text:
Melatonin: Pharmacology, Functions and Therapeutic Benefits

Abstract: Background: Melatonin synchronizes central but also peripheral oscillators (fetal adrenal gland, pancreas, liver, kidney, heart, lung, fat, gut, etc.), allowing temporal organization of biological functions through circadian rhythms (24-hour cycles) in relation to periodic environmental changes and therefore adaptation of the individual to his/her internal and external environment. Measures of melatonin are considered the best peripheral indices of human circadian timing based on an internal 24-hour clock. Methods: First, the pharmacology of melatonin (biosynthesis and circadian rhythms, pharmacokinetics and mechanisms of action) is described, allowing a better understanding of the short and long term effects of melatonin following its immediate or prolonged release. Then, research related to the physiological effects of melatonin is reviewed. Results: The physiological effects of melatonin are various and include detoxification of free radicals and antioxidant actions, bone formation and protection, reproduction, and cardiovascular, immune or body mass regulation. Also, protective and therapeutic effects of melatonin are reported, especially with regard to brain or gastrointestinal protection, psychiatric disorders, cardiovascular diseases and oncostatic effects. Conclusion: This review highlights the high number and diversity of major melatonin effects and opens important perspectives for measuring melatonin as a biomarker (biomarker of early identification of certain disorders and also biomarker of their follow-up) and using melatonin with clinical preventive and therapeutic applications in newborns, children and adults based on its physiological regulatory effects.

INTRODUCTION
Melatonin or 5 methoxy-N-acetyltryptamine ( Fig. 1) was discovered and isolated from bovine pineal in 1958 by Aaron Lerner [1]. Melatonin is the main hormone secreted by the pineal gland. Extrapineal sources of melatonin were reported in the retina, bone marrow cells, platelets, skin, lymphocytes, Harderian gland, cerebellum, and especially in the gastrointestinal tract of vertebrate species [2][3][4][5][6][7][8][9]. Indeed, melatonin is present but can also be synthesized in the enterochromaffin cells; the release of gastrointestinal melatonin into the circulation seems to follow the periodicity *Address correspondence to this author at the Hospital-University Department of Child and Adolescent Psychiatry, Guillaume Régnier Hospital, Rennes 1 University, Rennes 35000, France; Tel: +33-6-15-38-07-48; Fax: +33-2-99-64-18-07; E-mail: s.tordjman@yahoo.fr of food intake, particularly tryptophan intake [2,10]. It is noteworthy that the concentration of melatonin in the gastrointestinal tract surpasses blood levels by 10-100 times and there is at least 400 times more melatonin in the gastrointestinal tract than in the pineal gland [2]. Melatonin in the gastrointestinal tract of newborn and infant mammals is of maternal origin given that melatonin penetrates easily the placenta and is after secreted into the mother's milk [11][12][13]. It has even been suggested that melatonin is involved in the production of mekonium [2]. Melatonin in human breast milk follows a circadian rhythm in both preterm and term milk, with high levels during the night and undetectable levels during the day [14,15]. No correlation was found between gestational age and concentration of melatonin. It is noteworthy that bottle milk composition does not contain melatonin in powder formula. Also, human colostrum, during the first 4 or 5 days after birth, contains immune -competent cells (colostral mononuclear cells) which are able to synthesize melatonin in an autocrine manner [16]. Melatonin is mainly synthesized by the pinealocytes from the amino acid tryptophan, which is hydroxylated (by the tryptophan-5-hydroxylase) in 5-hydroxytryptophan, then decarboxylated (by the 5-hydroxytryptophan decarboxylase) in serotonin. Two enzymes, found mainly in the pineal gland, transform serotonin to melatonin [17,18]: serotonin is first acetylated to form N-acetylserotonin by arylalkylamine-N-acetyltransferase (AA-NAT, also called "Timezyme", is the rate-limiting enzyme for melatonin synthesis), and then N-acetylserotonin is methylated by acetylserotonin-Omethyltransferase (ASMT, also called hydroxyindole-Omethyltransferase or HIOMT) to form melatonin (Fig. 2). Both AA-NAT and ASMT activities are controlled by noradrenergic and neuropeptidergic projections to the pineal gland [19]. Norepinephrine, also called noradrenaline, activates adenylate cyclase which in turn promotes the melatonin biosynthesis enzymes, especially AA-NAT [20]. Once synthesized, melatonin is quickly released into the systemic circulation to reach central and peripheral target tissues.
Melatonin synthesis and secretion is enhanced by darkness and inhibited by light (Fig. 3) [21]. Luminous information is transmitted from the retina to pineal gland through the suprachiasmatic nucleus (SCN) of the hypothalamus. In humans, its secretion starts soon after sundown, reaches a peak in the middle of the night (between 2 and 4 in the morning) and decreases gradually during the second half of the night [22]. Nearly 80% of the melatonin is synthesized at night, with serum concentrations varying between 80 and 120 pg/ml. During daylight hours, serum concentrations are low (10-20 pg/ml) [23]. Serum concentrations of melatonin vary considerably with age, and infants secrete very low levels of melatonin before 3 months of age. Melatonin secretion increases and becomes circadian along with child development: Sadeh [24] reported an association between melatonin secretion and organization of sleep-wake rhythm from 6 months of age. However, more recent studies suggest that melatonin rhythm is set around 3 months of age in typical development, at the same time that infants begin to have more regular sleepwake cycles associated with nighttime sleep lasting 6-8 h [25]. In 3-years-old children, a stabilization of the sleepwake rhythm is observed, which corresponds to a regular melatonin secretion rhythm [26]. Nocturnal concentration peaks are the highest between the 4 th and 7 th years of age [23], and then decline progressively [27].

PHARMACOKINETICS
After intravenous administration, melatonin is rapidly distributed (distribution half-life of 0.5 to 5.6 minutes) and eliminated [28]. After oral administration, plasma concentration peak arises within 60 minutes [29]. Plasma concentrations diminution is biphasic, with a half-life of respectively 2 and 20 minutes [30]. Intake of an usual dose (i.e., 1 to 5 mg), allows within the hour after ingestion, melatonin concentrations 10 to 100 times higher than the physiological nocturnal peak to be obtained, with a return to basal concentrations in 4 to 8 hours.
A bioavailability study in four male healthy volunteers [31] showed a plasma melatonin peak varying between 2 and 395 nmol/L and an elimination half-life of 47± 3 min (mean ± SD) after oral administration of a 500 µg dose. Bioavailability varied from 10 to 56% (mean 33%).
After intravenous or oral administration, melatonin is quickly metabolized, mainly in the liver and secondarily in the kidney. However, after intravenous administration, the hepatic bio-degradation is less important due to the absence of hepatic first pass. It undergoes hydroxylation to 6hydroxymelatonin by the action of the cytochrome P450 enzyme CYP1A2, followed by conjugation with sulfuric acid (90%) or glucuronic acid (10%) and is excreted in the urine. About 5% of serum melatonin is excreted unmetabolized also in urine. The principal metabolite, the 6-sulfatoxymelatonin (6-SM), is inactive, and its urinary excretion reflects melatonin plasma concentrations [32]. Plasma levels can be also measured directly or indirectly assessed through salivary measures. A reverse relation between bioavailability of melatonin and the 6-SM concentrations area under the curve has been shown, the low bioavailability being explained by an important hepatic first pass [31].

