PTEN-β-Catenin Signaling Modulates Regulatory T Cells and Inflammatory Responses in Mouse Liver Ischemia and Reperfusion Injury
Introduction
Liver inflammatory injury induced by ischemia and reperfusion injury (IRI) is the major pathogenesis to cause hepatic dysfunction and failure following liver transplantation (1). Macrophages (Kupffer cells) play a pivotal role in triggering innate immune response and developing liver inflammation during liver IRI (2). Activated macrophages release TNF-α and IL-1, which initiate a complex inflammatory cascade that leads to activate CD4 T cells (3, 4). Moreover, the RORγt-expressing (RORγt) T cells are the main source of Th17-producing effector cells during the early phase of liver IRI. Inhibition of RORγt activity resulted in reduced IR-induced liver damage (5).
Phosphatase and tensin homolog deleted on chromosome 10 (PTEN) has been known to play an important role during inflammatory response. PTEN positively regulates LPS-induced TLR4 signaling and proinflammatory cytokine secretion by antagonizing phosphoinositide3-kinase (PI3K) signaling (6). Increased PTEN activity promotes tissue inflammation, while deletion of PTEN activates PI3K/Akt signaling and reduces inflammatory response (7). Our previous studies demonstrated that PTEN/PI3K regulated innate TLR4-driven inflammatory response, which was mediated by activating β-catenin signaling in IR-triggered liver inflammation (8).
Macrophage polarization plays an important role in the regulation of inflammatory responses. The innate and adaptive immunity regulated by macrophage PTEN/PI3K signaling might be involved in the expression of Arginase I (Arg1) (9). Activation of Wnt/β-catenin signaling promoted tissue damage repair by a signal transducer and activator of transcription 6 (STAT6)-mediated M2 polarization (10). Moreover, disruption of PPARγ in myeloid cells impairs alternative macrophage activation (11), suggesting PPARγ is required for the acquisition and maintenance of the anti-inflammatory macrophage phenotype.
Regulatory T cells (Tregs) are essential for the maintenance of immune homeostasis in liver inflammatory injury (12). Tregs constitutively express the transcription factor Foxp3, which is a key for Treg development and function (13). Increasing PTEN activity inhibits the development of CD4CD25 Tregs via a distinct IL-2 receptor (IL-2R) signaling, which is associated with downstream mediators of PI3K (14). PTEN deficiency increases nuclear accumulation of β-catenin (15) and promotes PI3K, leading to activation of downstream Akt and induction of Tregs (16). Moreover, stabilization of β-catenin enhances Treg survival and controls inflammatory responses (17). Activation of Jagged-1/Notch signaling instructs Treg differentiation (18). Thus, PTEN-β-catenin signaling might play an important role in promoting Treg induction during inflammatory response. However, it is still unclear how PTEN-β-catenin signaling may regulate Treg induction during liver inflammatory injury. Here, we report a novel regulatory mechanism of PTEN-β-catenin signaling on inflammatory response in liver injury. We have demonstrated that myeloid PTEN deficiency ameliorates IR-induced liver injury through the activation of β-catenin, which in turn promotes PPARγ-mediated Jagged-1/Notch signaling to induce Foxp3 Tregs while inhibiting Th17 cells. Our data documents that PTEN-β-catenin signaling is crucial for the modulation of innate and adaptive immunity in the mechanism of IR-induced liver injury.
Materials and Methods
Animals
Floxed PTEN (PTEN) mice (The Jackson Laboratory) and the mice expressing the Cre recombinase under the control of the Lysozyme M (LysM) promoter (LysM-Cre; The Jackson Laboratory) were used to generate myeloid-specific PTEN knockout (PTEN) mice, as described (19). All animals were maintained under specific pathogen free condition. The study protocols were approved by the Institutional Animal Care and Use Committee (IACUC) of University of California at Los Angeles.
Mouse liver IRI model
We used an established mouse model of warm hepatic ischemia followed by reperfusion, as described (20). Some animals were injected via tail vein with Notch1 siRNA or nonspecific (NS) siRNAs (2mg/kg, Santa Cruz Biotechnology) at 4 h prior to ischemia (8), or with γ-secretase inhibitor DAPT (10 mg/kg, Sigma-Aldrich) and DMSO vehicle at 30 min prior to ischemia. See Supplementary Materials.
Hepatocellular function assay
Serum alanine aminotransferase (sALT) levels, an indicator of hepatocellular injury, were measured by IDEXX Laboratories (Westbrook, ME).
Histology, immunohistochemistry, and immunofluorescence staining
Liver sections were stained with hematoxylin and eosin (H&E). The IRI severity was graded using Suzuki’s criteria (21). Liver CD11b macrophages were detected by immunohistochemistry staining with a rat anti-mouse CD11b mAb (BD Biosciences). Immunofluorescence staining was used to identify CD68 macrophages with a goat anti-mouse CD68 mAb (Santa Cruz Biotechnology). See Supplementary Materials.
TUNEL assay
The Klenow-FragEL DNA Fragmentation Detection Kit (EMD Chemicals) was used to detect the DNA fragmentation characteristic of oncotic necrosis/apoptosis in formalin-fixed paraffin-embedded liver sections (8). Results were scored semi-quantitatively by averaging the number of apoptotic cells/microscopic field at 400× magnification. Ten fields were evaluated/sample.
Quantitative RT-PCR analysis
Quantitative real-time PCR was performed as previously described (22). The primer sequences used for the amplification are shown in Supplementary Table 1. See Supplementary Materials.
Western blot analysis
Protein was extracted from liver tissue or cell cultures, as described (22). Monoclonal rabbit anti-mouse β-catenin, phos-Akt, Akt, PPAR-γ, Jagged-1, cleaved Notch1, Hes1, and β-actin Abs (Cell Signaling Technology) were used. The relative quantities of proteins were determined by densitometer, and expressed in absorbance units (AU).
BMM isolation and in vitro transfection
Murine bone marrow-derived macrophages (BMMs) were generated as previously described (22). Cells (1×10/well) were cultured for 7 days and then transfected with 100nM of siRNA (β-catenin siRNA, PPAR-γ siRNA or Jagged-1 siRNA (Santa Cruz Biotechnology). See Supplementary Materials.
