved insulin sensitivity and lower lipid accumulation in L-PTP1B−/− mice.
- endoplasmic reticulum (ER) stress
- insulin resistance
- metabolic syndrome
- protein tyrosine phosphatase 1B (PTP1B)
ER (endoplasmic reticulum) is a eukaryotic intracellular organelle required for synthesis and metabolism of many complex metabolites and biologically active proteins . ER can also sense and respond to alterations in cellular homoeostasis. Conditions interfering with the function of ER are collectively called ER stress. For example, accumulation of unfolded protein aggregates in the ER lumen leads to the activation of a pathway termed UPR (unfolded protein response) with the aim to return the ER to its normal physiological state. This complex cellular response is mediated initially by three molecules: PERK [PKR (double-stranded-RNA-dependent protein kinase)-like ER kinase], ATF6 (activated transcription factor 6) and IRE1 (inositol-requiring enzyme 1) [2,3]. The ER luminal domain of PERK, IRE1 and ATF6 interacts with the ER chaperone GRP78 (glucose-regulated protein 78)/BIP (binding immunoglobulin protein)].
Activated PERK phosphorylates eukaryotic initiation factor 2α [eIF2α (eukaryotic initiation factor 2α)], thereby reducing the rate of general protein translation, and thus protein load, on the ER [4,5]. Phosphorylation of eIF2α paradoxically increases translation of ATF4 mRNA to produce a transcription factor that activates expression of several UPR target genes [4,6]. Expression profiling studies have found that PERK, eIF2α and ATF4 are required for expression of genes involved in amino acid biosynthesis and transport, anti-oxidative stress and apoptosis, e.g. GADD34 (growth-arrest and DNA-damage-inducible protein 34) and CHOP (CCAAT/enhancer-binding protein homologous protein) . Activation of the ER protein kinase IRE1 triggers its endoribonuclease activity to induce cleavage of XBP-1 (X-box-binding protein-1) mRNA. XBP-1 mRNA is then ligated by an uncharacterized RNA ligase and translated to produce spliced XBP-1 protein . Spliced XBP-1 protein is a highly active transcription factor and one of the key regulators of ER-folding capacity . Some of the genes identified that require activation of the IRE1/XBP-1 pathway are components of the ERAD (ER-associated degradation) machinery, such as EDEM (ER degradation-enhancing α-mannosidade-like protein) [10,11], major ER chaperones such as GRP78/BIP and GRP94, and the ERDJ4 (ER-localized DnaJ homologue) [9,10]. Concurrently, ATF6 is released from GRP78 and transits to the Golgi body where it is cleaved to release a transcriptionally active fragment . Cleaved ATF6 acts in concert with spliced XBP-1 protein to induce expression of genes encoding protein chaperones and components of the ERAD machinery [13,14].
Obesity and Type 2 diabetes result in conditions that increase demand on the ER. This is particularly clear in the liver, adipose tissue and pancreas, where changes in tissue architecture, increases in protein synthesis and perturbations in cellular energy fluxes occur . Previous studies have demonstrated that ER stress is increased in adipose and liver tissues in both dietary and genetic obesity [16–18]. In cellular systems, the induction of ER stress leads to insulin resistance, at least in part through IRE-1-dependent activation of JNK (c-Jun N-terminal kinase). Modulation of ER-folding capacity through gain- and loss-of-function studies with XBP-1 showed a close link between ER function and insulin action in vitro and in vivo .
