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Role of microRNA in zinc depletion-induced growth inhibition in mouse fibroblast 3T3 cells He, Li 2014

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    ROLE OF MICRORNA IN ZINC DEPLETION-INDUCED GROWTH INHIBITION IN MOUSE FIBROBLAST 3T3 CELLS   by  LI HE  B.Sc., The Huazhong Agricultural University, 2009 M.Sc., The South China University of Technology, 2012     A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE   in   THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES   (Human Nutrition)     THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)     August 2014    © Li He, 2014 ii  Abstract  Zinc, an essential trace element for humans, exerts many physiological functions, including its indispensible role in growth. Dietary zinc deficiency in children results in growth retardation. However, the mechanism whereby zinc regulates growth remains unclear. MicroRNAs are a group of newly discovered, small, non-coding RNAs and have been demonstrated to play a regulatory role in cell proliferation. The expression of microRNA and process of producing mature microRNA can be influenced by cellular and tissue zinc status. Thus, the hypothesis for my research is that microRNA plays a role in mediating zinc-dependent cell proliferation. The overall objective was to determine whether microRNAs are involved in zinc-dependent cell proliferation in mouse fibroblast 3T3 cells. 3T3 cells were cultured in Dulbecco’s Modified Eagle Medium with 10% fetal bovine serum (FBS) for 3 days. To deplete intracellular zinc, cells were cultured in the same media containing 5% (v/v) dimethyl sulfoxide (DMSO, control) or N,N,N’,N’- tetrakis(2-pyridylmethyl)ethylenediamine (TPEN, 2.5 μM) for 24 or 48 h with or without induction of quiescence (aphidicolin, 0.5 μg/ml; 24 h). To establish zinc-dependent effect, zinc was replenished at the final concentration of 0 (TPEN only), 1.25, 2.5 or 5 μM for 24 or 48 h. Cell proliferation was measured by cell cycle analysis using flow cytometry. MicroRNA expression profile was assessed by microRNA microarray. In the absence of quiescence induction, zinc-depletion for 24 or 48 h inhibited cell proliferation by 10.4% and 16.0% compared to control. In the presence of quiescence induction, zinc-depletion inhibited cell proliferation by 52.9%. Regardless of the status of quiescence, zinc replenishment at 1.25 μM nearly brought cell proliferation back to the level iii  observed in the DMSO control, showing a zinc-dependent cell proliferation in 3T3 cells. Zinc-depletion increased the abundance of miR-132-3p, miR-212-3, and let-7e-3p, while zinc replenishment brought back the abundance of these three microRNAs to the level observed in the DMSO control. Interestingly, zinc-depletion decreased the abundance of miR-145b, and its abundance was increased after zinc replenishment. Overall, it appeared that microRNA played a role in zinc-mediated growth regulation in 3T3 cells; however, this role of microRNA remains to be affirmed by further investigation.          iv  Preface  This dissertation presents the findings of my master’s thesis research study and was prepared in accordance with the requirements of the University of British Columbia, Faculty of Graduate Studies. I was mainly responsible for the development of the zinc-depletion model used in this thesis research. All the experiments were designed and carried out by Li HE under the guidance and supervision of Dr. Zhaoming Xu. LC Science carried out the miRNA microarray profiling experiment, as well as assisted with subsequent data analysis.   v  Table of Contents  Abstract ..................................................................................................................................... ii Preface ..................................................................................................................................... iv Table of Contents ...................................................................................................................... v List of Tables .......................................................................................................................... vii List of Figures ........................................................................................................................ viii List of Abbreviations ............................................................................................................... ix Acknowledgements................................................................................................................. xii Introduction............................................................................................................................... 1 Chapter 1 Literature Review ..................................................................................................... 3 1.1 Zinc and zinc nutrition .................................................................................................... 3 1.1.1 Introduction............................................................................................................... 3 1.1.2 Food sources and bioavailability .............................................................................. 3 1.1.3 Physiological functions ............................................................................................. 5 1.1.4 Zinc homeostasis ...................................................................................................... 7 1.2 Zinc and growth ............................................................................................................ 12 1.3 Zinc and cell proliferation ............................................................................................. 14 1.3.1 Cell proliferation ..................................................................................................... 14 1.3.2 Role of zinc in cell proliferation ............................................................................. 16 1.4 MicroRNAs and cell proliferation ................................................................................ 21 1.4.1 miRNA .................................................................................................................... 21 1.4.2 miRNA biogenesis .................................................................................................. 23 1.4.3 Functions of miRNA............................................................................................... 25 1.4.4 miRNA and cell proliferation ................................................................................. 27 1.5 Zinc and miRNA ........................................................................................................... 31 1.6 Hypothesis ..................................................................................................................... 33 1.7 Overall objective and specific aims .............................................................................. 34 Chapter 2 Materials and Methods ........................................................................................... 35 2.1 Cell culture system ........................................................................................................ 35 2.2 Zinc depletion and zinc replenishment ......................................................................... 35 2.3 Quiescence induction .................................................................................................... 36 2.4 Cell cycle analysis ......................................................................................................... 36 vi  2.5 Total RNA isolation ....................................................................................................... 38 2.6 miRNA microarray assay .............................................................................................. 39 2.7 Statistics ........................................................................................................................ 41 Chapter 3 Results and Discussion........................................................................................... 42 3.1 Results ........................................................................................................................... 42 3.1.1 TPEN-induced zinc depletion caused a G1/S arrest ................................................ 42 3.1.2 TPEN-induced zinc depletion altered miRNA expression profile .......................... 44 3.2 Discussion ..................................................................................................................... 45 Chapter 4 Conclusions, Limitations, and Future Directions .................................................. 63 4.1 Conclusions ................................................................................................................... 63 4.2 Limitations .................................................................................................................... 64 4.3 Future directions ............................................................................................................ 65 Bibliography ........................................................................................................................... 68 Appendix................................................................................................................................. 96     vii  List of Tables  Table 3.1 Effects of zinc-depletion and zinc replenishment on cell cycle progression in 3T3 cells without the induction of quiescence*…………………...…………….…….59 Table 3.2 Effects of zinc-depletion and zinc replenishment on cell cycle progression in 3T3 cells upon the induction of quiescence*…………………...………………….….60 Table 3.3 Differential expression of miRNAs induced by 24 h zinc-depletion in 3T3 cells*…………………………………...……………………………………..…..61 Table 3.4 Differential expression of miRNAs induced by 24 h zinc replenishment in 3T3 cells*……………………………………………...……………………..………..62 Table A.1 Differential expression of miRNAs induced by 24 h zinc-depletion in 3T3 cells*…………………………………………...……………………..…………..96 Table A.2 Differential expression of miRNAs induced by 24 h zinc replenishment in 3T3 cells*……………………………………………...…………………………..…101   viii  List of Figures Figure 3.1 Effects of 24 h zinc-depletion and zinc replenishment on cell cycle progression in 3T3 cells without quiescence induction……………………….……………...…..52 Figure 3.2 Effects of 48 h zinc-depletion and zinc replenishment on cell cycle progression in 3T3 cells without quiescence induction…………………..………………...…….53 Figure 3.3 Effects of 24 h zinc-depletion and zinc replenishment on cell cycle progression in 3T3 cells upon quiescence induction……………………..……….…………..….54 Figure 3.4 Effects of zinc-depletion and zinc replenishment on the abundance of miR-132-3p in 3T3 cells…………………………………………………………..…...………55 Figure 3.5 Effects of zinc-depletion and zinc replenishment on the abundance of miR-212-3p in 3T3 cells…………………………………..……….………………………..…56 Figure 3.6 Effects of zinc-depletion and zinc replenishment on the abundance of let-7e-3p in 3T3 cells……………………………………..…………………………………...57 Figure 3.7 Effects of zinc-depletion and zinc replenishment on the abundance of miR-145b in 3T3 cells………………………………………………..……………………...…58    ix  List of Abbreviations  Ago Argonaute Ap4A Adenosine(5’)tetraphosphate(5’)adenosine  BCL2 B-cell lymphoma 2 gene BrdU 5-Bromo-2’-deoxy-uridine  DCIS Ductal carcinoma in situ  DGCR8 DiGeorge syndrome critical region gene 8  DMEM Dulbecco’s Modified Eagle’s Medium  DMSO Dimethyl sulfoxide  DTPA Diethylene triamine pentaacetic acid  EDTA  Ethylenediaminetetraacetic acid EGF Epidermal growth factor  ERK Extracellular-signal-regulated kinase FBS Fetal bovine serum  HEK-239 Human embryonic kidney 293 cells HOS Human osteosarcoma  HSD Honestly Significant Difference  IGF-1 Insulin-like growth factor 1  JNK Jun N- terminal kinase  LIPZ Labile intracellular pool of zinc LOWESS  Locally-weighted Regression MAPK Mitogen-activated protein kinases  x  MAPKK MAPK kinase  MAPKKK MAPK kinase kinase  MCF-7 Michigan Cancer Foundation-7 miRNAs MicroRNAs MRE Metal response elements  MTF-1 Metal-responsive-element binding transcription factor-1  MTs Metallothioniens  PBS Phosphate buffered saline  PC-3 Prostate cancer cell line-3 PI Propidium iodide  PIWI P-element induced wimpy testis in Drosophila pre-miRNA precursor miRNA  pri-miRNA primary miRNA  qRT-PCR Quantitative real-time PCR  Ran-GTP RAs-related Nuclear protein-guanosine-5’-triphosphate RBP2 Retinoblastoma binding protein 2  RDA Recommended Dietary Allowance RISC RNA-induced silencing complex  RNase Ribonuclease Saos-2 Sarcoma osteogenic-2 SGC7901 Gastric cancer cell 7901 SLC Solute-like carrier  xi  SOD Superoxide dismutase  T Thionein TPEN N,N,N’,N’-tetrakis(2-pyridylmethyl)ethylenediamine  UTR Untraslated region  xii  Acknowledgements   First of all, I would like to acknowledge my supervisor Dr. Zhaoming Xu for his patience, guidance, and encouragement during my entire program. I want to sincerely thank him for giving me the opportunity to study in his laboratory. In addition, I would like to thank my committee members Dr. Thomas Chang and Dr. Kevin Allen for their insightful advice and feedback on my research work, and Dr. Tim Green for his feedback on my thesis writing. I also want to thank my lab-mates, Alice Lin, Wendy Hempstock, and Melinda Bakker, as well as co-op students Joseph Tio for their support and help. I feel lucky to have you as colleagues and friends. I am very happy to know you and spend time together with you. I wish you all the best for everything in future.  Secondly, I want to express my gratitude to the Biochemical Research Centre, the University of British Columbia to offer me the access to the flow cytometry equipment for cell cycle analysis. Especially, I would like to thank Andy Johnson for his training and instructions on the flow cytometry.  Last but not the least, I want to thank my family and friends. Thanks to all of you being there for me, whenever I needed you.   1  Introduction  Zinc is an essential trace element required for human. One of the physiological functions of zinc is its indispensible role in growth (MacDonald, 2000). Dietary zinc deficiency in children results in growth retardation (Vallee and Falchuk, 1993).  Numerous studies have documented the essential role of zinc in cell proliferation and the suppressed cell proliferation caused by zinc deficiency. The previous work carried out in our laboratory has showed that DNA synthesis and cell proliferation were negatively impacted when mouse fibroblast 3T3 cells were cultured in a low-zinc medium. When the cells were replenished with zinc, DNA synthesis and cell proliferation are significantly elevated along with a significant increase in the abundance of labile intracellular pool of zinc (LIPZ; Paski and Xu, 2001). Available evidences suggest that zinc may influence cell proliferation at the level of DNA synthesis, cellular signalling pathways, and zinc-containing regulators. The exact mechanism whereby zinc regulates cell proliferation and growth remains unknown.  MicroRNAs (miRNAs) are a group of newly discovered, small, and non-coding RNAs that regulate gene expression at the post-transcriptional level (Cai et al., 2009). Although the physiological functions of the majority of miRNAs remain undefiend, miRNA has been demonstrated to play a regulatory role in cell proliferation, with either proliferative or anti-proliferative effects depending on the forms of miRNAs involved.  To date, very few studies have investigated the effects of zinc status on miRNA expression. It appears that zinc status can influence miRNA expression both in vivo (Ryu et al., 2011; Alder et al., 2012) and in vitro (Zheng et al., 2012; Hlavna et al., 2012). Thus, miRNAs might play a role in the zinc-mediated cell proliferation. The correlation between zinc status and miRNA expression reveals a potential relationship among zinc 2  nutrition, miRNA expression, and cell proliferation. This study is aimed to investigate the possible involvement of miRNAs in the zinc-dependent cell proliferation in 3T3 cells. Results obtained from this study can contribute to our understanding of the mechanisms involved in zinc-dependent cell proliferation.   3  Chapter 1 Literature Review  1.1 Zinc and zinc nutrition  1.1.1 Introduction  Zinc is an essential trace mineral for humans. The average amount of zinc in human adults is 2-3 grams with 0.1% of which are replenished daily (Stefanidou et al., 2006). Dietary requirement for zinc in humans varies with ages, genders and physiological states. The current Recommended Dietary Allowance (RDA) for zinc in adults is 8 mg/day for women and 11 mg/day for men (Institute of Medicine, 2001). In the human body, zinc is present as divalent cation (Zn2+).   