Spleen T cell isolation
The spleen T cells were purified using the EasySep mouse T cell isolation kit (STEMCELL Technologies) according to the manufacturer’s instructions. T cells were then stimulated with anti-CD3 (1 μg/ml) and anti-CD28 (2 μg/ml) (eBioscience).
Macrophage/T cell co-culture
The PTEN, PTEN, β-catenin siRNA, PPARγ siRNA or Jagged-1 siRNA-transfected macrophages (5×10 cells/ml) were cultured and stimulated with LPS (100 ng/ml) for 6 h. Splenic T cells were added at a ratio of 1:10 (macrophage:T cell). The co-cultured cells were incubated for 24 h.
Flow cytometry analysis
Spleen T cells from DMSO or DAPT-treated PTEN mice were stained with anti-mouse CD4-PE-Cyanine5, CD25-PE, RORγt-PE, and Foxp3-FITC mAbs (eBioscience) according to the manufacturer’s instructions. PE-labeled rat anti-mouse IgG2a isotypes were used as negative controls. Measurements were performed using a FACSCalibur flow cytometer (BD Biosciences). Data analysis was performed using CellQuest software.
Statistical analysis
Data are expressed as mean±SD and analyzed by Student’s t-tests. Per comparison two-sided p values less than 0.05 were considered statistically significant. Multiple group comparisons were performed using one-way ANOVA with a post-hoc test.
Animals
Floxed PTEN (PTEN) mice (The Jackson Laboratory) and the mice expressing the Cre recombinase under the control of the Lysozyme M (LysM) promoter (LysM-Cre; The Jackson Laboratory) were used to generate myeloid-specific PTEN knockout (PTEN) mice, as described (19). All animals were maintained under specific pathogen free condition. The study protocols were approved by the Institutional Animal Care and Use Committee (IACUC) of University of California at Los Angeles.
Mouse liver IRI model
We used an established mouse model of warm hepatic ischemia followed by reperfusion, as described (20). Some animals were injected via tail vein with Notch1 siRNA or nonspecific (NS) siRNAs (2mg/kg, Santa Cruz Biotechnology) at 4 h prior to ischemia (8), or with γ-secretase inhibitor DAPT (10 mg/kg, Sigma-Aldrich) and DMSO vehicle at 30 min prior to ischemia. See Supplementary Materials.
Hepatocellular function assay
Serum alanine aminotransferase (sALT) levels, an indicator of hepatocellular injury, were measured by IDEXX Laboratories (Westbrook, ME).
Histology, immunohistochemistry, and immunofluorescence staining
Liver sections were stained with hematoxylin and eosin (H&E). The IRI severity was graded using Suzuki’s criteria (21). Liver CD11b macrophages were detected by immunohistochemistry staining with a rat anti-mouse CD11b mAb (BD Biosciences). Immunofluorescence staining was used to identify CD68 macrophages with a goat anti-mouse CD68 mAb (Santa Cruz Biotechnology). See Supplementary Materials.
TUNEL assay
The Klenow-FragEL DNA Fragmentation Detection Kit (EMD Chemicals) was used to detect the DNA fragmentation characteristic of oncotic necrosis/apoptosis in formalin-fixed paraffin-embedded liver sections (8). Results were scored semi-quantitatively by averaging the number of apoptotic cells/microscopic field at 400× magnification. Ten fields were evaluated/sample.
Quantitative RT-PCR analysis
Quantitative real-time PCR was performed as previously described (22). The primer sequences used for the amplification are shown in Supplementary Table 1. See Supplementary Materials.
Western blot analysis
Protein was extracted from liver tissue or cell cultures, as described (22). Monoclonal rabbit anti-mouse β-catenin, phos-Akt, Akt, PPAR-γ, Jagged-1, cleaved Notch1, Hes1, and β-actin Abs (Cell Signaling Technology) were used. The relative quantities of proteins were determined by densitometer, and expressed in absorbance units (AU).
BMM isolation and in vitro transfection
Murine bone marrow-derived macrophages (BMMs) were generated as previously described (22). Cells (1×10/well) were cultured for 7 days and then transfected with 100nM of siRNA (β-catenin siRNA, PPAR-γ siRNA or Jagged-1 siRNA (Santa Cruz Biotechnology). See Supplementary Materials.
Spleen T cell isolation
The spleen T cells were purified using the EasySep mouse T cell isolation kit (STEMCELL Technologies) according to the manufacturer’s instructions. T cells were then stimulated with anti-CD3 (1 μg/ml) and anti-CD28 (2 μg/ml) (eBioscience).
Macrophage/T cell co-culture
The PTEN, PTEN, β-catenin siRNA, PPARγ siRNA or Jagged-1 siRNA-transfected macrophages (5×10 cells/ml) were cultured and stimulated with LPS (100 ng/ml) for 6 h. Splenic T cells were added at a ratio of 1:10 (macrophage:T cell). The co-cultured cells were incubated for 24 h.
Flow cytometry analysis
Spleen T cells from DMSO or DAPT-treated PTEN mice were stained with anti-mouse CD4-PE-Cyanine5, CD25-PE, RORγt-PE, and Foxp3-FITC mAbs (eBioscience) according to the manufacturer’s instructions. PE-labeled rat anti-mouse IgG2a isotypes were used as negative controls. Measurements were performed using a FACSCalibur flow cytometer (BD Biosciences). Data analysis was performed using CellQuest software.
Statistical analysis
Data are expressed as mean±SD and analyzed by Student’s t-tests. Per comparison two-sided p values less than 0.05 were considered statistically significant. Multiple group comparisons were performed using one-way ANOVA with a post-hoc test.
