Thioredoxin-Interacting Protein: Comparison
Please note this is a comparison between Version 2 by Peter Tang and Version 1 by Douglas Gordon Walker.

The development of new therapeutic approaches to diseases relies on the identification of key molecular targets involved in amplifying disease processes. One such molecule is thioredoxin-interacting protein (TXNIP), also designated thioredoxin-binding protein-2 (TBP-2), a member of the α-arrestin family of proteins and a central regulator of glucose and lipid metabolism, involved in diabetes-associated vascular endothelial dysfunction and inflammation. TXNIP sequesters reduced thioredoxin (TRX), inhibiting its function, resulting in increased oxidative stress. Many different cellular stress factors regulate TXNIP expression, including high glucose, endoplasmic reticulum stress, free radicals, hypoxia, nitric oxide, insulin, and adenosine-containing molecules. TXNIP is also directly involved in inflammatory activation through its interaction with the nucleotide-binding domain, leucine-rich-containing family, and pyrin domain-containing-3 (NLRP3) inflammasome complex. Neurodegenerative diseases such as Alzheimer’s disease have significant pathologies associated with increased oxidative stress, inflammation, and vascular dysfunctions.

  • oxidative stress
  • inflammation
  • Alzheimer’s disease
  • glucose metabolism
  • neuropathology

1. Introduction

Oxidative stress resulting from an imbalance in cellular redox can occur due to the production of excessive levels of reactive oxygen species (ROS) and/or a deficit of cellular antioxidant systems. Supplementation with various classes of antioxidants has been widely investigated for neurodegenerative diseases (for recent reviews, see [1,2][1][2]). Excess levels of ROS can occur from inflammatory responses by innate immune cells (e.g., neutrophils, monocytes/macrophages/microglia) due to infections or chronic autoimmune responses [2], deficiencies in mitochondrial oxidative phosphorylation pathways [3[3][4],4], and environmental factors such as UV light, toxic chemicals, or heavy metals. Oxidative stress is linked to many different chronic human diseases, particularly those with inflammatory and/or metabolic components (e.g., diabetes and diabetes-related retinopathy [5], stroke [6], cancers [7], vascular diseases, and neurodegenerative diseases [8,9][8][9]). Developing effective therapies for these diseases requires the identification of key pathological targets responsible for amplifying disease processes.

Thioredoxin-interacting protein (TXNIP) has emerged as a key pathological regulator of diseases, particularly those associated with glucose and lipid abnormalities and inflammation. TXNIP has been identified in disease mechanisms involved in various cancers [10[10][11][12],11,12], diabetes mellitus [13], cardiovascular disease [14], renal disease [15], and retinal disease [16], among others. Many of these diseases associated with TXNIP can arise from vascular complications of diabetes [17]. The involvement of TXNIP in diabetes, pancreatic beta-cell death, and glucose metabolism has been the focus of several recent review articles [14,17,18][14][17][18]. TXNIP was identified (and named) due to its interaction and inhibition of the key antioxidant proteins thioredoxin-1 (TRX1) and thioredoxin-2 (TRX2) [19,20,21][19][20][21], but more recent findings identified properties involved in cellular metabolism and transcription regulation [22], apoptosis and cell death [13], inflammation [23], and tumor suppression [11][24] [11,24] that might not be directly related to the modulation of TRX and oxidative stress.

2. Biochemistry of TXNIP: Overview

The gene for TXNIP is located on the human chromosome 1q21.1 and is transcribed to four messenger RNA (mRNA) splice variants [28][25]. TXNIP is a member of the α-arrestin protein family and has two arrestin-like domains: one is a PxxP sequence and the other is a PPxY sequence. One mRNA for TXNIP codes for 336 amino acids for a polypeptide of approximately 37.4 kDa, whereas another codes for 391 amino acids for a polypeptide of approximately 43.7 kDa. The two other splice variants have not been associated with proteins. Reported molecular weights of TXNIP vary depending on the expressing cell type, but the major protein band(s) detected by SDS-gel electrophoresis is/are 50–55 kDa. This would suggest a certain amount of post-translational modification occurring. Protein bands of approximately 37 kDa have been observed on immunoblots with certain antibodies and cell types, but 50–55 kDa polypeptides appear to be the major form. TXNIP was identified initially by several investigators as vitamin D3-upregulated protein-1 (VDUP1) in different cell types [19,20,21][19][20][21]. Interestingly, other studies have not confirmed TXNIP induction by vitamin D3 in additional cell types, and noticeably, there is a lack of a vitamin D3-responsive element in the TXNIP gene promoter [29,30][26][27]. It has been suggested that vitamin D3 stabilizes cellular TXNIP protein rather than inducing expression [29][26].

2.1. TXNIP and Thioredoxin System

Earlier studies provided the first link between VDUP1/TXNIP and oxidative stress by demonstrating binding to reduced, but not oxidized, thioredoxin (TRX) [19,20,21][19][20][21]. TXNIP can bind two cysteine residues (Cys32 and Cys35) present in the active catalytic site of TRX. The binding of oxidized TXNIP to reduced TRX involves the formation of disulfide bonds between TXNIP-Cys247 and TRX-Cys32 [31][28]. A mutation in TXNIP-Cys247 is sufficient to remove its ability to sequester TRX activity [31][28]. Although classified as belonging to the α-arrestin family, these cysteine residues are unique to TXNIP. TXNIP can bind and inactivate both TRX1 and TRX2 [19,32][19][29]. The TRX cellular disulfide oxidoreductase system is a highly conserved system found from prokaryotes to plants to mammals [33,34,35][30][31][32]. TRX plays a central role in protecting cells from oxidative stress. Oxidized TRX is reduced by NADPH through a reaction catalyzed by thioredoxin reductase complex (TRX-R). Reduced TRX can then directly reduce disulfides in target proteins [33] [30]. The main form of TRX is TRX1, which is located primarily in the cytosol but can be translocated to the plasma membrane and/or nucleus, particularly under inflammatory conditions [36,37][33][34]. Closely related in structure, TRX2 is specific to mitochondria [38,39][35][36]. Immunohistochemistry and in situ hybridization mRNA studies have shown that TRX was highly expressed in mammalian brains, particularly in subsets of neurons in areas of high metabolic activity and oxidative burden [40,41,42][37][38][39]. TRX mRNA expression was highest in the piliform cortex, dentate gyrus, CA3/CA4 region of the hippocampus, locus coeruleus, and nucleus of the hypothalamus and solitary tract [41][38]. Trx2 mRNA and protein were highly expressed in rat brains in neurons in the olfactory bulb, frontal cortex, hippocampus, some hypothalamic and thalamic nuclei, cerebellum, and numerous brainstem nuclei [42][39].

The significance of the TRX system for maintaining cellular health was demonstrated in transgenic mice overexpressing forms of TRX. Increased expression of human TRX in mice resulted in resistance to oxidative stress and increased lifespan [43][40]. Other studies have shown that TRX overexpression in transgenic mice was protective of transient ischemic brain damage [44][41]. This effect was also observed by intravenous administration of TRX protein [45][42]. These examples of the beneficial outcomes of increased TRX are used to introduce the widely appreciated pathological consequences of reduced TRX activity. The first study linking TRX to AD showed a significant reduction in TRX protein levels in most brain regions in AD cases compared to nondemented controls with accompanying increased TRX-R activity. This study also demonstrated the protective effect of TRX when added to cultures of neurons treated with toxic doses of amyloid beta (Aβ) peptide [46][43]. A further study showed that TRX1 levels were increased significantly in the cerebrospinal fluid (CSF) and plasma of AD cases compared to mild cognitive impairment (MCI) cases [47][44]. This suggested that TRX1 was being secreted from damaged neurons in AD brains. A noticeable alteration in the patterns of TRX1 and TRX2 cellular immunoreactivity in hippocampal neurons of AD cases was also observed.