MECHANISMS OF ACTION
Melatonin acts through different molecular pathways. The best characterized pathway is the activation of two types of membrane specific receptors: high affinity ML1 sites and low affinity ML2 sites [33,34]. The activation of ML1 receptors, which are G protein-coupled receptors [35], leads to an inhibition of the adenylate cyclase in target cells. The activation of ML2 receptors, currently called MT3, leads to phospho-inositides hydrolysis. MT3 is expressed in various brain areas and has been shown to be the enzyme quinone reductase 2 [36]. Two sub-types of the ML1 receptor have been described [37] Mel1a and Mel1b. Mel1a (or MT1) is encoded in human chromosome #4 (4q35.1) and consists of 351 amino acids. Mel1a is widely distributed in the pars tuberalis of the anterior pituitary and the SCN of the hypothalamus (which is the anatomic site of the circadian clock), and also in the cortex, thalamus, substantia nigra, nucleus accumbens, amygdala, hippocampus, cerebellum, cornea and retina [38]. Mel1b (or MT2) is encoded in human chromosome #11 (11q21-q22) and consists of 363 amino acids. Mel1b is distributed mainly in the retina and secondarily in the hippocampus, cortex, paraventricular nucleus, and cerebellum [39]. Melatonin has also an intracellular action by binding, on the one hand, to cytosolic calmodulin [40], and on the other hand, to two receptors of the Z retinoid nuclear receptors family [41]. Melatonin receptors have been found in several central but also in peripheral tissues, including heart and arteries, adrenal gland, kidney, lung, liver, gallbladder, small intestine, adipocytes, ovaries, uterus, breast, prostate, and skin [42]. Furthermore, they have also been detected in T and B lymphocytes. Evidence shows that there is a considerable variation in the density and location of the expression of melatonin receptors between species [43]. In a recent review, Liu et al. suggest, on the basis of affinity studies showing species-dependent fluctuations in vivo ligand selectivity, that there are considerable species differences in melatonin receptor pharmacology [44]. Models of mice with targeted disruption of either Mel1a or Mel1b receptor subtypes, or double mutants have been used, allowing the cellular mechanisms through which melatonin modulates circadian and photoperiodic rhythmicity to be understood [45,46].
The effects of melatonin depend on the localization and types of melatonin receptors; those effects are described in the next section.
For example, children with multiple developmental, neuropsychiatric and health difficulties often show melatonin deficiency [51]. When circadian rhythms are restored, behavior, mood, development, intellectual function, health, and even seizure control may improve [50,52]. It should be noted that according to several studies, circadian rhythms are important for typical (normal) neurodevelopment and their absence suppresses neurogenesis in animal models [53][54][55][56].
Finally, melatonin may as well be involved in early fetal development, with direct effects on placenta, glial and neuronal development, and could play an ontogenic role in the establishment of diurnal rhythms and synchronization of the fetal biological clock [57][58][59]. Iwasaki et al. [57] investigated the expression of the two enzymes involved in the conversion of serotonin to melatonin (AA-NAT and ASMT) and found that transcripts of these enzymes were present in the first-trimester human placenta. Moreover, they found also that melatonin significantly potentiated hcg secretion at optimal concentrations on cultured human trophoblast cells. These results suggest that melatonin regulates in a paracrine/autocrine way human placental function with a potential role in human reproduction. In vitro studies have shown that neural stem/progenitor cells express MT1 receptors, and melatonin induces glial cell-line derived neurotrophic factor (GDNF) expression in neural stem cells, suggesting an early role for melatonin in central nervous system development. Indeed, as indicated previously, melatonin of maternal origin crosses the placenta and can therefore influence fetal development. Studies in humans have repeatedly confirmed that the cycle of melatonin in maternal blood occurs also in fetal circulation [60,61]. The maturation and synchronization of the fetal circadian system have not been thoroughly studied. However, studies in nonhuman primate fetus have shown that maternal melatonin stimulates growth of the primate fetal adrenal gland and entrains fetal circadian rhythms, including SCN rhythms [62,63]. Furthermore, in mice, the suppression of melatonin rhythm by maternal exposure to constant light changes the rhythmic expression in fetal clock genes; these changes are reversed when melatonin is injected daily to the mother [64]. These results document that the fetal clock is imprinted by melatonin, which under normal circumstances is of maternal origin. In addition, some studies in humans and nonhuman primates show 24h rhythms in fetal heart rate and respiratory movements during the latter half of pregnancy. Whether the circadian system of the human fetus, particularly in late pregnancy, is under the influence of maternal SCN remains to be better ascertained [65].
Besides the well-known effects of melatonin on the regulation of sleep-wake rhythms, melatonin is considered as an endogenous synchronizer and a chronobiotic molecule, i.e. a substance that reinforces oscillations or adjusts the timing of the central biological clock located in the suprachiasmatic nuclei of the hypothalamus to stabilize bodily rhythms [66]. Furthermore, Pevet and Challet [67] view melatonin as both the master clock output and internal time-giver in the complex circadian clocks network: as a major hormonal output, melatonin distributes, through its daily rhythm of secretion, temporal cues to the numerous tissue targets with melatonin receptors, driving circadian rhythms in some tissue structures such as the adenohypophysis or synchronizing peripheral oscillators such as the fetal adrenal gland but also many other peripheral tissues (pancreas, liver, kidney, heart, lung, fat, gut, etc.). Circadian rhythms, and more precisely the circadian clocks network, allow temporal organization of biological functions in relation to periodic environmental changes and therefore reflect adaptation to the environment. Thus, the sleep-wake rhythm associated with biological circadian rhythms can be seen as an adaptation to the day-night cycle. Moreover, the synchronization by melatonin of peripheral oscillators reflects adaptation of the individual to his/her internal and external environment (for example, the synchronized effects of melatonin on cortisol and insulin secretion allow the individual to be fully awake at 8am and able to start the day by eating and getting some energy from food intake). Given the major synchronizing effects of melatonin on central and peripheral oscillators, measures of melatonin are considered the best peripheral indices of human circadian timing [68].
Futhermore, melatonin is involved in blood pressure and autonomic cardiovascular regulation, immune system regulation but also various physiological functions such as retinal functions, detoxification of free radicals and antioxidant actions through its action on MT3 receptors protecting the brain from oxidative stress [12,38,[68][69][70][71][72][73][74]. A through its action on MT3 receptors specific section is developed below on melatonin and brain protection. The antioxidant actions of melatonin protect also the gastrointestinal tract from ulcerations by reducing secretion of hydrochloric acid and the oxidative effects of bile acids on the intestinal epithelium, and by increasing duodenal mucosal secretion of bicarbonate through its action on MT2 receptors (this alkaline secretion is an important mechanism for duodenal protection against gastric acid); melatonin prevents also ulcerations of gastrointestinal tract by increasing microcirculation and fostering epithelial regeneration [2,66]. Concerning the role of melatonin in immune regulation, melatonin has direct immuno-enhancement effects in animals and humans [75,76]. Indeed, melatonin stimulates the production of cytokines and more specifically interleukins (IL-2, IL-6, IL-12) [77]. In addition, melatonin enhances T helper immune responses [76,78]. Furthermore, the melatonin antioxidant actions contribute to its immunoenhancing effects [77] and have also an indirect effect by reducing nitric oxide formation which facilitates the decrease of the inflammatory response [79]. As suggested by Esquifino et al. [80], melatonin might provide a time-related signal to the immune network.
In addition, effects of melatonin on body mass and bone mass regulation have been reported. Melatonin is known for its role in energy expenditure and body mass regulation in mammals by preventing the increase in body fat with age [81][82][83]. These effects are mediated by MT2 receptors in adipose tissue [84]. Moreover, melatonin increases bone mass by promoting osteoblast cell differentiation and bone formation [85,86]. In humans, melatonin stimulates bone cell proliferation and Type I collagen synthesis in these cells, and inhibits bone resorption through down-regulation of the RANKL-mediated osteoclast formation and activation [87,88]. Also, a deficit of melatonin has been found to be associated with animal scoliosis following pinealectomy and human idiopathic scoliosis [89,90].
Finally, melatonin has physiological effects on reproduction and sexual maturation in mammals through down-regulation of gonadotropin-releasing hormone (GnRH) gene expression in a cyclical pattern over a 24-hour period [91][92][93]. The rhythmic release of GnRH controls luteinizing hormone (LH) and follicule-stimulating hormone (FSH) secretion. The daily profile of melatonin secretion conveys internal information used for both circadian and seasonal temporal organization [67]. The melatonin rhythmic pattern entrains the reproductive rhythm via the influence of photoperiod on LH pulsatile secretion and therefore mediates the seasonal fluctuations of reproduction clearly observed in animals (seasonal breading as species-specific seasons for reproduction) and moderately observed in humans [94][95][96].

Melatonin in Disorders and Therapeutic Effects
Therapeutic effects of melatonin have been reported in several disorders such as certain tumors, cardiovascular diseases or psychiatric disorders. The part concerning melatonin and psychiatric disorders is in particular developed given our past and current work on this topic.
Concerning psychiatric disorders, secretion disturbances of the pineal gland have been described in child and adult psychiatry, with notably in most studies a decreased nocturnal melatonin secretion observed in major depressive disorder, bipolar disorder, schizophrenia or autism spectrum disorder [114][115][116][117].
Also, a phase-shift of melatonin has been reported in major depressive disorder and bipolar disorder, including in particular a delayed melatonin peak secretion [114,118,119]. It is noteworthy that increased melatonin levels (measures of blood melatonin and urinary excretion of 6-SM) were found when clinical therapeutic benefits were observed following the use of antidepressants [120][121][122]. Furthermore, significant improvement of major depressive disorder and anxiety was described following administration of 25mg per day of agomelatine, a MT1/MT2 melatonin agonist and selective antagonist of 5-HT 2C receptors [123][124][125].
Concerning autism spectrum disorder (ASD), abnormalities in the serotoninergic system and sleepwake rhythm disturbances observed in children with ASD suggest altered melatonin secretion in autism [126][127][128]. Sleep disorders (mostly increased sleep latency, reduced total sleep and nocturnal awakenings with insomnia) are observed in 50-80% of individuals with autism [129]. It is noteworthy that sleep problems are not specific of autism and are also observed in children with intellectual disability associated or not with autism [128]. However, melatonin measures in children with intellectual disability not associated with autism, such as some children with Down syndrome and Fragile X syndrome, showed respectively normal melatonin production despite delayed nocturnal melatonin peak secretion and increased levels of melatonin [130,131], whereas decreased nocturnal melatonin secretion was mostly observed in children with autism [128]. We reported in two different large samples of children with autism significant relationships between decreased nocturnal urinary excretion and severity of autistic impairments in social communication [127,132]. These results suggest that abnormalities in melatonin physiology might contribute not only to sleep problems in autism, but also to biological and psychopathological mechanisms involved in the development of ASD (for example, certain immunological abnormalities found in autism, such as a decrease number of T lymphocytes, might be explained by the hypo-functioning of the melatonin system).
Concerning schizophrenia, as suggested by Morera-Fumero and Abreu-Gonzalez [116], a possible explanation for the "low melatonin syndrome" present in some individuals with schizophrenia may stem from the study of the melatonin-synthesizing enzymes, the AA-NAT and ASMT. Furthermore, according to some authors, MT3 might be involved in the melatonin disturbances observed in schiozophrenia [38]. Finally, melatonin secretion was also studied in obsessive compulsive disorder but no abnormalities in melatonin levels were reported.

Melatonin and Brain Protection
Neurological and neuropsychological disabilities caused by brain injuries are a major public health concern. Thus, reducing deficits after a stroke is a major issue. In this line, a number of recent studies have reported the important role of melatonin in neuroprotection in animal models of stroke [133]. Indeed, melatonin administration after an experimental stroke in animals reduces infarction volume [134,135]. Such a protective effect can be seen in both gray and white matter [136] and melatonin reduces also inflammatory response [137], cerebral oedema formation [138], and blood-brain barrier permeability [139]. In addition, Kilic et al. [140] investigated how sub-acute delivery of melatonin, starting at 24 hours after stroke onset, and continuing for 29 days can influences neuronal survival, endogenous neurogenesis, motor recovery and locomotor activity in mice submitted to an occlusion of the middle cerebral artery during 30 minutes. Furthermore, melatonin improved neuronal survival and enhanced neurogenesis, even when applied one day after stroke. In addition, the authors showed both motor as well as behavioral improvements after melatonin administration. Indeed, the results indicate that cell survival was associated with a long-lasting improvement of motor and coordination deficits as well as with attenuation of hyperactivity and anxiety of the animals as revealed in open field tests. Its neuroprotective activity in animal models of ischemic stroke, as well as its lack of serious toxicity suggests that melatonin could be used for human stroke treatment in the future.
In addition to its protective effect after stroke, experimental data obtained in various independent animal models of brain lesions in neonates support the notion of a neuroprotective effect of melatonin in preterm neonates (see for a review, Biran et al., [141]). In infants, a major source of brain injury is preterm birth, often associated to long-term neurological, cognitive, educational, and social problems. Neurodevelopmental disorders are not only seen in extremely preterm birth [142] but also in late prematurity [143]. A large number of infants who survive very preterm birth develop cerebral palsy [144] with a high occurrence of associated motor, perceptual and cognitive deficiencies in childhood [145,146]. Nowadays, the most common brain damage observed in preterm children is diffuse white matter damage as well as reduced neural connectivity [147,148] in the context of infection, inflammation, and hypoxia-ischemia [149]. Although a number of treatments have been tested in preclinical animal models of perinatal brain injury, none of them had been proved to be efficient as a neuroprotector nor translated in clinical practice. Among the molecules proposed, melatonin is a very good candidate, given its effect on brain development, neuroprotection as well as regarding its absence of adverse effects [150]. As discussed by Biran et al., [141], in addition to its good safety profile, melatonin easily crosses the placenta as the blood-brain barriers and blocks oxidative, excitotoxic and inflammatory pathways, all involved in the pathogenesis of perinatal brain damage caused by preterm birth. However, only a few studies have looked at the synthesis of melatonin in preterm and term neonates. These studies point to a reduced urinary concentration of melatonin during the first 3 months after birth in preterm infants (see Biran et al. [141] for review and discussion). As these authors discussed, compared with term neonates, preterm neonates show a delayed secretion of melatonin which persists after correction for gestational age up to 8 to 9 months of age. In the absence of maternal melatonin, the appearance of circadian rhythms depends principally on neurological maturation, and very little on the environment [151].
Since, as above-mentioned, melatonin easily crosses the blood-brain and placental barriers, it can be administered antenatally in order to reduce or prevent the impact of brain lesions in preterm neonates. Currently, two therapeutic trials testing the neuroprotective properties of melatonin administration in the immediate prepartum period in very preterm infants are under way in France and in the United Kingdom (see Biran et al., [141] for description). The French trial aims to determine the dose of melatonin to be administered in prepartum by parenteral route to mothers at risk of preterm delivery, to decrease the extent of white matter damage detected by diffusion tensor imaging in infants born preterm. The objective of the English trial is to prove that melatonin is capable of reducing brain injury and white matter disease as defined by magnetic resonance imaging at term. These trials will probably lead to a clinical use of melatonin before preterm birth (in case of at risk mother) of just after birth in preterm neonates in order to prevent neurodevelopmental deficits in these children.
Interestingly, from a functional point of view, abnormalities in melatonin physiology associated with sleep disorders, and in particular sleep deprivation, are seen to endanger cerebral and more specifically hippocampal integrity, leading to cognitive dysfunction and contributing to the development of mood disorders [152]. The involvement of melatonin in the development of mood disorders was discussed in the previous section.
Finally, based on the brain protective role of melatonin against oxidative stress previously described in this article, there is also increased experimental evidence showing the therapeutic potential of melatonin in neurodegenerative conditions such as Alzheimer disease, Parkinson disease, Huntington's disease and amyotrophic lateral sclerosis (see Polimeni et al. [153] for a review). Additional studies and clinical trials are now required both in preterm neonates and aging adults to test the clinical efficacy of melatonin supplementation in such disorders, and to identify the specific therapeutic concentrations needed regarding the subject's age, disease and brain lesion as well as the short and long-term effects of melatonin both on physiological, functional and cognitive outcomes.

