Tau Loss of Function,by Deletion or Aggregation,Contributes to Peripheral Insulin Resistance

Animals

Ten-month-old male C57BL/6, homozygous P301 L (JNPL3) (Taconic farms) and Tau KO (The Jackson Laboratory) mice were used. The JNPL3 mouse model [56] expresses a frontotemporal dementia-associated human mutant tau transgene (P301 L) and has been shown to develop tau aggregates in different regions in the brain as early as 4.5 months. Tau KO mice do not express tau in the whole body. Mice were housed at the UT Health Science Center at Houston animal care facility and maintained according to USDA standards (12 h light/dark cycle, food and water ad libitum), in accordance with the Guide for the Care and Use of Laboratory Animals (National Institutes of Health).

Preparation of tau aggregates

Recombinant human tau 2N4 R was produced in E. coli BL21(DE3). Bacteria pellets were lysed in 20 mM PIPES pH 6.5, 500 mM NaCl, plus protease inhibitor cocktail complete (Roche) and sonicated with a 1/2” probe (S-4000, Misonix). Lysates were then heated at 95°C for 30 min followed by centrifugation at 15,000×g for 20 min at 4°C to remove debris. Protein was precipitated by adding ammonium sulfate at 55% w/v and incubated at room temperature for 1 h. Precipitated protein was recovered by centrifugation at 15,000×g at room temperature, pellets were stored at –20°C.

Tau protein was then purified by Cation Exchange Chromatography in an AKTA pure Chromatography System. Ammonium sulfate pellets were dissolved in 20 mM PIPES pH 6.5 and the solution was filtered through 0.2μm filter. The sample was applied to a Hitrap SP HP column and eluted in a linear salt gradient (50–1000 mM NaCl, 20 mM PIPES pH 6.5). The content of tau protein was followed at 260 nm absorbance and checked by SDS-PAGE and Blue Coomassie staining. Fractions containing tau protein were pooled and dialyzed overnight in 10 mM HEPES buffer pH 7.4, 100 mM NaCl. Tau protein was concentrated using centrifugal filters (10 kDa MW CO, Millipore), and then filtered in 100 kDa centrifugal filters to remove precipitated or aggregated protein. Protein concentration was determined by BCA protein Assay (ThermoScientific, Waltham, MA). Aliquots were prepared at 5 mg/ml and stored at –80°C.

Tau filaments were prepared by incubating monomeric tau protein at 50μM with 25μM heparin (average MW 18,000, Sigma-Aldrich, St. Louis, MO) in 10 mM HEPES buffer pH 7.4, 100 mM NaCl for 6–120 h at 37°C with constant shaking at 500 rpm in a thermomixer (Eppendorf). The formation of amyloid filaments was followed by Thioflavin T fluorescence (excitation/emission wavelengths: 435/485 nm). Tau oligomers were collected at 18 h, time at which ThT fluorescence was still at lag-phase levels and aggregates did not have a filamentous morphology by TEM imaging.

Characterization of tau aggregates

Glucose uptake cell-based assay

Basal as well as insulin-stimulated glucose uptake were measured using Glucose uptake cell-based assay kit from Cayman Chemical (Cayman chemical # 600470, Ann Arbor, MI) according to manufacturer’s instructions. Briefly, cells were seeded at 30,000 cells/well in a 96-well black, clear bottom plate and incubated overnight. The next day, cells were incubated with 100 nM insulin and 2-deoxy-2-[(7-Nitro-2, 1, 3-benzoxadiazol-4-yl) Amino]-D-glucose (2-NBDG, final concentration 200μg/ml) for 8 h in glucose-free Williams’ E culture medium (US biological, Salem, MA). Control cells were incubated with only 2-NBDG for 8 h. Wild type (C57BL/6) cells exposed to TauO treatment were treated with different concentrations of TauO (0.25, 0.5, 1.0μM) for 24 h, with the last 8 h exposed to 2-NBDG alone or with insulin. At the end of the treatment, plates were centrifuged for 5 min at 400 x g at room temperature, and the supernatants were aspirated. 200μl of cell-based assay buffer was added to each well carefully not to disturb the cell layer, followed by centrifugation again for 5 min at 400 x g at room temperature. Supernatants were aspirated and 100μl cell-based assay buffer was added to each well then cells were analyzed immediately using fluorescent filters (excitation/emission wavelengths: 485/535 nm). Fluorescence was measured using Gemini EM plate reader (Molecular Devices, San Jose, CA)