2.2. TXNIP Interacting Proteins

TXNIP has been shown to interact with other proteins besides TRX, including importin-α1 [48][45], human ecdysoneless (hEcd) [49][46], and NOD-, LRR-, and pyrin domain-containing protein-3 (NLRP3), a component of the inflammasome complex [23,50][23][47]. These interactions relate to additional identified properties of TXNIP. The significance of the interaction of TXNIP with inflammasome components and enhancement of inflammation will be considered.

3. Regulation of TXNIP Expression

Many different cellular stress factors positively or negatively regulate TXNIP expression. These include UV light, heat shock [51][48], hypoxia, ROS, nitric oxide [52][49], nicotinamide adenine dinucleotide [53][50], ATP, glutamine, nicotine, vascular endothelial growth factor, basic fibroblast growth factor [54][51], transforming growth factor β [55][52], estradiol [56][53], calcium channel blockers [57][54], activators of advanced glycation endoproduct receptor (RAGE) [58][55], insulin, and glucose [59][56]. Activation of the TXNIP promoter, which contains a carbohydrate response element (ChoRE), is regulated by transcription factors MondoA:Max-like protein X (MLx), nuclear factor Y (NF-Y), and the carbohydrate response element-binding protein (ChREBP) [45][42]. Transcription factors forkhead box O1 (FOXO1) and FOXO3a can bind to the TXNIP promoter and by competing with ChREBP can downregulate TXNIP transcription [46,47][43][44]. Activation of AMP protein kinase (AMPK) can also lead to inhibition of TXNIP mRNA transcription [48][45].

The mechanism whereby TXNIP expression is induced through the activation of the receptor for advanced glycation endproducts (RAGE) by the RAGE ligand S100b is of relevance for th consideration of AD as RAGE is a receptor for Aβ and is involved in Aβ toxicity and AD pathogenesis [58][55]. It was demonstrated in Schwann cells in vitro and injured sciatic nerve in vivo that binding of S100b to RAGE induced TXNIP with TXNIP being involved in the downstream activation of p38 mitogen-activated protein kinase (MAPK), cAMP response element-binding protein (CREB), and nuclear factor κB (NFκB). RAGE silencing blocked the induction of TXNIP, whereas the silencing of TXNIP inhibited the activation of these signaling pathways, preventing RAGE-induced fibronectin and IL1β synthesis and Schwann cell migration [58][55]. Induction of TXNIP by RAGE ligands in retinal endothelial cells through the activation of p38 MAPK and NFκB also led to enhanced expression of inflammation-associated genes, including cyclooxygenase 2 (Cox2), vascular endothelial growth factor (VEGFA), and intercellular adhesion molecule-1 (ICAM1). Expression of these genes was reduced when TXNIP expression was inhibited, and enhanced with TXNIP overexpression [60][57].

4. TXNIP, Hyperglycemia, and Oxidative Stress—NLRP3 Inflammasome Complex

Presentations of research data on TXNIP in relation to disease have frequently considered TXNIP/TRX interactions and the resulting oxidative stress as a separate mechanism from its role in the NLRP3 inflammasome activation, but these two features are interrelated. Although associated with enhanced inflammation, NLRP3 inflammasome activation occurs in all brain cell types, including neurons, astrocytes, and endothelial cells, not just microglia [81,102][58][59]. If one considers the events occurring under hyperglycemic conditions, excess glucose leads to increased levels of ROS by inducing the overproduction of NADH and increased mitochondrial-derived ROS that inhibits GAPDH, the enzyme that removes excess cellular glucose [103][60]. However, this inhibition that activates alternative glucose metabolic pathways leads to further ROS production. Excess ROS directly induces the expression of TXNIP and the inflammation-associated transcription factor NFκB. Excess TXNIP will further exacerbate oxidative stress by binding TRX and, in cooperation with NFκB, bind and activate the inflammasome complex. The inflammasome complex consists of an association of NLRP3, apoptosis-associated speck-like protein containing a caspase-binding domain (ASC) and pro-caspase-1 [104,105][61][62]. In the presence of excess ROS, TXNIP interaction with TRX can be reversed, which then allows released TXNIP to bind and activate NLRP3 inflammasome, resulting in enhanced inflammation. Activation of NLRP3-inflammasome promotes the formation of activated caspase-1, which processes interleukin (IL)-1β, IL-18, and IL-33 into bioactive forms [106][63], and also induces cell death through pyroptosis [107][64]. An earlier study also demonstrated that inflammasome activators such as uric acid crystals induced the dissociation of TXNIP from TRX in the presence of ROS, allowing it to bind and activate the NLRP3 complex and enhance caspase activation [50][47].

5. TXNIP Expression in the Brain

Studies focusing on the cellular distribution of TXNIP in brain cells are limited. Detailed immunochemical characterization of TXNIP/VDUP1 by comparing its distribution in Drosophila and rat brains showed relatively conserved patterns of expression. The antibodies employed in this study identified this protein in both of these species. Constitutive expression in subsets of neurons and astrocytes were identified using double immunohistochemistry staining with appropriate markers. This study did not report the constitutive microglial expression of TXNIP but observed nuclear immunoreactivity under hyperglycemic conditions [108][65]. A further study employed immunohistochemistry to compare the neuroanatomical distribution of different TRXs, TRRs, glutathione/glutaredoxin, peroxiredoxins, and TXNIP in rat brains. Expression of these antioxidants and related molecules was highest in brain regions susceptible to damage in conditions of hypoxia/ischemia including the cerebellum, cortex, hippocampus, substantia nigra, striatum, and spinal cord. Weak expression of TXNIP was detected in subsets of neurons, not glia, in these brain regions, except the spinal cord, and with the highest expression in the retina. The pattern of expression of TRX1 and TXNIP showed extensive overlap [109][66]. A direct link between metabolic activity and TXNIP was demonstrated in mice with the expression of TXNIP in medial hypothalamic neurons, which increased under conditions of nutrient excess and obesity [110][67]. A further study using directed TXNIP gene deletion and overexpression in Agrp hypothalamic neurons showed that the overexpression of TXNIP led to reduced energy expenditure and activity, resulting in obesity and fat accumulations, whereas the deletion of TXNIP in Agrp neurons had a reverse effect, with increased energy metabolism and activity [111][68]. Another study showed strong induction of TXNIP in the hypothalamus of mice in a state of lowered energy metabolism, though increased TXNIP expression was detected in ependymal-lining cells, not neurons, in the hypothalamus. Increased TXNIP expression was also detected in the liver and adipose tissues of these animals [112][69]. These studies highlighted physiological functions for TXNIP in regulating energy metabolism.

