Biomedical activities of plant extract-based synthesized ZnO nanoparticles.
\\n\\n
Released this past November, the list is based on data collected from the Web of Science and highlights some of the world’s most influential scientific minds by naming the researchers whose publications over the previous decade have included a high number of Highly Cited Papers placing them among the top 1% most-cited.
\\n\\nWe wish to congratulate all of the researchers named and especially our authors on this amazing accomplishment! We are happy and proud to share in their success!
\\n"}]',published:!0,mainMedia:null},components:[{type:"htmlEditorComponent",content:'IntechOpen is proud to announce that 179 of our authors have made the Clarivate™ Highly Cited Researchers List for 2020, ranking them among the top 1% most-cited.
\n\nThroughout the years, the list has named a total of 252 IntechOpen authors as Highly Cited. Of those researchers, 69 have been featured on the list multiple times.
\n\n\n\nReleased this past November, the list is based on data collected from the Web of Science and highlights some of the world’s most influential scientific minds by naming the researchers whose publications over the previous decade have included a high number of Highly Cited Papers placing them among the top 1% most-cited.
\n\nWe wish to congratulate all of the researchers named and especially our authors on this amazing accomplishment! We are happy and proud to share in their success!
\n'}],latestNews:[{slug:"intechopen-authors-included-in-the-highly-cited-researchers-list-for-2020-20210121",title:"IntechOpen Authors Included in the Highly Cited Researchers List for 2020"},{slug:"intechopen-maintains-position-as-the-world-s-largest-oa-book-publisher-20201218",title:"IntechOpen Maintains Position as the World’s Largest OA Book Publisher"},{slug:"all-intechopen-books-available-on-perlego-20201215",title:"All IntechOpen Books Available on Perlego"},{slug:"oiv-awards-recognizes-intechopen-s-editors-20201127",title:"OIV Awards Recognizes IntechOpen's Editors"},{slug:"intechopen-joins-crossref-s-initiative-for-open-abstracts-i4oa-to-boost-the-discovery-of-research-20201005",title:"IntechOpen joins Crossref's Initiative for Open Abstracts (I4OA) to Boost the Discovery of Research"},{slug:"intechopen-hits-milestone-5-000-open-access-books-published-20200908",title:"IntechOpen hits milestone: 5,000 Open Access books published!"},{slug:"intechopen-books-hosted-on-the-mathworks-book-program-20200819",title:"IntechOpen Books Hosted on the MathWorks Book Program"},{slug:"intechopen-s-chapter-awarded-the-guenther-von-pannewitz-preis-2020-20200715",title:"IntechOpen's Chapter Awarded the Günther-von-Pannewitz-Preis 2020"}]},book:{item:{type:"book",id:"6325",leadTitle:null,fullTitle:"Quality Control in Laboratory",title:"Quality Control in Laboratory",subtitle:null,reviewType:"peer-reviewed",abstract:"The book presents a qualitative and quantitative approach to understand, manage and enforce the integration of statistical concepts into quality control and quality assurance methods. 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Dr. Kucukkocaoglu received his BSBA and MBA Finance degrees form Harmon College of Business Administration-Central Missouri State University, USA, Ph.D. degree from Faculty of Political Sciences-Ankara University, Ankara, Turkey. He has published a number of papers and is serving as a referee for national and international journals. 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I am responsible for developing and maintaining strong relationships with all collaborators to ensure an effective and efficient publishing process and support other departments in developing and maintaining such relationships."}},relatedBooks:[{type:"book",id:"6000",title:"Accounting and Corporate Reporting",subtitle:"Today and Tomorrow",isOpenForSubmission:!1,hash:"aa4f840bdfa861be2f1c4b982a1e2cb5",slug:"accounting-and-corporate-reporting-today-and-tomorrow",bookSignature:"Soner Gokten",coverURL:"https://cdn.intechopen.com/books/images_new/6000.jpg",editedByType:"Edited by",editors:[{id:"70354",title:"Ph.D.",name:"Soner",surname:"Gokten",slug:"soner-gokten",fullName:"Soner Gokten"}],productType:{id:"1",chapterContentType:"chapter",authoredCaption:"Edited by"}},{type:"book",id:"6490",title:"Energy Management for Sustainable Development",subtitle:null,isOpenForSubmission:!1,hash:"a5e77ab7a4bfed7a2a1d524b8a3d6a90",slug:"energy-management-for-sustainable-development",bookSignature:"Soner 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Venkateswarlu",coverURL:"https://cdn.intechopen.com/books/images_new/371.jpg",editedByType:"Edited by",editors:[{id:"58592",title:"Dr.",name:"Arun",surname:"Shanker",slug:"arun-shanker",fullName:"Arun Shanker"}],productType:{id:"1",chapterContentType:"chapter",authoredCaption:"Edited by"}}]},chapter:{item:{type:"chapter",id:"62683",title:"The Hepatic Fate of Vitamin E",doi:"10.5772/intechopen.79445",slug:"the-hepatic-fate-of-vitamin-e",body:'Vitamin E is a lipophilic vitamin and thus naturally mainly occurring in high-fat plant products such as oils, nuts, germs, seeds, and in lower amounts in vegetables and some fruits. The term “vitamin E” comprises different structures that are classified as tocopherols (TOH), tocotrienols (T3), and “vitamin E-related structures”. However, α-TOH is considered as the most important representative of vitamin E in humans as the central vitamin E metabolizing organ, the liver, discriminates for this form [1]. Notwithstanding the classification as vitamin, the way vitamin E exactly contributes to human health is controversially discussed. Vitamin E deficiency has been linked to several disease states like ataxia with vitamin E deficiency (AVED) [2, 3] and Alzheimer’s disease [4, 5], indicating a role in the preservation of human health. AVED has severe neurological consequences and is caused by a defect in the α-TOH transfer protein (α-TTP); the protein responsible for the discrimination of α-TOH from the other vitamin E forms in the liver [2, 3]. This emphasizes the role of the liver as a central organ in human vitamin E handling. The liver further distributes vitamin E in the body [6] and metabolizes excess vitamin E in order to form products for excretion [6] or presumably to produce activated metabolites of vitamin E as known for other lipophilic vitamins [7]. Given the crucial role of the liver for vitamin E handling, this review aims to summarize the knowledge on the physiological hepatic handling of vitamin E as well as on factors influencing hepatic handling of vitamin E.
The liver is the central organ of vitamin E handling. While intestinal absorption efficiency is similar for all forms of vitamin E [8], the plasma concentrations of vitamin E forms differ a lot (e.g., 22.1 μM for α-TOH vs. 2.2 μM for γ-TOH [9]). The preference of α-TOH in the human body is mediated by several complex and interacting hepatic mechanisms.
Vitamin E is absorbed in the intestine along with lipids (for details, see [8]) and is packed into lipoproteins. These are transported via lymph or blood toward the liver (via chylomicron remnants, low density lipoproteins (LDL), and high density lipoproteins (HDL) [10, 11]). Different mechanisms facilitate the cellular uptake of vitamin E: (i) via lipid transfer proteins or lipases, (ii) receptor-mediated lipoprotein endocytosis, and (iii) selective lipid uptake [12]. The degradation of chylomicrons to chylomicron remnants by lipoprotein lipase (LPL) seems to be highly important for vitamin E uptake in the liver; when lipolysis of triglyceride-rich chylomicrons by LPL is inhibited, the α-TOH uptake in the liver is diminished [13]. The phospholipid transfer protein (PLTP) mediates the exchange of phospholipids between lipoproteins [14] and is also able to bind α-TOH in vitro [15]. PLTP-null mice have lower hepatic levels of vitamin E than the wild-type mice [16]; hence, the transfer of vitamin E between the lipoproteins seems to be important for its effective hepatic uptake. The chylomicron remnants and LDL are taken up by the liver via endocytosis, mainly mediated through the LDL receptor (LDLR) or LDLR-related proteins [6, 17]. In addition, the cholesterol transporter Niemann-Pick C1-like 1 (NPC1L1) is involved in hepatic vitamin E uptake; α-TOH binds to the N-terminal domain of NPC1L1, which mediates α-TOH uptake via endocytosis (mechanism similar to intestinal cholesterol uptake) [18]. The scavenger receptor B type I (SR-BI) is known to mediate the uptake of vitamin E in several tissues (e.g., intestine [19], epithelium [20], and hepatocytes [21]) by channeling the molecules into the cells (shown for cholesterol or triglycerides [22]). Furthermore, the scavenger receptor cluster of differentiation 36 (CD36) is likely involved in hepatic uptake of vitamin E [23].
Following its lipophilic nature, vitamin E is transported by intracellular carrier proteins [24]. The intestinally absorbed vitamin E is taken up via endocytosis [25] and follows endosomal fate. Here, the hepatic sorting of vitamin E forms starts as a specific protein, called α-TTP selectively recognizes and preferentially binds α-TOH, which is then extracted from endosomes and transported to the inner leaflet of the plasma membrane [26]. α-TTP is therefore considered to be a “gatekeeper”, which discriminates non-α-TOH forms [27] and regulates the plasma concentrations of α-TOH [1]. The affinity of α-TTP to the different forms of vitamin E differs greatly: it is defined as 100% for α-TOH, whereas β-TOH has 38%, γ-TOH 9%, δ-TOH 2%, and α-tocotrienol (T3) 12% affinity to α-TTP [28]. The regular function of α-TTP is crucial, since missense mutations lead to the disruption of α-TOH distribution and the development of a severe degenerative disease, termed AVED [29]. The transfer of α-TOH from endosomes to the plasma membrane is a multi-step process. First, it is speculated whether the ATP-binding cassette transporter A1 (ABCA1) enriches the outer layer of endosomes with α-TOH [30]. The cholesterol transporter NPC1 may also be involved, as a genetic missense mutation of the NPC1 gene leads to an accumulation of α-TOH in late endosomes [31]. Second, α-TTP extracts the α-TOH from endosomes, and third, α-TTP mediates its transport to the plasma membrane [24]. This process seems to depend on phosphatidylinositol phosphates (PIPs; preferentially PI(4,5)P2 and PI(3,4)P2) in the plasma membrane, as α-TTP binds to them, in turn targeting α-TOH to the plasma membrane and stimulating its release [32]. Chung et al. analyzed the localization of α-TTP depending on the cellular α-TOH concentration [33]. They found (i) perinuclear localization for α-TOH-depleted cells, (ii) a directional transport of α-TOH/α-TTP toward the plasma membrane, when depleted cells were pulsed with a low dose of α-TOH, and (iii) a homogenous cytosolic pattern under long-term and high-dose treatment of cells with α-TOH, which was suggested to be the picture of several α-TOH transport cycles [33]. Furthermore, the authors also postulated a bi-phasic concentration-dependent circulation of α-TTP: the PI(4,5)P2 gradient (low in endosomes and high in plasma membrane) forces the α-TTP-mediated transport of α-TOH toward the plasma membrane, whereas the α-TOH gradient (low in plasma membrane and high in endosomes) triggers the recycling of α-TTP toward the endosomes [33]. It has been proposed that once α-TOH is incorporated into the plasma membrane, it is mediated toward the outer leaflet of the membrane by a flippase, maybe ABCA1, and is then available for the uptake via very low density lipoproteins (VLDL) [34]. For more details on the process, please see Section 2.5 “Release of vitamin E”.
Intracellular storage of vitamin E is limited to the lipophilic sites of the cell, which are membranes and lipid droplets [33]. Not much is known about a specific localization of vitamin E accumulation in liver cells, apart from the observation that lysosomal membranes of rat livers seemed to have the highest concentration of all membranes [35, 36, 37]. However, it is known that one-third of the total body vitamin E is stored in the liver [38]. Within membranes, vitamin E is thought to stabilize the membrane bilayers due to colocalization with phosphatidylcholine [39] and cholesterol (leading to an association to lipid rafts) [40]. It was further hypothesized that vitamin E also colocalizes with poly-unsaturated fatty acids (PUFAs) in nonraft domains in order to provide protection from lipid peroxidation [41]. Newly added α-TOH in cell culture enriches in the same organelles as the endogenous α-TOH pool [42]. Hereby, the subcellular content of α-TOH was directly proportional to the lipid content [43].