1.1.2 Food sources and bioavailability  Zinc in foods is usually associated with proteins. Zinc content varies between animal- and plant-based foods. Generally, animal-based foods are the better sources of dietary zinc (Groff and Gropper, 2000). For example, red meat such as ground beef is rich in zinc with a content of 3.9-4.1 mg/100 g. Organ meat containing approximate 3.1-3.9 mg zinc per 100 g, is also a good source of zinc (Government of Canada, 2010). Seafood generally is an excellent source of zinc. For instance, 100 g of oyster contains 17-91 mg zinc, which is considerably higher than other sources (Groff and Gropper, 2000). Other animal-based sources, such as white meat and dairy products contain moderate amounts of zinc. For example, cooked turkey has a zinc content of 1.1-3.6 mg zinc/100 g while zinc content in milk is 1.0-1.1 mg zinc per 250 mL (Government of Canada, 2010). In 4  contrast, plant-based foods are poor source of zinc (Groff and Gropper, 2000). Even though moderate amounts of zinc is found in whole grains and legumes, such as whole wheat bread (1 mg zinc/100 g) and peas (1.5 mg zinc/100 g; Government of Canada, 2010), the bioavailability of zinc in plant-based foods is relatively low because of the presence of absorption inhibitors (Maret and Sandstead, 2006). Bioavailability of zinc is influenced by other food constituents. Zinc enhancers, such as amino acids and proteins, can increase the bioavailability of zinc (Solomons, 1982). Generally, sulphur-containing amino acids, cysteine and methionine, and nitrogen-containing amino acid such as histidine, have greater effects on facilitating zinc absorption compared to other amino acids (Lonnerdal, 1989). The type of protein in a meal also affects zinc bioavailability. Animal protein such as beef, pork, and eggs, has been shown to promote zinc absorption due to amino acids released from the protein which keep zinc soluble. Casein in milk has been shown to have a negative effect on zinc absorption, since the phosphorylated serine and threonine residues in undigested casein subunits can bind zinc and reduce zinc bioavailability. That is the reason why animal-based foods, which are high in protein content, are generally good sources for dietary zinc.  In contrast, the low zinc bioavailability in plant-based foods is attributed to the presence of non-digestible ligands, so-called zinc inhibitors, such as phytate, oxalate, polyphenols and fiber (Groff and Gropper, 2000). These inhibitors can form insoluble complexes with zinc, resulting in low zinc bioavailability. For example, the predominant presence of zinc in legumes is phytate-zinc. Oxalate, which is present in vegetables such as spinach, also can bind zinc and decrease the availability of zinc. Polyphenols, are able to bind zinc through phenolic groups and the amount of zinc-binding phenolic galloyl groups in foods roughly corresponds to the extent of inhibition of zinc absorption. For 5  example, beans containing various amounts of polyphenols were demonstrated to possess different inhibitory effects on zinc absorption (Solomons, 1986; Groff and Gropper, 2000). Therefore, zinc bioavailability is generally lower in plant-based food sources, including fruits and vegetables, than that in those animal-based foods. Furthermore, presence of some other minerals in foods can also reduce zinc bioavailability. For example, calcium can reduce zinc absorption, because calcium augments the inhibition of zinc absorption by phytate (Oberleas et al., 1966). High concentration of ferrous iron supplementation can also possibly decrease zinc bioavailability (Solomons, 1986). The bioavailability of zinc supplementation varies widely from very low, such as zinc oxide, to relatively high, such as zinc salts (i.e. zinc acetate; Maret and Sandstead, 2006).  1.1.3 Physiological functions  Zinc is required for a wide range of physiological functions. One of the most profound physiological functions of zinc is its role in growth. Zinc deficiency results in growth retardation in all species studied to date (Vallee and Falchuk, 1993). Clinical zinc deficiency in humans was first described in 1961, when the consumption of diets with low zinc bioavailability due to high phytic acid content was associated with adolescent nutritional dwarfism in the Middle East (Prasad et al., 1996). Zinc is also involved in many other physiological functions such as reproduction, immune system, behaviour, taste, and skins, etc (Maret and Sandstead, 2006). The symptoms of zinc deficiency include poor reproductive performance in male, immune dysfunction, cognitive impairment, loss of appetite, and skin lesions (Chasapic et al., 2012). In addition, zinc deficiency may also lead to poor healing of wounds (Henzel et al., 1970) and hair loss (Maret and Sandstead, 2006). 6  Zinc is present in a large number of proteins in the body. Almost 10% of the genes encode zinc-containing proteins, corresponding to about 3,000 zinc proteins in human (Andreini et al., 2006). There are three classes of zinc-containing proteins: zinc metallothioniens (MTs), zinc metalloenzymes, and zinc-finger proteins.  MTs, is a family of cysteine-rich, low molecular weight proteins (500 to 14,000 Da). MTs are capable of binding to zinc and exert as the intracellular zinc storage sites (Tapiero and Tew, 2003). MTs also play a critical role in controlling oxidative stress by chelating oxidatively active metals such as iron and copper, and scavenging hydroxyl radicals through cysteine residues (Kumari et al., 1998).  There are over 300 zinc-containing enzymes, which are collectively known as zinc metalloenzymes (Vallee and Auld, 1990). These zinc metalloenzymes are found in every enzyme classification, indicating an indispensible role of zinc in normal enzymatic function (Vallee and Falchuk, 1993). Zinc plays a catalytic, co-activator and structural role in these enzymes, which are involved in a wide range of physiological activities including DNA and RNA syntheses, cell proliferation and apoptosis, energy metabolism, cellular antioxidant defense, brain development, bone formation and immunity (Macdonald, 2000; Beyersmann and Haase, 2001; Stefanidou et al., 2006). Exerting its catalytic role, zinc ion directly participates in the bond-making or breaking step. For example, zinc is catalytically required for DNA and RNA polymerase (Prasad and Oberleas, 1974). Being a co-activator, there are several metals involving in enzyme activities, where one plays a catalytic role and the zinc ion enhances the catalytic activity. For example, Cu/Zn superoxide dismutase (SOD), an antioxidant enzyme that is involved in superoxide anion radicals’ removal, contains a cooper ion in its catalytic site while zinc is present at co-catalytic site (Vallee and Auld 1993; Tapiero and Tew, 2003). Finally, to exert its structural role, zinc ion stabilizes the structure of the enzymes in a manner 7  analogous to disulfide bonds and removal of the zinc leads to the loss of enzyme activities. For example, aspartate transcarbamoylase, which catalyzes the first step in the pyrimidine biosynthetic pathway, contains a single structural zinc atom (Simmer et al., 1990), which locates in the regulatory subunit and binds tetrahedrally to four cysteines (Vallee and Auld, 1992). Moreover, recently free zinc ion (Zn2+) has been identified to directly serve as a second messenger in some cellular signalling transductions (Yamasaki et al., 2007). In zinc-finger proteins, zinc is responsible for the formation of the zinc-finger motif, which is a small protein structural motif. Zinc-finger motif is characterized by the coordination of one or more zinc ions to stabilize the folding (Klug, 2010). Zinc is most commonly bound to four cysteine residues, or two cysteine and two histidine residues (Andreini et al., 2011). Zinc-finger proteins are zinc-containing transcriptional factors which are directly involved in the expression of the genes related to numerous cellular activities including cell proliferation and apoptosis (Dreosti, 2001).    1.1.4 Zinc homeostasis  Homeostasis is defined as the nutrient flow within a living organism in a state of equilibrium (Kirchgessner, 1993). Maintaining a constant internal environment, under varying external conditions, is critical for survival. As stated above, zinc is involved in numerous physiological functions. Therefore, achieving zinc homeostasis is essential for normal physiological functions.  8  1.1.4.1 Cellular zinc homeostasis  Quantitatively, most of the body zinc is present intracellularly (95-99%) (Vallee and Falchuk, 1993). Zinc deficiency is associated with a range of pathological conditions while excessive zinc is also toxic. The abnormal fluctuation of cellular zinc content, resulted from zinc deficiency or excessive rise, could have catastrophic consequences for the cellular physiological functions. Thus, cellular zinc status and its cellular distribution are tightly regulated to provide sufficient zinc for exerting its functions. Normal intracellular zinc concentration is approximately 100-500 μM (Stefanidous et al., 2006). For example, intracellular zinc content in human colon cancer HT-29 cells is 264 μM (Krezel and Maret, 2006). Most of intracellular zinc (~ 50%) is located in the cytosol, while 30-40% of the intracellular zinc is found in the nucleus. The remaining intracellular zinc is associated with membranes or presents as free zinc ion (Stefanidous et al., 2006). Intracellular free zinc ions generally is extremely low, ranging from 5-1000 pM (Maret, 2013).  The exact mechanism involved in cellular zinc homeostasis remains to be elucidated. However, available evidence shows that a coordinate interplay between zinc transportation and storage is important to cellular zinc homeostasis. The uptake of extracellular zinc and its transportation in and out of intracellular organelles require transporter ZnTs and Zips. These zinc transporters all have transmembrane domains and are encoded by two solute-like carrier (SLC) gene families: ZnT (SLC30) and Zip (SLC39) families. There are 10 ZnTs and 14 Zips transporters in humans. These two transporter families appear to have opposite roles in zinc homeostasis. ZnT transporters are involved in zinc efflux, while Zip transporters are involved in zinc influx (Lichten and Cousins, 2009). Zips are responsible for uptake of extracellular zinc into the cytosol and 9  release of zinc from subcellular organelle lumen (i.e. endoplasmic reticulum, Golgi and lysosomes) into the cytoplasm (Maret, 2011). Most Zip proteins have eight predicted transmembrane domains (Eide, 2004). It appears that gradient of HCO3-, which exists across the membrane drives the zinc uptake in mammalian cells (Gaither and Eide, 2000). On the other hand, ZnTs are responsible for sequestering zinc from cytosol into the extracellular space and into the subcellular organells (Liuzzi and Cousins 2004). However, ZnT5 is an exception, which serves as a bidirectional zinc transporter (Valentine et al., 2007). Through these two families of zinc transporters, intracellular zinc level is controlled within an appropriate range. Another important protein family involved in zinc homeostasis is MTs, which participates in this process by serving as a reservoir of cellular zinc. MT and thionein (T) couple safeguards zinc and controls the concentration of readily available zinc (Vasak and Kagi, 1994). An increase in the amount of free zinc ion (Zn2+) induces the synthesis of T which leads to the formation of MT. On the other hand, zinc is released from MT when intracellular zinc level is low or zinc is needed for the incorporation into zinc proteins (Krezel and Maret, 2006). Human MT proteins have a set of 17 gene products which can be grouped into four major isoforms MT-1, MT-2, MT-3 and MT-4, according to the similarities of their sequences. MT-1 and MT-2 are found widely across different tissues, while the expression of MT-3 is observed in brain and MT-4 is expressed in squamous epithelia (Krizkova et al., 2012). MTs have very high affinity for zinc because of the special structure. There are a total of twenty cysteine residues in each MT molecule. Zinc is bound to MT via the –SH groups of the cysteine residues. These cysteine residues are organized into two domains: the Zn3S9 and Zn4S11 domains. (Maret and Vallee, 1998). The Zn3S9 can bind upto 3 zinc ions (Zhang et al., 2002), while the Zn4S11 domain can bind up to 4 zinc ions (Krezel and Maret, 2007). Therefore, MTs provides a great buffer 10  when cellular zinc content is high. Zinc-induced MT expression is mediated by metal-responsive-element binding transcription factor-1 (MTF-1). In response to high cytosol zinc concentration, zinc binds to MTF-1 to form two to six zinc-fingers in MTF-1 (Gunther et al., 2012). Upon formation of these zinc-fingers, MTF-1 translocates from cytosol to the nucleus, where it activates MT expression by binding to the metal response elements (MRE) (Heuchel et al., 1994; Koizumi et al., 1999; Smirnova et al., 2000). Besides, MTF-1 also controls the expression of a number of other genes directly involved in zinc transportation such as Zip10 (Laity and Andrews, 2007).  1.1.4.2 Whole body zinc homeostasis   Zinc is present in all organs, with a higher concentration in liver and skin, and a moderate level in kidney, prostate, pancreas, bone, and muscle (He et al., 1991). There is no specific storage site for zinc in human body. The whole body zinc homeostasis is mainly maintained through the balance of absorption and excretion. Dietary zinc is obtained by hydrolysis of proteins during digestion (Gropper and Smith, 2012). As discussed before, the absorption can be influenced by the presence of zinc enhancers and inhibitors. Animal-based foods generally offer better absorption of zinc than plant-based foods. In addition, zinc absorption is enhanced when plant-based foods are consumed at the same time with animal-based food sources (Sandstroem et al., 1989). The efficiency of dietary zinc absorption is inversely related to dietary zinc intake. Excessive dietary zinc intake can decrease the efficiency of zinc absorption and increase zinc excretion (Solomons and Cousin, 1984). 11  The gastrointestinal tract is the major site for regulation of zinc homeostasis. It involves the adjustments of both absorption of dietary zinc and excretion of endogenous zinc into the feces (Wang and Zhou, 2010; Krebs, 2000). Zinc is absorbed in the small intestine with the major site at jejunum (Lee et al., 1989). Absorption can be considered as the process of influx into the enterocytes (Lee et al., 1989). The mechanism of zinc absorption involves ZnTs and Zips. Both ZnTs and Zips show tissue specific expression. In addition, differential responsiveness is observed to dietary zinc deficiency, dietary excessiveness, and different physiological stimulation (Reyes, 1996). For example, dietary zinc deficiency caused upregulation of Zip4 expression in intestine and pancreas and promoted zinc uptake. Recently, molecular characterization of zinc transporters has been carried out, which substantially improved our understanding of the relationship between cellular zinc absorption and excretion. However, it has not been applied to the whole body level yet. In addition to dietary zinc, there are several potential sources of the endogenous zinc, including pancreatic, biliary, and gastroduodenal secretions (Rink, 2011). Small intestine reabsorbs zinc from these endogenous sources. The reabsorption is considered to be important to maintain zinc homeostasis for the population with chronically low dietary zinc intake (Lee et al., 1993a). Currently, regulation of the zinc secretion of endogenous zinc is unknown. However, available evidence shows that the amount of endogenous zinc secreted with meal may be comparable to the amount of exogenous zinc, indicating that reabsorption of endogenous zinc is an important factor in zinc homeostasis (Matseshe et al., 1980). Upon digestion, zinc enters the enterocytes by Zip1 and Zip4 transporters. Subsequently, zinc is transported across the basolateral membrane into portal circulation by ZnT1 transporter (McMahon and Cousins, 1998). In blood, albumin acts as the major 12  transport protein for zinc. Zinc also binds to other proteins for its transportation, including transferrin, alpha 2-macroglobulin, histidine, and cysteine (Chesters and Will, 1981). In fact, zinc in blood circulation only accounts for 0.1% of the total amount of zinc in human body (Cousins, 1997). Fecal zinc excretion accounts for the largest amount of the zinc loss in the body (Wang and Zhou, 2010). Body also loses zinc through urine, sweat, skin, hair, menstruation and semen at very tiny amount (King et al., 2000) In summary, zinc homeostasis at the whole-body level is primarily achieved through regulation of absorption, transportation, and excretion. At the cellular level, zinc level is homeostatically regulated to meet the requirements for cellular activities and to prevent against intracellular zinc overload. The failure of maintenance of zinc homeostasis at both whole-body and cellular levels not only affects the physiological functions, but also leads to the development and progression of several types of diseases such as asthma, diabetes and Alzheimer's disease (Devirgiliis et al., 2007).  1.2 Zinc and growth  Zinc is essential for growth. In fact, one of the most profound effects of zinc deficiency is growth retardation. In early 1960’s severe dwarfism was reported among Egyptian adolescents. These adolescences consumed mainly whole-wheat flour-based diets with little or no intake of animal-based proteins. Plant-based foods contains low amount of zinc with low bioavailability (Sandstead et al., 1967). This low dietary zinc bioavailability caused zinc deficiency, which in turn led to the development of dwarfism among these Egyptian adolescents. In fact, this observation led to the establishment of zinc essentiality in humans (Sandstead et al., 1967).  