6. Physiological Consequences of Loss of TXNIP

6.1. Experimental Animals

A number of studies have reported the phenotypes of mice constructed to have total TXNIP gene deletion. The consequences of this have been both beneficial and detrimental depending on the disease model. In an initial report, the cause of hyperlipidemia in a mutant mouse strain (Hcb-19) was a mutation in the Txnip gene resulting in reduced expression. These mutant mice had hypertriglyceridemia, hypercholesterolemia, elevated plasma apolipoprotein B, and increased secretion of triglyceride-rich lipoproteins with increased ketone body synthesis [144][70]. In a TXNIP gene-deleted mouse model, they developed significantly increased fatty liver, with high levels of triacylglycerol, cholesterol ester, and total cholesterol, with higher-serum non-esterified fatty acids. Elevated fatty acid synthesis was the primary cause of liver lipogenesis in TXNIP knockout mice [145][71]. Gene deletion of TXNIP also resulted in impaired immunity with reduced numbers of natural killer cells and the impaired dendritic cell maturation of T cells [146,147,148][72][73][74]. TXNIP gene deletion can lead to lethality under fasting conditions due to a switch in metabolism to hyperlipidemia and hypoglycemia. Under these conditions, mice experienced enhanced glucose-induced insulin sensitivity and secretion and an increased expression of PPAR-target genes [149][75]. TXNIP gene-deletion mice fed a high-fat diet developed significantly greater fat deposits and body mass but with increased insulin sensitivity. This study correlated the loss of TXNIP with the augmented activation of PPAR-γ [150][76]. These effects on enhanced hyperlipidemia due to TXNIP gene deletion were detrimental to the affected organisms, but beneficial outcomes upon reversing the consequences of diabetes/hyperglycemia were also observed. Knockdown of TXNIP in vitro protected mesangial cells grown under high-glucose conditions from apoptosis and ASK1 and p38 MAP kinase signaling [151,152][77][78]. In wild-type and TXNIP knockout mice rendered diabetic with streptozotocin, the TXNIP knockout mice showed significant protection from diabetic nephropathy. The knockout diabetic mice did not show increased thickening of the basal membrane or increased glomerular TGF-β1, collagen IV, and fibrosis was seen in the wild-type mice [153][79]. Loss of TXNIP provided significant protection in mice fed a HFD, showing significant protection from retinal degeneration and microvascular dysfunction due to reduced activation by the retinal endothelial cells of TXNIP/NLRP3 inflammasome [94][80]. TXNIP gene-deletion mice have been shown in general to be protected from the vascular consequences of high-fat/obesity or high-sugar diets compared to wild-type mice. In a model that studied the consequence of HFD and TXNIP gene deletion, it was shown that wild-type mice had significantly impaired blood flow and vascular density due to impaired angiogenesis compared to the TXNIP knockout mice. In this model, there was significantly higher IL-1β, increased numbers of infiltrating macrophages, and increased vascular endothelial growth factor (VEGF) expression and VEGF receptor activation in wild-type mice only. The enhanced vascular inflammation was due to the high-fat diet activating the TXNIP/NLRP3 inflammasome, which did not occur in TXNIP knockout mice [154][81].

6.2. Human Subjects

From the above-described experimental studies, the loss of TXNIP in mice has been beneficial in reversing the consequences of hyperglycemia but with a detrimental effect on liver fatty acid metabolism. These animals were constructed with Txnip gene deletions, but the loss of TXNIP has been identified in rare genetic cases in human subjects. The published case report covers children of 4, 9, and 13 years of age [155][82]. This showed that the loss of TXNIP is nonlethal, but the affected subjects show lactic acidosis, low-serum methionine, and impaired oxidative phosphorylation in response to glucose and pyruvate [155][82]. There are no reports as yet if such individuals are resistant to diabetes or neurodegenerative diseases. These studies demonstrate that the pharmacological lowering of TXNIP levels is unlikely to be associated with significant side effects.

Table 1 lists the most significant studies related to the effects of TXNIP expression in the brain or brain-derived cells. This is a subjective list based on the theme of this article and is proposed to illustrate the different features of TXNIP in interactions with the nervous system.

Table 1. Summary of major studies of TXNIP in the brain or brain-derived cells.

7. Modulating TXNIP Expression

A number of different classes of agents can modulate TXNIP expression, including PPAR-γ agonists, which include classes of drugs with insulin-sensitizing properties used for treating diabetes [157][93]. There have been conflicting results on the interactions of PPAR-γ and TXNIP. One study using human kidney proximal tubule cells showed PPAR-γ agonists attenuated high-glucose-mediated TXNIP expression [158][94], whereas another using human macrophages showed that the PPAR-γ agonist GW929 enhanced TXNIP expression [71][95]. Using a rat insulinoma cell line INS-1E to represent pancreatic beta cells, it was shown that hyperglycemia activated TXNIP expression and inhibited the activation of AMPK. Activation of AMPK with metformin or aminoimidazole-carboxamide ribonucleotide (AICAR) reduced TXNIP expression, an effect also observed when cells were treated with the lipid palmitate [142][96]. AMPK activation inhibited glucose-stimulated ChREBP nuclear entry and binding to the TXNIP promoter, thereby inhibiting TXNIP mRNA expression. These investigators also demonstrated that the addition of insulin to high-glucose-treated INS-1E cells reduced TXNIP expression and prevented high-glucose-mediated apoptosis. Treatment of cells with nitric oxide (NO) stimulated insulin secretion and reduced the expression of TXNIP; this effect was reversed when cells were treated with a nitric oxide synthase inhibitor [159][97]. A recent study showed that insulin-like growth factor-1 (IGF-1) negatively regulated TXNIP expression in vitro and in vivo. Furthermore, this study demonstrated that oxidative stress and glucose-induced TXNIP expression could be reversed by the administration of IGF-1 [160][98]. The calcium channel blocker verapamil, which is in widespread clinical use for hypertension, has been demonstrated to significantly inhibit TXNIP expression with therapeutic benefits. In an animal model of diabetic cardiomyopathy, three weeks of treatment with verapamil had a significant therapeutic benefit, which correlated with the reduced expression of TXNIP in cardiomyocytes [161][99]. The mechanism of action of verapamil appeared to be mediated by the enhanced activation of the transcription repressor nuclear factor Y (NPY) [162][100]. As mentioned above, verapamil was also effective in inhibiting tau phosphorylation in vivo in 5xFAD AD model mice, inhibiting Aβ-induced tau phosphorylation in vitro, and inhibiting the expression of TXNIP and the activation of p38 MAPK [123][101].

In a recent study, we identified a novel fluorinated derivative of curcumin that was highly effective in reducing stress-induced TXNIP expression. Cellular models examined used the human retinal pigment epithelial cell line ARPE-19, which have high constitutive levels of TXNIP expression, and macrophages derived from the human THP-1 monocyte cell line, which have lower constitutive expression. In this study, the effectiveness of fluorinated curcumin derivative Shiga Y6 compared to its non-fluorinated derivative Shiga Y5 in reducing TXNIP protein and mRNA expression under constitutive, high-glucose, endoplasmic reticulum stress, and inflammatory activation was demonstrated [163][102]. This compound was also effective in inducing TRX protein and mRNA levels in these cellular models. These studies demonstrate that derivatized curcumin molecules were more effective than curcumin alone in lowering TXNIP expression. Other studies have shown the effectiveness of curcumin in vitro and in vivo in inhibiting TXNIP expression but at higher doses than we used for testing the fluorinated curcumin compound [163,164][102][103]. The effect of curcumin on TXNIP expression appears to be through the activation of AMPK [81,165][58][104].

