The present study demonstrates that 3,5,3′-tri-iodothyronine (T3) in physiological dose range inhibits tumor necrosis factor α(TNFα)/Fas-induced apoptosis in mouse hepatocytes. T3 pretreatment prevented Fas-induced early stage of apoptosis signs assessed by flow cytometry analysis of the annexin V positive cell population. T3 attenuated TNFα/Fas-induced cleavage of caspase-8 and DNA fragmentation. We found that T3 exerted its anti-apoptotic effects by mobilization of several non-genomic mechanisms independent of transcriptional activity. Inhibition of protein kinase A (PKA), extracellular signal-regulated kinase (ERK), and Na+/H+ exchanger blocked T3-dependent anti-apoptotic effects indicating an involvement of these intracellular targets into T3-induced signaling cascade. Furthermore, physiological concentrations of T3, but not reverse T3, caused increases in intracellular cAMP content and activated PKA. T3 markedly induced phosphorylation of ERK. We also detected T3-dependent intracellular alkalinization that abolished TNFα-induced acidification. PKA inhibitor KT-5720 blocked T3-induced activation of ERK and intracellular alkalinization confirming the upstream position of PKA signaling. We further detected that hepatocytes from hypothyroid mice are more sensitive to TNFα/Fas-induced apoptosis than euthyroid animals in vivo. Together, these findings imply that T3 triggers PKA- and ERK-regulated intracellular pathways capable of driving and ensuring hepatocytes survival in the presence of death receptor ligand-induced damage under chronic inflammatory conditions.
Thyroid hormones (THs) are widely known regulators of growth, metabolism, and development (Feng et al. 2000, Yen 2001, Davis & Davis 2002). Multiple studies demonstrated that liver is an important target organ for THs. THs exerted mitogenic effects on hepatic metabolism, on the level of gene expression (Francavilla et al. 1994, Feng et al. 2000, Yen 2001) and on cell-cycle progression (Alisi et al. 2005, Furumoto et al. 2005). Majority of the functions of THs were mediated by the c-erbA proto-oncogenes – THs nuclear receptors, members of the steroid/thyroid hormone nuclear receptor subfamily that also function as ligand-modulated transcription factors (Yen 2001).
Beyond the abundant genomic effects, THs have been shown to initiate fast non-genomic signaling (Yen 2001, Davis & Davis 2002, Incerpi 2005). It is important that most of plasma membrane and cytosol targets of THs have been strongly circumstantially linked to mitogenic and anti-apoptotic signaling pathways (Qi et al. 1999, Laoag-Fernandez et al. 2004, Verga Falzacappa et al. 2006). For instance, THs were able to activate the Na+/H+ exchanger (NHE) and reduce intracellular acidity produced by activated sodium currents in different cells (Wang et al. 2003, D’Arezzo et al. 2004). THs induced activation of protein kinase C (PKC), protein kinase A (PKA), and mitogen activated protein kinase (MAPK) via putative G-protein-coupled receptor (GPCR; Lin et al. 1999, Yen 2001, D’Arezzo et al. 2004). Activation of the MAPK cascade by physiological concentrations of THs caused translocation of phosphorylated extracellular signal-regulated kinase (ERK) to the cell nucleus resulting in serine phosphorylation of the nuclear THs receptor (TR) and transcriptional activity (Yen 2001, Davis & Davis 2002, Shih et al. 2004).
THs may also exert a fast permissive action on the target cells’ responsiveness to other active peptide and hormones. The status of THs modulated the steady-state levels of specific G-proteins induced by β-adrenergic agonists (Daza et al. 1997). It has also been shown that THs positively regulated epidermal growth factor receptor signaling pathway (Lin et al. 1999, Di Fulvio et al. 2000, Shih et al. 2004) – established survival regulator of Fas-induced apoptosis (Gibson et al. 1999). However, the effects of THs on Fas-induced apoptosis in hepatocytes have not yet been addressed.
Death receptor ligands (DRLs) have been demonstrated to impact hepatocellular damage (Zang et al. 2000, Ding & Yin 2004). Hepatocyte apoptosis initiated by DRLs like tumor necrosis factor-α (TNFα) or Fas contributed to a variety of liver diseases, including acute liver failure, viral and inflammatory hepatitis, alcoholic hepatitis, and ischemia/reperfusion injury (Luedde et al. 2002, Ding & Yin 2004). Several intriguing clues from clinical reports alluded to possible correlation between thyroid dysfunction and inflammation-related liver disorders. It was found that hepatitis sensitizes hepatocytes to TNFα apoptosis (Ding & Yin 2004) and, on the other hand, is often accompanied by autoimmune hypothyroidism via impaired THs secretion (Antonelli et al. 2004). However, the possibility that hypothyroidism impacts the DRLs-dependent liver damage has not yet been explored given the demonstrated importance of TH status for liver regeneration (Alisi et al. 2005, Malik et al. 2005). Supporting the protective role of THs during liver injury, it has been shown that THs regulated the expression and activity of the proteins involved in the control of cell cycle, apoptosis, and growth arrest in hepatocytes after partial hepatoectomy (Malik et al. 2005). The analysis of the phosphorylated forms of MAPK, as survival and growth markers, revealed that they are induced during hepatic regeneration in euthyroid and hyperthyroid rats whereas they are negatively regulated in hypothyroid rats (Alisi et al. 2004). Therefore, considering in vivo and in vitro data, we questioned the ability of THs to influence hepatocyte survival signaling pathways.