13  So far, marginal and moderate zinc deficiency in children with growth impairment has been reported in both the developed and developing countries (Prasad, 1996). In Iranian adolescents, severe growth retardation is greatly improved through zinc supplementation (Ronaghy et al., 1974). In Denver, school children received zinc supplementation grew about 10% taller than the controls (Hambidge et al., 1972; Hambidge, 1989). Similarly, zinc supplementation promotes growth in children with very low birth weight and significant amelioration of psychomotor development (Hamadani et al., 2001). In addition, zinc is also required for several other aspects of growth and development in children. For example, Castillo-Duran et al., (2001) reported that zinc supplementation promotes motor and mental development among Chilean infants. It was reported that maternal zinc deficiency could adversely impact the fetus intellectual development (Caulfield et al., 1998). Furthermore, zinc also appears to play a significant role in recognition, memory, reasoning and psychomotor functions (Sandstead et al., 1998). Clearly, zinc is important to growth and development in children. However, the exact mechanisms where by zinc influences growth remain unclear. One of the possible mechanisms involved is through its influence on the expression and function of the growth hormone and insulin-like growth factor 1 (IGF-1), two principle hormones involved in regulating growth (Underwood et al., 1994). Growth hormone is a peptide hormone secreted by pituitary which reaches target tissues through peripheral circulation to stimulate growth. Zinc concentration is very high in pituitary (Henkin, 1976). In fact, there is a zinc-binding site in growth hormone (Cunningham et al., 1991). The relationship among zinc, growth hormone, and animal growth indicates the direct involvement of zinc in the action of this hormone. It was reported that zinc treatment promoted the expression of growth hormone in mice (Palmiter et al., 1983). Zinc 14  administration was found to be able to increase the basal plasma growth hormone level in human (Imamoglu et al., 2005). In contrast, insufficient zinc supply reduces the concentrations of growth hormone and the abundance of its receptors in rats (Roth and Kirchgessner, 1997). Bone is one of the major targets of growth hormone. Bone growth stimulated by growth hormone was only observed in zinc-adequate rats, while zinc-deficient rats were resistant to growth hormone which leads to impaired bone growth (Cha and Rojhani, 1997).  IGF-1 is a hormone with a similar structure to insulin, which plays an important role in growth and shows anabolic effects in humans (Corpas et al., 1993). Serum IGF-1 in rats fed with zinc free fodder (27.3 μmol/kg) is lower than that in rats fed with zinc enriched fodder (500 μmol/kg) after a three-week treatment period (Dorup et al, 1991). The attempt of maintaining serum IGF-1 level by osmotic pump and energy intake within the normal ranges in zinc-deficient rats was not able to correct the impaired growth in rats completely. Since the impaired growth was associated with zinc deficiency (Browning et al., 1998). In humans serum IGF-1 level appears to be positively correlated with body zinc status (Cossack, 1984). The circulating concentration of IGF-1 level is lower in humans with zinc deficiency (Cossack, 1991).  1.3 Zinc and cell proliferation  1.3.1 Cell proliferation   Cell proliferation is the biological process that produces two cells from one, including cell growth followed by cell division (Evan and Littlewood, 1998). In normal tissues, cell proliferation is required to replenish the tissue. The stem cells, which exist in 15  most of the tissues, possess the replenishment function (Sylvester and Longaker, 2004). Cell proliferation involves DNA replication and cell division, which are organized in the processes known as cell cycle. Cell cycle consists of four stages: G1, S, G2, and M phases. The S phase is the DNA replication stage. DNA packaging, chromosome separation, and cell division occur in the M phase. S and M phases are separated by gap phases: the G1 and G2 phases. The G1 phase is the gap between M and S phases, which is the critical control point of cell cycle. The G2 is the gap between the S and M phases, which assures proper DNA replication and packaging before cell division (Schafer, 1998; Vermeulen et al., 2003).  Cell proliferation is tightly controlled for normal tissue replenishment and growth as well as prevention of excessive and unwanted cell replication. There are two classes of regulatory circuit: intrinsic and extrinsic regulations. The intrinsic pathways can be initiated by intracellular conditions such as oxidative stress, hypoxia, and other intracellular signals, while the extrinsic regulatory pathways function in response to extracellular conditions (Hartwell and Weinert, 1989).  One of the signalling pathways involving in the regulation of cell proliferation is the mitogen-activated protein kinases (MAPK) pathway. MAPK are a family of serine/threonine/tyrosine-specific protein kinases, which are responsible for signalling transduction and controlling intracellular events (Pearson et al., 2001). In mammalian cells, three MAPK families have been identified: extracellular signal-regulated kinases (ERK), C- Jun N- terminal kinase (JNK), and p38 kinase. Members of these MAPK families are an integral part of a number of protein kinase cascades. Each cascade consists of no less than three MAPK enzymes, which are activated in series. For example, a MAPK kinase kinase (MAPKKK) activates a MAPK kinase (MAPKK), which in turn activates a MAPK (Widmann et al., 1999). Upon activation, the MAPK activates its 16  downstream target through phosphorylation. In addition to cellular signalling pathway, one of the most important links between cell proliferation and cell death machineries is p53, a tumor suppressor, which promotes the arrest of cell proliferation (Levine, 1997). The p53 gene is the most frequently mutated gene in human cancers, indicating its critical role in maintaining normal cell cycle progression (Amaral et al., 2010). p53 has been demonstrated to regulate the expression of numerous genes (i.e. WAF1, p21, and MDM2) with a role in controlling cell proliferation (el-Deiry et al., 1993; Kastan et al., 1992). The essential role of p53 in controlling cell proliferation is to suppress cell proliferation at G1 phase, before DNA is replicated. Tumor cells with mutated p53 are more likely to progress to the S phase, resulting in unwanted proliferation (Brown and Wouters, 1999). p53 can directly increase the expression of p21, a protein with a molecular weight of 21 kDa. p21 prevents the activation of cyclin-dependent kinases, leading to G1 arrest and ultimately suppression of cell proliferation (Zhou et al., 2013).  In summary, cell proliferation is a highly regulated process. Signalling network is important for the regulation of cell proliferation. Cross-talk between different signalling pathways may take place at many levels from membrane to nucleus, including positive and negative feedback signals. In addition, some cell proliferative and anti-proliferative regulators are strongly implicated in the signalling pathways to assure the precise regulation of cell proliferation.  1.3.2 Role of zinc in cell proliferation  Numerous studies have documented the essential role of zinc in cell proliferation and suppressed cell proliferation during zinc deficiency. Zinc deficiency at the cellular level 17  can be reached by culturing cells in a low or no zinc medium or using zinc-specific chelator (Truong-Tran et al., 2000). N,N,N’,N’-tetrakis(2-pyridylmethyl)ethylenediamine (TPEN) is a membrane permeable, divalent cation chelator with a higher affinity towards Zn2+ than other divalent cations such as Ca2+, Fe2+, and Mg2+ (Arslan et al., 1985). Zinc deficiency has been shown to suppress cell proliferation. Culturing HUT 78 lymphoblasts in a low zinc medium (0.76 ± 0.24 μmol/L) significantly prolonged the cell doubling time, compared to the cells grown with regular medium (4.12 ± 0.02 μmol/L; Prasad et al., 1996). The previous work carried out in our laboratory has showed that DNA synthesis and cell proliferation are negatively impacted when mouse fibroblast 3T3 cells were cultured in a low-zinc medium (Paski and Xu, 2001). Since DMEM contains no zinc, zinc in the cell culture medium comes exclusively from FBS. To prepare a low-zinc medium, the regular FBS will be batch-treated with Chelex-100, a divalent cation chelator, for 24 hours. Then the Chelex-100 treated FBS will be added to DMEM at 10% to formulate the low-zinc medium with a zinc concentration of about 0.1 μmol/L. When the 3T3 cells are cultured in the low-zinc medium supplemented with zinc at 5, 10, or 20 μmol/L, DNA synthesis and cell proliferation are significantly elevated along with a significant increase in the abundance of LIPZ, compared to the cells cultured in the low-zinc medium. Further, zinc is also required to support a growth factor-stimulated DNA synthesis and cell proliferation in 3T3 cells (Paski and Xu, 2002) and cell cycle progression (Huang et al., 2000). Similarly, a lower intracellular zinc concentration is associated with reduced proliferation of pancreatic cancer cells (Donadelli et al., 2008). Wu (2003) has observed a reduced proliferation of human breast cancer MDA-MB-231 cells in response to TPEN-induced zinc depletion. Evidently, zinc is required for cell proliferation; however, the role of zinc in cell proliferation and regulation remains unclear. 18   1.3.2.1 Possible mechanism involved in zinc-dependent cell proliferation  Currently, our understanding of the mechanisms by which zinc regulates cell proliferation is quite limited. The available evidences suggest zinc may influence cell proliferation at the level of DNA synthesis, cellular signalling pathways, and zinc-containing regulators. DNA synthesis The effect of zinc on DNA synthesis is the most direct way to regulate cell proliferation. Zinc is present in the cell nucleus and is able to stabilize the structure of both DNA and RNA (Wu and Wu, 1987). A series of enzymes, involved in the DNA and RNA synthesis contain zinc, including DNA polymerase, RNA polymerase, and reverse transcriptase (Wu and Wu 1987, Wu et al., 1992). The zinc-finger domain, where zinc ion creates a bridge between cysteine and histidine, is considered essential to regulate the binding of transcription factors to their specific DNA sequences (Vallee and Auld, 1995).  Zinc is required for thymidine kinase which catalyzes the phosphorylation of deoxythymidine to deoxythymidine monophosphate in the pyrimidine salvage pathway. Thymidine kinase is responsible for one part of reaction chain to introduce deoxythymidine into DNA. Therefore, its activity is used as a marker for cell proliferation. Decreased incorporation of thymidine into DNA was observed in the livers, and kidneys in rats fed with zinc deficient diets for five days (Williams and Chesters, 1970). Unlike DNA and RNA polymerases, thymidine kinase is not a zinc-containing enzyme. However, the transcription of this enzyme is regulated by zinc. Chelation of zinc by the diethylene triamine pentaacetic acid (DTPA) results in decreased mRNA expression of thymidine kinase (Chesters et al., 1990) due to a weakened binding 19  between the zinc-containing transcriptional factors and the promoter of the thymidine kinase genes (Chesters et al., 1995). In addition, zinc plays a critical role in hormone receptors such as estrogen and androgen. Inadequate zinc supply may impair the responsiveness of the binding between hormone and its receptor (Bunce, 1994). Zinc is present as a bridge between the growth hormone and its receptor in human (Berg and Shi, 1996). The thymidine uptake was suppressed by the DPTA treatment while the uptake of thymidine, was restored by the addition of zinc (MacDonald et al., 1998). The mitogenic signalling can induce the transition of cells from G1 to S phase by adenosine(5’)tetraphosphate(5’)adenosine (Ap4A) pool regulated by zinc. Zinc serves as a putative second messenger of mitogenic induction and a trigger for DNA synthesis in baby hamster kidney cells (Grummt et al., 1986).  Cellular signalling pathways  Mediation of cell proliferation by growth factors requires the binding of ligands to their corresponding receptors, which leads to the initiation of the intracellular pathway. So far, several studies have been carried out to investigate the effects of zinc deficiency and zinc supplementation on MAPK signalling pathway using both human and rat experimental models (Cossack, 1991; Dorup et al., 1991). The inhibition of IGF-1 mitogenic action by zinc chelation using DTPA, is associated with a decreased MAPK activation in rat fibroblasts. This inhibition was alleviated by adding zinc sulfate back to the medium (Lefebvre et al., 1999). Furthermore, the elevated zinc level was observed to stimulate the tyrosine phosphorylation and to enhance the MAPK activity as well (Hansson 1996). In human epithelial cells, zinc also induces epidermal growth factor (EGF) receptor phosphorylation and MAPK activation (Wu et al., 1999). It was demonstrated that MAPK, ERK, JNK, and p38 which in turn regulate the cell proliferation were activated 20  by a high zinc concentration in both the mouse cortical cells and human bronchial epithelial cells (He and Aizenman, 2010; Samet et al., 1998). The enhancement of the MAPK activites was presumably attributed to the inhibition of protein phosphatases caused by zinc (Maret et al. 1999). Protein kinase C is a family of protein kinase enzymes which are involved in controlling the function of phosphorylation of other proteins. The activation of protein kinase C is associated with the induction of cell proliferation. Hence, protein kinase C is important in signal transduction cascades. Protein kinase C is a zinc metalloenzyme with two zinc-binding motifs in the regulatory region (Hubbard et al., 1991). It was reported by Csermely et al. (1988) that zinc at nanomolar concentration could activate protein kinase C and trigger its translocation to plasma membrane, where most cellular activation events were catalyzed by protein kinase C (Csermely et al., 1988).  QM, another zinc-binding protein, binds to the leucine region of C-Jun, which is a part of the JNK group of MAPK and mediates the cell cycle (Pelech and Charest, 1995; Inada et al, 1997). Zinc is required for the binding between QM and C-Jun. In other words, the presence of zinc is a prerequisite for the functionality of QM.  Zinc-containing regulators Tumor suppressor p53, which is down-regulated in a large range of cancers, is a zinc-finger transcription factor. The zinc-finger motif is formed via coordination of a zinc atom within a Cys-His cluster. p53 binds to DNA via sequence-specific DNA binding domain to execute its function (Meplan and Hainaut, 1999). p53 is activated by cellular stress signals to prevent cells from proceeding with new DNA synthesis or G2 checkpoint to undergo cell division (Ho and Ames, 2002). p53 can induce cell cycle arrest and inhibit cell proliferation by transcriptional activation of the p21 gene, whose protein product can inhibit cyclin and cyclin dependent kinase resulting in G1 block (Hainaut and Mann, 2001). In the past two decades, numerous 21  studies have been conducted to investigate how zinc regulates cell proliferation by manipulating the mRNA levels of p53, the accumulation of p53 protein, and its downstream targets. In lung fibroblasts, an increase in p53 expression was induced by zinc deficiency, leading to cell cycle arrest (Ho et al., 2003). The levels of p53 protein and p21 mRNA increase as zinc level decreases indicating the inhibition of cell proliferation in aortic endothelial cells (Fanzo et al., 2002). Zinc depletion-induced cell cycle arrest and inhibition of cell proliferation accompanied with an up-regulated p53 expression level were observed in both in HepG2 cells (Reaves et al., 2000) and human bronchial epithelial cells (Fanzo et al., 2001). Clearly, the expression of p53 is activated when the suppression of cell proliferation is induced by zinc deficiency. In summary, cell proliferation can be regulated by zinc. Zinc depletion has been shown to inhibit cell proliferation in a wide range of cells and tissues, which possibly involves regulation of DNA synthesis, cellular signalling pathways, and zinc-containing regulators. However, the zinc-mediated regulation of cell proliferation is complex and remains to be further elucidated.  1.4 MicroRNAs and cell proliferation  1.4.1 miRNA  MicroRNAs (miRNAs) are a newly discovered class of short, non-coding RNA, which regulate numerous cellular activities via post-transcriptional regulation (Cai et al., 2009). The first miRNA, lin-4, was identified in 1993 and the research in miRNA has been rapidly evolving since then (Lee et al., 1993b). In the beginning of the 21st century, a small regulatory RNA, let-7 was identified as an additional regulator of developmental 22  timing in C. elegans (Reinhart et al., 2000). Interestingly, it was demonstrated that both the sequence and expression pattern of let-7 are conserved in many animal species, indicating that the miRNA-mediated regulation is part of overall regulation of gene regulation (Pasquinelli et al., 2000). This new level of gene regulation has not attracted much more attention from the academic world, until hundreds of similar miRNAs were found and investigated in C. elegans, Drosophila, and mammals (Bartel, 2004). With over 2,500 mature miRNAs identified in human, the miRNA research has been prosperously developed from then on (miRBase, 2014).  