Results
Myeloid PTEN deficiency ameliorates hepatocellular damage and reduces macrophage trafficking in IR-induced liver injury
The hepatocellular damage was evaluated in mouse livers subjected to 90 min of warm ischemia followed by 6 h of reperfusion. Livers in PTEN mice showed severe edema, sinusoidal congestion, and necrosis (Figure 1A and 1B, score=3.4±0.5). However, livers in PTEN mice showed mild to moderate edema and sinusoidal congestion (Figure 1A and 1B, score=1.3±0.2, p<0.001). Consistent with the histopathological data, the serum ALT levels (IU/L) in PTEN mice were significantly lower than those in the PTEN controls (Figure 1C, 6905±1852 vs. 26265±2610, p<0.001). Moreover, PTEN reduced the frequency of TUNEL cells in ischemic livers compared to the PTEN controls (Figure 1D, 29.5±3.5 vs 67.9±5.6, p<0.001). PTEN deficiency in PTEN livers decreased CD68 macrophage infiltration (Figure 1E, 14.5±2.5) compared to the PTEN controls (26.5±4.5, p<0.001). To confirm inflammatory cell recruitment in ischemic livers, CD11b macrophages were detected by immunohistochemistry staining. Indeed, reduced CD11b macrophages were observed in PTEN but not PTEN mice (Supplementary figure 1, 12.3±2.2 vs. 31.5±4.6, p<0.001).
(A) Representative histological staining (H&E) of ischemic liver tissue (n=4–5/group, magnification x100). (B) Liver damage, evaluated by Suzuki’s score. ***p<0.001. (C) Hepatocellular function, assessed by serum ALT levels (IU/L). Results expressed as mean±SD (n=4–5/group), ***p<0.001. (D) Liver apoptosis by TUNEL staining. Results expressed as mean±SD (n=4–6/group, magnification x400), ***p<0.001. (E) Immunofluorescence staining of CD68 macrophages (short arrow) in ischemic liver lobes. Results expressed as mean±SD (n=4/group, magnification ×400), ***p<0.001.
Myeloid PTEN deficiency promotes β-catenin activation and Treg induction in IR-induced liver injury
We found that by 6 h of reperfusion after 90 min of ischemia, the protein expression of β-catenin (p=0.006) and phosphorylated Akt (p=0.026) was up-regulated in PTEN but not in PTEN livers (Figure 2A). The mRNA levels of proinflammatory genes coding for TNF-α (p=0.008), IL-1β (p=0.022), and IL-6 (p=0.034) were decreased, whereas TGF-β (p=0.027) expression was increased in PTEN livers compared to the PTEN controls (Figure 2B). Moreover, PTEN promoted M2 macrophage differentiation as evidenced by the increased arginase1 (Arg1) (p=0.026) and reduced M1 macrophage inducible nitric oxide synthase (iNOS) (p=0.006) expression in ischemic livers compared to the PTEN controls (Figure 2C). Interestingly, the expression of Foxp3 (p=0.021), a master regulator of Treg cells, was increased while the expression of RoRγt (p=0.006) and IL-17A (p=0.014) was decreased in PTEN but not in PTEN livers (Figure 2D). Consistent with this data, we found that PTEN activated Notch signaling by increased expression of Notch1 (p=0.026) and its downstream target gene, hairy and enhancer of split-1 (Hes1) (p=0.005), and transcription factor recombination signal sequence-binding protein jκ (RBP-J) (p=0.037) (Figure 2E), which was accompanied by augmented Foxp3 (p=0.013) and reduced RORγt (p=0.005) and IL-17A (p=0.023) in spleen T cells (Figure 2F).
(A) Western blot analysis and relative density ratio of β-catenin, p-Akt, and Akt in ischemic livers. *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (B) TNF-α, IL-1β, IL-6, TGF-β, (C) Arg1, iNOS, and (D) Foxp3, RORγt, IL-17A in ischemic livers or (E) Notch1, Hes1, RBP-J and (F) Foxp3, RORγt, IL-17A in spleen T cells from PTEN and PTEN mice. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01.
PTEN-β-catenin axis activates PPARγ and Jagged-1/Notch signaling pathway and induces Foxp3Tregs in vitro
To elucidate the putative mechanisms by which PTEN-mediated β-catenin regulates Notch signaling and adaptive Treg development, we disrupted β-catenin in BMMs from PTEN mice by using a small interfering RNA (siβ-cat). Indeed, PTEN increased the expression of β-catenin (p=0.004), PPARγ (p=0.036), and Jagged-1 (p=0.017) in LPS-stimulated BMMs after non-specific (NS) siRNA treatment. However, knockdown of β-catenin with siβ-cat pretreatment in PTEN-BMMs resulted in reduced PPARγ (p=0.034) and Jagged-1 (p=0.008) expression after LPS stimulation (Figure 3A). Moreover, siβ-cat treatment in PTEN-BMMs augmented the mRNA levels of TNF-α (p=0.021), IL-1β (p=0.017), and IL-6 (p=0.013) but reduced levels of TGF-β (p=0.037) in response to LPS stimulation compared to the NS siRNA-treated controls (Figure 3B). The decreased Arg1 (p=0.008) and increased iNOS (p=0.026) expression was observed in siβ-cat-treated PTEN-BMMs but not in NS siRNA-treated cells (Figure 3C). Furthermore, we used siβ-cat-pretreated BMMs from PTEN mice and then co-cultured them with spleen T cells after LPS stimulation. Indeed, knockdown of β-catenin in PTEN-BMMs decreased cleaved Notch1 (p=0.006) protein expression (Figure 3D) and mRNA levels coding for Notch1 (p=0.042), Hes1 (p=0.037), and RBP-J (p=0.024) (Figure 3E). This was accompanied by reduced Foxp3 (p=0.036) but augmented RORγt (p=0.041) and IL-17A (p=0.025) in spleen T cells (Figure 3F).
BMMs were transfected with β-catenin siRNA (siβ-cat), and then co-cultured with spleen T cells after LPS stimulation for 6 h. (A) Western blot analysis and relative density ratio of β-catenin, PPARγ and Jagged-1 in LPS-stimulated macrophages. *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (B) TNF-α, IL-1β, IL-6, TGF-β, (C) Arg1, iNOS in LPS-stimulated macrophages. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01. (D) Western blot analysis and relative density ratio of cleaved Notch1 in spleen T cells after co-culture *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (E) Notch1, Hes1, RBP-J and (F) Foxp3, RORγt, IL-17A in spleen T cells after co-culture. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01.