PTP (protein tyrosine phosphatase) 1B is the prototype for the superfamily of PTPs and has been implicated in multiple signalling pathways . Gene-targeting studies in mice have established PTP1B as a critical physiological regulator of metabolism and body mass by attenuating insulin and leptin signalling [16,20–25]. PTP1B-deficient mice exhibit resistance to diabetes and obesity, the two major metabolic diseases in industrialized societies. Not surprisingly, PTP1B is a highly regarded target of the pharmaceutical industry in the treatment of these disorders . Recently, we generated liver-specific PTP1B−/− (L-PTP1B−/−) mice and found that they exhibited improved glucose homoeostasis and lipid profiles, independently of changes in adiposity when fed on an HFD (high-fat diet). In addition, L-PTP1B deletion also partially attenuated some markers of the hepatic ER-stress response induced by chronic HFD feeding . The aim of the present study was to investigate in detail the contribution of PTP1B in the induction and regulation of the ER-stress response in liver using both in vitro and in vivo models of PTP1B deletion. The ER-stress response was induced either by chronic feeding of mice with an HFD and/or with pharmacological treatment of mice and cells, and subsequently activation of the PERK, IRE1 and ATF6 pathways were examined.
MATERIALS AND METHODS
All animal studies were performed under a project licence approved by the Home Office under the Animals (Scientific Procedures) Act 1986 and the principles of laboratory animal care (NIH publication no. 85-23, revised 1985; http://grants1.nih.gov/grants/olaw/references/phspol.htm) were followed. Mice were maintained on a 12 h light/12 h dark cycle in a temperature-controlled barrier facility, with free access to water and food. L-PTP1B−/− and fl/fl control mice have been described previously [16,22]. All mice studied were age-matched littermate males on the mixed 129Sv/C57Bl6 background. Mice were placed either on standard laboratory chow or an HFD (Harlan Teklad, Bicester, Oxon, U.K.; 55% fat) at weaning (21 days old), and weights were monitored weekly. The animals were kept on the diet for 16 weeks.
For tunicamycin treatment of animals, mice were given a single 1 μg/g of body weight intraperitoneal injection of 0.05 g/l suspension of tunicamycin (Sigma, Gillingham, U.K.) in saline. The control groups were injected with saline. After 3 h, mice were killed by decapitation, and livers were removed and frozen in liquid N2.
Glucose from tail blood was assessed using a glucometer (Accu-Check, Burgess Hill, U.K.). Serum insulin was determined by ELISA (CrystalChem, Downers Grove, IL, U.S.A.) as described previously [16,22,23].
The HepG2 (a human liver carcinoma cell line) was routinely cultivated in DMEM (Dulbecco's modified Eagle's Medium; Gibco, Paisley, U.K.) supplemented with 10% FBS (foetal bovine serum, Invitrogen, Paisley, U.K.), 2 mmol/l glutamine and 1% penicillin/streptomycin (Gibco), and were maintained at 37 °C in a humidified atmosphere with 5% CO2.
siRNA (small interfering RNA) duplexes specific for human PTP1B were obtained from Santa Cruz Biotechnology and control non-silencing siRNA was from Eurogentec (Southampton, U.K.). Transient transfection of cells was done using the Lipofectamine™ RNAiMAX transfection reagent (Invitrogen). Then, cells were treated with tunicamycin (5 mg/l) or vehicle (DMSO) for 7 h, followed by RNA or protein extraction for further analysis.
In another set of experiments, stable knockdown of PTP1B in HepG2 cells was performed using specific shRNA (short-hairpin RNA) constructs designed against human PTP1B using Gateway System (Invitrogen). The viruses were transduced into the cells in the presence of 5 g/l polybrene (Millipore, Chandlers Ford, Hampshire, U.K.). Then, cells were treated with thapsigargin (300 nM) or vehicle (DMSO) for 7 h, followed by RNA extraction for further analysis.
mRNA expression analysis
Total RNA was isolated from transfected/transduced HepG2 cells or mouse livers using TRI Reagent® (Ambion, Warrington, U.K.) according to the manufacturer's protocol. First-strand cDNA was synthesized from 1 μg of total RNA employing the Biorscript™ Preamplification System (Bioline, London, U.K.) and an oligo(dT)12–18 primer as the reverse primer. Then, target genes were amplified by real-time PCR using GoTaq™ qPCR Master Mix (Promega, Southampton, U.K.) in Roche LightCycler® 480 System (Roche Diagnostics, Burgess Hill, U.K.). Relative gene expression was calculated using the comparative Ct (2−ΔΔCt) method. The primer sequences used for PCR, real-time PCR and sequencing are listed in Supplementary Table S1 (at http://www.BiochemJ.org/bj/438/bj4380369add.htm).