The results of our experiments demonstrate that T3 serves as an important anti-apoptotic factor in hepatocyte survival after DRLs-initiated apoptosis. T3 mobilizes several fast-signaling mechanisms and abrogates Fas/TNF-induced nuclear fragmentation and activation of caspase-8 via increases in intracellular cAMP, induction of PKA and ERK, and maintenance of intracellular pH (pHin) in homeostasis.
Materials and Methods
3,5,3′-Tri′-iodothyronine (T3), reverse T3 (rT3), and 8-bromoadenosine 3′,5′-cyclic monophosphate (Br-cAMP) were purchased from Calbiochem (San Diego, CA, USA). MEK-1 inhibitor, U0126, was purchased from Cell Signaling Technologies (Beverly, MA, USA). bis carboxyethyl-5-(6)-carboxyfluorescein (BCECF/AM), 5-(N-ethyl-N-isopropyl)-amiloride (EIPA), actinomycin D (ActD), forskolin, amiodarone, KT-5720 (PKA inhibitor), cycloheximide, staurosporine (StSp), and digitonin were purchased from Sigma. Mouse TNFα and anti-Fas antibodies (clone Jo2) were purchased from Pharmingen (San Diego, CA, USA). Fas receptor, phospho-ERK (pERK), and ERK total antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Anti-Fas-associated death domain-like interleukin -1beta-connecting enzyme-inhibitory protein (FLIP) CT polyclonal antibodies were from StressGen Biotechnology (Victoria, BC, Canada). The anti-caspase-8 antibody that recognizes the 18 kDa active fragment of caspase-8 (p18) was from Oncogene Research Products (Boston, MA, USA). Unless otherwise specified, the rest of the chemicals used in this study were obtained from Sigma.
Animals and hepatocyte culture experiments
Mouse liver and primary hepatocyte cultures have been widely used for Fas-induced apoptosis studies (Fladmark et al. 1997, Ding & Yin 2004). Wild-type male mice 8-weeks of age were assigned to one of the following groups: (a) euthyroid (sham operated); (b) hypothyroid that had been thyroidectomized surgically, allowed to stabilize, and used only if they had gained no weight for a minimum of 1 week prior to hepatocyte preparation or further treatment. The Fas-injury model group received a single i.p. injection of 0.15 μg/g body weight Jo2 anti-Fas antibody (PharMingen, Franklin Lakes, NJ, USA) or TNFα (0.1 μg/g body weight) 6 or 10 h before liver extraction. Animals were placed under inhaled isoflurane anesthesia for isolation of blood and liver tissue. Plasma was prepared from whole blood by centrifugation at 4000 g for 30 min at 4 °C. Plasma levels of alanine aminotransferase (ALT) and aspartate aminotransferase (AST) were measured by routine clinical chemical methods using enzyme assay kits as described previously (Zang et al. 2000). The concentrations of THs (T4 and T3) in mouse blood were measured as previously described (Escobar-Morreale et al. 1996). Experiments with mice were conducted according to the National Research Council’s guidelines for the care and use of laboratory animals.
For in vitro experiments, hepatocytes were isolated from male wild-type mice by the collagenase perfusion method (Francavilla et al. 1994). Briefly, livers were first perfused in situ with oxygenated 0.5 mM EGTA in a calcium-free salt solution (8 ml/min, 37 °C for 5 min), and then with a 0.04% collagenase solution (Wako Pure Chemical Industries Co., Osaka, Japan) for 10 min. Livers were then gently minced and filtered through a polyamide mesh. Hepatocyte viability was >90%, as determined by Trypan blue dye exclusion. Hepatocytes were diluted to a concentration of 2 × 105 cells/ml and plated in 12-well plates (1 ml/well). The plates were previously coated with mouse collagen type I in Dulbecco’s modified Eagle’s medium (DMEM; GIBCO). The cells were cultured in DMEM containing 10% fetal bovine serum, 2 mM l-glutamine, 15 mM HEPES (pH 7.4), 100 U/ml penicillin, and 100 μg/ml streptomycin in a 5% CO2 incubator at 37 °C for 16 h before various treatments. Hepatocytes were pretreated with vehicle or different concentrations of T3 for various time periods. For apoptosis induction, cells were treated with ActD (0.2 μg/ml) and Jo2 (0.5 μg/ml) or recombinant mouse TNFα (10 ng/ml) as described previously (Ogasawara et al. 1993). We used a serum-free culture medium in in vitro experiments with hormones/inhibitors treatments so as to avoid the possible influences of various hormones contained in serum.
