Cytochrome P450 Aromatase
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Aromatase is the cytochrome P450 enzyme converting androgens into estrogen in the last phase of steroidogenesis. As estrogens are crucial in reproductive biology, aromatase is found in vertebrates and the invertebrates of the genus Branchiostoma, where it carries out the aromatization reaction of the A-ring of androgens that produces estrogens.

cytochrome P450 aromatase estrogens molecular evolution structural alignment substrate recognition sites conservation

1. Introduction

Aromatase is the enzyme that converts androgens into estrogens through a three-step reaction that allows the aromatization of the A-ring of the steroid molecule [1][2]. The enzyme belongs to the cytochrome P450 (P450s) superfamily that comprises thousands of enzymes involved in the metabolism of endogenous and exogenous substrates [3][4][5]. The origin of such a large number of enzymes is still controversial, even though the presence of a common ancient precursor, CYP51 (lanosterol 14alpha-demethylase), for both prokaryotes and eukaryotes has been hypothesized [6].

The P450 superfamily is composed of two groups of enzymes. Depending on their substrate recognition abilities, one group comprises P450s that catalyze specific reactions on specific endogenous substrates; a second group includes enzymes that have evolved towards broad substrate selectivity, usually employed for xenobiotic metabolism, as in the case of mammalian liver proteins. While for the second group, it can be hypothesized that evolution has widened their substrate selectivity, for the first one, it is not clear how molecular evolution has worked.

Aromatase belongs to the first group as it carries out the conversion of androgens into estrogens across different classes of living organisms. From an evolutionary point of view, its gene and activity have been found in invertebrates of the genus Branchiostoma, belonging to cephalochordates [7]. Indeed, aromatase, together with other P450 enzymes involved in steroidogenesis, have been found in the gonads of the invertebrate Branchiostoma belcheri, which is considered to be evolutionarily closer to vertebrates than other invertebrates [8][9].

The enzyme is present in all vertebrates as the product of expression of a single gene, with some exceptions represented by pigs and teleosts, where duplication events have produced three and two isoforms, respectively [10][11][12][13]. Furthermore, the protein is expressed in different tissues in vertebrates, where it plays an essential role in reproductive biology as estrogens are responsible for ovarian differentiation, development of the reproductive system, sex differentiation, and reproduction [14]. Moreover, a critical role of estrogens has also been demonstrated in brain, bone, skin, fat, and cardiovascular tissues [15][16][17][18][19][20]. In humans, tissue-specific regulation of aromatase gene expression is allowed by the presence of eleven promoters and alternative first exons [21]. However, a wide tissue distribution of the aromatase protein and a complex regulatory region in its gene is already present in fishes [22].

Vertebrates have been used as models to understand the roles of aromatase and estrogens in the different tissues where it is expressed. For example, in birds and mammals, it has been demonstrated that in the brain, there is a rapid modulation of aromatase activity through phosphorylation and that estrogens can be considered neurotransmitters [23]. Moreover, estrogens are involved in different processes, such as neurogenesis, neuroprotection, and cognition [22][24].

In reptiles and amphibians, temperature regulates aromatase expression and is responsible for temperature-dependent sex determination [25][26][27]. In some hermaphrodite fishes, sex changes occur in response to environmental cues related to social interactions, and aromatase is involved in the remodeling of the gonads during this process [28][29]. Due to the phenotypic effects as a consequence of androgen/estrogen unbalance, amphibians and fishes are widely used as model organisms to understand the possible effect of many compounds that also target human aromatase [30][31], known as endocrine-disrupting chemicals (EDCs) [32][33].

Among fishes, teleosts represent the only case where two isoforms are present (CYP19A1 and CYP19B1), and they are preferentially expressed in the gonads and brain, respectively. Interestingly, these isoforms have also been reported to have different catalytic activity in comparison to the human enzyme [34][35], indicating that functional differences can be present. Thus, it is interesting to understand the phylogenetic origins of these differences.