miRNAs are 18 to 25 nucleotides in length (Bartel, 2004). miRNAs regulate their target mRNAs by complementary base pairing and normally lead to silencing or degradation of the targeted mRNA (Ameres and Zamore, 2013). miRNAs play a role in a wide-range of physiological processes, including embryogenesis, development, and immunity (Chang and Mendell, 2007). Abnormal expression of miRNAs contributes to disease pathogenesis such as cancer initiation and progression.  Each miRNA is capable of regulating multiple target mRNAs and multiple miRNAs may coordinate to the regulation of a single mRNA transcript (Krek et al., 2005). Through this extensive and complex regulatory mechanism, the potential regulatory circuitry afforded by miRNA is enormous. In fact, it is estimated that more than 3% of human genes encode for miRNAs and up to 30% of human protein-coding genes may be regulated by miRNAs, suggesting the importance of miRNA in gene expression and regulation (Lewis et al., 2005). The targets of miRNAs include mRNAs encoding enzymes and transcriptional factors involved in regulating numerous key cellular activities such as cell proliferation, differentiation, and apoptosis (Kim, 2005). Mature miRNAs are generally very stable with an average half-life ranging from hours to days (Zhang et al., 2012; Ruegger and Grobhans, 2012). For example, the 23  majority of miRNAs (95%) in human embryonic kidney HEK-239T cells are stable for at least 8 hours (Bail et al., 2010). In mouse embryonic fibroblasts, the average half-life of miRNAs is 119 hours, which confirmed the inherent stability of miRNAs. However, it is demonstrated that some miRNAs turnover more rapidly than others. For example, miR-155 has a shorter half-life than miR-125b (Gantier et al., 2011). Thus, miRNAs are generally stable with a half-life ranging from 8 to 211 hours (~ 9 days) (Ruegger and Grobhans, 2012). Even though numerous efforts have been invested to elucidate the biological importance of miRNA, little is known about miRNA-mediated regulation of gene silencing, mechanisms by which miRNA expression and interactions with other regulators of gene expression, and signalling pathways etc. (Ameres and Zamore, 2013; Bartel, 2004). 1.4.2 miRNA biogenesis  Production of mature miRNAs is a multi-step process, including nuclear processing followed by cytoplasmic processing. Long primary miRNA (pri-miRNA) sequences are encoded within introns, exons, and intergenic regions (Kim and Nam, 2006). miRNAs are frequently clustered such that a single pri-miRNA contains multiple miRNA sequences (Saini et al., 2007). The transcriptions of miRNA genes are typically initiated by RNA polymerase II to form pri-miRNAs, which are capped, polyadenylated and usually have several kilobases in length. The pri-miRNA contains a hairpin-like structure of about 80-nucleotide in length, which consists of a double stranded RNA stem and an unpaired RNA loop. This loop is subsequently processed through cleavage to produce mature miRNA (Kim and Nam 2006). Two steps of ribonuclease processing reactions are generally required to trim a pri-24  miRNA transcript into a mature miRNA. The first processing event occurs in the nucleus by a nucleus multi-protein complex, known as the microprocessor (Denli et al., 2004; Gregory et al., 2004). The microprocessor-mediated cleavage of pri-miRNA requires a cofactor, called DiGeorge syndrome critical region gene 8 (DGCR8) for substrate recognition and recruitment of RNase enzyme Drosha (Han et al., 2004). The importance of DGCR8 cofactors has been confirmed using DGCR8-deficient mouse embryonic stem cells (Wang et al., 2007). In the absence of DGCR8, no mature miRNA is produced. Drosha cleaves the pri-miRNA into a short precursor miRNA (pre-miRNA) with 70 nucleotides in length (Lee et al., 2003). Pre-miRNA is rapidly exported to the cytoplasm by the nuclear export protein called exportin 5 with Ran-GTP as the cofactor (Yi et al., 2003; Bohnsack et al., 2004).  The second phase of miRNA production occurs in cytoplasma by Dicer, a miRNA generating complex containing an RNAase III enzyme to produce an 18-24 nucleotide duplex. The miRNA generating complex may contain additional proteins, including trans-activating response RNA binding protein, which may cleave the fully processed duplex to form a single-strand mature miRNA and another strand, known as passenger strand miRNA*, which is degraded subsequently (Finnegan and Pasquinelli, 2013). The thermodynamic stability of the miRNA duplex is an important factor of strand selection. Clearly, miRNA biogenesis is highly regulated. Despite the pathway described is thought to be universal, recently researches have revealed some exceptions. For example, some miRNAs undergo the processing that is independent of Drosha or Dicer (Czech and Hannon, 2011).  25  1.4.3 Functions of miRNA  The mature miRNA complexes with a protein to form a RNA-induced silencing complex (RISC) and acts as the guide that directs RISC to target mRNA, subsequently cleaved or translational silenced (Bartel, 2004). The RISC is composed of Dicer, trans-activating response RNA-binding protein, and Argonaute (Ago; Winter et al., 2009). Ago proteins are composed of four domains: the PAZ domain, which binds to the miRNA at the 3’ terminus, the PIWI domain, which can catalyze the cleavage of miRNA base-paired target, the MID-domain which binds to the 5’ end of the mature miRNA, and the N-domain (Kim and Nam, 2006). In humans, there are four Ago proteins mediating miRNA-targeted regulation of translation (Czech and Hannon, 2011). The RISC mediates degradation of mRNA based on complementarity between miRNA and its targeted mRNA. The mature miRNA contains a seed region of 6 to 8 nucleotides at the 5’ end of miRNAs which can bind to mRNA by complementary base paring. Although base paring at the 3’ end of miRNA seem to be less critical in target recognition, it may contribute to target selection when weak miRNA seed match occurs (Ameres and Zamore, 2013). The seed region may bind to any part of the mRNA. However in most cases, miRNA binding sites are located at the 3’ un-translated region (UTR; Carthew and Sontheimer, 2009). Binding of miRNA to the target mRNA leads to mRNA cleavage or translational attenuation based on the degree of complementarity. Perfect base paring allows Ago-catalyzed cleavage of mRNA strand, while imperfect base paring with some central mismatches leads to translation repression of mRNA (Lewis et al., 2005). The translational attenuation may occur at the beginning of the translation, post-initiational repression, or elongation stage depending on different experimental systems (Carthew and Sontheimer, 2009). In mammalian cells, it has been showed that the miRNA-26  meditated mRNA degradation is the primary type of gene silencing mechanism (Huntzinger and Izaurralde, 2011).  Through bio-informatical approaches, miRNA targets may be predicted based on conserved seed pairing and sequence features. Thus the predicted targets of mammalian miRNAs are primarily involved in the transcriptional regulation. However, miRNAs are also related to a broad range of other functions (Smalheiser and Torvik, 2006). For example, miRNAs are involved in development. As mentioned above, lin-4, the first discovered miRNA, regulates the timing of C. elegans development by inhibiting LIN-14 protein expression (Lee et al., 1993b). In addition, studies with defect in miRNA biogenesis pathway provide evidence that miRNAs also play important roles on stem cell regulation (Wang et al., 2007). Functional analysis indicates that miRNAs may have specific roles in controlling stem cell differentiation and renewal. For example, miR-290~295 cluster and miR-296 decreased during stem cell differentiation, while miR-21 and miR-22 were up-regulated in the same process (Gangaraju and Lin, 2009). In mammalian immune system, miRNAs serve as the critical regulators. Loss or deregulation of certain miRNAs can lead to compromised immune development as well as some immunity disorders (Petrocca and Lieberman, 2009). Typically, miRNA-mediated gene regulation is fulfilled in negative patterns, however, positive regulatory patterns have also been found in gene regulation. For example, miR-369-3 and let-7 up-regulate the translation of target mRNA in cell cycle arrest (Vasudevan et al., 2007). In fact, the miRNA-mediated gene regulation is a newly emerging research hotspot and the exact mechanisms involved are currently not well known. Some findings are even controversial (Ameres and Zamore, 2013). In summary, miRNAs have attracted considerable attention because of their important roles in development, normal physiology and even disease states. It will be 27  discussed with a focus on the regulatory role of miRNAs on cell proliferation in the following sections.  1.4.4 miRNA and cell proliferation  The earlier relevance of miRNAs in cell proliferation was established when investigating the miRNA biogenesis mechanism, the function of Dicer. Depletion of Dicer induces global miRNA down-regulation due to the repression of miRNA maturation (Bernstein et al., 2003). It is found that the Dicer ablation-induced miRNA down-regulation reduced cell proliferation with up-regulation of p53 signalling pathway in mouse embryonic cells (Kanellopoulou et al., 2005; Mudhasani et al., 2008). However, the overall decrease in miRNA expression in human colon cancer HCA-7 cells, human breast cancer MCF7 cells, and human osteosarcoma U2OS cells, has also been observed to promote cell proliferation, indicating that miRNAs may have either proliferative or anti-proliferative activities to influence cell proliferation (Kumar et al., 2007). In fact, miRNAs with proliferative activity are capable of promoting cell proliferation, even induce oncogenesis, while miRNAs with inhibitory/anti-proliferative activity can suppress cell proliferation, and even induce apoptosis under specific conditions. Some well-studied miRNAs were selected and discussed with a focus on their regulations of cell proliferation in this section. mir-17 cluster The miR-17 cluster, including six miRNAs: miR-17, miR-18, miR-19a, miR-19b, miR-20, and miR-92, targets multiple cell cycle regulators, including E2F family, and c-myc, which are key regulators of cell-cycle progression and cell proliferation (Iaquinta and Lees, 2007). In fact, E2Fs execute their functions by binding to DNA sequence; while c-myc, a helix-loop-helix leucine zipper transcription factor, is a 28  potent activator of E2Fs (DeGregori et al., 1997). Though members of the E2Fs family tend to bind to the DNA sequences by similar mechanisms; however, it is divided into two groups according to the activating effect. E2F1, E2F2 and E2F3 are involved in promoting cell proliferation; while E2F4 and E2F5 are involved in suppressing cell proliferation (Trimarchi and Lees, 2002). That is the reason why miR-17 family members might either promote or inhibit cell proliferation based on the different regulators they target. E2Fs mediate the cell proliferation during G1-to-S transition in cell cycle, which is called the critical point, after which cells start proliferation through DNA replication. The locus of miR-17 cluster is located in a region where it is frequently amplified in several types of lymphoma and breast tumors (Ota et al., 2004). Introducing miR-17 cluster into Eμ-myc mice, a strain of mice, which develops B-cell lymphomas during the late life stage, dramatically accelerated the development of diseases suggesting that these miRNAs act as the promoters of cell proliferation (O’Donnell et al., 2005). In prostate cancer cell line, induction of c-myc expression is positively correlated with the increased of miR-17 cluster expression, and in turn these miRNAs inhibit E2Fs translation (Sylvestre et al., 2007). This feedback system provides the possible mechanisms to regulate whether cellular proliferation is disfavoured or promoted depending on specific circumstance. For example, cell proliferation of some breast cancer cell lines is down-regulated by increased expression of miR-17 cluster (Yu et al., 2008; Hossain et al., 2006). Recently, amplification of miR-17 cluster has been reported in lung cancers and cell proliferation is accelerated by the overexpression of the cluster (Hayashita et al., 2005). let-7 family Early studies of let-7 in C. clegans revealed its crucial regulatory role in cell cycle exit (Reinhart et al., 2000). let-7 family in mammalian shows its inhibitory effect on cell proliferation. The let-7 family includes 12 homologs with most of their 29  genes locate in a region deleted in human cancers such as human lung cancer, ovarian cancer, and prostate cancer (Calin et al., 2004). Accumulated evidences have indicated that let-7 is involved in mediating suppression of cell proliferation. Transferring let-7 into a mouse model with lung adenocarcinoma reduces tumor number and size (Esquela-Kerscher et al., 2008). The abundance of the let-7 family is also reduced in breast tumor (Yu et al., 2007). RAS is a group of proteins belongs to the small GTPase families. RAS is involved in controlling cellular signalling pathways such as MAPK or its subfamilies and is a potent activator of cell proliferation (Aksamitiene et al., 2012). RAS mRNA has been identified as a target of let-7 in C. elegans and in humans (Johnson et al., 2005). In lung cancer patients, RAS protein levels are inversely correlated with let-7 expression level (Takamizawa et al., 2004). Overexpression of let-7 results in accumulation of the liver cancer cells HepG2 in the G0 to G1 phases. Several important cell cycle control genes (i.e. cyclin-dependent kinase and cyclins D1, D3 and A) are also reported to be targets of let-7 by microarray analysis (Schultz et al., 2008). Thus, compelling evidence demonstrates that let-7 miRNAs can suppress cell proliferation through targeting at RAS and cell cycle progression regulators. miR-15a/16 cluster miR-15a/16 exerts an inhibitory activity to cell proliferation. This cluster is located on human chromosome 13 in a region frequently deleted in choronic lymphocytic leukemia (Calin et al., 2002). The miR-15a/16 cluster is also observed to be deleted or down-regulated in gastric cancer and pituitary adenoma (Bottoni et al., 2005; Xia et al., 2008). BCL2 is a potent inhibitor of cell death. The levels of miR-15a/16 are inversely correlated with the BCL2 protein levels. Overexpression of miR-15a/16 cluster in leukaemic cells inhibits cell proliferation and ultimately triggers cell death, with a decreased BCL2 protein expression (Cimmino et al., 2005). Transferring miR-15a and miR-16 into colon carcinoma cells DLD-1 demonstrated a 30  down-regulation of cell-cycle related gene (i.e. cyclin D3 and cyclin D1), investigated by microarray (Linsley et al., 2007). Collectively, available evidence suggests that miR-15a/16 cluster plays an inhibitory role in cell proliferation with BCL2 being a target. miR-34 family The miR-34 family consisting miR-34a, miR-34b, and miR-34c is a group of miRNAs that are direct transcriptional targets of p53 and show inhibitory effect on cell proliferation (He et al., 2007). p53 can bind to the promoter of miR-34 to prevent cells from proceeding with newly synthesized DNA resulting in G1 arrest (Tazawa et al., 2007; Bommer et al., 2007). The locus encoding p53 is frequently deleted in a diverse cancer types (i.e. lung cancer, ovarian cancer and breast cancer etc.), indicating its tumor suppressor role (He et al., 2007). The complete spectrum of targets down-regulated by miR-34 awaits for future confirmation; however, numerous crucial cell-cycle regulators including cyclin D1, cyclin-dependent kinases, and proliferative proteins such as BCL2 have been validated (Tarasov et al., 2007; Sun et al., 2008).  miR-21 miR-21 is a potent oncogene with an apparent proliferative effect on cell proliferation. It is overexpressed in glioblastoma tumor tissue and cell lines; while inhibition of miR-21 expression in glioblastoma cells using oligonucleotide antisence transfection results in decreased cell proliferation by activation of the caspase pathway (Chan et al., 2005). In addition, an elevated expression of miR-21 was observed in breast cancer cells (Iorio et al., 2005). Inhibition of miR-21 in breast cancer MCF-7 cells suppresses the cell proliferation of breast cancer cells (Si et al., 2007). As described above, it has been well investigated that miRNAs play a critical role in the regulation of cell proliferation, with either proliferative or anti-proliferative effects depending on the form of miRNAs involved. The interaction between miRNAs and their corresponding targets has added a new layer of complexity to the regulation of cell proliferation. Since there are numerous miRNAs being demonstrated to possess a 31  regulatory role on cell proliferation, only the most well-studied miRNAs were selected and discussed in this section. According to the literature, it is possible and promising that more new miRNAs which are involved in the regulation of cell proliferation will be discovered and identified.  