References

  1. Ramli, N.Z.; Yahaya, M.F.; Tooyama, I.; Damanhuri, H.A. A Mechanistic Evaluation of Antioxidant Nutraceuticals on Their Potential against Age-Associated Neurodegenerative Diseases. Antioxidants 2020, 9, 1019, doi:10.3390/antiox9101019.
  2. Simpson, D.S.A.; Oliver, P.L. ROS Generation in Microglia: Understanding Oxidative Stress and Inflammation in Neurodegenerative Disease. Antioxidants 2020, 9, 743, doi:10.3390/antiox9080743.
  3. Pradeepkiran, J.A.; Reddy, P.H. Defective mitophagy in Alzheimer’s disease. Ageing Res. Rev. 2020, 64, 101191.
  4. Lee, D.; Jo, M.G.; Kim, S.Y.; Chung, C.G.; Lee, S.B. Dietary Antioxidants and the Mitochondrial Quality Control: Their Potential Roles in Parkinson’s Disease Treatment. Antioxidants 2020, 9, 1056, doi:10.3390/antiox9111056.
  5. López-Contreras, A.K.; Martínez-Ruiz, M.G.; Olvera-Montaño, C.; Robles-Rivera, R.R.; Arévalo-Simental, D.E.; Castellanos-González, J.A.; Hernández-Chávez, A.; Huerta-Olvera, S.G.; Cardona-Muñoz, E.G.; Rodríguez-Carrizalez, A.D. Importance of the Use of Oxidative Stress Biomarkers and Inflammatory Profile in Aqueous and Vitreous Humor in Diabetic Retinopathy. Antioxidants 2020, 9, 891, doi:10.3390/antiox9090891.
  6. Tian, Y.; Su, Y.; Ye, Q.; Chen, L.; Yuan, F.; Wang, Z. Silencing of TXNIP Alleviated Oxidative Stress Injury by Regulating MAPK-Nrf2 Axis in Ischemic Stroke. Neurochem. Res. 2020, 45, 428–436, doi:10.1007/s11064-019-02933-y.
  7. Su, C.; Shi, A.; Cao, G.; Tao, T.; Chen, R.; Hu, Z.; Shen, Z.; Tao, H.; Cao, B.; Hu, D.; et al. Fenofibrate suppressed proliferation and migration of human neuroblastoma cells via oxidative stress dependent of TXNIP upregulation. Biochem. Biophys. Res. Commun. 2015, 460, 983–988, doi:10.1016/j.bbrc.2015.03.138.
  8. Pereira, J.D.; Fraga, V.G.; Santos, A.L.M.; Carvalho, M.D.G.; Caramelli, P.; Gomes, K.B. Alzheimer’s disease and type 2 diabetes mellitus: A systematic review of proteomic studies. J. Neurochem. 2020, 1–24, doi:10.1111/jnc.15166.
  9. Saleem, U.; Sabir, S.; Niazi, S.G.; Nadeem, M.; Ahmad, B. Role of Oxidative Stress and Antioxidant Defense Biomarkers in Neurodegenerative Diseases. Crit. Rev. Eukaryot. Gene Expr. 2020, 30, 311–322, doi:10.1615/critreveukaryotgeneexpr.2020029202.
  10. Nishizawa, K.; Nishiyama, H.; Matsui, Y.; Kobayashi, T.; Saito, R.; Kotani, H.; Masutani, H.; Oishi, S.; Toda, Y.; Fujii, N.; et al. Thioredoxin-interacting protein suppresses bladder carcinogenesis. Carcinogenesis 2011, 32, 1459–1466, doi:10.1093/carcin/bgr137.
  11. Xie, M.; Xie, R.; Xie, S.; Wu, Y.; Wang, W.; Li, X.; Xu, Y.; Liu, B.; Zhou, Y.; Wang, T.; et al. Thioredoxin interacting protein (TXNIP) acts as a tumor suppressor in human prostate cancer. Cell Biol. Int. 2020, 44, 2094–2106, doi:10.1002/cbin.11418.
  12. Goldberg, S.F.; Miele, M.E.; Hatta, N.; Takata, M.; Paquette-Straub, C.; Freedman, L.P.; Welch, D.R. Melanoma metastasis suppression by chromosome 6: Evidence for a pathway regulated by CRSP3 and TXNIP. Cancer Res. 2003, 63, 432–440.
  13. Chen, J.; Saxena, G.; Mungrue, I.N.; Lusis, A.J.; Shalev, A. Thioredoxin-interacting protein: A critical link between glucose toxicity and beta-cell apoptosis. Diabetes 2008, 57, 938–944.
  14. Chong, C.-R.; Chan, W.P.A.; Nguyen, T.H.; Liu, S.; Procter, N.E.K.; Ngo, D.T.; Sverdlov, A.L.; Chirkov, Y.Y.; Horowitz, J.D. Thioredoxin-Interacting Protein: Pathophysiology and Emerging Pharmacotherapeutics in Cardiovascular Disease and Diabetes. Cardiovasc. Drugs Ther. 2014, 28, 347–360, doi:10.1007/s10557-014-6538-5.
  15. Du, C.; Wu, M.; Liu, H.; Ren, Y.; Du, Y.; Wu, H.; Wei, J.; Liu, C.; Yao, F.; Wang, H.; et al. Thioredoxin-interacting protein regulates lipid metabolism via Akt/mTOR pathway in diabetic kidney disease. Int. J. Biochem. Cell Biol. 2016, 79, 1–13, doi:10.1016/j.biocel.2016.08.006.
  16. Devi, T.S.; Hosoya, K.-I.; Terasaki, T.; Singh, L.P. Critical role of TXNIP in oxidative stress, DNA damage and retinal pericyte apoptosis under high glucose: Implications for diabetic retinopathy. Exp. Cell Res. 2013, 319, 1001–1012, doi:10.1016/j.yexcr.2013.01.012.
  17. Hu, J.; Yu, Y. The Function of Thioredoxin-Binding Protein-2 (TBP-2) in Different Diseases. Oxidative Med. Cell. Longev. 2018, 2018, 1–10, doi:10.1155/2018/4582130.
  18. Alhawiti, N.M.; Al Mahri, S.; Aziz, M.A.; Malik, S.S.; Mohammad, S. TXNIP in Metabolic Regulation: Physiological Role and Therapeutic Outlook. Curr. Drug Targets 2017, 18, 1095–1103, doi:10.2174/1389450118666170130145514.
  19. Nishiyama, A.; Matsui, M.; Iwata, S.; Hirota, K.; Masutani, H.; Nakamura, H.; Takagi, Y.; Sono, H.; Gon, Y.; Yodoi, J. Identification of Thioredoxin-binding Protein-2/Vitamin D(3)Up-regulated Protein 1 as a Negative Regulator of Thioredoxin Function and Expression. J. Biol. Chem. 1999, 274, 21645–21650, doi:10.1074/jbc.274.31.21645.
  20. Junn, E.; Han, S.H.; Im, J.Y.; Yang, Y.; Cho, E.W.; Um, H.D.; Kim, D.K.; Lee, K.W.; Han, P.L.; Rhee, S.G.; et al. Vitamin D3 Up-Regulated Protein 1 Mediates Oxidative Stress Via Suppressing the Thioredoxin Function. J. Immunol. 2000, 164, 6287–6295, doi:10.4049/jimmunol.164.12.6287.
  21. Yamanaka, H.; Maehira, F.; Oshiro, M.; Asato, T.; Yanagawa, Y.; Takei, H.; Nakashima, Y. A Possible Interaction of Thioredoxin with VDUP1 in HeLa Cells Detected in a Yeast Two-Hybrid System. Biochem. Biophys. Res. Commun. 2000, 271, 796–800, doi:10.1006/bbrc.2000.2699.