XBP-1 splicing assay
We analysed the splicing of XBP-1 from cDNA using Fast Start Taq DNA polymerase. The PCR conditions were as follows: 94 °C for 3 min; 35 cycles of 94 °C for 10 s, 65 °C for 30 s, 72 °C for 30 s and 72 °C for 10 min. We used the following primers: forward 5′-AAACAGAGTAGCAGCTCAGACTGC-3′ and reverse 5′-TCCTTCTGGGTAGACCTCTGGGAG-3′. To distinguish the unspliced from the spliced band, we resolved the PCR products on a 2% agarose gel.
Tissue lysates were prepared by extraction in RIPA as described previously [16,23]. Immunoblots were performed using antibodies against PTP1B (Millipore), SHP2 (Src homology 2 domain-containing protein tyrosine phosphatase 2), CHOP, ERK (extracellular-signal-regulated kinase) 1, ERK2, human PTP1B (Santa Cruz Biotechnology), lamin A/C, phospho-eIF2α (Ser51), BIP (GRP78), IRE1-α, GAPDH (glyceraldehyde-3-phosphate dehydrogenase) (Cell Signaling Technology), XBP-1 (spliced and unspliced forms) (Abcam, Cambridge, U.K.), ATF6 (full-length and cleaved forms) (IMGENEX, Cambridge, U.K.) and PTEN (phosphatase and tensin homologue deleted on chromosome 10) (Abgent, Oxford, U.K.). Proteins were visualized using enhanced chemiluminescence and quantified either by scanning densitometry (Image J) or Bio1D software on PeqLab Fusion gel imaging system (PeqLab, Fareham, U.K.).
Nuclear and cytosolic extraction
For nuclear extracts, we cut the liver tissue into small pieces and washed once with ice-cold PBS then lysed in NE-PER extraction reagent (ThermoScientific, Loughborough, U.K.) according to the manufacturer's instructions and as described in . Samples were subjected to SDS/PAGE, transferred on to a nitrocellulose membrane and immunoblots performed as described above.
For XBP-1/p85α co-immunoprecipitation, we subjected both nuclear and cytosolic liver extracts from HFD-fed mice to immunoprecipitation using Pierce Classic IP kit (Thermoscientific) according to the manufacturer's protocol with minor modification as described by Park et al. . Briefly, we added 5 μg of a goat anti-XBP-1 antibody to 400 μg of liver nuclear or cytosolic lysates and incubated overnight at 4 °C with gentle rotation. The next day, we added the antibody/lysate sample to Protein A/G Plus–agarose in spin columns and incubated with gentle end-over-end mixing or shaking for 2 h (Eppendorf, St Albans, U.K.). Proteins from the immunoprecipitates were resolved by SDS/PAGE, transferred on to a nitrocellulose membrane and immunoblots were performed as described above using rabbit anti-XBP-1 (Abcam) and p85α (Santa Cruz Biotechnology).
Results are expressed as means±S.E.M., and n represents the number of mice or biological replicates. Statistical analyses were performed using ANOVA (two-way or one-way, as appropriate), and Mann–Whitney U tests. P<0.05 was considered to be statistically significant.
