s2fieldsofstudy: Biology Chemistry

text:
Protein Targeting to Glycogen (PTG): A Promising Player in Glucose and Lipid Metabolism

Protein phosphorylation and dephosphorylation are widely considered to be the key regulatory factors of cell function, and are often referred to as “molecular switches” in the regulation of cell metabolic processes. A large number of studies have shown that the phosphorylation/dephosphorylation of related signal molecules plays a key role in the regulation of liver glucose and lipid metabolism. As a new therapeutic strategy for metabolic diseases, the potential of using inhibitor-based therapies to fight diabetes has gained scientific momentum. PTG, a protein phosphatase, also known as glycogen targeting protein, is a member of the protein phosphatase 1 (PP1) family. It can play a role by catalyzing the dephosphorylation of phosphorylated protein molecules, especially regulating many aspects of glucose and lipid metabolism. In this review, we briefly summarize the role of PTG in glucose and lipid metabolism, and update its role in metabolic regulation, with special attention to glucose homeostasis and lipid metabolism.

Introduction
Carbohydrates and lipids are the main sources of energy required by organisms to maintain life activities. The homeostasis imbalance of the regulatory network is also an important inducing factor for the occurrence and development of a variety of metabolic diseases [1][2][3]. Glucose and lipid metabolism disorder is the main clinical phenotype of obesity, type 2 diabetes and nonalcoholic fatty liver disease, and can also induce cardiovascular and cerebrovascular diseases [4]. Under the influence of genetic and/or environmental factors, the decrease of insulin secretion or the disturbance of signal transduction (insulin resistance) will lead to the decrease of energy supply from glucose sources and the enhancement of lipid decomposition and release, and can stimulate liver gluconeogenesis, promote liver glucose release, and participate in the occurrence and development of diabetes and its complications [5,6]. In addition, hormones such as estrogen, cortisol, and insulin, and excess nutrition, can promote the liver to synthesize more triglycerides, and excess glucose can also be transformed into triglycerides through a lipogenesis reaction [7]. These excessively synthesized triglycerides accumulate in the liver, which may lead to the formation of nonalcoholic fatty liver disease [3].The disorder of liver lipid metabolism can cause liver steatosis, obesity, insulin resistance, and can aggravate the process of diabetes [8,9].With the rapid increase of the prevalence of these metabolic diseases in the world, it is more and more important to explore the regulation of glucose and lipid metabolism and its function in the homeostasis.
The liver plays an important role in various organs involved in glucose and lipid metabolism [10]. The homeostasis of intracellular glucose and lipid mainly depends on the liver [11,12]. Glycogen and lipid are the main forms of energy storage, which are strictly regulated by hormones and metabolic signals and provide energy at different stages [13]. When the postprandial blood glucose concentration increases, glucose is firstly phosphorylated to glucose 6-phosphate after it is transported to hepatocytes with blood [14]. Glucose 6-phosphate can no longer penetrate the cell membrane and return to the blood. Instead, it acts as the source of carbohydrate metabolism in the liver, opening up metabolic pathways such as glycogen synthesis, glycolysis, pentose phosphorylation and lipogenesis. Under starvation, hepatocytes can also convert non-glucose substances such as lactic acid and glycogenic amino acids into glucose-6-phosphate through the process of gluconeogenesis [15,16]. The liver is one of the two major sites for the conversion of carbohydrates into fats in the human body, and is also the regulatory center for the digestion, absorption, synthesis and transportation of lipids [10]. It promotes the digestion and absorption of lipids by secreting bile, and is the most important place for the synthesis of lipids such as triglycerides, fatty acids, cholesterol, and ketones ( Figure 1). Under normal circumstances, these lipid metabolites are transported by the blood to extrahepatic tissues for storage or utilization after being synthesized in the liver [7,17].
Biomolecules 2022, 12, x FOR PEER REVIEW 2 of 13 strictly regulated by hormones and metabolic signals and provide energy at different stages [13]. When the postprandial blood glucose concentration increases, glucose is firstly phosphorylated to glucose 6-phosphate after it is transported to hepatocytes with blood [14]. Glucose 6-phosphate can no longer penetrate the cell membrane and return to the blood. Instead, it acts as the source of carbohydrate metabolism in the liver, opening up metabolic pathways such as glycogen synthesis, glycolysis, pentose phosphorylation and lipogenesis. Under starvation, hepatocytes can also convert non-glucose substances such as lactic acid and glycogenic amino acids into glucose-6-phosphate through the process of gluconeogenesis [15,16]. The liver is one of the two major sites for the conversion of carbohydrates into fats in the human body, and is also the regulatory center for the digestion, absorption, synthesis and transportation of lipids [10]. It promotes the digestion and absorption of lipids by secreting bile, and is the most important place for the synthesis of lipids such as triglycerides, fatty acids, cholesterol, and ketones ( Figure 1). Under normal circumstances, these lipid metabolites are transported by the blood to extrahepatic tissues for storage or utilization after being synthesized in the liver [7,17]. The enrichment of glucose-induced glucokinase (GCK), which phosphorylated glucose to glucose-6-phosphate (G-6-P); it is used as a substrate for glycolysis or glycogen synthesis, depending on the nutritional status. The product pyruvate is further decarboxylated to acetyl coenzyme A and enters the tricarboxylic acid cycle (TCA) in mitochondria. Acetyl coenzyme A or malonyl coenzyme A synthesize fatty acids from scratch and further process them into TAG. In mammals, FA synthesis is catalyzed by acetyl CoA carboxylase (ACC) and fatty acid synthase (FAS).
Phosphorylation and dephosphorylation of proteins are widely recognized as key regulators of cellular function [18,19] and are often referred to as "molecular switches" in the regulation of cellular physiological processes [20,21]. Protein phosphorylation and dephosphorylation can regulate signal transduction in a variety of ways, including by activating or inhibiting conformational changes to create binding sites for proteins containing specific domains, thereby affecting the conformation of proteins or proteinprotein interactions, and controlling their cellular localization [22,23]. Numerous studies have shown that the phosphorylation/dephosphorylation of related signaling molecules plays a key role in the regulation of hepatic glucose and lipid metabolism [24][25][26][27]. Glycogen targeting protein (PTG) is a member of the protein phosphatase 1 (PP1) family and can play a role by catalyzing the dephosphorylation of phosphorylated protein molecules. Previous studies have shown that it can affect glycogen metabolism in liver and skeletal muscles by regulating the dephosphorylation of ribosomal S6 protein kinase, glycogen synthase, and glycogen phosphorylase [28,29]. This article reviews the research progress of protein targeting to glycogen (PTG) in glucose and lipid metabolism in recent years, and provides new insights for the prevention and treatment of metabolism-related diseases and clinical transformation research.

Structure and Characteristics of PP1
Protein phosphorylation is an important cellular regulatory mechanism, and many enzymes and receptors are activated/deactivated by phosphorylation and dephosphorylation events through kinases and phosphatases [22,23]. It is generally believed that phosphorylation or dephosphorylation of proteins is one of the initial steps in the coordination of cell and organ functions, such as regulation of glycolipid metabolism, cell proliferation, apoptosis, inflammation, and other important physiological processes. Protein kinases and protein phosphatases catalyze this process [24][25][26][27]. Therefore, kinases and phosphatases are promising targets for regulating this reaction, and many inhibitors against these proteins have been developed [30]. Krzyzosiak et al. [31] discovered Raphin1, a serine/threonine phosphatase inhibitor that mainly inhibits PPP1R15B, which is reported to be a potential drug candidate for the treatment of Huntington's disease. Furthermore, some studies have pointed out that PTP1B inhibitors are considered to have obvious antidiabetic potential [32]. Therefore, the study of protein phosphatase is of great significance.
The PP1 regulatory subunit, also known as glycogen targeting regulatory subunit, coordinates glycogen synthesis by targeting the catalytic subunit of protein phosphatase 1 to glycogen particles [33][34][35]. They together constitute the PP1 holoenzyme, which regulates the activity of glycogen metabolizing enzymes, and plays a role through PP1 catalyzing the dephosphorylation of phosphorylated protein molecules, mainly dephosphorylating glycogen synthase phosphatase (GSP) and activating glycogen synthase (GS), which in turn stimulates glycogen production [33,36]. According to the GenBank database, there are seven genes encoding G subunit (PPP1R3C-PPP1R3G), all of which have PP1 binding domain and glycogen binding domain. Previous studies have found that many subtypes of ppp1r (PPP1R3A, B, G) can promote glycogen storage when overexpressed in hepatocytes [37,38]. PTG is a subtype of the protein phosphatase 1 family, encoding protein phosphatase regulatory subunit 3C, which can play a role by catalyzing dephosphorylation of phosphorylated protein molecules [39]. The PTG gene is located on human chromosome 10 and mouse chromosome 19. The full length of its cDNA is 1499 BP, encoding 285 amino acid residues [28]. It was found that PTG was expressed in muscle, liver, adipose tissue, heart, brain and other tissues of mice, especially highly expressed in liver and muscle, and has a variety of important biological roles [28,29,40,41]. Previous studies have found that PTG is also involved in the occurrence and development of colorectal cancer [42], and renal cell carcinoma [43].
At present, the prevalence of related diseases caused by disorders of glucose and lipid metabolism is increasing substantially, and a better understanding of the regulation of glucose and lipid metabolism is essential for the prevention and treatment of related diseases and the improvement of people's quality of life. In recent years, many researchers have conducted in-depth research on the role of PTG in glucose and lipid metabolism [29,40,44].

Overview of PTG and Its Regulatory Factors
Studies have shown that PTG was mainly expressed in insulin-sensitive tissues such as liver and muscle [39], and the expression of PTG in the liver of mice fed with a highfat diet was significantly increased [45]. More interestingly, Munro et al. [46] found that the phosphatase activities related to PP1 and PTG were down-regulated by 60-70% in streptozotocin-induced diabetes and recovered through insulin treatment, indicating that the phosphatase activities such as PTG were regulated by insulin. Allaman et al. [47] found that both norepinephrine and adenosine can up-regulate PTG mRNA levels in astrocytes and hepatocytes, while increasing glycogen synthesis. Mapping the major phosphorylation sites of PTG by mass spectrometry analysis, Vernia et al. [48] found that phosphorylation activation of AMPK Ser-8 and Ser-268 increases the formation of a complex of dual-specificity phosphatase and E3 ubiquitin ligase and down-regulates PTG activity, resulting in a decrease in its glycogen activity. This study further confirmed that AMPK can physically interact with PTG and change its basal phosphorylation state. Cheng et al. [41] found multiple forkhead box protein A2 (FoxA2) binding sites in the promoter region of PTG, and further constructed a luciferase reporter gene to confirm that FoxA2 transactivates the PTG promoter in H4IIE liver cancer cells. Likewise, nuclear extracts from mouse liver and H4IIE cells were able to bind FoxA2-specific probes derived from the PTG promoter region. Chromatin immunoprecipitation experiments further demonstrated that FoxA2 could bind to the PTG promoter in vivo. Furthermore, adenosine-3',5'-cyclic monophosphate (cAMP) treatment also activated the PTG promoter and significantly increased the level of PTG in H4IIE cells. In general, a variety of glucose metabolismrelated signaling molecules are involved in the regulation of PTG.