Our knowledge about the storage of vitamin E in lipid droplets is also limited. It was recently reported that newly endocytosed vitamin E was also found in lipid droplets, thus indicating endosome-lipid-droplet interactions [33].
The hepatic metabolism of vitamin E has not been fully characterized. However, the principle steps of vitamin E degradation, that is, the shortening of the side chain without the alteration of the chroman ring, are generally accepted. Hence, the metabolites are classified as α-, β-, γ-, and δ-metabolites according to their respective precursors.
In principle, TOHs and T3s are degraded like long branched chain fatty acids (TOH) or long unsaturated branched chain fatty acids (T3) via β-oxidation in peroxisomes. However, as TOHs and T3s do not bear a terminal carboxy function in their side chain, they are not susceptible to β-oxidation. Hence, the initial and rate-limiting step in vitamin E degradation is the introduction of a carboxy function to the ω-terminus of the side chain. This first step is carried out in the endoplasmic reticulum (ER) of liver cells [44]. Here, two representatives of the cytochrome P450 (CYP) protein family, namely, CYP4F2 [45] and CYP3A4 [46, 47], have been identified to catalyze the initial ω-hydroxylation step. The resulting 13′-hydroxychromanol (13′-OH) is then further metabolized via ω-oxidation, a step that most likely involves alcohol dehydrogenase and aldehyde dehydrogenase [44], leading to 13′-carboxychromanol (13′-COOH). The carboxylated side chain resembles a long branched chain fatty acid that is further degradable via β-oxidation. However, a transport mechanism for the carboxychromanol from the ER to the peroxisomes has not been identified so far. Nevertheless, two cycles of peroxisomal β-oxidation after the activation of α-13′-COOH to the respective CoA ester have been suggested [44], as the peroxisomal β-oxidation system has a higher affinity toward long branched chain fatty acids than the mitochondrial counterpart [48]. The proposed 11′- and 9′-COOH metabolites have indeed been identified in human and mouse samples [49] as well as in a hepatic cell line [45, 50]. Subsequently, three more cycles of β-oxidation are needed to form the final product of vitamin E degradation, namely, carboxyethyl hydroxychromanol (CEHC) or 3′-COOH. These steps, however, are assigned to mitochondrial β-oxidation, as CEHC has solely been found in the mitochondria of hepatic cells [44]. Again, the transport mechanisms of the long-chain metabolites (LCM) (13′- to 9′-COOHs) from peroxisomes to the mitochondria are not known. The respective products for each cycle of β-oxidation (7′-COOH, 5′-COOH, and 3′-COOH) have been identified in different human and murine tissues [49, 51, 52, 53, 54] as well as the hepatic cell line HepG2 [45, 47, 51]. Taken together, the hepatic metabolism of vitamin E is characterized by a series of β-oxidation steps after an initial introduction of a carboxy moiety at the ω-terminus of the phytyl-like side chain. The metabolism likely takes place in different cell compartments depending on the enzymatic systems needed for the different degradation steps. However, a concept of vitamin E degradation exclusively in mitochondria cannot be excluded [44]. T3 degradation is believed to follow the same route as TOH degradation but requiring further steps due to the unsaturated side chain. In line with this assumption is the identification of the respective unsaturated metabolites from 13′-carboxytrienol down to carboxymethylbutadienylhydroxychromanol (CMBenHC) in human and mouse samples [49]. According to these findings, the side chain of the T3 metabolites needs a saturation step before the shortening of the chain. Enzymes involved in the degradation of unsaturated fatty acids like 2,4-dienoyl-CoA reductase and 3,2-enoyl-CoA isomerase were suggested to contribute to the degradation of T3s [55].
Following the nature of the lipoprotein metabolism, hepatic release of vitamin E is mostly realized via VLDL. Thus, this section will focus on the packaging of vitamin E into VLDL particles, notwithstanding that the mechanism is not well understood. However, hepatic transfer of vitamin E to HDL has also been suggested [56]. Since it was shown that the expression of α-TTP is crucial for the maintenance of plasma α-TOH levels [57, 58] and that the liver is controlling plasma α-TOH levels [59], hepatic α-TTP is likely involved in the incorporation of vitamin E into lipoproteins. This concept is supported by the observation that nascent VLDL particles are preferentially enriched with RRR-α-TOH after oral administration of vitamin E ([60, 61]. In contrast, in the liver, no preferential retention of RRR-α-TOH was found, indicating that α-TTP is not involved in the delivery of vitamin E to the liver, but in the release from the liver [62]. Hence, efforts have been made to identify the intracellular location of VLDL enrichment with α-TOH mediated by α-TTP [30]. According to the assembly of VLDL, either the rough ER or the Golgi apparatus were assumed. However, the action of α-TTP in these compartments was not confirmed as the nascent VLDL particles contained equal amounts of SRR and RRR α-TOH forms [30]. Further, the inhibition of ER/Golgi action in cells overexpressing α-TTP did not prevent α-TOH secretion [63]. In conclusion, α-TTP is necessary for the hepatic release of vitamin E, but the enrichment of VLDL with RRR-α-TOH occurs after exocytosis.
Based on this, the hypothesis of α-TOH uptake by VLDL directly from the plasma membrane was developed. This idea was inspired by the proposed mechanism of the incorporation of free cholesterol into nascent VLDL [64], that is, the spontaneous transfer from membranes to lipoproteins [65]. The hypothesis involves also the α-TTP-mediated trafficking of vitamin E from late endosomes (where vitamin E occurs after cellular uptake and large parts of α-TTP are located [66]) to the plasma membrane. This process might involve ABCA1, which has been shown to transport α-TOH [67] and could thus present vitamin E to α-TTP at the outer leaflet of the endosomal membrane. After the transport to the plasma membrane, a yet unidentified flippase is required to transfer α-TOH to the appropriate site of the membrane for uptake by nascent VLDL [30]. This hypothesis is supported by findings of Chung et al. [33], which provided a model of α-TTP-facilitated trafficking of vitamin E from endosomes to the plasma membrane (the reader is referred to Section 2.2 “Intracellular trafficking of vitamin E”). Taken together, the release of α-TOH from hepatocytes depends on vesicular transport [21, 31, 63, 68, 69], but is independent from ER or Golgi [63]. Hence, lipoproteins are not loaded with TOH during their intracellular assembly, but rather after exocytosis, a mechanism is required for the presentation of α-TOH at the plasma membrane. Evidence has been provided that the trafficking of α-TOH to the plasma membrane is realized via α-TTP which is located at recycling endosomes in hepatocytes [33]. However, the mechanism of the loading of lipoproteins with α-TOH from the plasma membrane has not been elucidated yet, although the involvement of ABC transporters has been suggested [56, 67, 70]. However, ABC transporters are fueling HDL particles, which is in contrast to the assumption that the hepatic release of α-TOH is mediated via VLDL. In turn, two explanations have evolved: first, α-TOH translocates spontaneously from the membrane to VLDL like free cholesterol [65], and second, α-TOH is transported to HDL via ABCA1 and is then spontaneously transferred to VLDL [71]. However, both hypotheses need evaluation. A recent report on the self-assembly of α-TTP to form nanoparticles and transport vitamin E to tissues protected by endothelial barriers like the brain [34] opens another possible way for the distribution of vitamin E throughout the body starting from the liver.
Key factors in the hepatic handling of vitamin E have been outlined in the previous sections. This section will focus on the action of vitamin E on its own intracellular handling. As indicated above, the key enzyme for the intracellular trafficking of vitamin E is α-TTP, and the rate-limiting enzymes in vitamin E metabolism are CYP4F2 and CYP3A4. Hence, we will here focus on the known actions of vitamin E on these key players.
The key protein of the hepatic handling of vitamin E is α-TTP, with its implications in cellular trafficking, metabolism, and release of vitamin E. Hence, several studies have been conducted to elucidate a possible feedback regulation of α-TTP in response to vitamin E intake, resulting in alterations of the metabolism or the distribution of the vitamin. In principle, research is focused on three levels of regulation: mRNA expression, protein expression, and stabilization of α-TTP protein. However, contradictory results from rodent models have been reported. Fechner et al. found that hepatic α-TTP mRNA expression was strongly induced in rats depleted from vitamin E for 5 weeks after the intake of a TOH-supplemented diet for 24 h [72]. However, rats fed a vitamin E-depleted diet, control diet, or vitamin E-enriched diet for 20 weeks showed upregulation of α-TTP mRNA when vitamin E is deprived, but a downregulation when vitamin E was repleted. Hepatic α-TTP protein levels were comparable for depletion and control, but lowest in rats fed the repleted diet [73]. A similar study reported no differences in hepatic α-TTP mRNA levels of rats fed either a control diet or a diet rich in or low in vitamin E. However, in contrast to the aforementioned study, downregulation of α-TTP protein was reported in the vitamin E-depleted group, while high vitamin E intake did not alter the levels compared to control [74]. The lack of an effect of a vitamin E deficient diet for 290 days on hepatic α-TTP mRNA levels was also reported in another rat model [75]. In line with this, subcutaneous injection of vitamin E for up to 18 days did not alter α-TTP protein levels in rats [76]. However, mice fed a diet rich in vitamin E showed 20% higher hepatic α-TTP protein levels than mice fed a low vitamin E diet [77]. Taken together, some studies report elevated α-TTP levels due to a higher intake of vitamin E [72, 77], but some revealed no effect [74, 75, 76] or even lower levels [73]. Hence, further studies are needed to clarify the role of vitamin E in the regulation of α-TTP. In addition, an in vitro study suggested that vitamin E does not regulate α-TTP at the level of gene expression, but stabilizes α-TTP at the protein level upon binding and thus protects the protein from degradation, leading to higher α-TTP protein levels [78]. Reports on the hepatic mRNA levels might thus be of minor importance for the interpretation of the contribution of vitamin E to α-TTP action; however, the findings on α-TTP protein expression are also inconsistent.