CONCLUSION AND FUTURE DIRECTIONS
Regular physiological rhythms are important to ensure a stable internal environment by creating continuity from repetition of identical discontinuities. Several studies, based on animal models and human perinatal development, suggest that stable patterns of repeated stimuli in the form of maternal physiological rhythms, involving cross-modal perception such as regular cardiac rhythm, which provides the fetus with auditory and vibratory stimuli, allow the fetus to integrate sensory information facilitating prenatal perceptual learning and to develop a coherent representation of his or her internal and external environment [154][155][156]. Fluctuations in physiological rhythms (variants), such as variations in the maternal cardiac rhythm and also variations in hormone levels involved in the circadian rhythms that are already present during the fetus life (the fetus' circadian rhythms are the mother's ones), occurring in a background of regular repetition of identical sequences (invariants), may help the fetus to develop the ability to adapt to change in an environment characterized by high regularity. As previously emphasized by Tordjman [156], very early mother-infant relations provide a secure environment based on the repetition of invariants, while at the same time promoting adaptation to change through the presence of variants. It is through the regular repetition of identical sequences of discontinuity, such as circadian rhythms, that a continuum is constructed associated with the development of adaptation to changes. Melatonin plays a major role in the regularity and synchronization of central and peripheral oscillators allowing the development of harmonious internal functioning and adaptation of the internal environment to external environment. As highlighted by Arendt [50], melatonin might be the best peripheral biomarker of human circadian timing. This review shows the high number and diversity of melatonin effects opening important perspectives for measuring melatonin as a biomarker (biomarker of early identification of certain disorders and also biomarker of their follow-up) and using melatonin with therapeutic applications. Given the very large potential clinical applications of melatonin and its apparent absence of major side effects, it is amazing to see that the role of melatonin in human clinical trials has been mainly confined to sleep problems and focused on just a few disorders [157]. Future studies are required, such as studies of melatonin dose-response relationships, to extend the use of melatonin in clinical practice, prevention and treatment.


extfieldsofstudy: Biology Medicine

text:
Mecp2-mediated Epigenetic Silencing of miR-137 Contributes to Colorectal Adenoma-Carcinoma Sequence and Tumor Progression via Relieving the Suppression of c-Met

The molecular mechanisms underlying colorectal cancer (CRC) development remain elusive. In this study, we examined the miRNA and mRNA expressions in the adenoma-carcinoma sequence (ACS), a critical neoplastic progression in CRC development. We found that miR-137 was down-regulated in all adenoma and carcinoma tissues. Low miR-137 levels were correlated negatively with tumor progression and metastasis. Then we identified the inhibition effect of the miR-137 in CRC development, both in CRC cell lines and mouse models. MiR-137 was shown to control CRC cell proliferation, colony formation, migration and invasion and to control tumor growth and metastasis. We further confirmed the negative association between miR-137 and c-Met expression and thus validated this important oncogene as the target of miR-137 in CRC. In addition, we found a DNA methyl-CpG-binding protein, Mecp2, was up-regulated in ACS tissues via mRNA sequencing. Further experiment showed that miR-137 expression in CRC was subjected to epigenetic regulation mediated by Mecp2. We also confirmed c-Met expression can be up-regulated by silencing of miR-137 and suppressed by coexpression of Mecp2 and miR-137. These findings highlight the critical role of miR-137-c-Met nexus in CRC development and reveal Mecp2-regulated epigenetic silence causes the downregulation of miR-137 in colorectal adenoma and carcinoma.

body of research showed that miRNA alternations played a key role in the development of various types of cancer. However, little is known about the functional role of miRNA in consecutive colorectal ACS and CRC progressions. In this study, we examined the expression of miR-137 in ACS and explored its role in the regulation of CRC cell function. In addition, miRNA-137-mediated c-Met expression in cells and the underlying mechanism of miRNA-137 alternation in colorectal ACS were also investigated.

Results
MiR-137 is associated with ACS and CRC progression. A small RNA sequencing analysis of 18 colorectal ACS tissues from 6 patients was conducted to study the effect of miRNA profile in regulating human colorectal ACS and CRC progression. We identified 15 miRNAs that had at least 2-folds higher expression levels compared with other groups (Fig. 1a). MiR-137 was found to be consistently down-regulated in 6 pairs of adenoma and carcinoma tissues. QRT-PCR analysis of miR-137 expression in 30 colorectal adenoma tissues and in 70 CRCs showed miRNA-137 was not only differentially expressed in colorectal adenoma ( Fig. 1b; P = 0.041), but also significantly reduced in tumor tissues ( Fig. 1c; P < 0.001). When the clinicopathological implication of miR-137 was analyzed in CRC patients it is found that low miR-137 levels were negatively correlated to tumor TNM stage ( Fig. 1d; P = 0.019) and metastasis ( Fig. 1e; P = 0.017).
Over expression of miR-137 inhibits CRC cell proliferation, colony formation, migration, and invasion in vitro. The biological role of miR-137 was examined in vitro by functional assays. Expression of miR-137 in 6 CRC cell lines and colon mucosa cell line NCM640 was shown in Fig. 2a. A significant decrease in cell proliferation was observed in both miR-137 lentivirus (LV.miR-137)-infected HCT116 and LoVo cells comparing with the negative control (LV.NC) (Fig. 2b). In both cell lines, colony formation ability was inhibited by the overexpression of miR-137 (Fig. 2c). MiR-137 mimics were transfected into HCT116 and LoVo cell to transiently increase the miR-137 expression. The results from cell migration and invasion assays showed that the overexpression of miR-137 significantly inhibited HCT116 and LoVo cell migration and invasion via cell migration and invasion assays (Fig. 2d,e). Taken together, we have shown the tumor suppressor role of miR-137 in CRC development.

Overexpression of miR-137 inhibits colorectal tumor growth and hepatic metastasis in vivo.
To further investigate the in vivo effect of miR-137 on tumor formation and metastasis, HCT116 cells, which were stably transfected with either LV.miR-137 or LV.NC, were subcutaneously and intraspleenically injected into nude mice.
At 4 weeks post-injection, the average tumor size was found to be significant smaller in rats injected with cells that overexpressed miR-137 compared with the control group (Fig. 3a,b). Similarly, less hepatic metastasis nodes were found in rats with cells that overexpressed miR-137 (Fig. 3c,d). In coordination with in vivo study, overexpression of miR-137 was found to inhibit colorectal tumor progression and hepatic metastasis as well.

MiR-137 alterations affect ACS and tumor progression by down regulating c-Met expression.
Both Target Scan and Pictar systems were used to identify putative gene targets of miR-137. C-Met was selected for further analysis because of its involvement in various malignances and was predicted to be a target of miR-137 in melanoma 7 . In vitro and in vivo assays were conducted to determine if miR-137 could regulate c-Met expression in CRC. HCT116 and LoVo cells were transfected with miR-137 or control mimics respectively. Forced expression of miR-137 was associated with reduce c-Met mRNA levels in both cell lines (Fig. 4a,b). Anti-miR-137, and anti-miRNA control were then transfected into the above 2 cell lines. Lower expression of miR-137 increased c-Met mRNA levels in both cell lines (Fig. 4a,b). Cell lysate were analyzed with Western blot to determine the protein levels of c-Met in the aforementioned treated cells and xenograft tumors. We found that the expression of both c-Met was negatively correlated with miR-137 (Fig. 4c,d).
Mecp2-regulated epigenetic silencing of miR-137 contributes to colorectal ACS and tumor progression by relieving the suppression of c-Met. MiR-137 is constitutively expressed in colonic epithelium and epigenetic silencing of miR-137 is an early event in colorectal carcinogenesis 8 . To study the epigenetic profile and its role in regulating human ACS, a transcriptome analysis in 18 colorectal ACS tissues collected from the same 6 patients was performed. Thirty five epigenetic genes with a fold-change of > 2 in the between group expression levels were collected. Mecp2 was found to be consistently up-regulated in ACS tissues, compared to the normal tissues (Fig. 5a). Mecp2 is a DNA methyl-CpG-binding protein and was reported to epigenetically regulate miR-137 in adult neural stem cells 9 . In the current study, a ChIP assay was performed and we confirmed that Mecp2 could directly bind to the promoter of miR-137 (Fig. S1). In addition, we overexpressed Mecp2 in HCT116 and LoVo cells and found that miR-137 expression was significantly decreased (Fig. 5b). Furthermore, the expression of c-Met, the target of miR-137, was significantly increased because of inhibition of miR-137 (Fig. 5c). The results were consistent with the aforementioned Western blot assay data (Fig. 5d). To further confirm the deduction, Mecp2 plasmid and miR-137 mimics were transfected into HCT116 and LoVo cells. The expressions of c-Met were suppressed again when the miR-137 expression was recovered (Fig. 6a), (Fig. 6b,c). These results suggest a regulatory cascade from MeCP2, miR-137, to c-Met exists (Fig. 6d).