Nearly confluent plates of cells, plated at a density 2 × 106 cells per 160 mm dish, were treated with various agents and then harvested in 500 μl TNE media (40 mM Tris–HCl (pH 7.5), 140 mM NaCl, and 1.5 mM EDTA). Cells, collected into microcentrifuge tubes kept on ice, were pelleted (100 g for 5 min). The cell pellets were resuspended in 200 μl ice-cold cAMP extraction buffer (50 mM Tris–HCl (pH 7.5) and 4 mM EDTA). Extract aliquots were boiled for 10 min, and cell debris was pelleted. The [3H]cAMP RIA kit (Amersham) was used to measure the cAMP content of samples. Incubations were carried out at 4 °C for 3 h, and charcoal–dextran-treated samples were analyzed for radioactivity.
The cytosolic fractions that had been separated from cells after various treatments were used for protein kinase assays (Shih et al. 2004). The PKA activity was determined using a PKA Assay Kit, according to the manufacturer’s directions (Upstate Biotechnology, Lake Placid, NY, USA). Briefly, cells were lysed, sonicated, centrifuged, and total protein was determined. A mixture of the cytosolic fraction (5 μg protein), assay dilution buffer (pH 7.2), cAMP, PKC/CaMK inhibitor cocktail, Mg/ATP [γ-32P]ATP (5 μCi or 3000 Ci/mmol), and Kemptide (a synthetic peptide containing the consensus PKA phosphorylation sequence) as substrate was incubated at 30 °C for 10 min, in a total volume of 60 μl. A 25 μl aliquot of the reaction mixture was applied to phosphocellulose filter paper (Whatman p81) and washed repeatedly in 75 mM phosphoric acid, and dried. The radioactivity was quantitated by scintillation. The results were expressed in terms of nanomoles phosphate/minute per milligram protein. The PKA assay was also performed in the presence of the protein kinase inhibitor peptide of PKA. These values were similar to background counts obtained in the absence of Kemptide substrate, indicating the specificity of the assay (data not shown).
DNA fragmentation was demonstrated by harvesting total cellular DNA and 1.2% agarose gel electrophoresis stained with ethidium bromide. Briefly, after the indicated treatment, both adherent and detached cells were harvested, washed with PBS, and lysed in 50 mM Tris–EDTA (pH 7.4), 1% SDS, and 0.5 mg/ml proteinase K for 3 h at 50 °C. Samples were then extracted with phenol/chloroform and precipitated with ethanol. The pellet was resuspended in 20 mM Tris–EDTA (pH 8.0). After digesting RNA with RNase (0.1 mg/ml) at 37 °C for 1 h, DNA was separated by electrophoresis.
Cells were harvested and lysed by sonication in lysis buffer containing 50 mM Tris–HCl (pH 7.4), 10% glycerol, 0.05% Triton X-100, 150 mM NaCl, 1 mM dithiothreitol, 2 mM Na3VO4, 10 mM NaF, 1 mM EDTA, and protease inhibitors (Roche Molecular Biochemicals). Protein concentrations in cell homogenates were determined with DCA protein assay (Bio-Rad) using BSA as standard. Proteins were normalized prior to gel loading. Aliquots of cell lysates were resolved by 8–12% SDS-PAGE and transferred to Hybond-P membranes (Amersham). The membranes were then probed with appropriate antibodies according to the manufacturer’s standard method. The immunocomplexes were detected with an enhanced chemiluminescence PLUS kit (Amersham Pharmacia Biotech). Densitometry was performed on a Typhoon 9410 Mode Imager using the ImageQuant program (Amersham).
Annexin V staining and analysis
Annexin V-FITC binding to externalized phosphatidylserinehas been used for apoptosis detection. Cells were plated in culture plates and exposed to various pretreatments. After apoptosis induction by Fas/TNFα/ActD, the cells were harvested, washed with ice-cold PBS, and subsequently resuspended and incubated (in the dark) with 100 μl annexin V incubation reagent that includes FITC-conjugated annexin V (Trevigen, Gaithersburg, MD, USA) and propidium iodide (PI) for 30 min at room temperature. Flow cytometry analyses were conducted using Epics XL-MCL flow cytometer (Coulter, Hialeah, FL, USA) at 488 nm wavelength. The percentage of apoptotic cells was determined as a percentage of annexin V-FITC-positive cells.
