s2fieldsofstudy: Biology Chemistry

text:
Bisphenol a and human diseases. mechanisms of action

The review describes the molecular mechanisms and biological effects of bisphenol A exposure, which is a chemical (ecotoxicant) that destroys the endocrine system and has epigenetic toxicity.

BacKGRound
Many chemical and physical environmental factors, depending on the dose and duration of exposure, can result in anomalies in development in embryonal and postnatal periods and become the reason for a number of diseases of adults. Currently, special attention of researchers is focused on endocrine-disrupting chemicals (EDCs). They are rather widely spread in the environment and are natural or synthetic compounds, which, after entering the organism even in small doses, can prevent the biosynthesis, storage, release, transfer, and/or receptor interaction of endogenous hormones, changing their functions and destroying the system of internal regulation of the organism. This, in turn, results in increased number of pathologies connected to hormonal disorders. In particular, obesity, diabetes mellitus, and various oncological diseases (breast, ovarian, prostate, and testicular cancers) can occur along with changes in the reproductive system (cryptorchidism, hypospadias, low quality of seminal fluid, and female sterility), cognitive disorders, and deviations in behavior and neuromental development. According to the latest data, only a small part of the available 800 commercial EDCs has been checked for potential effects causing disorders in the functioning of the endocrine system [1,2].
EDCs primarily affect people and animals via oral, dermal, and inhalation ways. Their bioavailability depends on the mode of penetration. As not all absorbed EDCs can be metabolized, the original compounds become bioavailable. For the major part of EDCs, the most biologically active form is nonmetabolized compound also known as the final toxicant reacting with organic molecules. After entering the blood flow, the final toxicant is able to reach the target cell(s) and affect it [3,4]; the phenotypical consequences caused by EDC depend a lot on the gap of effect. The EDC effect is mostly crucial for the organism during pregnancy, babyhood, early childhood, and teenage stage. As the mechanism of detoxication in the developing fetus and the newborn is not finally formed, the organism is especially sensitive to EDC in these periods. In the prenatal and neonatal periods, the targets for the EDC effect are the primary germ cells. This, in turn, can result in not only to a disruption of gametogenesis in children whose mothers have been directly affected by EDC, but also to the transmission of different phenotypic anomalies Генетическая токсиколоГия (including predisposition to socially significant diseases) in a row of generations.
Currently, the molecular mechanisms of the EDC effect are unknown. Available data indicate that the mechanisms of the EDC effect are complicated; studies in this area are decisive for understanding the occurrence of unfavorable phenotypes as well as for the development of strategies of interference and/or preventive actions [5].
It is noteworthy that some chemicals due to their nature can have negative "biological effects," although according to up-to-date regulations they are attributed to nontoxic ones after appropriate tests, and the possible remote consequences of such chemicals effect are not considered [6]. In this regard, it is necessary to consider the new concept of toxicity, namely the "epigenetic toxi city" [7]. Epigenetic toxicity is a phenomenon in which exogenous chemicals affect the epigenome and unfavorably affect living organisms, which can explain the long-term effects and remote consequences of the chemical impact as well as the predisposition to diseases caused by harmful environmental factors. Due to the development of new and improvement of existing analytical technologies, the number of chemicals that have epigenetic toxicity is constantly growing [8], and the understanding of the molecular mechanisms of epigenetic toxicity is enhanced.
This review describes the molecular mechanisms and biological effects of ecotoxicant bisphenol A (BPA) exposure that is attributed to EDC and, as it becomes clear, has epigenetic toxicity.

chemical pRopeRties and occuRRence of Bpa
BPA (4,4'-dihydroxy-2,2-diphenylpropane) is one of the widespread organic synthetic compounds. BPA is used in the industry for the production of various plastic items and is available in the content of epoxy resins used as coating of water supply pipes and the inner surface of cans and packages for food and drinks [9][10][11]. BPA can be released from containers and enter food and drinks and then can be accumulated in humans and animals [12,13]. BPA can enter humans via the -gastrointestinal tract as well as through the skin, for example, in contact with thermal paper [14]. As the modern life is "surrounded" by plastic items, BPA exposure on living organisms occurs permanently and in different doses.

patholoGies connected to Bpa chRonic eXposuRe
For several decades, studies on the different doses of BPA exposure on health have been conducted around the world using laboratory animals and in clinical practice. Currently, it has been shown that BPA has hepatotoxicity, and its exposure can result in oncological diseases (breast cancer, prostate cancer, and cancer of thyroid glands) and pathologies of the nervous system (disturbance of neurogenesis, stroke, and Parkinson's disease), cardiovascular system (ischemic heart disease, hypertensive disease, and clotting defect), endocrine system (diabetes and obesity), and reproductive system (disturbance of sexual cycle, endometriosis, and changes in breast and prostate gland and testis). BPA exposure can be one of the reasons for chronic respiratory diseases (asthma) as well as arrested development and mental disorders (anxiety, depression, hyperactivity, and aggression) [15][16][17][18].
Currently, approximately 347 million people in the world have diabetes. Together with genetic factors, the possible reasons promoting the development of this disease include ways of life and intake of incorrect food as well as the inevitable chronic effect of xenobiotics. Experimental studies demonstrated that BPA affects the metabolism of glucose with the participation of different mechanisms, including resistance to insulin, dysfunction of beta-cells of the pancreatic gland, adipogenesis, inflammation, and oxidation stress, which prove the availability of the connection between BPA exposure and diabetes development [19,20]. It has been shown that BPA can stipulate the dysfunction of mitochondria due to oxidation stress (for example, in GC-2 cells) and lipid metabolism (for example, in HepG2 and INS-1 cells) [21]. It is supposed that BPA exposure can promote the effect of other risk factors of diabetes, which result in obesity, regulate eating behaviors, or change the differentiation of adipocytes.
BPA exposure is associated with chronic respiratory diseases such as asthma. For example, children with asthma demonstrated increased concentrations of BPA in urine [22]. Besides, it has been shown that BPA exposure in the prenatal period increases the risk of dyspnea development in children in the neonatal period, although a further negative effect of BPA is reduced in the first 3 years after birth [23].
At the present time, it has been shown that men and women exposed to BPA have a higher risk of developing of the coronary artery atherosclerosis. Thus, patients with severe stenosis of coronary arteries demonstrated increased BPA concentration in urine compared with people without atherosclerosis [24,25]. It also has been shown that carriers of some genetic polymorphisms are more sensitive to cardiovascular and respiratory diseases associated with reduced cell response to oxidation stress [26]. It should be noted that one of the possible molecular mechanisms of BPA exposure can be its impact on oxidation stress [27].
BPA can result in changes in the brain structure and mental and neurological disorders. For example, mice and rats exposed to BPA were more aggressive compared with controls. This was observed only in certain age periods and was not connected to the increase in testosterone concentration [28,29]. In studies of laboratory animals it was found that BPA exposure in the prenatal period affects brain development. Thus, large doses of BPA reduce the proliferation activity of multipotent neuronal stem cells; low doses, on the contrary, accelerate the differentiation GEnEtiC toxiColoGy and migration of neurons. This further results in abnormal neocortical architecture and corticothalamic projection and disturbed neurotransmitter system and behavior in postnatal period and adult age [30,31]. In addition, it was observed that BPA exposure in the early postnatal period leads to vacuolization, pycnosis, edema, degenerative changes, reduction of sizes and number of cells in the cerebral hemispheres and cerebellum, as well as results in the disturbance of hypothalamus sexual differentiation. In cultivated cells of the hypothalamus of rat embryos, BPA exposure caused the development of dendrites and synapses by increasing the level of presynaptic protein of synapsin I and microtubulin-associated protein 2 [27,32]. It has been shown that BPA can cause cognitive disorders, autism, schizophrenia, Parkinson's disease, and Alzheimer's disease [18,33,34].
Epidemiological studies demonstrated that BPA can cause disorders of the reproductive system and sexual behavior of men and women, although no deviations have been observed in the genitals and hormonal status [35][36][37]. However, according to the latest data, it cannot be considered that BPA exposure in small doses on adults actually affects reproductive health [38]. Probably, this is connected to the fact that different populations (and groups) have been studied in published works, different doses of BPA have been examined, and different schemes and methods of BPA measurement in biological fluids have been used.
Many works showed the connection between BPA exposure during pregnancy and pathologies in fetal development. Available data state that if the mother took food containing BPA, then this toxicant was detected in the blood serum, follicular fluid, and amniotic fluid as well as in embryonal serum. This indicates that BPA can penetrate the placenta (even in small doses) and has a negative effect during the entire prenatal period [39,40]. The analysis of BPA content in the organism demonstrated reduced ability to metabolize the chemical in mothers, often coinciding with fetal development defects [41]. For example, it was shown that BPA intrauterine exposure resulted in anomalies in the development of genitals of boys in approximately 37% of cases [42], and it was the reason for prematurity and the birth of children with small weight (especially male babies) [43]. Therefore, Welshos et al. [44] called the inborn defects and disturbances in development caused by BPA as "the large effects of small exposures." Moreover, presented data indicate that exposure on fetus of small doses of BPA changes cell proliferation and affects apoptosis and the time of breast development, which can further stipulate the predisposition to breast cancer in adult age [45,46]. BPA exposure during pregnancy, in combination with diet enriched with fats, significantly increases the risk of breast cancer development in offspring [47,48]. It is supposed that BPA can enhance the oncogenesis of the breast by the direct stimulation of estrogen-dependent growth of tumor cells and/or by molecular changes in fetal glands without associated morphological changes [47]. It should be noted that BPA can also affect the proliferation and apoptosis of ovary cells and terminate steroidogenesis in ovaries by changing steroidogenic enzymes, which in turn can promote the progression of ovarian tumor [48,49].
The latest data indicate that exposure of small, ecologically valuable doses of BPA in the embryonal period affects the cells of the prostate gland, enhancing the predisposition to premalignant lesions of this organ and hormonal disorders in adults. There is an opinion that cells of the prostate gland are more sensitive to BPA exposure in the embryonal period than in adult age. A number of researchers demonstrated that BPA could enhance the proliferation and migration of prostate cancer cells and induce DNA adducts in case of pathology [50,51].
Currently, the molecular mechanisms in which BPA affects the fetus and causes the development of ovarian, breast, and prostate cancers in adults are unclear, requiring further studies. Propositions have been made about the possible direct interaction of BPA with receptors of steroid hormones [estrogenic (ER) and androgenic (AR)], which play a decisive role in the origin and progression of these pathologies. In particular, ERα and ERβ start expression on the 12 th day of embryonal development in mesenchymes surrounding the embryonal anlage and regulate the growth of breast canals both before and after the birth. That is why BPA exposure in these periods can be crucial for the development of breast cancer in adult age [52]. The mechanisms of BPA exposure in prostate cancer are more complicated compared with that in breast and ovarian cancers, as the studies shown.
In general, considering the connection of BPA with different pathologies, two main conclusions can be made: (1) BPA is a typical xenoestrogen, and its estrogenic, estrogen-independent "steroid" activity is probably involved in the carcinogenesis of different organs and development of endocrine and/or hormone-dependent diseases, and (2) BPA exposure (even in small doses) in critical periods of ontogenesis (prenatal, neonatal, and teenage) can result in long-term negative effects in adults.