PPARγ mediates Jagged-1/Notch signaling pathway in vitro
Using macrophage (BMM)/spleen T cell co-culture system, we found that myeloid PTEN deficiency increased PPARγ (p=0.009) and Jagged-1 (p=0.016) expression in NS siRNA-treated macrophages after LPS stimulation (Figure 4A). However, knockdown of PPARγ with PPARγ siRNA (siPPARγ) treatment resulted in reduced Jagged-1 (p=0.007) expression in PTEN-deficient cells (Figure 4A), with increased expression of TNF-α (p=0.028), IL-1β (p=0.039), and IL-6 (p=0.016) and reduced levels of TGF-β (p=0.034) in response to LPS stimulation compared to the NS siRNA-treated controls (Figure 4B). siPPARγ treatment decreased Arg1 (p=0.005) but increased iNOS (p=0.014) expression in PTEN-deficient macrophages compared to the NS siRNA-treated cells (Figure 4C). Moreover, PPARγ knockdown in macrophages reduced cleaved Notch1 (p=0.026) protein expression (Figure 4D) and Notch1 (p=0.035) and RBP-J (p=0.042) mRNA levels (Figure 4E), with reduced Foxp3 (p=0.037) but increased RORγt (p=0.028) and IL-17A (p=0.008) (Figure 4F) in spleen T cells after co-culture.
BMMs were transfected with PPARγ siRNA (siPPARγ), and then co-cultured with spleen T cells after LPS stimulation for 6 h. (A) Western blot analysis and relative density ratio of PPARγ and Jagged-1 in LPS-stimulated macrophages. *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (B) TNF-α, IL-1β, IL-6, TGF-β, (C) Arg1, iNOS in LPS-stimulated macrophages. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01. (D) Western blot analysis and relative density ratio of cleaved Notch1 in spleen T cells after co-culture *p<0.05. Quantitative RT-PCR-assisted detection of (E) Notch1, RBP-J and (F) Foxp3, RORγt, IL-17A in spleen T cells after co-culture. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01.
Jagged-1/Notch signaling is essential for the Foxp3+Treg induction in the PTEN-β-catenin signaling-mediated immune regulation in vitro
We disrupted Jagged-1/Notch signaling in BMMs from PTEN mice with Jagged-1 siRNA (siJagged-1), and then co-cultured with spleen T cells. Pretreatment of LPS-stimulated BMMs with siJagged-1 diminished Jagged-1 expression compared to the NS siRNA-treated cells (Figure 5A, p=0.005). Moreover, unlike NS siRNA-treated controls, siJagged-1 pretreatment inhibited cleaved Notch1 (p=0.008) and Hes1 (p=0.006) protein expression in spleen T cells after co-culture (Figure 5B). The mRNA levels coding for Notch1 (p=0.032) and RBP-J (p=0.039) were reduced in siJagged-1 but not NS siRNA-treated cells (Figure 5C). Consistent with these data, knockdown of Jagged-1 resulted in reduced Foxp3 (p=0.032) while increasing RORγt (p=0.024) and IL-17A (p=0.042) expression in T cells (Figure 5D).
BMMs were transfected with Jagged-1 siRNA (siJagged-1), and then co-cultured with spleen T cells after LPS stimulation for 6 h. (A) Western blot analysis and relative density ratio of Jagged-1 in LPS-stimulated macrophages. *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (B) Western blot analysis and relative density ratio of cleaved Notch1 in spleen T cells after co-culture *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (C) Notch1, RBP-J and (D) Foxp3, RORγt, IL-17A in spleen T cells after co-culture. Each column represents the mean±SD (n=3–4/group). *p<0.05.
Blocking Jagged-1/Notch signaling pathway aggravates IR-induced liver damage and inhibits Foxp3Treg induction in vivo
We next investigated whether disruption of Notch signaling may affect local inflammatory responses in mouse liver IRI. DAPT, a γ-secretase inhibitor, can prevent the final cleavage step of the precursor form of Notch to the active Notch intracellular domain (NICD) (23). At 90 min of partial liver warm ischemia followed by 6 h of reperfusion, livers in PTEN mice treated with vehicle DMSO controls, showed mild to moderate edema without necrosis (Figure 6A and 6B, score=1.1±0.2). In contrast, livers in mice after receiving DAPT revealed significant edema, severe sinusoidal congestion/cytoplasmic vacuolization, and extensive (30–50%) necrosis (score=3.2±0.5, p<0.001). This data was consistent with hepatocellular function, which showed that DAPT treatment in PTEN mice increased sALT levels compared to the DMSO-treated controls (Figure 6C, 22274±3340 vs. 4914±1941, p<0.001). Liver cell apoptosis was analyzed by TUNEL staining. DAPT treatment in PTEN mice increased the frequency of apoptotic TUNEL cells in ischemic livers compared to the DMSO-treated controls (Figure 6D, 67.5±7.8 vs. 30.9±4.2, p<0.001). To further confirm the role of Notch1 signaling in liver IRI, we disrupted Notch signaling in PTEN mice using Notch1 siRNA or non-specific siRNA (NS siRNA). Livers in mice treated with NS siRNA showed mild to moderate edema without necrosis (Supplementary figure 2, score=1.1±0.2). In contrast, Notch1 siRNA-treated livers revealed significant edema, severe sinusoidal congestion/cytoplasmic vacuolization, and extensive (30–50%) necrosis (score=3.3±0.7, p<0.001). PTEN deficiency increased Hes1 expression in DMSO-treated PTEN mice (Figure 6E, p=0.006). However, blocking Notch signaling in PTEN mice resulted in reduced Hes1 expression (Figure 6E, p=0.026) after DAPT treatment. The expression of pro-inflammatory TNF-α (p=0.025), IL-1β (p=0.034), and IL-6 (p=0.008) was increased in DAPT but not in DMSO- treated PTEN mice (Figure 6F). Consistent with these data, the mRNA level of RBP-J was decreased (Figure 6G, p=0.007), leading to reduced Foxp3 (p=0.037) while augmented RORγt (p=0.024) and IL-17A (p=0.008) expression in DAPT-treated PTEN mice, as compared with that in DMSO-treated controls (Figure 6H). We then analyzed Foxp3 or RORγt expression in spleen T cells by flow cytometry analysis. As a result, we observed a significantly reduced percentage of CD4CD25Foxp3 Tregs in DAPT-treated PTEN mice but not in DMSO-treated controls (Figure 6I, 3.9±0.4 vs. 6.8±0.6, p<0.001). In contrast, DAPT treatment resulted in an increased percentage of RORγt T cells (IL-17A-producing T cells) compared to the control groups (Figure 6I, 3.6±0.5 vs. 1.5±0.4, p<0.001).