Measurement of intracellular pH
For fluorimetric determination, cells (108 cells/ml) were washed twice and then loaded with 1 μM BCECF (in dimethylsulfoxide) for 30 min at 37 °C in the dark with gentle shaking in sodium HEPES buffer (D’Arezzo et al. 2004). Then the cells were washed twice and resuspended in the same buffer at 106 cells/ml, and introduced in a thermostatic cuvette at 37 °C with constant stirring for the continuous recording of the pH-sensitive fluorescence on a Perkin-Elmer LS50B luminescence spectrometer (Perkin-Elmer Corp., Norwalk, CT, USA). pHin was estimated from the ratio of the 530 nm fluorescence signals obtained at 500 nm (pH sensitive) and 450 nm (isosbestic) excitation wavelengths. Routinelyat the end of each experiment, the calibration of fluorescence was carried out by established methods (Musgrove et al. 1986, D’Arezzo et al. 2004) in high potassium HEPES buffer containing the H+/K+ ionophore nigericin (10 μg/ml). The calculations of ion concentration were carried out as described in Musgrove et al.(1986).
For cytofluorimetric pHin determination, cells were treated according to the designed protocol with or without hormones or inhibitors. The cells were then washed with media containing RPMI 1640, containing 10 mM HEPES and 2.5% serum, loaded with BCECF (as mentioned above), and analyzed using Epics XL-MCL flow cytometer as described previously (Musgrove et al. 1986). Excitation of the probe was performed at 488 nm with an argon laser. Emission fluorescence was filtered through a 525 band pass (green) and a 610 long pass (red) fluorescence filters.
All data are expressed as means ± s.d. Unpaired Student’s t-tests were used for comparison between two groups. For multiple comparisons, results were analyzed by ANOVA followed by the Dunnet’s test. P values <0.05 were considered statistically significant.
Thyroid hormones suppress TNFα/Fas-induced apoptosis in mouse hepatocytes
In our experiments, mouse primary hepatocyte culture has been treated for 4 h with death receptor ligands (DRLs) – Fas (0.5 μg/ml) or TNFα (10 ng/ml) plus ActD (0.2 μg/ml; Fladmark et al. 1997, Ding & Yin 2004). Apoptosis was assessed by flow cytometry detecting annexin V labeling of phosphatidylserine. DRLs-treated hepatocytes exhibited significantly increased annexin V binding (Figs 1 and 2).
The major form of THs secreted from thyroid gland is l-thyroxine (T4); however, most of THs actions were induced by 3,5,3′-tri-iodothyronine (T3) – the product of T4 deiodination (Yen 2001). We have assayed the effect of both T4 and T3 on Fas-induced apoptosis in hepatocytes. To detect the effect of T3/T4 on DRLs-induced apoptosis hepatocytes were pretreated with one of the hormones (150 nM) or vehicle for 0.25–6 h before stimulation with Fas/or TNFα/ActD. THs exposure decreased Fas and TNFα/ActD-induced apoptosis (Fig. 1). The preventive effect was also dose dependent and has been detected as early as 30-min T3 pretreatment. T4 demonstrated effects similar to but lower than those induced by T3. No protective effect was observed when T3 was simultaneously applied or post-applied after Fas/TNFα/ActD treatments. Application of reverse T3 (rT3) did not prevent DRL-induced apoptosis (Figs 1D and 2).
Structurally similar to biologically active THs, the potent anti-arrhythmic drug amiodarone (1 μM) used as a TR inhibitor (Van Beeren et al. 1996), prevented the T3 effects (Figs 1D and 2). However, T3 did not protect hepatocytes from 1 μM StSp-induced apoptosis (Fig. 1C) that activated the mitochondrial death pathway independently from death receptors. Our results indicate that in hepatocytes, T3 selectively inhibits DRL-mediated apoptosis, and the observed protection does not result from a general inhibitory effect on cell death.
Next, we investigated whether new protein synthesis is required for the anti-apoptotic effect of T3. As shown in Figs 1D and 2, T3-mediated protection from TNFα-induced apoptosis occurred equally well in the presence or in the absence of cycloheximide (CHX; 60 μg/ml), indicating that the T3 protective effect was independent of new protein synthesis. It was also independent of activation of transcription because we have added ActD in all experiments along with application of TNFα/Fas to sensitize hepatocytes to apoptosis (Ding & Yin 2004). Therefore, the involvement of cytoplasmic, non-genomic signaling pathways has to be considered. Indeed, anti-apoptotic and mitogenic effects of THs were associated with the activation of several non-genomic mechanisms, including activation of PKA, ERK, and NHE in different cells (Yen 2001, Davis &Davis 2002, D’Arezzo et al. 2004, Furumoto et al. 2005, Incerpi 2005). In our experiments, the THs-induced anti-Fas protective effect was reversed by pretreatment (30 min) and co-incubation with PKA inhibitor KT-5720 (500 nM), ERK activation inhibitor U0126 (10 μM), and NHE inhibitor EIPA (60 μM; Figs 1 and 2). KT-5720, U0126, and EIPA have been tested separately and did not affect cell viability or DNA integrity during overnight treatment (data not shown).
Successful inhibition of T3 effects by KT-5720 indicated an involvement of cAMP/PKA into T3-induced anti-apoptotic signaling. To elucidate whether this anti-apoptotic effect of THs is causally linked to the increases in cAMP induced by T3, we examined the effect of 100 μM 8-bromoadenosine 3′,5′-cyclic monophosphate (Br-cAMP) on Fas-induced apoptosis. Pretreatment with Br-cAMP significantly reduced the number of apoptotic cells compared with TNFα/Fas-treated controls. However, we have not observed an additive effect of Br-cAMP and T3 (Fig. 1D).