moleculaR mechanisms of Bpa eXposuRe on liVinG oRGanisms: connection to chRonic diseases
The issue of the mechanisms in which BPA can negatively affect humans and animals is still open. Currently, it is accepted that BPA acts as mutagen as well as the endocrine-active compound that affects DNA methylation and histone modifications. These mechanisms do not conflict, although studies on the epigenetic mechanisms of BPA exposure are still insufficient; most likely, they depend on its endocrine activity [15,[53][54][55].  The main mechanisms of BPA exposure on vertebrates are presented in Fig. 1.

Genetic© damages© caused© by© BPA
Different cell lines of humans and animals have been used to demonstrate that BPA is genotoxic and cytotoxic. It causes disorders of the cell cycle (in both mitosis and meiosis) and results in the gene, chromosomal, and genome mutations [51,56,57]. The cases of aneuploidy due to the disorders of the segregation of chromosomes during cell division are described for Chinese hamster V79 (lung fibroblasts) and golden hamster SHE (embryonal cells) cell lines [58]. BPA is frequently a reason for DNA damage, formation of DNA adducts, and apoptosis. For example, cell cultivation in the presence of BPA caused apoptosis in ER-positive of breast adenocarcinoma MCF-7 cell line and ER-negative of the human embryonal kidney HEK293 cell line as well as in the line of male germ cells of mice GC-2 [59][60][61]. It has been reported that the speed of formation of DNA adducts depends on the dose of BPA exposure; in particular, the larger the dose is, the quicker is the formation of such compounds. In the cell line of human prostate gland after exposure to large doses of BPA, DNA adducts formed within 24 h, whereas under the effect of low doses of this ecotoxicant their formation took 2 months [51,62]. It is supposed that the mutagenic effect of BPA, as of any other xenobiotics, can be due to the formation of free radicals, electrophiles, nucleophiles, and redox reagents that are accumulated and damage the plasma membrane and cell components [63]. Genetic damages caused by BPA exposure can result in changes in proteome in the breast and can be the reason of inborn defects, miscarriage, female and male sterility, and the development of many other pathologies mentioned above.

Mechanisms©of©BPA©effect©as©a©substance©destroying© the© endocrine© system
BPA is a xenoestrogen rather than an estrogen imitator. Its effect on the organism as a synthetic hormone is explained by the fact that, like steroid hormones, it has phenol groups; therefore, the nuclear receptors of estrogen (ERα and ERβ as well as recently detected in bone ERγ) perceive BPA as the signal for the initiation of the estrogenic pathway of the activation of transcription of estrogensensitive genes. In vertebrates, this toxicant can change hormonal balance by directly interacting with receptors ERα, ERβ, and ERγ or affecting enzymes by ensuring the metabolism of these hormones. For example, BPA can affect the ERβ-mediated transcription of the target genes by inhibiting ERβ degradation and ubiquitination [64].
It should be noted that nuclear receptors ERα and ERβ are functionally and genetically different; they differ in their affinity and specificity and have different spatial time types of expression. In this regard, cells of different types can respond in a different way to the same estrogenic in-GEnEtiC toxiColoGy centives depending on the ratio and expression of the two subtypes of receptors in the cell; therefore, the "pathogenic" effect of BPA can be different in different types of tissue. Masking for natural germ hormones, BPA can disturb endocrine regulation and result in different changes in the target organs of estrogens, including the brain, ovaries, thyroid gland, breasts, and prostate gland. Thus, the interaction of BPA with receptors of steroid hormones can be the reason for hormone-associated oncological diseases of the ovaries, breasts, and prostate gland [52].
Currently, the existence of additional membrane receptors to estrogens in the brain is supposed (similar to catecholaminergic receptors detected in the pancreas), which can explain the mechanism of estrogen effect on cognitive functions, pain development, delicate motor functions, emotional behavior, neuroprotection action in Parkinson's and Alzheimer's diseases, multiple sclerosis, depression, schizophrenia, and cerebral thrombosis [16]. In this connection, the presence of BPA in the organism can have a negative effect on these processes.
Moreover, there are data about BPA effects directly on the expression of the genes-receptors of hormones, in particular estrogens. For example, this has been demonstrated in the culture of cells of the rat cerebellum and human neuroblastoma as well as on human cell lines H295R (suprarenal cortex, angiotensin II sensitive, and steroidproducing line), HEK293 (embryonic kidney cells), and HepG2 (hepatic carcinoma) [65][66][67].
BPA is related to endocrine destructors, as it can interact with classical and nonclassical membrane receptors of estrogens. BPA exposure on metabotropic receptors transfers chemical signals to receptors jointed with G-proteins (for example, GPR30) and receptors jointed with the fragments, thus resulting in the disturbance of the regulatory ways of androgens, glucocorticoids, thyroid hormone, prolactin, insulin, and the dopaminergic system [68][69][70]. Besides, BPA negatively affects the organism through "nonsteroid pathways" affecting the activity of the genes participating in cell and tissue differentiation [60,71].
BPA can cause functional effects not only through the activation of receptors of steroid hormones but also through signal pathways, such as nuclear factor-κB, STAT3, phosphatidylinositol-3-kinase/AKT (PI3K/AKT), and mitogenactivated protein kinase (МАРK) [72]. This xenobiotic can also affect sodium, calcium, and chlorine ion canals, ionotropic glutamate receptors, and nicotinic and GABA receptors, changing the excitability and signal transmission in the neurons [73,74]. In addition, BPA exposure can increase the activity of the markers of oxidizing stress and reduce the activity of antioxidant markers. In this connection, it was supposed that the hypothyroid condition induced by BPA in the neonatal period can affect the thyroid gland-brain axis by the formation of free radicals, which in turn can disturb plasma membrane and cell components resulting in the delay of brain development [27].
As there are data indicating that estrogenic proteins affect the epigenetic status of the target genes (both at the level of DNA methylation and chromatin proteins) changing the level of their transcriptional activity [75,76], it can be supposed that BPA exposure has similar epigenetic mechanisms. A limited number of works have been published regarding research of epigenetic consequences of BPA exposure on the developing organism [77][78][79]. The obtained data confirmed that this xenobiotic can actually cause changes in the status of the expressed gene DNA methylation.

