Abstract
LPS-induced TLR4 activation alters cellular bioenergetics and triggers proteolytic cleavage of AMPKα and HIF-1α expression in leukocytes. In human leukocytes, and more specifically neutrophils, AMPKα cleavage yields 55- and 35-kDa protein fragments. In this study, we address the mechanism by which AMPKα is cleaved and its relevance to human health. Our data indicate that AMPKα cleavage is linked to MMP9 expression and that both are required for mammalian target of rapamycin complex-1 and S6K1 activation and HIF-1α expression in LPS-stimulated human and mice leukocytes. Three key observations support this conclusion. First, no changes in AMPKα and TLR4 signaling intermediates (mammalian target of rapamycin complex-1/S6 kinase 1/HIF-1α) were detected in LPS-stimulated MMP9-deficient mice leukocytes. Second, rMMP9 cleaved human AMPKα ex vivo, producing degradation products similar in size to those detected following LPS stimulation. Third, MMP9 inhibitors prevented AMPKα degradation and HIF-1α expression in LPS-activated human leukocytes, whereas AMPK activators blocked MMP9 and HIF-1α expression. Significantly, AMPKα degradation, MMP9, and TLR4 signaling intermediates were all detected in leukocytes from patients with type 2 diabetes mellitus and patients following cardiopulmonary bypass surgery. Plasma from these two patient cohorts induced AMPKα cleavage and TLR4 signaling intermediates in healthy donor leukocytes and either a TLR4 inhibitor or polymyxin prevented these outcomes. Detection of AMPKα degradation, MMP9 expression, and TLR4 signaling intermediates described in this study in leukocytes, the most readily available human cells for clinical investigation, may provide a powerful tool for further exploring the role of TLR4 signaling in human diseases and lead to identification of new, context-specific therapeutic modalities for precision medicine.
Introduction
Toll-like receptors are a family of transmembrane pattern-recognition receptors that respond to ligands derived from pathogens and host tissues. These receptors are central to innate immune cells activation. In humans, there are 10 functional TLRs. TLR4, the most extensively studied TLR in humans (1–4), is activated by LPS (endotoxin), derived from Gram-negative bacteria, and a variety of endogenous ligands. These ligands include proteins derived from damaged tissues (4–8), proteins released by tissues that are stressed by excess nutrients (9), and saturated fatty acids (10). Despite significant progress in our understanding of TLR4 signaling, complete understanding of molecular mechanisms that underlie the earliest TLR4 induced responses in human leukocytes, and how chronic low-grade activation of these responses might contribute to human diseases, is lacking.
Analyses of tissue biopsies from critically ill patients revealed reduced ATP levels, suggesting that severe inflammation contributes to a decline in cellular bioenergetics (11, 12). Subsequent studies showed that in addition to well-described systemic responses (2, 13), LPS triggers within minutes after administration to human subjects profound metabolic changes in leukocytes, in part, through transcriptional regulation of proteins that contribute to mitochondrial ATP production (14). The transcriptional changes occur in conjunction with a decline in cellular ATP levels and dramatic changes in expression of two key regulators of cellular metabolism: AMPK and HIF-1α (15).
AMPK is a heterotrimer α–β–γ serine/threonine kinase that monitors the cellular energy status (16). A decline in cellular ATP levels and an increase in AMP/ATP and ADP/ATP ratio promote AMPK activation (17). Activated AMPK switches on ATP-generating catabolic pathways while switching off ATP-consuming anabolic pathways, such as protein synthesis (16). AMPK suppresses protein synthesis through inactivation of mammalian target of rapamycin complex 1 (mTORC1), a multisubunit complex that includes the catalytic subunit mTOR and the regulatory-associated protein of mTOR (Raptor) (18). AMPK inactivates mTORC1 by indirect and direct mechanisms. AMPK can phosphorylate TSC2, which then acts on Ras homolog enriched in brain (RHEB) to inhibit mTORC1 (19). AMPK can also phosphorylate Raptor at Ser792 (20). Phosphorylated Raptor binds 14-3-3 proteins. This interaction inhibits the binding of Raptor to mTOR, thus preventing mTORC1 activation (20). mTORC1 regulates the rate of protein synthesis through interactions with two proteins, p70 ribosomal protein S6 kinase 1 (S6K1) and eIF4E-binding protein. mTORC1 phosphorylates S6K1 at Thr389, a site required for S6K1 activation (21, 22). By shifting the balance from energy-consuming processes to energy production, AMPK improves the efficiency of cellular energy homeostasis.
AMPK activation requires phosphorylation of Thr172 within AMPKα. Liver kinase B1 (LKB1) and Ca2+/calmodulin-dependent protein kinase kinase (CaMKK) are two serine/threonine kinases that activate AMPK (23–27). Phosphatases PP2C and PP2A dephosphorylate AMPKα Thr172 triggering AMPK inactivation (28, 29). We reported recently that LPS-induced changes in cellular bioenergetics are accompanied by rapid, and yet transient, proteolytic cleavage of AMPKα (15, 30) followed by an increase in HIF-1α expression. These observations suggested the possibility that the proteolytic cleavage of AMPKα is an alternative mechanism employed by TLR4 to regulate cellular bioenergetics and HIF-1α expression.
HIF-1 is a transcription factor that regulates cellular bioenergetics by upregulating glycolysis while reducing mitochondrial activity (31, 32). HIF-1 is a heterodimer composed of HIF-1α and HIF-1β subunits. HIF-1β is expressed constitutively. Although HIF-1α is degraded rapidly under normoxic conditions, it is stabilized under hypoxic conditions. However, LPS/TLR4 stabilize the expression of HIF-1α in leukocytes under either hypoxic or normoxic conditions (31–33). The expression of HIF-1α is below detection level in mice myeloid cells (which are predominantly neutrophils). Nonetheless, myeloid cells genetically deficient in HIF-1α exhibited dramatically reduced ATP levels, indicating that HIF-1 plays a role in these cells irrespective of their activation state (34). HIF-1α deficiency also impaired cytokine production and myeloid cell migration, invasion, and bacterial-killing capacity, establishing that HIF-1 is a key regulator of both cellular bioenergetics and inflammatory responses (34, 35).
Given these data, and the immediate response of leukocytes when faced with danger signals, it seemed plausible that TLR4 might use a unique regulatory mechanism to inactivate AMPKα and induce HIF-1 activation in these cells. Indeed, our data show that AMPKα cleavage is regulated by intracellular MMP9 and that both AMPKα cleavage and MMP9 expression are required for mTORC1 activation and HIF-1α expression. Furthermore, we show that leukocytes from two patients’ cohorts express a signaling phenotype that is similar to that induced by LPS and that this phenotype is induced by a soluble TLR4 ligand present in patient’s blood. The TLR4-like signaling phenotype described in this study could provide a tool for future determination of TLR4 signaling in human diseases.