(A) Representative histological staining (H&E) of ischemic livers. (n=4–5/group, magnification x100). (B) The severity of liver IRI was evaluated by the Suzuki’s histological grading. ***p<0.001. (C) Hepatocellular function was evaluated by sALT levels (IU/L). Results expressed as mean±SD (n=4–5/group). ***p<0.001. (D) Liver apoptosis by TUNEL staining. Results expressed as mean±SD (n=4–6/group, magnification x400), ***p<0.001. (E) Western blot analysis and relative density ratio of Hes1 in ischemic livers. *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (F) TNF-α, IL-1β, IL-6, (G) RBP-J, and (H) Foxp3, RORγt, IL-17A in ischemic livers. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01. (I) Foxp3 and RORγt expression in spleen T cells were evaluated by flow cytometry. Results expressed as mean±SD (n=3/group, p<0.001).
Myeloid PTEN deficiency ameliorates hepatocellular damage and reduces macrophage trafficking in IR-induced liver injury
The hepatocellular damage was evaluated in mouse livers subjected to 90 min of warm ischemia followed by 6 h of reperfusion. Livers in PTEN mice showed severe edema, sinusoidal congestion, and necrosis (Figure 1A and 1B, score=3.4±0.5). However, livers in PTEN mice showed mild to moderate edema and sinusoidal congestion (Figure 1A and 1B, score=1.3±0.2, p<0.001). Consistent with the histopathological data, the serum ALT levels (IU/L) in PTEN mice were significantly lower than those in the PTEN controls (Figure 1C, 6905±1852 vs. 26265±2610, p<0.001). Moreover, PTEN reduced the frequency of TUNEL cells in ischemic livers compared to the PTEN controls (Figure 1D, 29.5±3.5 vs 67.9±5.6, p<0.001). PTEN deficiency in PTEN livers decreased CD68 macrophage infiltration (Figure 1E, 14.5±2.5) compared to the PTEN controls (26.5±4.5, p<0.001). To confirm inflammatory cell recruitment in ischemic livers, CD11b macrophages were detected by immunohistochemistry staining. Indeed, reduced CD11b macrophages were observed in PTEN but not PTEN mice (Supplementary figure 1, 12.3±2.2 vs. 31.5±4.6, p<0.001).
(A) Representative histological staining (H&E) of ischemic liver tissue (n=4–5/group, magnification x100). (B) Liver damage, evaluated by Suzuki’s score. ***p<0.001. (C) Hepatocellular function, assessed by serum ALT levels (IU/L). Results expressed as mean±SD (n=4–5/group), ***p<0.001. (D) Liver apoptosis by TUNEL staining. Results expressed as mean±SD (n=4–6/group, magnification x400), ***p<0.001. (E) Immunofluorescence staining of CD68 macrophages (short arrow) in ischemic liver lobes. Results expressed as mean±SD (n=4/group, magnification ×400), ***p<0.001.
Myeloid PTEN deficiency promotes β-catenin activation and Treg induction in IR-induced liver injury
We found that by 6 h of reperfusion after 90 min of ischemia, the protein expression of β-catenin (p=0.006) and phosphorylated Akt (p=0.026) was up-regulated in PTEN but not in PTEN livers (Figure 2A). The mRNA levels of proinflammatory genes coding for TNF-α (p=0.008), IL-1β (p=0.022), and IL-6 (p=0.034) were decreased, whereas TGF-β (p=0.027) expression was increased in PTEN livers compared to the PTEN controls (Figure 2B). Moreover, PTEN promoted M2 macrophage differentiation as evidenced by the increased arginase1 (Arg1) (p=0.026) and reduced M1 macrophage inducible nitric oxide synthase (iNOS) (p=0.006) expression in ischemic livers compared to the PTEN controls (Figure 2C). Interestingly, the expression of Foxp3 (p=0.021), a master regulator of Treg cells, was increased while the expression of RoRγt (p=0.006) and IL-17A (p=0.014) was decreased in PTEN but not in PTEN livers (Figure 2D). Consistent with this data, we found that PTEN activated Notch signaling by increased expression of Notch1 (p=0.026) and its downstream target gene, hairy and enhancer of split-1 (Hes1) (p=0.005), and transcription factor recombination signal sequence-binding protein jκ (RBP-J) (p=0.037) (Figure 2E), which was accompanied by augmented Foxp3 (p=0.013) and reduced RORγt (p=0.005) and IL-17A (p=0.023) in spleen T cells (Figure 2F).
(A) Western blot analysis and relative density ratio of β-catenin, p-Akt, and Akt in ischemic livers. *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (B) TNF-α, IL-1β, IL-6, TGF-β, (C) Arg1, iNOS, and (D) Foxp3, RORγt, IL-17A in ischemic livers or (E) Notch1, Hes1, RBP-J and (F) Foxp3, RORγt, IL-17A in spleen T cells from PTEN and PTEN mice. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01.
PTEN-β-catenin axis activates PPARγ and Jagged-1/Notch signaling pathway and induces Foxp3Tregs in vitro
To elucidate the putative mechanisms by which PTEN-mediated β-catenin regulates Notch signaling and adaptive Treg development, we disrupted β-catenin in BMMs from PTEN mice by using a small interfering RNA (siβ-cat). Indeed, PTEN increased the expression of β-catenin (p=0.004), PPARγ (p=0.036), and Jagged-1 (p=0.017) in LPS-stimulated BMMs after non-specific (NS) siRNA treatment. However, knockdown of β-catenin with siβ-cat pretreatment in PTEN-BMMs resulted in reduced PPARγ (p=0.034) and Jagged-1 (p=0.008) expression after LPS stimulation (Figure 3A). Moreover, siβ-cat treatment in PTEN-BMMs augmented the mRNA levels of TNF-α (p=0.021), IL-1β (p=0.017), and IL-6 (p=0.013) but reduced levels of TGF-β (p=0.037) in response to LPS stimulation compared to the NS siRNA-treated controls (Figure 3B). The decreased Arg1 (p=0.008) and increased iNOS (p=0.026) expression was observed in siβ-cat-treated PTEN-BMMs but not in NS siRNA-treated cells (Figure 3C). Furthermore, we used siβ-cat-pretreated BMMs from PTEN mice and then co-cultured them with spleen T cells after LPS stimulation. Indeed, knockdown of β-catenin in PTEN-BMMs decreased cleaved Notch1 (p=0.006) protein expression (Figure 3D) and mRNA levels coding for Notch1 (p=0.042), Hes1 (p=0.037), and RBP-J (p=0.024) (Figure 3E). This was accompanied by reduced Foxp3 (p=0.036) but augmented RORγt (p=0.041) and IL-17A (p=0.025) in spleen T cells (Figure 3F).