Hepatocyte apoptosis initiated by DRLs-like TNFα or Fas elicits the recruitment of several cellular pathways resulting in the activation of caspase-8 and caspase-3, translocation of Bid into mitochondria with subsequent cytochrome c release (Van Beeren et al. 1996). The cytosolic cytochrome c accumulation results in the formation of the apoptosome and DNA degradation (Ding & Yin 2004). Therefore, DNA fragmentation and caspase activation have been described as apoptosis markers in hepatocytes challenged with Fas/TNFα/ActD (Ding & Yin 2004). In our experiments, T3 prevented Fas-induced caspase-8 activation (Fig. 2). The protective effect of T3 was inhibited in the presence of KT-5720, U0126, and EIPA indicating the necessity of PKA, ERK, and NHE activation to overbalance Fas-induced apoptosis and prevent caspase-8 activation. Protective T3 effects have also been confirmed using the DNA fragmentation assay (Fig. 2B).
T3-induced increases in cAMP and PKA activation are important for prevention of Fas-induced apoptosis
In hepatocytes, cAMP protected against apoptosis induced by TNF and Fas ligands (Van Beeren et al. 1996, Fladmark et al. 1997, Li et al. 2000). To confirm the cAMP participation in T3-mediated effects, we next examined how T3 affects the cellular cAMP content. Dose–response studies revealed that low, physiological concentrations of T3, but not rT3 caused a sixfold increase in the cellular cAMP content (Fig. 3). The effect was blocked by pretreatment with amiodarone. Used as a positive control, forskolin (5 μM) increased cAMP nine times above the basal level. Intracellular cAMP levels returned to the control level by 6 h despite continued exposure to hormonal ligands.
To assess whether protein and RNA synthesis were required for the stimulatory effect of T3 on intracellular cAMP concentration, cells were pretreated with CHX and ActD for 60 min prior to the addition of the hormone. CHX/ActD-treated cells showed no inhibition of T3-induced stimulation of cAMP, although the inhibitors prevented protein and RNA synthesis by >95% (data not shown). This implies that T3-induced stimulation of cAMP accumulation does not require RNA or protein synthesis. The early time course of cAMP increase is also consistent with the apparent absence of a requirement for RNA synthesis.
In classic signaling cascades, cAMP activates PKA, a cAMP-dependent serine threonine kinase, which in turn regulates intracellular pathways via phosphorylation of signaling intermediates (Kopperud et al. 2003). Since T3 increased intracellular cAMP, we asked whether the changes in cAMP were significant enough to affect the activation of PKA. We have measured PKA activity stimulated by T3. As shown in Fig. 3, T3 induced activation of PKA that decreased to baseline levels by 24 h. The specificity of this response was confirmed by the finding that amiodarone blocked T3 effects and rT3 did not stimulate PKA activity (Fig. 3C). The T3-induced increase in PKA activity was not affected by TNFα (Fig. 3D). These data suggest that T3 signaling is a potential source of PKA activation that might be required for inhibition of DRL-dependent apoptosis in hepatocytes.
T3 activates ERK to suppress Fas-mediated apoptosis
ERK is a member of the MAPK family that participates in mitogenic signaling through complex phosphorylation cascades converting cell surface signals into nuclear transcription programs. Induction of ERK signaling by cAMP/PKA-dependent pathway has been shown (Kopperud et al. 2003, Rudolph et al. 2004). Importantly, ERK is also known to attenuate Fas-mediated apoptosis in hepatocytes (Sautin et al. 2001). We have detected that the inhibition of ERK activation by U0126 suppressed the T3-dependent protection against Fas-induced apoptosis (Figs 1 and 2).
To confirm the ERK-dependent mechanisms of T3-mediated prevention of Fas-induced apoptosis, we addressed the activation of MAPK as a potential target for T3 action. In our experiments, T3 markedly induced the phosphorylation of ERK in hepatocytes, starting at 30 min and lasting up to 10 h (Fig. 4A). The stimulation of ERK phosphorylation by T3 also occurred in the presence of the apoptotic trigger (Fas/TNFα; Fig. 4B). We have not detected any significant increases in ERK activation after time-course treatment (15 min to 4 h) with TNFα or Fas/ActD in primary mouse hepatocytes (data not shown). U0126, used as a positive control, effectively inhibited T3-dependent ERK activation (Fig. 4B). PKA inhibition by KT-5720 also prevented T3-induced ERK activation suggesting PKA activation is upstream of ERK signaling. Together, these data supported our findings that T3 inhibits Fas-mediated apoptosis in hepatocytes through activation of the ERK signaling pathway by a PKA-dependent mechanism.