ER-stress induction leads to increased PTP1B protein and mRNA expression levels
We aimed to investigate the impact of ER-stress induction on hepatic PTP1B expression in cultured hepatocytes and mice. In HepG2 cells, tunicamycin treatment led to a ~2-fold increase in PTP1B mRNA expression and protein levels, but not in those transfected with siRNA against PTP1B (Figures 1A and 1B). Chronic ER-stress , caused by prolonged HFD feeding for 16 weeks, led to an increase in hepatic-PTP1B protein levels compared with the mice fed a standard chow diet (Figure 1C). Interestingly, acute pharmacological induction of ER-stress in mice with tunicamycin treatment also resulted in an increase in hepatic PTP1B protein expression (Figure 1C). The increase in mouse liver PTP1B protein levels was due to increased mRNA levels with both HFD feeding and tunicamycin treatment (Figures 1D and 1E). These results indicate that pharmacological induction of ER stress by tunicamycin treatment leads to increased PTP1B mRNA and protein levels, both in vitro and in vivo. Importantly, tunicamycin-induced ER-stress did not affect expression of other phosphatases such as SHP2 or PTEN (Figures 1F and 1G), although there was a trend towards an increase in SHP2 protein levels with HFD feeding in comparison with chow-fed animals (Figure 1F).
PTP1B knockdown protects against pharmacologically induced ER stress in HepG2 cells
Since PTP1B siRNA decreased PTP1B mRNA expression in cells by ~75% compared with scrambled control siRNA (Figures 1A and 1B), we used these cells to analyse ER-stress activation. Interestingly, as shown in Figure 2(A), tunicamycin treatment of HepG2 cells transfected with control siRNA enhanced eIF2α phosphorylation, whereas PTP1B silencing prevented this. In addition, tunicamycin treatment led to XBP-1 splicing in scrambled siRNA- and PTP1B siRNA-transfected cells (Figure 2B); however, PTP1B-silenced cells exhibited lower amounts of spliced XBP-1 (Figure 2B). Interestingly, the tunicamycin-induced increase in CHOP protein levels was lower in PTP1B-silenced cells compared with controls (Figure 2C). Furthermore, we analysed mRNA expression levels of the main markers of ER-stress response by real-time PCR. As expected, we found that treatment of control cells with either tunicamycin or thapsigargin led to enhanced expression of ER-stress response markers regulated through the PERK/eIF2α pathway (CHOP, ATF4 and GADD34) and through IRE1-α and ATF6 pathways (BIP, GRP94 and ERDJ4) (Figures 2D–2K). Importantly, silencing of PTP1B in these cells partially (CHOP and BIP) or completely (ATF4, GADD34, GRP94 and ERDJ4) protected against the effects of tunicamycin, as well as thapsigargin, on the main ER-stress response components, suggesting that PTP1B may be required for the full induction of ER-stress response signalling (Figures 2D–2K).
In vivo liver PTP1B deficiency and glucose/lipid homoeostasis
To better understand the physiological role of hepatic PTP1B in ER-stress response signalling, we used an in vivo mouse model for the rest of our present study. We used the L-PTP1B−/− mice, as described previously [16,22]. L-PTP1B−/− and control mice were weaned either on to normal chow diet (4.5% fat) or HFD (55% kcal from fat). The absence of PTP1B in hepatocytes had no effect on the body weight on either diet (results not shown), and as published previously [16,22].
To analyse this mouse model further, we investigated whether L-PTP1B played a role in fasting/re-feeding control of glucose homoeostasis. We fasted the mice overnight (at 8 weeks on the HFD), and allowed them free access to food for 1 h afterwards. After an overnight fast, L-PTP1B−/− exhibited markedly lower blood glucose levels (Figure 3A), circulating insulin levels (Figure 3B) and HOMA (homoeostasis model assessment)-IR index (Figure 3C), consistent with some of our previous observations . PTP1B, however, does not seem to play an important role in the fasting/re-feeding response of glucose maintenance, as glucose levels increased to the same level in both groups of mice (Figure 3A) and, although L- PTP1B−/− had a moderately lower increase in re-feeding circulating insulin levels (Figure 3B) and HOMA-IR index (Figure 3C) in comparison with controls, this was not statistically significant.