Epigenetic© effects© of© BPA© and© gene© expression
At the present time, studies have been conducted regarding the links between xenobiotic effect and changes in epigenome [80]. Three main epigenetic regulations of gene activity are known, which can be involved in the occurrence of pathologies connected to the EDC effect. These are DNA methylation, hydroxymethylation, different posttranslational modifications of histones (methylation, acetylation, phosphorylation, ubiquitination, sumoylation, and ADP-ribosylation of histones) and noncoding RNA. It should be underlined that these epigenetic mechanisms do not work in isolation from each other, but together in a complicated regulatory network. Different combinations of these modifications can significantly affect the chromatic status and result in transcriptional silencing and, on the contrary, increase the activity of transcription [81][82][83]. These covalent modifications do not cause classic genetic mutations, are rather liable, and are the most sensitive targets for the direct and indirect (metabolism products) effect of ecotoxicants on the epigenome of living organisms even in low doses. Disorders in any of the mentioned epigenetic regulatory mechanisms are connected to the elevated risk of disease [84]. The incorrect epigenetic regulation that occurs in primary germ cells ensures the mechanism of epigenetic inheritance of abnormal phenotypes in the number of generations, including the inheritance of predisposition to the number of the socially significant diseases [80].
The first studies of epigenetic changes caused by xenobiotics were conducted using the model of changing the color of the mouth fur Agouti viable yellow (Avy). It was demonstrated that the mother's diet with different content of sources of methyl groups (for example, folic acid) affects the degree of methylation of retrotransposon IAP located upstream of the Agouti gene, affects the level of gene transcription, and results in changes in the descendants' fur co lor [85,86]. Such effect was found in BPA exposure on pregnant females. It turned out that this ecotoxicant reduces IAP methylation of Avy and CapbiAP genes [87]. Besides, the hypomethylation of imprinted igf2r, Peg3, and H19 genes was observed in mice with increased concentration of BPA, resulting in increased mRNA level of these proteins, which in turn suppressed the maturation of oocytes due to the abnormal structure of the spindle during meiosis [88]. The authors Генетическая токсиколоГия came to the conclusion that BPA exposure in the embryonal period can change the cell processes and pathways of development through epigenetic mechanisms by changing the phenotype of the descendants. Exposure to low doses of BPA in the preimplantation period in laboratory mice can disturb DNA methylation during cleavege and at later stages of embryonal development. BPA caused the dose-dependent reduction of DNA methylation level in the 1-cell and 2-cell embryos and in blastocysts, which was accompanied by inhibition of cleavage. In germ cells on the 9 th day of development, i. e., during early organogenesis, a small increase in the level of genome-wide DNA methylation was observed. At the same time, on the 12 th day of embryonal development, both DNA hypomethylation and hypermethylation were detected depending on the body part (tissue type) and the germ weight [77][78][79]. The obtained data confirmed that preimplantation development is a highly sensitive period to BPA exposure. This was due to active reprogramming processes connected to the different pattern of changes in DNA methylation in the whole genome. Probably, this process involves different repeated DNA sequences that are also important for the regulation of gene activity, chromosome arrangement, and nuclear architecture.
It appeared that BPA can reduce the genome-wide DNA methylation in combination with the reduction of expression of DNA methyltransferase Dnmt1, which is stipulated by the disturbance of estrogenic mechanisms [89]. The reduction of the level of the genome-wide methylation of sequences LINE1 of DNA in sperm of males after BPA exposure was reverse proportional to the BPA level in urine, but it was not observed for methylation LINE1 in blood cells [90]. This indicated that epigenetic changes explained by DNA methylation are one of the possible mechanisms of the unfavorable effect of BPA on hematogenesis and fertility.
Futhermore, BPA exposure in the prenatal period caused disturbance in gene expression crucial for brain development, including the main (spiral-turn-spiral) transcriptional factors, which could be connected to epigenetic changes in CpG islands associated with the promoters of these genes [30]. For example, BPA exposure on developing neurons in the brain cortex of mice, rats, and humans reduced the level of mRNA of chlorine potassium transporter 2 gene (Kсс2). Probably, this was connected to the increased activity of bonding of methyl-CpG-bonding protein 2 (MECP2, MBD2) with "cytosine-phosphate-guanine coasts" of promoter of gene Kсс2 and due to the reduction of interaction with acetylated histone H3K9 surrounding the site of transcription initiation. Wherein it were observed sex differences: the BPA effect was stronger in females than in males [91]. The reduction of expression of DNA methyltransferases and hypomethylation of genes connected to lipid synthesis was also detected after BPA exposure in Hepa1-6 (mouth hepatoma) and BeWo (human trophoblasts, chorion carcinoma) cell lines [60,92,93].
BPA can result in DNA hypomethylation as well can cause the increase in the level of DNA methylation. Hypermethylated DNA was found in tissues of tail of the mouth successors, which was exposed to low BPA doses in the perinatal period, i. e., in successors of the second generation [94]. Experiments with laboratory animals demonstrated that this toxicant results in the stable expression of certain genes, including lactoferrin, epidermal growth factor, and proto-oncogenes (c-fos and c-jun), inhibiting the methylation process [53]. It was shown that BPA exposure in the neonatal period causes hypermethylation of the promoter of the receptor gene of estrogen in rat testicles [95]. Exposure to low doses of BPA on the cells of the primary culture of epithelium of the human breast results in the increase in methylation of CpG islands of DNA of the lysosomal-bound membrane protein 3 gene (lAMP3) and the suppression of transcription of this gene, which indicates the role of BPA in the enhancement of breast cancer development risk [96]. The epigenetic mechanism of regulation of BPA effects in breast carcinogenesis is also indicated by the elevated expression of trimethylated histone H3 as per lysine EZH2 after exposure to this xenoestrogen [97]. BPA can also increase the level of transcription of the cytokine gene of the family of tumor necrosis factor (tnFSF11 and RAnKl) and the family of genes coding secreted signal proteins (Wnt-4) required in embryogenesis regulate proliferation, participate in carcinogenesis of stem cells of breast, and play an important role in the metabolism of the bone tissue [98]. It was reported that BPA can increase the expression level of microРНK-146a, which is important in immune response [94]; therefore, the regulation of the epigenetic program and microRNA can become one of the areas of study and probably cancer therapy associated with BPA exposure.
It was demonstrated that BPA exposure during pregnancy (in mice and rats) can induce in the brain the successors of the first generation in puberty the sex-dependent, dose-dependent, and area-specific (in brain areas) changes in expression of genes coding receptors of estrogen (ERα, ERβ, and ERRγ). Together with the changes in estrogen-associated receptors, the dose-dependent changes of the mRNA level of DNA methyltransferases DnMt1 and DnMt3A genes were observed in juvenile cortex (male) and hypothalamus (female) as well as the level of methylation of ERα gene . Besides, such successors (male) demonstrated changes in the regulation of glucocorticoid, namely increased DNA methylation in Fkbp5 gene and reduction of the level of this protein in the hippocampus, which resulted in anomalies in behavior and response to stress of these animals [100]. BPA exposure in the prenatal and neonatal periods also disturbs the expression of methyl-CpG binding protein 2 in hypothalamic cells, which can be the reason for the disorders of the normal development of hypothalamus and its functions [99].
Thus, all these data indicate the interaction of two regulation systems, epigenetic and receptor (hormonal), and GEnEtiC toxiColoGy underline the importance of the study of BPA exposure effects on the health of humans and animals. Obviously, the methodological differences in research of BPA exposure on living organisms (studies in vivo and in vitro, different objects of study, different ways of exposure and experimental doses of BPA, and exposure of individual compounds and mixtures) explain an alternative hypothesis about the molecular mechanisms of this xenoestrogen effect. For example, mice and rats are different models for understanding the mechanisms of the human disease occurrence. Moreover, it should be noted that the same dose of BPA can result in DNA hypomethylation and hypermethylation or does not change it depending on the gender differences in response of the organism to the exposure, stage of development, cell differentiation, and tissue type.

conclusion
Data on epidemiological studies indicate potentially harmful chronic exposure of BPA on human and animal ontogenesis; therefore, BPA penetration into the organism (even in small doses) should be limited as much as possible, especially during pregnancy, taking into account its possible remote negative effects on health. It should be noted that some individuals have a low risk of pathology development under the effect of harmful environmental factors, whereas others are more sensitive to such effects. This is explained by genetic features, although individual epigenome differences should not be excluded at the current stage. Results of many studies indicate that the molecular mechanisms of xenobiotic effect go far beyond the limits of interaction with the DNA sequence. Apparently, additional research and development of new test systems are required for the assessment of the actual ratios of the dose and the effect and the mechanisms of ecotoxicant action in pathology development, as stated in the review. The development of preventive measures for the negative effect of xenobio tics requires research of the features of epigenomic/epigenetic modifications and DNA methylation first of all. Such studies shall be preferably conducted at different levels of arrangement -from molecular (DNA and chromatin), cell, and tissue to the entire organism -in experimental models in vivo and in vitro, taking into account different sensitivities to unfavorable BPA consequences.
One more very important aspect that should be given attention is the fact that epimutations caused by BPA in early embryogenesis result in changes in gene normal expression that can be kept in adults and transferred to the next generations through germ cells resulting in the intergeneration inheritance of abnormal phenotypes. Besides, it should be kept in mind that we actually are exposed to a mixture of pollutants; as a result, adaptive and synergetic effects take place, including BPA with other widespread compounds. Finally, it should be underlined that the approaches used in ecotoxicology based only on the analysis of nucleotide DNA sequence are currently insufficient for the complete explanation of the risks of diseases that can be modulated by nongenetic or extragenetic mechanisms.


extfieldsofstudy: Chemistry Biology

text:
Spire stimulates nucleation by Cappuccino and binds both ends of actin filaments.