T3 prevents TNFα-induced intracellular acidification
Low pH-dependent (acidic) nucleases are often involved in intranucleosomal genome degradation. Fas-ligands have been shown to induce not only DNA degradation, but also cytosole acidification via inhibition of NHE (Gottlieb et al. 1996). In our experiment, NHE inhibitor EIPA abrogated T3-dependent anti-apoptotic effects indicating the importance of pHin regulation for cell survival (Figs 1 and 2). Our data suggested that T3 effects could be mediated by activation of the NHE and subsequent intracellular alkalinization.
To assess whether T3 regulates intracellular pH in hepatocytes, we performed conventional BCECF fluorescence measurements. We analyzed the distribution of TNFα/Fas- and vehicle-treated cells according to their pHin using flow cytometry and fluorimetry. Treatment of hepatocytes with TNFα or Fas led to the appearance of a sub-population of cells with a more acidic pHin than vehicle-treated cells (Fig. 5) as it was noted previously that exposure to DRL-stimulated decreases in pHin (Gottlieb et al. 1996, D’Arezzo et al. 2004). In contrast, shift of the main cell population towards alkalinized pH was observed after the addition of physiological concentrations of T3 indicating significant rise in the pHin (Fig. 5). Using a fluorimetric assay, we confirmed these data and found that T3 induced gradual increases of hepatocyte pHin. EIPA caused suppression of the T3-induced increases in pHin. Treatment with rT3 had no effect on hepatocyte pHin. Our experiments were carried out in HCO3-free media, which renders the NHE as the main mechanism for regulation of pHin (Gottlieb et al. 1996). Control application of EIPA alone induced slow acidification over 3-h treatment (data not shown).
As evident from flow cytometry analysis of mouse hepatocytes, T3 induced cytosolic alkalinization contrary to Fas stimulation that was marked by cytosolic acidification. T3 pretreatment, but not rT3, prevented Fas effects and markedly increased pHin when compared with TNFα/Fas-treatment alone (Fig. 5B). EIPA abrogated T3-induced pH effects. These results suggest that T3 can compensate the DRLs-dependent inhibition of the NHE and prevent acute cytosolic acidification in mouse hepatocytes.
Hypothyroidism enhances Fas-induced apoptosis in vivo
The liver of mice is particularly sensitive in vivo to anti-Fas antibodies, which cause liver apoptosis with a phenotype that resembles fulminant hepatitis marked by activation of caspases (Ogasawara et al. 1993). We addressed the activation of caspase-8 in livers extracted from hypothyroid (thyroidectomized) and euthyroid mice. The thyroidectomized animals injected with placebo were severely hypothyroid with decreased T4/T3 levels (5 ± 2.4 and 0.2 ± 0.38 ng/ml plasma). T4 and T3 blood concentrations in euthyroid mice were 60 ± 14 and 1.5 ± 0.4 ng/ml plasma respectively. Anti-Fas antibodies (0.15 μg/g body weight) or TNFα (0.1 μg/g body weight) at non-lethal concentrations were injected 6 or 10 h before liver extraction. At 6 h after injection of Fas in vivo, as shown in Fig. 6, we could detect 20 ± 7% increases in activation of caspase-8 in hepatocytes from euthyroid animals. TNFα did not induce detectable caspase-8 activation under these experimental conditions. However, Fas/TNFα effect was accelerated during 10-h exposure. Nevertheless, the group of animals survived well under the experimental conditions and was relatively resistant to Fas-mediated apoptosis. Hypothyroid animals demonstrated significant adverse response to TNFα and anti-Fas treatment. We detected an increased sensitivity to Fas-mediated death by increased caspase-8 cleavage in hypothyroid when compared with euthyroid mice. The increased sensitivity to Fas antibodies suggested an intrinsic change in the Fas cell death pathway. The increased damage of hepatocytes in hypothyroid mice after anti-Fas treatment in vivo was further confirmed by the significantly elevated serum AST and ALT levels when compared with control euthyroid mice (Table 1). However, as the time passed, ASTand ALT levels increased slightly further in the euthyroid mice as well, but to a significantly lesser degree than in hypothyroid animals. These data implied that livers from hypothyroid mice are more sensitive to an otherwise non-lethal dose of anti-Fas antibodies.
These findings were not related to an enhanced cell surface expression of Fas (Fig. 6). Neither did we find a difference in the hepatic expression of cFLIP (Fig. 6B), which interferes with the autoproteolytical maturation of Fas-FADD-bound caspase-8 proximal in the Fas apoptotic pathway (Irmler et al. 1997).