2. Multiple Sequence Alignment

2.1. Structural Conservation

In order to identify the most conserved structural elements in aromatase, 365 sequences, ranging from invertebrates to mammals, were used for multiple sequence alignment. Out of the 365 sequences aligned, 66 were from mammals, 8 from birds, 12 from reptiles, 18 from amphibians, 259 from fishes, and 2 from the invertebrates of the genus Branchiostoma.

For all the analyses performed in this work, the residue numbers refer to the sequence of human aromatase (CYP19A1, Uniprot ID P11511).

When the positions of the most conserved regions were analysed in the crystal structure of the human enzyme, they resulted as part of helix A (65–78), the β-sheet formed by strands β1 (83–88) and β2 (93–97), helix E (187–205), part of helix F (221–224), the central part of helix I (residues 302–318 in human aromatase), helix K (354–366), the K-β3 loop and the β3 strand (368–376), the β6 strand (393–396), and helix L and part of the L-K’’ loop (427–448) (Figure 1). Helices C, D, F, and H carry conserved amino acids oriented toward the core of the protein and nonconserved amino acids exposed to the solvent. Thus, the conserved structural core in cytochrome P450 is formed by a four-helix bundle formed by helices D, E, I, and L that is conserved among aromatase sequences; an exception is made for the residues of helix D, exposed to the solvent (Figure 1) [36]. Helix G is not conserved, whereas the F-G loop and the first part of helix F, known to be important for opening the access channel in cytochrome P450, are conserved.

Figure 1. Crystal structure of human aromatase (PDB ID 4KQ8), colored according to the conservation. The violet areas correspond to the more conserved regions, whereas the dark green ones correspond to the most variable. Heme is shown in red and the substrate androstenedione in light brown. (a) Overall structure of human aromatase. (b) The core structure of aromatase, carrying the most conserved regions. The residues important for substrate binding are also shown.

The key cysteine residue coordinating heme iron is obviously conserved in all the sequences, and it is within a consensus sequence formed by FGFGPRX1CX2GK/R, where X1 is variable (G, A, S, T, or N) whereas X2 is A, V, L, or I. This consensus sequence is also well-conserved in cytochrome P450 (FXXGX(H/R)XCXG), together with the meander region, a loop preceding the cysteine residue [36], which is also well-conserved in most aromatase sequences.

The three Arg residues involved in salt bridges with heme propionyl groups (R115, R145, and R435 in human aromatase) are also present in all the sequences, together with Trp141, and are involved in an H-bond with the heme propionyl group.

A highly conserved motif in cytochrome P450 is the EX1X2R motif located in helix K and involved in salt bridge interactions that are important for its tertiary structure and the correct incorporation of the heme cofactor [36]. This motif is conserved in all sequences; X1 is a serine residue, whereas X2 is L or M in most aromatase sequences.

2.2. Functional Conservation

The level of conservation of amino acids that are relevant for substrate binding and catalysis was then verified in the multiple alignments. A highly conserved alcohol–acid pair is present on helix I in cytochrome P450, and it is part of the proton relay network that allows the formation of the reactive intermediate (Compound I) in the catalytic cycle. In aromatase, the alcohol–acid pair is formed by an aspartic acid residue (D309 in human aromatase) and a threonine residue (T310) that are conserved (exception is made for two fish sequences), and they are preceded by a proline residue (P308) in all the sequences analyzed. When compared to other P450s, this proline residue is unique to aromatase, and it is responsible for the shift of the I-helix axis observed in the crystal structure of the human enzyme [37]. Such a shift is important as it allows the 3-keto moiety of the substrate androstenedione to be accommodated near the fifth turn of the I-helix that is formed by M303 and A307. These two residues are conserved, with some exceptions. The methionine is substituted by an isoleucine in five fish sequences, one amphibian sequence, and one mammal sequence; there is an alanine residue that is a glycine residue in 4.5% of fish sequences and in two invertebrates. Moreover, the shift of the I-helix allows the formation of a hydrogen bond between D309 and the 3-keto oxygen of the substrate. Such an aspartic acid residue has never been changed into a glutamic acid during evolution due to its important role in substrate binding and catalysis [38]. All these residues (303–310) are located on helix I, and they are part of one of six substrate recognition sites (SRSs), namely, SRS-4. The residues involved in androstenedione binding are highly conserved, with some exceptions represented by few fish sequences (Table 1).