The rate-limiting enzymes of vitamin E metabolism are CYP4F2 and CYP3A4. The latter was reported to be under transcriptional control of pregnane-X-receptor (PXR) [79, 80]. Hence, vitamin E might regulate its metabolism by binding to PXR and subsequent alteration of the expression of the enzymes involved in the first catabolic step. Indeed, studies using cells transfected with reporter genes provided evidence for an activation of PXR by different vitamin E structures (i.e., TOHs, T3s, and metabolites) [81, 82]. Interestingly, α-, δ-, and γ-TOH as well as α- and γ-T3 activated PXR in HepG2 liver cells transfected with human PXR and chloramphenicol acetyl transferase linked to two PXR responsive elements [81], while α- and γ-TOH as well as their metabolites α- and γ-CEHC did not in transfected colon carcinoma cells [82]. However, the LCM α-13′-COOH activated PXR in the latter cellular system and so did γ-T3 [82]. This finding implicates that the LCM of TOH are the responsible mediators of reported TOH actions via PXR. Hence, the findings in hepatic HepG2 cells [81] might be due to a higher catabolic rate of TOH and in turn the more efficient formation of the LCM than in colon cells. However, these findings were made in artificial cellular reporter systems and might not resemble the actual (hepatic) situation in vivo. Further, the specificity of PXR might depend on the species, as γ-T3 (the vitamin E form that activated PXR in both of the aforementioned studies) fails to bind murine PXR [83]. However, results obtained in vivo support the regulation of Cyp3a11 (the murine orthologue of CYP3A4) by vitamin E via PXR. Mice supplemented with α-TOH show elevated hepatic expression of Cyp3a11, while their PXR-deficient counterparts as well as mice with humanized PXR showed no upregulation of Cyp3a11 in response to α-TOH [84]. The same finding was made for Cyp4f13, the murine orthologue of CYP4F2, in this model [84]. These findings suggest that both enzymes are under the control of PXR and murine, but not human PXR is susceptible to α-TOH (or its metabolites as outlined above). Further studies reporting upregulation of hepatic Cyp3a in rodent models with α-TOH supplementation support this finding [76, 83, 85]. Interestingly, in these studies, γ-TOH and γ-T3 had no effect on Cyp3a expression [83, 85], supporting the suggested specificity of murine PXR for α-TOH. In line with this, γ-TOH did not alter the expression of Cyp4f13 in mice [85]. However, subcutaneous application of α-TOH in rats did not induce Cyp4F2 levels [76], which is in contrast to the above mentioned induction of Cyp4f13 in mice via PXR [84]. The reported induction of CYP4F2 activity in HepG2 cells by α-TOH further complicates the interpretation of the data on the effect of vitamin E on CYP4F2 [45]. Taken together, there is evidence for the regulation of CYP4F2 and CYP3A4 via PXR by vitamin E in the human liver. However, several aspects need further clarification, for instance, species and vitamin E isoform specificity of PXR, the regulation of CYP4F2 by vitamin E or the relevance of the α-LCM as true mediators of α-TOH effects via PXR.
Several key enzymes determine the rate of vitamin E catabolism (the reader referred to Section 2.4 “Hepatic metabolism of vitamin E”) and, as outlined in the previous section, there is evidence that vitamin E in general might regulate its own metabolism. However, there are differences in the ability to regulate the metabolism depending on structural properties of the vitamin E isomers (i.e., methylation of the chroman ring, saturation of the side-chain, and stereochemistry). In principle, high intake of vitamin E, independent from the isomer, leads to enhanced formation of the respective metabolites [49]. However, the catabolic rates of the different forms of vitamin E clearly differ: the γ-isoforms are more susceptible to metabolization than the α-isoforms. Subjects supplemented with γ-T3 and α-T3 (125 mg or 500 mg) showed four to six times higher urinary excretion of the catabolic end product γ-CEHC and an induction of α-CEHC only after high dose (500 mg), but not after low dose supplementation (125 mg) [86]. In line with this, equimolar supplementation with 50 mg of α- and γ-TOH leads to a twofold increase of plasma γ-CEHC, but no alterations in α-CEHC [87]. These data indicate that there might be a threshold for the intake of α-TOH and α-T3 (or plasma levels, respectively) that needs to be exceeded to accelerate catabolism of α-TOH and α-T3 to form α-CEHC, as suggested by Schuelke et al. [88]. Interestingly, already in 1985, Handelman et al. reported that high α-TOH levels in human plasma are related to low γ-TOH levels [89]. After supplementation of α-TOH, the plasma α-TOH levels were, as expected, twofold to fourfold higher, but the γ-TOH level decreased to between one-third and one-half of the initial level [89]. Hence, α-TOH intake seems to boost γ-TOH catabolism. Supporting data were generated in a rat model, where the combined supplementation of α- and γ-TOH leads to higher excretion of γ-CEHC than the supplementation of γ-TOH alone [90], as well as the reported stimulation of γ-TOH catabolism by α-TOH in HepG2 liver cells [91]. Although the underlying mechanisms are not fully unraveled, there is evidence that α-TOH induces the activity of enzymes involved in the metabolism of vitamin E, leading to the degradation of non-α-forms, while α-TOH remains protected (please refer to Section 3.1.1 “Intracellular handling of vitamin E”).
Sesamin is a lignan, a group of natural compounds derived from vegetable sources, like sesame seeds [92]. Sesamin is known as a natural inhibitor of the metabolism of TOH [93, 94, 95, 96, 97]. The cell regulatory actions of sesamin have been initially investigated in in vitro models, where Parker et al. showed that sesamin acts as a selective inhibitor of CYP3A4, an initial enzyme of TOH metabolism [46]. In this study, the authors compared the inhibitory potential of sesamin on TOH metabolism in human HepG2 cells to the well-characterized CYP3A4 inhibitor ketoconazole. HepG2 cells were treated with one of the mentioned compounds in combination with either 25 μM α-TOH or 25 μM γ-TOH. Afterwards, the concentration of the corresponding CEHC was determined as a marker for TOH metabolism in cell culture media. It became apparent that ketoconazole (1 μM) and sesamin (1 μM) inhibited the formation of α- and γ-CEHC. This result provides evidence that sesamin is able to modulate TOH metabolism via the inhibition of CYP3A4 [46]. In addition to the in vitro data, Uchida and coworkers investigated the inhibitory effects of sesamin on vitamin E metabolism in rats. Vitamin E-deficient rats (vitamin E free diet for 4 weeks) were treated with 50 mg/kg RRR-α-TOH alone or in combination with 200 g/kg sesame seeds [95]. Next, the concentration of α-TOH in different tissues as well as the urinary excretion of α-CEHC was measured. The urinary excretion of α-CEHC in the sesamin group was significantly lower compared to the α-TOH control group. Further, the combination of α-TOH and sesamin provoked a significant increase of hepatic α-TOH concentrations compared to α-TOH treated animals [95]. These observations have been confirmed in other animal studies [93, 94]. Beside the investigations in animal models, there are also a few results originating from studies in humans. In 2004, Frank and colleagues used muffins enriched with sesame oil (94 mg sesamin/muffin) or corn oil (control) to investigate the effect of a single dose sesamin application on urinary excretion of γ-CEHC as well as blood levels of γ-TOH in 10 healthy volunteers [97]. Both, control and intervention group, received the muffins together with a capsule containing deuterium-labeled γ-TOH (50 mg) in a crossover design. Blood and urine samples were collected over 72 hours after the application of the muffins and capsules. While the urinary excretion of γ-CEHC was significantly lowered, the sesamin treatment did not affect γ-TOH concentrations in blood compared to the corn oil control group [97]. Unfortunately, the study does not provide data on the elevation of the hepatic γ-TOH concentration in response to the reduced urinary excretion of γ-CEHC. Taken together, in vitro and in vivo studies provide evidence that the dietary intake of sesamin leads to an increase of the hepatic concentration of TOH via the inhibition of vitamin E metabolism, but further experiments are needed to characterize the interaction of sesamin and vitamin E metabolism in more detail.
The pharmacological modification of the enzymatic activity of CYP3A4 represents an effective way to influence vitamin E homeostasis in the human body. Mechanistically, the direct or indirect interference of vitamin E metabolism is usually just a side effect of the pharmacological inhibition or induction of CYP3A4 by various chemical compounds. Thus, it is not surprising that the first evidence for the involvement of CYP3A4 in vitamin E metabolism was provided in an experimental subset using ketoconazole as a specific inhibitor for CYP3A4 [46, 98]. In HepG2 liver cells, different concentrations of ketoconazole (1 mmol/l or 0.25 mmol/l) inhibited the metabolic conversion of γ- and δ-TOH (25 μmol/l cell culture media) to γ- or δ-CEHC by almost 90% [46]. This finding has been confirmed by the reproduction of the same experiment with sesamin, the natural inhibitor of CYP3A4, revealing comparable results [46]. The inhibitory effect of ketoconazole on vitamin E metabolism has further been observed in an in vivo model. Here, rats were supplemented with ketoconazole (50 mg/kg body weight) together with α-TOH (10 mg/kg body weight), γ-TOH (10 mg/kg body weight) or mixture of different T3s (29.5 mg/kg body weight). Ketoconazole significantly reduced the catabolism of all applied vitamin E forms resulting in impaired urinary excretion of the respective CEHCs [99]. Beside its inhibition, the pharmacological induction of CYP3A4 represents another way to modulate vitamin E metabolism. Birringer and coworkers demonstrated that 50 μmol/L rifampicin, an inducer of CYP3A4 activity [100], induced the degradation of all-rac-α-TOH in HepG2 cells fivefold [47]. In this study, the cell culture medium has been preconditioned with 100 μmol/L α-TOH for 10 days, as the standard medium was deficient for α-TOH [47]. Further, an indirect approach for the modulation of vitamin E metabolism via the modification of CYP3A4 expression could be realized by triggering PXR, a nuclear receptor that regulates the expression of metabolic enzymes and transporters involved in the metabolism of xenobiotics and endobiotics [101, 102]. Landes and coworkers showed that γ-T3 as well as rifampicin acts as PXR agonists, thus upregulating CYP3A4 mRNA expression in HepG2 liver cells [81]. Given the fact that enhanced mRNA expression of CYP3A4 results in enhanced enzymatic activity, the stimulation of PXR by various pharmacological agonists or antagonists could also modulate the hepatic metabolism of vitamin E. In summary, the direct or indirect regulation of CYP3A4 by various pharmacological means represents an effective way to modify the hepatic vitamin E metabolism.
The handling of vitamin E is also influenced by nonmodifiable factors. These are aging, gender, and individual genetics. Published data in this area are sparse but interesting.
The aging process is characterized by nine hallmarks: genomic instability, telomere attrition, epigenetic alterations, loss of proteostasis, deregulated nutrient sensing, mitochondrial dysfunction, cellular senescence, stem cell exhaustion, and altered intercellular communication [103]. In particular, the mitochondrial dysfunction leads to higher formation of reactive oxygen species (ROS) and enhanced oxidative damage [104]. Both processes can be diminished by the antioxidant function of vitamin E [105]. Consequently, two questions arise: (i) can vitamin E modulate the aging processes or prevent age-related diseases? This has been subject of several reviews [106, 107, 108, 109]. (ii) And how is the concentration, distribution, and function of vitamin E modulated by the aging process? In humans, age-dependent changes of α-TOH plasma concentrations are known. In healthy aged humans, the α-TOH plasma concentrations are higher than in younger individuals [110, 111, 112, 113]. However, this might be due to the age-related increase of plasma cholesterol concentrations, as the age-related increase in α-TOH plasma concentrations disappear after adjustment for cholesterol plasma concentrations [112] or serum lipids [113]. Traber et al. suggested that α-TOH plasma concentrations are more dependent on control mechanisms for plasma lipids rather than on α-TOH absorption [113]. Hospitalized elderly patients [114] as well as older persons with cognitive impairments (dementia or Alzheimer’s disease [115, 116]) have low α-TOH plasma concentrations [117]. However, an unfavorable nutrient status of the hospitalized patients was discussed as the cause of the lower α-TOH plasma concentrations.
Several studies analyzed the age-dependent changes of α-TOH tissue concentrations and handling in mice [37, 117, 118, 119] and rats [120]. In brain [37, 117, 118] and kidney [37, 117], epididymal adipose tissue [117] and aortic vessel wall [120], a consistent increase in α-TOH was found with age. In old rats, however, an age-dependent increase in intestinal absorption was found [121]. This was considered as a “self-protective age-dependent adaption” [120], which is thought to counteract increased oxidative stress during aging. In the liver and heart, however, data are conflicting: while some found increased concentrations [37, 119, 120], Takahashi et al. found decreased values [117]. Two studies also analyzed the age-dependent regulation of genes, known to be involved in vitamin E handling, which are α-TTP, ABCA1, and Cyp4f14 (murine orthologue of CYP4F2) [117] as well as NPC1, NPC2, and LPL [37]. Takahashi et al. found increasing (mice with the age of 3–12 month) and then decreasing (12—24 months) α-TTP protein levels in the liver, while mRNA expression was stable over age [117]. Overall, Cyp4f14 mRNA expression decreased during aging (60% decrease in mRNA expression at the age of 24 months compared to the age of 3 weeks), while ABCA1 mRNA expression slightly increased (20% in the same age range as measured for Cyp4f14) [117]. The authors concluded that the age-related changes of hepatic α-TOH levels cannot be explained by the metabolism of α-TOH via Cyp4f14. König et al. analyzed protein expression in kidney tissue or its lysosomal membranes and found a significant decrease of NPC1 and NPC2, but a prominent increase in LPL (361% compared with the tissue from younger mice) [37]. The increased expression of LPL may explain the accumulation of α-TOH in aged mice. Furthermore, NPC1 and NPC2 may be responsible for the transport of α-TOH from the endosomes to the cytosol [69] and their reduced expression may explain the accumulation of α-TOH in lysosomal membranes [37]. In summary, there are age-dependent changes in α-TOH tissue and plasma concentrations and also in the expression of genes responsible for vitamin E handling; however, the underlying regulatory processes are not unraveled completely yet.