Materials and Methods
Abs, reagents, and inhibitors
The following Abs were used at the indicated dilution: Actin (A2066; 1:1000) from Sigma-Aldrich; HIF-1α (sc-10790; 1:250), AMPKα (sc-25792; 1:1000), and MMP9 (sc-10737; 1:1000) from Santa Cruz Biotechnology. p-Raptor (Ser792) (#2083; 1:1000), Raptor (#2280; 1:200), p-p70 S6 kinase (Thr389) (#9205; 1:1000), and p-AMPKα (Thr172) (#2535; 1:1000) were from Cell Signaling Technology. The source of reagents and final concentrations used are as follows: LPS (from Escherichia coli 0111:B4, Sigma-Aldrich). Where indicated, blood samples were treated with LPS at 10 ng/ml. Mice were challenged with LPS at 3 mg/kg body weight. The following were also used: CLI-095 (TAK-242; Invivogen; 3 μM), Polymyxin (Invivogen; 50 μg/ml), LY294002 (Cayman Chemical; 10 μM), rapamycin (Tocris Bioscience; 100 nM), A769662 (LC Labs; 100 nM), and metformin (Sigma-Aldrich; 10 μM). MMP2/MMP9 inhibitor I (In. 1; 240 nM), MMP2/MMP9 inhibitor IV (In. 2; 27 nM), and MMP9 inhibitor I (In. 3; 50 nM) were all from Millipore.
Human subjects
The Rutgers Health Sciences Institutional Review Board approved the study. Written informed consent was obtained from all participants prior to inclusion in the study. Patients with type 2 diabetes mellitus (T2DM) and nondiabetic patients were recruited from endocrinology clinics at Rutgers Robert Wood Johnson Medical School (RWJMS). Patient demographics are presented in Table I. Patients weighing <110 lbs, with an autoimmune disease, who have undergone major surgery in the past 3 mo, with a current infection, with a symptomatic heart disease, and patients >89 y old were excluded from the study. The cardiac surgery patients were recruited from Rutgers RWJMS and Robert Wood Johnson University Hospital. Patient demographics are presented in Table II. Premenopausal women, patients on insulin, with reoperative surgery, on preoperative steroid therapy, on nonsteroidal anti-inflammatory medication other than aspirin, and patients maintained on immunosuppressive medications or chemotherapeutic agents were excluded from the study. LPS (0.1 ng/kg) was administered in vivo as previously described (15). For in vitro studies, blood drawn into EDTA-containing tubes was separated into aliquots and treated with LPS (10 ng/ml) or the specified inhibitors for the indicated time. Leukocytes were isolated as described (15). Lysates containing equal protein amounts were analyzed by immunoblotting. Neutrophils were isolated using Ficoll-Hypaque (Sigma-Aldrich) centrifugation followed by dextran (m.w. 500,000) sedimentation. For the mixing experiments, patient blood samples were sedimented at unit gravity for 1.5 h. The upper plasma fraction was recovered, leaving the cellular fraction intact. The plasma fraction was next centrifuged for 10 min at 1800 × g to remove residual cells. The plasma was stored at −70°C. Healthy donor’s blood was either not treated or treated for 1 h with DMSO (vehicle; 0.5%), the TLR-4 inhibitor CLI-095 (TAK-242; 3 μM), the AMPKα activator A769662 (100 nM), the MMP2/MMP9 inhibitor I (In. 1; 240 nM), or the PI3K inhibitor LY294002 (10 μM). The samples were then centrifuged for 5 min at 1800 × g at 4°C. The upper plasma fraction was removed and replaced with an equal volume of patient’s plasma. The samples were rocked gently for 2 h. The healthy donor’s leukocytes were then isolated and analyzed by immunoblotting. In another set of experiments, patient plasma was treated for 2 h with polymyxin at 50 μg/ml before mixing it with healthy donor’s leukocytes.
Characteristics . | Nondiabetic . | T2DM . |
---|---|---|
Total no. | 10 | 13 |
Age (y) | 61.4 ± 4.1 | 61.1 ± 2.1 |
Sex (Male/female) | 5/5 | 10/3 |
BMI (kg/m2) | 34 ± 3.3 | 29.5 ± 0.9 |
FPG (mg/dl) | 101 ± 5.3 | 147.0 ± 14.1* |
HbA1C (%) | 5.9 ± 0.1 | 7.7 ± 0.3** |
Cholesterol (mg/dl) | 157 ± 14 | 171.0 ± 16.0 |
HDL cholesterol (mg/dl) | 49.0 ± 4.5 | 49.7 ± 2.3 |
LDL cholesterol (mg/dl) | 89.0 ± 10.4 | 96.0 ± 13.7 |
Triglyceride (mg/dl) | 94.4 ± 20.3 | 126.7 ± 11.7 |
Alanine aminotransferase (IU/l) | 28.0 ± 8.6 | 30.3 ± 6.4 |
Aspartate aminotransferase (IU/l) | 18.0 ± 2.4 | 27.3 ± 3.2 |
Characteristics . | Nondiabetic . | T2DM . |
---|---|---|
Total no. | 10 | 13 |
Age (y) | 61.4 ± 4.1 | 61.1 ± 2.1 |
Sex (Male/female) | 5/5 | 10/3 |
BMI (kg/m2) | 34 ± 3.3 | 29.5 ± 0.9 |
FPG (mg/dl) | 101 ± 5.3 | 147.0 ± 14.1* |
HbA1C (%) | 5.9 ± 0.1 | 7.7 ± 0.3** |
Cholesterol (mg/dl) | 157 ± 14 | 171.0 ± 16.0 |
HDL cholesterol (mg/dl) | 49.0 ± 4.5 | 49.7 ± 2.3 |
LDL cholesterol (mg/dl) | 89.0 ± 10.4 | 96.0 ± 13.7 |
Triglyceride (mg/dl) | 94.4 ± 20.3 | 126.7 ± 11.7 |
Alanine aminotransferase (IU/l) | 28.0 ± 8.6 | 30.3 ± 6.4 |
Aspartate aminotransferase (IU/l) | 18.0 ± 2.4 | 27.3 ± 3.2 |
Data are mean ± SEM.
*p = 0.024, **p = 0.002.
BMI, body mass index; FPG, fasting plasma glucose; HbA1C, hemoglobin A1C; HDL, high-density lipoprotein; LDL, low-density lipoprotein.