1.5 Zinc and miRNA   Zinc is required for cell proliferation while miRNAs have been shown to play a role in regulating cell proliferation. It is plausible that miRNAs are involved in zinc-mediated regulation of cell proliferation. Even though it is a newly emerging area of research with very limited studies, the interference of zinc status on miRNAs expression has been documented by a few studies to date. One human study focusing on the effects of low dietary zinc on serum miRNAs expression was carried out in young male (Ryu et al., 2011). The subjects consumed a low-zinc diet (0.3 mg/d) for 10 days and the miRNAs expression level in blood was profiled by miRNA microarray. There were 20 miRNAs whose expression levels are altered among the total 88 miRNAs tested, with 18 miRNAs being down-regulated and 2 miRNAs being up-regulated. Among the 18 miRNAs, miR-204 and miR-296-5p are most severely down-regulated by zinc depletion. These two miRNAs have been demonstrated to suppress oncogene BCL2 expression (Lee et al., 2010; Wei et al., 2011). After dietary zinc replenishment, the expression levels of 9 of the 18 miRNAs which are down-regulated in response to zinc depletion are increased, suggesting that these miRNAs may serve as the potential biomarkers of zinc status (Ryu et al., 2011). In addition, it has been reported that chronic zinc deficiency (3-4 ppm) for 23-week in rats results in altered miRNA expression profile in all the tissues studied, including 32  esophagus, skin, lung, pancreas, liver, prostate, and peripheral blood mononuclear cells of the rats with esophagus cancer (Alder et al., 2012). In particular, consuming a 23-week zinc deficiency diet, miR-21 and miR-31, which are oncogenic miRNAs, are the top two up-regulated miRNAs in these tissues studied. The expression of these two miRNAs responded in an opposite manner during zinc replenishment. The development of esophagus cancer is alleviated with the reduced level of miR-21 and miR-31 due to the dietary zinc replenishment (Alder et al., 2012). The regulatory effect of zinc on miRNA expression has also been demonstrated in vitro (Zheng et al., 2012). Zinc supplementation, created by using Clioquinol which acts as a zinc ionophore by transporting zinc into cells to raise the intracellular zinc concentration to a cytotoxic level of 50 μM, was found to be associated with a global down-regulation of miRNA expression in human breast cancer MCF-7 cells. The mechanism involved in the down-regulation of miRNAs expression is that zinc cytotoxicity reduced Dicer and Ago2 expression. Dicer is a key enzyme to miRNA maturation; while Ago2 is a key protein for miRNA stability (Zheng et al., 2012). A recent study has been conducted to determine the expression levels of selected miRNAs after zinc treatment in human prostate cancer cell lines 22Rv1, PC-3, LNCaP, and normal prostate cell line PNT1A. The enhanced expression of miR-23a and miR-375 was observed in all these tumor lines, compared to non-tumor PNT1A cell line, suggesting the potential role of these two miRNAs in prostate cancer diagnosis. Additionally, the significant up-regulation of miR-224 expression in 22Rv1 cancerous cell line in comparison to other cell lines indicated that miR-224 may be suitable for the classification of tumors (Hlavna et al., 2012). Decreased zinc level in prostate is a hallmark of prostate cancer since zinc uniquely concentrates in the healthy prostate tissue. Mihelich et al. (2011) investigated that altered 33  miRNA expression plays a role in zinc homeostasis in prostate cancer with reduced zinc status. Array profiling of tissue suggested that the expression levels of miR-183, miR-96 and miR-182 were higher in prostate cancer tissue than those in normal prostate tissue. Overexpression of the entire miR-183-96-182 cluster reduced intracellular zinc level by decreasing expression of zinc transporters involved in zinc uptake. The regulation of zinc homeostasis by this cluster was also observed in embryonic kidney cells HEK-293, indicating a universal mechanism that dysregulation of miRNA expression may contribute to altered zinc homeostasis (Mihelich et al., 2011).  In summary, zinc is required for cell proliferation and zinc restriction can result in suppression of cell growth. However, the exact mechanisms involved remain to be further elucidated. On the other hand, miRNAs has been shown to play an important role in the regulation of cell proliferation. The correlation between zinc status and miRNA expression reveals a potential relationship among zinc nutrition, miRNA expression, and cell proliferation. Currently, very few studies have been carried out to investigate the importance of nutrients’ role in the regulation of miRNA expression. Especially no studies on the role of miRNAs in zinc-regulated cell proliferation have been reported. Further investigation is necessary to determine the functional significance between zinc status and miRNA expression in the context of cell proliferation.  1.6 Hypothesis  It is hypothesized that miRNA plays a role in zinc-dependent cell proliferation in mouse fibroblast 3T3 cells.  34  1.7 Overall objective and specific aims  The overall objective of this project was to determine whether miRNA is involved in zinc-dependent cell proliferation in mouse fibroblast 3T3 cells. 3T3 cell is a well-established in vitro model for studying the regulatory role of zinc in cell proliferation. The study of Chester et al. (1990) has first shown that inadequate zinc impairs DNA synthesis in 3T3 cells. The previous work carried out in our laboratory has demonstrated an intracellular zinc-dependent DNA synthesis and cell proliferation in 3T3 cells (Paski and Xu, 2001). However, the mechanism of how zinc exerts its regulatory role in cell proliferation remains unclear. The specific aims of this research project were:     (1) To determine whether TPEN-induced zinc depletion has an effect on cell proliferation by analyzing cell cycle progression in 3T3 fibroblasts; and (2) To obtain the total miRNA expression profile by determining the abundance of individual miRNA using miRNA microarray in response to zinc-depletion in 3T3 fibroblasts   35  Chapter 2 Materials and Methods  2.1 Cell culture system  3T3-Swiss albino embryo fibroblasts (American Type Culture Collection, Manassas, VA, USA), a cell line established in 1963 whose original growing procedure includes being transferred (T) every 3 days followed by plating at 300,000 cells per plate (Todaro and Green, 1963) were used in this research project. 3T3 cells were routinely maintained in Dulbecco’s Modified Eagle’s Medium (DMEM; Gibco, Grand Island, New York) supplemented with fetal bovine serum (FBS; Gibco, Grand Island, New York; 10%), sodium pyruvate (Gibco; 1 mM), and 50 units/mL and 50 μg/mL penicillin-streptomycin respectively (Gibco) at 37 oC, 5% CO2. This culture medium was referred as the regular medium in this thesis. Cells were cultured in Petri dishes (100 mm x 15 mm, Fisher Scientific, Dubuque, IA, USA) with an initial density of 2 x 104 cells per dish. Cells were harvested for passage when the confluence reached around 85–90%. Cells with passage 116 to 123 were used in this project.   2.2 Zinc depletion and zinc replenishment   TPEN (molecular weight = 424.54; Sigma, St Louis, MO, USA) was used to deplete intracellular zinc. TPEN is a membrane permeable intracellular divalent cation chelator with a very high affinity towards zinc (Arslan et al., 1985). Therefore, TPEN was selected as an intracellular Zn2+ chelator in this study. It has been demonstrated that TPEN can effectively deplete intracellular zinc in HeLa cells (Chimienti et al, 2001) and human breast cancer cell lines (Wu, 2003).  36  TPEN stock solution (20 mM) was prepared by dissolving TPEN powder in dimethyl sulfoxide (DMSO; Sigma, St Louis, MO, USA). The stock solution was stored at -20 oC and thawed at room temperature before use. Prior to zinc depletion, 3T3 cells were cultured in the regular medium for 72 hours with an initial seeding density of 2 x 104 cells per Petri dish (100 mm x 15 mm) at 37 oC, 5% CO2. At the end of the 3-day initial growth phase, the cells were treated by TPEN at a final concentration of 2.5 μM with or without quiescence induction. To replenish zinc, zinc working solution (ZnSO4) was added to the TPEN-treated cell culture medium (0.5% v/v) with a final zinc concentration of 0 (TPEN only), 1.25, 2.5, and 5 μM. The control cells were treated with an equal volume of DMSO. The cells were then cultured at 37 oC, 5% CO2 for 24 or 48 hours respectively depending on the experiment.   2.3 Quiescence induction  Aphidicolin, a reversible inhibitor of nuclear DNA replication, blocks cell cycle at early S phase and is commonly used for quiescence induction in cell culture (Macdonald-Bravo and Bravo, 1985). In order to induce quiescence, 3T3 cells were treated with 0.1% FBS DMEM for 24 hours followed by aphidicolin treatment at 0.5 μg/mL for 24 hours. At the end of the aphidicolin treatment period, the cells were rinsed with cold phosphate buffered saline (PBS) for 3 times before starting TPEN treatment.  2.4 Cell cycle analysis  For cell cycle analysis, 3T3 cells were cultured in the regular medium with an initial seeding density of 2 x 104 cells per Petri dish (100 mm x 15 mm) at 37 oC, 5% CO2 for 72 37  hours to achieve the initial growth of cells. In the set of experiments without the induction of quiescence, cells were treated with DMEM (overall control), DMSO (control), TPEN alone (2.5 μM), or TPEN + 1.25, 2.5 or 5 μM of zinc immediately after 72 hours of the initial growth period. After 24 or 48 hours, the cells were used for cell cycle analysis (n=6).  In the set of experiments involving the induction of quiescence, the cells were induced quiescence using the procedures described above. Then, the cells were treated with DMEM (overall control), DMSO (control), TPEN alone (2.5 μM), or TPEN + 1.25, 2.5, or 5 μM of zinc immediately after 72 hours of the initial growth period and 48 hours of quiescence. After 24 hours, the cells were used for cell cycle analysis (n=6). Cell proliferation was assessed using a flow cytometric-based cell cycle analysis assay (Cecchini et al., 2012). This assay measures the total DNA content using propidium iodide (PI; Sigma, St Louis, MO, USA), and DNA incorporation using 5-bromo-2’-deoxy-uridine (BrdU; Sigma, St Louis, MO, USA). PI is a fluorescent dye, which binds to DNA. The population of the cells at replication state can be determined based on the incorporation of BrdU into thymidine. BrdU-incorporated cells are then measured using monoclonal antibody against BrdU and a fluorochrome-conjugated second antibody. Briefly, one hour prior to the end of TPEN treatment, BrdU (10 μM) was added to the culture medium and the cells were continued to be cultured at 37 oC, 5% CO2. The cells were then rinsed twice with warm PBS (37 oC) before harvest. Adherent cells were harvested by addition of warm 0.25% trypsin-EDTA (ethylenediaminetetraacetic acid; Gibco) and incubated for 3 min at 37 oC, 5% CO2 until the cells became unattached, followed by addition of the regular medium with an equal volume to neutralize the trypsin. The harvested cells were stored in a 15 ml conical tube. Cells were centrifuged (350 x g, 4 oC) for 4 min and the supernatant was removed. The cells were fixed by 38  slowly adding 0.4 mL cold PBS (4 oC), followed by 0.6 mL of 100% ethanol (-20 oC) while vortex for 15 seconds, and kept in dark overnight at 4 oC. After fixation, cells were stained using anti-BrdU mouse antibody and second goat anti-mouse-FITC antibody (Santa Cruz Biotechnology, Dallas, TX, USA) followed by 4-min centrifugation (350 g, 20 oC) and removal of the supernatant. The cell pellet was then re-suspended in PI staining solution (50 μg/mL; Sigma) with a total volume of approximate 1 mL for 30 min. Ribonuclease A (Sigma) was added to obtain a final concentration of 20 μg/mL 30 min prior to reading using flow cytometry. Finally, the cell suspension was transferred to a 5 mL Falcon tube at room temperature and kept in dark. The cell cycle distribution of stained cells was determined by flow cytometry (BD FACSCalibur Flow Cytometer; BD Biosciences, San Jose, CA, USA) installed with BD FACSDIVA Software. A total of 10,000 events were counted for each sample. The percentage of cell cycle distribution was analyzed by FlowJo Software (Tree Star, Ashland, OR, USA).  2.5 Total RNA isolation  3T3 cells were cultured under the same conditions as described in Section 2.1 for the initial growth followed by induction of quiescence. The cells were treated with DMSO (overall control), TPEN alone (2.5 μM) or TPEN + zinc at 1.25 μM for 24 hours. At the end of the treatment period, total RNA was isolated using miRVana miRNA isolation kit (Ambion, Life Techonologies Corporation, Burlington, ON, Canada) according to the manufacture’s instruction. Briefly, cells were rinsed with 3 mL cold PBS and 600 μL of lysis buffer was added to all the Petri dishes. Cells were scraped from the plates and transferred into a 2-mL microcentrifuge tube. Homogenate additive solution (60 μL) was 39  added to the cell lysate and the mixture was vortexed gently, followed by 10-min incubation on ice. Subsequently, acid-phenol chloroform (600 μL) was added to the mixture, followed by 2-min vortex. The mixture was then centrifuged (10,000 x g, 4 oC) for 5 min with the upper aqueous phase was removed and transferred into a 2-mL microcentrifuge tube. Finally, ethanol was added to the tube at 1.25 times of the volume of the aqueous phase transferred and mixed thoroughly by vortexing at room temperature. The total RNA was precipitated twice using the filter cartridge included in the miRVana isolation kit, followed by purification using the ready-to-use wash solution provided in the kit. Finally, total RNA was collected using 50 μL of pre-heated 95 oC nuclease-free water. The purity of the collected RNA was assessed based on the 260/280 and 260/230 ratios using Nanodrop spectrophotometer (Thermo Fisher Scientific, Wilmington, DE, USA). The RNA integrity was assessed by 1% agarose gel electrophoresis.  2.6 miRNA microarray assay  The global miRNA expression was profiled through miRNA microarray carried out by LC Science (Houston, TX, USA). Profiling was based on all mature miRNAs included in the miRBase, consisting of 1892 mouse miRNAs. Three groups: DMSO (control), TPEN (2.5 μM), and TPEN+zinc (1.25 μM) groups were selected for miRNA microarray analysis.  The microarray assay is carried out using a chip which can analyze a large amount of biological materials at one time on a solid substrate usually a glass slide. Briefly, total RNA sample, approximately, 4-8 μg, were extended with a poly(A) tail and labeled with fluorescent dye. The labeled RNAs were hybridized to a μParaflo microfluidic chip, on 40  which miRNA-specific probes have been arrayed (Zhu et al., 2007; Atactic Technologies, Houston, Texas). The microfluidic platform, which consists of thousands of three-dimensional chambers, can produce a uniform distribution of the sample solution and enhance binding reactions. Each detection probe was composed of chemically modified oligonucleotide, which was complementary to target miRNA or other RNA controls. Saline-sodium phosphate-EDTA buffer (0.90 M NaCl, 60 mM Na2HPO4, 6 mM EDTA, pH 6.8) with 25% formamide was used in hybridization at 34 oC in a total volume of 100 μL. The cyanine 3 dye was circulated thoroughly in the chip for fluorescent staining. The distribution and intensity of the fluorescence were detected by a scanner (Molecular Devices, Sunnyvale, CA, USA) and image digitization was generated by Array-Pro image analysis software (Media Cybernetics, Rockville, MD, USA). Cylic LOWESS (Locally-weighted Regression) method was employed for data normalization (Bolstad et al., 2003). Relative signal intensity was derived from normalization, which distributed from 0 to 65,535. The z-scores of log-transformed signal intensities were used for all data analyses. According to the manufacturer’s instructions and detailed investigation, several factors were considered to select proper miRNAs as the potential targets. Firstly, a minimum intensity of 500 in at least one sample group was required to ensure that the intensity was detectable in the subsequent analysis. Secondly, to identify miRNAs whose expression was affected by TPEN or TPEN plus zinc replenishment, a minimum of 1.5-fold change was used as the cutoff point to balance between false positive and false negative in identifying potential targets. In addition, p-value less than 0.05 was a prerequisite to obtain a true biological difference between the treatment group and the control.  