Cell viability assay

Wild type primary mouse hepatocytes were seeded in 96-well plate and incubated overnight. The next day, they were exposed to different concentrations of TauO (0.25, 0.5, 1μM) for 24 h followed by assessment of cell viability to evaluate TauO toxicity at the concentrations used. The cell viability was determined using MTT reagent (Thiazolyl Blue Tetrazolium Bromide, # M5655, Sigma). Briefly, cells were incubated with MTT reagent for 2 h at 37°C followed by incubation with solubilization solution overnight at 37°C. Absorbance was measured at 562 nm with SpectraMax Plus 384 plate reader (Molecular Devices, San Jose, CA). Cells viability was corrected for the culture medium background absorbance. All measurements were performed in six replicates and cell viability was calculated as the percentage of the optical density (OD) value of treated cells relative to untreated controls.

Cell toxicity assay

Wild type primary mouse hepatocytes were seeded in 96-well plate and incubated overnight. The next day, they were exposed to different concentrations of TauO (0.25, 0.5, 1μM) for 24 h followed by assessment of cell toxicity at TauO concentrations used. The cell toxicity was determined based on LDH release using The CytoTox 96® Non-Radioactive Cytotoxicity Assay (Promega #G1781, Madison, WI) according to manufacturer’s instructions. Absorbance was measured at 490 nm with SpectraMax Plus 384 plate reader (Molecular Devices, San Jose, CA). Cells toxicity was corrected for the culture medium background absorbance. All measurements were performed in six replicates and cell toxicity was calculated as the percentage of the OD value of untreated or treated cells relative to maximum LDH release controls of lysed cells.

Protein assay

Total protein concentrations were measured using BCA protein assay kit with bovine serum albumin as a protein standard (Pierce #23225, ThermoFisher Scientific, Waltham, MA).

Western blot analysis

40μg each of cytosolic (for phosphorylated Akt and GSK3β) or membrane fractions (for GLUT4 and phosphorylated IRβ) were resolved in a pre-cast NuPAGE 4–12% Bis-Tris Gel for SDS-PAGE (Invitrogen, Waltham, MA) at 80 volts for the first 10 min then 110 volts in MOPS-SDS running buffer and subsequently electro-transferred onto nitrocellulose membranes (Bio-Rad, Hercules, CA). After blocking for 1 h at room temperature with 5% nonfat dried milk/Tris-Buffered Saline with 0.1% Tween 20 (TBST), membranes were probed overnight at 4°C with antibodies against the following insulin signaling proteins; phospho-Akt (Ser473) (Cell Signaling, Catalog No. 9271), phospho-GSK3_(Ser9) (Cell Signaling, Catalog No. 9323), GLUT4 (EMD Millipore, Catalog No. 07-1404), phospho-IGF1Rβ (Tyr1135/1136)/insulin receptor β (Tyr1150/1151) (Cell Signaling, Catalog No. 3024). Immunoreactivity was detected with horseradish peroxidase (HRP)-conjugated anti-rabbit or anti-mouse IgG secondary antibodies (1:5000, Sigma). β-actin was used as a loading control. For signal generation, Amersham ECL prime western blotting detection reagent (Cytiva) was used. Protein band density was quantified using ImageJ software and normalized to that of β-actin. All densitometry results represent the mean and standard errors of the mean.