Characteristic . | n = 26 . |
---|---|
Age (y) | 68.0 ± 2.3 |
Sex (Male/female) | 15/12 |
BMI (kg/m2) | 30.0 ± 1.3 |
FPG (mg/dl) | 120.0 ± 4.6 |
HbA1C (%) | 5.9 ± 0.1 |
Cholesterol (mg/dl) | 179.0 ± 11.0 |
HDL cholesterol (mg/dl) | 55.0 ± 4.1 |
LDL cholesterol (mg/dl) | 94.8 ± 6.4 |
Triglyceride (mg/dl) | 122.7 ± 21.9 |
Alanine aminotransferase (IU/l) | 32.6 ± 6.9 |
Aspartate aminotransferase (IU/l) | 34.7 ± 9.3 |
OR time (h) | 3.3 ± 0.1 |
CPB time (h) | 1.2 ± 0.1 |
Characteristic . | n = 26 . |
---|---|
Age (y) | 68.0 ± 2.3 |
Sex (Male/female) | 15/12 |
BMI (kg/m2) | 30.0 ± 1.3 |
FPG (mg/dl) | 120.0 ± 4.6 |
HbA1C (%) | 5.9 ± 0.1 |
Cholesterol (mg/dl) | 179.0 ± 11.0 |
HDL cholesterol (mg/dl) | 55.0 ± 4.1 |
LDL cholesterol (mg/dl) | 94.8 ± 6.4 |
Triglyceride (mg/dl) | 122.7 ± 21.9 |
Alanine aminotransferase (IU/l) | 32.6 ± 6.9 |
Aspartate aminotransferase (IU/l) | 34.7 ± 9.3 |
OR time (h) | 3.3 ± 0.1 |
CPB time (h) | 1.2 ± 0.1 |
Data are mean ± SEM.
BMI, body mass index; FPG, fasting plasma glucose; HDL, high-density lipoprotein; LDL, low-density lipoprotein; OR, operating room.
In vitro MMP9 degradation assay
Recombinant Human MMP9 (purchased from R&D Systems) was activated by incubation with p-aminophenylmercuric acetate (AMPA; final concentration 1 μM; Sigma-Aldrich) for 24 h at 37°C. For controls, whole blood was divided. One part was stimulated with LPS (10 ng/ml) for 2 h. Leukocytes were isolated and lysed in RIPA buffer. The other part was used for unstimulated leukocyte isolation. The leukocytes were lysed in buffer containing 25 mM Tris-HCl (pH 7.5), 100 mM NaCl, and 0.1% (v/v) Nonidet P-40. The lysates were incubated for 30 min with AMPA at a final concentration of 1 μM, or with 2 μg rMMP9 preactivated with AMPA. The reactions were stopped with the addition of sample buffer.
Immunoprecipitation
Leukocytes were lysed with lysis buffer containing 24 mM Tris/HCl (pH 7.5), 50 mM KCl, 2 mmol MgCl2, 1 mM EDTA, 0.5% Triton X-100, 0.5 mM PMSF, and Complete protease inhibitor mixture (Roche). The lysate was precleared with 50 μl normal rabbit serum (Santa Cruz Biotechnology) and protein A/G agarose beads (Santa Cruz Biotechnology) (20 μl). AMPKα Ab (1:150) (Santa Cruz Biotechnology) or MMP9 Ab (1:1:500) (Santa Cruz Biotechnology) were added to the cell lysate and incubated with gentle rocking overnight at 4°C. Afterwards, protein A/G agarose beads (20 μl) were added to the lysates, and the samples were incubated with gentle rocking for 3 h at 4°C. Lysate was spun down, and the pellet was washed five times with 500 μl cell lysis buffer. The pellet was resuspended in 20 μl 4× NaDodSO4 sample buffer and heated to 97°C for 5 min. Subsequently, the samples were analyzed by Western blotting. The polyvinylidene difluoride membrane was blocked with 5% skim milk and incubated overnight with the specified Abs. The specific signal was amplified by HRP-conjugated secondary Abs, developed by ECL substrate (Pierce) and visualized by autoradiography.
Animal studies
Animal studies were approved by the Rutgers RWJMS Institutional Animal Care and Use Committee. Normal C57/BL6 mice and MMP9 null mice were purchased from The Jackson Laboratory. Animals were challenged with a bolus dose of LPS (i.p. injection, 3 mg/kg in 300 μl saline). The mice were sacrificed by CO2 inhalation. Blood was obtained by heart puncture, and leukocytes were isolated using the protocol used for human leukocytes (30).
Results
LPS induces AMPKα proteolytic cleavage, Raptor Ser792 dephosphorylation, S6K1 Thr389 phosphorylation, and HIF-1α and MMP9 expression in human and mice leukocytes
AMPK can inhibit mTORC1 by direct phosphorylation of its critical subunit Raptor at Ser792 (20), whereas mTORC1 phosphorylates S6K1 at Thr389, a site required for S6K1 activation (21, 22). Recently, we reported that administration of LPS to human subjects triggers proteolytic cleavage of AMPKα (63 kDa) in leukocytes, yielding two new protein bands ∼50 and 35 kDa in size (15). We surmised that AMPKα degradation contributes to AMPK inactivation. If true, AMPKα degradation should correlate with mTORC1 and S6K1 activation. Analyses of human leukocytes treated with LPS in vitro confirmed this possibility because AMPKα degradation, Raptor dephosphorylation at Ser792, and S6K1 phosphorylation at Thr389 were all detected as early as 10 min post–LPS stimulation (Fig. 1A). Though S6K1 phosphorylation at Thr389 was also seen by 10 min, it peaked 30–60 min posttreatment. HIF-1α expression, in contrast, was first detected by 90 min.
LPS-induced changes in AMPKα expression correlate with Raptor dephosphorylation at Ser792 and S6K1 phosphorylation at Thr389, both indicative of mTORC1 activation, as well as increases in MMP9 and HIF-1α expression in human leukocytes and neutrophils. In vitro and in vivo LPS-induced changes in human leukocytes and neutrophils were characterized by Western blotting. Actin served as a loading control throughout. Leukocytes were isolated from blood samples activated with LPS in vitro for the indicated time (A), or blood samples obtained from subjects 0–24 h after these subjects were challenged with LPS (B). (C) Neutrophils were isolated and then stimulated with LPS in vitro for the indicated time.
LPS-induced changes in AMPKα expression correlate with Raptor dephosphorylation at Ser792 and S6K1 phosphorylation at Thr389, both indicative of mTORC1 activation, as well as increases in MMP9 and HIF-1α expression in human leukocytes and neutrophils. In vitro and in vivo LPS-induced changes in human leukocytes and neutrophils were characterized by Western blotting. Actin served as a loading control throughout. Leukocytes were isolated from blood samples activated with LPS in vitro for the indicated time (A), or blood samples obtained from subjects 0–24 h after these subjects were challenged with LPS (B). (C) Neutrophils were isolated and then stimulated with LPS in vitro for the indicated time.
Neutrophils are the most abundant leukocyte cell type in human blood, constituting 50–65% of all circulating leukocytes at steady state. By 2 h post–LPS infusion to humans, neutrophils constitute ∼90% of all circulating blood leukocytes (15). Based on this information, it seemed reasonable that the changes detected in leukocytes challenged with LPS in vivo for 2 h (Fig. 1B) (15) reflect changes that unfold in neutrophils. Indeed, purified neutrophils stimulated with LPS in vitro (Fig. 1C) exhibited temporal changes in AMPKα and HIF-1α expression, which reproduced those seen in leukocytes challenged with LPS in vitro and in vivo (Fig. 1A, 1B).