PTG and Glucose Metabolism
The liver is essential for maintaining normal glucose homeostasis-it produces glucose when fasting and stores it after meals. The maintenance of this homeostasis is multifaceted, mainly through the regulation of gluconeogenesis, glycogen synthesis, glycogenolysis, and glycolysis [13]. The liver has a remarkable ability to produce glucose, which is released into the systemic circulation and utilized by other tissues. Glucose production in the liver results from glycogenolysis and de novo synthesis of glucose (gluconeogenesis). Normally, the production and uptake of glucose are balanced to maintain glucose homeostasis in human body [10]. After a meal, the pancreas β cells secrete postprandial insulin, and the increased insulin signal activates AKT in the cells. Subsequently, glycogen synthase kinase-3β (GSK-3β) is phosphorylated and inactivated, while glycogen synthase is activated, inhibiting the decomposition of glycogen, promoting the synthesis and storage of glycogen, and finally reducing blood glucose levels [49,50]. Additionally, glucagon and adipose free fatty acid release were also indirectly inhibited, resulting in the inhibition of the gluconeogenesis process [51]. Conversely, during fasting, pancreatic α cells can secrete glucagon to act on liver tissue, inactivating glycogen synthase, thereby inhibiting glycogen synthesis. During short-term fasting, glycogenolysis is the main source of glucose released into the blood, and stored glycogen is further hydrolyzed into glucose to increase glucose production in the liver. However, during prolonged periods of fasting, glycogen stores are gradually depleted and glycogenolysis decreases [13]. Under the action of phosphoenolpyruvate carboxy kinase (PEPCK) and glucose-6-phosphatase (G6Pase), non-carbohydrate precursors can resynthesize glucose in the liver, known as gluconeogenesis [52]. In addition, insulin levels also decreased, and the proportion of peripheral tissues using glucose as fuel decreased to maintain blood glucose levels [53]. With the prolongation of fasting time, the contribution of gluconeogenesis to hepatic glucose production gradually increased to increase blood glucose [11].

PTG and Glycogen Synthesis
Glycogen synthesis is dynamically regulated by insulin through synergistic dephosphorylation of the serine/threonine phosphatase PP1 and the glycogenolytic enzyme glycogen phosphorylase (GP) on glycogen synthase (GS), leading to the activation of GS and the deactivation of GP [54]. In the liver, PTG and PPP1R3B(GL)are expressed at roughly equivalent levels [55], and they jointly promote hepatic glycogen mobilization and storage. PTG overexpression significantly increased glycogen content, mainly due to its ability to promote the redistribution of PP1 and glycogen synthase to glycogen granules, significantly increasing GS activity and glycogen synthesis ( Figure 2) [28,29]. Studies also noted that PTG recruited phosphatases to where glycogen synthesis occurs, allowing the activation of glycogen synthase and deactivation of glycogen phosphorylase, thereby increasing glycogen synthesis and reducing its degradation [55]. Further study also found that overexpression of the PP1 subunit protein targeting PTG could significantly increase the level of cell glycogen. At the same time, on the fourth day after PTG knockdown, the glycogen level decreased by more than 85%, which reduced the glycogen targeted PP1 protein level, and correspondingly reduced the activity of glycogen synthase and phosphorylase targeted PP1 in cells, and the glycogen decomposition rate increased significantly under PTG knockdown, indicating that PTG mainly played a role in inhibiting glycogen decomposition [56]. This notion is supported by the results of Crosson et al. [44], in which mice with heterozygous deletion of the PTG gene had reduced glycogen storage in adipose tissue, liver, heart and skeletal muscle. Accordingly, the activity of glycogen synthase and the rate of glycogen synthesis decreased. Furthermore, Greenberg et al. [57] found that overexpression of PTG in 3T3-L1 adipocytes discretely stimulated the activity of PP1 on glycogen synthase and phosphorylase, resulting in increased uptake and storage of glucose as glycogen. Similarly, Printen et al. [29] found that PTG also forms complexes with phosphorylase kinase, phosphorylase a and glycogen synthase in 3T3-L1 adipocytes. The latter are the main enzymes involved in the hormone regulation of glycogen metabolism. Further studies by Lu et al. [45] found that feeding mice with a high-fat diet could increase hepatic glycogen, which was due to the increased expression of the glycogen scaffolding protein PTG. Activation of mTORC1 and its downstream target SREBP1 resulted in increased PTG promoter activity and increased glycogen levels in mice and cells. In contrast, deletion of the PTG gene in mice prevented HFD-induced hepatic glycogen accumulation. López-Soldado et al. [14] further bred liver PTG-overexpressing mice, which could maintain relatively high hepatic glycogen levels even when fasting, thereby maintaining hepatic energy status. Some researchers hybridized Akita mice with PTG overexpressing mice in the liver to construct Akita PTG OE mice with increased liver glycogen content. The blood glucose level of Akita PTG OE mice decreased gradually with the progress of the disease. Akita PTG OE mice have stronger glycogen production ability, which shifts blood glucose to glycogen synthesis, so the above results suggest that long-term enhancement of hepatic glycogen may have the effect of reducing hyperglycemia [58]. It can be seen that PTG, as a protein phosphatase with dephosphorylation, plays a very important role in regulating energy metabolism in the liver.
PTG also plays an important role as a regulator in the regulation of glycogen metabolism in cells other than the liver. Researchers found that the gene encoding PTG is abundantly expressed in astrocytes of the central nervous system, and some evidence suggests that PTG plays an important role in the regulation of glycogen metabolism in this cell line [59]. Allaman et al. [47] found that in astrocytes, overexpression of PTG resulted in a 100-fold increase in glycogen, a downregulation of endogenous PTG expression by siRNA resulted in a two-fold decrease in glycogen, and knockdown of PTG expression (PTG-KO astrocytes) resulted in an 80 % decrease in glycogen. Interestingly, the reduction in glycogen content measured in PTG-KO astrocytes (80%) was similar to that observed in the brains of PTG-KO mice (70%), further indicating the close parallel relationship between glycogen metabolism in cultured astrocytes and brain glycogen metabolism. These observations suggest that PTG is also a master regulator of astrocyte glycogen, a central element of glycogen regulation in vivo. Furthermore, changes in PTG expression in the brain have been shown to be associated with different behavioral conditions related to glycogen mobilization, such as sleep deprivation and learning and memory [60][61][62]. Astrocytes are the most widely distributed type of cells in the mammalian brain and the largest type of glial cells. It is also the main storage site of glycogen in the brain. When neurons are highly active and the blood glucose provided through the blood brain barrier cannot meet the needs, glycogen in glial cells can be decomposed into glucose to provide energy for neurons under the effect of neurotransmitters [63]. In cultured astrocytes isolated from a mouse model of schizophrenia, a strong reduction in PTG expression was also observed, which is associated with altered glycogen metabolism [59]. The above results suggest that PTG can control brain glycogen metabolism under physiological and pathological conditions (such as psychiatric disorders), thereby controlling the key role of astrocyte function.
x FOR PEER REVIEW 6 of 1 Figure 2. Possible PTG mediated crosstalk mechanism of glucose and lipid metabolism. The effe of PTG on fat metabolism is still controversial, but it can promote fat synthesis and promote th expression of related genes, such as SREBP1, FAS, SCD1, ACC. In addition to influencing glycoge synthesis, recent studies also found that PTG can promote the expression of PEPCK and G6Pas key enzymes of gluconeogenesis, and then participate in gluconeogenesis.
PTG also plays an important role as a regulator in the regulation of glycogen metab olism in cells other than the liver. Researchers found that the gene encoding PTG is abun dantly expressed in astrocytes of the central nervous system, and some evidence sugges that PTG plays an important role in the regulation of glycogen metabolism in this cell lin [59]. Allaman et al. [47] found that in astrocytes, overexpression of PTG resulted in a 100 fold increase in glycogen, a downregulation of endogenous PTG expression by siRN resulted in a two-fold decrease in glycogen, and knockdown of PTG expression (PTG-K astrocytes) resulted in an 80 % decrease in glycogen. Interestingly, the reduction in glyco gen content measured in PTG-KO astrocytes (80%) was similar to that observed in th brains of PTG-KO mice (70%), further indicating the close parallel relationship betwee glycogen metabolism in cultured astrocytes and brain glycogen metabolism. These obse vations suggest that PTG is also a master regulator of astrocyte glycogen, a central elemen of glycogen regulation in vivo. Furthermore, changes in PTG expression in the brain hav Possible PTG mediated crosstalk mechanism of glucose and lipid metabolism. The effect of PTG on fat metabolism is still controversial, but it can promote fat synthesis and promote the expression of related genes, such as SREBP1, FAS, SCD1, ACC. In addition to influencing glycogen synthesis, recent studies also found that PTG can promote the expression of PEPCK and G6Pase, key enzymes of gluconeogenesis, and then participate in gluconeogenesis.
A study of skeletal muscle glycogen levels showed that an overexpression of interferon regulatory factor 4 (IRF4) in mice exhibited decreased exercise capacity and decreased glycogen content. Furthermore, PTG expression increased in the absence of IRF4, and decreased when IRF4 was overexpressed. This indicates that IRF4 regulates the glucose metabolism of skeletal muscle by regulating PTG, and the overexpression of IRF4 leads to the low expression of PTG, thereby reducing the content of glycogen in skeletal muscle. However, opinions on the regulation of muscle glycogen levels by PTG are divided. Studies have shown that heterozygous PTG deletion does not result in impaired glycogen synthesis and reduced glycogen levels in the gastrocnemius muscle. This study also found that PTG in C2C12 myoblasts did not significantly change the content of glycogen. These results indicate that the effect of PTG on skeletal muscle glycogen depends on IRF4 [64].