The actin nucleators Spire and Cappuccino synergize to promote actin assembly, but the mechanism of their synergy is controversial. Together these proteins promote the formation of actin meshes, which are conserved structures that regulate the establishment of oocyte polarity. Direct interaction between Spire and Cappuccino is required for oogenesis and for in vitro synergistic actin assembly. This synergy is proposed to be driven by elongation and the formation of a ternary complex at filament barbed ends, or by nucleation and interaction at filament pointed ends. To mimic the geometry of Spire and Cappuccino in vivo, we immobilized Spire on beads and added Cappuccino and actin. Barbed ends, protected by Cappuccino, grow away from the beads while pointed ends are retained - as expected for nucleation-driven synergy. We found that Spire is sufficient to bind barbed-ends and retain pointed-ends of actin filaments near beads and we identified Spire's barbed-end binding domain. Loss of barbed-end binding increases nucleation by Spire and synergy with Cappuccino in bulk pyrene assays and on beads. Importantly, genetic rescue by the loss-of-function mutant indicates that barbed-end binding is not necessary for oogenesis. Thus increased nucleation is a critical element of synergy both in vitro and in vivo. [Media: see text] [Media: see text] [Media: see text] [Media: see text] [Media: see text] [Media: see text] [Media: see text] [Media: see text] [Media: see text].

the N-terminal Spir-KIND domain binds the C-terminal Capu-tail with ∼100 nM affinity (Figure 1; Quinlan et al., 2007;Pechlivanis et al., 2009;Vizcarra et al., 2011;Zeth et al., 2011). However, our understanding of the functional consequences of the interaction remains incomplete. Capu, like all formins, is a dimer. It can bind two copies of Spir and, thereby, accelerate actin assembly by Spir (Quinlan et al., 2007;Vizcarra et al., 2011). In contrast, Spir's KIND domain inhibits nucleation by Capu and competes with barbed ends for Capu binding, effectively inhibiting Capu's ability to accelerate actin assembly (Quinlan et al., 2007;Vizcarra et al., 2011). However, when a human Spire-1 construct, containing both the KIND domain and four tandem WH2 domains, is mixed with the C-terminal half of Fmn-2, actin assembly is greatly enhanced (Montaville et al., 2014). So-called ping-ponging-Spir and Fmn-2 alternately binding to the barbed end of filaments-is observed and was proposed to account for synergistic actin assembly (Montaville et al., 2014).
Spir is enriched at the cortex of the Drosophila oocyte and localizes to Rab11-positive vesicles in mouse oocytes, and the C-terminal mFYVE domain of mSpire-1 binds phospholipids (Quinlan et al., 2007;Pfender et al., 2011;Quinlan, 2013;Tittel et al., 2015). While Fmn-2 is also observed on Rab11-positive vesicles in mouse oocytes, GFP-Capu appears diffuse throughout the Drosophila oocyte (Dahlgaard et al., 2007;Schuh, 2011;Quinlan, 2013). In the mouse oocyte, Rab11-positive vesicles that contain mSpire, Fmn-2, and myosin V are at nodes of the actin mesh that fills the oocyte (Schuh, 2011). Mesh dynamics contribute to nucleus positioning and long distance transport of the Rab11-positive vesicles toward the cell cortex (Holubcová et al., 2013;Almonacid et al., 2015;Ahmed et al., 2018). A model to account for the long distance transport includes mSpire/Fmn-2-nucleated actin filaments growing with their barbed ends remaining near the vesicles, creating tracks that myosin V on neighboring vesicles can walk along, pulling vesicles together (Schuh, 2011). When ping-ponging was observed in vitro, Montaville et al. (2014) expanded on this model, proposing that barbed ends of filaments and/or prenuclei are recruited by vesiclebound mSpire and handed off to Fmn-2 to stimulate elongation.
To learn more about how Spir and Capu interact to build actin filaments and structures, we combined biochemistry and fly genetics. We developed a bead-based assay, in which we attached the N-terminal half of Spir (SpirNT) to beads and added the unconjugated C-terminal half of Capu (CapuCT), based on reported localization data (see Figure 1A for construct definitions). Through this work we found that beads decorated with SpirNT and CapuCT (CapuCT/SpirNT-beads) nucleate filaments that grow with their barbed ends away from beads, protected by CapuCT, opposite to the polarity expected based on the ping-pong model. We found that SpirNT alone on beads (SpirNT-beads) nucleates filaments oriented the same way. We also found that SpirNT-beads retain the pointed ends of filaments at the bead with a dwell time of ∼100 s while neighboring SpirNT-beads capture and cap the barbed ends of filaments for several hundreds of seconds. Next, we identified the domain necessary for Spir's high-affinity barbed-end binding. Surprisingly, barbed-end binding is not necessary for oogenesis and loss of barbed-end binding increases Spir/Capu synergy in vitro. These genetic and biochemical data indicate that ping-ponging at the barbed end is neither the dominant source of synergy nor necessary in vivo. Instead, enhanced nucleation by the Spir/Capu complex is critical.

Barbed ends project away from CapuCT/SpirNT-beads
To learn more about how Spir and Capu interact to build actin filaments and structures, we developed a bead-based assay. Spir is enriched at the cortex of the Drosophila oocyte, localizes to Rab11positive vesicles in mouse oocytes, and the C-terminal mSpire-mFYVE domain binds phospholipids (Quinlan et al., 2007;Quinlan, 2013;Pfender et al., 2011;Tittel et al., 2015). While Fmn2 is also observed on vesicles in the mouse oocyte, GFP-Capu appears diffuse throughout the Drosophila oocyte (Schuh, 2011;Quinlan, 2013). On the basis of these data, we immobilized SpirNT on beads and added unconjugated CapuCT. To do so, we biotinylated a Cterminal Avitag on SpirNT and mixed it with CapuCT. We then attached these complexes to streptavidin-coated beads under saturating conditions ( Figure 1B). We introduced the decorated beads to a sparsely biotinylated flow chamber to visualize them with total internal reflection fluorescence (TIRF) microscopy. Any unbound SpirNT or CapuCT was washed away. When we added actin (20% OG-actin) and profilin to the flow chamber, extensive and rapid polymerization followed (Figure 2A and Supplemental Video 1).
Two groups proposed that Spir/Capu-coated vesicles have filaments with barbed ends apposed to the vesicle and pointed ends growing away (Figure 1B;Schuh, 2011;Montaville et al., 2014). We tested this model by examining the orientation of filaments growing off of CapuCT/SpirNT-beads. By lowering the concentration of actin (600 nM), we decreased nucleation such that we could track individual filaments. Under these conditions, we observed filaments growing from but "retained" at the beads (Supplemental Video 2).
We carefully analyzed nine individual filaments. In every case, fiducial marks that do not move with respect to the bead, and increased fluorescence intensity at filament ends (i.e., ends brighter due to the addition of unbleached monomers [Kovar and Pollard, 2004]), away from the bead, indicated that barbed ends grow away from the bead surface ( Figure 2B). Filaments growing under these conditions elongated away from the bead at a rate of 50 ± 16 s −1 μM −1 (mean ± SD, n = 11), consistent with Capu-mediated elongation. We expected to see the opposite orientation of filaments with Capu directly conjugated to the beads. For unknown reasons, Capu was not functional when attached to beads through a number of different linkers. Instead, we conjugated the formin mDia1 to beads. In this case, fiducial marks were displaced at the rate of filament elongation (∼17 s −1 μM −1 ) and the addition of FIGURE 2: Barbed ends project away from CapuCT/SpirNT-beads. (A) Actin assembles off of CapuCT/SpirNT-beads. See also Supplemental Video 1. (B) A single filament is nucleated and retained by a bead. Kymograph (bottom) shows fiducial marks, such as bleached filament regions, which do not move with respect to the bead (cyan triangles). New, labeled actin is added away from the bead, indicating growth away from the bead. See Supplemental Video 2 for another example of retention by CapuCT/SpirNT-beads. (C) A single filament is nucleated and elongated by a bead coated with mDia1. Fiducial marks (highlighted with cyan triangles) in the kymograph are displaced from the bead at the rate of elongation, indicating growth at the bead. See also Supplemental Video 3.
unbleached monomers was no longer observed at the filament end away from the bead ( Figure 2C and Supplemental Video 3), consistent with barbed-end growth at the bead surface. Together these data demonstrate that CapuCT/SpirNT-beads nucleate filaments with barbed ends growing away from the bead and suggest that CapuCT separates from SpirNT to elongate the filament.