Prior studies showed that LPS triggers a rapid increase in MMP9 transcripts and protein expression in neutrophils (36), whereas others linked AMPK activation to the regulation of MMP9 expression in mice fibroblasts (37). Hence, we next asked whether the increase in intracellular MMP9 expression is sufficiently rapid to potentially account for the change in AMPKα expression. Intracellular MMP9 was detected within 10 min in both LPS-stimulated leukocytes (Fig. 1A) and neutrophils (Fig. 1C), and its levels continued to rise, reaching a peak by 60 min. Intracellular MMP9 was also expressed in leukocytes challenged with LPS in vitro (Fig. 1B). These data establish that the onset of AMPKα cleavage and the increase in intracellular MMP9 expression are both rapid and overlap.
Recently, we reported that administration of LPS to mice triggers temporal changes in ATP levels, AMPKα, and HIF-1α expression in leukocytes (30). However, AMPKα proteolytic fragments of 50 and 35 kDa, which are seen in human leukocytes, are not detected in leukocytes from LPS-challenged mice. In this study, we examined and confirmed that as seen in human leukocytes, the changes in AMPKα expression correlate with Raptor Ser792 dephosphorylation and S6K1 Thr389 phosphorylation in mice leukocytes challenged with LPS in vivo (Fig. 2A).
Characterization of changes in protein expression/activation in leukocytes from wild-type (WT) and MMP9 knockout (KO) mice challenged with LPS in vivo. WT (A) and MMP9 KO (B) mice were challenged with LPS (3 mg/kg) for the indicated time. Each lane represents a sample obtained from a single mouse. Leukocytes obtained from a WT mouse challenged with LPS for 4 h served as a positive control (B, lane 9).
Characterization of changes in protein expression/activation in leukocytes from wild-type (WT) and MMP9 knockout (KO) mice challenged with LPS in vivo. WT (A) and MMP9 KO (B) mice were challenged with LPS (3 mg/kg) for the indicated time. Each lane represents a sample obtained from a single mouse. Leukocytes obtained from a WT mouse challenged with LPS for 4 h served as a positive control (B, lane 9).
AMPKα degradation and the increase in intracellular MMP9 expression are related events
Because AMPKα is cleaved as soon as intracellular MMP9 expression reaches detection level (Fig 1), we wanted to know whether MMP9 is required for AMPKα degradation. To this end, we challenged MMP9-deficient mice with LPS. Strikingly, AMPKα degradation, Raptor Ser792 dephosphorylation, S6K1 Thr389 phosphorylation, and HIF-1α expression were all absent in LPS-challenged MMP9-deficient mice leukocytes (Fig. 2B). These data establish that intracellular MMP9 is required for AMPKα degradation, mTORC1 and S6K1 activation, as well as HIF-1α expression in mice leukocytes activated with LPS in vivo.
As human neutrophils are not amenable to genetic manipulations, we undertook four complementary approaches to determine whether the increase in MMP9 expression and AMPKα degradation are related events. Firstly, we tested whether activated MMP9 could cleave AMPKα in vitro. An unstimulated human leukocyte lysate sample was incubated for 30 min with rMMP9, which was preactivated by incubation with p-aminophenylmercuric acetate (AMPA). A second sample was incubated with AMPA alone. Controls included unstimulated and LPS-stimulated leukocyte lysates. Activated rMMP9, but not AMPA alone, triggered AMPKα degradation and appearance of AMPKα degradation products indistinguishable in size from those seen in LPS-stimulated leukocytes (Fig. 3A, lanes 2 and 4). Secondly, we asked whether there was an association between AMPKα and MMP9. Lysates from leukocytes treated with LPS in vitro for the indicated time were immunoprecipitated with antisera directed against AMPKα, MMP9, or control antisera. A very low level of MMP9 was detected in AMPKα immunoprecipitates of leukocytes exposed to LPS for 0 or 0.5 h (Fig. 3B). Though the level of full-sized AMPKα declined with time, the abundance of MMP9 in the immunoprecipitates increased at 1 to 2 h post–LPS stimulation, likely reflecting an increase in AMPKα/MMP9 interaction as the levels of intracellular MMP9 increased. The MMP9 immunoprecipitates showed a significant increase in MMP9 expression 0.5–2 h post–LPS stimulation, but no intact AMPKα was detected in these samples. These data suggest that only a small fraction of total intracellular MMP9 interacts with AMPKα. Because the polyclonal AMPKα Abs used in this study were raised against a C-terminal end peptide, it appears that MMP9 remains associated with the C-terminal end of AMPKα even after AMPKα is cleavage. Thirdly, we examined the effect of three distinct MMP9 inhibitors on LPS/TLR4 signaling. Two of three MMP9 inhibitors prevented AMPKα cleavage and the increase in HIF-1α expression (Fig. 3C). Fourthly, we examined the effect of the PI3K inhibitor LY294002 and mTORC1 inhibitor rapamycin on LPS-induced responses. LY294002 prevented LPS-induced AMPKα cleavage and the increases in MMP9 and HIF-1α expression, whereas rapamycin suppressed HIF-1α expression (38), having no effect on AMPKα or MMP9 expression (Fig. 3D). Taken together, all four approaches support the possibility that intracellular MMP9 contributes to AMPKα degradation either directly or indirectly.