THs have been shown to modulate fundamental processes associated with development, including cell proliferation, differentiation, and apoptosis in nearly all tissues (Feng et al. 2000, Yen 2001, Furumoto et al. 2005). Regarding hepatic homeostasis, it has recently been detected that thyroid status strongly impacted liver regeneration altering the expression and activity of proteins involved in the cell-cycle control (Francavilla et al. 1994, Feng et al. 2000, Alisi et al. 2005). Although THs are known to exert many of their effects via classical regulation of gene transcription, a large number of their actions occur rapidly and are unaffected by inhibitors of transcription and protein synthesis (Yen 2001, Davis & Davis 2002, Incerpi 2005). Analyzing available data indicated that thyroid status influences sensitivity to apoptosis (Lin et al. 1999, Qi et al. 1999, Antonelli et al. 2004, Laoag-Fernandez et al. 2004, Verga Falzacappa et al. 2006), we questioned whether T3-activated signaling could suppress DRLs-induced apoptosis. In this study, we found that THs counteract apoptotic effects of Fas/TNF and prevent hepatocyte apoptosis in a rapid non-genomic manner independent of transcriptional activity (Figs 1 and 2). T3 effects resulted in inhibition of DRLs-induced annexin V labeling, caspase-8 activation, and DNA degradation. As T3-mediated protection is independent of new protein synthesis, it is probable that the observed inhibition of apoptosis upstream of caspase-8 was due to post-translational mechanisms. We further analyzed the intracellular signaling by which THs modulated DRLs-induced apoptosis in hepatocytes. We found that T3 affected several intracellular targets, such as production of cAMP, PKA, ERK, and pHin.
We detected that T3 dose- and time-dependently increased cellular content of cAMP and activated PKA in hepatocytes (Fig. 3). Our observations are consistent with previous reports regarding enhanced intracellular cAMP content in different cells after THs application (Yen 2001, Shih et al. 2004). THs modulated the activity of phosphodiesterase (Morgan et al. 1982) and mimicked the effects of cAMP-inducing hormones (Muller & Seitz 1983, Daza et al. 1997). Furthermore, cAMP-induced PKA activation has been shown to mediate THs effects (Lin et al. 1999), such as potentiation of growth hormone receptor signaling in cancer cells (Shih et al. 2004). Consistently, we found that the activation of PKA is necessary for T3-initiated apoptosis prevention in hepatocytes. In our experiments (Fig. 2), the treatment of hepatocytes with Fas resulted in apoptosis induction and generation of the p18 active form of caspase-8, which are strongly inhibited in the presence of T3 or cAMP. Pretreatment of the cells with the PKA inhibitor KT-5720 abolished the ability of T3 to reduce caspase-8 activation and inhibit apoptosis. These results have been confirmed in annexin V and DNA fragmentation assays (Figs 1 and 2).
PKA and its activator cAMP are established mediators of differential sensitivity to apoptosis (Van Beeren et al. 1996, Fladmark et al. 1997, Kopperud et al. 2003, Li et al. 2000). Increases in intracellular cAMP have been shown to prevent Fas-induced apoptosis in hepatocytes (Van Beeren et al. 1996, Fladmark et al. 1997). Moreover, activated PKA induced a prolonged stimulation of hepatocyte proliferation (Kopperud et al. 2003). Beyond PKA signaling, an increase in intracellular cAMP induced distinct mechanisms of anti-apoptosis, such as phosphorylation of Bad (Harada et al. 1999), glycogen synthase kinase-3 (Feng et al. 2000), and Akt pathway (Kopperud et al. 2003). Therefore, we cannot exclude the possibility that cAMP/PKA might also regulate additional steps in the T3 signaling cascade.
There are many possible targets for a modulation of the DRL signaling, although only a few protein synthesis-independent signal-based mechanisms that regulate DRL-dependent apoptosis have been reported. In this case, MAPK/ERK signaling was shown to phosphorylate DRL receptors directly, thereby causing inhibition of its cytotoxic ability (Cottin et al. 1999). We detected ERK activation in response to different concentrations of T3 (Fig. 4). Inhibition of ERK activation by MEK inhibitor U0126 significantly prevented T3 anti-apoptotic effects. Our results are consistent with the recently published in vivo data on the analysis of the phosphorylated forms of ERK. It has been revealed that ERK phosphorylation is induced during hepatic regeneration in euthyroid and hyperthyroid rats, whereas it is negatively regulated in hypothyroid animals (Alisi et al. 2004, 2005). Supportingly, several groups found that THs initiated non-genomic induction of ERK via putative membrane GPCR (Lin et al. 1999, Shih et al. 2004). Whether T3 prevents DRLs-induced apoptosis through GPCR remains to be determined.
ERK activity might be sufficient to trigger the THs protective effect as was shown for other ERK-activating agents (Sautin et al. 2001). However, T3-induced activation of ERK was blocked by PKA inhibition (Fig. 4B). It appears that PKA activation is required for THs-promoted ERK activation and apoptosis prevention. Therefore, we suggest that the increases in intracellular cAMP and the activation of PKA are possible upstream mediators of ERK activation in response to T3. Indeed, ERK activation has been linked to cAMP production (Rudolph et al. 2004) and PKA activation (Saxena et al. 1999). Furthermore, ERK was proposed as a downstream effector of cAMP signaling linked to hepatocyte survival via activation of the small GTPases, Rap1/2 (Caron 2003).