Spir-beads retain the pointed ends of nucleated actin filaments
A dense collection of filaments emanated radially from CapuCT/ SpirNT-beads. Because we concluded that CapuCT-mediated elongation proceeded away from the beads, the density of actin near the bead surface suggested that the pointed ends of filaments could be retained by Spir. We repeated the experiments with SpirNT-beads, without added CapuCT or profilin, and observed similar patterns ( Figure 3A and Supplemental Video 4). We found that nucleation by SpirNT-beads was suppressed enough to observe individual filaments by adding profilin ( Figure 3B). We again observed filaments retained at and growing away from the bead surface ( Figure 3C and Supplemental Video 5). Filaments growing away from SpirNT-beads elongated at a rate of 16.0 ± 0.3 s −1 μM −1 (mean ± SD, n = 12), as expected for free barbed ends. The filaments near the beads were released over time, demonstrating that they were not attached to the surface. The distribution of filament dwell times was well fit by a single exponential with an off rate of 0.007 ± 0.003 s −1 (n = 70; Figure 3D). We note that retention FIGURE 3: SpirNT-beads retain the pointed ends of nucleated actin filaments. (A) SpirNT-beads, in the absence of Capu and profilin, also potently polymerize actin. The highlighted region (gold) is magnified and pseudocolored (right) to emphasize the enrichment of actin filaments between beads. See also Supplemental Video 4. (B) Nucleation by SpirNT-beads is suppressed by profilin. When added at 1 μM (gold box), profilin permits the observation of several, individual filaments per bead. (C) SpirNT-beads also retain nucleated filaments. G, growth; B, bead. Fiducial marks (e.g., cyan triangle) in the kymograph are not displaced as the filament elongates, indicating growth away from the bead. See also Supplemental Video 5. (D) Dwell times of filament pointed ends on beads. Nucleated and retained filaments were tracked from SpirNT-beads in the presence of 1 μM actin and 1 μM profilin. All bead-associated filaments were counted unless obviously captured from solution or >1 μm in length when first visible. The data are well fit by a single exponential (k off = 0.007 ± 0.003 s −1 , n = 70 filaments; four independent experiments).
of filaments by SpirNT-beads was independent of profilin (Supplemental Figure S1G and Supplemental Video 6). Furthermore, neither filament end was observed to interact with beads coated with Spir-KIND and CapuCT (Supplemental Figure S1F), indicating that retention is specifically mediated by Spir-WH2 domains.
We did not assume that filaments were oriented with their barbed ends out when nucleated by SpirNT-beads because Spir has been reported to bind both the barbed and pointed ends of filaments (Quinlan et al., 2005;Bosch et al., 2007;Ito et al., 2011). mSpire1 caps barbed ends with high affinity and SpirNT binds pointed ends weakly (nM vs μM K d 's; Quinlan et al., 2005;Bosch et al., 2007;Montaville et al., 2014). We closely analyzed 10 of the 70 filaments included in the retention data set. All 10 filaments, growing from SpirNT-beads, displayed fiducial marks that did not move with respect to the bead and typically had brighter filament ends away from the beads, consistent with barbed ends being oriented away from the beads ( Figure 3C). Thus, we conclude that barbed ends grow away from SpirNT-beads and that SpirNT is sufficient to retain the pointed ends of actin filaments near beads for >100 s.

SpirNT-beads capture and cap the barbed ends of actin filaments
When we examined the dense actin networks emanating from beads, we also observed apparent connections between neighboring beads ( Figure 3A). This pattern suggested to us that the barbed ends of actin filaments could also be associated with beads. Under conditions where individual filaments could be tracked, we observed the "capture" of filaments ( Figure 4). The association of captured filaments with Spir-beads often outlasted the single-filament imaging window (10+ min), precluding measurement of an off rate and suggesting that the interaction is distinct from retention.
In addition to seeing filaments spanning two beads, we often observed capture of free filaments. We probed the orientation of captured filaments using two colors of actin. We initiated elongation in the presence of Cy3B-actin and stabilized these filaments with phalloidin after several minutes. Adding a limiting pool of OG-actin (600 nM) plus profilin to favor barbed-end growth, we observed that, whether the filament had both ends free (as shown) or one end retained by a bead, only the barbed ends of filaments were captured by beads ( Figure 4A and Supplemental Video 7). Filaments captured by beads did not grow measurably, suggesting that they are capped at their barbed ends ( Figure 4A). Consistent with this conclusion, the fluorescence intensity of the filament near the bead steadily declined after binding ( Figure 4B). Occasionally, we observed the release of a barbed end, following capture. In these cases, the filament resumed elongation at the same rate as before capture ( Figure 4A). As noted above, barbed ends do not interact with CapuCT/Spir-KIND-beads (Supplemental Figure S1F). These data indicate that the barbed ends are capped by Spir-WH2 domains, as opposed to being nonspecifically stuck to the beads or KIND domain.
FIGURE 4: SpirNT-beads capture and cap the barbed ends of actin filaments. (A) Nucleated filaments (green) grow only at their barbed ends (magenta). A free filament (dashed outline) diffuses and grows until it is captured by a SpirNT-bead (solid triangle). The filament resumes growth once released by the bead (open triangle). The kymograph shows that the growing, barbed end (magenta) is captured/capped by Spir and resumes growth after release. See also Supplemental Video 7. (B) Intensity of filament ends decrease after bead capture. Images of a filament immediately before attachment and 2 min later. The intensity of the first 1 μm of this filament is plotted vs. time, with 0 s set to the capture event.
(C) Two filaments (cyan triangles) nucleated by a SpirNT/CapuCT-bead (A) are retained for more than 5 min. Their barbed ends are captured (gold triangles) by other CapuCT/SpirNT-beads (B and C). Accelerated growth of these filaments (∼50 subunits/s*μM) in the presence of profilin and capping protein indicates that CapuCT is elongating and protecting their barbed ends. Filaments did not measurably grow, following capture, suggesting that CapuCT is displaced and that the barbed end is bound by SpirNT upon capture. See also Supplemental Video 8.
To further study the phenomenon of filament capture, we added capping protein and CapuCT to the experiment. Under these conditions we observed sustained and accelerated growth (51 ± 11 s −1 μM −1 , mean ± SD, n = 3) of retained filaments or absence of growth (n = 2), confirming that CapuCT can separate from SpirNT and protect growing filament barbed ends from capping protein ( Figure 4C). Growing filaments were still captured by beads, suggesting that SpirNT displaces CapuCT from the barbed end ( Figure  4C and Supplemental Video 8). Together these data demonstrate that, with or without CapuCT present, Spir-beads can nucleate and retain the pointed ends of actin filaments, while neighboring beads can capture and cap their barbed ends.

Spir's WH2-A binds filament barbed ends and reduces actin nucleation
Each of Spir's four WH2 domains contributes differently to nucleation and they bind actin monomers with a range of affinities (from ∼100 nM to 1.1 μM; Quinlan et al., 2005;Rasson et al., 2015). We reasoned that the N-terminal-most WH2 domain, WH2-A, binds barbed ends of filaments, based on our earlier observation that isolated WH2-A is the only Spir-WH2 domain that slows filament growth (Rasson et al., 2015). To test the contribution of WH2-A to barbed-end binding, we first established that wild-type SpirNT caps barbed ends in a seeded elongation assay, as has been shown for mSpire-1 (Bosch et al., 2007). In the presence of actin seeds, monomers, and profilin, SpirNT potently inhibited elongation ( Figure 5A). The dose dependence of inhibition indicates an apparent affinity of SpirNT for barbed ends of 20 ± 3 nM, similar to that reported for mSpire-1 ( Figure 5B). We then mutated three conserved, hydrophobic residues in WH2-A of SpirNT (SpirNT(A*); Figure 1A) to remove actin monomer binding by this domain alone. These mutations were previously shown to dramatically decrease, if not abolish, actin monomer binding (Quinlan et al., 2005). When added to the seeded elongation assay, SpirNT(A*) was essentially unable to inhibit elongation ( Figure 5, A and B). Thus, a functional WH2-A domain is necessary for high-affinity, barbed-end capping.
In standard pyrene-actin assays, we found that SpirNT(A*) assembled actin more potently than wild-type SpirNT ( Figure 5C and Supplemental Figure S1C). The plateau was consistently lower for wild type, suggesting that barbed-end binding by SpirNT is sufficient to change the critical concentration under these conditions. We also noted that the plateau of SpirNT(A*) pyrene traces did not decrease at high concentrations as is seen for wild-type SpirNT, suggesting that this mutant does not potently sequester actin monomers ( Figure  5C; Quinlan et al., 2005). We next attached biotinylated SpirNT(A*) to beads and added actin and profilin. Consistent with increased activity in pyrene assays, we observed faster accumulation of filaments from SpirNT(A*)-beads compared with wild-type SpirNTbeads ( Figure 5D and Supplemental Video 9). To compare nucleation rates on beads, we measured the integrated intensity of actin within a band 1.6 μm away from the beads. At this proximity, we are minimally sensitive to filament elongation and interpret the intensity as proportional to the number of filaments; that is, nucleation. We measured time courses and calculated the rates of increase in OG-actin signal. The difference in rates indicates that nucleation by SpirNT(A*) is approximately three times stronger than wild-type SpirNT in the presence of profilin ( Figure 6, D and E). We also measured the dwell times of filaments retained by SpirNT(A*)-beads. The off rate was not significantly different than that of wild type (k off(WT) = 0.007 ± 0.003 s −1 , k off(A*) = 0.011 ± 0.003 s −1 , Student's t test, p = 0.98; Figure 5E), indicating that WH2-A does not play an important role in filament retention. Notably, we rarely observed capture events by SpirNT(A*)-beads. In several cases, filaments appeared to be close enough to a neighboring bead for capture for tens of seconds, but sustained association of the barbed end with beads was rare. Taken together, these data are consistent with the requirement of functional WH2-A for high-affinity barbed-end binding and demonstrate that this domain is not critical for pointed-end binding.
Synergy between Spir and Capu does not require barbed-end binding Previously, synergistic actin assembly by mSpire-1 and Fmn-2 was described: the rate of actin assembly by mSpire-1 + Fmn-2 is greater than the sum of assembly rates of the individual nucleators (Montaville et al., 2014). We confirmed that SpirNT and CapuCT exhibit a similar synergistic effect in pyrene-actin assembly assays ( Figure 6A and Supplemental Figure S1D). In the presence of profilin, the assembly rate of SpirNT + CapuCT (based on the t 1/2 at ≥40 nM SpirNT) was 6× and 15× faster than CapuCT or SpirNT, respectively ( Figure 6B). As seen for the mammalian paralogues, direct interaction was required for synergy: SpirNT(Y232K), a mutant that does not bind Capu, does not synergize with CapuCT (Supplemental Figure S1E; Vizcarra et al., 2011;Montaville et al., 2014). Montaville et al. (2014) concluded that actin assembly was enhanced by mSpire-1 and Fmn-2 alternately binding the barbed end, which they dubbed the ping-pong mechanism. Given that SpirNT(A*) does not bind to or capture barbed ends, we assume that ping-ponging is not possible with this mutant. However, when we tested SpirNT(A*) in the pyrene synergy assay, we found approximately three times enhanced synergy (based on the t 1/2 at ≥40 nM SpirNT), as opposed to loss of activity ( Figure 6, A and B, and Supplemental Figure S1D). The dose dependence of synergy is indistinguishable for SpirNT and SpirNT(A*) ( Figure 6B). We speculate that dose dependence is a function of Spir-KIND/Capu-tail binding, which should not be affected in SpirNT(A*). These data lead us to propose that enhanced nucleation, not just elongation, drives synergy when SpirNT and CapuCT interact.
To directly test whether nucleation is increased by SpirNT and CapuCT collaborating, we returned to the bead assay. When CapuCT/SpirNT-beads were mixed with actin and profilin, we observed potent nucleation ( Figure 6C). Consistent with nucleation being a significant element of Spir/Capu synergy, the rate of filament formation was 6.5× greater than in the absence of Capu ( Figure 6, C-E). Nucleation of profilin-actin by CapuCT/SpirNT(A*)beads was even stronger than wild type (4×) and the synergy more pronounced (9.5× vs. SpirNT(A*)-beads in the absence of Capu) with the actin signal saturating at the measured 1.6 μm radius within ∼2 min (Figure 6, C and D). Thus, nucleation is potently enhanced when SpirNT and CapuCT are combined and loss of barbed-end binding by SpirNT leads to even stronger nucleation in the presence and absence of CapuCT. These data indicate that enhanced nucleation is a major source of synergy in vitro.