Discussion
The tumor miRNA profile may resemble that of their antecedent stem cells and thus reflect development lineage 10 . A study conducted by Lu et al. 6 examined the expression of miRNAs across normal tissues, primary tumors as well as cell lines and indicted that tumors display a miRNA expression profile reminiscent of that in the tissues from which they were derived. Therefore, the miRNA may play a role in determination and/or maintenance of lineage during tumor development. In this sense, we collected 18 ACS samples from 6 patients and employed a small RNA sequencing to examine the differential expressions of miRNAs in them. To control the tumor heterogeneity, 3 consecutive ACS samples were collected from one patient. We found miR-137, a key molecular in cross talk between microRNAs and epigenetic regulation in the carcinogenesis 9 , was significantly down-regulated in colorectal adenomas and carcinomas, comparing to normal tissues. It has been reported that miR-137 functioned as a tumor suppressor in cancer types such as head-neck cancer and gastric cancer 11,12 . However, its effect in colorectal ACS remains obscure. In this study, we focus our investigation on the role of miRNAs in colorectal ACS via small RNA sequencing analysis and found low expression of miR-137 may be involved in. The miR-137 whose expression was down-regulated in colorectal adenomas and carcinomas was further validated. In addition, the miR-137 level was significantly correlated with CRC progression. To identify the role of the miR-137 in CRC development, functional assays were performed and aberration of miR-137 expression was shown to alter cell proliferation, colony formation, migration, and invasion in CRC cell lines as well as tumor growth and liver metastasis in mouse models.
MiRNAs functions as post-transcriptional regulation of gene expression by complementary base pairing with the 3′UTR of target mRNAs, causing their degradation or suppressing mRNA translation 13 . Previous studies found that miR-137 is negatively correlated with mRNAs 7,8 . We selected c-Met as a candidate target of miR-137 for its role in the progression of a wide variety of human malignancies 14 . The findings strongly suggested miR-137 could directly target c-Met and regulate c-Met expression in vitro and in vivo. Furthermore, the results obtained from qRT-PCR and Western blot analysis confirmed both mRNA and protein of c-met are negatively associated with miR-137 expression. Takeuchi et al. 15 found the mRNA expression of c-Met in CRC specimens was significantly higher than that in colorectal normal mucosa tissues. Additionally, colorectal adenomas showed different  15 . The c-Met pathway was implicated to be related to colorectal normal mucosa, colorectal adenoma, and colorectal carcinoma progression 15 .
Aberrant epigenetic regulation of miR-137 promoter, such as DNA hypermethylation, may represent a key mechanism for miR-137 down-regulation in several human cancers 16 . Epigenetic silencing through promoter methylation of miR-137 is an early event in colorectal adenomas and the following carcinogenesis 8 . Results from silico analysis have found several epigenetic genes altered in ACS. In this study, we focused on Mecp2 which was reported to epigenetically regulate specific miRNAs in adult neural stem cells 9 . Our results suggested miR-137 expression in CRC was subjected to epigenetic regulation mediated by Mecp2. We also found c-Met expression could be up-regulated by silencing of miR-137 and suppressed by coexpression of Mecp2 and miR-137. Identification of these gene alterations has provided new sights into the molecular process of developmental course from colorectal adenoma to carcinoma. In addition, in the study by Plummer et al. 17 , they indicated that Mecp2 could directly bind to the c-Met promoter and activate its transcription. In the current study, our results suggested Mecp2 could upregulate c-Met expression by silencing miR-137 expression (Fig. 6d). Thus, the relationship between Mecp2 and c-Met and the regulation mechanism needed to be further investigated.
Taken together, our results identified a new regulatory network of the miR-137/c-Met nexus, which is not reported in ACS but functions as an oncogenic miRNA in CRC development. More than that, an important funding regarding the significant roles that crosstalk between miRNAs and epigenetics might played in ACS and tumor progression was illustrated. Further understanding in pathogenic significance for epigenetic regulation of miRNAs and its functional targets will inspire novel thinking in diagnosis and treatment of CRC.

Materials and Methods
Human tissue samples. Human tissue samples were collected from 6 patients with both colorectal adenoma and carcinoma for small RNA and mRNA sequencing, 30 patients with colorectal adenomas and 70 patients with CRCs. All tissue samples were snap frozen in liquid nitrogen at the time of collection and then stored at -80 °C for later use. The diagnosis of colorectal adenoma and carcinoma was confirmed through pathological examination. All clinicopahtological data were collected (Tables S1 and S2). This study was conducted under the principles of the World Medical Association Helsinki agreement. Ethical approval was obtained from the Ethics Committee of Zhongshan Hospital, Fudan University. Written informed consents were obtained for experimentation with human subjects prior to recruiting participants and conducting procedures.
Cell culture and Proliferation assay. HCT116 and LoVo human CRC cells were obtained from Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences. HCT116 cells were cultured in McCoy's 5 A medium supplemented with 10% fetal bovine serum (FBS). LoVo cells were cultured in RPMI1640 medium supplemented with 10% FBS. All cells were cultured in a humidified incubator at 37 °C with 5% CO 2 and plated on 96-well plates at 1 × 10 3 cells/well. The proliferation of cancer cells 5 days after treatment were measured by the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) assay.
Colony Formation Assay. Colony Formation Assay was carried out according to standard procedures. Cells were mixed with culture medium containing 0.6% agar to achieve a final agar concentration of 0.4%. 1 mL of the cell suspension was immediately plated onto 6-well plates that were coated with 0.6% agar in tissue culture media at 1 mL/well. Cell colonies were counted in triplicates at day 15.
Cell migration and invasion assays. 24-well transwell plate (8-mm pore size, Corning) was used to measure the migratory and invasive ability of each cell line. For migration assay, 5 × 10 4 cells were plated onto the top chamber lined with a non-coated membrane. For invasion assay, chamber inserts were coated with 200 mg/ml of Matrigel, which were allowed to dry overnight under sterile conditions. 1 × 10 5 cells were plated onto the top chamber. Both assays were carried out according to standard procedures.
Overexpression and knockdown of miR-137. Cells were seeded onto 6-well plates at a density of 2 × 10 5 per well and were given at least 16 hours to allow cell attachment to the well surface before consucting experiments. siRNA or negative control was transfected with lipofectamine 2000 (Invitrogen). Stable cell lines were established by lentiviral infection (LV). LV-miR-137, si-miR-137, and miR-137 mimics were purchased from GeneChem, Shanghai China.
Chromatin immunoprecipitation. HCT116 cells were grown on 3-6 confluent 6 cm cell culture plates and were fixed by 1% formaldehyde to culture medium for 10 min at room temperature. Cell lysates were pelleted by centrifugation at 3,000 rpm for 5 min, resuspended again in 1 ml of cold cell lysis buffer for 5 min on ice, and then repelleted to collect nuclei. ChIP was performed as described previously 9 . The Pierce Agarose ChIP Kit (Thermo) was used. For qPCR followed by ChIP, the Premix Taq ™ Kit (Takara) was used. Primers spanning 200-250 bp of miR-137 promoter were used for the ChIP qPCR (Supplementary Table S3). Primer sequences spaced at 250 bp intervals spanning 1.0 kb from upstream of miR-137 open reading frame (ORF) were designed using Primer 5.0 software (Premier). Immunodeficient mouse model. All animal experiments were approved by the animal care and use committee of Fudan University and performed in accordance with the guidelines of the National Institutes. HCT116 cells infected with LV-miR-137 or LV-NC were harvested and injected into female nude (nu/nu) mice (5 × 10 6 viable tumor cells/mouse for the subcutaneous xenograft tumor model and 1 × 10 6 viable tumor cells/mouse for the intraspleenic liver metastasis model). The animals were equally divided into a treatment group and a control group (8 mice per group for the subcutaneous xenograft tumor model and 5 mice per group for the intraspleenic liver metastasis model). Based on length and width measurements, xenograft tumor volume (V) was calculated by V = (length × width2)/2 every 4 days. During this period, in the study of subcutaneous xenograft tumor model, there were 2 mice dead in the control group with and 1 mouse dead in the treatment group. Subcutaneous tumor volume and hepatic metastasis nodes were calculated after the mice were sacrificed 28 days post-injection.


text:
Reverse Gyrase Functions in Genome Integrity Maintenance by Protecting DNA Breaks In Vivo

Reverse gyrase introduces positive supercoils to circular DNA and is implicated in genome stability maintenance in thermophiles. The extremely thermophilic crenarchaeon Sulfolobus encodes two reverse gyrase proteins, TopR1 (topoisomerase reverse gyrase 1) and TopR2, whose functions in thermophilic life remain to be demonstrated. Here, we investigated the roles of TopR1 in genome stability maintenance in S. islandicus in response to the treatment of methyl methanesulfonate (MMS), a DNA alkylation agent. Lethal MMS treatment induced two successive events: massive chromosomal DNA backbone breakage and subsequent DNA degradation. The former occurred immediately after drug treatment, leading to chromosomal DNA degradation that concurred with TopR1 degradation, followed by chromatin protein degradation and DNA-less cell formation. To gain a further insight into TopR1 function, the expression of the enzyme was reduced in S. islandicus cells using a CRISPR-mediated mRNA interference approach (CRISPRi) in which topR1 mRNAs were targeted for degradation by endogenous III-B CRISPR-Cas systems. We found that the TopR1 level was reduced in the S. islandicus CRISPRi cells and that the cells underwent accelerated genomic DNA degradation during MMS treatment, accompanied by a higher rate of cell death. Taken together, these results indicate that TopR1 probably facilitates genome integrity maintenance by protecting DNA breaks from thermo-degradation in vivo.