41  2.7 Statistics  The data of cell cycle analysis were presented as mean ± SD (n = 6 biological replications). The differences in the proportion of cells at each phase of the cell cycle between the DMEM and DMSO groups, and DMSO and TPEN groups were tested using Students’ t-test (p < 0.05). The difference in the proportion of cells at each phase of the cell cycle among the TPEN group and TPEN plus zinc replenishment groups were subjected to one-way ANOVA followed by post-hoc analysis using Tukey’s Honestly Significant Difference (HSD) test (p < 0.05). The value of miRNA microarray were presented as mean ± SD (n = 4 biological replications). The effects of treatments in miRNA microarray were analyzed using one-way ANOVA followed by Students’ t-test (p < 0.05), performed by LC science.   42  Chapter 3 Results and Discussion   3.1 Results   3.1.1 TPEN-induced zinc depletion caused a G1/S arrest   TPEN treatment in the absence of quiescence Cell cycle distribution in cells treated with TPEN for 24 hours without the induction of quiescence was shown in Figure 3.1 and Table 3.1. After culturing the cells in the growth medium (DMEM group) for 24 hours, there were 45.4% and 45.9% of the cells in the G1 and S phase, respectively. Inclusion of DMSO in the culture medium had no effect on the proportion of cells in the G1 and S phases, showing that DMSO had no effect on the cell cycle progression. Treating the cells with TPEN for 24 hours resulted in 9.4% increase in the proportion of cells in the G1 phase (p < 0.05) and 10.4% decrease in the proportion of cells in the S phase (p < 0.05), compared to the DMSO group. However, treating the cells with TPEN + 1.25 μM of zinc reduced proportion of cells in the G1 phase by 4.3% (p < 0.05), while increased the proportion of cells in the S phase by 5.1% (p < 0.05), compared to the TPEN group. Similarly, treating the cells with TPEN + 2.5 μM and 5 μM of zinc reduced the proportion of cells in the G1 phase by 5.1% and 5.7%, respectively; while the proportion of cells in the S phase was increased by 7.2% and 7.7%, respectively, compared to the TPEN group. The proportion of cells in the G1 and S phases was not affected by increase in the level of zinc replenishment from 1.25 to 5 μM. The proportion of cells in the G2/M phase was not affected by treatments. Similarly, after culturing the cells for 48 hours, the proportion of cells in the G1 and S phases was not affected by the inclusion of DMSO as compared to the DMEM group, as showed in Figure 3.2 and Table 3.1. Treating the cells with TPEN increased the 43  proportion of cells in the G1 phase by 7.4%, but reduced the proportion of cells in the S phase by 16.0%, compared to the DMSO group. However, treating the cells with TPEN and zinc at 1.25, 2.5, or 5 μM reduced the proportion of cells in the G1 phase by 5.4%, 6%, and 5.1%, respectively (p < 0.05), and increased the proportion of cells in the S phase by 14.1%, 16.8%, and 15.9%, respectively (p < 0.05), compared to the TPEN group. Further increase in the level of zinc replenishment from 1.25 μM to 2.5 μM or 5 μM had no further effect on the proportion of cells in the G1 and S phases compared to the TPEN group. The proportion of cells in the G2/M phase was not affected by treatments. TPEN treatment in the presence of quiescence Cell cycle distribution in cells treated with TPEN with the induction of quiescence was shown in Figure 3.3 and Table 3.2. After culturing the cells in the growth medium (DMEM group) for 24 hours, there were 47.2% and 40.2% of the cells in the G1 and S phase, respectively. Inclusion of DMSO in the culture medium had no effect on the proportion of cells in the G1 and S phases, showing that DMSO had no effect on the cell cycle progression. Treating the cells with TPEN for 24 hours resulted in 41.0% increase in the proportion of cells in the G1 phase (p < 0.05) and 52.9% decrease in the proportion of cells in the S phase (p < 0.05), compared to the DMSO group. However, treating the cells with TPEN + 1.25 μM of zinc reduced proportion of cells in the G1 phase by 25.3% (p < 0.05); while the proportion of cells in the S phase was increased by 101.6%, compared to the TPEN group. Similarly, treating the cells with TPEN + 2.5 and 5 μM of zinc reduced the proportion of cells in the G1 phase by 26.1% and 24.4% respectively (p < 0.05); while the proportion of cells in the S phase was increased by 103.2% and 98.9% respectively (p < 0.05), compared to the TPEN group. The proportion of cells in the G1 and S phases was not affected by the increase in the level of zinc replenishment from 1.25 to 5 μM. The proportion of cells in 44  the G2/M phase was not affected by the treatments.  3.1.2 TPEN-induced zinc depletion altered miRNA expression profile   Treating 3T3 cells with TPEN altered the expression of a total of 184 isoforms of miRNAs, compared to the DMSO group (Table A.1). Among these 184 isoforms of miRNA, only 28 miRNAs were detected to have the signal intensity higher than the cut-off level of 500 for at least one individual sample group. Among the 28 miRNAs, three of them were down-regulated; while 25 miRNAs were up regulated. The differential expression of miRNAs induced by TPEN-induced zinc depletion with a minimal fold change of 1.5 were presented in Table 3.3. The miR-464a-3p showed the greatest fold change of 4.01 in the TPEN-treated groups compared to control. The only one miRNA with a negative fold change is miR-145b, of which the signal intensity in both DMSO control and TPEN treated groups were relatively high. The expression of 158 miRNAs was significantly affected in the TPEN-treated group with zinc replenishment compared to the TPEN-induced zinc depletion (Table A.2). The majority of the affected miRNAs were demonstrated to have a low signal intensity (< 500). Only 33 miRNAs showed the signal intensity higher than the cut-off level of 500 for at least one individual sample group. 12 miRNAs out of these 33 were down-regulated; while the rest were up regulated. As showed in Table 3.4, the differential expression of miRNAs with a minimal fold change of 1.5 was presented for the evaluation of the effect of zinc replenishment on the miRNAs expression. Applied to the criteria with a minimal fold change of 1.5 compared to the control, miR-212-3p showed the greatest change with a negative fold change, due to zinc replenishment. In addition, zinc replenishment was observed to cause an expression decrease of the other two 45  miRNAs, miR-132-3p and let-7e-3p, with a fold change of 2.88 and 2.01, respectively. miR-293-3p showed the greatest increase of its expression in the TPEN-treated group with zinc replenishment. Among the miRNAs with increased expression levels induced by zinc replenishment, miR-1929-5p, miR-3084-5p, miR-423-5p, miR-504-3p, miR-3099-3p, miR-145b, and miR-6368 showed relatively high abundance (signal intensity > 1000) in zinc replenishment groups. In summary, four miRNAs, with expression level significantly affected, were in common when compared among the DMSO control, TPEN-treated groups with and without zinc replenishment (Table 3.3 and 3.4). Therefore, these four miRNAs were selected as the potential targets for the subsequent validation experiment. Zinc depletion increased the abundance of miR-132-3p, miR-212-3p and let-7e-3p with a fold change of 2.92, 1.73, and 1.63; while zinc replenishment decreased the abundance of these miRNAs with a fold change of 2.88, 3.04, and 2.01, respectively (Figure 3.4, 3.5 and 3.6). On the other hand, TPEN-induced zinc depletion negatively impacted the expression level of miR-145b with a fold change of 1.93; while zinc replenishment brought back the expression level with an increased fold change of 1.71 (Figure 3.7).   3.2 Discussion  TPEN is a membrane permeable divalent cation chelator with a higher affinity for zinc and a lower affinity for other divalent cations, such as calcium, cooper, and iron (Arslan et al., 1985). It has been documented that Zinquin is a membrane permeable zinc specific fluorescent probe with an emission florescence at 482-488 nm. Upon binding to zinc, Zinquin emits a florescence and the intensity of Zinquin-dependent fluorescence is a measurement of the abundance of the labile intracellular zinc. Treating cells (i.e. MDA-46  MB-231 cells) with TPEN reduces the intensity of the Zinquin-dependent fluorescence (Wu, 2003). However, this TPEN-induced reduction in the intensity of the Zinquin-dependent fluorescence can be removed by the addition of zinc, but not calcium or other divalent cations (Zalewski et al., 1993; Andrews et al., 1995). These observations indicate that TPEN chelation results in a reduction in abundance of LIPZ; while zinc replenishment to zinc-depleted cells increases the abundance of LIPZ. In this study, 3T3 cells were treated with TPEN to deplete the intracellular zinc, which was different from the preparation of low-zinc medium adopted by the study of Paski and Xu (2001) in which FBS was treated with Chelex-100 followed by adding Chelex-100 treated FBS to the medium. While TPEN treatment was directly applied to the 3T3 cells in this study to chelate the intracellular zinc. In response to TPEN treatment, the proportion of cells in S phase was significantly reduced; while the proportion of cells in the G1 phase was increased when compared to the control cells (DMSO group). Further, zinc replenishment resulted in a significant reduction in the proportion of cells in the G1 phase and a significant increase in the proportion of cells in the S phase, compared to the cells treated with TPEN alone. These observations collectively suggest that TPEN-induced zinc depletion inhibited cell proliferation in 3T3 fibroblast through causing G1/S arrest. In cell cycle, the S phase is where DNA replication is taken place. Since zinc depletion considerably decreased the proportion of cells in the S phase, the results obtained in my thesis research project indicated the great importance of zinc in DNA synthesis. This result is consistent with the previous reports. Chester and co-worker (1990; 1995) reported that chelation of zinc by the DTPA, an extracellular zinc chelator, resulted in a decreased mRNA level of thymidine kinase and its enzymatic activity, leading to the down-regulated DNA synthesis. In addition, the results obtained in this 47  study are also in accordance with the findings obtained from our previous study showing that DNA synthesis and cell proliferation of 3T3 cells are negatively impacted when cells were cultured in a low-zinc medium; while the addition of zinc (5, 10, 20 μmol/L) to the low zinc medium resulted in a zinc dependent stimulation of DNA synthesis and cell proliferation (Paski and Xu, 2001). However, in this study, cell cycle distribution was determined by evaluating the cell proliferation status, which provides a more precise indication of the effect of zinc on cell proliferation than cell counting that was the technique used in the previous study. Further, this zinc depletion-induced inhibition of cell proliferation of 3T3 fibroblasts is also supported by the earlier zinc-depletion studies using TPEN. For example, TPEN treatment has been shown to reduce cell proliferation in lymphoblasts (Prasad et al., 1996) and pancreatic cancer cells (Donadelli et al., 2008). It is interesting to note that, in this study, induction of quiescence resulted in a greater inhibition in cell proliferation in response to TPEN treatment compared to the cells treated with TPEN without the induction of quiescence. The possible reason is that induction of quiescence was employed to maintain all the cells at the same starting line prior to the treatment, which ensured all the cells equally exposed to the treatments. As for the cutoff of fold change in miRNA microarray, the cutoff fold change is normally set at 2; however, there is only one miRNA in common between the treatments with a fold change larger than 2. Therefore, the criterion was broaden to 1.5 to include more potential targets. miRNA expression was evaluated by miRNA microarray assay, which illustrated that the abundance of miR-132-3p was enhanced by zinc-depleted suppression of cell proliferation with a fold change of 2.92; while the expression of miR-132-3p was decreased when zinc was replenished with a fold change of 2.88. Decreased expression of miR-132-3p has been previously demonstrated in cancers and the tumor suppressive role of miR-132-3p in these cancers have also been reported. For example, 48  miR-132-3p was down-regulated in ductal carcinoma in situ (DCIS), a common type of non-invasive breast cancer, compared to the expression level of miR-132-3p in the normal breast tissue (Li et al., 2013). In addition, decreased cell viability was observed in the MCF-7, MDA-MB-231, and BT-549 breast cancer cell lines when those cell lines were transfected with pre-miR-132 mimics, a precursor of miR-132. Thus, miR-132 appears to inhibit cell proliferation in breast cancer cells. In prostate cancer, significant down-regulation of miR-132-3p expression was also detected (Formosa et al., 2013). Decreased miR-132-3p expression was associated with metastasis and cancer progression. Transfection of PC-3 prostate cancer cells with pre-miR-132 mimic resulted in the detachment of cells from the culture dishes at 72 hours, indicating decreased cell proliferation which was followed by increased apoptosis at 96 hours. In this study, increased expression of miR-132-3p was observed during zinc-depleted suppression of cell proliferation, showing that miR-132-3p may potentially play an anti-proliferative role in zinc-mediated cell proliferation in 3T3 fibroblasts. The microarray results also indicated that zinc depletion-induced cell growth arrest was associated with an increased expression of miR-212-3p with a fold change of 1.73. The abundance of miR-212-3p was lessened with a fold change of 3.04 when zinc replenishment substantially alleviated the cell cycle arrest. As for miR-212, a significant expression decrease was examined in gastric carcinoma cells (Wada et al., 2010). Transfection of the precursor miR-212 was demonstrated to induce a decreased growth of gastric carcinoma cells. The study carried out by Jiping et al. (2013) also showed that overexpression of miR-212 significantly inhibited the cancer cell proliferation by decreasing expression of retinoblastoma binding protein 2 (RBP2) which is highly expressed in gastric cancer, whereas, knockdown of miR-212 promoted RBP2 expression resulting in enhanced cell proliferation. In hepatocellular carcinoma, overexpression of 49  miR-212 up-regulated cyclin-dependent kinase inhibitor which coincided with inhibited cell proliferation and induced cellular senescence (Liang et al., 2013). These observations collectively suggest that miR-212 plays an anti-proliferative role in cell proliferation which was in agreement with the finding obtained by this study that miR-212-3p was up-regulated in the zinc depletion-induced suppression of 3T3 cell proliferation. Abundance of let-7e-3p was elevated with a fold change of 1.63 by zinc depletion; while reduced abundance of let-7e-3p was observed when zinc was added back. let-7e-3p has been reported to be related to reduced cell viability and induction of apoptosis in KPL-4 and JIMT-1 breast cancer cell lines (Aure et al., 2013). It has been demonstrated that let-7e has the tumor suppressive function in neuroblastoma cells (Buechner et al., 2011). Overexpressed let-7e by transfection of miRNA mimic was able to inhibit proliferation and clonogenic growth of neuroblastoma cells. Meanwhile, let-7e was proven to be a strong negative regulator of MYCN expression which is an oncogene (Buechner et al., 2011). In our study, let-7e-3p was showed to be up-regulated during cell proliferation suppression by zinc depletion suggesting the tumor suppressive role of let-7e-3p. Unlike miR-132-3p, miR-212-3p, and let-7e-3p, the expression level of miR-145b was restrained when zinc in the culture medium was depleted by TPEN; while the miR-145b expression was recovered when zinc was supplemented in the TPEN-treated culture medium to eliminate the zinc-depleting effect. Interestingly, it is reported by several studies that miR-145 inhibited cell proliferation and served as an anti-proliferative role in cancer cells, including human gastric cancer cells SGC7901, MKN45 and BCG823 (Qiu et al., 2014), non-small cell lung cancer cell A549 (Hu et al., 2014), prostate cancer cell PC-3 (Xie et al., 2014), human osteosarcoma cells (HOS, Saos-2, MG-63, and U2OS; Li et al., 2014), and renal carcinoma cells (Lu et al., 2014). Reduced abundance of miR-50  145b observed in this study is different from the reported findings by others. It can be possibly explained by that the cell line used in my experiments was normal fibroblasts instead of the cancerous cells which were used by the studies discussed above. In summary, zinc depletion suppressed cell proliferation in mouse fibroblast 3T3 cells. Zinc depletion increased the abundance of miR-132-3p, miR-212-3p, and let-7e-3p; while the expression of miR-145b was negatively impacted by zinc depletion. Based on the previously demonstrated role of miR-132-3p, miR-212-3p, and let-7e-3p in suppressing cell proliferation, these miRNAs possibly involve in mediation of cell proliferation suppression induced by zinc depletion. However, further research is needed to verify their exact roles in zinc-mediated cell proliferation as well as the detailed mechanisms. Interestingly, miR-145b displayed a proliferative effect based on our results, which is inconsistent with the findings reported from other cancer cells. Taken together, based on the expression profile of miRNAs, this study provided the evidence showing a potential association between miRNA expression and zinc-depletion-induced cell proliferation in mouse fibroblast 3T3 cells. Considering the technical limitations associated with the miRNA microarray, the effects of zinc-depletion and zinc replenishment on the expression levels of miRNAs need to be validated using more quantitative techniques, such as qRT-PCR. For example, the target specific cDNA was generated from mature miRNA using target specific reverse transcription primers, followed by real-time PCR, which was carried out using target specific forward and reverse primers together with TaqMan probes designed for the target miRNA. The reaction mixture contained a fluorescent reporter dye and quencher dye. During the extension of the amplified DNA, the probe is cleaved, when the quencher is separated from the reporter dye resulting in the emission of a detectable fluorescent signal. In addition, their specific functional roles in the regulation of zinc-mediated cell 51  proliferation in mouse fibroblast 3T3 cells remains to be elucidated.  