Barbed-end binding is not necessary for oogenesis
Finally, we asked whether barbed-end binding is necessary for Drosophila oogenesis. To do so, we used full-length Spir (CG10076, splice variant "PB") with a C-terminal monomeric enhanced green fluorescent protein (GFP) tag in a genetic rescue assay. We mutated three conserved residues of WH2-A as done in vitro. We previously demonstrated that expression of wild-type Spir-GFP, driven by germline-specific nanos-Gal4-vp16, is sufficient to rescue fertility in flies that lack endogenous Spir (Table 1; Quinlan, 2013). We found that fertility was largely rescued when Spir(A*)-GFP was expressed (hatch rate 40% compared with 59% for wild type; Table 1). Consistently, the mesh was present in 11 of 20 oocytes and streaming was properly regulated in about half of the oocytes observed ( Figure 7, D and D′). Thus, barbed-end binding is not necessary for mesh formation or oogenesis, though the fertility decrease, relative to wild-type rescue, suggests that it may contribute under normal conditions. has no effect (purple). (B) Dose-dependent elongation rates are plotted for three experiments. The data are fit with a quadratic equation, indicating that SpirNT binds the barbed end with a K d of 20 ± 3 nM. The line is a fit to all data points, and the K d is the average of three independent experiments. Inhibition by SpirNT(A*) is negligible. (C) SpirNT(A*) assembles actin more potently than wild-type SpirNT in a pyrene-actin assembly assay. Plateaus that are independent of Spire concentration suggest that SpirNT(A*) does not sequester actin like SpirNT (compare solid lines). (D) SpirNT(A*)-beads nucleate more potently than wild type. See also Supplemental Video 9. (E) Quantification of filament pointed-end dwell times on beads (k off = 0.011 ± 0.003 s −1 , n = 33 filaments; three independent experiments). The off rate is not significantly different from wild type (data from Figure 3D).  Figure 5D. Fluorescence around beads is expressed as an average percentage of initial values over time (n ≥ 4 beads, n ≥ 2 independent experiments per condition). See Materials and Methods for the description of data transformation. Data plotted were obtained using 1-μm-diameter beads, 1 μM actin, and 1 μM profilin, with or without CapuCT (solid or dashed lines, respectively). S, SpirNT-beads; P, profilin; C, CapuCT. SpirNT-bead data are plotted in blue and SpirNT(A*)bead data in magenta. (E) Rates of fluorescence increase, from D. Mean ± SEM are indicated for each condition. The initial 50 s of images (t = ∼30-80 s) were used to determine rates. A Student's t test was performed on each pair of conditions indicated (black brackets). ***, p < 0.0001. The difference in activity by SpirNT(A*/WT)-beads (without Capu) was not statistically significant (unequal variance; p = 0.13).

WH2 domains are necessary for Drosophila oogenesis
The Spir-KIND domain inhibits CapuCT in pyrene-actin assays (Quinlan et al., 2007;Vizcarra et al., 2011). However, genetics indicate that Spir and Capu both play positive roles in oocyte mesh formation (Manseau and Schupbach, 1989;Dahlgaard et al., 2007;Quinlan, 2013). Further, the N-terminal half of Spir (containing only the KIND domain and WH2s) is sufficient to build the mesh (Dahlgaard et al., 2007;Quinlan, 2013). Combined, these data suggest that Spir's WH2 domains contribute to in vivo function. We decided to test this hypothesis directly, in part because Spir(A*)-GFP rescues. In each of Spir's four WH2 domains, we mutated three conserved, hydrophobic residues that contact actin ( Figure 1A). These mutations were previously shown to dramatically decrease, if not abolish, actin monomer binding (Quinlan et al., 2005). In vitro, SpirNT with all of these mutations (SpirNT(A*B*C*D*)) does not accelerate actin assembly (Supplemental Figure S1A; Quinlan et al., 2005). nanos-Gal4-vp16-driven expression of Spir(A*B*C*D*)-GFP failed to rescue fertility (hatch rate <2%; Table 1). In oocytes expressing Spir(A*B*C*D*)-GFP, the actin mesh was absent and streaming was premature, consistent with loss of actin assembly activity and loss of fertility (Figure 7, A-C). Thus, we conclude that actin binding by at least one of Spir's WH2 domains is necessary for actin assembly and oogenesis.

Spir binds both ends of actin filaments
The bead assay facilitated observation of several steps in actin assembly by Spir and Capu. We found that filaments nucleated on CapuCT/SpirNT-beads grow with their barbed ends away from the bead with enhanced rates, accelerated by CapuCT. Thus, SpirNT and CapuCT separate after nucleation, consistent with genetics results (Quinlan, 2013). We also found that CapuCT-bound barbed ends would subsequently stop growing if they encountered CapuCT/SpirNT-beads, suggesting that the barbed end was passed from CapuCT to SpirNT. To our surprise, we found that SpirNT-beads were sufficient to bind both ends of actin filaments. There are conflicting reports regarding filament-end binding for two classes of tandem-WH2 nucleators: Spir and VopF/L (Liverman et al., 2007;Yu et al., 2011;Pernier et al., 2013Pernier et al., , 2016Burke et al., 2017). The N-termini of WH2 domains bind actin monomers between subdomains 1 and 3 of actin-the surface exposed at the barbed ends of filaments (Hertzog et al., 2004;Chereau et al., 2005). It was, therefore, reasonable to expect tandem-WH2 nucleators to associate with filament barbed ends and surprising when both VopL and Spir were reported to bind filament pointed ends (Quinlan et al., 2005;Namgoong et al., 2011;Yu et al., 2011). Spir nucleates filaments with free barbed ends and inhibits depolymerization of gelsolin-capped filaments, albeit weakly (Quinlan et al., 2005). Spir also caps barbed ends of growing filaments but with nanomolar affinity (Bosch et al., 2007;Montaville et al., 2014). Here we present evidence that these apparently conflicting data are all correct. This is not unprecedented. Namgoong et al. (2011) and Burke et al. (2017) reported that VopL/F can interact with both ends of actin filaments, depending on the conditions. Earlier, we did not observe barbed-end binding by Spir when assayed by inhibition of depolymerization (Quinlan et al., 2005). We now report high-affinity barbed-end binding in inhibition of elongation assays. Possibly, Spir binds barbed ends only in the presence of actin monomers as was described for N-Wasp (Co et al., 2007). This contrasts with VopL/F that primarily bind barbed ends of preformed filaments in the absence of free monomer (Burke et al., 2017). The first of Spir's tandem WH2 domains, WH2-A, is necessary to cap barbed ends. It is curious that removing function of this WH2 domain increases the activity of SpirNT. In earlier work, we did not observe a significant difference in actin assembly, with or without WH2-A (Quinlan et al., 2005). The original data were acquired with higher concentrations of both SpirNT and actin, which may have masked the difference. In addition, the data were acquired with a slightly longer construct: 1-520 versus 1-490. One possible explanation for enhanced actin nucleation by SpirNT(A*) is that loss of capping leads to fewer so-called SA 4 complexes (Spir bound to four actin monomers). We and others observed SA 4 complexes when Spir is mixed with actin under polymerizing conditions (Quinlan et al., 2005;Bosch et al., 2007). We originally interpreted these structures as prenuclei. In contrast, Bosch et al. (2007) proposed that they are stable structures that sequester actin monomers. Whether there are two paths (i.e., nucleation vs. SA 4 complex), or one (nucleation, with formation of the nucleus from the SA 4 complex being a rate-limiting step), removing capping would likely destabilize the SA 4 structure and could favor nucleation.
How do tandem-WH2-domain nucleators bind filament pointed ends? Single-molecule observations showed that VopL/F remains associated with the pointed ends of filaments it nucleates for ∼100 s (Burke et al., 2017). In the absence of actin monomers, VopL/F binds pointed ends for shorter times (∼25 s). Higher affinity bind-ing is likely due to the contribution of the VopL/F C-terminal domain, which dimerizes and binds the pointed end. In the case of Spir, weak inhibition of depolymerization may be mediated by a linker or a domain straddling the WH2 domains. Side binding by the C-terminal half of tandem WH2 domains may enhance relatively weak pointed-end binding of both Spir and VopL/F. Side binding by Spir is consistent with the fact that it can sever filaments (Bosch et al., 2007;Chen et al., 2012). The resolution of our bead assay is not sufficient to distinguish between direct binding to the pointed end and binding to the filament side near the pointed end. Additionally, pointed-end elongation is difficult to detect with low actin concentrations and is essentially nonexistent in the presence of profilin. Thus, we speculate that the ability to retain the pointed ends of filaments nucleated by Spir on beads reflects enhanced binding, perhaps to both filament sides and the pointed end, due to clustering on beads ( Figure 8A). Multiple cases of emergent behavior of clustered WH2 domains have been reported. Spir nucleates more potently when clustered on gold particles (Ito et al., 2011). VopL retains the pointed end of ∼10% of nucleated filaments for several minutes (compared with seconds for ∼80% of filaments) and accelerates barbed-end elongation of ∼8% of filaments when clustered on Qdots (Namgoong et al., 2011); and Ena/VASP are potent elongation factors when clustered on beads (Breitsprecher et al., 2008;Winkelman et al., 2014). In sum, WH2 domains can associate with both ends of actin filaments and they are likely to function as part of a larger context that dictates their activity (Dominguez, 2016).