PTG and Gluconeogenesis
The liver plays a major role in maintaining normal glucose homeostasis by controlling the balance between hepatic glucose production and storage [15]. During fasting, an increase in circulating glucagon triggers the gluconeogenesis by activating the cAMP pathway [65]. It has been reported that norepinephrine could stimulate PTG expression in mouse cortical astrocytes via the cAMP pathway [66]. PTG was significantly up-regulated during 3T3-L1 adipocyte differentiation, which involved the cAMP signaling pathway [67]. FoxA2, a transcriptional regulator of hepatic gluconeogenesis genes, mediates cAMPstimulated PTG transcription by binding to promoters in hepatocytes [41]. Recent studies have found that the phosphorylation/dephosphorylation of related signaling molecules plays a key role in the regulation of hepatic gluconeogenesis, which is mainly regulated by phosphorylase and dephosphorylase. Various hormones can regulate glucose metabolism by affecting the phosphorylation and dephosphorylation status of liver enzymes [68]; glucagon activates adenylate cyclase to produce cAMP, which, on the one hand, activates the cAMP-dependent protein kinase, which causes the phosphorylation of pyruvate kinase to inhibit the effect of pyruvate kinase, and on the other hand, it can also promote the phosphorylation of fructose 2,6-bisphosphatase, which in turn inhibits the glycolytic pathway, stimulates gluconeogenesis, and promotes glucose production, while insulin exhibits the opposite effect [6,69]. In addition, glucagon can also activate peroxisome proliferatoractivated receptor gamma coactivator 1α (PGC-1α) to dephosphorylate. Dephosphorylated PGC-1α will be transferred to the nucleus and combined with hepatocyte nuclear factor 4α (HNF4α) to form a complex to activate it. In this way, the transcription of G6Pase and PEPCK-encoding genes is initiated, which affects gluconeogenesis [65,69]. Previous studies also found that glucagon can promote the nuclear translocation and stability of FOXO1 through cAMP-dependent and protein kinase α-dependent phosphorylation of FOXO1, thereby affecting gluconeogenesis [70]. Uebi et al. [71] found that PP1 can dephosphorylate and activate the Ser171 site of CREB-regulated transcriptional coactivator 2 (TORC2). Phosphorylated TORC2 remains inactive in the cytosol, while dephosphorylated TORC2 is translocated to the nucleus. It binds with phosphorylated cAMP response element binding protein (CREB) to form a CREB-TORC2 complex, which increases the expression of key gluconeogenesis enzymes such as G6Pase and PEPCK. Qi et al. [72] found that follicle-stimulating hormone can promote the membrane translocation of G proteincoupled receptor kinase 2 (GRK2), resulting in the phosphorylation of serine 485 of AMPK α and the dephosphorylation of threonine 172. In the nucleus, it promotes gluconeogenesis, while TORC2 translocates into the nucleus and promotes gluconeogenesis. As a member of the PP1 family, PTG has a dephosphorylation effect, suggesting that PTG may be closely related to hepatic gluconeogenesis. Ji et al. [73] found that overexpression of PTG in primary mouse hepatocytes or wild-type mouse liver promoted hepatic glucose production and expression of gluconeogenesis genes. Conversely, PTG knockout reduced hepatic gluconeogenesis and suppressed cAMP-stimulated gluconeogenic gene expression and TORC2 dephosphorylation. Animal studies showed that PTG knockdown in the liver of db/db mice significantly improved blood glucose levels and reduced the expression of key genes of gluconeogenesis ( Figure 2). However, other researchers constructed PTG OE hybridized mice with PTG overexpression. The results showed that, compared with normal mice, PTG OE mice showed decreased liver gluconeogenesis and increased glycolysis, and PGC1 α and PEPCK expression levels decreased, further reducing the blood glucose level of the mice [58]. The possible reason for the difference is that, first of all, the models of the two research teams are from different strains of mice, and the differences in genetic background may lead to abnormal results. Secondly, the methods of overexpression and knockdown of PTG adopted by the two groups were inconsistent, which may also be the reason for the difference in the results. In addition, the influence of living environment and intestinal flora on the results cannot be ruled out. Previous studies have shown that hepatic glycogen synthesis is closely related to the process of gluconeogenesis, and that, in addition to glucose uptake, hepatic gluconeogenesis flux also determines the amount of glycogen formed, especially in the fasted state [1,74,75]. Overexpression of PTG in the liver stimulates glycogen synthesis mainly from gluconeogenic precursors through an indirect pathway. The roles and mechanisms of PTG in gluconeogenesis have not been fully elucidated, and more in-depth studies are still needed.

PTG and Lipid Metabolism
Liver is the central organ that controls lipid homeostasis by precisely regulating various biochemical, signaling and cellular pathways. Hepatic de novo lipogenesis is a fundamental biosynthetic pathway in the liver that helps hepatocytes store synthetic lipids [76]. This process is an extension of the complex metabolic network within the liver, providing substrates primarily through glycolysis and carbohydrate metabolism. Longterm unhealthy lifestyle and overnutrition can lead to disorders of hepatic lipid metabolism, leading to serious lipid-related diseases, including obesity, NAFLD, and T2DM [77,78].

PTG and Fatty Acid Metabolism
The liver is the main site of fatty acid metabolism and the main source of ketogenesis. Fasting will increase the synthesis of TG in the liver, and the promotion of liver steatosis is due to excessive intake of circulating fatty acids produced by fat decomposition of adipose tissue [79]. It was found that prolonged fasting led to liver TG level, plasma βhydroxybutyric acid, FFA concentration and plasma FGF21 level being significantly increased in the normal feeding of mice, but the increase was significantly reduced in PTG overexpression mice. In the further investigation, prolonged fasting also promoted the expression of the fatty acid metabolism related genes HMGCS2, CD36, PPAR α, CPT1 α and FGF21, while PTG overexpression significantly weakened this effect [14]. This suggests that PTG may be involved in fatty acid uptake or oxidation in the liver. Further research needs to be carried out to clarify this metabolic effect.

PTG and Fat Synthesis
The main feature of liver fat metabolism disorder is excessive accumulation of TG in the liver [80]. Under normal circumstances, the liver only stores a small amount of fatty acids as triglycerides. However, in addition to increasing the intake of free fatty acids, increased fat synthesis can also lead to liver steatosis [81]. In the case of over nutrition and obesity, the metabolism of fatty acids in the liver changes, which usually leads to the accumulation of triglycerides in the liver cells, resulting in liver steatosis and various metabolic related diseases, such as nonalcoholic fatty liver disease, insulin resistance, and T2DM [3,77]. Studies have shown that circulating hyperglycemia enters the liver and is converted into glycogen for storage. When there is too much glycogen, glucose enters the glycolytic pathway, and the pyruvate and acetyl coenzyme A produced will generate malonyl coenzyme A under the action of ACC, and then FAS catalyzes the production of long-chain saturated fatty acids to promote fatty acid synthesis [12]. Moreover, glucose can also promote the expression of SREBP-1 [82], which can up-regulate the expression of ACC and FAS genes and increase the synthesis of fatty acids [83]. Previous studies have found that PTG mainly plays an important regulatory role in glycogen synthesis ( Figure 2) [28,29,40]. The latest evidence has shown that PTG-silencing mice can prevent the accumulation of liver glycogen induced by high fat, reduce fasting blood glucose and insulin levels, and improve insulin sensitivity; mTORC1/SREBP1 can act on PTG promoter, regulate PTG transcription and affect glycogen metabolism, and glycogen accumulation can regulate SREBP1 expression through feedback [45]. More interestingly, the study by Wang et al. [84] found that AMPK inhibits mTORC1 activity by phosphorylating mTORC1 pathway members TSC2 and Raptor, thereby preventing protein and lipid synthesis. In the adipose tissue of mice PTG was overexpressed, using the fatty acid binding protein 4 (FABP4) promoter, and the results showed that glycogen levels in the adipose tissue were 200 to 400 times higher than in wild-type mice, which shows that adipocytes have the steric capacity to accommodate high levels of glycogen (Figure 2), and the importance of glycogen in fat deposition is demonstrated. The study found that transgenic overexpression of PTG in adipose tissue increased glucose flux into the glyco-gen synthesis pathway, indicating that adipocytes are capable of storing high levels of glycogen [54]. Cui et al. [85] found that PTG can promote the adipocyte differentiation of 3T3-L1 preadipocytes, and found that knockdown of SREBP1 gene expression led to downregulation of PTG gene expression, speculating that the effect of PTG on adipogenesis may be regulated by SREBP1 in 3T3-L1 to be realized. Lu et al. [45] found that PTG silencing in mice could prevent high-fat-induced accumulation of hepatic glycogen, reduce hepatic lipid deposition, reduce fasting glucose and insulin levels, and improve insulin sensitivity, while Rapamycin complex 1 (mTORC1) and cholesterol regulatory element binding protein 1 (SREBP1) can act on the PTG promoter to regulate the transcription of PTG and affect the metabolism of glycogen. Moreover, the accumulation of glycogen can regulate SREBP1 through feedback, thereby affecting fat metabolism, establishing a dialogue between hepatic glucose and lipid metabolism, and regulating energy balance, indicating that PTG plays an important role as a bridge in glycogen and lipid energy balance. It was found that the reduced liver adipogenesis in normal Akita mice was restored in Akita PTG OE mice, and the expression of FAS and SCD1, the key regulatory factors of adipogenesis, was up-regulated. Increased lipogenesis may be due to increased hepatic glucokinase, as glucokinase overexpression has been reported to lead to increased hepatic glycolytic flux, resulting in increased concentrations of glycerol-3-phosphate and malonyl-CoA, the latter of which are a nascent fat generated substrate. More importantly, Akita PTG OE mice had reduced hepatic lipolytic capacity. Thus, the increase in hepatic TG observed in Akita PTG OE mice was associated with higher lipogenesis and lower lipolysis in the liver [58].

Conclusions
In conclusion, PTG is a pleiotropic and promising protein phosphatase which seems to play an important role in every link of glucose and lipid metabolism, and plays different key roles in different physiological conditions and pathological abnormalities. It may be an important protein that leads to the interaction of abnormal liver glucose and lipid metabolism. PTG may become a new therapeutic target for metabolic diseases in the near future, with great clinical value. Of course, it is worth further exploring gluconeogenesis and lipid metabolism and its clinical application in transformation.
Author Contributions: Conceptualization, G.Y.; first draft, X.D. and C.W.; writing, review and editing, G.Y. and X.D.; map: C.W. and Y.X. All authors have read and agreed to the published version of the manuscript.