We also showed previously that liver cholesterol and TAG (triacylglycerol) levels were significantly lower in L-PTP1B−/− mice after 5 weeks of HFD feeding . Since the ER-stress response is known to play a major role in lipid regulation , we wanted to investigate expression of genes involved in synthesis of fatty acids. De novo synthesis of fatty acids requires SREBP1 (sterol-regulatory-element-binding protein 1), and PPARγ (peroxisome-proliferator-activated receptor γ) . Analysis of SREBP-1c expression levels, as well as its target gene FAS (fatty acid synthase), and PPARγ revealed that HFD-fed L-PTP1B−/− mice had lower expression levels of these genes relative to control littermates (Figures 3D–3F). Expression of genes involved in cholesterol synthesis, such as HMGCS1 (3-hydroxy-3-methylglutaryl-CoA synthase 1) was also significantly decreased in HFD-fed L-PTP1B−/− mice (Figure 3G), as expected from our published phenotype of these mice. Interestingly, however, fatty acids are catabolized to acetyl-CoA for use in energy production mainly by β-oxidization in mitochondria and peroxisomes and PPARα is a known master transcriptional regulator of genes involved in β-oxidization in mitochondria and peroxisomes . We found that HFD-fed L-PTP1B−/− mice also had significantly lower mRNA expression levels of PPARα when compared with control mice (Figure 3H). Collectively, these results indicate that, in our hands, L-PTP1B−/− mice exhibit improved glucose and lipid homoeostasis compared with their control littermates, and that these mice are a useful tool to dissect out the in vivo role of PTP1B in ER-stress response signalling.
Liver PTP1B deficiency decreases the ER-stress response regulated through the PERK/eIF2α and ATF6 pathways
In light of the present findings in cells that hepatic PTP1B may be required for full induction of the ER-stress response, we used L-PTP1B−/− mice to delineate the in vivo role of hepatic PTP1B in the ER-stress response regulated through different branches of the pathway. Mice fed chow or a HFD for 16 weeks were killed and livers harvested for analysis of ER-stress response components. First, we analysed the involvement of L-PTP1B in regulation of ER-stress response via the PERK/eIF2α pathway. Although HFD feeding led to an increase in phosphorylation of eIF2α at Ser51 in control mice in comparison with chow-fed mice (Figure 4A), L-PTP1B deletion protected against this. Since phosphorylation of eIF2α is regulated through PERK activation, this suggests that there is a lower activity of the PERK/eIF2α pathway in L-PTP1B−/− mice (Figure 4A). In addition, HFD feeding, as expected, enhanced mRNA expression levels of several components of the ER-stress response under the control of the PERK/eIF2α pathway: CHOP, ATF4 and GADD34 (Figures 4C–4E). Importantly, L-PTP1B deletion partially or completely prevented the HFD-induced increase in protein expression of CHOP (Figure 4B) and mRNA expression of CHOP, ATF4 and GADD34 (Figures 4C–4E).
The second arm of the ER-stress response pathway is mediated by ATF6. To investigate ATF6 activation, we analysed the expression of the full-length (90 kDa) and the active cleaved (50 kDa) forms of ATF6 by Western blotting. HFD feeding enhanced cleavage of ATF6 and reduced expression of its full-length form in control animals (Figure 5). Importantly, L-PTP1B deletion protected against HFD-induced ATF6 cleavage (Figure 5), suggesting that L-PTP1B is also required for maximal cleavage of ATF6 with HFD feeding.
Liver PTP1B deficiency decreases the ER-stress response regulated through the IRE1-α sensor
Next we analysed the involvement of hepatic-PTP1B in the IRE1α pathway. HFD feeding enhanced the expression of IRE1-α mRNA in livers from both L-PTP1B−/− and control mice compared with chow animals (Figure 6A). Its phosphorylation was also unaffected by L-PTP1B deficiency (Figure 6B). However, we observed that chronic (Figure 6C) induction of ER stress by HFD feeding increased the protein expression of the spliced form of XBP-1 (XBP-1s) in control fl/fl mice, whereas L-PTP1B−/− mice were protected (Figure 6C). In addition, tunicamycin-injected fl/fl mice also had increased protein expression of XBP-1s, whereas L-PTP1B−/− mice were completely protected against this (Figure 6D).