BMMs were transfected with β-catenin siRNA (siβ-cat), and then co-cultured with spleen T cells after LPS stimulation for 6 h. (A) Western blot analysis and relative density ratio of β-catenin, PPARγ and Jagged-1 in LPS-stimulated macrophages. *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (B) TNF-α, IL-1β, IL-6, TGF-β, (C) Arg1, iNOS in LPS-stimulated macrophages. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01. (D) Western blot analysis and relative density ratio of cleaved Notch1 in spleen T cells after co-culture *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (E) Notch1, Hes1, RBP-J and (F) Foxp3, RORγt, IL-17A in spleen T cells after co-culture. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01.
PPARγ mediates Jagged-1/Notch signaling pathway in vitro
Using macrophage (BMM)/spleen T cell co-culture system, we found that myeloid PTEN deficiency increased PPARγ (p=0.009) and Jagged-1 (p=0.016) expression in NS siRNA-treated macrophages after LPS stimulation (Figure 4A). However, knockdown of PPARγ with PPARγ siRNA (siPPARγ) treatment resulted in reduced Jagged-1 (p=0.007) expression in PTEN-deficient cells (Figure 4A), with increased expression of TNF-α (p=0.028), IL-1β (p=0.039), and IL-6 (p=0.016) and reduced levels of TGF-β (p=0.034) in response to LPS stimulation compared to the NS siRNA-treated controls (Figure 4B). siPPARγ treatment decreased Arg1 (p=0.005) but increased iNOS (p=0.014) expression in PTEN-deficient macrophages compared to the NS siRNA-treated cells (Figure 4C). Moreover, PPARγ knockdown in macrophages reduced cleaved Notch1 (p=0.026) protein expression (Figure 4D) and Notch1 (p=0.035) and RBP-J (p=0.042) mRNA levels (Figure 4E), with reduced Foxp3 (p=0.037) but increased RORγt (p=0.028) and IL-17A (p=0.008) (Figure 4F) in spleen T cells after co-culture.
BMMs were transfected with PPARγ siRNA (siPPARγ), and then co-cultured with spleen T cells after LPS stimulation for 6 h. (A) Western blot analysis and relative density ratio of PPARγ and Jagged-1 in LPS-stimulated macrophages. *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (B) TNF-α, IL-1β, IL-6, TGF-β, (C) Arg1, iNOS in LPS-stimulated macrophages. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01. (D) Western blot analysis and relative density ratio of cleaved Notch1 in spleen T cells after co-culture *p<0.05. Quantitative RT-PCR-assisted detection of (E) Notch1, RBP-J and (F) Foxp3, RORγt, IL-17A in spleen T cells after co-culture. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01.
Jagged-1/Notch signaling is essential for the Foxp3+Treg induction in the PTEN-β-catenin signaling-mediated immune regulation in vitro
We disrupted Jagged-1/Notch signaling in BMMs from PTEN mice with Jagged-1 siRNA (siJagged-1), and then co-cultured with spleen T cells. Pretreatment of LPS-stimulated BMMs with siJagged-1 diminished Jagged-1 expression compared to the NS siRNA-treated cells (Figure 5A, p=0.005). Moreover, unlike NS siRNA-treated controls, siJagged-1 pretreatment inhibited cleaved Notch1 (p=0.008) and Hes1 (p=0.006) protein expression in spleen T cells after co-culture (Figure 5B). The mRNA levels coding for Notch1 (p=0.032) and RBP-J (p=0.039) were reduced in siJagged-1 but not NS siRNA-treated cells (Figure 5C). Consistent with these data, knockdown of Jagged-1 resulted in reduced Foxp3 (p=0.032) while increasing RORγt (p=0.024) and IL-17A (p=0.042) expression in T cells (Figure 5D).
BMMs were transfected with Jagged-1 siRNA (siJagged-1), and then co-cultured with spleen T cells after LPS stimulation for 6 h. (A) Western blot analysis and relative density ratio of Jagged-1 in LPS-stimulated macrophages. *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (B) Western blot analysis and relative density ratio of cleaved Notch1 in spleen T cells after co-culture *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (C) Notch1, RBP-J and (D) Foxp3, RORγt, IL-17A in spleen T cells after co-culture. Each column represents the mean±SD (n=3–4/group). *p<0.05.