Table 1. Conservation of the residues involved in substrate binding and catalysis in human aromatase. The scores are normalized so that the average score for all residues is zero and the standard deviation is one. The lowest score represents the most conserved position in a protein. For reference, the lowest score associated with a fully conserved residue was −1.103, whereas the highest score obtained for a nonconserved residue in human aromatase was +2.844.

Residue Location Conservation Score Notes
C437 K″-L helix loop −1.095  
I305 I-helix −0.936 L/V only in invertebrate Branchiostoma
A306 I-helix −1.002 T in the mammal Capra hircus
D309 I-helix −1.058 Q in CYP19B1 of the fish Halichoeres tenuispinis
T310 I-helix −1.011 I in the fish Maylandia zebra
F221 F-helix −0.805  
W224 F-helix −0.896  
I133 B-C loop −1.038 M in pig aromatase isoform 3
F134 B-C loop −1.073  
V370 K-helix—β3 loop −1.001  
L372 K-helix—β3 loop −0.202 Phe in fishes
V373 K-helix—β3 loop −0.583 S/ T in most fishes and in CYP19A1 of zebrafish and goldfish
M374 β3 −1.031  
L477 β8–β9 loop −1.011  
S478 β8–β9 loop −0.828 A in many sequences, starting from mammals to amphibians. S in fishes.
R192 Helix E −0.974 C or H in some mammals, birds and fishes, including the two isoforms of zebrafish
E483 β9–β10 loop −0.761 Conserved in the two isoforms of zebrafish and goldfish

Two other residues are important for aromatase function; they are predicted to be part of the proton relay network that allows the formation of the reactive Compound I in the typical P450 catalytic cycle: R192 and E483. These residues form a salt bridge in the same position as the one found in the crystal structure of the bacterial cytochrome P450cam [39]. The residues R192 and E483 are highly conserved, starting from the sequences of aromatase from invertebrates. The crystal structure of the bacterial camphor-hydroxylating P450cam from Pseudomonas putida shows that this salt bridge is broken when the P450cam interacts with the redox partner that stabilizes the open conformation of the enzyme, exerting an effector role [39][40][41]. For human aromatase, the redox partner cytochrome P450 reductase (CPR) has been shown to promote substrate binding, acting as an effector [42], and the presence of the R192-E483 salt bridge in the same structural position as P450cam suggests that a similar effect can be exerted by its redox partner CPR.

3. Homology Modeling of Evolutionarily Old Aromatase

Based on the sequence alignment, homology modeling was applied to two aromatase sequences as it was found that they carry significant insertions, in addition to mutations, in key positions.

The invertebrate aromatase sequence from Branchiostoma floridae was selected as it shows an amino acid insertion, 40% of identity, and 60% of homology with the human one. Thus, a homology model was built to study where the main differences between the two aromatase enzymes are located.

A six-amino-acid insertion is present in the invertebrate sequence compared to all the other sequences analyzed (between M276 and D277 in human aromatase), and the model shows that such an insertion elongates the loop connecting helices H’ and the H loop (Figure 2). Moreover, the analysis of the location of the substitutions shows that they are all on the protein surface and on structural elements such as helix G, which are the least conserved ones in aromatase. There are no mutations in the core structure of the protein and the active site, indicating that the main structural scaffold of aromatase was already present in this old protein. Moreover, many mutations are located in the SRSs, indicating that these areas have evolved in vertebrates.