  22. Farrell, M.R.; Rogers, L.K.; Liu, Y.; Welty, S.E.; Tipple, T.E. Thioredoxin-interacting protein inhibits hypoxia-inducible factor transcriptional activity. Free Radic. Biol. Med. 2010, 49, 1361–1367, doi:10.1016/j.freeradbiomed.2010.07.016.
  23. Schroder, K.; Zhou, R.; Tschopp, J. The NLRP3 Inflammasome: A Sensor for Metabolic Danger? Science 2010, 327, 296–300, doi:10.1126/science.1184003.
  24. Zhang, P.; Gao, J.; Wang, X.; Wen, W.; Yang, H.; Tian, Y.; Liu, N.; Wang, Z.; Liu, H.; Zhang, Y.; et al. A novel indication of thioredoxin-interacting protein as a tumor suppressor gene in malignant glioma. Oncol. Lett. 2017, 14, 2053–2058, doi:10.3892/ol.2017.6397.
  25. Vionnet, N.; Hani, E.H.; Dupont, S.; Gallina, S.; Francke, S.; Dotte, S.; De Matos, F.; Durand, E.; Leprêtre, F.; Lecoeur, C.; et al. Genomewide Search for Type 2 Diabetes–Susceptibility Genes in French Whites: Evidence for a Novel Susceptibility Locus for Early-Onset Diabetes on Chromosome 3q27-qter and Independent Replication of a Type 2–Diabetes Locus on Chromosome 1q21–q24. Am. J. Hum. Genet. 2000, 67, 1470–1480, doi:10.1086/316887.
  26. Abu El Maaty, M.A.; Almouhanna, F.; Wölfl, S. Expression of TXNIP in Cancer Cells and Regulation by 1,25(OH)₂D₃: Is It Really the Vitamin D₃ Upregulated Protein? Int. J. Mol. Sci. 2018, 19, 796, doi:10.3390/ijms19030796.
  27. Ludwig, D.L.; Kotanides, H.; Le, T.; Chavkin, D.; Bohlen, P.; Witte, L. Cloning, genetic characterization, and chromosomal mapping of the mouse VDUP1 gene. Gene 2001, 269, 103–112, doi:10.1016/s0378-1119(01)00455-3.
  28. Patwari, P.; Higgins, L.J.; Chutkow, W.A.; Yoshioka, J.; Lee, R.T. The interaction of thioredoxin with Txnip. Evidence for formation of a mixed disulfide by disulfide exchange. J. Biol. Chem. 2006, 281, 21884–21891.
  29. Saxena, G.; Chen, J.; Shalev, A. Intracellular Shuttling and Mitochondrial Function of Thioredoxin-interacting Protein. J. Biol. Chem. 2010, 285, 3997–4005, doi:10.1074/jbc.m109.034421.
  30. Luthman, M.; Holmgren, A. Rat liver thioredoxin and thioredoxin reductase: Purification and characterization. Biochemistry 1982, 21, 6628–6633, doi:10.1021/bi00269a003.
  31. Holmgren, A.; Fagerstedt, M. The in vivo distribution of oxidized and reduced thioredoxin in Escherichia coli. J. Biol. Chem. 1982, 257, 6926–6930.
  32. Berstermann, A.; Vogt, K.; Follmann, H. Plant Seeds Contain Several Thioredoxins of Regular Size. Eur. J. Biochem. 1983, 131, 339–344, doi:10.1111/j.1432-1033.1983.tb07267.x.
  33. Zhu, Z.; Chen, X.; Sun, J.; Li, Q.; Lian, X.; Li, S.; Wang, Y.; Tian, L. Inhibition of nuclear thioredoxin aggregation attenuates PM(2.5)-induced NF-κB activation and pro-inflammatory responses. Free Radic. Biol. Med. 2019, 130, 206–214.
  34. Obikane, H.; Abiko, Y.; Ueno, H.; Kusumi, Y.; Esumi, M.; Mitsumata, M. Effect of endothelial cell proliferation on atherogenesis: A role of p21(Sdi/Cip/Waf1) in monocyte adhesion to endothelial cells. Atherosclerosis 2010, 212, 116–122, doi:10.1016/j.atherosclerosis.2010.05.029.
  35. Bodenstein-Lang, J.; Buch, A.; Follmann, H. Animal and plant mitochondria contain specific thioredoxins. FEBS Lett. 1989, 258, 22–26, doi:10.1016/0014-5793(89)81606-0.
  36. Bodenstein, J.; Follmann, H. Characterization of Two Thioredoxins in Pig Heart Including a New Mitochondrial Protein. Zeitschrift für Naturforschung C 1991, 46, 270–279, doi:10.1515/znc-1991-3-418.
  37. Rozell, B.; Hansson, H.A.; Luthman, M.; Holmgren, A. Immunohistochemical localization of thioredoxin and thioredoxin reductase in adult rats. Eur. J. Cell Biol. 1985, 38, 79–86.
  38. Lippoldt, A.; Padilla, C.A.; Gerst, H.; Andbjer, B.; Richter, E.; Holmgren, A.; Fuxe, K. Localization of thioredoxin in the rat brain and functional implications. J. Neurosci. 1995, 15, 6747–6756, doi:10.1523/JNEUROSCI.15-10-06747.1995.
  39. Rybnikova, E.; Damdimopoulos, A.E.; Gustafsson, J.Å.; Spyrou, G.; Pelto-Huikko, M. Expression of novel antioxidant thioredoxin-2 in the rat brain. Eur. J. Neurosci. 2000, 12, 1669–1678, doi:10.1046/j.1460-9568.2000.00059.x.
  40. Mitsui, A.; Hamuro, J.; Nakamura, H.; Kondo, N.; Hirabayashi, Y.; Ishizaki-Koizumi, S.; Hirakawa, T.; Inoue, T.; Yodoi, J. Overexpression of Human Thioredoxin in Transgenic Mice Controls Oxidative Stress and Life Span. Antioxid. Redox Signal. 2002, 4, 693–696, doi:10.1089/15230860260220201.
  41. Takagi, Y.; Mitsui, A.; Nishiyama, A.; Nozaki, K.; Sono, H.; Gon, Y.; Hashimoto, N.; Yodoi, J. Overexpression of thioredoxin in transgenic mice attenuates focal ischemic brain damage. Proc. Natl. Acad. Sci. USA 1999, 96, 4131–4136, doi:10.1073/pnas.96.7.4131.
  42. Hattori, I.; Takagi, Y.; Nakamura, H.; Nozaki, K.; Bai, J.; Kondo, N.; Sugino, T.; Nishimura, M.; Hashimoto, N.; Yodoi, J. Intravenous Administration of Thioredoxin Decreases Brain Damage Following Transient Focal Cerebral Ischemia in Mice. Antioxid. Redox Signal. 2004, 6, 81–87, doi:10.1089/152308604771978372.
  43. Lovell, M.A.; Xie, C.; Gabbita, S.P.; Markesbery, W.R. Decreased thioredoxin and increased thioredoxin reductase levels in alzheimer’s disease brain. Free Radic. Biol. Med. 2000, 28, 418–427, doi:10.1016/s0891-5849(99)00258-0.
  44. Arodin, L.; Lamparter, H.; Karlsson, H.; Nennesmo, I.; Björnstedt, M.; Schröder, J.; Fernandes, A.P. Alteration of Thioredoxin and Glutaredoxin in the Progression of Alzheimer’s Disease. J. Alzheimers Dis. 2014, 39, 787–797, doi:10.3233/jad-131814.