Blocking Jagged-1/Notch signaling pathway aggravates IR-induced liver damage and inhibits Foxp3Treg induction in vivo
We next investigated whether disruption of Notch signaling may affect local inflammatory responses in mouse liver IRI. DAPT, a γ-secretase inhibitor, can prevent the final cleavage step of the precursor form of Notch to the active Notch intracellular domain (NICD) (23). At 90 min of partial liver warm ischemia followed by 6 h of reperfusion, livers in PTEN mice treated with vehicle DMSO controls, showed mild to moderate edema without necrosis (Figure 6A and 6B, score=1.1±0.2). In contrast, livers in mice after receiving DAPT revealed significant edema, severe sinusoidal congestion/cytoplasmic vacuolization, and extensive (30–50%) necrosis (score=3.2±0.5, p<0.001). This data was consistent with hepatocellular function, which showed that DAPT treatment in PTEN mice increased sALT levels compared to the DMSO-treated controls (Figure 6C, 22274±3340 vs. 4914±1941, p<0.001). Liver cell apoptosis was analyzed by TUNEL staining. DAPT treatment in PTEN mice increased the frequency of apoptotic TUNEL cells in ischemic livers compared to the DMSO-treated controls (Figure 6D, 67.5±7.8 vs. 30.9±4.2, p<0.001). To further confirm the role of Notch1 signaling in liver IRI, we disrupted Notch signaling in PTEN mice using Notch1 siRNA or non-specific siRNA (NS siRNA). Livers in mice treated with NS siRNA showed mild to moderate edema without necrosis (Supplementary figure 2, score=1.1±0.2). In contrast, Notch1 siRNA-treated livers revealed significant edema, severe sinusoidal congestion/cytoplasmic vacuolization, and extensive (30–50%) necrosis (score=3.3±0.7, p<0.001). PTEN deficiency increased Hes1 expression in DMSO-treated PTEN mice (Figure 6E, p=0.006). However, blocking Notch signaling in PTEN mice resulted in reduced Hes1 expression (Figure 6E, p=0.026) after DAPT treatment. The expression of pro-inflammatory TNF-α (p=0.025), IL-1β (p=0.034), and IL-6 (p=0.008) was increased in DAPT but not in DMSO- treated PTEN mice (Figure 6F). Consistent with these data, the mRNA level of RBP-J was decreased (Figure 6G, p=0.007), leading to reduced Foxp3 (p=0.037) while augmented RORγt (p=0.024) and IL-17A (p=0.008) expression in DAPT-treated PTEN mice, as compared with that in DMSO-treated controls (Figure 6H). We then analyzed Foxp3 or RORγt expression in spleen T cells by flow cytometry analysis. As a result, we observed a significantly reduced percentage of CD4CD25Foxp3 Tregs in DAPT-treated PTEN mice but not in DMSO-treated controls (Figure 6I, 3.9±0.4 vs. 6.8±0.6, p<0.001). In contrast, DAPT treatment resulted in an increased percentage of RORγt T cells (IL-17A-producing T cells) compared to the control groups (Figure 6I, 3.6±0.5 vs. 1.5±0.4, p<0.001).
(A) Representative histological staining (H&E) of ischemic livers. (n=4–5/group, magnification x100). (B) The severity of liver IRI was evaluated by the Suzuki’s histological grading. ***p<0.001. (C) Hepatocellular function was evaluated by sALT levels (IU/L). Results expressed as mean±SD (n=4–5/group). ***p<0.001. (D) Liver apoptosis by TUNEL staining. Results expressed as mean±SD (n=4–6/group, magnification x400), ***p<0.001. (E) Western blot analysis and relative density ratio of Hes1 in ischemic livers. *p<0.05, **p<0.01. Quantitative RT-PCR-assisted detection of (F) TNF-α, IL-1β, IL-6, (G) RBP-J, and (H) Foxp3, RORγt, IL-17A in ischemic livers. Each column represents the mean±SD (n=3–4/group). *p<0.05, **p<0.01. (I) Foxp3 and RORγt expression in spleen T cells were evaluated by flow cytometry. Results expressed as mean±SD (n=3/group, p<0.001).
Discussion
This study documents for the first time that PTEN-β-catenin signaling is crucial for orchestrating inflammatory responses, PPARγ activation, Jagged-1/Notch signaling, and Treg induction in liver inflammatory injury due to IR. First, PTEN, a key negative regulator of the PI3K/Akt signaling pathway, is essential for IR-induced liver injury. Myeloid-specific PTEN knockout reduced liver damage and macrophage trafficking in ischemic livers. Second, myeloid-specific PTEN knockout promoted β-catenin activation, which in turn induced PPARγ activation, increased anti-inflammatory M2 macrophage differentiation and decreased pro-inflammatory cytokines. Third, PTEN-β-catenin axis activated PPARγ-mediated Jagged-1/Notch signaling, leading to increased Foxp3 Tregs. Fourth, disruption of Jagged-1/Notch signaling pathway resulted in aggravated IR-induced liver damage and reduced Foxp3 Treg induction. Our results highlight the role of PTEN-β-catenin signaling in regulating innate and adaptive immune responses during IR-triggered liver inflammation.
The molecular mechanisms of IR-induced liver damage involved in the activation of innate and adaptive immunity may be through multiple cellular and molecular signaling pathways. PTEN, a multifunctional phosphatase, is shown to be essential for controlling innate immunity in liver injury through regulation of its downstream PI3K/Akt signaling (24, 25). Activation of Akt increases β-catenin activity and inhibits TLR4 and NLRP3-mediated innate immune response during liver IRI (8, 22). Consistent with the role of Akt/β-catenin signaling cascade in the inflammatory response, our current in vivo study has shown that myeloid-specific PTEN deficiency promoted β-catenin activation and diminished inflammatory injury, as evidenced by ameliorated IR-induced liver damage, reduced macrophage activation, and pro-inflammatory cytokines while increasing anti-inflammatory M2 macrophage differentiation. Interestingly, myeloid-specific PTEN deficiency augmented liver Foxp3 and reduced RORγt/IL-17A expression, implying the important role of PTEN-β-catenin signaling in the regulation of Foxp3 Treg induction during liver IRI.
The transcription factor Foxp3 is crucial for the ability of Treg cells to inhibit inflammatory response (26). Using the macrophage and T cell co-culture system, we found that myeloid-specific PTEN deficiency activated β-catenin, PPARγ, Jagged-1/Notch signaling, and increased Foxp3 Treg induction. However, knockdown of β-catenin in PTEN-deficient macrophages inhibited PPARγ and Jagged-1/Notch, which led to reduced Foxp3 Tregs. Indeed, β-catenin signaling was required for the control of innate and adaptive immunity during inflammatory response (8). Activation of β-catenin increased anti-inflammatory mediators and Treg induction while inhibiting inflammatory effect T cells (27). PPARγ was shown to transcriptionally regulate macrophage activation and function (28). Ligand of PPARγ inhibited T cell and pro-inflammatory cytokine activation during the regulation of inflammatory activities (29). In agreement with these findings, we found that PPARγ knockdown inhibited M2 macrophage differentiation as evidenced by reduced Arg1 and augmented iNOS expression. Moreover, PPARγ knockdown in PTEN-deficient macrophages resulted in reduced Jagged-1/Notch1 activation. This may imply the mechanistic links between PTEN-β-catenin axis and the PPARγ-mediated Jagged-1/Notch signaling pathway in the regulation of Foxp3 Treg induction in liver inflammation.