Herpes simplex virus type 2 inhibits TNF-α-induced NF-κB activation through viral protein ICP22-mediated interaction with p65

Herpes simplex virus type 2 (HSV-2) is a prevalent human pathogen and the main cause of genital herpes. After initial infection, HSV-2 can establish lifelong latency within dorsal root ganglia by evading the innate immunity of the host. NF-κB has a crucial role in regulating cell proliferation, inflammation, apoptosis, and immune responses. It is known that inhibition of NF-κB activation by a virus could facilitate it to establish infection in the host. In the current study, we found that HSV-2 inhibited TNF-α-induced activation of NF-κB-responsive promoter in a dose-dependent manner, while UV-inactivated HSV-2 did not have such capability. We further identified the immediate early protein ICP22 of HSV-2 as a vital viral element in inhibiting the activation of NF-κB-responsive promoter. The role of ICP22 was confirmed in human cervical cell line HeLa and primary cervical fibroblasts in the context of HSV-2 infection, showing that ICP22 deficient HSV-2 largely lost the capability in suppressing NF-κB activation. HSV-2 ICP22 was further shown to suppress the activity of TNF receptor-associated factor 2 (TRAF2)-, IκB kinase α (IKK α)-, IKK β-, IKK γ-, or p65-induced activation of NF-κB-responsive promoter. Mechanistically, HSV-2 ICP22 inhibited the phosphorylation and nuclear translocation of p65 by directly interacting with p65, resulting in the blockade of NF-κB activation. Furthermore, ICP22 from several alpha-herpesviruses could also inhibit NF-κB activation, suggesting the significance of ICP22 in herpesvirus immune evasion. Findings in this study highlight the importance of ICP22 in inhibiting NF-κB activation, revealing a novel mechanism by which HSV-2 evades the host antiviral responses.

Introduction
HSV-2 is a large dsDNA virus, a member of the Herpesviridae family, belonging to genus Simplexvirus. The WHO reported that about 13% of the world's population aged 15 to 49 years were living with HSV-2 infection in 2016 (1). HSV-2 is mainly sexually transmitted and infects epithelial cells, causing genital herpes (2). It also infects leukocytes and neuronal cells (3)(4)(5), leading to encephalitis and disseminated diseases that affect other organ systems (6,7). After initial infection, HSV-2 can establish life-long latency in the dorsal root ganglia (8).
It is known that HSV (HSV-1/2) has evolved countermeasures to evade the host innate immune responses. However, most of the studies to date have been focusing on HSV-1 (9). For instance, the UL2, UL24, UL42, RL2 and US3 of HSV-1 can interfere with the NF-kB signaling pathway by interacting with p65 or p50, the key component of NF-kB heterodimer (10)(11)(12)(13)(14), while HSV-1 UL36 suppresses NF-kB activation by cleaving the polyubiquitin chains of IkB a, an inhibitor of NF-kB activation (15). The RL2, RL1 and UL48 of HSV-1 suppress the production of type I interferons (IFN) by acting on IRF3 (16,17), whereas the UL54, US11 and UL46 of HSV-1 act on TBK1 to block the production of type I IFN (18)(19)(20). In addition, the RL2 and UL41 of HSV-1 were reported to suppress the production of IFN-stimulated genes (ISGs) by degrading IFI16 (21). So far, little is known concerning how HSV-2 evades the host innate immune system. We previously demonstrated that the immediate early protein ICP22 of HSV-2 not only suppresses IFN-b production by blocking the association of IRF-3 with IFN-b promoter (22), but also inhibits the production of ISGs by directly degrading IFNstimulated gene factor 3 (ISGF3) (23). Although ICP22 is a key mediator of HSV-2 immune evasion in type I IFN production and signaling, it remains to be determined whether HSV-2 ICP22 could inhibit the activation of NF-kB signaling pathway.
TNF receptor-associated factor 2 (TRAF2), IkB kinase (IKK) a (IKK a), IKK b, IKK g and p65 are key components of the NF-kB signaling pathway. Under foreign stimuli, the IKK complex is first activated, resulting in the phosphorylation of IkB proteins and its subsequent degradation by the proteasome. Once IkB protein is detached, released NF-kB dimers are phosphorylated and then translocate to the nucleus to activate the transcription of target genes (24)(25)(26). Given that a number of viruses were previously shown to block the activation of NF-kB by acting on different components of NF-kB signaling pathway (27)(28)(29)(30)(31)(32)(33), we asked whether and how HSV-2 inhibits NF-kB activation.
In this study, we first revealed that HSV-2 blocks TNF-ainduced activation of NF-kB-responsive promoter. We further demonstrated that the immediate early protein ICP22 of HSV-2 inhibits NF-kB activation by directly interacting with p65, resulting in the blockade of p65 phosphorylation and nuclear translocation.
Dual luciferase reporter assay HEK 293T cells seeded in 24-well plates overnight were cotransfected with Firefly luciferase reporter plasmid pNF-kB-Luc, Renilla luciferase reporter plasmid phRL-TK and empty vector or plasmid encoding indicated viral protein. Transfections were performed using Lipofectamine 2000 (Invitrogen, 11668-027) according to the manufacturer's instructions. At 24 h posttransfection, cells were mock-treated or treated with TNF-a (20 ng/ml) for 6 h. In some cases, HEK 293T cells were cotransfected with pNF-kB-Luc, phRL-TK, plasmid expressing pFlag-p65, pHA-TRAF2, pHA-IKK a, pHA-IKK b or pHA-IKK g and empty vector or plasmid expressing indicated viral protein for 30 h. For HeLa cells, after co-transfection with reporter plasmids pNF-kB-Luc and phRL-TK for 4 h, cells were infected or mock infected with HSV-2, UV-inactivated HSV-2 or us1 del HSV-2 for 20 h, followed by stimulation with or without TNF-a (20 ng/ml) for 6 h. For primary human cervical fibroblasts, after co-transfection with reporter plasmids pNF-kB-Luc and phRL-TK for 4 h, cells were infected with HSV-2 or us1 del HSV-2 for 20 h, followed by stimulation with or without TNF-a (20 ng/ml) for 6 h. Cells were subsequently harvested and lysed to measure Firefly and Renilla luciferase activities using a Dual Luciferase Reporter (DLR) Assay System (Promega, E1980) according to the manufacturer's instructions.

Western blot
The proteins from transfected or infected cells were prepared using Lysis Buffer supplemented with protease inhibitor cocktail (Roche, 11697498001). Prepared cell lysates or immunoprecipitates were subjected to 10% SDS-PAGE and transferred to 0.45 µm polyvinylidene difluoride (PVDF) membranes (Millipore, IPVH00010 PORE). The membrane was blocked using 5% nonfat milk in TBST (20 mM Tris-HCl buffer [pH 7.4] containing 37 mM NaCl and 0.1% Tween 20) at 4°C for 1 h, followed by incubation with a primary Ab overnight at 4°C. After three washes with TBST, the membrane was probed with a HRPconjugated secondary Ab (Proteintech, SA00001-1 or SA00001-2) at room temperature for 1 h, and then washed five times with TBST. The bands were visualized by exposure to ChemiDoc MP Imaging System after the addition of chemiluminescent substrate.

Co-immunoprecipitation assay
HeLa cells seeded in 6-well plates were transfected with Flagtagged ICP22-expressing plasmid or empty vector. At 24 h posttransfection, cells were mock-treated or treated with TNF-a (20 ng/ mL) for 6 h. Cells were subsequently harvested and lysed on ice for 30 min using 200 mL lysis buffer (50 mM Tris [pH 8.0], 150 mM NaCl, 1% NP40) containing protease inhibitor cocktail (Roche, 11697498001). 3 mg mouse anti-Flag Ab or mouse normal IgG (BOSTER, BA1051) was added to fresh Dynabeads protein G (Invitrogen, 10003D), and mixed with cell lysate, respectively. In some cases, 3 mg rabbit anti-p65 Ab or rabbit normal IgG (Beyotime, A7016) was mixed with fresh Dynabeads protein G, and the mixture was then added to cell lysates. After incubation with rotation overnight at 4°C, the Ag-Ab-dynabead complexes were washed three times with PBST, and then the target antigens (Ags) were subjected to western blot analysis after elution followed by boiling.
The kinetics of binding was performed on a Forte-Bio Octet Red System. After Protein A Biosensors (Fortebio, 18-5010, USA) were soaked in 1× PBS, 5 mg/mL anti-p65 Ab was diluted and captured by the Biosensors. The Abs-captured Biosensors were used to bind p65, and then immersed in different concentration of ICP22 (62.5, 125, 250, 500 or 1000 nM) for association and disassociation. The response in nm shift was recorded as a function of time.

Immunofluorescence analysis
HeLa cells seeded in 35-mm glass-bottom dishes were transfected with Flag-tagged ICP22-expressing plasmid. At 24 h post-transfection, cells were treated with or without 20 ng/mL TNF-a for 6 h, and then fixed with 4% paraformaldehyde at room temperature for 10 min. After permeabilized with 0.2% Triton X-100 at room temperature for 10 min, cells were blocked in PBS containing 3% BSA at 4°C overnight. Thereafter, cells were incubated with the rabbit anti-human p65 polyclonal Ab (pAb) and the mouse anti-Flag mAb for 1 h at 37°C. After three washes with PBS, cells were then incubated with Alexa Fluor 488-labeled Goat anti-Mouse IgG (H+L) (Invitrogen, A-10667) and Alexa Fluor 647-labeled Goat anti-Rabbit IgG (H+L) (Invitrogen, A27018) for 1 h at 37°C. Cells were subsequently washed and incubated with DAPI for 10 min at 37°C. After washes, cells were observed under a fluorescence microscope (Nikon A1R/MP).

Statistical analysis
All experiments were repeated at least three times and the data were presented as mean ± SD unless otherwise specified. Data analyses were performed with GraphPad Prism 7.0 software (GraphPad). Comparison between two groups was analyzed by Student t-test, whereas comparisons among more than two groups were analyzed by one-way ANOVA with the Tukey's test. P < 0.05 was considered statistically significant.