  45. Nishinaka, Y.; Masutani, H.; Oka, S.-I.; Matsuo, Y.; Yamaguchi, Y.; Nishio, K.; Ishii, Y.; Yodoi, J. Importin α1 (Rch1) Mediates Nuclear Translocation of Thioredoxin-binding Protein-2/Vitamin D(3)-up-regulated Protein 1. J. Biol. Chem. 2004, 279, 37559–37565, doi:10.1074/jbc.m405473200.
  46. Suh, H.-W.; Yun, S.; Song, H.; Jung, H.; Park, Y.-J.; Kim, T.-D.; Yoon, S.R.; Choi, I. TXNIP interacts with hEcd to increase p53 stability and activity. Biochem. Biophys. Res. Commun. 2013, 438, 264–269, doi:10.1016/j.bbrc.2013.07.036.
  47. Zhou, R.; Tardivel, A.; Thorens, B.; Choi, I.; Tschopp, J. Thioredoxin-interacting protein links oxidative stress to inflammasome activation. Nat. Immunol. 2010, 11, 136–140, doi:10.1038/ni.1831.
  48. Kim, K.-Y.; Shin, S.M.; Kim, J.K.; Paik, S.G.; Yang, Y.; Choi, I. Heat shock factor regulates VDUP1 gene expression. Biochem. Biophys. Res. Commun. 2004, 315, 369–375, doi:10.1016/j.bbrc.2004.01.047.
  49. Schulze, P.C.; Liu, H.; Choe, E.; Yoshioka, J.; Shalev, A.; Bloch, K.D.; Lee, R.T. Nitric Oxide–Dependent Suppression of Thioredoxin-Interacting Protein Expression Enhances Thioredoxin Activity. Arter. Thromb. Vasc. Biol. 2006, 26, 2666–2672, doi:10.1161/01.atv.0000248914.21018.f1.
  50. Yu, F.-X.; Goh, S.-R.; Dai, R.-P.; Luo, Y. Adenosine-Containing Molecules Amplify Glucose Signaling and Enhance Txnip Expression. Mol. Endocrinol. 2009, 23, 932–942, doi:10.1210/me.2008-0383.
  51. Ng, M.K.C.; Wu, J.; Chang, E.; Wang, B.; Katzenberg-Clark, R.; Ishii-Watabe, A.; Cooke, J.P. A Central Role for Nicotinic Cholinergic Regulation of Growth Factor–Induced Endothelial Cell Migration. Arter. Thromb. Vasc. Biol. 2007, 27, 106–112, doi:10.1161/01.atv.0000251517.98396.4a.
  52. Masaki, S.; Masutani, H.; Yoshihara, E.; Yodoi, J. Deficiency of Thioredoxin Binding Protein-2 (TBP-2) Enhances TGF-β Signaling and Promotes Epithelial to Mesenchymal Transition. PLoS ONE 2012, 7, e39900, doi:10.1371/journal.pone.0039900.
  53. DeRoo, B.J.; Hewitt, S.C.; Peddada, S.D.; Korach, K.S. Estradiol Regulates the Thioredoxin Antioxidant System in the Mouse Uterus. Endocrinology 2004, 145, 5485–5492, doi:10.1210/en.2004-0471.
  54. Saitoh, T.; Tanaka, S.; Koike, T. Rapid induction and Ca(2+) influx-mediated suppression of vitamin D3 up-regulated protein 1 (VDUP1) mRNA in cerebellar granule neurons undergoing apoptosis. J. Neurochem. 2001, 78, 1267–1276, doi:10.1046/j.1471-4159.2001.00505.x.
  55. Sbai, O.; Devi, T.S.; Melone, M.A.B.; Féron, F.; Khrestchatisky, M.; Singh, L.P.; Perrone, L. RAGE-TXNIP axis is required for S100B-promoted Schwann cell migration, fibronectin expression and cytokine secretion. J. Cell Sci. 2010, 123, 4332–4339, doi:10.1242/jcs.074674.
  56. Parikh, H.; Carlsson, E.; Chutkow, W.A.; Johansson, L.E.; Storgaard, H.; Poulsen, P.; Saxena, R.; Ladd, C.; Schulze, P.C.; Mazzini, M.J.; et al. TXNIP Regulates Peripheral Glucose Metabolism in Humans. PLoS Med. 2007, 4, e158, doi:10.1371/journal.pmed.0040158.
  57. Perrone, L.; Devi, T.S.; Hosoya, K.-I.; Terasaki, T.; Singh, L.P. Thioredoxin interacting protein (TXNIP) induces inflammation through chromatin modification in retinal capillary endothelial cells under diabetic conditions. J. Cell. Physiol. 2009, 221, 262–272, doi:10.1002/jcp.21852.
  58. Li, Y.; Li, J.; Li, S.; Li, Y.; Wang, X.; Liu, B.; Fu, Q.; Ma, S. Curcumin attenuates glutamate neurotoxicity in the hippocampus by suppression of ER stress-associated TXNIP/NLRP3 inflammasome activation in a manner dependent on AMPK. Toxicol. Appl. Pharmacol. 2015, 286, 53–63, doi:10.1016/j.taap.2015.03.010.
  59. Koka, S.; Xia, M.; Chen, Y.; Bhat, O.M.; Yuan, X.; Boini, K.M.; Li, P.-L. Endothelial NLRP3 inflammasome activation and arterial neointima formation associated with acid sphingomyelinase during hypercholesterolemia. Redox Biol. 2017, 13, 336–344, doi:10.1016/j.redox.2017.06.004.
  60. Yan, L.-J. Pathogenesis of Chronic Hyperglycemia: From Reductive Stress to Oxidative Stress. J. Diabetes Res. 2014, 2014, 1–11, doi:10.1155/2014/137919.
  61. Yu, J.-W.; Wu, J.; Zhang, Z.; Datta, P.K.; Ibrahimi, I.M.; Taniguchi, S.; Sagara, J.; Fernandes-Alnemri, T.; Alnemri, E.S. Cryopyrin and pyrin activate caspase-1, but not NF-κB, via ASC oligomerization. Cell Death Differ. 2006, 13, 236–249, doi:10.1038/sj.cdd.4401734.
  62. Gao, P.; He, F.-F.; Tang, H.; Lei, C.-T.; Chen, S.; Meng, X.-F.; Su, H.; Zhang, C. NADPH Oxidase-Induced NALP3 Inflammasome Activation Is Driven by Thioredoxin-Interacting Protein Which Contributes to Podocyte Injury in Hyperglycemia. J. Diabetes Res. 2015, 2015, 1–12, doi:10.1155/2015/504761.
  63. Li, H.; Willingham, S.B.; Ting, J.P.-Y.; Re, F. Cutting Edge: Inflammasome Activation by Alum and Alum’s Adjuvant Effect Are Mediated by NLRP3. J. Immunol. 2008, 181, 17–21, doi:10.4049/jimmunol.181.1.17.
  64. Luo, B.; Huang, F.; Liu, Y.; Liang, Y.; Wei, Z.; Ke, H.; Zeng, Z.; Huang, W.; He, Y. NLRP3 Inflammasome as a Molecular Marker in Diabetic Cardiomyopathy. Front. Physiol. 2017, 8, 519, doi:10.3389/fphys.2017.00519.
  65. Levendusky, M.C.; Basle, J.; Chang, S.; Mandalaywala, N.V.; Voigt, J.M.; Dearborn, R.E.J. Expression and regulation of vitamin D3 upregulated protein 1 (VDUP1) is conserved in mammalian and insect brain. J. Comp. Neurol. 2009, 517, 581–600, doi:10.1002/cne.22195.