The sex-dependent differences in vitamin E handling were described recently by Schmölz et al. [6] and will be summarized here briefly for humans only. While intake of vitamin E in total is higher in men than in women [122], the intake per kcal is higher for women than for men [123]. The absorption of α-TOH seems not to be influenced by sex, but is mainly regulated by downstream regulatory processes (likely by hepatic sorting or metabolism) [113]. The data on serum concentrations of vitamin E are inconsistent: while some researchers reported elevated α-TOH serum concentrations for women compared to men [124, 125], others found contradictory results [123]. Sex-dependent regulation of vitamin E metabolism is specific for the different forms of vitamin E. Women degrade γ-TOH to a higher degree than men, while the metabolism of α-TOH seems to be independent [87]. Two mechanisms may be relevant for sex-dependent regulation of vitamin E metabolism: the hormonal status of individuals and the activation of the CYP enzymes involved in vitamin E metabolism [6]. Further studies could illuminate gender-specific differences in more detail. In the light of the discovery of vitamin E as a factor that limits female fertility, this is of special interest.
The influence of genetics on vitamin E handling was summarized in detail in a recent review (for more details, please see [6]). Therefore, only a short overview will be provided here. Interindividual differences in the handling of vitamin E can be caused by individual genetic constitutions. Polymorphisms in genes, which are responsible for vitamin E handling such as CYP4F2 [126], NPC1L1 [127], and CD36 [128] are likely to contribute to variations in vitamin E status. The best-studied gene in this context is α-TTP, as its genetic variability may cause AVED. Two genetic variants are known, which are located in or nearby the proposed tocopherol-binding domain and cause reduced α-TOH serum concentrations [129]. Furthermore, mutations in the promoter region of α-TTP (with increased or decreased activity) were also reported [130]. In summary, vitamin E handling is influenced by several mechanisms, one of which is the variability of genes involved in these processes. This might held responsible for interindividual differences in vitamin E serum concentrations.
Nonalcoholic fatty liver disease encompasses a histological spectrum ranging from simple steatosis to nonalcoholic steatohepatitis (NASH). NASH is a clinical symptom characterized by a pattern of steatosis, inflammation, and hepatocyte ballooning, which can result in the development of cirrhosis and liver cancer [131]. Although the molecular mechanisms of NASH development remain poorly understood, studies provide evidence for a critical role of oxidative stress together with an impaired antioxidative response [132, 133]. In line with this, Erhardt and coworkers observed significantly lower plasma levels of α-TOH and other antioxidants in NASH patients compared to healthy controls [134]. Given the fact that an induction of CYP3A4 or CYP4F2 results in decreased vitamin E concentrations in the human body, it has been expected that NASH leads to an enhanced activity or expression of these enzymes. Thus, Woolsey and coworkers investigated the enzymatic activity as well as the mRNA expression of CYP3A4 in NASH patients [135]. The authors used liver biopsies for mRNA analyses and determined the concentration of 4β-hydroxycholesterol in plasma as an endogenous biomarker for CYP3A4 activity. Interestingly, NASH patients showed a 37% reduced enzymatic activity of CYP3A4 as well as a 69% lower CYP3A4 mRNA expression compared to healthy controls [135]. Unfortunately, there is no further data on the activity or the expression of CYP4F2 in NASH patients. However, Athinarayanan and coworkers investigated the influence of two different CYP4F2 genotypes (V433 M and W12G) on vitamin E plasma concentrations in NASH patients [136, 137, 138]. The V433 M genotype was associated to higher baseline levels of vitamin E, indicating lower enzymatic activity compared to the W12G genotype [136, 137, 138]. Thus, the authors hypothesized that the W12G genotype in NASH patients could explain the lower vitamin E plasma concentrations. However, this hypothesis has been disproved by the finding that the vitamin E plasma concentrations of NASH patients did not differ between the two CYP4F2 genotypes [136, 137, 138]. Based on the available data, CYP4F2 and CYP3A4 seem to have no influence on vitamin E plasma concentrations during the NASH development. Next to the CYPs, α-TTP could also be involved in a potential mechanism explaining the observation of Erhardt and coworkers mentioned above. In line with this, Ban and coworkers used a rat model to investigate whether an exposure to hyperoxia (>95% O2 for 48 h), an established stimulus for ROS production [139], could alter the expression of hepatic α-TTP [140]. Indeed, hyperoxia decreased the expression of α-TTP mRNA in rat liver, while α-TTP protein expression remained unchanged [140]. As oxidative stress and ROS formation are crucial factors for NASH development, lowering α-TTP expression by ROS could explain the lower vitamin E levels in NASH patients. In summary, the concentration of vitamin E and other antioxidants is reduced in NASH patients by yet not fully understood molecular mechanisms, potentially involving α-TTP. Nevertheless, recent human intervention trials provide evidence that vitamin E treatment could improve primary NASH outcomes (i.e., steatosis, inflammation, hepatocellular ballooning, and fibrosis) [137, 138].
The current data on vitamin E as a potential agent for cancer therapy are inconsistent. While in vitro and early epidemiological studies provided evidence for cell growth-inhibiting, anti-proliferative and pro-apoptotic effects of vitamin E in cancer treatment [141, 142, 143, 144, 145], more recent investigations reported contradictory results [146, 147, 148]. These findings were further sustained by the “Selenium and Vitamin E Cancer Prevention Trial (SELECT),” a randomized intervention study to determine the long-term effect of a supplementation of vitamin E (400 IU/d all-rac-α-tocopheryl-acetate) and selenium (200 μg/d L-selenomethionine) on the risk of prostate cancer in healthy men. Interestingly, the authors observed an increased incidence for prostate cancer in subjects supplemented with vitamin E [149]. Beside the investigations on beneficial effects of vitamin E in cancer therapy, almost nothing is known about the influence of cancer on human vitamin E homeostasis. An early study by Knekt, who investigated the association of vitamin E serum concentrations and the risk for different types of female cancer, showed an inverse relation between α-TOH serum concentrations and cancer risk [150]. Thus, women with the lowest α-TOH levels were at enhanced risk for cancer compared to those with higher α-TOH levels. Indeed, this association was restricted to cancer outcomes in tissues and organs, which were not exposed to estrogens [150]. Thus, Knekt hypothesized that low vitamin E levels could represent a potential risk factor for several, but not all types of cancer [150]. Nevertheless, the molecular mechanisms underlying this impairment of vitamin E serum concentrations in cancer patients remain unclear. The enhanced metabolic conversion of vitamin E might represent a mechanistic explanation. In line with this, investigations of tissues from cancer patients showed elevated expression of CYP3A4 [151] and CYP4F2 [152], the two major enzymes of vitamin E catabolism. Unfortunately, vitamin E serum concentrations have not been determined in these studies. Further, in vitro studies provided evidence that cancer also affects transporters for vitamin E, such as the tocopherol-associated protein (TAP) [153]. Tissue samples from prostate cancer patients showed significantly lower TAP mRNA expression compared to healthy controls, indicating that cancer may affect the intracellular transport of vitamin E. In addition, the overexpression of TAP in prostate cancer cells leads to a significant reduction of cell growth, while a TAP knockdown by small interfering RNA increased their growth [153]. Interestingly, these effects appeared without additional vitamin E treatment, indicating that TAP not only mediates vitamin E transport but also functions as a vitamin E-independent tumor suppressor gene [153]. In summary, the promising cancer preventive effects of vitamin E shown in vitro have not been confirmed in recent in vivo trials. Nevertheless, cancer could probably be associated with reduced vitamin E concentrations in the human body, because of an enhanced vitamin E catabolism and/or the alteration of its intracellular transport. However, further investigations are required to validate these results.
After its intestinal absorption, the transport of vitamin E, including its transfer to and its export from the liver as well as the subsequent distribution of vitamin E in the human body, strictly depends on different lipoproteins [7]. Thus, disorders of the lipoprotein metabolism can lead to disturbances of vitamin E homeostasis. Abetalipoproteinemia or Bassen-Kornzweig syndrome is a rare form of neurodegenerative ataxia with a strong impact on the hepatic handling of vitamin E. Abetalipoproteinemia is caused by mutations in the gene encoding for the microsomal triglyceride transfer protein (MTP), which is required for the assembly and secretion of the apolipoprotein B (apoB) forms in the liver and the intestine [154]. The apoB forms are the primary apolipoproteins associated to chylomicrons or VLDL, IDL, and LDL, respectively, and are thus essential for the distribution of vitamin E in the human body [7, 155]. As a result of the disturbed intestinal absorption and hepatic excretion of all lipid soluble molecules, patients with abetalipoproteinemia show vitamin E deficiency as well as low serum concentrations of cholesterol and triglycerides [156]. Next, the hepatic handling of vitamin E can be affected by familial hypobetalipoproteinemia. This lipoprotein disorder is caused by mutations in the APOB gene, leading to disturbances of translation of the apoB proteins and/or impaired secretion of VLDL [157]. Thus, familial hypobetalipoproteinemia displays the same clinical features as abetalipoproteinemia. In summary, lipoprotein disorders exert clear impact on the hepatic and systemic handling of vitamin E.
AVED is a neurological disorder, which has for the first time been described in a 12-year-old boy with cerebellar ataxia and low serum vitamin E concentrations. Interestingly, the boy showed no lipid malabsorption or a lack of lipoproteins, like it has been observed in abetalipoproteinemia [158]. Subsequent studies identified a mutation in the TTPA gene, the gene encoding for α-TTP, as the disease causing factor [159]. Thus, AVED patients have impaired expression of α-TTP, leading to impaired incorporation of vitamin E (α-TOH) into VLDL as well as a higher metabolic conversion and excretion of vitamin E [154]. In addition, AVED patients show very low plasma vitamin E concentrations together with normal absorption rates for vitamin E in the absence of intestinal malabsorption and abetalipoproteinemia [2, 154]. In summary, AVED represents a clinical condition that includes altered hepatic handling of vitamin E without affecting lipoprotein homeostasis.
In the last decades of vitamin E research, the liver appeared as the central organ for the uptake, distribution, metabolism, and storage of vitamin E. Thus, it is also a starting point for various strategies for the modulation of the vitamin E homeostasis. Based on current knowledge, we identified physiological, nonphysiological as well as pathophysiological factors influencing the hepatic handling of vitamin E, verifying the crucial role of the liver in vitamin E homeostasis (a brief schematic overview is provided in Figure 1). Nevertheless, further studies are needed to unravel the molecular mechanisms underlying the described disturbances of hepatic vitamin E handling by various factors.
The crucial role of the liver in vitamin E homeostasis.
The work of S.L. is supported by grants from the Federal Ministry of Education and Research (01EA1411A), the Deutsche Forschungsgemeinschaft (DFG; RTG 1715), and the German Ministry of Economics and Technology (AiF 16642 BR) via AiF (German Federation of Industrial Research Associations) and FEI (Research Association of the German Food Industry) and by the Free State of Thuringia and the European Social Fund (2016 FGR 0045). The work of L.S. is supported by the Free State of Thuringia and the European Social Fund (2016 FGR 0045). The work of M.W. is also funded by the DFG (Wa 3836/1-1).