Introduction
DNA topoisomerases impose changes to the topological states of circular DNA and play important roles in DNA replication, recombination, and repair as well as in RNA transcription [1,2]. Thermophiles encode a unique type of topoisomerase named reverse gyrase (TopR) that introduces positive supercoils to circular DNAs [3]. Because reverse gyrase is the only protein that is exclusively conserved in thermophilic organisms, the enzyme has been implicated in genome stability maintenance in thermophiles [4]. Indeed, genetic studies in Thermococcus kodakaraensis, a hyperthermophilic euryarchaeon, show that growth of cells lacking TopR enzyme is significantly retarded at high temperatures (from 65 to 90 • C) [5]. Further, crenarchaea encode two TopR proteins, but attempts to isolate TopR deletion mutants from S. islandicus Rey15A was not successful [6], suggesting an essential function for these enzymes in this crenarchaeon. Therefore, TopR enzymes must have important roles in thermophilic life.
Likewise, base alkylation agents, such as methyl methanesulfonate (MMS), generate DNA lesions that can also be converted to AP sites and SSBs [10]. If not repaired in a timely fashion, AP sites and SSBs induce collapse of DNA replication, giving rise to double strand DNA breaks (DSBs). DSBs are a more severe type of DNA lesion that must be repaired by homologous recombination repair (HRR), an energy-consuming pathway [11,12]. As a result, severe DSB damage on chromosomal DNA often leads to cell death. Therefore, one of the great challenges in thermophilic life is to deal with numerous DNA lesions that result from spontaneous DNA decomposition in order to prevent the formation of DSBs.
Since DNA lesions produced by MMS treatment must be repaired in a similar fashion as DNA lesions generated from spontaneous base decomposition in DNA, MMS treatment provides a useful means for investigating DNA damage repair mechanisms in thermophilic organisms. Indeed, investigation on the effect of MMS on genome integrity maintenance in Sulfolobus solfataricus has revealed that TopR1 is subjected to degradation upon a lethal MMS treatment and that MMS-induced TopR1 degradation coincides with genomic DNA degradation [13]. Moreover, it has been shown that the Archaeoglobus fulgidus TopR binds to DNA nicks in vitro and inhibits heat-induced degradation of nicked DNA, and the DNA protection mechanism is independent of supercoiling [14]. This raises an interesting question as to whether TopR proteins could protect damaged DNA from thermo-degradation in vivo.
In the article, we employed S. islandicus, a genetic model for which very versatile genetic tools have been developed [15], to investigate the function of TopR1 in genome integrity maintenance. We found that MMS treatment induced a series of events including: (a) massive breaks on the sugar-phosphate backbone of genomic DNA, (b) gradual DNA content reduction and TopR1 degradation, and (c) DNA-less cell formation and chromatin protein degradation. Moreover, we constructed a TopR1 knockdown strain (pKD-topR1/E233S1) with a CRISPR-based RNA interference tool (CRISPRi) [16]. The strain exhibits a higher sensitivity to MMS and an early onset of DNA degradation during MMS treatment, indicating that TopR1 protects DNA breaks from thermo-degradation in vivo.

Determination of Moderate and Lethal Dosages of Methyl Methanesulfonate (MMS) for S. islandicus
To investigate the effects of MMS on S. islandicus, exponentially growing S. islandicus cultures were treated with 0.6, 1.3, or 2.6 mM of MMS, and the treated samples were analyzed for culture growth and cell viability. While 0.6 mM MMS did not exert any apparent effect on culture growth (data not shown), 1.3 mM and 2.6 mM MMS yielded growth retardation to the S. islandicus. While a strong growth inhibition was observed for the 2.6 mM MMS-treated culture, since the absorbance at 600 nm (A 600 ) of the culture stopped increasing immediately after drug addition, only a moderate growth inhibition was attained by the 1.3 mM MMS-treated culture ( Figure 1). Furthermore, in the 2.6 mM MMS-treated culture, colony formation units (CFU) declined by more than 500-fold in the first 30 min of drug treatment, and cell viability did not increase in the following 24 h of incubation. In contrast, about a 50% drop in cell viability was observed for the 1.3 mM MMS-treated culture in the first 30 min, and the cell viability started to increase 2 h after drug treatment ( Figure 1). Therefore, the two drug concentrations (1.3 and 2.6 mM) represent a moderate and a lethal dosage of MMS for exponentially growing S. islandicus cells, respectively. respectively. The cultures without any treatment were also grown at the same time as control (CK). The optical density (left panel) and cell viability (right panel) were analyzed during the treatment at indicated time points. The data are representative of three independent replicates. (CK, black squares; 1.3 mM, white squares; 2.6 mM, black triangles). OD, the optical density measured at a wavelength of 600 nm; CFU, colony formation unit.

MMS Treatment Yields Immediate Genomic DNA Breakage and Subsequent DNA Degradation in S. islandicus
To reveal whether and how MMS induces genome instability in Sulfolobus, total DNAs were prepared from cell samples collected during the MMS treatments and analyzed by pulsed field gel electrophoresis (PFGE). As shown in Figure 2, whereas the circular chromosomal DNA of untreated cells was retained in the loading wells over the 24 h incubation, 1.3 and 2.6 mM of MMS induced the accumulation of considerable amounts of genomic DNA fragmentation, which peaked at 15 min after drug treatment, indicating that MMS immediately induced extensive DNA backbone breaks on the chromosome ( Figure 2). Noticeably, the higher dosage of MMS yielded more and shorter DNA fragments than the moderate dose, indicating that the number of DNA backbone breaks is correlated to MMS dosage, and, furthermore, a DNA band close to the loading well which could correspond to the linear chromosomal DNA is present in 1.3 mM MMS-treated samples but absent from 2.6 mM MMS-treated samples. Subsequently, the intensity of DNA fragments gradually declined from 2 h in both 1.3 and 2.6 mM MMS treatment. Noticeably, from 6 to 24 h during 1.3 mM MMS treatment, most genomic DNA was retained in the loading wells, similar to that of the untreated samples, suggesting reestablishment of genome integrity after the repair of DNA breaks during the moderate treatment.  respectively. The cultures without any treatment were also grown at the same time as control (CK). The optical density (left panel) and cell viability (right panel) were analyzed during the treatment at indicated time points. The data are representative of three independent replicates. (CK, black squares; 1.3 mM, white squares; 2.6 mM, black triangles). OD, the optical density measured at a wavelength of 600 nm; CFU, colony formation unit.

MMS Treatment Yields Immediate Genomic DNA Breakage and Subsequent DNA Degradation in S. islandicus
To reveal whether and how MMS induces genome instability in Sulfolobus, total DNAs were prepared from cell samples collected during the MMS treatments and analyzed by pulsed field gel electrophoresis (PFGE). As shown in Figure 2, whereas the circular chromosomal DNA of untreated cells was retained in the loading wells over the 24 h incubation, 1.3 and 2.6 mM of MMS induced the accumulation of considerable amounts of genomic DNA fragmentation, which peaked at 15 min after drug treatment, indicating that MMS immediately induced extensive DNA backbone breaks on the chromosome ( Figure 2). Noticeably, the higher dosage of MMS yielded more and shorter DNA fragments than the moderate dose, indicating that the number of DNA backbone breaks is correlated to MMS dosage, and, furthermore, a DNA band close to the loading well which could correspond to the linear chromosomal DNA is present in 1.3 mM MMS-treated samples but absent from 2.6 mM MMS-treated samples. Subsequently, the intensity of DNA fragments gradually declined from 2 h in both 1.3 and 2.6 mM MMS treatment. Noticeably, from 6 to 24 h during 1.3 mM MMS treatment, most genomic DNA was retained in the loading wells, similar to that of the untreated samples, suggesting reestablishment of genome integrity after the repair of DNA breaks during the moderate treatment.

MMS Treatment Yields Immediate Genomic DNA Breakage and Subsequent DNA Degradation in S. islandicus
To reveal whether and how MMS induces genome instability in Sulfolobus, total DNAs were prepared from cell samples collected during the MMS treatments and analyzed by pulsed field gel electrophoresis (PFGE). As shown in Figure 2, whereas the circular chromosomal DNA of untreated cells was retained in the loading wells over the 24 h incubation, 1.3 and 2.6 mM of MMS induced the accumulation of considerable amounts of genomic DNA fragmentation, which peaked at 15 min after drug treatment, indicating that MMS immediately induced extensive DNA backbone breaks on the chromosome ( Figure 2). Noticeably, the higher dosage of MMS yielded more and shorter DNA fragments than the moderate dose, indicating that the number of DNA backbone breaks is correlated to MMS dosage, and, furthermore, a DNA band close to the loading well which could correspond to the linear chromosomal DNA is present in 1.3 mM MMS-treated samples but absent from 2.6 mM MMS-treated samples. Subsequently, the intensity of DNA fragments gradually declined from 2 h in both 1.3 and 2.6 mM MMS treatment. Noticeably, from 6 to 24 h during 1.3 mM MMS treatment, most genomic DNA was retained in the loading wells, similar to that of the untreated samples, suggesting reestablishment of genome integrity after the repair of DNA breaks during the moderate treatment.  Next, we set out to analyze the DNA content distributions in the cells of MMS-treated cultures by flow cytometry (Figure 3). The results showed that DNA content distributions of untreated samples remained unchanged in the first 9 h of incubation. Then, the population of G1 and S phase cells decreased at 24 h, which could reflect that the cells were in a late exponential stage at the time. For the moderate MMS treatment, G1 and S phase cells were accumulated in the first 6 h of incubation, and a small fraction of DNA-less cells appeared at 9 h. By 24 h after MMS addition, most cells showed a DNA content of one or more than one chromosomes. Together, these data suggest that, while the majority of cells were recovered from MMS-mediated DNA damage, some cells underwent cell death. In the lethal MMS treatment, however, the DNA content distribution remained the same in the first 1 h, suggesting that although MMS induce immediate DNA backbone breakage, the DNA breaks were protected from degradation at this time. Then, DNA content gradually decreased as the DNA content peaks moved leftwards from 1 to 4 h during the drug treatment, and eventually a peak corresponding to DNA-less cells occurred at 6 h. Additionally, at 24 h, most cells became DNA-less cells. Thus, we concluded that lethal MMS-induced DNA breaks eventually lead to chromosomal DNA degradation, while DNA breaks after a moderate MMS treatment are largely repairable such that most cells are eventually recovered from the MMS-mediated DNA damage. Next, we set out to analyze the DNA content distributions in the cells of MMS-treated cultures by flow cytometry (Figure 3). The results showed that DNA content distributions of untreated samples remained unchanged in the first 9 h of incubation. Then, the population of G1 and S phase cells decreased at 24 h, which could reflect that the cells were in a late exponential stage at the time. For the moderate MMS treatment, G1 and S phase cells were accumulated in the first 6 h of incubation, and a small fraction of DNA-less cells appeared at 9 h. By 24 h after MMS addition, most cells showed a DNA content of one or more than one chromosomes. Together, these data suggest that, while the majority of cells were recovered from MMS-mediated DNA damage, some cells underwent cell death. In the lethal MMS treatment, however, the DNA content distribution remained the same in the first 1 h, suggesting that although MMS induce immediate DNA backbone breakage, the DNA breaks were protected from degradation at this time. Then, DNA content gradually decreased as the DNA content peaks moved leftwards from 1 to 4 h during the drug treatment, and eventually a peak corresponding to DNA-less cells occurred at 6 h. Additionally, at 24 h, most cells became DNA-less cells. Thus, we concluded that lethal MMS-induced DNA breaks eventually lead to chromosomal DNA degradation, while DNA breaks after a moderate MMS treatment are largely repairable such that most cells are eventually recovered from the MMS-mediated DNA damage.