HFD feeding also increased protein levels of BIP and mRNA expression of EDEM1, ERDJ4 and GRP94 (Figures 6E–6H) in fl/fl mice compared with their respective chow groups; however, L-PTP1B−/− mice had dramatically lower protein levels of BIP and mRNA expression levels of EDEM1 and ERDJ4, but not GRP94, in comparison with their controls (Figures 6E–6H).
Overall, these results suggest that L-PTP1B also plays a role in the HFD feeding and pharmacologically induced ER-stress response pathway downstream of the IRE1-α sensor, but may not be required for its full induction.
L-PTP1B deficiency increases XBP-1 nuclear translocation through p85α interaction
Recently, the PI3K (phosphoinositide 3-kinase) regulatory subunit p85 was identified as an interacting partner with XBP-1 and a main modulator of its translocation to the nucleus to resolve ER stress [27,30]. L-PTP1B deficiency is associated with increased IRS (insulin receptor substrate) 1 and IRS2 association with p85 ; thus we assumed that PTP1B may also play a role in the liver regulation of the p85α–XBP1 interaction. We immunoprecipitated XBP-1 (using a goat anti-XBP-1 antibody) from cytosolic and nuclear fractions, followed by blotting for p85α. Interestingly, in liver lysates from L-PTP1B−/− mice, XBP-1s nuclear translocation was enhanced (Figure 6I) and p85α co-immunoprecipitated with XBP-1 both in the nuclear and cytosolic fractions. However, the amount of p85α interacting with XBP-1 was greater in liver lysates from L-PTP1B−/− mice (Figure 6I) in comparison with controls. It is important to note here that in order to capture this interaction, we used a rabbit anti-XBP-1 antibody for immunoblotting, in order to avoid interference with the goat heavy chain immunoglobulin that runs closely to the molecular mass of spliced XBP-1 (~54 kDa) (Supplementary Figure S1 at http://www.BiochemJ.org/bj/438/bj4380369add.htm). Importantly, the total levels of p85α were unchanged between genotypes (Figure 6J), in concurrence with our previously published data . To assess the purity of nuclear and cytosolic fractions we blotted for a nuclear marker, lamin A/C, and for ERK1 as total loading control (Figure 6K).
The pathogenesis of obesity and associated insulin resistance is thought to involve overactivation of the ER-stress response signalling [17,18,31,32]. Whole-body and tissue-specific knockout studies in mice have revealed that PTP1B is a major regulator of insulin sensitivity and adiposity via regulation of insulin and leptin signalling pathways in muscle, liver, fat and hypothalamus [16,20–22]. In the present study, we demonstrate that the ER-stress response and PTP1B expression are interlinked and that directly down-regulating PTP1B expression in liver can relieve overactivation of the ER-stress response associated with HFD feeding, obesity and insulin resistance.
Previous studies have shown that PTP1B is overexpressed in multiple insulin- and leptin-responsive tissues in mice with diet-induced or genetic obesity or in cells with pro-inflammatory cytokine or NEFA (non-esterified fatty acid; free fatty acid) treatment [33–35]. We now show that in addition to chronic HFD feeding, acute pharmacological induction of the ER-stress response leads to an elevation of PTP1B mRNA and protein levels in mouse livers and hepatic cell lines. The mechanism(s) for this elevation may involve ATF6, since overexpression of ATF6 in hepatic cells reportedly increases mRNA expression of PTP1B  and we show in the present study that the active (cleaved) form of ATF6 is increased in HFD-fed mice. These findings suggest that overactivation of the ER-stress response with diet-induced obesity promotes insulin resistance via elevation of PTP1B. However, PTP1B also appears to contribute to the ER-stress response since liver-specific deficiency in mice or siRNA knockdown in hepatic cells blunts the full activation of all three ER-stress pathways, namely IRE1-α/XBP-1, PERK/eIF2α and ATF6 ([16,37] and the present study). Collectively, these studies suggest that there may be a positive-feedback loop between PTP1B expression and full activation of the ER-stress response.