There are many conventional zinc oxide (ZnO) nanostructure synthesis routes employing the chemical and physical methods, which require particular set-up, high cost, high temperature-pressure conditions, and nonecological chemicals [1]. However, high-energy consumption of these routes and released toxic chemicals after the process can be hazardous to the environment and human health. In recent years, the green synthesis approach has been gaining attention, which eliminates the use of toxic chemicals and applies environmentally friendly routes. These strategies handle the use of plant-extracts, microorganisms, biomolecules, and ionic liquids by applying hydrothermal, microwave-assisted, sonochemical, and low-temperature processes (Figure 1).
\nGreen synthesis strategies of ZnO nanostructures with various morphologies.
The aim of these developments is to allow the use of toxic chemicals and reduce energy consumption by using simple, rapid, and safe routes. Green synthesis strategies for the ZnO nanostructures could be summarized as biosynthesis (natural extract-based, microorganism-based, and biomolecule-based) and nontoxic chemical synthesis (water-based, calcination, solvent-free, and ionic liquid).
\nNatural extracts (mainly phytochemicals) obtained from plants, leaves, fruit peels, flowers, and seeds have been utilized for the green synthesis of metal oxide nanoparticles for years. After the plants are collected from different sources, they are washed with water and basic extraction procedures are applied to obtain plant extracts in which leaves are ground and immersed in water by stirring at room temperature for a while. Then, the solutions are filtered and the eluted extract solution is separated for further use in ZnO synthesis (Figure 2). The eluent solution could be used directly for ZnO synthesis or could be dried for the concentration of solid extracts. Afterward, zinc precursors and plant extracts are reacted under various pH and temperature conditions [2]. If the extract is used as an aqueous solution, the zinc precursors are added into the solution. Otherwise, the zinc precursor and powder form of leaf extract are mixed in distilled water. The key mechanism is the oxidation and reduction of metal ion ‘zinc’ by phytochemicals, which are found in natural extracts. The leaf extracts behave as reducing and capping agents. Under favor of plant extracts, the synthesis procedure can be accomplished without using any chemical stabilizers. Finally, the obtained powders are washed with methanol or ethanol and annealed at high temperatures to attain purity [3].
\nSynthesis route of ZnO nanostructures from leaf extracts.
The green synthesized ZnO nanoparticles have been used in various fields such as biomedical application due to their significant antibacterial activities, photocatalysis, and metal ion adsorption purposes [4]. Moreover, nanoparticles synthesized by the green route exhibit better antibacterial performances due to the functional groups on their surfaces that come from phytochemicals. Here, we will describe the main applications of natural extract-based green synthesized ZnO nanoparticles.
\nThe advantage of using natural extracts for the synthesis of ZnO nanoparticles is that coating of nanoparticles with various pharmacologically active biomolecules on the metal oxide surface allows the conjugation of nanoparticles with receptors of the bacterial membrane. These molecules might be flavones, aldehydes, amides, polysaccharides, etc. and the green synthesized nanoparticles exhibit better biomedical activity than the chemically synthesized ones [1]. Inorganic metal oxides have widely emerged as antibacterial, antioxidant, antifungal, and anticancer agents in the last decades. Moreover, because of their specific targeting and nominal toxicity, the metal oxide nanoparticles could be used in personalized medicine applications. In the area of nanoscaled metal oxides, ZnO has shown promising activity in the biomedical field due to its unique electronic, optical, and medicinal properties. The ZnO nanoparticles show antibacterial activity against a broad spectrum of pathogenic bacteria, and these nanoparticles adopt various mechanisms such as reactive oxygen species (ROS) generation, cell membrane integrity disruption, biofilm formation, or enzyme inhibition [5]. Under UV irradiation, ROS such as superoxide ions, hydroxyl ions, singlet oxygen species, and peroxide molecules are formed. The formed peroxide ions could easily penetrate through the cell membrane and result in cell death. Figure 3 shows the possible ROS generation mechanism and its effect on the bacterial cell wall.
\nROS mechanism of ZnO nanoparticles [6].
Cell membrane integrity disruption is another significant mechanism for the antibacterial effect of ZnO nanoparticles. Penetration of ZnO nanoparticles results in cell death by the loss of phospholipid bilayer integrity and leakage of intracellular components of the cell. While the Gram-positive bacteria have a thick layer of peptidoglycan, teichoic acid, and lipoteichoic acid in their cell membrane, Gram-negative bacteria have a triple layer of peptidoglycan. The different structure of cell membranes of these two types of bacteria results in a different mechanism of nanoparticle penetration through the cell membranes. In this part, we focused on the biomedical activity of ZnO nanoparticles, and in Table 1, the used plant extracts, zinc precursors, biomedical applications and related biomolecules are summarized.
\nPlant type | \nZinc precursor type | \nSize of ZnO (nm) | \nTreated biomolecule | \nBiomedical field | \nRef. | \n
---|---|---|---|---|---|
Momordica charantia | \nNitrate | \n21 | \nR. microplus, P. humanus capitis, An. stephensi, and Cx. Quinquefasciatus | \nAntiparasitic | \n[30] | \n
Rosa canina | \nNitrate | \n11 | \nS. typhimurium, S. aureus, L. monocytogenes, and E. coli | \nAntibacterial | \n[31] | \n
Ulva fasciata | \nChloride | \n16 | \nB. cereus, S. aureus, S. thermophilus, E. coli, and P. aeruginosa | \nAntibacterial | \n[32] | \n
Eclipta alba | \nAcetate | \n6 | \nE. coli | \nAntibacterial | \n[33] | \n
Terminalia arjuna | \nAcetate | \n21 | \nE. coli and S. aureus | \nAntibacterial | \n[34] | \n
Stevia | \nAcetate | \n10–90 | \nE. coli and S. aureus | \nAntibacterial Antiparasitic | \n[35] | \n
Glycosmis pentaphylla | \nAcetate | \n32–36 | \nB. cereus, S. aureus, S. paratyphi, S. dysenteriae, C. albicans, and A. niger | \nAntibacterial | \n[36] | \n
L. leschenaultiana | \nAcetate | \n— | \nL. sericata | \nAntiparasitic | \n[37] | \n
Adhatoda vasica | \nAcetate | \n10–12 | \nS. epidermidis, E. coli, P. aeruginosa, T. rubrum, and M. audouinii | \nAntimicrobial | \n[38] | \n
Vitex negundo | \nNitrate | \n38 | \nHuman Serum Albumin | \nProtein binding | \n[39] | \n
Anchusa italica | \nAcetate | \n8–14 | \nB. megaterium, S. aureus, E. coli, S. typhimurium | \nAntimicrobial | \n[40] | \n
Jacaranda mimosifolia | \nGluconate | \n2–4 | \nE. faecium and E. coli | \nAntibacterial | \n[41] | \n
Heritiera fomes and Sonneratia apetala | \nChloride | \n40–50 | \nS. aureus, S. flexneri, V. cholera, S. epidermidis, B. subtilis, and E. coli | \nAntibacterial Anti-inflammatory | \n[42] | \n
Nyctanthes arbor-tristis | \nAcetate | \n12–32 | \nA. alternata, A. niger, B. cinerea, F. oxysporum, and P. expansum | \nAntifungal | \n[43] | \n
Nocardiopsis sp | \nNitrate | \n500 | \nE. coli and P. mirabilis | \nAntibacterial | \n[44] | \n
Ceropegia candelabrum | \nNitrate | \n12–35 | \nS. aureus, B. subtilis, E. coli, and S. typhi | \nAntibacterial | \n[45] | \n
Pongamia pinnata | \nAcetate | \n21 | \nC. maculatus | \nAnti-pesticide | \n[46] | \n
Capsicum annuum | \nNitrate | \n30–40 | \nE. coli and S. aureus | \nAntibacterial | \n[47] | \n
Euphorbia petiolata | \nNitrate | \n— | \nE. coli | \nAntibacterial | \n[48] | \n
Tradescantia pallida | \nAcetate | \n25 | \nHeLa cervical cancer cell | \nAnticancer | \n[49] | \n
Punica granatum | \nNitrate | \n5 | \nP. vulgaris, E. coli, and S. aureus | \nAntibacterial | \n[50] | \n
Swertia chirayita | \nNitrate | \n5 | \nS. enterica, E. coli, and S. aureus | \nAntibacterial | \n[51] | \n
Vitex negundo | \nNitrate | \n38 | \nE. coli and S. aureus | \nAntibacterial | \n[52] | \n
Aegle marmelos | \nNitrate | \n20 | \nP. aeruginosa, E. coli, S. aureus, A. niger, and F. solani | \nAntimicrobial | \n[53] | \n
Azadirachta indica | \nAcetate | \n9–25 | \nS. pyogenes, E. coli, and S. aureus | \nAntibacterial | \n[54] | \n
Vaccinium arctostaphylos | \nNitrate | \n15 | \nLipid, insulin, and fasting blood sugar | \nAntidiabetic | \n[55] | \n
Coffee powder | \nAcetate | \n25–30 | \nProteinase K | \nEnzymatic | \n[56] | \n
Ziziphus nummularia | \nNitrate | \n17 | \nHeLa cancer cell and Candida spp. | \nAnticancer Antifungal | \n[57] | \n
Chelidonium majus | \nNitrate | \n10 | \nP. aeruginosa, E. coli, S. aureus, C. albicans, A. niger, and T. rubrum | \nAntibacterial | \n[58] | \n
Prunus yedoensis Matsumura | \nNitrate | \n10–40 | \nB. linens and S. epidermidis | \nAntibacterial | \n[59] | \n
Sechium edule | \nAcetate | \n36 | \nB. subtilis and K. pneumoniae | \nAntibacterial | \n[60] | \n
Catharanthus roseus | \nAcetate | \n50–90 | \nS. aureus, S. pyogenes, B. cereus, P. aeruginosa, P. mirabilis, and E. coli | \nAntibacterial | \n[61] | \n
Albizia saman | \nNitrate | \n15–80 | \nD. indica | \nGenotoxicity | \n[62] | \n
Parthenium hysterophorus | \nNitrate | \n16–45 | \nB. subtilis, S. aureus, K. pneumonia, E. coli, and Enterobacteraerogens | \nAntibacterial | \n[63] | \n
Sarcopoterium spinosum | \nAcetate | \n26–115 | \nE. coli, P. aeruginosa, S. aureus, S. pneumoniae, B. subtilis, E. faecalis, C. glabrata, and C. albicans | \nAntibacterial | \n[64] | \n
Ruta graveolens | \nNitrate | \n28 | \nK. aerogenes, P. aeruginosa, E. coli, and S. aureus | \nAntibacterial | \n[65] | \n
Pichia kudriavzevii | \nAcetate | \n10–61 | \nB. subtilis, S. epidermidis, S. aureus, E. coli, and S. marcescens | \nAntibacterial | \n[66] | \n
Azadirachta indica | \nSulfate | \n84 | \nXanthomonas oryzae pv. oryzae | \nAntibacterial | \n[67] | \n
Coptidis Rhizoma | \nNitrate | \n3–25 | \nB. megaterium, B. cereus, E. coli, and B. pumilus | \nAntibacterial | \n[68] | \n
Polygala tenuifolia | \nNitrate | \n33–73 | \nRAW 264.7 cells | \nAntibacterial | \n[69] | \n
Sargassum muticum | \nAcetate | \n10–15 | \nWEHI-3 cell | \nAnticancer | \n[70] | \n
Scadoxus multiflorus | \nAcetate | \n31 | \nAedes aegypti, A. niger, and A. flavus | \nAntifungal | \n[71] | \n
Eclipta prostrata | \nNitrate | \n29 | \nHep-G2 cell | \nAnticancer | \n[72] | \n
Azadirachta indica | \nNitrate | \n30–60 | \nE. coli and S. aureus | \nAntibacterial | \n[73] | \n
Calotropis gigantea | \nAcetate | \n25 | \nDNA | \nAnticancer | \n[74] | \n
Andrographis paniculata | \nNitrate | \n57 | \nα-Amylase and BSA | \nAntioxidant | \n[75] | \n
Tamarindus indica, Moringa oleifera | \nNitrate | \n27–54 | \nα-Amylase and α-glucosidase | \nAntidiabetic Antioxidant | \n[76] | \n
Bauhinia tomentosa | \nSulfate | \n22–94 | \nB. subtilis, S. aureus, E. coli, and P. aeruginosa | \nAntibacterial | \n[77] | \n
Egg albumen | \nAcetate | \n16 | \nC. albicans | \nAnticandidal | \n[78] | \n
Moringa oleifera | \nAcetate | \n40–45 | \nA. saloni, S. rolfii, S. aureus, and E. coli | \nAntibacterial | \n[79] | \n
Cassia fistula | \nNitrate | \n5–15 | \nK. aerogenes, E. coli, P. desmolyticum, and S. aureus | \nAntibacterial | \n[80] | \n
Biomedical activities of plant extract-based synthesized ZnO nanoparticles.