MMS-Induced TopR1 Degradation is Prior to Chromatin Protein Degradation
To investigate the dynamics of chromatin architecture proteins during MMS-induced genomic DNA degradation in S. islandicus, we analyzed the level of TopR1, Sul7, and Cren7 in S. islandicus cells by western blot during 1.3 and 2.6 mM MMS treatment ( Figure 4). In the moderate MMS

MMS-Induced TopR1 Degradation is Prior to Chromatin Protein Degradation
To investigate the dynamics of chromatin architecture proteins during MMS-induced genomic DNA degradation in S. islandicus, we analyzed the level of TopR1, Sul7, and Cren7 in S. islandicus cells by western blot during 1.3 and 2.6 mM MMS treatment ( Figure 4). In the moderate MMS treatment, Cren7 and Sul7 contents remained more or less the same in the cells whereas TopR1 level decreased from 2 to 6 h. Then, at 9 h after drug addition, TopR1 level started to increase, indicative of full recovery of the cells from the MMS-mediated DNA damage. In the lethal MMS treatment, after the onset of TopR1 degradation at 2 h after drug addition, the level of the protein continuously decreased along with the incubation, which coincided with cellular DNA content reduction. Finally, cellular TopR1 was no longer detectable after 19 h of drug treatment. In addition, lethal MMS treatment also induced chromatin protein degradation at 6 h. The sequential degradation of TopR1 and chromatin proteins suggests that they play different roles in genome stability maintenance in this archaeon. treatment, Cren7 and Sul7 contents remained more or less the same in the cells whereas TopR1 level decreased from 2 to 6 h. Then, at 9 h after drug addition, TopR1 level started to increase, indicative of full recovery of the cells from the MMS-mediated DNA damage. In the lethal MMS treatment, after the onset of TopR1 degradation at 2 h after drug addition, the level of the protein continuously decreased along with the incubation, which coincided with cellular DNA content reduction. Finally, cellular TopR1 was no longer detectable after 19 h of drug treatment. In addition, lethal MMS treatment also induced chromatin protein degradation at 6 h. The sequential degradation of TopR1 and chromatin proteins suggests that they play different roles in genome stability maintenance in this archaeon.

Knockdown of TopR1 Transcripts Accelerates MMS-Induced Genomic DNA Degradation
To further investigate the functions of TopR1 in genome stability maintenance in S. islandicus, we employed a CRISPR-based RNA interference (CRISPRi) method previously developed in our laboratory [16] to reduce the level of TopR1 in the archaeal cells. The CRISPRi plasmid (pKD-topR1) contained an artificial mini-CRISPR locus with a single copy of two spacers that target two different positions on the topR1 mRNA. After transformation of S. islandicus E233S1 with the plasmid, the expression of TopR1 in the transformants (pKD-topR1/E233S1) was analyzed by western blot, and we found that the cellular TopR1 level was indeed much lower than that in the cells containing the reference plasmid pSeSD1 (pSeSD1/E233S1) ( Figure 5A). On the other hand, the growth rates of the two strains were similar at around 78 °C (data not shown). These results indicate that reduction of TopR1 did not impair the growth of this archaeon.
These strains were then employed to investigate the roles of TopR1 in genome stability upon MMS treatment. Cultures of pKD-topR1/E233S1 and pSeSD1/E233S1 were treated with different concentrations of MMS for 24 h during which cell samples were taken and analyzed for TopR1 content, cell viability, and DNA content distributions. As shown in Figure 5A, after 3 h of MMS treatment, TopR1 was partly degraded in both pKD-topR1/E233S1 and pSeSD1/E233S1, although the former expressed much less TopR1 protein. In addition, in both strains, increasing MMS concentrations did not lead to further TopR1 degradation at 3 h. Meanwhile, the survival rate of pKD-topR1/E233S1 was about four times lower than pSeSD1/E233S1 during 1.3 and 2 mM MMS treatment and about 90 times lower during 2.6 mM MMS treatment ( Figure 5B), indicating that knockdown of topR1 increased MMS sensitivity.
Flow cytometry analysis showed that a small population of DNA-less cells (about 8%) was formed in pKD-topR1/E233S1 cultures even without MMS treatment. Further, its S-phase cell population moderately increased compared to pSeSD1/E233S1 ( Figure 5C). The results indicate that, although the growth curve of pKD-topR1/E233S1 was similar to that of pSeSD1/E233S1, reduced TopR1 level also affected genome stability and/or DNA replication in pKD-topR1/E233S1 cells. At 24 h, the population of S phase cells in the pKD-topR1/E233S1 culture also reduced, suggesting that the culture also entered a late exponential stage similar to the pSeSD1/E233S1 culture.

Knockdown of TopR1 Transcripts Accelerates MMS-Induced Genomic DNA Degradation
To further investigate the functions of TopR1 in genome stability maintenance in S. islandicus, we employed a CRISPR-based RNA interference (CRISPRi) method previously developed in our laboratory [16] to reduce the level of TopR1 in the archaeal cells. The CRISPRi plasmid (pKD-topR1) contained an artificial mini-CRISPR locus with a single copy of two spacers that target two different positions on the topR1 mRNA. After transformation of S. islandicus E233S1 with the plasmid, the expression of TopR1 in the transformants (pKD-topR1/E233S1) was analyzed by western blot, and we found that the cellular TopR1 level was indeed much lower than that in the cells containing the reference plasmid pSeSD1 (pSeSD1/E233S1) ( Figure 5A). On the other hand, the growth rates of the two strains were similar at around 78 • C (data not shown). These results indicate that reduction of TopR1 did not impair the growth of this archaeon.
These strains were then employed to investigate the roles of TopR1 in genome stability upon MMS treatment. Cultures of pKD-topR1/E233S1 and pSeSD1/E233S1 were treated with different concentrations of MMS for 24 h during which cell samples were taken and analyzed for TopR1 content, cell viability, and DNA content distributions. As shown in Figure 5A, after 3 h of MMS treatment, TopR1 was partly degraded in both pKD-topR1/E233S1 and pSeSD1/E233S1, although the former expressed much less TopR1 protein. In addition, in both strains, increasing MMS concentrations did not lead to further TopR1 degradation at 3 h. Meanwhile, the survival rate of pKD-topR1/E233S1 was about four times lower than pSeSD1/E233S1 during 1.3 and 2 mM MMS treatment and about 90 times lower during 2.6 mM MMS treatment ( Figure 5B), indicating that knockdown of topR1 increased MMS sensitivity.
Flow cytometry analysis showed that a small population of DNA-less cells (about 8%) was formed in pKD-topR1/E233S1 cultures even without MMS treatment. Further, its S-phase cell population moderately increased compared to pSeSD1/E233S1 ( Figure 5C). The results indicate that, although the growth curve of pKD-topR1/E233S1 was similar to that of pSeSD1/E233S1, reduced TopR1 level also affected genome stability and/or DNA replication in pKD-topR1/E233S1 cells. At 24 h, the population of S phase cells in the pKD-topR1/E233S1 culture also reduced, suggesting that the culture also entered a late exponential stage similar to the pSeSD1/E233S1 culture.
At 24 h after treatment, 1.3 and 2 mM MMS induced a moderate increase of DNA-less cells in the pKD-topR1/E233S1 culture relative to that of pSeSD1/E233S1, while 2.6 mM MMS induced almost 100% DNA-less cells for both strains ( Figure 5C). To further reveal the roles of TopR1 in the response to MMS, we compared DNA content distributions between the two strains before DNA-less cell formation, and the results show that the DNA content in pKD-topR1/E233S1 cells was less than that in pSeSD1/E233S1 from 3 h to 9 h during 2 mM MMS treatment and at 3 h during 2.6 mM MMS treatment ( Figure 5D), indicating that the cells with a reduced TopR1 level underwent accelerated genomic DNA degradation during MMS treatment. At 24 h after treatment, 1.3 and 2 mM MMS induced a moderate increase of DNA-less cells in the pKD-topR1/E233S1 culture relative to that of pSeSD1/E233S1, while 2.6 mM MMS induced almost 100% DNA-less cells for both strains ( Figure 5C). To further reveal the roles of TopR1 in the response to MMS, we compared DNA content distributions between the two strains before DNA-less cell formation, and the results show that the DNA content in pKD-topR1/E233S1 cells was less than that in pSeSD1/E233S1 from 3 h to 9 h during 2 mM MMS treatment and at 3 h during 2.6 mM MMS treatment ( Figure 5D), indicating that the cells with a reduced TopR1 level underwent accelerated genomic DNA degradation during MMS treatment.

MMS Resistance of S. islandicus is Concomitant with the Association of TopR1 with Chromosomal DNAs
Previous research showed that 0.7 mM MMS is a lethal dosage for S. solfataricus cells, which is much lower than the lethal dosage (2.6 mM) for S. islandicus identified in this work. To verify the results, we compared the MMS sensitivity of S. solfataricus and S. islandicus. The two species were