Figure 2. Homology models of evolutionarily old aromatase. (a) Homology model for aromatase from the invertebrate Branchiostoma floridae (green) superimposed onto the crystal structure of human aromatase (blue). The nonconserved regions are shown in orange, and the grey shadow shows the location of the insertion. (b) Zoomed-in view of the active site showing the conserved (green) and nonconserved residues (dark green) involved in substrate binding and catalysis. (c) Homology model for aromatase from the pufferfish Takifugu rubripes (magenta) superimposed to the crystal structure of human aromatase (blue). The grey shadow shows the location of the long insertion (violet). (d) Zoomed-in view of the active site showing the conserved (magenta) and nonconserved residues (dark purple) involved in substrate binding and catalysis. Heme is shown in red and the substrate androstenedione in light brown.

The multiple sequence alignment also shows the presence of some important mutations together with a long insertion in aromatase from some fish species, including the one from pufferfish Takifugu rubripes. In this case, the fish sequence shares 52% of identity and 70% of homology with the human one. A homology model was built in order to predict the possible effect of the substitutions found in the active site. Figure 2 shows the model carrying a long insertion between N421 and V422, which corresponds to the loop connecting helix K’’ and helix L.

Since this long insertion is modeled as a long loop, secondary structure prediction tools were used to verify a possible elongation of the K’’ helix. However, both PsiPred and I-Tasser servers did not predict any secondary structure formation for the amino acids present in that loop. Such a result justifies the absence of such a long and not-necessary loop in the other aromatase sequences.

Concerning the active site, while the substitution of L372 with a phenylalanine does not seem to affect the polarity and dimensions of the catalytic pocket, the substitution of V373 with the polar threonine residue, which in some species is a serine, can be predicted to affect the polarity of the active site (Figure 2). As the substrate carries at least two keto- (as in androstenedione) groups or one keto- group and one hydroxyl group (as in testosterone), the presence of a serine/threonine residue can be predicted to possibly affect the orientation and positioning of the substrate in the active site of the enzyme. Indeed, the Thr/Ser residue could form a hydrogen bond with the substrate. Thus, this substitution seems to be important to properly orient the substrate in the active site for efficient catalysis.

4. Discussion

Aromatase is a unique enzyme carrying out a three-step reaction on the androgen substrate, with the third step leading to the aromatization of the A-ring of the steroid molecule. This intriguing reaction has been the subject of many studies aimed at understanding the mechanism of the third aromatization step [43][44]. Moreover, the crystal structure of the human enzyme has indicated the amino acids within the protein matrix involved in substrate binding and catalysis, and their role has been confirmed by site-directed mutagenesis [45][46].

In this work, sequence and structural alignments were performed with aromatase sequences available on databases. Unfortunately, the number of sequences for the different classes of vertebrates is very different as most of the sequences are available from fishes and mammals and, therefore, a bias is introduced in the conservation score. However, we performed a qualitative analysis in order to see the effect of mutations in key positions using the conservation score as an indicator for the level of conservation.

The multiple alignment shows that the enzyme structural scaffold and the key functional residues have been highly conserved during evolution, with only few exceptions in the aromatase sequences from fishes and invertebrates. Thus, the structural core elements of the protein carrying the residues involved in substrate binding are evolutionarily old and this is reasonable as they guarantee the specific function that aromatase has in species conservation. On the other hand, while some SRSs have also been well-conserved during evolution, SRS-3 has shown the lowest level of conservation (15% of the residues are highly conserved). SRS-3 is located on helix G, a flexible element, which, together with helix F and the F-G loop, is known to be involved in the opening and closure of the access channel for the substrate. Interestingly, helix F and the F-G loop are much more conserved as they belong to SRS-2, which shows 40% of the residues to be highly conserved. Out of the conserved residues, we could identify the ones unique to aromatase, thanks to a structural alignment with the other human P450 enzymes. The data show that some conserved and unique amino acids, such as N135 and Y220, are involved in H-bond networks and have a structural role that supports the positioning of the residues involved in substrate binding in the active site.