Evidence that AMPKα and MMP9 function within a feedback loop in human leukocytes. (A) To determine whether activated MMP9 could cleave MMP9 in vitro, unstimulated leukocytes were lysed (UN; lane 1), or treated for 30 min with AMPA (lane 3), or with AMPA-activated rMMP9 (2 μg; lane 4). As a control, whole-blood leukocytes were treated with LPS (10 ng/ml) for 2 h and then isolated and lysed (lane 2). (B) To ask whether AMPKα and MMP9 interact, leukocytes stimulated with LPS in vitro for the indicated time were lysed and analyzed by immunoblotting (top panel) or subjected to immnoprecipitation (IP) with antisera directed against AMPKα, MMP9, or control normal sera (NRS). MMP9 was clearly visible in AMPKα immunoprecipitates from leukocytes exposed to LPS for 1 or 2 h, but not in control NRS immunoprecipitates. Intact AMPKα was not detected in the MMP9 immunoprecipitates, suggesting that only a small fraction of total intracellular MMP9 interacts with AMPKα. (C) To examine the effect of MMP9 inhibitors on TLR4 signaling, human leukocytes were untreated (control [con]) or treated for 2 h with DMSO (vehicle; 0.5%; lanes 2 and 3), MMP2/MMP9 inhibitor I (In. 1; 240 nM; lanes 4 and 7), MMP2/MMP9 inhibitor IV (In. 2; 27 nM; lanes 5 and 8), or MMP9 inhibitor I (In. 3; 50 nM; lanes 6 and 9), and then left unstimulated (−; lanes 1, 2, and 7–9) or stimulated with LPS for 90 min (+; lanes 3–6). In. 1 and In. 2 suppressed the cleavage of AMPKα and HIF-1α expression. (D) To examine the effect of mTORC1 and PI3K inhibitors on TLR4 signaling, human leukocytes were treated for 2 h with DMSO (0.5%; lanes 1–3), rapamycin (RAPA; 100 nM; lanes 4–6), or LY294002 (LY; 10 μM; lanes 7–9), and then stimulated with LPS for 0–2 h. The mTORC1 inhibitor rapamycin inhibited the expression of HIF-1α, whereas the PI3K inhibitor LY prevented the changes in AMPKα, MMP9, and HIF-1α expression. To determine whether AMPK activation contributes to MMP9 expression, human leukocytes were untreated or pretreated with two AMPK activators, A769662 (100 nM) (E) and metformin (Met; 10 μM) (F). (E) Leukocytes were untreated (lanes 1 and 6) or treated for 2 h with A769662 and then with LPS (10 ng/ml) for the indicated time. (F) Untreated leukocytes were isolated and lysed at time 0 (lane 1) and 7 h later (lane 8). Leukocytes were treated for 4 (lanes 2–4) or 7 h (lanes 5–7) with LPS alone, LPS plus metformin (Met), or metformin alone. A769662 and metformin induced robust AMPKα phosphorylation at Thr172. Neither AMPKα cleavage nor MMP9 and HIF-1α expression were detected in leukocytes treated with A769662 plus LPS or metformin plus LPS. (G) A working model of TLR4 signaling in leukocytes. We propose that upon engagement of TLR4, PI3K is activated and contributes to an increase in MMP9 expression and AMPKα cleavage. This leads to dephosphorylation of Raptor at Ser792, enabling mTORC1 activation. Activated mTORC1 phosphorylates S6K1 at Thr389 and induces HIF-1α expression.
Evidence that AMPKα and MMP9 function within a feedback loop in human leukocytes. (A) To determine whether activated MMP9 could cleave MMP9 in vitro, unstimulated leukocytes were lysed (UN; lane 1), or treated for 30 min with AMPA (lane 3), or with AMPA-activated rMMP9 (2 μg; lane 4). As a control, whole-blood leukocytes were treated with LPS (10 ng/ml) for 2 h and then isolated and lysed (lane 2). (B) To ask whether AMPKα and MMP9 interact, leukocytes stimulated with LPS in vitro for the indicated time were lysed and analyzed by immunoblotting (top panel) or subjected to immnoprecipitation (IP) with antisera directed against AMPKα, MMP9, or control normal sera (NRS). MMP9 was clearly visible in AMPKα immunoprecipitates from leukocytes exposed to LPS for 1 or 2 h, but not in control NRS immunoprecipitates. Intact AMPKα was not detected in the MMP9 immunoprecipitates, suggesting that only a small fraction of total intracellular MMP9 interacts with AMPKα. (C) To examine the effect of MMP9 inhibitors on TLR4 signaling, human leukocytes were untreated (control [con]) or treated for 2 h with DMSO (vehicle; 0.5%; lanes 2 and 3), MMP2/MMP9 inhibitor I (In. 1; 240 nM; lanes 4 and 7), MMP2/MMP9 inhibitor IV (In. 2; 27 nM; lanes 5 and 8), or MMP9 inhibitor I (In. 3; 50 nM; lanes 6 and 9), and then left unstimulated (−; lanes 1, 2, and 7–9) or stimulated with LPS for 90 min (+; lanes 3–6). In. 1 and In. 2 suppressed the cleavage of AMPKα and HIF-1α expression. (D) To examine the effect of mTORC1 and PI3K inhibitors on TLR4 signaling, human leukocytes were treated for 2 h with DMSO (0.5%; lanes 1–3), rapamycin (RAPA; 100 nM; lanes 4–6), or LY294002 (LY; 10 μM; lanes 7–9), and then stimulated with LPS for 0–2 h. The mTORC1 inhibitor rapamycin inhibited the expression of HIF-1α, whereas the PI3K inhibitor LY prevented the changes in AMPKα, MMP9, and HIF-1α expression. To determine whether AMPK activation contributes to MMP9 expression, human leukocytes were untreated or pretreated with two AMPK activators, A769662 (100 nM) (E) and metformin (Met; 10 μM) (F). (E) Leukocytes were untreated (lanes 1 and 6) or treated for 2 h with A769662 and then with LPS (10 ng/ml) for the indicated time. (F) Untreated leukocytes were isolated and lysed at time 0 (lane 1) and 7 h later (lane 8). Leukocytes were treated for 4 (lanes 2–4) or 7 h (lanes 5–7) with LPS alone, LPS plus metformin (Met), or metformin alone. A769662 and metformin induced robust AMPKα phosphorylation at Thr172. Neither AMPKα cleavage nor MMP9 and HIF-1α expression were detected in leukocytes treated with A769662 plus LPS or metformin plus LPS. (G) A working model of TLR4 signaling in leukocytes. We propose that upon engagement of TLR4, PI3K is activated and contributes to an increase in MMP9 expression and AMPKα cleavage. This leads to dephosphorylation of Raptor at Ser792, enabling mTORC1 activation. Activated mTORC1 phosphorylates S6K1 at Thr389 and induces HIF-1α expression.
Studies showed that activated AMPKα suppresses the expression of MMP9 in fibroblasts (37). To examine whether this scenario is relevant to leukocytes, leukocytes were treated with two AMPK activators, A769662 and metformin (39). As expected, both A769662 and metformin triggered AMPKα phosphorylation at Thr172 (Fig. 3E, 3F). No Thr172-phosphorylated protein band was detected when leukocytes were treated with LPS alone (Fig. 3E, lane 6, and Fig. 3F, lanes 2 and 5). Significantly, phosphorylated AMPKα remained intact, and neither MMP9 nor HIF-1α expression was detected in leukocytes pretreated with A769662 or metformin and then with LPS (Fig. 3E, 3F). Collectively, these data indicate that AMPK and intracellular MMP9 function within a negative signaling feedback loop. Our working model is presented in Fig. 3G.