ELISA

ELISA kits used for Akt (phospho-Ser⁴⁷³, CBP1490), GSK3β (phospho-Ser⁹, CBP1567), and IRS1 (phospho-Ser³⁰⁷, CBP1090) were all colorimetric cell-based ELISA kits from Assay Biotechnology. GLUT4 colorimetric ELISA kit was from Novus Biologicals (NBP2-82175). ELISAs were performed according to manufacturer’s instructions. For cell-based ELISA kits, 20,000 cells were seeded per well into each ELISA plate and incubated overnight. After treatments, cells were washed with TBS, and fixed using 4% formaldehyde. After washing, the plates were stored at 4°C for 5 days. Fixed cells were incubated with quenching buffer for 20 min at room temperature, followed by washing. Plates were incubated with blocking buffer for 1 h at room temperature, after which, primary antibodies were added and the plates were incubated overnight at 4°C. The next day, plates were washed and immunoreactivity was detected using HRP-conjugated secondary antibodies for 1.5 h at room temperature. Finally, plates were washed and incubated with substrate solution and the reaction was stopped with stop solution. The OD was measured at 450 nm. Following the colorimetric measurement of the HRP activity, plates were washed and crystal violet cell staining solution was added for 30 min at room temperature. This staining step is necessary to stain cells in order to determine cell density for normalization. Crystal violet solution was washed well, then SDS was added and incubated for 1 h at room temperature, followed by measuring absorbance at 595 nm. Each sample was measured in duplicates. Absorbance values for samples were normalized to crystal violet absorbance values. For GLUT4 ELISA, membrane fractions were added to plate wells and incubated for 90 min at 37°C, followed by incubation with biotinylated antibody solution for 1 h at 37°C. Following washing, samples were incubated with HRP-conjugated secondary antibody solution for 30 min at 37°C. Finally, plates were washed and incubated with substrate solution for 15 min at 37°C, then the reaction was stopped with stop solution. OD was measured at 450 nm. Each sample was measured in duplicates. ELISA results were analyzed using Graph Pad Prism 9.0 software, as stated under each figure.

ROS quantification

Primary hepatocytes were plated on 96-well plates and incubated overnight. Intracellular ROS levels were determined the next day using the fluorogenic probe dichlorodihydrofluorescein diacetate (H₂DCF-DA) (Invitrogen, Eugene, OR). Briefly, cells were loaded with 50 mM H₂DCF-DA or with H₂DCFDA and 20μM SB203580 and incubated for 45 min. After incubation, DCF-DA-loaded cells were washed twice with PBS and changes in DCF fluorescence were recorded using Gemini EM plate reader (Molecular Devices, San Jose, CA) at 485 nm excitation and 528 nm emission. Results are expressed as percent of wild type mice or untreated controls. All measurements were performed in six experimental replicates.

Statistical analysis

All data are presented as mean±SEM for the number of replicates indicated under each figure. All data were analyzed using GraphPad prism 9.0 software (GraphPad Software Inc., La Jolla, CA, USA). Statistical analysis was carried out as indicated under figure legends. Results were considered significant if the p value was <0.05. Data were checked for normality using Shapiro-Wilk test. Data passed the normality test.

Tau deletion or loss of function decreases hepatocytic insulin-stimulated glucose uptake

Tau deletion has been shown to impair insulin signaling in the brain [47], and to cause impairment in glucose metabolism [15]. To address the influence of tau loss of function on peripheral insulin signaling, we evaluated the impact of tau deletion as well as tau aggregation on peripheral insulin sensitivity using primary hepatocytes isolated from Tau KO and P301 L mice, respectively, as well as wild type mice, and compared the responsiveness of these cells towards insulin. Glucose-uptake assay was utilized to test the uptake of 2-deoxyglucose (2-NBDG) with or without treatment with 100 nM insulin for 5 min. As shown in Fig. 2, primary hepatocytes isolated from 10-month-old Tau KO as well as P301 L mice show no response to insulin stimulation (Fig. 2A) as well as a significant reduction in insulin-stimulated glucose uptake compared to wild type mice (Fig. 2B), indicating insulin resistance in these cells.