52                    (A)                            (B)                            (C)                                                        (D)                            (E)                            (F)                             Figure 3.1 Effects of 24 h zinc-depletion and zinc replenishment on cell cycle progression in 3T3 cells without quiescence induction Figures shown above are the representative flow cytometry plots for each of the treatment groups. (A) overall control (DMEM growth medium only), (B) control (DMSO only), (C) TPEN (2.5 μM), (D) TPEN + zinc at 1.25 μM, (E) TPEN + zinc at 2.5 μM, (F) TPEN + zinc at 5 μM for 24 hours without induction of quiescence.      53                           (A)                            (B)                            (C)                                                     (D)                            (E)                            (F)                            Figure 3.2 Effects of 48 h zinc-depletion and zinc replenishment on cell cycle progression in 3T3 cells without quiescence induction Figures shown above are the representative flow cytometry plots for each of the treatment groups. (A) overall control (DMEM growth medium only), (B) control (DMSO only), (C) TPEN (2.5 μM), (D) TPEN + zinc at 1.25 μM, (E) TPEN + zinc at 2.5 μM, (F) TPEN + zinc at 5 μM for 48 hours without induction of quiescence.                             (A)                            (B)                            (C) 54                                                         (D)                            (E)                            (F)                            Figure 3.3 Effects of 24 h zinc-depletion and zinc replenishment on cell cycle progression in 3T3 cells upon quiescence induction Figures shown above are the representative flow cytometry plots for each of the treatment groups. (A) overall control (DMEM growth medium only), (B) control (DMSO only), (C) TPEN (2.5 μM), (D) TPEN + zinc at 1.25 μM, (E) TPEN + zinc at 2.5 μM, (F) TPEN + zinc at 5 μM for 24 hours upon induction of quiescence. 55   Figure 3.4 Effects of zinc-depletion and zinc replenishment on the abundance of miR-132-3p in 3T3 cells 3T3 cells were treated with DMSO only (control), TPEN (2.5 μM in DMSO), or TPEN plus zinc at 1.25 μM for 24 h with quiescence induction. Values are means ± SD (n=4). Lower case letters indicate the significant difference between DMSO control and TPEN groups. Upper case letters indicate the significant difference between TPEN and TPEN plus zinc groups. Different letters indicate significant difference between the treatments (p < 0.05).   ba;AB06001,2001,8002,4003,000  Control  TPEN  TPEN + ZnSignal intensityTreatment groups 56   Figure 3.5 Effects of zinc-depletion and zinc replenishment on the abundance of miR-212-3p in 3T3 cells 3T3 cells were treated with DMSO only (control), TPEN (2.5 μM in DMSO), or TPEN plus zinc at 1.25 μM for 24 h with quiescence induction. Values are means ± SD (n=4). Lower case letters indicate the significant difference between DMSO control and TPEN groups. Upper case letters indicate the significant difference between TPEN and TPEN plus zinc groups. Different letters indicate significant difference between the treatments (p < 0.05).   ba;AB0200400600800  Control  TPEN  TPEN + ZnSignal intensityTreatment groups 57   Figure 3.6 Effects of zinc-depletion and zinc replenishment on the abundance of let-7e-3p in 3T3 cells 3T3 cells were treated with DMSO only (control), TPEN (2.5 μM in DMSO), or TPEN plus zinc at 1.25 μM for 24 h with quiescence induction. Values are means ± SD (n=4). Lower case letters indicate the significant difference between DMSO control and TPEN groups. Upper case letters indicate the significant difference between TPEN and TPEN plus zinc groups. Different letters indicate significant difference between the treatments (p < 0.05).   ba;AB0200400600800  Control  TPEN  TPEN + ZnSignal intensityTreatment groups 58   Figure 3.7 Effects of zinc-depletion and zinc replenishment on the abundance of miR-145b in 3T3 cells 3T3 cells were treated with DMSO only (control), TPEN (2.5 μM in DMSO), or TPEN plus zinc at 1.25 μM for 24 h with quiescence induction. Values are means ± SD (n=4). Lower case letters indicate the significant difference between DMSO control and TPEN groups. Upper case letters indicate the significant difference between TPEN and TPEN plus zinc groups. Different letters indicate significant difference between the treatments (p < 0.05).   ab;BA01,2002,4003,6004,8006,000  Control  TPEN  TPEN + ZnSignal intensityTreatment groups 59  Table 3.1 Effects of zinc-depletion and zinc replenishment on cell cycle progression in 3T3 cells without the induction of quiescence*   Treatment                            Distribution of cells in the cell cycle (%)                  24 hours        48 hours G1 S G2/M G1 S G2/M  DMSO  44.6 ± 0.6 b 46.2 ± 0.6 a 7.3 ± 0.7 63.4 ± 0.8 b 26.9 ± 0.9 a 7.9 ± 0.7 TPEN  48.8 ± 0.7 a;A 41.4 ± 0.5 b;C 7.3 ± 0.7 68.1 ± 0.5 a;A 22.6 ± 0.4 b;B 7.4 ± 0.5 TPEN + Zn 1.25 46.7 ± 1.2 B 43.5 ± 0.8 B 7.2 ± 0.6 64.4 ± 0.8 B 25.8 ± 1.0 A 7.6 ± 0.6 TPEN + Zn 2.5 46.3 ± 1.3 B 44.4 ± 0.4 AB 7.1 ± 0.4 64.0 ± 0.8 B 26.4 ± 0.8 A 7.6 ± 0.6 TPEN + Zn 5 46.0 ± 0.5 B 44.6 ± 0.6 A 7.1 ± 0.5 64.6 ± 0.6 B 26.2 ± 0.2 A 7.3 ± 0.4  *3T3 cells were treated with DMSO only (control), TPEN (2.5 μM), or TPEN + zinc at 1.25 (TPEN + Zn 1.25), 2.5 (TPEN + Zn 2.5) or 5 (TPEN + Zn 5) μM for 24 or 48 hours without the induction of quiescence. The values are means ± SD (n = 6). Lower case letters indicate the significant difference between DMSO control and TPEN treated groups. Upper case letters indicate the significant difference among TPEN treated and TPEN plus zinc (1.25, 2.5, and 5 μM) groups. Different letters indicate significant difference between comparable treatments (p < 0.05).   60  Table 3.2 Effects of zinc-depletion and zinc replenishment on cell cycle progression in 3T3 cells upon the induction of quiescence*  Treatment Distribution of cells in the cell cycle (%) G1 S G2/M  DMSO 47.3 ± 1.0 b 39.5 ± 0.8 a 10.3 ± 0.6 TPEN 66.7 ± 1.1 a;A 18.6 ± 1.9 b;B 10.6 ± 0.6 TPEN + Zn 1.25 49.8 ± 1.1 B 37.5 ± 0.9 A 9.8 ± 0.7 TPEN + Zn 2.5  49.3 ± 1.6 B 37.8 ± 1.8 A 10.1 ± 0.7 TPEN + Zn 5  50.4 ± 0.6 B 37.0 ± 0.5 A 10.3 ± 0.5  *Upon the induction of quiescence, 3T3 cells were treated with DMSO only (control), TPEN (2.5 μM), or TPEN + zinc at 1.25 (TPEN + Zn 1.25), 2.5 (TPEN + Zn 2.5) or 5 (TPEN + Zn 5) μM for 24 hours with the induction of quiescence. The values are means ± SD (n = 6). Lower case letters indicate the significant difference between DMSO control and TPEN treated groups. Upper case letters indicate the significant difference among TPEN treated and TPEN plus zinc (1.25, 2.5, and 5 μM) groups. Different letters indicate significant difference between comparable treatments (p < 0.05).   61  Table 3.3 Differential expression of miRNAs induced by 24 h zinc-depletion in 3T3 cells*  miRNAs DMSO TPEN Fold change p-value Signal intensity mmu-miR-465a-3p 17 ± 8 266 ± 207 4.01 4.78E-03 mmu-miR-686 39 ± 26 413 ± 283 3.42 3.12E-03 mmu-miR-1963 33 ± 10 329 ± 252 3.31 6.26E-03 mmu-miR-3083-3p 37 ± 22 367 ± 267 3.30 4.91E-03 mmu-miR-323-5p 40 ± 19 391 ± 162 3.29 6.30E-04 mmu-miR-7089-5p 39 ± 10 347 ± 325 3.14 3.09E-02 mmu-miR-7656-3p 33 ± 21 291 ± 302 3.14 2.44E-02 mmu-miR-7662-3p 33 ± 13 283 ± 218 3.11 1.54E-02 mmu-miR-6959-3p 49 ± 6 400 ± 242 3.03 1.23E-02 mmu-miR-193b-5p 32 ± 21 253 ± 189 2.97 6.82E-03 mmu-miR-7648-3p 52 ± 15 403 ± 257 2.95 1.65E-02 mmu-miR-132-3p 220 ± 23 1666 ± 943 2.92 3.18E-02 mmu-miR-3067-3p 67 ± 56 359 ± 142 2.41 9.82E-03 mmu-miR-7653-5p 79 ± 28 355 ± 251 2.16 2.18E-02 mmu-miR-6906-5p 88 ± 44 374 ± 186 2.09 8.56E-03 mmu-miR-212-3p 143 ± 37 474 ± 229 1.73 1.37E-02 mmu-let-7e-3p 177 ± 161  548 ± 209 1.63 3.80E-02 mmu-miR-714 196 ± 35 566 ± 222 1.53 1.01E-02 mmu-miR-145b 5621 ± 112 1476 ± 867 -1.93 1.50E-02  *Upon the induction of quiescence, 3T3 cells were treated with DMSO only (control), or TPEN (2.5 μM) for 24 hours with the induction of quiescence. The signal intensity values are means ± SD (n=4). Cut-off criteria was set at magnitude fold change > 1.5, signal intensity >500 in at least one individual sample group, and p < 0.05.   62  Table 3.4 Differential expression of miRNAs induced by 24 h zinc replenishment in 3T3 cells*  miRNAs TPEN TPEN + Zn Fold change p-value Signal intensity mmu-miR-293-3p 92 ± 93 633 ± 257 2.78 3.56E-02 mmu-miR-505-5p 74 ± 90 461 ± 119 2.63 4.14E-02 mmu-miR-1929-5p 221 ± 211 1343 ± 747 2.60 3.27E-02 mmu-miR-1948-3p 166 ± 152 920 ± 484 2.47 3.88E-02 mmu-miR-3084-5p 340 ± 267 1720 ±982 2.34 2.82E-02 mmu-miR-7039-5p 77 ± 67 387 ± 201 2.33 3.81E-02 mmu-miR-3087-5p 147 ± 116 683 ± 202 2.22 2.60E-02 mmu-miR-6240 191 ± 117 855 ± 327 2.17 1.62E-02 mmu-miR-1945 98 ± 55 420 ± 264 2.09 3.25E-02 mmu-miR-128-1-5p 136 ± 84 567 ± 245 2.06 1.62E-02 mmu-miR-1231-5p 191 ± 115 735 ± 210 1.95 1.40E-02 mmu-miR-423-5p 950 ± 623 3555 ± 1516 1.90 2.13E-02 mmu-miR-504-3p 273 ± 189 1008 ± 261 1.89 3.53E-02 mmu-miR-3572-5p 209 ± 207 771 ± 168 1.88 4.38E-02 mmu-miR-3099-3p 510 ± 359 1858 ± 630 1.86 2.14E-02 mmu-miR-6989-5p 134 ± 92 483 ± 105 1.85 3.28E-02 mmu-miR-7080-5p 162 ± 106 553 ± 124 1.77 3.85E-02 mmu-miR-145b 1476 ± 867 4837 ± 277 1.71 2.04E-02 mmu-miR-6368 324 ± 196 1045 ± 387 1.69 2.81E-02 mmu-miR-212-3p 474 ± 229 58 ± 31 -3.04 1.53E-03 mmu-miR-132-3p 1666 ± 943 226 ± 41 -2.88 3.34E-02 mmu-let-7e-3p 548 ± 209 136 ± 23 -2.01 4.81E-03  *Upon the induction of quiescence, 3T3 cells were treated with TPEN (2.5 μM) or TPEN + zinc at 1.25 μM (TPEN + Zn) for 24 hours with the induction of quiescence. The signal intensity values are means ± SD (n=4). Cut-off criteria was set at magnitude fold change > 1.5, signal intensity >500 in at least one individual sample group, and p < 0.05.   63  Chapter 4 Conclusions, Limitations, and Future Directions  4.1 Conclusions  Zinc is essential for cell proliferation. In this study, cell proliferation upon TPEN-induced zinc depletion was assessed. It was demonstrated that TPEN-induced zinc depletion caused G1/S arrest; while the subsequent zinc replenishment significantly alleviated cell proliferation suppression by bringing back the S phase population to the same level of the control (DMSO control). The effect of zinc depletion on the global miRNA expression profile in 3T3 cells was investigated in this study. According to the literature, this is the first study to investigate the effect of zinc depletion-induced suppression of cell proliferation on the miRNA expression in vitro. TPEN-induced zinc depletion altered the miRNA expression profile in 3T3 fibroblasts. There were four miRNAs of which the expression levels were significantly affected. Zinc depletion increased the abundance of miR-132-3p, miR-212-3p, and let-7e-3p with a fold change of 2.92, 1.73, and 1.63 respectively; while zinc replenishment reduced the abundance of these miRNAs with a fold change of 2.88, 3.04, and 2.01 respectively. On the other hand, TPEN-induced zinc depletion negatively impacted the expression level of miR-145b with a fold change of 1.93; while zinc replenishment brought back the expression level with an increased fold change of 1.71. Zinc is known to exert a broad range of physiological functions, including its great essence in growth (MacDonald, 2000). However, the diverse physiological functions of zinc have not been thoroughly understood from the molecular perspective. The main findings derived in this study can serve as novel evidence to support the influence of cellular zinc status on the miRNA expression in 3T3 fibroblasts. miRNAs are a group of 64  gene regulators targeting at thousands of proteins and enzymes which are responsible for normal cellular functions. Thus, the results obtained from our study provide new and innovative insights for future research on the mechanisms of zinc’s effect on cell proliferation and the regulative role of miRNAs in the zinc-mediated cell proliferation.   4.2 Limitations  There are several limitations of this study. The first limitation is the in vitro system with only one cell line examined. In addition, being a cell line, it lacks the complexity as that of the whole tissue or whole body levels. Therefore, it is essential to validate the observations using other cell lines or at the whole body level using experimental rodent models. The second limitation is that the miRNA expression level cannot be determined accurately and solely by miRNA microarray assay due to the high inherent background noise associated with this assay (Koshiol et al., 2010; Pritchard et al., 2012). miRNAs, unlike mRNAs, lack a common sequence, such as poly A which can be used for selective enrichment. A common sequence is of great importance for the accurate determination of miRNAs because miRNAs represent only a small fraction of the total RNA and there are considerable background interferences which consist of different kinds of noise caused by other RNA hybridizing. In addition, miRNAs within a family can differ from each other by as little as a single nucleotide, making the ability to identify the miRNAs with such slight differences important. This type of inherent background noise results in both false positive and false negative results. The two major causes of the inaccuracy associated with miRNA microarray assay are complicated sample preparation process and cross-hybridization of probes with related transcripts or similar sequence. Another challenge for 65  profiling hundreds of miRNAs in parallel is that, due to their short length, the variance in miRNA GC content leads to a wide variance in melting temperatures for annealing reactions, creating miRNA-specific biases (Pritchard et al., 2012). Therefore, it is essential and critical to have the results to be validated by at least one of the alternative methods. For example, quantitative real-time PCR, possesses greater sensitivity and higher specificity than miRNA microarray assay, because the larger dynamic range of stem-loop is adopted by quantitative real-time PCR and the used stem-loop primers can distinguish between miRNAs which differ by one nucleotide. In addition, the studies carried out to determine the possible regulation of miRNAs in zinc-mediated cell proliferation have only focused on the theoretical investigation so far. No practical applications of the findings have been addressed.  4.3 Future directions  As described in Section 4.2, the expression levels of the four target miRNAs identified by the miRNA microarray needs to be further validated. Quantitative real-time PCR (qRT-PCR) can be a potential and useful technique to confirm the expression levels of miR-132-3p, miR-212-3p, let-7e-3p, and miR-145b, since the sensitivity and specificity of qRT-PCR are substantially higher than those of miRNA microarray assay. RNA sequencing, possessing high accuracy in distinguishing miRNAs of very similar sequences, including isoMIRNAs, can be another alternative technique for the validation of the expression levels of target miRNAs. In addition to the confirmation of miRNAs expression levels, more research attempts are required to investigate the functions of specific miRNAs to assess their involvement in zinc-mediated cell proliferation by establishing the role of miRNA up-regulation or 66  down-regulation in 3T3 fibroblasts. Some of the miRNAs which significantly altered by zinc depletion in our research have been reported to serve as the regulators in other types of cells. However, the functional significances of these miRNAs in 3T3 fibroblasts are still unclear. It is possible to overexpress or knockdown the selected miRNA by transfection precursor or anti-sense miRNA of the selected miRNA. The cell proliferation status of the transduced 3T3 fibroblasts can be assessed to identify whether the selected miRNA has the promoting or suppressive effect on cell proliferation. For instance, it would be interesting to investigate the role of miR-132-3p up-regulation in 3T3 cells in higher proliferation status, since miR-132-3p has been reported to be associated with decreased cell viability and cell proliferation in breast cancer cells and prostate cancer cells. Finally, it would be worthwhile to examine the pathways involved in miRNA expression during zinc-depletion induced suppression of cell proliferation. It has been demonstrated that cytotoxic intracellular zinc level reduced the expression of the proteins responsible for the miRNA biogenesis and stability. Therefore, future work might focus on the possible relationship between the miRNAs whose expression levels altered by zinc depletion and the proteins responsible for the miRNA biogenesis and stability. In addition, since miRNAs are closely related to the expression of numerous proteins involving in diverse cellular activities, it would be promising to identify the proteins of which the expression levels are affected correspondingly by the altered miRNA during zinc-regulated cell proliferation. In summary, the results of this study can be used as a reference for future investigation on the possible and potential interaction between zinc and miRNAs. Our study provides novel insights that miRNAs probably involves in the zinc-regulated cell proliferation, which creates new perspective and direction for future research on the 67  mechanism of the importance of zinc in growth.  