AMPKα proteolytic cleavage, Raptor Ser792 dephosphorylation, S6K1 Thr389 phosphorylation, and HIF-1α and MMP9 expression are all detected in leukocytes from patients with type 2 diabetes mellitus and patients after cardiopulmonary bypass surgery
To determine whether the TLR4-signaling phenotype identified in LPS-stimulated leukocytes is relevant to human health, we studied leukocytes from two patient cohorts. The first cohort included 23 patients recruited from endocrinology clinics at RWJMS. Of these, 13 had T2DM, a disease that is associated with mild, chronic activation of the immune system, and 10 had no diabetes (patient demographics are presented in Table I). Patients with T2DM were defined by hemoglobin A1C (HbA1C) ≥6.4 (40) or current treatment for diabetes. Eight of 13 (61%) patients with T2DM showed AMPKα cleavage, increases in MMP9 and HIF-1α expression, as well as dephosphorylation of Raptor at Ser792 and S6K1 phosphorylation at Thr389 (Fig 4). One patient initially classified as nondiabetic who exhibited leukocyte activation markers (patient 16) was found subsequently to have an HbA1C of 6.0% and fasting glucose of 117 mg/dL, placing this patient within the range of prediabetes (40).
Leukocytes from patients with T2DM exhibit a TLR4-like signaling phenotype. Blood samples were obtained from nondiabetic patients (N) and patients with T2DM (2) (patient characteristics are presented in Table I). The single asterisk (*, control lanes) denotes a nondiabetic patient sample used repeatedly as a control. The researchers were blinded to the patient demographics while the samples were being analyzed. Leukocyte lysates were analyzed by immunoblotting. Arrowhead denotes a nondiabetic patient with an HbA1C of 6.0% and fasting plasma glucose of 117 mg/dL.
Leukocytes from patients with T2DM exhibit a TLR4-like signaling phenotype. Blood samples were obtained from nondiabetic patients (N) and patients with T2DM (2) (patient characteristics are presented in Table I). The single asterisk (*, control lanes) denotes a nondiabetic patient sample used repeatedly as a control. The researchers were blinded to the patient demographics while the samples were being analyzed. Leukocyte lysates were analyzed by immunoblotting. Arrowhead denotes a nondiabetic patient with an HbA1C of 6.0% and fasting plasma glucose of 117 mg/dL.
Because TLR4 is activated by a variety of endogenous ligands, we reasoned that patients scheduled for elective surgery could serve as a model for assessing TLR4 responses when endogenous danger signals are likely to be present. After obtaining informed consent, we enrolled and studied a cohort of 26 patients scheduled for elective cardiopulmonary bypass (CPB) surgery for coronary or valvular heart disease (patient demographics are presented in Table II). Blood samples were obtained on the morning of the surgery, in the recovery room, and daily for up to 4 d postsurgery. Leukocytes from 20 out of 26 (77%) patients showed the signaling phenotype that is characteristic to cells treated with LPS on day 1 postsurgery (Fig. 5A, 5B). Neutrophils displayed an identical signaling profile (Fig. 5B).
Leukocytes from patients after cardiac surgery with CPB exhibit a TLR4-like signaling phenotype. Blood samples were obtained from patients (n = 26) (patient characteristics are presented in Table II) scheduled for elective cardiac surgery with CPB on the morning of the surgery (presurgery; P), in the recovery room (R), and on days 1 and 2 postsurgery (D1, D2). (A) Twenty of twenty-six patients studied exhibited AMPKα cleavage, MMP9 expression, Raptor dephosphorylation at Ser792, and HIF-1α expression on day 1 postsurgery. Patient 2 was one of six patients who did not exhibit TLR4 activation markers in their leukocytes. Actin served as a loading control throughout. (B) Leukocytes and neutrophils obtained from three patients on the morning of the surgery (P), day 1 (D1), and day 2 (D2) postsurgery exhibited an identical protein expression pattern.
Leukocytes from patients after cardiac surgery with CPB exhibit a TLR4-like signaling phenotype. Blood samples were obtained from patients (n = 26) (patient characteristics are presented in Table II) scheduled for elective cardiac surgery with CPB on the morning of the surgery (presurgery; P), in the recovery room (R), and on days 1 and 2 postsurgery (D1, D2). (A) Twenty of twenty-six patients studied exhibited AMPKα cleavage, MMP9 expression, Raptor dephosphorylation at Ser792, and HIF-1α expression on day 1 postsurgery. Patient 2 was one of six patients who did not exhibit TLR4 activation markers in their leukocytes. Actin served as a loading control throughout. (B) Leukocytes and neutrophils obtained from three patients on the morning of the surgery (P), day 1 (D1), and day 2 (D2) postsurgery exhibited an identical protein expression pattern.
A soluble and transmissible leukocyte activator is present within the plasma of patients with T2DM and CPB
We then sought to determine whether the signaling pattern of leukocytes in patients after CPB surgery was cell intrinsic or reflected presence of a circulating, and potentially transmittable, TLR4-activating ligand. To this end, we prepared plasma and cellular fractions from patients’ blood samples and conducted a series of mixing experiments. In these experiments, plasma derived from patient blood was added to and mixed with cellular fractions isolated from a healthy donor’s blood. As proof-of-concept, we mixed plasma from samples from eight patients after CPB surgery obtained on day 1 postsurgery with healthy donor’s blood cells (Fig. 6A). Four of these eight CPB patients (marked +) showed a TLR4-like signaling phenotype on day 1 postsurgery, and four did not (marked −). Following mixing, the normal donor’s leukocytes reproduced the entire TLR4-like signaling phenotype initially detected in patient leukocytes. Significantly, only plasma from CPB patients who exhibited the characteristic TLR4-like signaling phenotype in their own leukocytes induced expression of TLR4 signaling components in healthy donor’s leukocytes.
Plasma from patients after cardiac surgery with CPB and from patients with T2DM contains a soluble factor that activates TLR4 signaling in healthy donor’s leukocytes. We used a mixing approach to determine presence of TLR4 ligands in patient blood. In these experiments, blood from patients and healthy donors was separated into leukocyte and plasma fractions. The plasma fractions recovered from the patient samples were then mixed with healthy donor leukocytes. The signaling responses detected in healthy donor leukocytes reflected presence of a stimulating ligand in the patient blood/plasma. (A) As proof-of-principal, plasma fraction from eight different CPB patients was mixed with leukocytes from a healthy donor. (+) and (−) indicate, respectively, which patient did or did not show the TLR4-signaling phenotype in their own leukocytes. These data demonstrate that only plasma from (+) patients induced activation of healthy donor’s leukocytes. (B–E) Blood samples obtained from a CPB patient presurgery (P), in the recovery room (R), and on 4 consecutive d postsurgery (D1–D4) were separated into plasma and cellular fractions. (B) The patient leukocytes exhibited a protein expression pattern similar to that induced by LPS on days 1–3, but not on day 4 or prior to surgery. (C) Plasma obtained from the blood samples analyzed in (B) were mixed with healthy donor leukocytes. Following mixing, the health donor’s leukocytes exhibited the signaling phenotype first seen in the patient leukocytes. (D) Preincubation of healthy donor’s leukocytes with pharmacologic agents (P.A.) including an AMPKα activator (A769662; lane 1), an MMP2/MMP9 inhibitor (In. 1; lane 2), or a PI3K inhibitor (LY294002), prior to mixing with patient’s plasma from the sample shown in (B) lane 3, inhibited all patient plasma-induced responses. Also shown are the phenotypes of healthy donor’s leukocytes treated with DMSO alone (0.5% vehicle control; lane 4) or not treated (N.T.; lane 5). (E) Incubation of the healthy donor’s leukocytes with a TLR4 inhibitor, CLI-095, prior to mixing with patient plasma inhibited all patient plasma-induced responses. (F) Patient plasma from the samples shown in (B) were pretreated with polymyxin and then mixed with healthy donor’s leukocytes. (E and F, lane 7). Leukocytes from the sample shown in (B), lane 3, served as a positive control. (G) Blood samples obtained from a healthy donor and two patients with T2DM who exhibited a TLR4-like signaling phenotype in their own leukocytes were used in a mixing experiment. Healthy donor’s leukocytes were untreated (lanes 1–3) or were pretreated for 1 h with DMSO (0.5%; lanes 4 and 6) or CLI-095 (TAK-242; lanes 5 and 7) prior to mixing with patient’s plasma.