A lower level of conservation is found in some of the amino acids that form the positively charged proximal side and in some other residues that are involved in the interaction with the redox partner through the formation of H-bonds. This finding is very interesting as CPR is shared between many P450 enzymes within the same organism. Moreover, we have recently demonstrated that human CPR has an effector role as it facilitates substrate binding by stabilizing the aromatase open conformation, which is optimal for substrate access to the active site [42]. Thus, the data suggest that one of the driving forces for evolution has been the optimization of the interface between aromatase and CPR in order to make aromatase more competitive for the same shared redox partner. Such an optimization involves the introduction of positively charged residues as well as amino acids that form H-bonds and facilitate CPR binding, which, in turn, promotes catalysis.

The other interesting finding is the poor conservation of some residues known to be involved in post-translational modifications. Phosphorylation is a rapid way to modulate enzyme activity compared to regulation at the gene level. Aromatase activity is affected by phosphorylation, and some of the residues that can undergo this post-translational modification have been identified [47][48][49][50]. Phosphorylation of S118 has been reported to decrease aromatase activity in human cell lines [50]. The residue S118 is highly conserved in aromatase sequences from vertebrates, together with R115, which forms the consensus sequence for PKA. This consensus is missing in invertebrates and in few fish sequences (3%).

Another consensus sequence for PKA, as well as for PKG, involves S267 and/or Thr268. These residues are not present in fishes, whereas the consensus sequence for PKA is present in amphibians. On the other hand, the consensus for PKG, which includes two or three basic residues, has appeared late during evolution as it is present in only 15% of the mammal aromatase sequences. Interestingly, this consensus sequence includes R264 in human aromatase that is mutated into a Cys or His in some polymorphisms that are also reported to alter aromatase activity when used in combination with polymorphic variants of CPR [51]. Moreover, they have been associated with an increased risk for estrogen-dependent pathologies such as breast cancer and polycystic ovary syndrome [52][53][54][55].

The other residue known to be phosphorylated is Y361, which appears in amphibians but is not fully conserved even within mammals. Aromatase phosphorylation in this position has been associated with tumor progression in breast cancer cell lines [48]. Indeed, short exposure to estradiol was found to increase aromatase activity through phosphorylation of a tyrosine residue (Y361) by c-Src kinase in estrogen-dependent MCF-7 breast cancer epithelial cells. The authors hypothesized the presence of a positive nongenomic autocrine loop between estradiol and aromatase in MCF-7 breast cancer cells [48]. Moreover, it was also demonstrated that estradiol impairs the ability of the tyrosine phosphatase PTP1B to dephosphorylate aromatase, resulting in increased aromatase activity and estrogen production [56]. The multiple sequence alignment shows that the tyrosine residue in position 361, located on helix K, which is one of the most conserved structural elements in aromatase, appears in few fish species, but it is poorly conserved even among mammals, where it is substituted by an asparagine residue, as in most fishes.

Taken together, the results of the conservation of the phosphorylation sites show that evolution has introduced and is still introducing amino acids in key surface positions that can be phosphorylated and consensus sequences in order to modulate aromatase activity. Thus, the need for quickly and locally altering the estrogen concentration in cells seems to be the other driving force for the evolution of this enzyme. This finding is supported by the fact that a rapid regulation of aromatase activity is known to occur in neurons [57][58] and teleost fishes express aromatase only in glial cells, indicating that the ability to synthesize estrogens in neurons has been acquired during evolution [59][60]. In the brain, the acquisition of phosphorylable sites may be explained by the need to modulate estrogen production in higher vertebrate neurons, where rapid changes in estrogen levels, as a consequence of aromatase phosphorylation, have been associated with important physiological and behavioral responses [58].