Sathishkumar et al. synthesized ZnO nanoflakes using Couroupita guianensis Aubl leaf extract and demonstrated the bactericidal activity against various types of bacteria. They reported that the main constituent of the extract, phenol, reduced the zinc acetate precursor into ZnO nanostructures. In history, Aloe (Aloe barbadensis Miller) extract has been used in therapeutic applications due to its antifungal, antidiabetic, anticancer, and antibacterial properties. Ali et al. also synthesized small-sized ZnO nanoparticles using Aloe vera extract and showed their antibacterial activity and cell damage of Escherichia coli and MRSA cells before and after ZnO nanoparticle treatment [7]. On the other hand, Gunalan et al. synthesized ZnO nanoparticles with Aloe vera extract and compared their results with the chemically synthesized ZnO nanoparticles. The results proved the enhanced antibacterial activity against various pathogens, and variation in the particle size is responsible for the significant bactericidal activity [8]. The effect of Aloe vera extract on the synthesis of ZnO nanoparticles and their antibacterial activity was investigated. Moreover, the antioxidant activity of the particles was evaluated by using five different free radical scavenging assays and the anticancer activity was tested against three cancerous cell lines [9].
\nIn a very recent work, Zare et al. presented the effect of Cuminum cyminum leaf extract on the synthesis of ZnO nanoparticles by using zinc nitrate precursors. The resulting nanoparticle diameter is around 7 nm, and the nanoparticles show high sensitivity to Gram-negative bacteria [10]. The nanoparticle formation of zinc nitrate precursors has been investigated by using several types of plant extracts such as Limonia acidissima [11, 12], Cochlospermum religiosum [13], Tabernaemontana divaricata [14], Conyza Canadensis, Citrus maxima [15], Aristolochia indica [16], Echinacea [17], Mentha [18, 19], Salvadora oleoides [20], Boswellia ovalifoliolata [21], and Costus pictus [22]. The synthesized ZnO nanostructures have shown an enhanced antibacterial effect for a broad spectrum of bacterial cultures (see Table 1). Zinc acetate precursor is another choice for the plant-based ZnO nanoparticle synthesis, and Santoshkumar et al. synthesized ZnO nanoparticles using Passiflora caerulea extract against urinary tract infection pathogens. The ZnO nanostructures have a particle size of around 37 nm and show good zone of inhibition to various pathogens [23]. Hibiscus sabdariffa [24] and Acalypha indica leaves [25] were used in ZnO nanoparticle synthesis, and the nanoparticles showed an enhanced antibacterial activity against E. coli and S. aureus.
\nZnO nanoparticles are currently under investigation due to their utilization in cancer treatment and diagnostic applications [26]. Since the treatment of cancer by chemotherapy is limited because of the adverse effect of tumor drugs and drug resistance by cancer cells, natural plant-based drug researches have focused on overcoming these limitations. Vijayakumar et al. investigated the anticancer activity of ZnO nanoparticles, which were synthesized by Laurus nobilis extract-mediated synthesis. The nanoparticles showed anti-lung cancer activity against human A549 lung cancer cells [27]. Toxicity is an important parameter for the in-vivo and in-vitro activity of nanoparticles because some nanoparticles can generate free radicals even under dark conditions. Anacardium occidentale extract was used in the synthesis of ZnO nanoparticles, and the resulting nanostructures exhibited concentration-dependent cytotoxicity against pancreatic cancer cells [28]. Yuvakkumar et al. utilized the Rambutan peels (Nephelium lappaceum) for the synthesis of ZnO nanoparticles and explored the effect of these particles on HepG2 liver cancer cells [29].
\nOn the other hand, zinc acts as an actuator for several enzymes, and blood sugar regulation is significantly affected in the presence of zinc element. Thus, the enzymatic and anti-diabetic activity of ZnO nanoparticles must be mentioned in their biomedical applications. Bayrami et al. synthesized ZnO nanoparticles by using Vaccinium arctostaphylos extract via a microwave-assisted method. The biosynthesized ZnO nanoparticles showed an enhanced efficiency for the treatment of diabetic problems and reduced the fasting blood glucose level effectively [55]. Rehana et al. demonstrated the antidiabetic activity of ZnO nanoparticles by using several types of plant extracts. The results showed that Tamarindus indica extract-based ZnO nanoparticles exhibited enhanced activity for α-amylase and α-glucosidase due to the presence of amino acids in the plant extract [76]. As an environmentally benign material, coffee powder extract was utilized in the biosynthesis of ZnO nanoparticles. Koupaei et al. studied the reduced effect of ZnO nanoparticles on proteinase K activity [56].
\nPhotocatalytic degradation of organic pollutants is a promising approach for the removal of dyes in wastewaters. ZnO nanoparticles have been involved in photocatalytic applications due to their optical and electronic properties (Table 2). When the ZnO nanoparticles are irradiated with UV light, valence band electrons are excited to the conduction band, which leaves holes behind. Then the generated holes create hydroxyl radicals by oxidizing \n
Plant type | \nZinc precursor type | \nSize of ZnO (nm) | \nTreated analyte | \nEfficiency (%) − time (min) | \nRef. | \n
---|---|---|---|---|---|
Lycopersicon esculentum | \nNitrate | \n9–20 | \nMB | \n97 − 180 | \n[86] | \n
Monsonia burkeana | \nChloride | \n20 | \nMB | \n48 − 45 | \n[87] | \n
Ulva lactuca | \nAcetate | \n10–50 | \nMB | \n90 − 120 | \n[88] | \n
Conyza canadensis | \nNitrate | \n— | \nMO MB | \n94 − 45 85 − 20 | \n[89] | \n
Allium sativum | \nNitrate | \n14–70 | \nMB | \n100 − 180 | \n[90] | \n
Garcinia mangostana | \nNitrate | \n21 | \nMB | \n99 − 180 | \n[91] | \n
Plectranthus amboinicus | \nNitrate | \n50–180 | \nMR | \n92 − 180 | \n[92] | \n
Calotropis procera | \nNitrate | \n15–25 | \nMO | \n81 − 100 | \n[93] | \n
Citrus paradisi | \nSulfate | \n12–72 | \nMB | \n56 − 360 | \n[94] | \n
Lantana camara | \nAcetate | \n340–520 | \nMB RhB | \n92 − 25 75 − 40 | \n[95] | \n
Chlamydomonas reinhardtii | \nAcetate | \n13 | \nMO | \n92 − 120 | \n[96] | \n
Lycopersicon esculentum | \nNitrate | \n7–20 | \nMB | \n97 − 150 | \n[97] | \n
Corymbia citriodora | \nNitrate | \n64 | \nMB | \n83 − 90 | \n[98] | \n
Catharanthus roseus | \nAcetate | \n38 | \nPR | \n100 − 480 | \n[99] | \n
Photocatalytic activities of plant-extract based synthesized ZnO nanoparticles (MB = methylene blue, MO = methyl orange, MR = methyl red, RhB = Rhodamine B, and PR = phenol red).
Schematic diagram of dye degradation by ZnO nanostructures.
Nava et al. addressed the effect of different amounts of Camellia sinensis extract on the synthesis of ZnO nanoparticles. The synthesized nanoparticles were studied in photocatalytic degradation of methylene blue (MB) dye where the nanoparticles presented MB degradation over 84% in 120 min [81]. In another study, Parkia roxburghii extracts have been used for the synthesis of ZnO nanoparticles, and they were found to be efficient in degradation with nearly 98% efficiency in 8 min for both MB and Rhodamine B (RhB) dyes [82]. The degradation of Congo Red dye has also been studied for the ZnO photocatalysis applications. Prasad et al. and Vidya et al. studied the degradation of Congo Red in aqueous solutions by ZnO nanoparticles. The dye was degraded with 90% efficiency in 35 and 60 min by using lemon juice [83] and Artocarpus heterophyllus [84] extracts in the nanoparticle synthesis, respectively. The aqueous leaf extract of Coriandrum sativum was used for the nanoparticle synthesis and the resulting materials have been used for the photocatalytic degradation of anthracene with 96% efficiency in 240 min [85].
\nHeavy metal ion pollutants in wastewaters create a problem worldwide because of their serious effects on both human health and environment. ZnO nanostructures have also been used as an adsorbent material due to their low toxicity and became more effective than the other adsorbent materials [100]. The plant-based synthesis of ZnO nanostructures enhances metal ion adsorption capacity due to the chemical interactions between ions and functional groups of plant extracts. Fazlzadeh et al. synthesized ZnO nanoparticle-loaded activated carbon by using Peganum harmala for the removal of chromium from aqueous systems. Peganum harmala acted as a stabilizing agent and enhanced the chromium uptake up to 68.48 mg g−1 [101]. The lead ion removal was studied by Azizi et al. using Zerumbone extract-mediated ZnO nanoparticles. They reported that the lead ion adsorption capacity reached up to 19.65 mg g−1 [100]. Sensing application of ZnO nanostructures is another field where the glucose sensing mechanism of ZnO nanostructures was studied by Muthuchamy et al. They fabricated a glucose sensor using ZnO nanoparticles and peach juice as a carbon source. The fabricated ZnO sensor showed high sensitivity (231.7 μA mM−1 cm−2) and low detection limit (6.3 μM) [102]. Sharma et al. also studied silymarin detection capability of ZnO nanostructures by using Carica papaya extract. The results showed that ZnO-modified sensors have 2-fold greater electrochemical signals than the neat ones [103].
\nSynthesis of ZnO nanostructures by using microorganisms has gained considerable interest, and numerous microorganisms can be utilized for their synthesis. Bacteria, fungus, and algae are the possible microorganisms in a green synthesis of ZnO nanostructures. Because of their easy genetic manipulation and easy handling, bacteria are preferred microorganisms [104]. Jayaseelan et al. used Aeromonas hydrophila bacteria as green capping agent, and the synthesized ZnO nanoparticles showed antibacterial activity against Pseudomonas aeruginosa and Aspergillus flavus [104]. In another study, Pseudomonas aeruginosa was used as a capping agent in ZnO nanoparticle synthesis, and the resulting particles demonstrated antioxidant activity [105]. Kundu et al. used Rhodococcus pyridinivorans as metabolically versatile Actinobacteria in the fabrication of self-cleaning, UV-blocking, and antibacterial textile fabrics with ZnO nanoparticles. Besides, the ZnO nanoparticles showed photocatalytic activity against malachite green and anticancer activity against HT-29 cancerous cells [106]. Tripathi et al. synthesized ZnO nanoflowers by using Bacillus licheniformis and assessed their photocatalytic activity against methylene blue [107]. However, bacterial utilization in ZnO green synthesis could be somewhat problematic because of the uncontrolled growth of bacteria and unavoidable contaminations [108].