One of the downstream targets of PKA and ERK activation is the regulation of intracellular pHin. It is well known that plasma membrane of hepatocytes has a system of membrane transporters that supports neutral pHin (D’Arezzo et al. 2004). Fas-induced apoptosis has been reported to induce cytosolic acidification via inhibition of NHE (Gottlieb et al. 1996). Acidic pHin stimulates endonuclease activity and DNA fragmentation (Gottlieb et al. 1996). Our presented results confirmed that Fas/TNF are able to induce cytosolic acidification. In contrast, T3-treated cells respond with a significant increase in pHin responsible for the recovery and/or maintenance of normal pHin (Fig. 5). We have observed that Fas-induced caspase-8 activation and DNA degradation can be prevented by T3 that is also able to block Fas-induced intracellular acidification. The T3 effect on the intracellular pH was completely blocked by EIPA – the specific NHE antiport inhibitor.
Cytosolic alkalinization and activation of NHE are known as non-genomic anti-apoptotic mechanisms. For instance, cAMP and PKA have been shown to activate NHE (Borgese et al. 1992, Caron 2003). Our results are in agreement with these studies and indicate that THs signaling is an important modulator of pHin that impacted hepatocyte survival. The raise in pHin induced by T3 should be sufficient to stop DNA degradation and/or prevent further activation of the endonuclease. We also found that T3-induced activation of PKA and ERK signaling are necessary for regulation of intracellular pHin in hepatocytes because the PKA and ERK inhibitors, KT-5720 and U0126, blocked T3-initiated pHin effects (Fig. 5).
Our results are in agreement with findings that NHE might be stimulated via the mechanism of direct phosphorylation of NHE either by ERK itself or by p90rsk, a newly described ERK-activated kinase (Moor & Fliegel 1999). Furthermore, THs were shown to activate the NHE by a rapid, non-genomic mechanism in rat skeletal muscle (D’Arezzo et al. 2004), l-6 myoblasts (D’Arezzo et al. 2004), and cat atrial myocytes (Wang et al. 2003). This T3 effect is thought to be mediated non-genomically via activation of ERK (Wang et al. 2003, D’Arezzo et al. 2004).
DRL-induced hepatic apoptosis is considered as an important cause for a variety of liver diseases, such as viral hepatitis, cholestasis, and ischemic or septic liver injury (Luedde et al. 2002). For instance, playing a primary role in inflammation TNFα mediates severe hepatocellular damage (Zang et al. 2000, Ding & Yin 2004). Therefore, the suppression of TNFα-induced hepatocyte apoptosis in clinical aspects is critical. Our in vivo data demonstrated that euthyroid mice are more resistant to TNFα/Fas-induced apoptosis when compared with hypothyroid animals (Fig. 6). The level of caspase-8 activation was increased in hypothyroid compared with euthyroid mice after Fas/TNFα injections. The significantly elevated serum AST and ALT levels in hypothyroid mice confirmed the increased hepatocyte sensitivity to Fas-induced apoptosis (Table 1). Taken together, all our results demonstrate that the THs-initiated inhibition of the DRL apoptotic signal occurs at the very proximal stages of apoptotic signaling because T3-induced signaling seems to suppress apoptosis before activation of caspase-8. Taken together, our in vitro and in vivo data indicate that T3-activated signaling pathway mediated by activation of PKA, ERK, and balanced pHin strongly enable survival of hepatocytes. Therefore, THs could be an important target in the therapeutic intervention that can block unwanted hepatocellular apoptotic cell death during inflammation and/or septic liver failure.
Effect of a single i.p. injection of anti-Fas antibodies on serum alanine aminotransferase (ALT) and aspartate aminotransferase (AST) levels in hypothyroid and euthyroid mice. Values are expressed as means ± s.d. (n = 6 for the saline-treated controls, while n = 10 for the anti-Fas-treated groups).
|Treatment||ALT (U/ml)||AST (U/ml)|
|Note: All mice were treated intraperitoneally with either saline or anti-Fas antibodies (0.15 μg/g body weight). *Statistically different from the anti-Fas-treated hypothyroid mice as determined by non-paired t-test (P<0.05).|
|Euthyroid||Saline||87 ± 43||172 ± 84|
|Anti-Fas 6 h||2178 ± 1023*||1567 ± 927*|
|Anti-Fas 10 h||4108 ± 2256||3212 ± 1986|
|Hypothyroid||Saline||116 ± 41||398 ± 276|
|Anti-Fas 6 h||12 049 ± 5698||8696 ± 3955|
|Anti-Fas 10 h||15 634 ± 5994||10 226 ± 4899|
This work was supported partially by the Edmund S Muskie/Freedom Support Act Fellowship Program (US Department of State; to O A S), Fogarty International Centre, National Institutes of Health (5D43TW00636 to D O C), and by the NYS Institute for Health and Environment. The authors declare that there is no conflict of interest that would prejudice the impartiality of this scientific work.