It is interesting to note that if, on the one hand, the introduction of phosphorylation sites can be evolutionarily beneficial, as in the case of brain aromatase, on the other hand, phosphorylation of residues that increases aromatase activity can strengthen the negative effects of estrogens, as in the case of breast cancer.

In conclusion, this study on aromatase shows that molecular evolution has worked to maintain a high selectivity for a substrate-specific human cytochrome P450 such as aromatase. However, based on the mutations introduced in key sites, it has been observed that evolution has introduced residues that optimize the interaction with the redox partner and phosphorylation sites that give the possibility of rapidly modulating its activity through phosphorylation. It will be interesting to extend the study to other P450s that are highly substrate-selective to understand how molecular evolution has worked for this group of P450s.

References

  1. Thompson, E.A.; Siiteri, P.K. Utilization of oxygen and reduced nicotinamide adenine dinucleotide phosphate by human placental microsomes during aromatization of androstenedione. J. Biol. Chem. 1974, 249, 5364–5372.
  2. Simpson, E.R.; Mahendroo, M.S.; Means, G.D.; Kilgore, M.W.; Hinshelwood, M.M.; Graham-Lorence, S.; Amarneh, B.; Ito, Y.; Fisher, C.R.; Michael, M.D.; et al. Aromatase cytochrome P450, the enzyme responsible for estrogen biosynthesis. Endocr. Rev. 1994, 15, 342–355.
  3. Coon, M.J. Cytochrome P450: Nature’s most versatile biological catalyst. Annu. Rev. Pharmacol. Toxicol. 2005, 45, 1–25.
  4. Guengerich, F.P. Cytochrome p450 enzymes in the generation of commercial products. Nat. Rev. Drug. Discov. 2002, 1, 359–366.
  5. Di Nardo, G.; Gilardi, G. Natural compounds as pharmaceuticals: The key role of cytochromes p450 reactivity. Trends Biochem. Sci. 2020, 45, 511–525.
  6. Nelson, D.R.; Goldstone, J.V.; Stegeman, J.J. The Cytochrome P450 Genesis Locus: The Origin and Evolution of Animal Cytochrome P450s. Philos. Trans. R. Soc. Lond B Biol. Sci. 2013, 368, 20120474.
  7. Callard, G.V.; Tarrant, A.M.; Novillo, A.; Yacci, P.; Ciaccia, L.; Vajda, S.; Chuang, G.-Y.; Kozakov, D.; Greytak, S.R.; Sawyer, S.; et al. Evolutionary origins of the estrogen signaling system: Insights from amphioxus. J. Steroid Biochem. Mol. Biol. 2011, 127, 176–188.
  8. Callard, G.V.; Pudney, J.A.; Kendall, S.L.; Reinboth, R. In vitro conversion of androgen to estrogen in amphioxus gonadal tissues. Gen. Comp. Endocrinol. 1984, 56, 53–58.
  9. Mizuta, T.; Kubokawa, K. Presence of sex steroids and cytochrome P450 genes in amphioxus. Endocrinology 2007, 148, 3554–3565.
  10. Kishida, M.; Callard, G.V. Distinct cytochrome P450 aromatase isoforms in zebrafish (Danio rerio) brain and ovary are differentially programmed and estrogen regulated during early development. Endocrinology 2001, 142, 740–750.
  11. Tchoudakova, A.; Kishida, M.; Wood, E.; Callard, G.V. Promoter characteristics of two cyp19 genes differentially expressed in the brain and ovary of teleost fish. J. Steroid Biochem. Mol. Biol. 2001, 78, 427–439.