Productive HSV-2 infection suppresses TNF-a-induced NF-kB activation
It is known that HSV-2 can evade the host innate immunity to establish lifelong infection. Considering the critical role of NF-kB in the innate immunity, we examined the effect of HSV-2 infection on NF-kB activation. Given that human genital epithelial cells are the main targets of HSV-2 primary infection, we used human cervical epithelial cell line HeLa for the initial experiment. HeLa cells were co-transfected with the reporter plasmids pNF-kB-Luc and phRL-TK for 4 h, followed by infection with HSV-2 or UV-inactivated HSV-2 at an MOI of 1, 0.6, 0.3, or 0.1 for 20 h. After 6 h stimulation with TNF-a, the activities of luciferase were detected. As shown in Figure 1A, HSV-2 significantly inhibited the activation of NF-kBresponsive promoter, whereas UV-inactivated HSV-2 did not have such capability. We further confirmed that the viral proteins were barely detectable by WB after HSV-2 was inactivated by UV ( Figure 1B), indicating that productive infection is necessary for HSV-2-mediated inhibition of NF-kB activation. Given that the chemokine CXCL10 could be induced via NF-kB activation (38, 39), we detected whether HSV-2 infection affects CXCL10 mRNA production. As showed in Figure 1C, HSV-2 infection indeed inhibited NF-kB activation-induced CXCL10 mRNA production, further suggesting the inhibitory effect of HSV-2 on NF-kB activation. These results indicate that HSV-2 infection suppresses TNF-ainduced NF-kB activation and that productive HSV-2 infection is necessary for such suppression.

HSV-2 ICP22 inhibits TNF-a-induced NF-kB activation
Our previous studies show that the HSV-2 ICP22 not only suppresses IFN-b production by blocking the association of IRF-3 with IFN-b promoter (22), but also inhibits the production of ISGs by directly degrading ISGF3 (23). Considering the key role of ICP22 in HSV-2-mediated immune evasion, we next assessed the involvement of HSV-2 ICP22 in interfering with NF-kB signaling pathway. HEK 293T cells were co-transfected with the reporter plasmids pNF-kB-Luc and phRL-TK together with ICP22-expressing plasmid. At 24 h post-transfection, cells were stimulated with TNF-a for 6 h. As shown in Figure 2A, HSV-2 ICP22 significantly inhibited the activation of NF-kBresponsive promoter. Moreover, HSV-2 ICP22 also suppressed CXCL10 mRNA production ( Figure 2B). To confirm the role of ICP22 in the inhibition of NF-kB activation in the context of virus infection, HeLa cells or primary human cervical fibroblasts were transfected with the reporter plasmids pNF-kB-Luc and phRL-TK for 4 h, followed by infection with HSV-2 or us1 del HSV-2, which was constructed as described previously (22), for 20 h, and a stimulation with TNF-a for another 6 h. The results showed that ICP22 knockout significantly impaired the capability of HSV-2 in inhibiting NF-kB activation ( F i gu r e s 2 C , D ) and CXCL10 mRNA production ( Figures 2E, F) in both HeLa cells and primary human cervical fibroblasts. To address whether the inhibitory effect of HSV-2 ICP22 on NF-kB activation was virus specific, we assessed the effects of the ICP22s from several alpha-herpesviruses on NF-kB activation. The results showed that the ICP22s of alphaherpesviruses HSV-1, PRV and VZV all significantly inhibited NF-kB activation, whereas HSV-2 UL46 had no such effect ( Figure 2G), suggesting the significance of ICP22 in herpesvirus immune evasion. These results collectively indicate that HSV-2 ICP22 inhibits TNF-a-induced NF-kB activation.

HSV-2 ICP22 inhibits NF-kB activation by acting on the downstream of p65
IkB protein inhibits the activation of NF-kB by trapping NF-kB in the cytoplasm (40). Under foreigner stimuli, IKK complex phosphorylates IkB, resulting in the phosphorylation, and subsequent degradation of IkB. Once IkB is detached from NF-kB, NF-kB is phosphorylated and activated (41). To understand how HSV-2 ICP22 antagonizes NF-kB activation, HEK 293T cells were co-transfected with the reporter plasmids pNF-kB-Luc and phRL-TK, and the plasmid expressing TRAF2, IKK a, IKK b, IKK g or p65, together with ICP22-expressing plasmid or empty vector for 30 h. As showed in Figures 3A-E, overexpression of TRAF2, IKK a, IKK b, IKK g or p65 resulted in the activation of NF-kB-responsive promoter, whereas HSV-2 ICP22 significantly inhibited TRAF2, IKK a, IKK b, IKK g or HSV-2 ICP22 inhibits NF-kB activation by acting on the downstream of p65. (A-E). HSV-2 ICP22 inhibits TRAF2, IKK a, IKK b, IKK g and p65induced NF-kB activation. HEK 293T cells were seeded in 24 well plates overnight and co-transfected with the reporter plasmids pNF-kB-Luc and phRL-TK, and plasmid expressing TRAF2, IKK a, IKK b, IKK g or p65, together with empty vector or ICP22-expressing plasmid. At 30 h posttransfection, the reporter activities were determined by DLR assay. (F). HSV-2 ICP22 has no effect on the expression of TRAF2, IKK a, IKK b, IKK g and p65 or degradation of IkB a. HEK 293T cells were seeded in 6 well plates overnight and transfected with plasmid expressing ICP22 or empty vector. At 24 h post-transfection, cells were stimulated with or without TNF-a (20 ng/ml) for 6 h. The expressions of TRAF2, IKK a, IKK b, IKK g, IkB a, p65 and phospho-p65 were detected by western blot. For graphs, data shown are mean ± SD of three independent experiments with each condition performed in triplicate. For images, one representative experiment out of three is shown. *p<0.05, **p<0.01, ***p<0.001, ns, not significantly.
p65-induced NF-kB activation, without affecting the expression of TRAF2, IKK a, IKK b, IKK g, and p65 ( Figure 3F). As showed in Figure 3F, in the condition of TNF-a stimulation, the inhibitory factor IkB a was degraded in both vector-and ICP22-transfected cells indicating that HSV-2 ICP22 did not affect the degradation of IkB a, while the total level of phospho-p65 was decreased in ICP22-tranfected cells. These results collectively indicate that HSV-2 ICP22 inhibits the activation of NF-kB by acting on the downstream of p65.

HSV-2 ICP22 inhibits the phosphorylation and nuclear translocation of p65
Given that HSV-2 ICP22 inhibits the activation of NF-kB by acting on the downstream of p65, we next investigated the influence of ICP22 on p65 phosphorylation and nuclear translocation. HeLa cells were transfected with ICP22expressing plasmid, followed by stimulation with TNF-a for 6 h. Cytoplasmic and nuclear proteins were subsequently isolated and detected by western blot to determine the distribution of p65. As showed in Figure 4A (Lane 1-4), the phosphorylation of p65 in the cytoplasm was inhibited by ICP22, although the expression of total p65 was not affected by ICP22 ( Figure 3F). Meanwhile the phosphorylated p65 in the nucleus also decreased in ICP22-transfected cells ( Figure  HSV-2 ICP22 inhibits the phosphorylation and nuclear translocation of p65. (A, B). HSV-2 ICP22 inhibits the phosphorylation of p65. HeLa cells seeded in the 6 well plates were transfected with empty vector or ICP22-expressing plasmid. At 4 h post-transfection, cells were mock infected or infected with HSV-2 or us1 del HSV-2 at an MOI of 1. At 20 h post-infection, cells were stimulated with or without TNF-a (20 ng/ml) for 6 h. The phosphorylated p65 in the cytoplasm and nucleus were detected by western blot. b-actin and PCNA were used as loading controls for cytoplasmic and nuclear proteins, respectively. (C, D). HSV-2 ICP22 significantly inhibits the nuclear translocation of p65. HeLa cells were transfected with ICP22-expressing plasmid or empty vector. At 24 h post-transfection, cells were stimulated with or without TNF-a (20 ng/ml) for 6 h. Cells were stained using the mouse anti-Flag and the rabbit anti-p65 Ab. Alexa Fluor 488-conjugated goat anti-mouse (green) and Alexa Fluor 647-conjugated goat anti-rabbit (red) were used as secondary antibodies. Cell nuclei (blue) were stained with DAPI. The images were obtained by fluorescence microscopy using a 60× objective. The percentage of p65-positive nuclei was quantified in a number of fields (D). The scale bar indicates 20 mm. For graphs, data shown are mean ± SD of three independent experiments with each condition performed in triplicate. For images, one representative experiment out of three is shown. ***p < 0.001. phosphorylation and nuclear translocation of p65 (Lane 5-6), whereas ICP22 knockout obviously impaired the inhibitory effect of HSV-2 on p65 phosphorylation and nuclear translocation (Lane 9-10). Immunofluorescence assay further showed that p65 translocated from the cytoplasm into the nucleus in the majority of cells after stimulation with TNF-a, and such translocation was significantly blocked in ICP22transfected cells (Figures 4C, D). These results together indicate that HSV-2 ICP22 inhibits the phosphorylation and nuclear translocation of p65.

HSV-2 ICP22 directly interacts with p65
To address how ICP22 suppresses the phosphorylation of p65, co-immunoprecipitation assay was performed to assess the interaction of HSV-2 ICP22 with p65. As showed in Figures 5A, B, HSV-2 ICP22 was found to interact with endogenous p65 in both pull-down experiments using the anti-Flag or anti-p65 antibody. To further confirm the results, recombinant HSV-2 ICP22 and human p65 were used to measure the binding kinetics of ICP22 with p65, showing that HSV-2 ICP22 indeed directly interacts with p65 ( Figure 5C). p65 contains a conserved Rel homology domain (RHD) at the N terminus, which is responsible for nuclear localization, dimerization and DNA binding (24). To map the functional region of p65 interacting with ICP22, we constructed three truncation mutants D1, D2 and D3 (36). As showed in Figure 5D, D1 (19-306aa) retains complete RHD of p65, while D2 (19-300aa) is deficient in the nuclear localization signal (NLS) domain of RHD. D3 (19-187aa) only retains the DNA binding domain of RHD. Subsequently, His-tagged full-length p65 or its truncated mutants (named D1, D2, and D3) were used to identify the functional domain of p65 interacting with HSV-2 ICP22. Co-immunoprecipitation assay showed that full-length p65 and its three truncation mutants D1, D2 and D3 all interacted with ICP22 ( Figure 5E). The interaction of ICP22 with p65 seemed to be weakened when the dimerization domain of p65 was deleted, indicating that the dimerization domain of p65 likely plays a more important role in the interaction. These results together inform that HSV-2 ICP22 directly interacts with p65, resulting in the blockade of p65 phosphorylation and nuclear translocation.