\nFungus-based green synthesis of ZnO nanoparticles is generally preferred over the bacteria based synthesis because of their large-scale production and better tolerance property [109]. Raliya et al. synthesized ZnO nanoparticles via an environmental method by using Aspergillus fumigatus as a stabilizing agent and investigated the effect of ZnO nanoparticles on phosphorus mobilizing enzymes in rhizosphere and gum contents in clusterbean grains [110]. In another study, ureolytic bacterium (Serratia ureilytica)-mediated green synthesis of ZnO nanoparticles was reported. The cotton fabrics were coated with the synthesized nanoparticles and their killing efficiency against S. aureus and E. coli bacteria was revealed [111].
\nAlgae are the members of a diverse group of aquatic photosynthetic organisms and they have been utilized sometimes in the synthesis of ZnO nanostructures. Sargassum muticum extract, which is a brown marine macroalga, was used in the biosynthesis of ZnO nanoparticles [112]. Nagajaran et al. used the seaweed extracts of green Caulerpa peltata, red Hypnea Valencia and brown Sargassum myriocystum in the synthesis of ZnO nanoparticles. The results revealed that among three seaweeds, only S. myriocystum could stabilize and reduce ZnO nanoparticles of size 36 nm. Also, the nanoparticles showed antimicrobial activity against a wide spectrum of bacterial cultures [109].
\nSynthesis of ZnO nanoparticles with controlled morphologies and using environmentally friendly chemicals could be possible in biomolecule-based synthesis routes, and utilization of amino acids, polysaccharides, gums, and enzymes is highly preferable. Gharagozlou et al. synthesized ZnO nanoparticles by using alanine amino acid, and a Schiff base complex was obtained at the end of the study [113]. Bovine skin gelatin has also been used in the synthesis of ZnO nanostructures, and Alnarabiji et al. demonstrated the environmentally friendly synthesis route for ZnO nanoparticles [114]. Arabic gum and gum tragacanth-based green synthesis of ZnO nanoparticles have been demonstrated by Fardood et al. [115] and Daraoudi et al. [116], respectively. Thus, they demonstrated an alternative method for the synthesis of ZnO nanoparticles instead of conventional ZnO reduction methods by using hazardous polymers or surfactants [115, 116]. Casein is another biomolecule that can be used as a capping and reducing agent in the ZnO nanoparticle synthesis. Somu et al. synthesized ZnO nanoparticles, which show heavy metal ion adsorption, dye adsorption, and antibacterial activity in wastewater treatment at the same time [117]. The resulting nanoparticles effectively remove Cd(III), Pd(II), and Co(II) ions, methylene blue, and Congo red dye from the wastewaters. Also, they demonstrated high antibacterial activity against E. coli cultures. Subramanian et al. synthesized ZnO nanoflowers, which comprise nanorods and ellipsoids as subunits, by using
Combating the major drawbacks of common ZnO nanostructure synthesis methods, mainly identified as the generation of pollutants, toxic materials, and side products during reactions, green chemical techniques using only nontoxic and biologically compatible materials were developed. Gharagozlou et al. [113] reported a novel method to synthesize ZnO nanoparticles without any pollutant or combustible side product in the process. Water was used as a solvent with a biologically compatible nitrogen source, amino acid instead of toxic amines, alanine and sodium salicylaldehyde-5-sulfonate, and zinc acetate to prepare the zinc Schiff-base complex and then subsequently heated to obtain ZnO nanoparticles. This work showed that the solid-state decomposition process applied at moderate temperatures has yielded nanoparticles ranging from 5 to 110 nm with fewer defects yet interestingly high crystallinity.
\nBiomimetic and bioinspired synthesis has also been regarded among the most attractive strategies in fabricating novel functional materials, and biological materials like eggshell membrane, oyster shells, nacre, diatoms, cuttlefish bone, DNA chains, and sea urchin spines have been actually employed as templates or bioreactive substrates. Silk fibroin fibers (SFFs) extracted from silkworm Bombyx mori cocoons were used for their capping and directing functions to control the morphology of ZnO crystals. Acting at the same time, zinc ions are anchored on the SFF and in-situ react with \n
Extrusion dripping is another novel technique using environmentally friendly, cost-effective, degradable, and renewable biomass materials. Generally, the process yields monodispersed spherical particles by controlled dripping of working solution into a biopolymer solution after extruding it through a narrow tube, thanks to the effect of viscous-surface tension forces and impact-drag forces that help to preserve the spherical shape of the drop [122]. Goes et al. have reported that spherical uniform sized ZnO nanoparticles were obtained by dropwise addition of alginate solution to zinc nitrate solution under a long slow magnetic stirring to ensure the ion-exchange to happen and stabilize ZnO nanoparticles; the heated ZnO outcome was used to fabricate a polymeric nanocrystalline microfilm, exhibiting interesting photodegradation results, as ZnO on the surface is likely to accept photons and generate holes promoting the oxidative decomposition of the dye [123].
\nChemical bath deposition and soft-template sol-gel methods are two wet organic solvent-free routes studied by a group led by Leone et al [124]. to obtain a nanostructured ZnO employed as a reservoir of clotrimazole for pharmaceutical purposes. Identifying the synthesis of carriers and active pharmaceutical ingredient loading as the main steps in which the waste of organic solvents occur, ZnO nanostructures have been introduced as green alternative carriers for their intrinsic biological properties, low toxicity, and high biocompatibility [125]. For the chemical bath deposition approach, a nanosheet-like zinc carbonate hydroxide hydrate was transformed into ZnO using solutions containing urea and different zinc salts [126]. As for the sol-gel method, pluronic F127 was used as a soft template forming an opalescent solution with zinc acetate in water, and then dried and calcined at 500°C to sacrifice the template and obtain the ZnO nanostructure [124].
\nIn our previous work, we successfully enhanced the antibacterial activity of ZnO nanowires by modifying the cooling route. Zinc acetate was calcined in a muffle oven, followed by a rapid cooling; the three resulting samples were compared to a free cooled batch synthesized under the same conditions, revealing noticeable effects on ZnO nanowire morphology in addition to the improvement of surface area due to the limiting time for crystallite growth [127].
\nIonic liquids (ILs) are an area of chemistry, which has received important attention in both academia and industry, because of the cost effectiveness coupled with being environmentally friendly [128]. ILs usually act as solvents and reactants as well as templates for inorganic nanomaterial synthesis and scavenging agents [129]. As a subdivision, room temperature ionic liquids (RTILs) are particularly doted of special considerations as nontoxic solvents with a wide liquid temperature range, remarkable chemical stability, negligible vapor pressures, and high fire resistance [130]. ILs have been great templates for the synthesis of nanomaterials as it was shown that only by modifying the structure of their cations or anions, it is possible to alter their properties in order to control the size, morphology, and thus the properties of nanomaterials [131]. Sabbaghan et al. have synthesized different morphologies of ZnO nanostructures using zinc acetate as the metal source in a basic media to react with different symmetrical imidazolium-based ILs, yielding nanoparticles, nanoparticle-like, spherical-like, nanosheet in different sizes ranging from 16 to 30 nm and different band gaps between 2.98 and 3.17 eV, demonstrating through this work the relation morphology-IL [128].
\nZnO nanostructures with nanosheet morphology have been successfully fabricated in another work by refluxing the mixture of zinc acetate and the ionic liquid in water according to Menshutkin reaction [132]. A comparative study has shown that when zinc is used with ILs as a template, ZnO nanoparticles with smaller crystallite size were formed compared to the yield without ILs. This work revealed as well that the template and the pH control the direction of growth of ZnO crystals and the shape of nanomaterials obtained. The final band gap values of ZnO with different morphology ranged around 2.88–3.16 eV [133]. Alammar et al. have studied the effect of five different ILs on ZnO morphologies, and claimed that the habitus and morphology come as the system naturally tends to reduce the total surface energy during formation; it is to note that the anion of the IL is proposed to be interacting with the ZnO surface during the growth [134]. Yet, the best performance was for ZnO nanoparticles that are obtained by use of IL with a long alkyl chain, reaching 95% in 9 h for methyl orange decomposition, proposing that along with high surface area, oxygen vacancies and polar plans that act like electron traps are the main factors for such interesting photocatalytic activity [135, 136]. It was also reported by Amde et al. [137] that common techniques for the determination of fungicide concentration in water are usually non-environmentally friendly, organic solvent, and time consuming; the group has prepared ZnO nanofluids by a green two-step method, dispersing the as-synthesized sol-gel ZnO nanoparticles in 1-hexyl-3-methylimidazolium hexafluorophosphate, hand shaking it to attain a homogeneous distribution, then sonicating it to break NPs clusters. The as-prepared ZnO nanofluids are applied in a modified, simple, versatile, and inexpensive liquid-liquid microextraction technique; this technique, called single drop microextraction not only reduces the amount of extraction solvent radically but also offers other functionalities such as high enrichment factor, different extraction modes, and full automation of the process [138]. It is proclaimed that the preferences of ZnO-based nanofluids for the investigation were driven from the fact that ZnO dotes on surface charges that enable to form stable suspensions, unlike many metallic nanofluids, and without the need of any additional stabilizer intervention.
\nZnO nanostructures can be synthesized by following different approaches, and each has its own distinctive advantages and downsides. In this section, we report some interesting methods and findings having the common main aim to avoid drawbacks like the use of toxic reagents, promoter, and stabilizer organic additives, lowering the reaction time, as well as high temperature and pressure. These methods have plenty of scopes to provide both qualitative and quantitative support for nanosized ZnO synthesis along with being simple, fast, efficient, and convenient.
\nHydrothermal method has gained particular interest as an efficient method for high quality and mass production of ZnO nanostructures [139]; it is indeed environmentally friendly as there is no need to control pH and subsequently no release of unwanted by-products. A work led by Guo et al. [140] reported a controllable hydrothermal synthesis of ZnO nanorods reacting with zinc carbonate hydroxide hydrate powder and H2O2 at various temperatures for different periods of time. The group claims that the formation mechanism of ZnO starts by the formation of ZnO2 when subjecting it to hydrothermal treatment at 170°C for more time (3–6 h), and then the thermally unstable ZnO2 would decompose into ZnO and O2. ZnO nanorods exhibited an optical band gap of 3.3 eV [140].
\nIt has also been reported that flower shaped ZnO nanoparticles were synthesized by hydrothermal method, where zinc nitrate and hexamethylenetetramine solutions were prepared separately in double distilled water, while NaOH solution was added dropwise to adjust the pH to 10. The obtained milky solution was refluxed at 80°C for 7 h, washed, and dried. XRD asserted the formation of hexagonal crystal structure ZnO that had a flower-shape, formed by agglomeration during the hydrothermal process. Dynamic light scattering (DLS) data have affirmed the average diameter of ZnO between 600 and 800 nm. The effect of different pH values from 2 to 10 on the removal performance of ZnO has been studied and the results showed a maximum increase of 80% removal efficiency when pH reached 6; increasing the temperature of the process also improved the removal efficiency from 68 to 97%. The contact time had a sharp rise at the value of 15 min of initiation of the experiment, and higher stirring had an enhancing effect as well. Jamal Al-Sabahi and his group have treated for the first time the degradation of HPAM polymer in oil produced water with supported ZnO nanorods synthesized via a microwave-assisted hydrothermal method in an aqueous solution [141, 142, 143]. Placing a prepared microscope glass substrate (25 mm × 75 mm) on a hotplate (350°C), 10 mM zinc acetate dihydrate solution is sprayed, and then the plate is immersed in an equimolar solution of zinc nitrate hexahydrate and hexamethylenetetramine, then heated in a domestic microwave oven for 45 min, and then cooled down for 15 min; the produced ZnO nanorod-covered substrate was afterward annealed in air at 350°C for 1 h. The morphology was of a typical ZnO array and the average length was about 4 μm while the average diameter reached around 95 nm [144].