We have shown previously that liver-specific deficiency of PTP1B enhanced insulin-stimulated IRS1- and IRS2-associated p85α protein, without changes in p85α expression levels . In the present paper, we report that L-PTP1B deletion increases the interaction between p85α and XBP-1 in both cytoplasmic and nuclear fractions from liver, and increases the nuclear translocation of XBP-1s in association with increased interaction with p85α. This is despite finding that mRNA levels of XBP-1s are decreased in L-PTP1B−/− mice under HFD- and tunicamycin-treated conditions (the present study and ). These findings are consistent with recent studies that have revealed a novel link between the IRE1-α pathway and insulin signalling [27,30]. These studies identified the p85 regulatory subunit of PI3K as an interacting partner with XBP-1 and a major modulator of its translocation to the nucleus to resolve ER stress [27,30]. Thus it appears that PTP1B may play a direct role in inhibiting insulin-stimulated XBP-1 nuclear translocation via p85 to promote ER-stress. However, further work is required to elucidate whether this is the case or secondary to its role in regulating glucose and lipid homoeostasis.
The activation of the IRE1-α/XBP-1 and PERK/eIF2α pathways in obese and insulin-resistant states has been well described [16–18,38]. Moreover, Gu et al.  showed that PTP1B plays an essential role in potentiating IRE-1-mediated ER-stress signalling pathways in vitro. Consistent with these and our previous findings, we show in the present study that L-PTP1B deletion decreases eIF2α phosphorylation in HFD-fed mice, indicating a reduced activation of the PERK/eIF2α axis, confirmed further by reduced mRNA levels of the target genes ATF4, CHOP and GADD34. Moreover, we now also show that HFD feeding results in increased processing of ATF6 into its cleaved active form and that this is reduced to chow levels in HFD-fed L-PTP1B−/− mice. Downstream, GRP78/BIP is co-ordinately up-regulated in control fl/fl HFD-fed mice, but completely reduced to chow diet levels in HFD-fed L-PTP1B−/− mice. This is consistent with ATF6α being solely responsible for transcriptional induction of ER chaperones, e.g. GRP78/BIP . Kammoun et al.  also found increased levels of cleaved ATF6 in livers of ob/ob mice. However, in other studies, cleaved ATF6 levels were found to be reduced in ob/ob, db/db and diet-induced obese mice and p85α-deficient cell lines with reduced nuclear XBP-1 [30,40]. Thus further work is required to elucidate the exact role of ATF6 in insulin-resistant states.
PTP1B overexpression has been reported in other insulin-sensitive tissues following HFD feeding or in cells after treatment with ER-stress inducers. It has been shown that treatment of C2C12 muscle cells with palmitate, a known inducer of insulin resistance and ER stress, enhanced both mRNA expression and protein levels of PTP1B . Moreover, PTP1B overexpression was observed in liver and white adipose tissue in ob/ob mice and in muscle and white adipose tissue in Zucker fatty rats (fa/fa), as well as in livers and white adipose tissue in mice fed HFD . In addition, Zabolotny et al.  showed that treatment of cultured hepatocytes and 3T3-L1 adipocytes with TNFα (tumour necrosis factor α), reported to induce ER stress , enhanced PTP1B expression. Also, Bettaieb et al.  suggested that PTP1B is involved in palmitate-induced ER-stress in MIN6 insulinoma β-cells. Altogether, these findings suggest a close relationship between chronic or acute ER-stress induction and PTP1B overexpression in other insulin-sensitive tissues; however, the involved mechanisms are not yet elucidated.