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Fig. 2

Tau loss of function by deletion or aggregation induces insulin resistance in primary mouse hepatocytes. A) Primary mouse hepatocytes isolated form 10-month-old wild type mice show increased glucose uptake after insulin stimulation, while tau KO and P301L primary hepatocytes do not respond to insulin. B) Primary mouse hepatocytes isolated from tau KO and P301L mice show significantly reduced insulin-stimulated glucose uptake compared to wild type hepatocytes. Data represent mean±SEM of 8 mice, each with six experimental replicates. WB: primary hepatocytes isolated form 10 months Tau KO and P301L mice show reduced basal (C) and insulin-stimulated (D) expression of GLUT4 at the membrane. Data represent mean±SEM of 4 mice. E) Corresponding gel for C and D. F) primary hepatocytes isolated form Tau KO and P301L mice show reduced insulin-stimulated expression of P-Tyr¹¹⁵⁰IRβ at the membrane. The antibody specifically detected P-IRβ Tyr¹¹⁵⁰ phosphorylation. Data represent mean±SEM of 4 mice. G) Corresponding gel for F. β-actin was used as a loading control and band intensities were normalized to β-actin bands intensities. *p < 0.05, **p < 0.01, ***p < 0.001; One-way ANOVA, Dunnett’s post hoc test.

Tau deletion or loss of function results in impairment of insulin signaling as shown by the reduction in hepatocytic GLUT4, P-Tyr¹¹⁵⁰ IRβ, P-Ser⁴⁷³AKT, and P-Ser⁹GSK3β

Insulin stimulation of hepatocytes results in translocation of the insulin-sensitive glucose transporter GLUT4 to the membrane. Therefore, to investigate if the reduction in glucose uptake we see in Tau KO and P301 L mice hepatocytes compared to wild type hepatocytes is due to reduction of GLUT4 at the membrane of these cells, we analyzed the expression of GLUT4 at the membrane before and after insulin stimulation using western blot analysis, and we found a significant reduction in both basal as well as insulin-stimulated expression of GLUT4 at the membrane in Tau KO and P301 L mice hepatocytes compared to wild type (Fig. 2C-E). To evaluate if the GLUT4 reduction at the membrane at the distal end of insulin signaling is a result of a proximal end impairment, we analyzed the tyrosine phosphorylation of IR at the proximal end of the pathway. Using western blot analysis, we could see that both Tau KO and P301 L hepatocytes exhibit reduction in tyrosine phosphorylation of the IR compared to wild type mice after insulin stimulation (Fig. 2F, G).

Since GLUT4 translocates to the membrane from intracellular pools upon activation of the insulin signaling as a consequence of Akt activation, we investigated the insulin signaling markers Akt and GSK3β. As shown in Fig. 3, there is a significant reduction in phosphorylation and activation of Akt in hepatocytes isolated from Tau KO and P301 L compared to those isolated from wild type mice using western blot analysis (Fig. 3A, B), which was confirmed with ELISA (Fig. 3E). This was also reflected in the downstream substrate of Akt, GSK3β, where its Ser⁹ phosphorylation was also significantly reduced in Tau KO and P301 L hepatocytes compared to wild type ones using western blot analysis (Fig. 3C, D) as well as ELISA (Fig. 3F). Phosphorylation of GSK3β by Akt results in inhibition of the enzyme activity, and reduced phosphorylation would, therefore, be expected to result in an increase in its activity. These data show downregulation of insulin signaling as seen by the significant reduction in phosphorylation and activation of the IRβ upon insulin stimulation, accompanied by reduction in activation of Akt resulting in reduction in expression of GLUT4 at the membrane of these cells.