MMS Resistance of S. islandicus is Concomitant with the Association of TopR1 with Chromosomal DNAs
Previous research showed that 0.7 mM MMS is a lethal dosage for S. solfataricus cells, which is much lower than the lethal dosage (2.6 mM) for S. islandicus identified in this work. To verify the results, we compared the MMS sensitivity of S. solfataricus and S. islandicus. The two species were grown and treated with MMS in the same conditions, and cell viability and DNA content distributions of the treated samples were analyzed. First, during 1.3, 2, and 2.6 mM MMS treatment, S. solfataricus exhibited about 4, 23, and over 100-times lower survival rate than that of S. islandicus, respectively, confirming that S. solfataricus is indeed more sensitive to MMS than S. islandicus ( Figure 6A). Further, the reduction of DNA content in S. solfataricus was also faster than that in S. islandicus, indicating that MMS induced more intensive DNA degradation in S. solfataricus ( Figure 6B and Figure S1). We also noticed that most S. solfataricus cells became DNA-less cells after treatment of 2 mM MMS in this study, which is much higher than the lethal dosage for S. solfataricus cells reported previously [13]. The difference between the two studies could be due to different growth and treatment conditions in experiments, or unknown strain differences.
respectively, confirming that S. solfataricus is indeed more sensitive to MMS than S. islandicus ( Figure  6A). Further, the reduction of DNA content in S. solfataricus was also faster than that in S. islandicus, indicating that MMS induced more intensive DNA degradation in S. solfataricus (Figures 6B and S1). We also noticed that most S. solfataricus cells became DNA-less cells after treatment of 2 mM MMS in this study, which is much higher than the lethal dosage for S. solfataricus cells reported previously [13]. The difference between the two studies could be due to different growth and treatment conditions in experiments, or unknown strain differences.
It was also reported that SsoTopR1 mainly exists in the cytoplasm of S. solfataricus and is not recruited into chromosomes during MMS treatment [13]. Further, AfuTopR1 binds to DNA nicks to protect nicked DNA from thermo-degradation in vitro [14]. Therefore, we analyzed the subcellular location of TopR1 in S. islandicus cells as well as in S. solfataricus cells. Cell lysates (total protein, TP) of S. solfataricus and S. islandicus were fractionated into soluble supernatant (S, cytoplasm fraction) and pellet fractions (P, DNA-rich fraction) as previously described [17]. The abundance of TopR1 and PCNA3 in TP, S, and P fractions was analyzed by western blot. As shown in Figure 6C, while most TopR1 in S. solfataricus existed in the cytoplasm fraction, more than a half of TopR1 in S. islandicus was bound to chromosomes, suggesting that the chromosome-associated TopR1 may be responsible for the higher resistance of S. islandicus to MMS. We further analyzed the dynamics of TopR1 protein from the cytoplasm fraction and chromatinrich fraction during MMS treatment in S. islandicus Rey15A. Treated S. islandicus Rey15A cells were collected and fractionated as previously described [17], and levels of TopR1 were analyzed in each fraction. The results indicate that TopR1 levels were reduced in both cytoplasm fraction and chromatin-rich fraction (Figure 7), suggesting that MMS-induced degradation occurs to TopR1 from both fractions in S. islandicus Rey15A. It was also reported that SsoTopR1 mainly exists in the cytoplasm of S. solfataricus and is not recruited into chromosomes during MMS treatment [13]. Further, AfuTopR1 binds to DNA nicks to protect nicked DNA from thermo-degradation in vitro [14]. Therefore, we analyzed the subcellular location of TopR1 in S. islandicus cells as well as in S. solfataricus cells. Cell lysates (total protein, TP) of S. solfataricus and S. islandicus were fractionated into soluble supernatant (S, cytoplasm fraction) and pellet fractions (P, DNA-rich fraction) as previously described [17]. The abundance of TopR1 and PCNA3 in TP, S, and P fractions was analyzed by western blot. As shown in Figure 6C, while most TopR1 in S. solfataricus existed in the cytoplasm fraction, more than a half of TopR1 in S. islandicus was bound to chromosomes, suggesting that the chromosome-associated TopR1 may be responsible for the higher resistance of S. islandicus to MMS.
We further analyzed the dynamics of TopR1 protein from the cytoplasm fraction and chromatin-rich fraction during MMS treatment in S. islandicus Rey15A. Treated S. islandicus Rey15A cells were collected and fractionated as previously described [17], and levels of TopR1 were analyzed in each fraction. The results indicate that TopR1 levels were reduced in both cytoplasm fraction and chromatin-rich fraction (Figure 7), suggesting that MMS-induced degradation occurs to TopR1 from both fractions in S. islandicus Rey15A. Figure 7. Western blot analysis of TopR1 level from cytoplasm fraction and chromatin-rich fraction during MMS treatment. S. islandicus Rey15A cells was treated with 2.6 mM MMS and cells were collected at 0, 2, and 4 h to prepare total protein (TP), cytoplasm fraction (supernatant, S) and chromatin-rich fraction (pellet, P). Antisera against PCNA3 and Cren7 were used as a loading control.

Discussion
In this study, we analyzed the effects of MMS on the genome integrity of S. islandicus, a thermophilic crenarchaeon and investigated the functions of TopR1 in genome stability maintenance. Several lines of experimental evidence indicate that SisTopR1 protects MMS-induced DNA breaks from thermo-degradation in a chromosome-associated manner in vivo.
First, we have demonstrated that MMS treatment immediately induces genomic DNA breakage in Sulfolobus. MMS does not directly introduce DNA breaks to genomic DNA in yeast and mammalian cells; however, MMS-induced DNA lesions could be converted into DNA breaks by base excision repair (BER) processing or DNA replication collapse [10]. In addition, MMS-induced DNA lesions are heat-labile in vitro and are converted to single stranded DNA breaks (SSBs) at 50 °C; two close SSBs will results in the formation of a DSB [18]. Therefore, the DNA breaks observed in this study can be derived from two main sources: (i) thermo-degradation of MMS-induced lesions at the physiologic growth temperature of S. islandicus and (ii) processing of the lesions by BER pathway. Considering the rapid occurrence of genomic DNA breakage, it is more likely that thermodegradation of MMS-induced lesions constitutes the main source of MMS-induced DNA breaks. As a result, this suggests that MMS can function as a direct DNA break agent in thermophiles.
Second, lethal MMS-induced genomic DNA degradation is evidenced by the decline of DNA fragments as shown by PFGE analysis and gradual reduction of cellular DNA content as shown in flow cytometry analysis. The degradation could be directly triggered by MMS-induced massive SSBs, since an SSB can accelerate thermo-degradation of genomic DNA in vitro: first, supercoiled DNA is not likely to be denatured, while nicks initiate dsDNA denaturation [7]; second, the exposed ssDNA is even more fragile, exhibiting a much higher depurination rate and breakage rate, leading to dsDNA breakage and further degradation [8,9]. More strikingly, we demonstrate that even though MMS induces immediate genomic DNA backbone breakage, cellular DNA content does not start to reduce until TopR1 degradation occurs (Figures 2-4), supporting the previous hypothesis that TopR functions in protecting damaged DNA from degradation [13,14]. Further, chromatin protein degradation occurs after TopR1 degradation and coincides with DNA-less cell formation, a common cell death consequence of the treatment of different DNA damage agents (submitted). The sequential degradation of TopR1 and chromatin proteins suggest that after MMS introduces DNA breaks, TopR1 is first subjected to degradation, leading to genomic DNA degradation, which triggers a cell death signal and induces chromatin protein degradation and DNA-less cell formation. The findings also suggest that TopR1 and chromatin proteins play distinct roles in protecting genomic DNA. We reason that TopR1 might bind to the regions that are loosely compacted or not compacted by chromatin proteins. Nevertheless, the detailed mechanisms for MMS-induced DNA degradation and the roles of these chromatin architecture proteins remain to be clarified.
Lastly, the roles of TopR1 in genome stability maintenance have been demonstrated by CRISPRi of TopR1 expression. The strain with decreased TopR1 levels (pKD-topR1/E233S1) is more sensitive to MMS treatment than the reference strain (pSeSD1/E233S1). Further, MMS-induced DNA degradation is strongly accelerated in pKD-topR1/E233S1, suggesting that damaged DNA is prone Figure 7. Western blot analysis of TopR1 level from cytoplasm fraction and chromatin-rich fraction during MMS treatment. S. islandicus Rey15A cells was treated with 2.6 mM MMS and cells were collected at 0, 2, and 4 h to prepare total protein (TP), cytoplasm fraction (supernatant, S) and chromatin-rich fraction (pellet, P). Antisera against PCNA3 and Cren7 were used as a loading control.

Discussion
In this study, we analyzed the effects of MMS on the genome integrity of S. islandicus, a thermophilic crenarchaeon and investigated the functions of TopR1 in genome stability maintenance. Several lines of experimental evidence indicate that SisTopR1 protects MMS-induced DNA breaks from thermo-degradation in a chromosome-associated manner in vivo.
First, we have demonstrated that MMS treatment immediately induces genomic DNA breakage in Sulfolobus. MMS does not directly introduce DNA breaks to genomic DNA in yeast and mammalian cells; however, MMS-induced DNA lesions could be converted into DNA breaks by base excision repair (BER) processing or DNA replication collapse [10]. In addition, MMS-induced DNA lesions are heat-labile in vitro and are converted to single stranded DNA breaks (SSBs) at 50 • C; two close SSBs will results in the formation of a DSB [18]. Therefore, the DNA breaks observed in this study can be derived from two main sources: (i) thermo-degradation of MMS-induced lesions at the physiologic growth temperature of S. islandicus and (ii) processing of the lesions by BER pathway. Considering the rapid occurrence of genomic DNA breakage, it is more likely that thermo-degradation of MMS-induced lesions constitutes the main source of MMS-induced DNA breaks. As a result, this suggests that MMS can function as a direct DNA break agent in thermophiles.
Second, lethal MMS-induced genomic DNA degradation is evidenced by the decline of DNA fragments as shown by PFGE analysis and gradual reduction of cellular DNA content as shown in flow cytometry analysis. The degradation could be directly triggered by MMS-induced massive SSBs, since an SSB can accelerate thermo-degradation of genomic DNA in vitro: first, supercoiled DNA is not likely to be denatured, while nicks initiate dsDNA denaturation [7]; second, the exposed ssDNA is even more fragile, exhibiting a much higher depurination rate and breakage rate, leading to dsDNA breakage and further degradation [8,9]. More strikingly, we demonstrate that even though MMS induces immediate genomic DNA backbone breakage, cellular DNA content does not start to reduce until TopR1 degradation occurs (Figures 2-4), supporting the previous hypothesis that TopR functions in protecting damaged DNA from degradation [13,14]. Further, chromatin protein degradation occurs after TopR1 degradation and coincides with DNA-less cell formation, a common cell death consequence of the treatment of different DNA damage agents (submitted). The sequential degradation of TopR1 and chromatin proteins suggest that after MMS introduces DNA breaks, TopR1 is first subjected to degradation, leading to genomic DNA degradation, which triggers a cell death signal and induces chromatin protein degradation and DNA-less cell formation. The findings also suggest that TopR1 and chromatin proteins play distinct roles in protecting genomic DNA. We reason that TopR1 might bind to the regions that are loosely compacted or not compacted by chromatin proteins. Nevertheless, the detailed mechanisms for MMS-induced DNA degradation and the roles of these chromatin architecture proteins remain to be clarified.
Lastly, the roles of TopR1 in genome stability maintenance have been demonstrated by CRISPRi of TopR1 expression. The strain with decreased TopR1 levels (pKD-topR1/E233S1) is more sensitive to MMS treatment than the reference strain (pSeSD1/E233S1). Further, MMS-induced DNA degradation is strongly accelerated in pKD-topR1/E233S1, suggesting that damaged DNA is prone to thermos-degradation upon an insufficiency of TopR1. The data, together with the fact that MMS induces SSBs at high temperature in vitro [18] and rapid genomic DNA breakage in Sulfolobus ( Figure  2), indicate that TopR1 functions in the thermo-protection of SSBs in vivo. Conceivably, TopR1 should also protect the DNA breaks derived from spontaneous DNA hydrolysis at high temperatures, which could represent a general function for reverse gyrase in thermophiles.
In addition, knockdown of TopR1 by CRISPRi in S. islandicus cells also leads to a moderate accumulation of S phase cells. This either suggests that (1) TopR1 has a role in DNA replication or that (2) the reduced level of protection for spontaneous genomic DNA breaks in TopR1-CRISPRi cells yields DNA replication fork collapse. Indeed, it has been suggested that TopR1 is involved in cell duplication in S. solfataricus [19]. On the other hand, we observed that moderate MMS treatment also induces accumulation of S phase cells (Figure 3), supporting that unrepaired DNA breaks arrest DNA replication in TopR1-CRISPRi cells. Currently, it is not possible to distinguish the two possible mechanisms experimentally.
Moreover, the association of SisTopR1 to chromatin and the higher MMS resistance of S. islandicus suggest that the DNA protection role of TopR1 is related to its association to DNA. The hypothesis is consistent with the fact that AfuTopR protects nicked DNA possibly by binding to the breaks in vitro [14]. On the other hand, even though TopR1 proteins from S. islandicus and S. solfataricus exhibit high sequence similarity ( Figure S2), most SsoTopR1 protein exists in the cytoplasm fraction. Further, SsoTopR1 is recruited to the chromatin-rich fraction upon ultraviolet (UV) treatment, suggesting translocation of TopR1 upon DNA damage [17]. A similar translocation mechanism might be adopted in S. islandicus under normal growth conditions. TopR1 degradation has been proved to be very specific. It was only observed during MMS treatment, while other stressful conditions, including hydroxyurea (HU) and UV treatment, fail to induce the phenomena [13], even though UV also induces DNA breaks and arrests DNA replication in S. solfataricus [20]. We reason that MMS and UV induce different types of DNA breaks: MMS-induced heat-labile DNA lesions are immediately converted to massive SSBs at the physiologic growth temperature of Sulfolobus, while UV-induced DNA lesions may be converted to DNA breaks by DNA replication fork collapse. The findings hint that TopR1 is specifically induced by massive SSBs.
Nevertheless, the question regarding the biological meaning of TopR1 degradation remains to be answered. Possibly, TopR1 degradation represents an important step in the archetypal programmed cell death, as hypothesized previously [13], such that TopR1 degradation results in thermo-degradation of damaged DNA, leading to cell death. However, we have shown that TopR1 is degraded to a similar extent from 2 to 6 h in the moderate and the lethal MMS treatment (Figures 4 and 5A). The difference between the treatments is that TopR1 level recovered from 6 to 24 h during moderate MMS treatment but continuously decreased upon lethal MMS treatment. Therefore, we assume that TopR1 degradation also plays a role in DNA repair to increase the local access to DNA repair machineries of the damaged site upon moderate treatment similar to that in Eukaryotes [21,22]. Indeed, TopR-coated dsDNA exhibits limited accessibility in vitro [14]. Furthermore, the two hypotheses are not mutually exclusive: upon moderate MMS treatment, TopR1 degradation increases the local access to DNA repair machineries and facilitates DNA repair and cell survival; upon lethal MMS treatment, DNA repair machineries may fail to repair massive DNA breaks in a timely fashion. As a result, the decrease of cellular TopR1 content would result in exposure of more DNA backbone breaks from thermo-degradation, leading to cell death.