Plasma from patients after cardiac surgery with CPB and from patients with T2DM contains a soluble factor that activates TLR4 signaling in healthy donor’s leukocytes. We used a mixing approach to determine presence of TLR4 ligands in patient blood. In these experiments, blood from patients and healthy donors was separated into leukocyte and plasma fractions. The plasma fractions recovered from the patient samples were then mixed with healthy donor leukocytes. The signaling responses detected in healthy donor leukocytes reflected presence of a stimulating ligand in the patient blood/plasma. (A) As proof-of-principal, plasma fraction from eight different CPB patients was mixed with leukocytes from a healthy donor. (+) and (−) indicate, respectively, which patient did or did not show the TLR4-signaling phenotype in their own leukocytes. These data demonstrate that only plasma from (+) patients induced activation of healthy donor’s leukocytes. (B–E) Blood samples obtained from a CPB patient presurgery (P), in the recovery room (R), and on 4 consecutive d postsurgery (D1–D4) were separated into plasma and cellular fractions. (B) The patient leukocytes exhibited a protein expression pattern similar to that induced by LPS on days 1–3, but not on day 4 or prior to surgery. (C) Plasma obtained from the blood samples analyzed in (B) were mixed with healthy donor leukocytes. Following mixing, the health donor’s leukocytes exhibited the signaling phenotype first seen in the patient leukocytes. (D) Preincubation of healthy donor’s leukocytes with pharmacologic agents (P.A.) including an AMPKα activator (A769662; lane 1), an MMP2/MMP9 inhibitor (In. 1; lane 2), or a PI3K inhibitor (LY294002), prior to mixing with patient’s plasma from the sample shown in (B) lane 3, inhibited all patient plasma-induced responses. Also shown are the phenotypes of healthy donor’s leukocytes treated with DMSO alone (0.5% vehicle control; lane 4) or not treated (N.T.; lane 5). (E) Incubation of the healthy donor’s leukocytes with a TLR4 inhibitor, CLI-095, prior to mixing with patient plasma inhibited all patient plasma-induced responses. (F) Patient plasma from the samples shown in (B) were pretreated with polymyxin and then mixed with healthy donor’s leukocytes. (E and F, lane 7). Leukocytes from the sample shown in (B), lane 3, served as a positive control. (G) Blood samples obtained from a healthy donor and two patients with T2DM who exhibited a TLR4-like signaling phenotype in their own leukocytes were used in a mixing experiment. Healthy donor’s leukocytes were untreated (lanes 1–3) or were pretreated for 1 h with DMSO (0.5%; lanes 4 and 6) or CLI-095 (TAK-242; lanes 5 and 7) prior to mixing with patient’s plasma.
In a second series of experiments, we examined the signaling phenotype of leukocytes obtained from the blood of a patient prior to surgery as well as the 4 consecutive d following the CPB surgery. AMPKα cleavage, increased MMP9 and HIF-1α expression, and S6K1 phosphorylation were detected in blood leukocytes obtained on days 1–3 following CPB, but not on day 4 or prior to surgery (Fig. 6B), suggesting that the changes in leukocyte signaling reflect an acute response that ends by day 4. Furthermore, after mixing with plasma from the CPB patient samples shown in Fig. 6B, the healthy donor’s leukocytes expressed a similar signaling phenotype (Fig. 6C). To ask whether the signaling pathway summarized in the model shown in Fig. 3G is relevant to responses induced by patient plasma, the healthy donor’s leukocytes were pretreated with pharmacologic agents described earlier (see Fig. 3C–F), including the AMPKα activator (A769662), In. 1, and PI3K inhibitor (LY294002) prior to exposure to patient plasma obtained on day 1 postsurgery. All three pharmacologic agents, but not the DMSO control, inhibited the patient’s plasma-induced responses (Fig. 6D). Then, to determine whether the component in T2DM and CPB patient plasma that regulates leukocyte activation is a TLR4 ligand, healthy donor’s cellular fractions were treated with CLI-095 (TAK-242), a specific TLR4 signaling inhibitor (41), prior to mixing and incubation with CPB patient plasma samples from the samples shown in Fig. 6B. CLI-095 prevented the CPB patient plasma-induced changes in healthy donor’s leukocytes (Fig. 6E). To address the possibility that endogenous LPS is present in patient plasma, CPB patient’s plasma from the samples shown in Fig. 6B were treated with the antibiotic polymyxin prior to mixing with the healthy donor’s leukocytes (Fig. 6F). Polymyxin is a natural polypeptide antibiotic that binds the lipid A moiety of LPS and thus preventing LPS binding to TLR4 (42). Polymyxin prevented CPB patient plasma-induced changes in the healthy donor’s leukocytes (Fig. 6E). A mixing experiment in which plasma from two patients with T2DM was mixed with healthy donor’s leukocytes showed that the patient leukocyte phenotype could be transferred to healthy donor’s leukocytes (Fig. 6G, lanes 4 and 5) and that the TLR-4 inhibitor CLI-095 prevented these responses (Fig. 6G, lanes 5 and 7). These data link the characteristic pattern of activation of multiple indicators of TLR4 signaling to presence of a TLR4-activating factor/ligand, and possibly LPS itself, in patient plasma.