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Fig. 3

Tau loss of function, by deletion or aggregation, induces downregulation of the insulin signaling pathway in primary mouse hepatocytes. WB: A) primary hepatocytes isolated form 10 months Tau KO and P301L mice show significantly reduced insulin-stimulated P-Ser⁴⁷³AKT expression compared to wild type hepatocytes. Data represent mean±SEM of 3 mice; B) Corresponding gel for A. C) primary hepatocytes isolated form 10 months Tau KO and P301L mice show significantly reduced insulin-stimulated P-Ser⁹GSK3β expression compared to wild type hepatocytes. Data represent mean±SEM of 5 (WT, TauKO), or 4 (P301L) mice. D) Corresponding gel for C. The antibodies specifically detected AKT-Ser⁴⁷³ and GSK3β-Ser⁹ phosphorylation. ELISA: primary hepatocytes isolated form Tau KO and P301L mice show significantly reduced insulin-stimulated P-Ser⁴⁷³AKT (E) and P-Ser⁹GSK3β (F) compared to wild type hepatocytes. Data represent mean±SEM of 3 mice, each with two experimental replicates. *p < 0.05, **p < 0.01, ****p < 0.0001; One-way ANOVA, Dunnett’s post hoc test.

In accordance with impaired insulin signaling in the brain of tau KO mice [47], our data support that tau deletion leads to insulin resistance but in the periphery, besides showing that tau aggregation in P301 L mice also results in insulin resistance.

Exposure to sub-toxic concentrations of tau aggregates results in dose-dependent decrease in glucose uptake

The peripheral insulin resistance shown in tau KO and P301 L mouse models suggests that loss of functional tau, either by deletion or aggregation, induces downregulation of insulin signaling. To analyze this further, we used wild type hepatocytes to confirm the involvement of tau in peripheral insulin signaling. To characterize tau aggregates species before treatment, we used Thioflavin T fluorescence. ThT is a dye that binds to β-sheet containing amyloid proteins [60–63]. Fibrillar species contain β-sheet and therefore would show fluorescence when bound to ThT, while oligomers do not contain β-sheet so they should not emit fluorescence upon exposure to ThT dye. As seen in Fig. 4A, aggregation of tau until 18 h still does not show an increase in fluorescence of ThT. At 24 h of aggregation, we start seeing an increase in fluorescence and it increases steeply after that until it reaches a plateau. We also ran dot blot analysis of tau aggregated for different time points using MC1 antibody, and as shown in Fig. 4B, MC1 does not detect tau aggregates until species aggregated for 30 h and after, indicating the presence of fibrillar species. To confirm the presence of oligomeric species before the 24 h and the fibrillar species after 120 h of aggregation, we performed transmission electron microscopy. Tau aggregated for 12 h shows only oligomeric species (Fig. 4C), while aggregation for 18 h shows mainly oligomeric tau but with some short fibrillar species as shown in Fig. 4D. Aggregation of tau for 120 h leads to formation of mainly fibrils (Fig. 4E). Based on the characterization done, we chose to use tau aggregated for 18 h but after sonication to break down short fibrils into oligomers, as shown in Fig. 4F, for treatment of cells. Tau oligomer (TauO) has been shown by many researchers to be the toxic form of tau that is responsible for neurodegeneration in AD [64–69].

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Fig. 4

Tau aggregates characterization. A) Tau aggregates were prepared by shaking 50μM 2N4R recombinant Tau at 37°C for different times in the presence of 25μM Heparin. ThT fluorescence was used to detect the presence of fibrillar aggregates at different time points. Data represent mean±SEM of 3 samples at each time point. B) Dot blot analysis of tau aggregates after different time points of aggregation using MC1 antibody to detect fibrillary species. C-F) Electron Microscopy images of tau aggregates after 12 h (C), 18 h (D), and 120 h (E) show the presence of only oligomeric (arrows) species after 12 h aggregation, while aggregation for 18 h shows the presence of mostly oligomeric (arrows) with some fibrillar (arrow heads) species, and aggregates formed after 120 h show the presence of mainly fibrillar species. F) Sonication of the aggregates formed after 18 h results in mainly oligomeric species with no fibrillary aggregates present. Scale bar = 200 nm.