To further elucidate the regulatory network by which PTEN-β-catenin signaling may regulate Foxp3 Treg induction through a Jagged-1/Notch pathway-dependent manner in live inflammation, we knocked down Jagged-1 in PTEN-deficient macrophages and then co-cultured with T cells. We found that knockdown of Jagged-1 inhibited Notch1 activation, which resulted in reducing its target gene Hes1 and transcription factor RBP-J expression in T cells. Hes1 is a basic helix-loop-helix–type transcriptional repressor and negatively regulates gene transcription. It is known for cell proliferation and differentiation (30). Hes1 may regulate innate TLR4 activation by targeting RBP-J via a feedback inhibitory loop (31). Moreover, Hes1 inhibits inflammatory response by modulating transcription elongation (32). Loss of Hes1 gene results in impairing T cell development in the selective expansion of early T cell precursors (33). Notch signaling might directly bind to Foxp3 promoter in Hes1- and RBPJ-dependent mechanisms (34). Activation of Notch-Hes1 may mediate Treg immunosuppressive functions via a TGF-β signaling (35), which is known to be crucial for Treg development (36). Consistent with these results, we found that myeloid PTEN knockout increased Notch1 and Hes1 expression in T cells. Blocking Jagged-1/Notch signaling suppressed Hes1, leading to reduced Foxp3 Tregs while increasing RORγt/IL-17A expression. This indicates that Hes1 is a key mediator for the induction of Foxp3 Tregs during liver IRI.
Further evidence of Jagged-1/Notch signaling-mediated modulation of Tregs in liver IRI was obtained from myeloid-specific PTEN knockout (PTEN) mice. We found that, PTEN mice treated with DAPT or Notch1 siRNA (Supplementary figure 2) exacerbated liver damage. Indeed, DAPT is a γ-secretase inhibitor. γ-secretase mediates cleavage of the Notch receptor and releases NICD, which translocates into the nucleus and directly participates in a transcriptional complex with the DNA-binding proteins to activate transcription of target genes (23). DAPT has been shown to inhibit Notch signaling in studies of autoimmune and lymphoproliferative diseases (37). Inhibition of γ-secretase activity by DAPT treatment increased LPS-induced inflammation by activating NF-κB signaling (38). In agreement with these findings, we found that blocking Notch signaling by DAPT treatment suppressed Hes1 expression and increased pro-inflammatory mediators. Notably, DAPT treatment resulted in reduced CD4CD25Foxp3 Tregs and increased RORγt-mediated IL-17A-producing T cells in PTEN livers, which suggests that Jagged-1/Notch signaling most likely contributed to the CD4CD25Foxp3 Treg induction during liver IRI.
Figure 7 depicts putative molecular mechanisms by which PTEN-β-catenin signaling may regulate Treg development in liver IRI. Myeloid-specific PTEN deficiency promotes β-catenin activation via Akt phosphorylation. After translocating to the nucleus, β-catenin activates PPARγ-mediated Jagged-1/Notch signaling pathway by γ-secretase cleavage of Notch to NICD, which interacts with Hes1 and RBP-J, leading to promote Foxp3 Treg induction and inhibit inflammatory response in the liver.
Schematic illustration of molecular mechanisms of PTEN-β-catenin axis in the regulation of regulatory T cells and inflammatory responses in liver IRI.
In conclusion, we demonstrate that PTEN-β-catenin signaling regulates Foxp3 Treg induction via activating PPARγ-mediated Jagged-1/Notch signaling pathway in liver IRI. Myeloid-specific PTEN ablation promotes β-catenin, which in turn activates PPARγ and Jagged-1/Notch signaling, leading to augmented Foxp3 Treg induction while inhibited RORγt/IL-17A in IR-triggered liver inflammation. By identifying molecular mechanisms of PTEN-β-catenin signaling in the modulation of innate immune response and adaptive Treg development, our study provides potential therapeutic targets in liver IRI followed by liver transplantation.
Supplementary Material
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Acknowledgments
Funding sources: NIH Grant R21AI112722 (B.Ke), National Natural Science Foundation of China 81100270, 1310108001, 81210108017, National Science Foundation of Jiangsu Province BK20131024, BE2016766, 863 Young Scientists Special Fund grant SS2015AA0209322 and the Foundation of Jiangsu Collaborative Innovation Center of Biomedical Functional Materials (L. Lu), 81273262 (F. Zhang), and The Dumont Research Foundation.
Abstract
The phosphatase and tensin homolog deleted on chromosome 10 (PTEN) plays an important role in regulating T cell activation during inflammatory response. Activation of β-catenin is crucial for maintaining immune homeostasis. This study investigates the functional roles and molecular mechanisms by which PTEN-β-catenin signaling promotes regulatory T cell (Treg) induction in a mouse model of liver ischemia and reperfusion injury (IRI). We found that mice with myeloid specific PTEN knockout (PTEN) exhibited reduced liver damage as evidenced by decreased levels of serum ALT, intrahepatic macrophage trafficking, and pro-inflammatory mediators compared to the PTEN-proficient (PTEN) controls. Disruption of myeloid PTEN activated β-catenin, which in turn promoted PPARγ-mediated Jagged-1/Notch signaling and induced Foxp3 Tregs while inhibiting Th17 cells. However, blocking of Notch signaling by inhibiting γ-secretase reversed myeloid PTEN deficiency-mediated protection in IR-triggered liver inflammation with reduced Foxp3 and increased RORγt-mediated IL-17A expression in ischemic livers. Moreover, knockdown of β-catenin or PPARγ in PTEN-deficient macrophages inhibited Jagged-1/Notch activation and reduced Foxp3 Treg induction, leading to increased proinflammatory mediators in macrophage/T cell co-cultures. In conclusion, our findings demonstrate that PTEN-β-catenin signaling is a novel regulator involved in modulating Treg development and provides a potential therapeutic target in liver IRI.
Footnotes
The authors declare no conflicts of interest.
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