Discussion
HSV-2 is one of the most common sexually transmitted viruses worldwide, causing neonatal herpes and genital ulcer disease (42). HSV-2 and HSV-1 are closely related but exhibit substantial differences in latency and reactivation patterns (10)(11)(12)(13)(14)43). NF-kB is a key regulator of a broad range of cellular responses, involved in the induction of inflammation (24,(44)(45)(46). Although a number of studies report that HSV-1 has evolved multiple countermeasures to subvert the activation of NF-kB signaling pathway (10-12, 15, 16, 20, 47), our current understanding of HSV-2 immune evasion against the activation of NF-kB is limited.
In this study, we found that HSV-2 infection inhibited TNFa-induced activation of NF-kB-responsive promoter, whereas UV-inactivated HSV-2 did not have such inhibition, indicating that productive HSV-2 infection is necessary for the inhibition of NF-kB activation. Subsequent studies indicated that ICP22 has a significant inhibitory effect on the activation of NF-kBresponsive promoter, which was further confirmed in the context of viral infection using ICP22 deficient HSV-2. Moreover, we found that the ICP22s from several alphaherpesviruses including HSV-1, PRV and VZV all inhibited NF-kB activation, which share 62%, 33% and 31% identity, respectively, with the amino acid sequence of HSV-2 ICP22, highlighting the significance of ICP22 in herpesvirus immune evasion. Given that HSV-2 ICP22 can also suppress the production of type I IFN and ISGs (22,23), our findings collectively informed that ICP22 is a key viral element counteracting not only type I IFN production and signaling but also NF-kB activation.
It is known that NF-kB activation is an attractive target for common human viral pathogens to evade host antiviral responses (34,(48)(49)(50). The activation of NF-kB signaling cascade includes the phosphorylation and nuclear translocation of p65, a major component of NF-kB heterodimer. We found that HSV-2 ICP22 significantly blocked TRAF2, IKK a, IKK b, IKK g and p65-induced NF-kB activation, but did not affect the expression of TRAF2, IKK a, IKK b, IKK g and p65 or the degradation of IkB a, indicating that HSV-2 ICP22 likely affects p65 activation. We previously demonstrated that HSV-2 ICP22 functions as a novel E3 ubiquitin protein ligase to degrade ISGF3, resulting in the inhibition of type I IFN signaling (23). However, in the current study, HSV-2 ICP22 appears to inhibit NF-kB activation independent of its E3 ubiquitin protein ligase activity, and instead, it suppresses the phosphorylation and nuclear translocation of p65, leading to the inhibition of NF-kB activation.
Mechanistically, we found that HSV-2 ICP22 directly interacts with endogenous p65. In accordance with previous findings, several other viral proteins of HSV-1 have also been shown to interact with p65 (10)(11)(12)(13)(14).The NF-kB family shares the RHD at the N-terminus, which contains 300 amino acids and has three functions: sequence specific DNA-binding, dimerization and inhibitory protein binding (24,41). By assessing three truncated p65, we found that full-length p65 and its three truncation mutants D1, D2 and D3 all interacted with ICP22. The p65 truncation mutant D3 (19-187aa) only HSV-2 ICP22 directly interacts with p65. (A) HSV-2 ICP22 interacts with endogenous p65. (B) Endogenous p65 interacts with HSV-2 ICP22. HeLa cells seeded in the 6 well plates were transfected with empty vector or ICP22-expressing plasmid. At 24 h post-transfection, cells were mock-treated or treated with TNF-a (20 ng/ml) for 6 h. Cell lysates were then subjected to co-immunoprecipitation assays using the anti-Flag (A) or anti-p65 Ab (B). The mouse (A) or rabbit (B) non-specific antibody was used as negative control. ICP22 and p65 were detected by western blot using the anti-Flag or anti-p65 Ab, respectively. (C) HSV-2 ICP22 directly interacts with p65. The kinetics of binding was performed on a Forte-Bio Octet Red System. 5 mg/mL rabbit anti-p65 Ab was coupled to Protein A biosensors. 25 mg/mL recombinant p65 was bound to Biosenors and immersed in different concentration of ICP22 (62.5, 125, 250, 500 or 1000 nM) for association and disassociation. The response in nm shift was recorded as a function of time. KD (M) = 2.18E-07. (D) Schematic representation of p65 truncations. (E) HSV-2 ICP22 interacts with the three truncation mutants D1, D2 and D3 of p65. Empty vector or ICP22-expressing plasmid and plasmid expressing full-length or truncated p65 were co-transfected into HEK 293T cells. At 24 h post-transfection, cells were mock-treated or treated with TNF-a (20 ng/ml) for 6 h. Cell lysates were then subjected to co-immunoprecipitation assays using the anti-Flag mAb. ICP22, truncated p65 were detected by western blot using the anti-Flag or anti-p65 Ab, respectively. Asterisk indicated the locations of proteins. One representative experiment out of three is shown. retains the DNA binding domain of RHD. In agreement, our previous study showed that HSV-2 ICP22 also interacts with the DNA binding domain of IRF-3 (22) to suppress IFN-b p r o d u c t i o n , w h i c h i n d i r e c t l y s u p p o r t s o u r c oimmunoprecipitation results in this study. An interaction of ICP22 with the DNA binding domain of p65 likely blocks its association with phosphorylase, resulting in the suppression of p65 phosphorylation and nuclear translocation. Given that HSV-2 ICP22 interacts with the DNA binding domain of p65, it likely facilitates viral immune evasion by interfering with the binding of NF-kB with the promoters of regulatory genes in the nucleus.
It is known that HSV-1 ICP22 is an immediate-early protein and a multifunctional viral regulator. HSV-1 ICP22 not only interacts with RNA polymerase II (51-57) and P-TEFb (58) to regulate viral replication (59), but is also involved in posttranslational modification as viral protein kinases (60)(61)(62)(63). To date, little is known about the functions of HSV-2 ICP22. We previously revealed that HSV-2 ICP22 functions as a novel E3 ubiquitin protein ligase to degrade ISGF3 (23) and a key viral element contributing to HSV-2 immune evasion (22). Although beyond the scope of this study, future work is warranted to explore the structural characteristics of HSV-2 ICP22 for its multiple functions.
In conclusion, we demonstrated that HSV-2 ICP22 blocks TNF-a-induced activation of NF-kB by directly interacting with p65. The findings highlight the significance of ICP22 in inhibiting NF-kB activation. We proposed a model as described in Figure 6. When the host recognizes foreign PAMPs, the innate immune system is activated by secreting cytokines such as TNF-a. The secreted TNF-a engages TNF receptor (TNFR), resulting in the activation of the IKK complex. Subsequently, the inhibitory protein of NF-kB, IkB a, is phosphorylated, degraded and detached from NF-kB. NF-kB dimers are then released and phosphorylated, and subsequently translocate to the nucleus to activate the expression of immunomodulatory genes. In the case of HSV-2 infection, the viral immediate early protein ICP22 directly interacts with p65 to suppress p65 phosphorylation and nuclear translocation, leading to the blockade of NF-kB activation, which would facilitate viral immune escape. A schematic model of the mechanism by which HSV-2 ICP22 blocks TNF-a-induced NF-kB activation. The host recognizes foreign PAMPs and activates the innate immune system to secret cytokines such as TNF-a. TNF-a subsequently binds TNF receptor (TNFR), resulting in the activation of the IKK complex. The inhibitory protein of NF-kB, IkB a, is then phosphorylated, degraded and detached from NF-kB. NF-kB dimers are then released and phosphorylated, and subsequently translocate to the nucleus to activate the expression of immunomodulatory genes. In the case of HSV-2 infection, the viral immediate early protein ICP22 directly interacts with p65 to block p65 phosphorylation and nuclear translocation, leading to an inhibition of NF-kB activation.

Data availability statement
The raw data supporting the conclusions of this article will be made available by the authors, without undue reservation.

Author contributions
HH, MZ and QH conceived the study. HH and MZ conducted most experiments. CL constructed the truncation mutants of p65. MF, BZ and YL provided help in western blot experiments. HH, MZ and QH analyzed the data. HH, MZ and QH wrote the manuscript. All authors contributed to the article and approved the submitted version.

Funding
This work was supported by National Natural Science Foundation of China (82171736, 81772192 and 31970172), and the National Mega-Projects against Infectious Diseases (2018ZX10301406-002).


s2fieldsofstudy: Biology Medicine

text:
Antibacterial Dental Resin Composites: A Narrative Review

Lack of antibacterial properties in resin-based composites (RBCs) is one of the flaws that cause the failure of filling clinically. Several agents have been incorporated to endow RBCs with antibacterial properties. In this review, we summarize the recent antibacterial agents between 2015 and 2020 using keywords of antibacterial or antimicrobial dental resin composites by PubMed databases. The most effective strategies are concerned with polymerizable monomers (50%), followed by filler particles (39%) and leachable agents (11%). A recent modification of the antibacterial agent is either by combining two agents from the same category or mixing agents from different categories in one. More than two methods were used in one study to assess antibacterial efficacy. The most common method was biofilm colony-forming units (CFUs) counting method (40%), followed by live/dead bacteria staining assay of biofilms (25%), metabolic activity assay of biofilms using MTT assay (16%), lactic acid production assay of biofilms (8%), agar diffusion test (8%), and other methods (3%) such as minimal inhibitory concentration (MIC) and minimal bactericidal concentration (MBC).

Introduction
The progressive developments in RBCs over 50 years endow composites with adequate mechanical and high aesthetic properties, making them the preferred restoration materials used clinically for anterior or posterior teeth [1]. However, clinical trials have shown that the risk of failure of RBCs is twice as high as that of silver amalgam, mainly due to the marginal microleakage caused by polymerization shrinkage during the curing process, which makes it easier for bacteria to invade and lead to secondary caries [2] [3]. In addition, the cured composites have higher biofilm and plaque aggregation rates than silver amalgam and glass ionomer [4] [5] [6].
In recent years, massive modifications have been done to composite monomer systems to reduce polymerization shrinkage stress by over 70% [4] [5] [7]. On the other hand, numerous studies have tended to endow RBCs with antibacterial properties by incorporating antibacterial additives to reduce biofilm formation and prevent caries. Several studies have reported that various antibacterial additives were added into resin-based material [6], resin composite [8], glass ionomer cement [9], and dentine bonding system [10], and their antibacterial activities were subsequently evaluated. These antibacterial additives were mainly classified into releasing and non-releasing additives, incorporated into either resin matrix or filler particles.
This review summarizes the recent antibacterial agents added to RBCs, which included articles from January 2015 to May 2020, using antibacterial or antimicrobial dental resin composites as keywords by PubMed databases. The dental resin composites used for restoration purposes were included, whereas those used for orthodontic, endodontic, or sealing purposes were excluded. Adhesives and resin-modified glass ionomer cement were also excluded.