\nSingh et al. have adopted a cost-effective and environmentally friendly method to fabricate a 3D self-assembled wool ball like spherical ZnO with high porosity by combining a urea-glycerol assisted hydrothermal approach with calcination under air atmosphere [145], where hydrated zinc carbonate was synthesized hydrothermally in a Teflon-lined stainless steel autoclave reacting zinc nitrate with urea in a triplet (glycerol, ethyl alcohol, and water (7:7:10)) solvent system. After drying the outcome, the white powder intermediate product was calcined at 450°C for 3 h to get hierarchical 3D porous ZnO. The group carried out a series of experiments to investigate the effect of synthesis parameters over the morphology, while 7 h and 140°C were about the optimum duration of synthesis and temperature to get the best ZnO with W-ball like spherical morphology. This ZnO photocatalyst has shown 98% of highly toxic Rhodamine B degradation in 60 min of UV photolysis catalysts at a pH of 4.
\nA green hydrothermal method was used recently to fabricate ZnO nanorods without any organic solvent or surfactant, by starting with ZnO powder and H2O2 aqueous solution in the sealed autoclave. Then the precipitates were washed and dried. In this work, Lam et al. reported as well that the hydrothermal treatment of ZnO at 100°C promotes the slow conversion of ZnO2 to ZnO and O2 without using any toxic reactant, nor releasing any pollutant by-product. XRD indicated the high purity of the ZnO wurtzite phase [146]. This environmentally friendly method has generated highly performing ZnO nanorods that completely degrade the resorcinol in aqueous solution after 120 min. The same group has used a similar hydrothermal method to fabricate ZnO nanotubes (NTs) with a diameter of around 10 nm, a wall thickness of 3.5 nm, and average lengths of up to 200 nm by scrolling of the ZnO2 layer nanosheet, which transforms to ZnO NTs with a symmetrical layer in both shell-tube structures. XRD asserted the hexagonal phase and then confirmed the purity of ZnO NTs that showed a ferromagnetic behavior because of the grain boundaries and developed free surfaces [147]; the band gap energy was measured to be around 3.21 eV, and the degradation of methylparaben over the surface of ZnO NTs had an efficiency of 87.6% in 105 min [148].
\nMicrowave-assisted sol-gel synthesis is based on subjecting samples to frequencies ranging from 300 MHz to 300 GHz [149]; besides its high selectivity of specific morphologies, dramatic reduction of reaction time (minutes), and remarkable increase of product yield, it generates localized superheating at the reaction sites promoting metal ion reduction in the solution [150]. It presents a promising green method for metal oxide nanostructures production. Azizi et al. [151] have worked on a green microwave-assisted combustion approach to synthesize ZnO-nanoparticles, presenting combustion as a fast, low cost, homogenous and highly pure outcome, as well a distinguished surface area at low temperature. Using fruit, seed, and pulp extracts of Citrullus colocynthis (L.) as biofuels with zinc nitrate as the zinc source, an in vitro cytotoxicity study has been made showing that smaller nanoparticles were more efficient penetrating in the cells membranes, while the optical band gap increased with the rear of the particle size from 3.25 to 3.40 eV. An interesting nontoxic and eco-friendly single-step, and green synthesis method of ZnO nanoparticles with excellent reproducibility was reported using coffee powder extract as a reducing material under microwave heating at 540 W for 5 min and the precipitate was dried in a hot air oven. SEM asserted a size of 80–120 nm for the nanoparticles. Afterward, a thin nanocomposite film was prepared using the as-prepared ZnO nanoparticles with natural graphite powder. The nanocomposite film showed a remarkable photovoltaic efficiency of 3.12% [152]. ZnO sub-micrometer particles and nanowires were synthesized by microwave assisted sol-gel reaction. Zinc acetate and N,N-dimethylacetamide were stirred into a beaker, and then, the solution was cooled rapidly to 15°C. After only a couple of minutes of the microwave, a white suspension was obtained. The average diameter of the particles prepared given by DLS analyses was around 275–352 nm [153]. Salari et al. reported a microwave-assisted synthesis of biogenic nanoparticles using Lavandula vera leaf extract as a reducing agent in the presence of zinc sulfate; the method led to simple and fast formation of microstructures that exhibited high antioxidant cytotoxic activity [154]. In a separate work, vertically aligned ZnO nanorods were grown at 90°C on a Si substrate by microwave synthesis and compared with nanorods made by the traditional heated waterbath method changing the pH from 10.07 to 10.9. The microwave synthesis was performed at a power of 100 W [155]. The same group observed that the increase of ammonia led in both methods to sparser and longer nanorods with the larger diameter, as well as an increase of oxygen percentages in the samples. The microwave synthesized samples exhibit a uniform distribution of nanorods as well as a better crystalline structure with fewer defects than the heated water bath-grown samples, which can be beneficial for band-edge transition optoelectronic devices.
\nMicrowave synthesis and ultrasound sonication have proven numerous advantages and actually become amid the most frequently sought nanosized material synthesis methods [156, 157]. Zak et al. have used ultrasonication to synthesize ZnO nanostructures at room temperature without any specific conditions or organic solvents, starting from an as-prepared zinc solution where zinc acetate was dissolved in ammonia solution, and sodium hydroxide was dropped in the solution. Deionized water was wisely added till attaining a concentration of 1 M zinc. The ultrasonication performed at different durations was sufficient to stimulate the formation of nanostructures. XRD asserted the presence of hexagonal pure ZnO and the band gap energies estimated by UV-Vis spectra are 3.3, 3.22, and 3.2 eV for ZnO seeds, nanorods, and nanoflowers, respectively [158]. Ultrasound was conducted as well for the synthesis of different ZnO nanostructures without any organic solvents, surfactants, or templating agents; zinc acetate and sodium hydroxide in ionic liquids (ILs) are reported to be a green, fast, and effective, yet highly selective route to 0D, 1D, and 2D nanostructures of ZnO [134]. A facile calcination-free ultrasound assisted approach has been reported by Bhatte et al. involving zinc acetate as a metal source and 1, 3-propane diol as a solvent, base, stabilizer, and template for the growth of nanocrystalline ZnO [159]. The mixture of both materials has been sonicated under 22 kHz frequency for 2 h with a 5 s interval on-off pulse. After sonication, the formed ZnO was collected, washed and dried. XRD confirmed the successful formation of ZnO without any impurities [159].
\nFor the large-scale production of pure ZnO nanocrystallites, low thermal processes are one of the most efficient methods minimizing the generated waste yet implementing sustainable processes: simple, cheap, and nonpolluting, addressing the key issues that draw much consideration in a green solid-state synthetic method, by eliminating the use of nontoxic materials and reducing energy consumption. ZnO2 nanocrystallites were employed as the precursor for ZnO production, because of their facile preparation, the absence of unwanted by-products, and low-temperature decomposition reaction [160]. Zinc acetate and hydrogen peroxide were used first to synthesize ZnO2 nanocrystallites hydrothermally at 100°C for 12 h in an alkaline aqueous solution; the product was subjected to 180°C in air for 12 h yielding pure ZnO phase shaped nanocrystallites of 8–10 nm and blue shift at around 350 nm in the UV-Vis spectra [161].
\nZnO nanosized structures were also synthesized starting from zinc acetate and urea in a 1:1 stoichiometry, where the decomposition of urea helped the formation of ZnO. Then two different heating methods were applied: microwave hydrothermal (MH) method and waterbath heating. XRD proved the purity of the wurtzite phase for both methods, while FE-SEM showed a difference in shape regularity in favor of the MH process; thus, the MH method contributes to the production of spherical and uniform particles after a short processing time by enhancing the interface mobility and the diffusivity in the medium [162].
\nRaja et al. have reported a laboratory procedure based on sol-gel for the preparation of nano-ZnO particles. Zinc acetate solution was stirred at room temperature while adding sodium hydroxide until reaching a pH of 14 and the solution went through a microemulsion. The suspension obtained was transferred for thermal treatment at 180°C for 3 h, and then the white precipitate was collected, washed, centrifuged, and dried under vacuum to reveal well-shaped uniform ZnO nanoparticles of 35 nm on average [163].
\nAnother group has proposed a simple, eco-friendly approach to synthesize ZnO nanoparticles by using carboxylic curdlan (cc) as a reducing and stabilizing agent. A solution of zinc acetate was blended with cc aqueous solution and stirred at 70°C for 6 h, and then the outcome was freeze-dried [164]. The carboxyl group is in charge of chelating and reducing zinc ions for the sake of the formation of ZnO nanoparticles, thanks to the numerous negatively charged carboxyl groups it contains. The average diameter of cc-ZnO nanoparticles is around 58 nm exhibiting a band gap energy of 3.3 eV. Furthermore, the interaction between the as-prepared nanoparticles and bovine serum albumin (BSA) at room temperature was investigated, which suggested the formation of a certain complex revealed by a blue shift of the fluorescence peak by about 8 nm with increasing nanoparticle concentration, due to the binding of cc-ZnO nanoparticles and BSA [164].
\nThe authors declare no conflict of interest.
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",metaTitle:"Conflicts of Interest Policy",metaDescription:"As an Open Access publisher, IntechOpen is dedicated to maintaining the highest ethical standards and principles in publishing. In addition, IntechOpen promotes the highest standards of integrity and ethical behavior in scientific research and peer-review.",metaKeywords:null,canonicalURL:"/page/conflicts-of-interest-policy",contentRaw:'[{"type":"htmlEditorComponent","content":"In each instance of a possible Conflict of Interest, IntechOpen aims to disclose the situation in as transparent a way as possible in order to allow readers to judge whether a particular potential Conflict of Interest has influenced the Work of any individual Author, Editor, or Reviewer. IntechOpen takes all possible Conflicts of Interest into account during the review process and ensures maximum transparency in implementing its policies.
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\n\nA Conflict of Interest is a situation in which a person's professional judgment may be influenced by a range of factors, including financial gain, material interest, or some other personal or professional interest. For IntechOpen as a publisher, it is essential that all possible Conflicts of Interest are avoided. Each contributor, whether an Author, Editor, or Reviewer, who suspects they may have a Conflict of Interest, is obliged to declare that concern in order to make the publisher and the readership aware of any potential influence on the work being undertaken.
\n\nA Conflict of Interest can be identified at different phases of the publishing process.
\n\nIntechOpen requires:
\n\nCONFLICT OF INTEREST - AUTHOR
\n\nAll Authors are obliged to declare every existing or potential Conflict of Interest, including financial or personal factors, as well as any relationship which could influence their scientific work. Authors must declare Conflicts of Interest at the time of manuscript submission, although they may exceptionally do so at any point during manuscript review. For jointly prepared manuscripts, the corresponding Author is obliged to declare potential Conflicts of Interest of any other Authors who have contributed to the manuscript.
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\n\nEditors can also have Conflicts of Interest. Editors are expected to maintain the highest standards of conduct, which are outlined in our Best Practice Guidelines (templates for Best Practice Guidelines). Among other obligations, it is essential that Editors make transparent declarations of any possible Conflicts of Interest that they might have.
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I am also a member of the team in charge for the supervision of Ph.D. students in the fields of development of silicon based planar waveguide sensor devices, study of inelastic electron tunnelling in planar tunnelling nanostructures for sensing applications and development of organotellurium(IV) compounds for semiconductor applications. I am a specialist in data analysis techniques and nanosurface structure. 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