Primary mouse hepatocytes isolated from wild type C57BL/6 mice were treated with different concentrations of TauO. We used sub-toxic concentrations of tau, based on MTT (Fig. 5C) and LDH (Fig. 5D) assays, to make sure that the effect we see on insulin signaling is not due to a toxic effect of TauO on cells’ viability, but rather a specific response on insulin signaling. We treated hepatocytes isolated from wild type mice with different concentrations of TauO for 24 h and measured the effect of TauO on glucose-uptake in these cells. As shown in Fig. 5, TauO resulted in a dose-dependent reduction in basal (Fig. 5A), as well as insulin-stimulated (Fig. 5B) glucose uptake into primary wild type hepatocytes. These findings suggest TauO induction of insulin resistance in primary mouse hepatocytes. For the rest of experiments, we decided to use 0.5μM TauO for treatment.

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Fig. 5

Exposure to sub-toxic concentrations of TauO results in downregulation of insulin signaling in wild type primary mouse hepatocytes. 8-month-old wild type (C57BL/6) primary mouse hepatocytes were treated with TauO at different concentrations (0.25, 0.5, and 1μM) for 24 h followed by an insulin challenge (100 nM) for 5 min. Glucose-uptake Assay: TauO results in a dose-dependent decrease in basal (A) as well as insulin-stimulated (B) glucose uptake in wild type primary mouse hepatocytes. Cell Viability and Toxicity Assays: Cell viability (MTT assay) (C) as well as cell toxicity (LDH release) (D) assays show no toxic effect of the different concentrations of TauO on primary wild type hepatocytes after 24 h of incubation. Data represent mean±SEM of 6 mice, each with six experimental replicates. ****p < 0.0001; One-way ANOVA, Dunnett’s post hoc test. ELISA: E, F) Glucose transporter GLUT4 : 0.5μM TauO treatment results in significant reduction of basal (E) as well as insulin-stimulated (F) expression of GLUT4 at the membrane in wild type primary mouse hepatocytes. Data represent mean±SEM of 6 (E), or 7 (F) mice, each with two experimental replicates. Treatment with 0.5μM TauO results in a significant increase in phosphorylation and inactivation of IRS1-Ser³⁰⁷ (G) as well as a significant decrease in phosphorylation and activation of AKT-Ser⁴⁷³ (H). Data represent mean±SEM of 8 (G) or 4 (H) mice, each with two experimental replicates. *p < 0.05, **p < 0.01, ****p < 0.0001; Unpaired t-test.

Exposure to sub-toxic concentrations of tau aggregates results in a decrease in GLUT4 expression, reduction in phosphorylation and activation of AKT, as well as inactivation of IRS1

To confirm TauO-induced downregulation of insulin signaling pathway in primary mouse hepatocytes, we investigated the effect of TauO treatment on the glucose transporter GLUT4 and insulin signaling proteins IRS1 and Akt. We found that treatment with 0.5μM TauO resulted in significant reduction in basal (Fig. 5E) as well as insulin-stimulated (Fig. 5F) expression of GLUT4 at the membrane.

To evaluate if the reduction in membrane expression of GLUT4 is a result of downregulation of insulin signaling, we examined IRS1 phosphorylation on Ser³⁰⁷ residue, which has been shown to result in IRS1 inhibition, and we found an increase in P-Ser³⁰⁷IRS1 as seen in Fig. 5 G. To validate that this inhibition of IRS1 at the proximal end of the insulin signaling extends towards the distal end, we looked at Akt phosphorylation and activation and, in line with impairment of insulin signaling at the proximal end, we found a significant reduction in P-Ser⁴⁷³Akt levels (Fig. 5H), indicating downregulation of insulin signaling pathway. These data imply that exposure of primary mouse hepatocytes to sub-toxic concentrations of TauO results in downregulation of the insulin signaling pathway and therefore, reduction in glucose uptake into these cells upon insulin stimulation, demonstrating insulin resistance.