  66. Aon-Bertolino, M.L.; Romero, J.I.; Galeano, P.; Holubiec, M.; Badorrey, M.S.; Saraceno, G.E.; Hanschmann, E.-M.; Lillig, C.H.; Capani, F. Thioredoxin and glutaredoxin system proteins—Immunolocalization in the rat central nervous system. Biochim. Biophys. Acta Gen. Subj. 2011, 1810, 93–110, doi:10.1016/j.bbagen.2010.06.011.
  67. Blouet, C.; Schwartz, G.J. Nutrient-Sensing Hypothalamic TXNIP Links Nutrient Excess to Energy Imbalance in Mice. J. Neurosci. 2011, 31, 6019–6027, doi:10.1523/jneurosci.6498-10.2011.
  68. Blouet, C.; Liu, S.-M.; Jo, Y.-H.; Chua, S.; Schwartz, G.J. TXNIP in Agrp Neurons Regulates Adiposity, Energy Expenditure, and Central Leptin Sensitivity. J. Neurosci. 2012, 32, 9870–9877, doi:10.1523/jneurosci.0353-12.2012.
  69. Hand, L.E.; Saer, B.R.C.; Hui, S.T.; Jinnah, H.A.; Steinlechner, S.; Loudon, A.S.I.; Bechtold, D.A. Induction of the Metabolic Regulator Txnip in Fasting-Induced and Natural Torpor. Endocrinology 2013, 154, 2081–2091, doi:10.1210/en.2012-2051.
  70. Bodnar, J.S.; Chatterjee, A.; Castellani, L.W.; Ross, D.A.; Ohmen, J.; Cavalcoli, J.; Wu, C.; Dains, K.M.; Catanese, J.; Chu, M.; et al. Positional cloning of the combined hyperlipidemia gene Hyplip1. Nat. Genet. 2002, 30, 110–116, doi:10.1038/ng811.
  71. Donnelly, K.L.; Margosian, M.R.; Sheth, S.S.; Lusis, A.J.; Parks, E.J. Increased lipogenesis and fatty acid reesterification contribute to hepatic triacylglycerol stores in hyperlipidemic Txnip-/- mice. J. Nutr. 2004, 134, 1475–1480.
  72. Lee, K.N.; Kang, H.-S.; Jeon, J.-H.; Kim, E.-M.; Yoon, S.-R.; Song, H.; Lyu, C.-Y.; Piao, Z.-H.; Kim, S.-U.; Han, Y.-H.; et al. VDUP1 Is Required for the Development of Natural Killer Cells. Immunity 2005, 22, 195–208, doi:10.1016/j.immuni.2004.12.012.
  73. Okuyama, H.; Yoshida, T.; Son, A.; Oka, S.-I.; Wang, D.; Nakayama, R.; Masutani, H.; Nakamura, H.; Nabeshima, Y.-I.; Yodoi, J. Thioredoxin Binding Protein 2 Modulates Natural Killer T Cell-Dependent Innate Immunity in the Liver: Possible Link to Lipid Metabolism. Antioxid. Redox Signal. 2009, 11, 2585–2593, doi:10.1089/ars.2009.2691.
  74. Son, A.; Nakamura, H.; Okuyama, H.; Oka, S.-I.; Yoshihara, E.; Liu, W.; Matsuo, Y.; Kondo, N.; Masutani, H.; Ishii, Y.; et al. Dendritic cells derived from TBP-2-deficient mice are defective in inducing T cell responses. Eur. J. Immunol. 2008, 38, 1358–1367, doi:10.1002/eji.200737939.
  75. Oka, S.-I.; Yoshihara, E.; Bizen-Abe, A.; Liu, W.; Watanabe, M.; Yodoi, J.; Masutani, H. Thioredoxin Binding Protein-2/Thioredoxin-Interacting Protein Is a Critical Regulator of Insulin Secretion and Peroxisome Proliferator-Activated Receptor Function. Endocrinology 2008, 150, 1225–1234, doi:10.1210/en.2008-0646.
  76. Chutkow, W.A.; Birkenfeld, A.L.; Brown, J.D.; Lee, H.-Y.; Frederick, D.W.; Yoshioka, J.; Patwari, P.; Kursawe, R.; Cushman, S.W.; Plutzky, J.; et al. Deletion of the α-Arrestin Protein Txnip in Mice Promotes Adiposity and Adipogenesis while Preserving Insulin Sensitivity. Diabetes 2010, 59, 1424–1434, doi:10.2337/db09-1212.
  77. Shi, Y.; Ren, Y.; Zhao, L.; Du, C.; Wang, Y.; Zhang, Y.; Li, Y.; Zhao, S.; Duan, H. Knockdown of thioredoxin interacting protein attenuates high glucose-induced apoptosis and activation of ASK1 in mouse mesangial cells. FEBS Lett. 2011, 585, 1789–1795, doi:10.1016/j.febslet.2011.04.021.
  78. Ren, Y.; Shi, Y.; Wang, Y.; Li, Y.; Wu, S.; Li, H.; Zhang, Y.; Duan, H. p38 MAPK pathway is involved in high glucose-induced thioredoxin interacting protein induction in mouse mesangial cells. FEBS Lett. 2010, 584, 3480–3485, doi:10.1016/j.febslet.2010.07.010.
  79. Shah, A.; Xia, L.; Masson, E.A.Y.; Gui, C.; Momen, A.; Shikatani, E.A.; Husain, M.; Quaggin, S.; John, R.; Fantus, I.G. Thioredoxin-Interacting Protein Deficiency Protects against Diabetic Nephropathy. J. Am. Soc. Nephrol. 2015, 26, 2963–2977, doi:10.1681/asn.2014050528.
  80. Mohamed, I.N.; Sheibani, N.; El-Remessy, A.B. Deletion of Thioredoxin-Interacting Protein (TXNIP) Abrogates High Fat Diet-Induced Retinal Leukostasis, Barrier Dysfunction and Microvascular Degeneration in a Mouse Obesity Model. Int. J. Mol. Sci. 2020, 21, 3983, doi:10.3390/ijms21113983.
  81. Elshaer, S.L.; Mohamed, I.N.; Coucha, M.; Altantawi, S.; Eldahshan, W.; Bartasi, M.L.; Shanab, A.Y.; Lorys, R.; El-Remessy, A.B. Deletion of TXNIP Mitigates High-Fat Diet-Impaired Angiogenesis and Prevents Inflammation in a Mouse Model of Critical Limb Ischemia. Antioxidants 2017, 6, 47, doi:10.3390/antiox6030047.
  82. Katsu-Jiménez, Y.; Vázquez-Calvo, C.; Maffezzini, C.; Halldin, M.; Peng, X.; Freyer, C.; Wredenberg, A.; Giménez-Cassina, A.; Wedell, A.; Arnér, E.S.J. Absence of TXNIP in Humans Leads to Lactic Acidosis and Low Serum Methionine Linked to Deficient Respiration on Pyruvate. Diabetes 2019, 68, 709–723, doi:10.2337/db18-0557.
  83. Mendsaikhan, A.; Takeuchi, S.; Walker, D.G.; Tooyama, I. Differences in Gene Expression Profiles and Phenotypes of Differentiated SH-SY5Y Neurons Stably Overexpressing Mitochondrial Ferritin. Front. Mol. Neurosci. 2018, 11, 470, doi:10.3389/fnmol.2018.00470.
  84. Feng, L.; Zhang, L. Resveratrol Suppresses Aβ-Induced Microglial Activation Through the TXNIP/TRX/NLRP3 Signaling Pathway. DNA Cell Biol. 2019, 38, 874–879, doi:10.1089/dna.2018.4308.
  85. Mohamed, I.N.; Hafez, S.S.; Fairaq, A.; Ergul, A.; Imig, J.D.; El-Remessy, A.B. Thioredoxin-interacting protein is required for endothelial NLRP3 inflammasome activation and cell death in a rat model of high-fat diet. Diabetologia 2014, 57, 413–423, doi:10.1007/s00125-013-3101-z.