68  Bibliography Aksamitiene, E., Kiyatkin, A., & Kholodenko, B. N. (2012). Cross-talk between mitogenic Ras/MAPK and survival PI3K/Akt pathways: A fine balance. Biochemical Society Transactions, 40(1), 139-146.  Alder, H., Taccioli, C., Chen, H., Jiang, Y., Smalley, K. J., Fadda, P., et al. (2012). 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Methods in Molecular Biology (Clifton, N.J.), 382, 287-312.   96  Appendix  Table A.1 Differential expression of miRNAs induced by 24 h zinc depletion in 3T3 cells* *Upon the induction of quiescence, 3T3 cells were treated with DMSO only (control), or TPEN (2.5 μM) for 24 hours with the induction of quiescence. The signal intensity values are means ± SD (n=4) and p < 0.05. miRNAs DMSO TPEN Fold change p-value Signal intensity Mean SD Mean SD mmu-miR-323-5p 40 19 391 162 3.29 6.30E-04 mmu-miR-686 39 26 413 283 3.42 3.12E-03 mmu-miR-465a-3p 17 8 266 207 4.01 4.78E-03 mmu-miR-3083-3p 37 22 367 267 3.30 4.91E-03 mmu-miR-1963 33 10 329 252 3.31 6.26E-03 mmu-miR-193b-5p 32 21 253 189 2.97 6.82E-03 mmu-miR-421-3p 277 26 452 79 0.71 8.33E-03 mmu-miR-6906-5p 88 44 374 186 2.09 8.56E-03 mmu-miR-3067-3p 67 56 359 142 2.41 9.82E-03 mmu-miR-714 196 35 566 222 1.53 1.01E-02 mmu-miR-6959-3p 49 6 400 242 3.03 1.23E-02 mmu-let-7d-3p 359 122 814 269 1.18 1.29E-02 mmu-miR-212-3p 143 37 474 229 1.73 1.37E-02 mmu-miR-145b 5,621 112 1,476 867 -1.93 1.50E-02 mmu-miR-7662-3p 33 13 283 218 3.11 1.54E-02 mmu-miR-7648-3p 52 15 403 257 2.95 1.65E-02 mmu-miR-7653-5p 79 28 355 251 2.16 2.18E-02 mmu-miR-7656-3p 33 21 291 302 3.14 2.44E-02 mmu-miR-5100 522 83 908 238 0.80 2.52E-02 mmu-miR-7089-5p 39 10 347 325 3.14 3.09E-02 mmu-miR-132-3p 220 33 1,666 943 2.92 3.18E-02 mmu-miR-199b-5p 5,644 470 4,770 381 -0.24 3.46E-02 mmu-miR-151-5p 1,254 84 1,062 104 -0.24 3.50E-02 mmu-miR-361-5p 758 52 855 45 0.17 3.67E-02 mmu-miR-3960 1,444 29 1,828 233 0.34 3.80E-02 mmu-let-7e-3p 177 161 548 209 1.63 3.80E-02 mmu-miR-709 20,878 1,004 24,181 2,051 0.21 3.85E-02 mmu-miR-7671-3p 128 57 337 181 1.40 4.94E-02 Following transcripts are statistically significant but have low signals (signal < 500) mmu-miR-466d-3p 294 25 155 19 -0.92 3.77E-04 mmu-miR-6966-3p 17 4 225 91 3.72 4.91E-04 mmu-miR-129-5p 32 7 289 131 3.19 7.56E-04 mmu-miR-211-5p 6 2 26 7 2.21 9.01E-04 mmu-miR-8109 34 8 91 16 1.42 9.02E-04 mmu-miR-8120 32 13 313 160 3.28 1.04E-03 97  miRNAs DMSO TPEN Fold change p-value Signal intensity Mean SD Mean SD mmu-miR-3061-5p 17 4 67 25 2.00 1.17E-03 mmu-miR-7087-5p 11 2 27 5 1.26 1.40E-03 mmu-miR-1947-3p 11 2 47 17 2.15 1.68E-03 mmu-miR-679-5p 40 4 17 4 -1.21 2.35E-03 mmu-miR-1931 18 3 47 15 1.42 2.40E-03 mmu-miR-3057-3p 43 16 165 59 1.93 2.40E-03 mmu-miR-7089-3p 7 6 52 24 2.82 2.71E-03 mmu-miR-1934-5p 14 6 58 19 2.07 3.08E-03 mmu-miR-1943-5p 24 7 233 140 3.27 3.39E-03 mmu-miR-218-2-3p 13 5 56 22 2.09 3.69E-03 mmu-miR-7032-5p 12 5 53 25 2.12 4.16E-03 mmu-miR-7669-3p 45 5 25 5 -0.85 4.38E-03 mmu-miR-6971-3p 45 9 103 26 1.19 4.69E-03 mmu-miR-551b-3p 6 4 56 37 3.29 4.82E-03 mmu-miR-194-2-3p 13 4 45 17 1.78 4.84E-03 mmu-miR-383-3p 22 8 67 23 1.62 5.12E-03 mmu-miR-7237-3p 13 5 120 86 3.24 5.20E-03 mmu-miR-3063-5p 9 4 48 27 2.49 5.24E-03 mmu-miR-33-3p 53 7 35 3 -0.58 5.32E-03 mmu-miR-1899 20 3 158 89 3.01 5.38E-03 mmu-miR-139-5p 15 6 53 22 1.79 5.67E-03 mmu-miR-3076-3p 20 15 191 159 3.29 5.72E-03 mmu-miR-1936 31 8 64 15 1.08 6.03E-03 mmu-miR-3077-3p 25 15 139 88 2.48 6.06E-03 mmu-miR-216a-3p 4 2 38 19 3.14 6.18E-03 mmu-miR-7228-3p 23 15 176 121 2.96 6.26E-03 mmu-miR-212-5p 14 4 34 6 1.31 6.29E-03 mmu-miR-335-5p 40 6 18 5 -1.16 6.35E-03 mmu-miR-6340 38 8 67 10 0.84 6.36E-03 mmu-miR-744-3p 88 10 64 5 -0.45 6.91E-03 mmu-miR-135a-2-3p 8 3 28 14 1.75 7.11E-03 mmu-miR-139-3p 12 2 45 16 1.92 7.28E-03 mmu-miR-6896-5p 30 22 237 152 2.99 7.97E-03 mmu-miR-3470b 24 4 59 18 1.28 8.16E-03 mmu-miR-6978-3p 22 4 69 27 1.64 8.32E-03 mmu-miR-717 8 4 112 89 3.80 8.53E-03 mmu-miR-106b-3p 218 7 133 20 -0.72 8.64E-03 mmu-miR-7036-5p 30 8 14 3 -1.14 8.79E-03 mmu-miR-6979-3p 25 5 172 88 2.76 8.88E-03 mmu-miR-344h-3p 16 4 55 27 1.77 8.97E-03 mmu-miR-7023-5p 34 13 178 103 2.39 9.13E-03 mmu-miR-6947-5p 36 5 23 3 -0.62 9.17E-03 mmu-miR-8116 28 9 81 31 1.52 9.22E-03 98  miRNAs DMSO TPEN Fold change p-value Signal intensity Mean SD Mean SD mmu-miR-6403 15 2 221 168 3.88 9.52E-03 mmu-miR-7119-5p 5 3 21 11 2.13 9.82E-03 mmu-miR-6941-3p 26 16 139 73 2.41 9.86E-03 mmu-miR-7035-3p 20 3 39 10 0.95 1.04E-02 mmu-miR-129b-3p 12 2 53 30 2.19 1.06E-02 mmu-miR-6977-5p 13 4 47 22 1.86 1.06E-02 mmu-miR-6406 8 4 39 23 2.27 1.08E-02 mmu-miR-341-3p 47 23 293 187 2.64 1.08E-02 mmu-miR-702-5p 35 6 78 25 1.13 1.08E-02 mmu-miR-16-1-3p 42 5 28 5 -0.59 1.16E-02 mmu-miR-6769b-5p 54 10 22 8 -1.29 1.25E-02 mmu-miR-216c-3p 10 6 62 48 2.63 1.27E-02 mmu-miR-466n-5p 102 30 28 14 -1.88 1.29E-02 mmu-miR-6244 3 2 35 13 3.56 1.30E-02 mmu-miR-450b-5p 26 11 8 2 -1.72 1.32E-02 mmu-miR-331-5p 24 10 102 58 2.11 1.32E-02 mmu-miR-3968 99 33 34 13 -1.54 1.35E-02 mmu-miR-6970-5p 31 6 202 132 2.68 1.38E-02 mmu-miR-7676-3p 29 4 12 4 -1.28 1.38E-02 mmu-miR-7000-3p 62 9 97 18 0.65 1.41E-02 mmu-miR-1953 4 2 27 19 2.81 1.44E-02 mmu-miR-7666-5p 20 11 98 57 2.31 1.45E-02 mmu-miR-6975-5p 61 17 31 6 -0.98 1.47E-02 mmu-miR-219b-5p 8 4 25 10 1.68 1.48E-02 mmu-miR-3071-5p 61 28 18 8 -1.73 1.54E-02 mmu-miR-691 191 46 110 17 -0.79 1.56E-02 mmu-miR-6408 4 2 32 23 2.91 1.59E-02 mmu-miR-6972-3p 6 4 32 9 2.56 1.60E-02 mmu-miR-1934-3p 32 3 72 23 1.16 1.70E-02 mmu-miR-1247-3p 23 10 5 3 -2.37 1.82E-02 mmu-miR-598-5p 6 3 24 14 1.91 1.84E-02 mmu-miR-5617-3p 25 6 42 9 0.76 1.84E-02 mmu-miR-6361 35 5 109 41 1.63 1.84E-02 mmu-miR-124-3p 23 11 7 4 -1.83 1.86E-02 mmu-miR-7671-5p 20 10 126 85 2.66 1.98E-02 mmu-miR-450a-5p 89 7 53 12 -0.75 1.99E-02 mmu-miR-7033-5p 69 10 49 7 -0.51 2.00E-02 mmu-miR-6977-3p 25 8 82 36 1.71 2.02E-02 mmu-miR-3086-3p 31 12 178 140 2.50 2.03E-02 mmu-miR-6241 21 4 191 104 3.18 2.04E-02 mmu-miR-5118 10 4 27 12 1.37 2.06E-02 mmu-miR-6963-5p 27 9 53 9 0.96 2.09E-02 mmu-miR-21c 40 23 9 5 -2.20 2.24E-02 99  miRNAs DMSO TPEN Fold change p-value Signal intensity Mean SD Mean SD mmu-miR-452-3p 24 10 9 3 -1.42 2.45E-02 mmu-miR-669b-3p 106 27 47 19 -1.19 2.50E-02 mmu-miR-1668 11 5 24 7 1.14 2.51E-02 mmu-miR-673-5p 44 7 21 9 -1.10 2.52E-02 mmu-miR-8098 32 12 80 31 1.31 2.57E-02 mmu-miR-672-3p 35 11 5 3 -2.72 2.59E-02 mmu-miR-146a-5p 310 51 222 26 -0.48 2.67E-02 mmu-miR-423-3p 44 18 21 3 -1.08 2.72E-02 mmu-miR-3068-5p 88 20 152 38 0.78 2.74E-02 mmu-miR-383-5p 7 3 46 46 2.73 2.75E-02 mmu-miR-7115-5p 6 2 33 27 2.53 2.79E-02 mmu-miR-666-5p 47 14 82 18 0.80 2.92E-02 mmu-miR-6392-3p 11 3 25 10 1.23 2.94E-02 mmu-miR-340-5p 95 8 61 13 -0.63 2.98E-02 mmu-miR-344f-5p 10 4 44 33 2.09 3.03E-02 mmu-miR-872-5p 280 18 241 19 -0.21 3.12E-02 mmu-miR-7036b-3p 13 2 26 9 1.01 3.14E-02 mmu-miR-6938-3p 11 12 105 50 3.26 3.15E-02 mmu-miR-10b-3p 55 5 26 10 -1.04 3.16E-02 mmu-miR-200a-3p 48 76 1 1 -6.12 3.17E-02 mmu-miR-497-5p 159 20 310 105 0.96 3.18E-02 mmu-miR-5128 34 5 111 53 1.70 3.23E-02 mmu-miR-8111 23 13 58 21 1.35 3.27E-02 mmu-miR-7648-5p 49 25 231 159 2.25 3.31E-02 mmu-miR-1198-3p 23 15 3 2 -2.93 3.35E-02 mmu-miR-466e-5p 159 31 63 31 -1.34 3.37E-02 mmu-let-7c-1-3p 44 51 221 154 2.33 3.38E-02 mmu-miR-703 77 19 31 17 -1.33 3.39E-02 mmu-miR-671-5p 141 8 102 17 -0.46 3.40E-02 mmu-miR-135a-1-3p 11 1 41 23 1.85 3.43E-02 mmu-miR-467c-5p 83 21 36 17 -1.20 3.45E-02 mmu-miR-466b-3p 254 26 143 44 -0.83 3.52E-02 mmu-miR-1967 62 14 33 10 -0.90 3.62E-02 mmu-miR-130b-5p 29 7 43 5 0.56 3.70E-02 mmu-miR-466a-5p 152 28 61 30 -1.31 3.78E-02 mmu-miR-5107-3p 36 10 79 41 1.13 3.84E-02 mmu-miR-6349 80 4 66 8 -0.27 3.85E-02 mmu-miR-7083-3p 61 12 86 3 0.51 3.86E-02 mmu-miR-298-5p 139 42 68 27 -1.03 3.88E-02 mmu-miR-6342 31 8 123 89 2.01 3.94E-02 mmu-miR-6990-5p 12 4 42 25 1.79 3.94E-02 mmu-miR-200b-5p 63 17 195 114 1.63 3.97E-02 mmu-miR-5622-3p 423 51 329 44 -0.36 4.00E-02 100  miRNAs DMSO TPEN Fold change p-value Signal intensity Mean SD Mean SD mmu-miR-7689-5p 90 21 57 14 -0.67 4.00E-02 mmu-miR-5134-5p 12 5 27 11 1.15 4.14E-02 mmu-miR-669i 65 12 23 14 -1.53 4.17E-02 mmu-miR-467e-5p 70 19 41 10 -0.80 4.20E-02 mmu-miR-7238-5p 22 6 131 117 2.58 4.31E-02 mmu-miR-5130 107 18 158 38 0.56 4.31E-02 mmu-miR-339-5p 62 10 42 9 -0.56 4.32E-02 mmu-miR-883a-3p 33 5 46 9 0.47 4.34E-02 mmu-miR-3967 43 31 6 2 -2.86 4.38E-02 mmu-miR-877-5p 48 11 183 113 1.92 4.46E-02 mmu-miR-3067-5p 30 24 91 17 1.59 4.50E-02 mmu-miR-7076-5p 29 8 68 29 1.23 4.50E-02 mmu-miR-7077-3p 37 10 90 45 1.30 4.51E-02 mmu-miR-30a-3p 178 17 134 23 -0.41 4.51E-02 mmu-miR-466k 23 17 5 4 -2.30 4.67E-02 mmu-miR-6902-3p 42 1 23 9 -0.84 4.76E-02 mmu-miR-582-5p 154 45 90 13 -0.77 4.79E-02 mmu-miR-1306-3p 25 2 43 14 0.81 4.83E-02 mmu-miR-344-3p 65 15 38 15 -0.80 4.92E-02 mmu-miR-7077-5p 16 1 57 44 1.82 4.95E-02 mmu-miR-6912-3p 22 6 60 34 1.42 4.99E-02                           101  Table A.2 Differential expression of miRNAs induced by 24 h zinc replenishment in 3T3 cells* *Upon the induction of quiescence, 3T3 cells were treated with TPEN (2.5 μM) or TPEN + zinc at 1.25 μM (TPEN + Zn) for 24 hours with the induction of quiescence. The signal intensity values are means ± SD (n=4) and p < 0.05. miRNAs TPEN TPEN + Zn Fold Change p-value Signal intensity Mean SD Mean SD mmu-miR-212-3p 474 229 58 31 -3.04 1.53E-03 mmu-let-7e-3p 548 209 136 23 -2.01 4.81E-03 mmu-miR-3960 1,828 233 1,330 128 -0.46 1.09E-02 mmu-let-7d-3p 814 269 379 50 -1.10 1.12E-02 mmu-miR-690 26,179 2,062 20,995 1,788 -0.32 1.24E-02 mmu-miR-1231-5p 191 115 735 210 1.95 1.40E-02 mmu-miR-5126 2,200 68 1,674 177 -0.39 1.50E-02 mmu-miR-128-1-5p 136 84 567 245 2.06 1.62E-02 mmu-miR-6240 191 117 855 327 2.17 1.62E-02 mmu-miR-6944-5p 285 124 777 293 1.45 1.91E-02 mmu-miR-145b 1,476 867 4,837 277 1.71 2.04E-02 mmu-miR-423-5p 950 623 3,555 1,516 1.90 2.13E-02 mmu-miR-3099-3p 510 359 1,858 630 1.86 2.14E-02 mmu-miR-3087-5p 147 116 683 202 2.22 2.60E-02 mmu-miR-6368 324 196 1,045 387 1.69 2.81E-02 mmu-miR-497-5p 310 105 601 176 0.95 2.82E-02 mmu-miR-3084-5p 340 267 1,720 982 2.34 2.82E-02 mmu-miR-1945 98 55 420 264 2.09 3.25E-02 mmu-miR-1929-5p 221 211 1,343 747 2.60 3.27E-02 mmu-miR-6989-5p 134 92 483 105 1.85 3.28E-02 mmu-miR-132-3p 1,666 943 226 41 -2.88 3.34E-02 mmu-miR-3473b 616 106 450 44 -0.45 3.49E-02 mmu-miR-504-3p 273 189 1,008 261 1.89 3.53E-02 mmu-miR-762 970 187 606 169 -0.68 3.55E-02 mmu-miR-293-3p 92 93 633 257 2.78 3.56E-02 mmu-miR-7045-5p 700 113 507 34 -0.47 3.75E-02 mmu-miR-7039-5p 77 67 387 201 2.33 3.81E-02 mmu-miR-7080-5p 162 106 553 124 1.77 3.85E-02 mmu-miR-1948-3p 166 152 920 484 2.47 3.88E-02 mmu-miR-505-5p 74 90 461 119 2.63 4.14E-02 mmu-miR-3572-5p 209 207 771 168 1.88 4.38E-02 mmu-miR-6239 20,248 2,215 16,928 1,292 -0.26 4.43E-02 mmu-miR-705 719 128 536 44 -0.42 4.97E-02 Following transcripts are statistically significant but have low signals (signal < 500) mmu-miR-221-5p 123 14 240 25 0.97 2.87E-04 mmu-miR-23b-5p 35 9 115 31 1.71 8.39E-04 mmu-miR-343 11 2 29 6 1.39 1.60E-03 mmu-miR-3473e 431 26 320 27 -0.43 2.34E-03 102  miRNAs TPEN TPEN + Zn Fold Change p-value Signal intensity Mean SD Mean SD mmu-miR-7064-5p 13 4 56 22 2.06 2.67E-03 mmu-miR-7005-5p 189 50 64 19 -1.57 3.66E-03 mmu-miR-7007-5p 37 9 79 9 1.12 3.83E-03 mmu-miR-125b-1-3p 126 37 309 64 1.29 3.93E-03 mmu-miR-7070-5p 59 13 168 56 1.50 4.61E-03 mmu-miR-7000-3p 97 18 175 31 0.86 5.29E-03 mmu-miR-3072-5p 268 67 116 30 -1.21 5.46E-03 mmu-miR-8100 92 26 264 83 1.52 5.77E-03 mmu-miR-7015-3p 17 7 56 17 1.73 5.98E-03 mmu-miR-6998-5p 12 5 45 13 1.93 7.39E-03 mmu-miR-3620-5p 169 55 54 19 -1.65 7.55E-03 mmu-miR-7094-1-5p 18 2 35 8 0.94 7.87E-03 mmu-miR-202-3p 12 5 41 11 1.75 8.97E-03 mmu-miR-1898 55 30 229 68 2.06 9.32E-03 mmu-miR-6937-5p 248 83 95 17 -1.38 9.61E-03 mmu-miR-671-5p 102 17 64 10 -0.68 1.02E-02 mmu-miR-704 24 4 60 22 1.34 1.11E-02 mmu-miR-6988-3p 54 24 138 43 1.36 1.23E-02 mmu-miR-6996-5p 68 44 334 156 2.30 1.25E-02 mmu-miR-351-3p 54 13 29 7 -0.91 1.25E-02 mmu-miR-212-5p 34 6 9 5 -1.99 1.27E-02 mmu-miR-6966-3p 225 91 75 24 -1.59 1.31E-02 mmu-miR-7221-3p 83 26 32 13 -1.38 1.31E-02 mmu-miR-5136 66 36 257 126 1.95 1.37E-02 mmu-miR-3076-5p 22 12 81 35 1.87 1.38E-02 mmu-miR-3474 43 8 22 7 -0.95 1.39E-02 mmu-miR-7038-3p 68 26 157 51 1.22 1.51E-02 mmu-miR-3081-5p 36 23 160 81 2.17 1.67E-02 mmu-miR-883a-3p 46 9 99 35 1.11 1.68E-02 mmu-miR-330-3p 20 9 62 19 1.63 1.70E-02 mmu-miR-7071-3p 64 38 14 7 -2.18 1.71E-02 mmu-miR-30e-3p 150 13 181 9 0.27 1.74E-02 mmu-miR-3088-3p 9 8 54 22 2.57 1.79E-02 mmu-miR-541-3p 39 7 24 5 -0.72 1.80E-02 mmu-miR-216b-5p 11 7 62 15 2.48 1.82E-02 mmu-miR-6901-5p 41 25 162 77 1.98 1.86E-02 mmu-miR-1906 124 37 246 63 0.99 1.93E-02 mmu-miR-7050-5p 93 23 47 14 -0.99 1.94E-02 mmu-miR-7083-5p 71 45 297 118 2.07 2.02E-02 mmu-miR-29c-5p 60 29 194 95 1.70 2.06E-02 mmu-miR-667-5p 31 8 53 8 0.79 2.06E-02 mmu-miR-671-3p 48 4 116 41 1.28 2.15E-02 mmu-miR-7235-5p 188 21 135 24 -0.47 2.24E-02 103  miRNAs TPEN TPEN + Zn Fold Change p-value Signal intensity Mean SD Mean SD mmu-miR-6236 32 7 54 12 0.75 2.24E-02 mmu-miR-344h-3p 55 27 22 7 -1.33 2.25E-02 mmu-miR-7047-5p 271 95 123 25 -1.14 2.37E-02 mmu-miR-346-3p 46 12 26 4 -0.84 2.38E-02 mmu-miR-222-5p 93 25 179 51 0.93 2.43E-02 mmu-miR-6997-5p 30 12 69 9 1.18 2.47E-02 mmu-miR-6715-3p 165 58 71 15 -1.21 2.52E-02 mmu-miR-6929-5p 36 7 75 25 1.07 2.52E-02 mmu-miR-759 52 32 171 76 1.73 2.54E-02 mmu-miR-339-3p 38 23 152 87 1.99 2.56E-02 mmu-miR-7049-3p 40 14 75 17 0.92 2.57E-02 mmu-miR-3093-5p 20 10 103 59 2.37 2.61E-02 mmu-miR-7652-5p 59 33 178 70 1.59 2.63E-02 mmu-miR-1892 164 28 111 22 -0.57 2.63E-02 mmu-miR-7042-5p 86 24 178 64 1.05 2.64E-02 mmu-miR-743a-5p 22 8 42 8 0.93 2.65E-02 mmu-miR-6349 66 8 44 8 -0.57 2.68E-02 mmu-miR-6340 67 10 169 67 1.33 2.68E-02 mmu-miR-21a-3p 92 66 345 130 1.91 2.69E-02 mmu-miR-551b-3p 56 37 15 4 -1.87 2.71E-02 mmu-miR-16-1-3p 28 5 52 17 0.89 2.71E-02 mmu-miR-6955-5p 24 16 93 37 1.93 2.73E-02 mmu-miR-7068-5p 29 27 211 131 2.88 2.76E-02 mmu-miR-99a-3p 67 23 123 30 0.89 2.82E-02 mmu-miR-21c 9 5 28 13 1.66 2.85E-02 mmu-miR-301b-3p 168 52 305 53 0.86 2.90E-02 mmu-miR-6939-5p 55 12 31 10 -0.86 2.92E-02 mmu-miR-7118-5p 230 37 134 38 -0.77 2.92E-02 mmu-miR-1193-3p 102 24 63 13 -0.69 2.93E-02 mmu-miR-25-5p 53 16 120 49 1.17 2.96E-02 mmu-miR-8119 126 62 310 97 1.30 3.00E-02 mmu-miR-6950-5p 27 8 9 6 -1.56 3.06E-02 mmu-miR-181c-5p 201 59 124 21 -0.69 3.25E-02 mmu-miR-192-5p 96 6 62 15 -0.64 3.32E-02 mmu-miR-409-3p 143 19 242 70 0.76 3.34E-02 mmu-miR-3472 17 10 69 19 2.02 3.36E-02 mmu-miR-216a-3p 38 19 115 62 1.61 3.40E-02 mmu-miR-451b 25 7 68 37 1.44 3.44E-02 mmu-miR-6983-5p 37 33 153 15 2.04 3.47E-02 mmu-miR-1188-5p 97 33 51 18 -0.92 3.49E-02 mmu-miR-2139 38 15 79 23 1.07 3.52E-02 mmu-miR-3473g 32 9 19 5 -0.77 3.53E-02 mmu-miR-652-5p 57 10 41 6 -0.48 3.54E-02 104  miRNAs TPEN TPEN + Zn Fold Change p-value Signal intensity Mean SD Mean SD mmu-miR-7040-5p 39 9 19 7 -1.03 3.56E-02 mmu-miR-125b-2-3p 80 73 281 84 1.81 3.59E-02 mmu-miR-215-3p 24 27 1 1 -5.33 3.61E-02 mmu-miR-8105 138 54 65 20 -1.08 3.70E-02 mmu-miR-7221-5p 6 5 27 14 2.10 3.71E-02 mmu-miR-105 14 10 56 26 1.99 3.73E-02 mmu-miR-7072-5p 148 29 99 22 -0.58 3.74E-02 mmu-miR-8104 163 27 112 25 -0.55 3.75E-02 mmu-miR-218-1-3p 45 11 28 6 -0.65 3.79E-02 mmu-miR-431-3p 20 9 58 31 1.50 3.83E-02 mmu-miR-6988-5p 39 25 138 36 1.84 3.84E-02 mmu-miR-7220-3p 16 12 63 32 1.97 3.85E-02 mmu-miR-6715-5p 22 10 45 9 1.00 3.90E-02 mmu-miR-146b-3p 17 7 35 9 1.04 3.91E-02 mmu-miR-7001-5p 45 16 96 33 1.10 3.99E-02 mmu-miR-7021-5p 38 37 198 68 2.37 4.01E-02 mmu-miR-7216-5p 47 19 91 18 0.94 4.03E-02 mmu-miR-325-5p 6 8 61 42 3.46 4.06E-02 mmu-miR-6984-3p 172 20 141 8 -0.29 4.06E-02 mmu-miR-6942-5p 200 35 138 33 -0.54 4.12E-02 mmu-miR-6931-5p 262 38 203 17 -0.36 4.14E-02 mmu-miR-6986-5p 26 23 131 44 2.35 4.28E-02 mmu-miR-6941-5p 19 11 59 31 1.65 4.30E-02 mmu-miR-6388 20 6 44 19 1.11 4.32E-02 mmu-miR-30b-3p 49 36 135 45 1.46 4.33E-02 mmu-miR-3057-3p 165 59 27 18 -2.64 4.36E-02 mmu-miR-3966 23 1 42 14 0.86 4.38E-02 mmu-miR-7053-5p 53 24 108 34 1.02 4.41E-02 mmu-miR-744-3p 64 5 35 12 -0.87 4.41E-02 mmu-miR-145a-3p 263 25 308 9 0.23 4.62E-02 mmu-miR-7687-5p 28 18 76 37 1.45 4.76E-02 mmu-miR-7233-5p 25 10 59 27 1.26 4.85E-02 mmu-miR-6541 74 71 356 105 2.27 4.86E-02 mmu-miR-547-5p 36 38 4 2 -3.01 4.87E-02 mmu-miR-3058-5p 53 41 15 10 -1.86 4.99E-02  

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