Construction of the TopR1 Knockdown Strain
The strategy for construction of the TopR1 (gene locus: SiRe_1581) knockdown plasmid was similar to that described in [16]. Specifically, we chose two unique sequence regions preceded by a GAAAG motif at the 5 end from the antisense strand as spacers to abolish DNA cleavage activity [23]. The oligonucleotides for them are shown in the Supplementary Table S1. The spacer fragments were prepared by annealing and extension of each pair of oligonucleotides. Then, the two spacer fragments were mixed and the mixture was used as a template for further amplification with S1-forward and S2-rev-SalI as primers. The obtained fragments were inserted into pSeSD1, an expression vector widely used in S. islandicus [24], between StuI and SalI. Then, the generated plasmid was transferred into Escherichia coli DH5α, and the colonies were screened by sequencing. At last, a plasmid containing one copy of each spacer was transferred into S. islandicus E233S1 and three transformants (pKD-topR1/E233S1) were picked and grown for further analysis. In the meantime, pSeSD1 was also transferred into the genetic host as control (pSeSD1/E233S1).

Culture Growth and Treatment
The genetic host, S. islandicus E233S1, was grown in SCVU medium, containing 0.2% sucrose (S), 0.2% Casamino acids (C), 5 µL/mL of vitamin mixture solution (V) and 20 µg/mL uracil (U), as describe previously [25], while other stains, including pSeSD1/E233S1, pKD-TopR1/E233S1, S. islandicus Rey15A, and S. solfataricus P2 were grown in SCV medium that lacks uracil compared to SCVU. Before treatment, the cultures were grown at exponential phase for at least 72 h, and then early exponential phase cultures (The optical density at a wavelength of 600 nm is about 0.2) were supplemented with MMS up to the indicated concentrations. Aliquots without the drug were set as controls. During treatment, the cultures were grown at 78 • C in the dark and the samples for OD measuring, cell viability assays, flow cytometry, PFGE, and western blot were taken at indicated time points.

Cell Viability
Cells were collected from 1 mL of untreated or treated cultures by centrifugation and resuspended in 1 mL of fresh medium of the same composition. Then, resuspended cells were diluted by 10 2 and 10 4 times, and for each dilution as well as the original cell suspension, 10 µL and 100 µL of cell suspension were mixed with pre-warmed top layer gel and spread onto SCV plates as previously described (SCVU for S. islandicus E233S1) [25]. The plates were incubated at 75 • C for 6 days; then, plates with suitable colony densities were selected for colony counting. The cell viability was determined in three independent experiments.

Flow Cytometry
To analyze DNA content and cell size distributions, 300 µL of samples was taken from a culture to which 700 µL of absolute ethanol was added. The samples were stored at 4 • C for at least 12 h to fix cells. Then, fixed cells were collected by centrifugation at 2800 rpm for 20 min and washed with 1 mL of 10 mM Tris-NaCl, pH 7.5, and 10 mM MgCl 2 . Cells were collected again and resuspended in 140 µL of staining dye containing 40 µg/mL ethidium bromide (Sigma-Aldrich, St. Louis, MO, USA) and 100 µg/mL mithramycin A (Apollo chemical, Tamworth, UK). Stained cell samples were analyzed by an Apogee A40 cytometer (Apogeeflow, Hertfordshire, UK) equipped with 405 nm laser. A dataset of at least 60,000 cells was collected for each sample.

Western Blot
Cells from 10 mL of the untreated or treated cultures were collected and resuspended in bufferA (50 mM Tris-NaCl, pH 8.0, 100 mM NaCl). Then, 5× SDS-loading buffer was added into the solution, followed by heating at 95 • C for 10 min, to disrupt the cells. The final volume of the samples was calculated according to the optical density to make sure that the number of cells for each line was same. Then, the proteins were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to a polyvinylidene difluoride (PVDF) membrane (Bio-Rad, Hercules, CA, USA) and the membrane was incubated with antibodies for Cren7 [26], Sul7d [27], TopR1, and PCNA [28]. Afterwards, the membrane was incubated with anti-rabbit horseradish peroxidase (HRP) Secondary antibody (Thermo Fisher Scientific, Waltham, MA, USA) and the enhanced chemiluminescence (ECL) western blot substrate (Thermo Fisher Scientific, Waltham, MA, USA) was used to detect the signals. The antiserum against TopR1 was raised in rabbits using purified recombinant TopR1 protein.

Pulsed-Field Gel Electrophoresis (PFGE)
About 6 × 10 9 cells (given that 1 mL culture of OD 600 = 1.0 contains 10 9 cells) were collected from 30 mL of untreated or treated S. islandicus culture by centrifugation. The cells were washed with 50 mL of TEN buffer (50 mM Tris-Cl, pH 8, 50 mM EDTA, pH 8, 100 mM NaCl) and resuspended in 150 µL prewarmed TEN buffer (37 • C). Then, the cell suspension was mixed with 150 µL of 0.8% low-melting-point agarose (precooled to 37 • C) and transferred to a plug mold with 85 µL per well. After solidification at 4 • C for 30 min, the plugs were released from the mold into a 1.5 mL microfuge tube containing 1 mL of NDS solution (0.5 M EDTA, 0.12% Tris, 0.55 M NaOH, pH 8.5, 1% N-lauroyl sarcosine sodium salt) and 1 mg/mL proteinase K (Sigma-Aldrich, St. Louis, MO, USA) and incubated for two days at 30 • C. The old proteinase K-containing NDS buffer was replaced by a fresh solution of the same composition after 1 day's incubation. The plugs then were washed three times with NDS and stored at 4 • C. For PFGE, about one fourth of a plug was transferred into each well of an agarose gel, and electrophoresis was conducted with a CHEF-DR III pulsed-field electrophoresis system (Bio-Rad, Hercules, CA, USA) under the running condition of 3.5 V/cm with the switch time from 5 to 100 s for 40 h. Then, the gel was stained with a nucleic staining dye SYBR gold (Thermo Fisher Scientific, Waltham, MA, USA), and the image was captured by using Typhoon FLA 7000 (GE Healthcare, Chicago, IL, USA).

Isolation of DNA-Rich Fraction
The procedure to isolate DNA-rich fraction from Sulfolobus was modified from that described in [29]. Specifically, about 8 × 10 9 cells from different Sulfolobus species were harvested by centrifugation, washed with extraction buffer (50 mM Tris-HCl pH 7, 15 mM MgCl 2 , 50 mM NaCl, 1 mM DTT, 0.4 M sorbitol) and resuspended with 400 µL of the same buffer. Then, half volume of the cell solution was mixed with 5× SDS-loading buffer (total protein, TP), while the rest was supplemented with 0.5% Triton X-100, placed on ice for 30 min, and subjected to centrifugation at 13,000 rpm for 10 min at 4 • C. The supernatant (S, cytoplasm fraction) derived from the centrifugation was then mixed with 5× SDS-loading buffer, while the pellet (P, DNA-rich fraction) was resuspended with 200 µL of extraction buffer and also mixed with 5× SDS-loading buffer. At last, all three fractions were heated at 95 • C for 10 min and 10 µL of each fraction were run on SDS-gels and analyzed by western blot.

Conclusions
Lethal MMS treatment induces immediate genomic DNA breakage, followed by genomic DNA degradation. The latter co-occurs with TopR1 degradation. Further, cells with a lower TopR1 level are more sensitive to MMS and undergo accelerated genomic DNA degradation during MMS treatment. Together, the data indicate that TopR1 could protect DNA breaks from degradation in vivo.