Inclusion Criteria
1) All studies which added new antibacterial additives to restorative resin composites; 2) Scientific papers were published in English from January 2015 to May 2020, where full text was available.

Exclusion Criteria
1) Studies about the antibacterial resin composite used for orthodontic, intracanal post cementation, core build-up restoration, and sealer; 2) Studies that did not include Streptococcus mutans (S. mutans, which are the primary bacteria responsible for dental caries formation) in the antibacterial test; 3) Studies whose antibacterial analysis was not clear; 4) Studies not in the range from January 2015 to May 2020.

Search Strategy
A search using keywords of antibacterial or antimicrobial dental resin compo- Three main antibacterial strategies depend upon the antibacterial mechanisms of the antibacterial constituents incorporated into the RBCs. Leachable agents can be released into the local environment around restorations under oral conditions. In contrast, non-leachable agents or polymerizable monomers can be immobilized in the dental resin matrix, while bacterial filler particles added to fillers or resin matrix can release small ions to create antibacterial effects ( Table  1). The summery of antibacterial modifications of RBCs is shown in Figure 2.   -Surface pre-reacted glass-ionomer (S-PRG) -SPRG [37] Each study involved in this review included at least one of the above three strategies: leachable agents, polymerizable monomers, and filler particles. When the antibacterial additives used in a study were in the same category, the count for that category increased by one. When a study included antibacterial additives referring to various categories, the counts for different categories increased by one. Therefore, the total number of different additives from these three strategies found among the 32 studies was counted as 100%, as seen in Figure 3.  Leachable agents are soluble antibacterial agents incorporated into the resin matrix and released under the oral environment. The foremost commonly utilized leachable antibacterial agents are benzalkonium chloride (BAC) and chlorhexidine [10]. The main disadvantage of these materials is a short-lasting effect (burst effect), resulting in large amounts of leachate in the surrounding environment and showing an antibacterial effect within a few days, followed by a dramatic decrease in the drug concentration.  Triclosan (TCN) is a common leachable antibacterial agent used as a component of dental toothpaste, mouthwashes, and RBCs [38] [39]. Likewise, due to the short-term burst effect, TCN is usually incorporated in particular "vehicles" known as nanotubes [40]. A recent study accomplished by Cunha et al. [11] used biocompatible nanomaterial halloysite nanotubes (HNT), which were previously used as a reinforcing nanofiller and reservoir for controlled discharge of an assortment of therapeutic drugs [41] [42]. TCN was successfully encapsulated into halloysite nanotubes (HNT/TCN) which were incorporated at 8% w/w to prepare the micro-hybrid dental resin composite, showing enhancements of the mechanical properties and no significant difference in antibacterial properties over 5 days.  Chlorhexidine (CHX) is another common leachable antibacterial agent used in limited concentrations due to cytotoxicity toward human fibroblasts [43]. Therefore, it is incorporated into mouthwashes as well as glass ionomer cement (GICs), resin composite, and resin-modified glass ionomer cement (RMGICs) materials in low concentration, exhibiting antibacterial activity with short-term CHX release [44] [45] [46]. Montmorillonite (MMT) is a common ingredient in pharmaceutical products, which is used as an excipient and active substance due to its good adsorptive ability, drug-loading, and cationic interchange capacities [47] [48]. Previous studies have prepared CHX-copper (II)/MMT nanocomposites and chitosan/MMT composite films containing CHX to get long and effective antibacterial properties with low cytotoxicity [49]. In the same way, a recent study was done by Boaro et al. [12] developed a composite modified by CHX-loaded MMT. The composite showed inhibition of bacterial adhesion and constant CHX release without a change in the mechanical properties or cytotoxic effect. To improve the antibacterial and remineralization properties, our group has recently synthesized CHX with amorphous calcium phosphate in core-shell structure (CHX/ACP), and then we merged CHX/ACP nanoparticles into the experimental composite resin [13]. The modified composite could continuously release CHX with calcium and phosphate (Ca and P) ions and improve antibacterial and remineralization properties.  Benzalkonium chloride (BC) is a common leachable antibacterial agent incorporated into dental materials [50]. Wang J et al. [14] converted BC to deep eutectic solvent (DES) by blending BC with acrylic acid (AA) to serve as the donor of hydrogen bond, which is essential in DES formulation. Then this DES was merged into the resin composite to produce antibacterial activity. The results from the DES-modified composite resin showed better mechanical properties and antibacterial inhibition compared with the BC-modified composite.  Chitosan is a natural polysaccharide polymer with a wide spectrum of antimicrobial activity [51]. Chitosan has been added to adhesives, glass-ionomer cement, and sealants to enhance its mechanical and antimicrobial properties [52] [53] [54]. Methacrylate chitosan (CH-MA) was prepared and incorporated into the adhesive that showed comparable bond strengths to the control system [55]. Stenhagen et al. [15] prepared dental composite and adhesive containing CH-MA and confirmed that the antibacterial effect was correlated with CH-MA amounts. Different synthesis methods are applied to chitosan powder to modify its properties, creating nanofiber and microspheres of chitosan [54] [56] [57]. Chitosan microspheres could encapsulate other bioactive compounds. For example, Tanaka et al. [16] synthesized novel chitosan microspheres encapsulate dibasic calcium phosphate anhydrous (DCPA) using the electrospray technique, which was incorporated into an experimental composite. The composites containing 0.5 wt% chitosan/DCPA showed an effective antimicrobial property compared to the control group. Polymerizable monomers Polymerizable antibacterial monomers are immobilized into a resin matrix based on copolymerization among the resin monomers to overcome the short-lasting release of the antibacterial agents. Their antibacterial effects occur through the contact of bacteria with the composite surface. Cationic groups like quaternary ammonium, pyridinium, and phosphonium are commonly found in the functional groups of polymerizable antibacterial monomers.
Polymerizable monomers used alone • A series of quaternary ammonium compounds (QACs) monomers with one or multiple methacrylate groups was considered the most effective immobilized antimicrobial monomer [58] [59]. Zhang et al. [60] synthesized quaternary ammonium methacrylates (QAMs) with different chain lengths (CL) varying from 3 to 18 and merged them into the amorphous calcium phosphate (NACP) composite. This study demonstrated that as the CL increased, the antibacterial effects increased, with the strongest result achieved with a CL of 16. In contrast, the antibacterial property was reduced as the CL was increased to 18.  Quaternary ammonium polyethyleneimine (QPEI) nanoparticles are a potential antimicrobial polymer incorporated into RBCs, exhibiting a powerful antibacterial capability [61]. Pietrokovski et al. [26] proved that the RBC containing QPEI nanoparticles had considerable antibacterial effects against Streptococcus mutans and Actinomyces viscosus.  A composite with antibacterial and remineralization capabilities was synthesized by integrating a strong antibacterial compound dimethyl amino hexadecyl methacrylate (DMAHDM, a kind of QACs) with NACP [17]. The best result was achieved when a 3% mass fraction of DMAHDM was integrated into the NACP resin composite without adversely affecting the mechanical properties. Similarly, adding DMAHDM into the rechargeable NACP composite has been reported by Al-Dulaijan et al. [18], the rechargeable NACP-DMAHDM composite showed Ca and P ions release with persisting remineralization and a potent antibacterial effect.
 Zhang et al. [19] have integrated 2-methacryloyloxyethyl phosphorylcholine (MPC, a kind of QACs) with DMAHDM in an attempt to synthesize an anti-biofilm and protein-repellent dental composite. After water aging for six months, resin composite modified by 3% MPC combined with 1.5% DMAHDM exhibited higher resistance to bacterial adhesion than the control group. The protein-repellent and antibacterial effects were durable and showed no loss in water aging from 1 to 180 days, with mechanical properties matching a commercial composite. Similarly, antibacterial DMAHDM monomer and MPC were incorporated into the rechargeable NACP composite [20]. Compared to a commercial control group, the composite with 3% MPC and 3% DMAHDM impaired bacterial growth and decreased the CFU count of biofilm by three orders of magnitude.
 A recent study reported by Lee et al. [21] added MPC to S-PRG filler to modify a resin-based composite to get the benefits of both materials (antibacterial ability, anti-biofouling function, acid resistance, and prevention of demineralization). The authors reported that as the percentage of MPC increased, the number of ions released from the S-PRG filler increased. So, the RBC containing S-PRG filler and 5% MPC had a significant anti-biofilm formation effect and improved release of ions and acid neutralization properties.  To overcome the drawback of mono-methacrylate QAMs monomers, QAMs monomers with dimethacrylate were prepared and synthesized, such as dimethyl hexadecyl methacryloxyethyl ammonium iodide (DHMAI) and ionic dimethacrylates (IDMAs) [62]. Cherchali et al. [22] have assessed the antibacterial activity of an experimental dental composite, including DHMAI. DHMAI was added to MPC to test both the new composite's antibacterial activity and mechanical properties. The above study showed that the composite incorporated with 7.5% DHMAI had a strong antibacterial effect with a reduction in CFU (by 98%), metabolic activity (by 50%), and acceptable me-chanical properties. However, the joint addition of both DHMAI and MPC monomers to composite didn't significantly improve antibacterial activity, but resulted in worse mechanical properties.  IDMAs have been applied in dentistry, with an antibacterial effect equivalent to methacryloyloxydodecyl pyrimidinium bromide (MDPB) [58]. Bienek et al. [23] synthesized purity-enhanced IDMA1 and IDMA2, and then assessed the biological, physicochemical, mechanical, and antibacterial properties of the IDMAs-modified resin composites. The authors concluded that IDMAs showed minimal or no cellular toxicity, and incorporation of IDMAs improved the degree of vinyl conversion (DVC) of the resins without affecting their wettability.  A series of urethane dimethacrylate quaternary ammonium monomers (UDM-QAs) have been synthesized, such as UDMQA-12, used at 30% to 40% in BisGMA/TEGDMA resin systems with significant antibacterial activity [63] [64]. A recent study reported by Huang et al. [24] prepared a photo-polymerized resin matrix with 30% UDMQA-12, mixed with silanated glass fillers at a mass ratio of 30:70. The new composite showed a significant antibacterial effect against S. mutans, better than commercial composite but still worse than glass ionomer cement (GIC). Moreover, its mechanical properties were similar to commercially available resin composites.  A new dental resin system without Bis-GMA used Tricyclodecane dimethanol diacrylate (SR833s) and diurethane dimethacrylate (UDMA) monomers as a base resin, then N, N-bis [2-(3-(methacryl oyloxy) propanamide)-ethyl]-N-methylhexadecyl ammonium bromide (IMQ-16) was incorporated to obtain an antibacterial dental resin [65]. UDMA/SR833s/IMQ-16 resin system showed higher physicochemical properties compared to Bis-GMA/TEGDMA formulation. Incorporating IMQ-16 into this system at 17% or 20% produced a considerable antibacterial resin system.
Polymerizable monomers in combination with leachable agents  To overcome the drawback of short-term release of antibacterial agents, they were immobilized with cationic polymers to create dental resins with a dual antibacterial mode that possesses both contacts and release antibacterial capabilities. The first study in this field was reported by Cao et al. [66] has developed photocurable core-shell silver bromide (AgBr)/cationic quaternary ammonium methacrylates (BHPVP) nanocomposites, releasing the active Ag + ions for a long term and possess the high antibacterial potency due to cationic polymers and Ag + ions.  [67] have synthesized and incorporated Triclosan methacrylate monomer (TM) into RBCs. The modified composite showed low biofilm accumulation and comparable mechanical properties without a significant difference from the control group. Antibacterial filler particles Antibacterial filler particles are usually incorporated into the RBCs, mainly metal, metal oxide, and bioactive glass filler. They are water-insoluble, but a small number of ions can be released into the surrounding environment. Silver is the most common antibacterial filler particle used for dental material [68].
Metal filler  Silver nanoparticles (AgNPs), a kind of metal, are a broad-spectrum antibacterial agent incorporated into the RBCs to produce a high antibacterial activity by releasing Ag + ions [69] [70]. The problem faced by adding nanoparticles is the aggregation and incomplete dispersion in the polymeric matrix, which affects various properties of the composites [71]. Some studies have overcome this drawback [72] [73]. Surface modification on nano-scale fillers with mussel-inspired dopamine (DA) has recently been highlighted in the preparation of organic-inorganic composites. The catechol group in DA can reduce silver ions to AgNPs and firmly bond the nanoparticles [74] [75] [76].
Ai et al. [27] synthesized hydroxyapatite (HA) nanowires using the hydrothermal technique, followed by surface modification via mussel-inspired dopamine (DA) to prepare polydopamine (PDA)-coated HA (HA-PDA) nanowires. The HA-PDA nanowires were further loaded with AgNPs to prepare the target product HA-PDA-Ag nanowires, which were finally incorporated into the resin composite. The authors reported that the composite reinforced by HA-PDA-Ag nanowires showed long-lasting antibacterial efficacy and no cytotoxicity.
 Barot et al. [28] used halloysite nanotubes (HNT) to load AgNPs. The HNT/Ag nanotubes were incorporated into BisGMA/TEGDMA-based dental resin composite, showing a high antibacterial activity on S. mutans and improved mechanical properties when 1 -5 wt% of HNT/Ag nanotubes were added. Another study [29] mixed silver sulfadiazine, a kind of metal salt, with commercial barium borosilicate glass powders to obtain antibacterial glass powders, which were added to BisGMA-based dental resins. The target composite showed a potent antimicrobial effect persisting for more than eight weeks and no changes in mechanical properties. Metal oxide filler  Another way to achieve antibacterial activity is by adding metal oxides [77].
One metal oxide is zinc oxide (ZnO) nanoparticles added to resin-based restorative materials, resulting in high anti-biofilm effects [78] [79]. The smaller ZnO particles in size have higher antibacterial capacity than the larger ones. The rod-shaped or wire-shaped ZnO particles have better antibacterial results than spherical ones [80] [81]. Many other shapes of ZnO have been tested, and it is concluded that the antibacterial effect of ZnO is shape-dependent  [83]. According to this conclusion, Dias et al. [31] synthesized ZnO particles with 3D microstructures and incorporated them as antimicrobial fillers in resin composites. The resin composite modified by 0.5 wt% of ZnO microrods exhibited a significant decrease in the bacterial accumulation on the composite surface without compromising its mechanical properties. Recently, Wang et al. [32] prepared cellulose nanocrystal/zinc oxide (CNC/ZnO) nanohybrids and incorporated them into dental resin composites. When 2% CNC/ZnO nanohybrids were added, the modified composite showed significant antibacterial properties without statistically different mechanical properties compared to the control composite.  Some efforts have been focused on filler components, morphology, drug-loading, and size to get dental composites with perfect physical-mechanical properties. Porous mesoporous filler has been introduced into the dental composites to increase the resin-filler interfacial bonding, aiming to improve the mechanical performance of the composites [84] [85]. For example, mesoporous SiO 2 has been introduced to enhance the micromechanical properties of resin matrix via the formation of interlocking structures in dental composites [86] [87]. Chen et al. [33] inserted an antibacterial agent into mesoporous filler to form a mesoporous core-shell structure filler (ZnO@m-SiO 2 ) and used it as a functional filler in the dental composite. The composite modified by 70 wt% of ZnO@m-SiO 2 filler demonstrated the best mechanical properties compared to the control composite and a superior antimicrobial activity (Antibacterial ratio > 99.9%.)  Titanium dioxide nanoparticles (TiO 2 NPs), another kind of metal oxide, are often used as an antibacterial agent to modify resin composite. A recent study was done by Dias et al. [34] added pure TiO 2 NPs and Ag-decorated TiO 2 NPs into the resin composite, and then the antibacterial activities of the synthesized resin composites were evaluated. The two modified composites significantly reduced biofilm formation without differences between them.  Many other strategies also demonstrated a high and long-lasting antibacterial efficiency. Like silver-decorated TiO 2 NPs, the synergetic antibacterial effect of other nanohybrid materials was also highlighted, such as ZnO-Ag and ZnO-Au NPs. For example, a study done by Dias et al. [30] modified commercial resin composite with Ag-decorated ZnO nanoparticles. The modified resin composite showed biofilm inhibition on the surface without compromising its compressive strength. Bioactive filler  Bioactive glass (BG) is a biocompatible filler that can release Ca and P ions and possess a remineralizing effect. In addition to the cytocompatibility of BG resin composites, their mechanical properties are similar to commercial composites [88] [89]. Korkut et al. [35] reported that the antibacterial activity of resin composite modified by BG lasted for about 90 minutes. Its compressive and flexural strengths presented a decreased trend and a concentration-dependent effect on BG contents. In addition, several studies have been A. S. Alansy et al. done to develop antibacterial and bioactive restorative materials [90]. However, these materials showed retrograded mechanical properties or color changes, limiting their clinical use [91]. Chatzistavrou et al. [36] synthesized a silver-doped bioactive glass modified resin composite (Ag-BG), which showed a homogeneous dispersion of Ag-BG particles within the resin composite. The enhanced remineralizing properties and the long-lasting biofilm inhibition were correlated to the amount of Ag-BG. There were no significant differences in mechanical properties compared to the control samples.  A surface pre-reacted glass-ionomer (S-PRG) filler has been incorporated into dental materials, making them have the capacity to release multiple ions, such as fluoride, aluminum (Al 3+ ), sodium (Na + ), and strontium (Sr 2− ) ions [92] [93] [94]. Therefore, the modified composites effectively prevented the demineralization of dentin, imparted acid resistance to enamel, and promoted mineralization [95] [96] [97]. Resin composite modified by S-PRG filler also showed less bacterial attachment and plaque accumulation [98] [99].
In 2016, Miki et al. [37] evaluated and demonstrated the ability of resin composites modified by S-PRG filler to impede the growth of S. mutans on the surface.