p38 MAPK inhibition rescues tau aggregates-induced downregulation of insulin signaling in wild type mice hepatocytes

Previous studies have shown that p38 MAPK stress-response pathway mediates ROS-induced impairment of the insulin signaling pathway in primary mouse hepatocytes [59]. p38 MAPK has also been reported to upregulate PTEN resulting in insulin resistance [70, 71], while Marciniak et al. showed that neuronal tau reduces PTEN activity [47]. Therefore, to analyze if tau loss-of-function-induced insulin resistance may be mediated through activation of p38 stress-response pathway, we used a p38 MAPK pharmacological inhibitor, SB203580, to explore if it will rescue hepatocytes from TauO-induced insulin resistance, especially since we noticed a reduction in oxidative stress levels, accompanied by increase in glucose uptake, in Tau KO and P301 L hepatocytes upon treatment with the p38 MAPK inhibitor (Supplementary Figure 1). Furthermore, we also noticed increase in phosphorylation and activation of p38 MAPK in Tau KO and P301 L mice (data not shown). Therefore, cells were treated with 20μM SB203580 for 30 min prior to their TauO or TauO-insulin treatments, in the continued presence of the inhibitor. Pretreatment with SB203580 rescued the TauO-induced reduction in glucose uptake (Fig. 6A) accompanied by rescuing of the reduction in GLUT4 expression at the membrane using ELISA (Fig. 6B), which was also shown by western blot analysis (Fig. 6C, D). This effect was associated with a significant rescuing of TauO-induced Ser³⁰⁷ phosphorylation and inactivation of IRS-1 (Fig. 6E) as well as relief of TauO-induced inhibition of Ser⁴⁷³ phosphorylation and activation of Akt (Fig. 6F). These results suggest that the TauO-induced inhibition of insulin signaling might be, at least partially, mediated through the activation of p38MAPK. This requires further studying to investigate if this was due to a direct effect between tau and p38 MAPK or through TauO-induction of oxidative stress, or maybe via another mediator, such as PTEN as shown previously in the brain [47], especially that upregulation of PTEN expression has been shown to result from p38 activation of its downstream transcription factor-2 (ATF-2) that binds to the PTEN promoter region, leading to increased PTEN transcription [71].

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Fig. 6

Exposure to the p38 MAPK inhibitor, SB203580, rescues TauO-induced downregulation of insulin signaling. Glucose-uptake: A) Treatment with 0.5μM TauO results in significant reduction of insulin-stimulated glucose uptake into primary wild type hepatocytes that is rescued by pretreatment with the p38 inhibitor, SB203580. Data represent mean±SEM of 7 (untreated, TauO treated) or 4 (SB-TauO-treated) mice. Western blot analysis: C, D) Pretreatment with SB203580 rescues TauO-induced reduction of insulin-stimulated expression of GLUT4 at the membrane in wild type primary mouse hepatocytes. Data represent mean±SEM of 4 (0, TauO) or 3 (SB-TauO, DMSO-TauO) mice. DMSO used as a vehicle control shows no effect on wild type primary hepatocytes. This effect was confirmed by ELISA (B), Data represent mean±SEM of 6 (0, TauO) or 3 (SB-TauO) mice. ELISA: E, F) Treatment with 0.5μM TauO results in a significant increase in phosphorylation and inactivation of IRS1-Ser³⁰⁷ (E) as well as a significant decrease in phosphorylation and activation of AKT-Ser⁴⁷³ (F). These effects were reversed by pretreatment with the p38 MAPK inhibitor, SB203580. DMSO used as a vehicle control shows no effect on wild type primary hepatocytes. The values are expressed as the percent of insulin-treated control. Data represent mean±SEM of 6 (E), or 7 (0, TauO, SB-TauO in F), 3 (DMSO in F) mice, each with two experimental replicates, *p < 0.05, **p < 0.01, ***p < 0.001; One-way ANOVA, Dunnett’s post hoc test.