How do Spir and Capu synergize?
Synergy of actin assembly can be explained by enhanced nucleation and/or elongation. In the cases of Bud6/Bni1 and APC/mDia1, nucleation is enhanced (Graziano et al., 2011;Breitsprecher et al., 2012). We propose that increased nucleation is also the dominant source of synergy for Spir/Capu. The ping-pong model suggests that Spir and Capu (Fmn2) enhance elongation by dynamic exchange at the barbed ends of filaments. A key element of the pingpong model is that Fmn2 binds barbed ends with a slow on-rate. Thus, by recruiting Fmn2 to barbed ends, Spir increases the fraction of time that elongation is enhanced by the formin. We found that synergistic assembly of profilin-actin is actually improved in the absence of Spir binding to barbed ends (CapuCT+SpirNT(A*)). We also directly observed increased nucleation in bead assays: CapuCT/SpirNT-beads nucleate 6× and 10× more potently than SpirNT-beads, with or without a functional WH2-A, respectively ( Figures 5D and 6, C-E). A full comparison (i.e., nucleation by SpirNT-beads and CapuCT-beads, vs. CapuCT/SpirNT-beads) was precluded by the inactivity of CapuCT-beads, regardless of the coupling method. Finally, we see a fourfold increase in nucleation by CapuCT/SpirNT(A*)-beads compared with wild-type CapuCT/ SpirNT-beads, similar to the approximately three times decrease in t 1/2 in bulk pyrene assays ( Figure 6B). Thus, we conclude that nucleation is the major source of enhanced actin assembly mediated by Spir/Capu synergy.
Ping-ponging may still take place and contribute to mesh assembly. We observe transfer of Capu-bound barbed ends to Spir ( Figure  4C and Supplemental Video 8), consistent with one half of pingpong. We do not see transfer in the other direction, but our beadbased assay is not well suited to studying this process, as we do not have excess Spir or Capu in solution. (The original observation of ping-pong was made with all proteins in solution [Montaville et al., 2014].) Importantly, the intermediate phenotype we observe in flies expressing Spir(A*)-GFP (Figure 7, D and D′, and Table 1) could indicate that, while not necessary, ping-ponging may enhance mesh assembly in the oocyte. Perhaps Spir helps Capu fall off of filaments when they reach their target. Additionally, Spir could pass the filament to another protein that anchors it at the vesicle even more stably, influencing the dynamics of the mesh. It is also possible that capping serves an important role in vivo, independent of the processes we are studying here.
The classical nucleation promoting factors (NPFs), N-Wasp and Scar, bind actin monomers and the Arp2/3 complex to stimulate nucleation by the Arp2/3 complex (Machesky et al., 1999;Welch and Mullins, 2002). Similarly, the yeast NPF, Bud6, binds an actin monomer and the formin Bni1 to stimulate nucleation by Bni1 (Graziano et al., 2011;Tu et al., 2012;Park et al., 2015). In each of these cases, the NPF has negligible independent activity. In contrast, APC and Spir can nucleate alone, as well as synergize with a formin (Quinlan et al., 2005;Moseley et al., 2007;Okada et al., 2010;Montaville et al., 2014). We note that Spir's nucleation activity in the presence of profilin correlates with synergy: in the absence of nucleation by SpirNT(A*B*C*D*), no synergy is observed; when nucleation is augmented (SpirNT(A*) nucleation is approximately three times stronger than SpirNT), synergy with Capu is also enhanced over wild type (approximately four times). These are only two data points, but they suggest that Spir's nucleation contributes to synergy, as opposed to Spir acting as a passive actinbinding protein, or NPF. In fact, if monomer delivery were Spir's main role, one might expect SpirNT(A*) to be a weaker Capu activator because it has fewer functional WH2 domains. Thus, we propose that Spir and Capu collaborate by a mechanism similar to that of APC/mDia1; Spir nucleates seeds that are elongated by Capu ( Figure 8B).

In vivo implications
The filament orientation we observe (barbed ends away from the bead) reflects the fact that Spir is attached to the beads and Capu is bound to Spir in our assays. As we and others have shown, when a formin is bound to a bead, filaments grow with their barbed ends at the bead surface. Both mSpire and Fmn-2 are enriched on Rab-11 vesicles. We speculate that Fmn2 is affiliated by binding mSpire as opposed to an independent association with the vesicle. We base this on the finding that mSpire binds directly to membranes and the fact that, in flies, GFP-Capu is diffuse throughout the oocyte (Quinlan, 2013;Tittel et al., 2015). If we are correct, then the geometry in our assays mimics the in vivo geometry and indicates that the actin filaments are oriented opposite to what was originally proposed by others (Schuh, 2011;Montaville et al., 2014). Long distance transport is still possible with this orientation of filaments ( Figure 8C). Instead of myosin V capturing filaments from a neighboring vesicle, Spir or another protein could capture a growing filament, and myosin V, on the vesicle from which the filament originated, could walk along the filament pulling the vesicle toward its neighbor ( Figure 8C′). In this case, pointed-end attachment would not have to be as long lasting as barbed-end capture. These conditions are surprisingly well satisfied in our simplified system.
Increased Capu expression is sufficient to build a mesh in the absence of Spir or in the absence of direct interaction with Spir (Dahlgaard et al., 2007;Quinlan, 2013). In these cases, the mesh is not as dense or as long lived. No mesh is built when exogenous Spir expression is driven in the absence of Capu (Dahlgaard et al., 2007). Thus, we propose that Spir's primary role in vivo is as an NPF, enhancing Capu's actin assembly activity. NPFs offer an additional level of control in the cell and may relieve the nucleation-dampening effect of profilin on formins like Capu. In the case of the fly oocyte, we also note that the cytoplasm is enriched with microtubules that potently inhibit Capu by binding to its tail, where the Spir-KIND domain also binds (Roth-Johnson et al., 2014). Thus, Spir is well suited to simultaneously protect Capu from inhibitory factors and amplify its activity. The fact that Spir(A*)-GFP expression largely rescues loss of Spir indicates that barbedend binding by Spir is not necessary for oogenesis. We note, however, that actin nucleation by SpirNT(A*) alone and with Capu is enhanced, which could compensate for loss of ping-ponging or other barbed-end binding roles of Spir.