The Measuring Methods of Antibacterial Efficacy
 The measuring methods for antibacterial efficacy are collected and categorized. Generally, more than one method is used in one study to confirm the results and support the conclusion. The most commonly used test method in this review was the biofilm colony-forming units (CFUs) counting method (40%), followed by live/dead bacteria staining assay of biofilms (25%), metabolic activity assay of biofilms using MTT assay (16%), lactic acid production assay of biofilms (8%), agar diffusion test (8%), and other methods (3%) such as minimal inhibitory concentration (MIC), minimal bactericidal concentration (MBC) (Figure 4).   [101].  The second most common method is the live/dead bacteria staining assay, which indirectly measures bacteria viability. This method differentiates viable and dead bacteria through cell membrane integrity, using the combination of SYTO 9 and propidium iodide dye. Therefore, it's more suitable for antibacterial agents which act on the cell membranes, such as quaternary ammonium compounds [101].
Metabolic activity assay of biofilms has various indicators to measure, such as a snapshot of the bulk metabolic function at a given time, the evaluation of gene expression, and the measurement of metabolic byproducts. About 16% of the studies used the MTT assay to measure the metabolic activity of biofilms, which depends on the enzymatic reduction of MTT from yellow tetrazole to purple formazan. The metabolic activity is not equal to the biofilm cell viability because some biofilm cells remain viable, but their metabolism is inactive. Therefore, this method has been used as a reinforcing and supplementary assessment to other methods [100] [102]. Because of the advantages and disadvantages of each technique, researchers should completely understand the mechanism, limitations, and operation procedures of each method and then select the appropriate methods to avoid misinterpreting the results. There are no universal protocols for all studies to follow, helping to make comparisons among different studies.

Conclusions
This review covered the antibacterial agents incorporated into the resin composite from January 2015 to May 2020, including 32 articles focused on modifying the RBCs using existing antibacterial agents to increase their antibacterial efficacy or prolong their antibacterial period. Generally, most studies were concerned with polymerizable monomers (50%), followed by filler particles (39%) and leachable agents (11%). The problem mostly faced is that the antibacterial effects are in a concentration-dependent manner depending on the contents of antibacterial agents. However, higher contents of antibacterial agents may cause cytotoxicity or interfere with the mechanical properties of the resin composite.
Consequently, most antibacterial agents are added in small amounts and modified to have significant antibacterial effects, minimal cytotoxicity, and no effects on mechanical characteristics of the experimental/commercial resin composite.
The recent antibacterial agents focused combination of leachable agents, polymerizable monomers, and metal oxide filler agents. This combination is of two agents from the same category (DMAHDM + MPC, DHMAI + MPC, Ag-decorated