  86. Kaya, B.; Erdi, F.; Kılınc, I.; Keskin, F.; Feyzıoglu, B.; Esen, H.; Karataş, Y.; Uyar, M.; Kalkan, E.; Kilinc, I.; et al. Alterations of the thioredoxin system during subarachnoid hemorrhage-induced cerebral vasospasm. Acta Neurochir. 2015, 157, 793–800, doi:10.1007/s00701-015-2390-z.
  87. Cho, M.J.; Yoon, S.-J.; Kim, W.; Park, J.; Lee, J.; Park, J.-G.; Cho, Y.-L.; Kim, J.H.; Jang, H.; Park, Y.-J.; et al. Oxidative stress-mediated TXNIP loss causes RPE dysfunction. Exp. Mol. Med. 2019, 51, 1–13, doi:10.1038/s12276-019-0327-y.
  88. Yao, A.; Storr, S.J.; Al-Hadyan, K.; Rahman, R.; Smith, S.; Grundy, R.; Paine, S.; Martin, S.G. Thioredoxin System Protein Expression Is Associated with Poor Clinical Outcome in Adult and Paediatric Gliomas and Medulloblastomas. Mol. Neurobiol. 2020, 57, 2889–2901, doi:10.1007/s12035-020-01928-z.
  89. Ding, R.; Ou, W.; Chen, C.; Liu, Y.; Li, H.; Zhang, X.; Chai, H.; Ding, X.; Wang, Q. Endoplasmic reticulum stress and oxidative stress contribute to neuronal pyroptosis caused by cerebral venous sinus thrombosis in rats: Involvement of TXNIP/peroxynitrite-NLRP3 inflammasome activation. Neurochem. Int. 2020, 141, 104856, doi:10.1016/j.neuint.2020.104856.
  90. Su, C.-J.; Feng, Y.; Liu, T.-T.; Liu, X.; Bao, J.-J.; Shi, A.-M.; Hu, D.-M.; Liu, T.; Yu, Y.-L. Thioredoxin-interacting protein induced α-synuclein accumulation via inhibition of autophagic flux: Implications for Parkinson’s disease. CNS Neurosci. Ther. 2017, 23, 717–723, doi:10.1111/cns.12721.
  91. Li, L.; Ismael, S.; Nasoohi, S.; Sakata, K.; Liao, F.-F.; McDonald, M.P.; Ishrat, T. Thioredoxin-Interacting Protein (TXNIP) Associated NLRP3 Inflammasome Activation in Human Alzheimer’s Disease Brain. J. Alzheimers Dis. 2019, 68, 255–265, doi:10.3233/jad-180814.
  92. Fertan, E.; Rodrigues, G.J.; Wheeler, R.V.; Goguen, D.; Wong, A.A.; James, H.; Stadnyk, A.; Brown, R.E.; Weaver, I.C.G. Cognitive Decline, Cerebral-Spleen Tryptophan Metabolism, Oxidative Stress, Cytokine Production, and Regulation of the Txnip Gene in a Triple Transgenic Mouse Model of Alzheimer Disease. Am. J. Pathol. 2019, 189, 1435–1450, doi:10.1016/j.ajpath.2019.03.006.
  93. Yamagishi, K.; Yamamoto, K.; Mochizuki, Y.; Nakano, T.; Yamada, S.; Tokiwa, H. Flexible ligand recognition of peroxisome proliferator-activated receptor-gamma (PPARgamma). Bioorg. Med. Chem. Lett. 2010, 20, 3344–3347.
  94. Qi, W.; Chen, X.; Holian, J.; Tan, C.Y.R.; Kelly, D.J.; Pollock, C.A. Transcription Factors Krüppel-Like Factor 6 and Peroxisome Proliferator-Activated Receptor-γ Mediate High Glucose-Induced Thioredoxin-Interacting Protein. Am. J. Pathol. 2009, 175, 1858–1867, doi:10.2353/ajpath.2009.090263.
  95. Billiet, L.; Furman, C.; Larigauderie, G.; Copin, C.; Page, S.; Fruchart, J.; Brand, K.; Rouis, M. Enhanced VDUP-1 gene expression by PPARgamma agonist induces apoptosis in human macrophage. J. Cell. Physiol. 2008, 214, 183–191, doi:10.1002/jcp.21179.
  96. Shaked, M.; Ketzinel-Gilad, M.; Cerasi, E.; Kaiser, N.; Leibowitz, G. AMP-Activated Protein Kinase (AMPK) Mediates Nutrient Regulation of Thioredoxin-Interacting Protein (TXNIP) in Pancreatic Beta-Cells. PLoS ONE 2011, 6, e28804, doi:10.1371/journal.pone.0028804.
  97. Shaked, M.; Ketzinel-Gilad, M.; Ariav, Y.; Cerasi, E.; Kaiser, N.; Leibowitz, G. Insulin counteracts glucotoxic effects by suppressing thioredoxin-interacting protein production in INS-1E beta cells and in Psammomys obesus pancreatic islets. Diabetologia 2009, 52, 636–644, doi:10.1007/s00125-009-1274-2.
  98. Nagaraj, K.; Lapkina-Gendler, L.; Sarfstein, R.; Gurwitz, D.; Pasmanik-Chor, M.; Laron, Z.; Yakar, S.; Werner, H. Identification of thioredoxin-interacting protein (TXNIP) as a downstream target for IGF1 action. Proc. Natl. Acad. Sci. USA 2018, 115, 1045–1050.
  99. Chen, J.; Cha-Molstad, H.; Szabo, A.; Shalev, A. Diabetes induces and calcium channel blockers prevent cardiac expression of proapoptotic thioredoxin-interacting protein. Am. J. Physiol. Metab. 2009, 296, E1133–E1139, doi:10.1152/ajpendo.90944.2008.
  100. Cha-Molstad, H.; Xu, G.; Chen, J.; Jing, G.; Young, M.E.; Chatham, J.C.; Shalev, A. Calcium channel blockers act through nuclear factor Y to control transcription of key cardiac genes. Mol. Pharmacol. 2012, 82, 541–549, doi:10.1124/mol.112.078253.
  101. Melone, M.A.B.; Dato, C.; Paladino, S.; Coppola, C.; Trebini, C.; Giordana, M.T.; Perrone, L. Verapamil Inhibits Ser202/Thr205 Phosphorylation of Tau by Blocking TXNIP/ROS/p38 MAPK Pathway. Pharm. Res. 2018, 35, 44.
  102. Buyandelger, U.; Walker, D.G.; Taguchi, H.; Yanagisawa, D.; Tooyama, I. Novel fluorinated derivative of curcumin negatively regulates thioredoxin-interacting protein expression in retinal pigment epithelial and macrophage cells. Biochem. Biophys. Res. Commun. 2020, 532, 668–674, doi:10.1016/j.bbrc.2020.08.114.
  103. Ren, Y.; Yang, Z.; Sun, Z.; Zhang, W.; Chen, X.; Nie, S. Curcumin relieves paraquat induced lung injury through inhibiting the thioredoxin interacting protein/NLR pyrin domain containing 3 mediated inflammatory pathway. Mol. Med. Rep. 2019, 20, 5032–5040, doi:10.3892/mmr.2019.10612.
  104. Lu, X.; Wu, F.; Jiang, M.; Sun, X.; Tian, G. Curcumin ameliorates gestational diabetes in mice partly through activating AMPK. Pharm. Biol. 2019, 57, 250–254, doi:10.1080/13880209.2019.1594311.
More