DNA constructs
CapuCT (aa 467-1059) constructs were expressed from a modified pET15b vector with an N-terminal hexahistidine tag (Vizcarra et al., 2011). All other proteins were expressed from a modified pET20b(+) vector with no affinity tag. A native polyhistidine region within the Spir-KIND domain is sufficient for binding of these constructs to TALON resin (Clontech).
Gibson cloning was employed for the scar-free introduction of Avidity's 45 base pair Avitag (translated sequence: GLN-DIFEAQKIEWHE) to all proteins requiring biotinylation for bead conjugation. This tag was introduced to the C-terminus of all constructs.
Expression was induced in Rosetta (DE3) cells (EMD Biosciences). Bacteria were grown in Terrific Broth medium supplemented with 100 mg/l ampicillin and 32 mg/l chloramphenicol. Cells were grown to an optical density of 0.6 at 37°C, cooled to 18°C for 1 h, induced with 250 μM isopropyl β-d-1-thiogalactopyranoside (IPTG), and shaken for 18 h at 18°C. Bacteria were harvested by centrifugation. Pellets were washed once with ice-cold phosphate-buffered saline (PBS), flash-frozen in liquid nitrogen, and stored at −80°C.
All purification steps were carried out at 4°C or on ice. Thawed cells were diluted at least twofold with lysis buffer (50 mM sodium phosphate, pH 8.0, 1 mM β-ME, 300 mM NaCl) supplemented with 1 mM phenylmethylsulfonyl fluoride and 2 μg∕ml DNaseI, and then lysed by microfluidizing, two to three times. Cell debris was removed by centrifugation at 20,000 × g for 20 min at 4°C. Clarified lysates were then rocked with TALON resin for 1 h at 4°C (4 ml slurry for every 1 l culture pellet). The TALON resin was washed with 20 column volumes of lysis buffer, followed by washing with 20 column volumes of wash buffer (lysis buffer, at pH 7.0). Proteins were eluted with elution buffer (wash buffer, plus 200 mM imidazole), until little or no protein remained on the column, as determined by a Coomassie-stained dot-blot of the eluates.
TALON eluates were pooled and dialyzed twice for 2 h and once overnight against 1 l volumes of 10 mM Tris, 1 mM dithiothreitol (DTT), pH 8.0; or 5 ml of the most concentrated eluates was buffer exchanged into the same, using a PD-10 desalting column (GE Life Sciences). Protein was loaded onto a MonoQ anion exchange column (GE Life Sciences) and eluted using a gradient of 50-500 mM KCl over 60 column volumes for Spir-KIND, 50-250 mM KCl over 100 column volumes for CapuCT, or 0-500 mM KCl over 60 column volumes for SpirNT. Pooled fractions from the MonoQ column were again dialyzed or buffer exchanged as described above. Unless tagged with an Avitag, 50% glycerol was added to the overnight dialysis step (1:1 glycerol:buffer). The pro-tein was flash-frozen in liquid nitrogen in 10-50 μl aliquots and stored at −80°C.
If tagged with an Avitag, 2 ml of the protein was added to 223 μl Biomix B and 5 μl (5 μg) BirA (both from Avidity) and rocked overnight at 4°C. The reaction was again loaded onto a MonoQ anion exchange column and eluted as described above. Pooled fractions from the MonoQ column were dialyzed or buffer exchanged as described above, with 50% glycerol added to the overnight dialysis step and flash-frozen in liquid nitrogen.
Purified, SNAP-tagged mDia1 protein was generously provided by the Kovar lab (University of Chicago).
Actin seeds were prepared by polymerizing 5 μM actin at 25°C for 1 h in KMEH. The filaments were dispensed in 5 μl aliquots and allowed to reequilibrate for 2-3 h at 25°C. SpirNT was incubated with filaments for 3 min at 25°C. During this incubation time, monomeric actin was converted to Mg-actin. Using a cut pipette tip to prevent shearing, polymerization buffer was added to Mg-actin and then mixed with seeds plus SpirNT. The slope of the pyrene fluorescence trace between 200 and 500 s was considered the elongation rate.

SpirNT-bead preparation
Streptavidin-coated microspheres were either colorless or Flash Red fluorescent, with mean diameters of ∼100 nm or ∼1 μm (Bangs Laboratories), respectively. Spheres were washed twice with ∼20 volumes of KMEH, resuspended in 1 volume of KMEH + 1 mg/ml bovine serum albumin, and added in 10 μl aliquots to 40 μl protein mixtures in KMEH. The protein mixture always included a saturating concentration of SpirNT-biotin (400 nM). When included, 200 nM (dimeric) CapuCT was preincubated with the SpirNT-biotin on ice for 10 min to allow the formation of SpirNT-CapuCT complexes. Protein mixtures were incubated with spheres for 10 min on ice and then spun for 10 min at 10,000× rpm and 4°C. Buffer and excess proteins (i.e., unbound SpirNT-biotin and CapuCT) were aspirated and pellets were gently resuspended in 30 μl KMEH. Pellets were briefly sonicated (∼5 s) if clumped and not well dispersed when visualized by TIRF microscopy.

TIRF microscopy assays
Coverslips were prepared and functionalized with polyethylene glycol (final surface composition, 97% methoxy-PEG and 3% biotin-PEG; JenKem Technology, Allen, TX) as previously described (Bor et al., 2012). Biotinylated coverslips were stored in a sealed container at 4°C for up to 2 mo before use. Flow cells with volumes of ∼10-15 μl were assembled using thin strips of double-stick tape.
Time zero was defined as the moment the final 25 μl mix, including actin, was entirely added. Polymerization was visualized immediately (typically, t = ∼30 s) on a DMI6000B TIRF microscope (Leica, Wetzlar, Germany), controlled by LAS X (Leica software). Images were acquired with a DU897 EMCCD camera (Andor) and 100×/1.47 HCX PL APO objective (Leica) at ∼25°C. All analyses were performed on raw data in FIJI. The brightness and contrast of figure images were minimally adjusted for clarity of image features. Filament lengths, elongation rates, and kymographs were analyzed/ prepared with JFilament incorporated in FIJI (Smith et al., 2010). Plots were made and statistical analyses conducted in Kaleidagraph. Filament dwell times were obtained by manually tracking individual, retained filaments until clearly released from a bead. Some filaments did not become clearly visible in the TIRF plane until some growth had already occurred. In these cases, time was added to the measured retention period, proportional to the initial length of the filament and the average filament elongation rate observed in the experiment.
For two-color experiments, 1 μM Cy3B-labeled actin was introduced to the functionalized, bead-bound flow cell. Following this (step 5 of the aforementioned protocol), and after allowing beads to briefly (1-2 mins) polymerize the Cy3B-actin, the following components were flowed in: 6) TIRF buffer supplemented with 1 μM phalloidin, with 2 min incubation; 7) TIRF buffer supplemented with GCC mix, 600 nM 20% labeled OG-actin, and 300 nM profilin. Polymerization was visualized as described above.
For the plots generated in Figure 6, D and E, circles of 10 and 11 pixel (px) radii (1.6 and 1.76 μm, respectively) were drawn around the centroids of each bead. Actin fluorescence was measured within these circles over time. The integrated density measurements of the 10 px circles were subtracted from those of their concentric, 11 px circles, to measure 1-px-wide (160 nm) bands of actin fluorescence, 1.6 μm away from beads. A script that performs these operations in FIJI is available upon request.
Fluorescence measurements in Figure 6, D and E, are expressed as a fraction of their initial values (i.e., a 10% increase = 1.1). Discrepancies in initial fluorescence values for different bead types were observed, in part, due to real differences in assembly rates, because actin polymerization could not be observed instantly. To scale these differences in assembly before image collection, the average starting values of each bead type were compared. The larger ratio of these two values was used as a multiplier for the more active bead type. For example, if measurements of SpirNT(A*)-and SpirNTbeads were 110% their initial values, these quantities would each be plotted as "1.1." However, if the average, initial values of SpirNT(A*)beads were twice those of SpirNT-beads, the quantities would be plotted as "2.2" and "1.1," respectively, reflecting the twofold greater assembly by SpirNT(A*)-beads before image collection.

Drosophila stocks
The following stocks were obtained from the Bloomington Drosophila Stock Center: spir 1 and capu 1 (Manseau and Schüpbach, 1989); Df(2L)Exel 6046 (Exelixis); and nos-GAL4-vp16 (Van Doren et al., 1998). Mutant SpirB-GFP transgenes were generated by inserting the coding region of spir (CG10076-RB) with point mutations created by QuikChange mutagenesis, between the KpnI and SpeI sites of pTIGER (Ferguson et al., 2012) with mEGFP inserted between the BamHI and XbaI sites. pTIGER plasmids were integrated at the attP2 landing site by BestGene.

Fertility assays
Approximately 100 test females were crossed to 40 wild-type males and kept on apple plates for two nights at 25°C. Flies were precleared on a fresh plate with yeast paste for at least 1.5 h; the plate was changed and eggs laid over the next 3 h were collected. Typically, 100 eggs were laid in this time period. Eggs were transferred to a fresh plate and stored at 25°C. The number of eggs that hatched after 24 h was recorded. Each trial was repeated at least three times with independent crosses.

Fly oocyte microscopy and staining
The visualization of cytoplasmic flows and the actin mesh were performed on a Leica SPE I inverted confocal microscope. Flies were kept at 25°C and fed yeast paste for ∼24 h before an experiment. Flows were visualized by imaging autofluorescent yolk granules of egg chambers teased apart in Halocarbon oil 700. The actin mesh was stained as described by Dahlgaard et al. (2007) with modifications. Briefly, ovaries were dissected, teased apart, and fixed in 10% paraformaldehyde/PBS (pH 7.4) for a total of less than 20 min. Samples were stained with 1 μM Alexa Fluor 488-phalloidin diluted in 0.3% Triton X-100/PBS for 25 min at room temperature. Samples were then washed extensively and mounted in ProLong Gold with 4′,6-diamidino-2-phenylindole (DAPI). Images were recorded within 24 h of staining because phalloidin staining quality degraded over time, as has been reported (Dahlgaard et al., 2007).