Further to our previous findings , we demonstrate in the present study that PTP1B deletion in liver decreases hepatic SREBP-1c and SREBP-1a gene expression levels, which is counterintuitive to what would be expected from the liver insulin-receptor-knockout phenotype and the enhanced insulin sensitivity observed in L-PTP1B−/− mice. It has been shown that silencing PTP1B in ob/ob mice resulted in a similar decrease in lipogenic gene expression, including SREBP1 [44,45]. Moreover, studies using high-fructose-diet treatment in rats showed that this led to the development of insulin resistance with concomitant increases in PTP1B and SREBP1 gene expression in the liver. Further investigation revealed that PTP1B may regulate SREBP1a and SREBP1c mRNA expression via PP2A (protein phosphatase 2A) activity . PTP1B tethers to the ER via its C-terminal tail , and changes in the intracellular localization of PTP1B induced by truncation of its C-terminal tail did not affect its negative regulation of insulin signalling . Importantly, Shi et al.  observed that overexpression of C-terminal truncated PTP1B in rat Fao cells did not induce SREBP1 gene expression. Furthermore, truncated PTP1B failed to bind PP2A, resulting in impaired PP2A activation. Therefore it seems that PTP1B may affect SREBP1 gene expression via a pathway distinct from the insulin signalling where its location within the ER membrane appears critical.
Altogether, our present findings demonstrate that PTP1B plays a critical role in hepatic glucose and lipid homoeostasis and in the positive regulation of the ER-stress response during diet-induced obesity. Therapeutically, inhibiting the activity of PTP1B in the liver as demonstrated previously with antisense oligonucleotide administration should result in improved glucose and lipid homoeostasis. The mechanism of this improvement appears not only to involve improved insulin sensitivity through the effects on the insulin-receptor signalling, but also alleviation of ER stress. Thus developing L-PTP1B inhibitors continues to be an attractive option for treatment of metabolic and cardiovascular diseases such as insulin resistance, Type 2 diabetes and dyslipidaemia.
The experiments were designed by Mirela Delibegović and performed by Abdelali Agouni, Nimesh Mody, Carl Owen, Alicja Czopek, Derek Zimmer, Mohamed Bentires-Alj and Kendra Bence. Data analysis of the experiments was done by Abdelali Agouni and Mirela Delibegović. The paper was written by Abdelali Agouni, Mirela Delibegović, Nimesh Mody and Kendra Bence.
This work was supported by Diabetes UK [project grant number BDARD08/0003597 (to M.D.)], and also by RCUK (Research Councils U.K.) Fellowship, British Heart Foundation, Tenovus Scotland and the Royal Society (to M.D.). N.M. is funded by a career development fellowship from the British Heart Foundation. C.O. is funded by the BBSRC (Biotechnology and Biological Sciences Research Council) postdoctoral training studentship. K.K.B. is funded by the USDA (United States Department of Agriculture) and National Institutes of Health [grant number RO1DK082417].
Abbreviations: ATF, activated transcription factor; BIP, binding immunoglobulin protein; CHOP, CCAAT/enhancer-binding protein homologous protein; EDEM, ER degradation-enhancing α-mannosidade-like protein; eIF2α, eukaryotic initiation factor 2α; ER, endoplasmic reticulum; ERAD, ER-associated degradation; ERDJ4, ER-localized DnaJ homologue; ERK, extracellular-signal-regulated kinase; FAS, fatty acid synthase; GADD34, growth-arrest and DNA-damage-inducible protein 34; GRP, glucose-regulated protein; HFD, high-fat diet; HOMA, homoeostasis model assessment; HMGCS, 3-hydroxy-3-methylglutaryl-CoA synthase; IRE1, inositol-requiring enzyme 1; IRS, insulin receptor substrate; L-PTP1B−/−, liver-specific PTP1B−/−; PI3K, phosphoinositide 3-kinase; PERK, PKR (double-stranded-RNA-dependent protein kinase)-like ER kinase; PPAR, peroxisome-proliferator-activated receptor; PP2A, protein phosphatase 2A; PTEN, phosphatase and tensin homologue deleted on chromosome 10; PTP, protein tyrosine phosphatase; shRNA, short-hairpin RNA; siRNA, short interfering RNA; SREBP1, sterol-regulatory-element-binding protein 1; UPR, unfolded protein response; XBP-1, X box-binding protein-1; XBP-1s, spliced form of XBP-1
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