Discussion
In this study, we set to identify the mechanism by which AMPKα is cleaved in response to LPS stimulation and how this event is linked to HIF-1α expression, a key regulator of cellular bioenergetics and myeloid cell functions (34, 35). In considering which protease might cleave AMPKα, MMP9 came to mind because LPS triggers within minutes a robust increase in MMP9 transcripts and protein expression in human neutrophils (36). In addition, MMPs are known to contribute to all phases of inflammation by cleaving substrates such as cytokines and chemokines (43). The striking absence of AMPKα cleavage, Raptor Ser792 dephosphorylation, S6K1 Thr389 phosphorylation, and HIF-1α expression in MMP9-deficient mice leukocytes challenged with LPS in vivo provided the first concrete evidence that MMP9 is required for AMPKα cleavage and TLR4 signaling. We propose that intracellular MMP9 also regulates the expression of AMPKα in human leukocytes/neutrophils based on the following lines of evidence: 1) the increase in intracellular MMP9 protein expression in leukocytes treated with LPS in vivo or in vitro is sufficiently rapid (10 min) to account for AMPKα cleavage; 2) MMP9 is detected in AMPKα immunoprecipitates, indicating that the two proteins interact; 3) activated rMMP9 cleaves human AMPKα in vitro, producing two proteolytic fragments similar in size to those detected in LPS-treated leukocytes and neutrophils; 4) two of three MMP9 inhibitors, as well as the PI3K inhibitor LY294002, blocked both AMPKα degradation and HIF-1α expression; and 5) AMPK activators that triggered AMPKα phosphorylation at Thr172 prevented the expression of MMP9 in LPS-treated leukocytes. We conclude that AMPKα and MMP9 interact/function within a negative-feedback loop. When expressed, MMP9 contributes either directly or indirectly to AMPKα degradation, whereas, as seen in fibroblasts (37), activated AMPKα suppresses the expression of MMP9.
MMPs, including MMP9, are secreted and degrade extracellular matrix proteins and other extracellular substrates. The possibility that intracellular MMP9 has an intracellular function and degrades intracellular protein(s) in leukocytes is surprising, but not without precedent. Studies from Schulz (44) and Sariahmetoglu et al. (45) have demonstrated that MMP2, the MMP with the closest sequence similarity to MMP9 (46, 47), degrades a number of cytoskeletal proteins in cardiac myocytes subjected to ischemia reperfusion injury (48, 49). MMPs are commonly activated when their propeptide is removed by cleavage. However, MMP-1, -2, -8, and -9 are all also activated in the presence of peroxnitrite (ONOO−) (50, 51). Because activated leukocytes/neutrophils produce large amounts of peroxnitrite, it is plausible that this potent oxidizing compound is used not only to kill microorganisms but also to activate intracellular MMP9 once expressed in leukocytes. It is particularly interesting to note that peroxnitrite-activated MMPs can revert from an active to an inactive state in the presence of reducing agents (50). These data highlight the possible existence of a signaling mechanism that links presence of reactive oxygen species to mTORC1, S6K1, and HIF-1 activation.
The relationship between HIF-1α and mTORC1 is complex. Under chronic hypoxic conditions, HIF-1α suppresses mTORC1 through REDD1, BNIP3, or PML, which act at distinct points upstream of mTORC1 (52–56). A reversed signaling pathway, positioning mTORC1 upstream to HIF-1α, was uncovered in PC-12 cells exposed to very short periods (15–30 s) of hypoxia (referred to as intermittent hypoxia) (57). Intermittent hypoxia contributed to NADPH oxidase activation and generation of reactive oxygen species in PC-12 cells (57). When taken as a whole, the model that emerges is one in which reactive oxygen species activate MMP9, triggering AMPKα degradation, mTORC1 activation, and then HIF-1α expression. O’Neill and Hardie (58) have recently proposed that AMPK activation limits inflammation by suppressing the expression of HIF-1α. Our data support this possibility and highlight a novel mechanism by which the negative regulatory effects of AMPK relative to HIF-1α expression are switched off to enable leukocytes to fully commit to their inflammatory functions.
Leukocytes synthesize and secret a variety of inflammatory mediators, including pro- and anti-inflammatory cytokines, shortly after activation (59). Our data show that the proteolytic cleavage of AMPKα is associated with mTORC1 and S6K1 activation. These data suggest that AMPKα inactivation has two roles: 1) to enable HIF-1 activation; and 2) to enable a rapid increase in protein synthesis.
This study also investigated whether the TLR4 signaling phenotype described in this study is clinically relevant. Our data demonstrate that leukocytes from two disparate patient cohorts, patients with T2DM and patients after CPB, exhibit a signaling phenotype that is similar to that induced by LPS. In addition, plasma from these patients appears to contain a soluble and transferable ligand(s) that can trigger TLR4-like signaling in healthy donor leukocytes. Furthermore, pharmacologic response modifiers that inhibited LPS-induced responses in vitro also prevented patient plasma-induced responses in healthy donor’s leukocytes, suggesting involvement of common signaling mediators in both scenarios. Whether these or other pharmacological agents could reverse the signaling pathway in patient leukocytes once the pathway is already activated and in the presence of the activating ligand(s) is currently undetermined.
The data highlight the likely involvement of TLR4 in physiologic stress responses that are induced by endogenous agonists and/or gut-derived LPS. We were unable to detect presence of LPS in patient blood using commercially available Limulus amebocyte lysate–based assays (detection limit 0.1 EU/ml). However, whether LPS can be detected in human serum is currently controversial (60, 61). We reported that an LPS concentration as low as 0.1 ng/kg triggers onset of TLR4 signaling in healthy subject leukocytes in the absence of systemic responses (15). LPS used in those studies had an activity of 10 EU/ng. Estimated blood volume per kg body weight is 75 ml for men and 65 ml for women. Administration of 0.1 ng/kg would have equaled to 1 EU/75 ml for men and 1 EU/65 ml blood for women or 0.013–0.015 EU/ml. Furthermore, the effect of even lower LPS concentrations might be augmented by presence of additional TLR4 ligands, such as fibronectin fragments and/or saturated fatty acids (4–10) in patients’ blood.
In conclusion, this study describes a novel TLR4-signaling arm that is functional in leukocytes/neutrophils and required for regulating AMPK, mTORC1, S6K1, and HIF-1. The study also demonstrates that this signaling pathway is clinically relevant and significant because it is expressed in patient leukocytes. Analyses of leukocytes from larger and more diverse patient cohorts could provide the basis for better understanding of TLR4 signaling in chronic diseases and opportunities for appropriate therapeutic interventions.
Footnotes
The study was supported in part by the National Institutes of Health National Institute on Environmental Health Sciences–sponsored Rutgers Center for Environmental Exposures and Disease Grant NIEHS P30ES005022, and funds from the New Jersey Health Foundation (to B.H. and L.Y.L.).
Abbreviations used in this article:
- AMPA
p-aminophenylmercuric acetate
- CPB
cardiopulmonary bypass
- In. 1
MMP2/MMP9 inhibitor I
- In. 2
MMP2/MMP9 inhibitor IV
- In. 3
MMP9 inhibitor I
- mTORC1
mammalian target of rapamycin complex 1
- Raptor
regulatory-associated protein of mTOR
- RWJMS
Robert Wood Johnson Medical School
- S6K1
S6 kinase 1
- T2DM
type 2 diabetes mellitus.
References
Disclosures
The authors have no financial conflicts of interest.