  12. Conley, A.J.; Corbin, C.J.; Hughes, A.L. Adaptive evolution of mammalian aromatases: Lessons from Suiformes. J. Exp. Zool. Part A: Ecol. Genet. Physiol. 2009, 311, 346–357.
  13. Corbin, C.J.; Hughes, A.L.; Heffelfinger, J.R.; Berger, T.; Waltzek, T.B.; Roser, J.F.; Santos, T.C.; Miglino, M.A.; Oliveira, M.F.; Braga, F.C.; et al. Evolution of suiform aromatases: Ancestral duplication with conservation of tissue-specific expression in the collared peccary (Pecari tayassu). J. Mol. Evol. 2007, 65, 403–412.
  14. Hamilton, K.J.; Hewitt, S.C.; Arao, Y.; Korach, K.S. Estrogen hormone biology. Curr. Top. Dev. Biol. 2017, 125, 109–146.
  15. McEwen, B.S.; Milner, T.A. Understanding the broad influence of sex hormones and sex differences in the brain. J. Neurosci. Res. 2017, 95, 24–39.
  16. Galea, L.A.M.; Frick, K.M.; Hampson, E.; Sohrabji, F.; Choleris, E. Why estrogens matter for behavior and brain health. Neurosci. Biobehav. Rev. 2017, 76, 363–379.
  17. Almeida, M.; Laurent, M.R.; Dubois, V.; Claessens, F.; O’Brien, C.A.; Bouillon, R.; Vanderschueren, D.; Manolagas, S.C. Estrogens and androgens in skeletal physiology and pathophysiology. Physiol. Rev. 2017, 97, 135–187.
  18. Brincat, M.P.; Baron, Y.M.; Galea, R. Estrogens and the skin. Climacteric 2005, 8, 110–123.
  19. Knowlton, A.A.; Lee, A.R. Estrogen and the cardiovascular system. Pharmacol. Ther. 2012, 135, 54–70.
  20. Bernasochi, G.B.; Rupasinghe, T.T.W.; Bell, J.R.; Roessner, U.; Boon, W.C.; Delbridge, L.M.D. A novel mass spectrometric methodology facilitates quantification of testosterone and progesterone, but not estrogens in cardiac and adipose tissues. J. Mol. Cell. Cardiol. 2020, 140, 11–12.
  21. Wang, H.; Li, R.; Hu, Y. The alternative noncoding exons 1 of aromatase (Cyp19) gene modulate gene expression in a posttranscriptional manner. Endocrinology 2009, 150, 3301–3307.
  22. Piferrer, F.; Blázquez, M. Aromatase distribution and regulation in fish. Fish Physiol. Biochem. 2005, 31, 215–226.
  23. Cornil, C.A.; Ball, G.F.; Balthazart, J. The dual action of estrogen hypothesis. Trends Neurosci. 2015, 38, 408–416.
  24. Coumailleau, P.; Pellegrini, E.; Adrio, F.; Diotel, N.; Cano-Nicolau, J.; Nasri, A.; Vaillant, C.; Kah, O. Aromatase, estrogen receptors and brain development in fish and amphibians. Biochim. Biophys. Acta 2015, 1849, 152–162.
  25. Pieau, C.; Dorizzi, M. Oestrogens and temperature-dependent sex determination in reptiles: All is in the gonads. J. Endocrinol. 2004, 181, 367–377.
  26. Matsumoto, Y.; Buemio, A.; Chu, R.; Vafaee, M.; Crews, D. Epigenetic control of gonadal aromatase (cyp19a1) in temperature-dependent sex determination of red-eared slider turtles. PLoS ONE 2013, 8, e63599.
  27. Flament, S. Sex Reversal in Amphibians. Sex Dev. 2016, 10, 267–278.
  28. Nakamura, M.; Kobayashi, T.; Chang, X.-T.; Nagahama, Y. Gonadal Sex Differentiation in Teleost Fish. J. Exp. Zool. 1998, 281, 362–372.
  29. Sunobe, T.; Nakamura, M.; Kobayashi, Y.; Kobayashi, T.; Nagahama, Y. Aromatase immunoreactivity and the role of enzymes in steroid pathways for inducing sex change in the hermaphrodite gobiid fish Trimma Okinawae. Comp. Biochem. Physiol. 2005, 141, 54–59.
  30. Baravalle, R.; Ciaramella, A.; Baj, F.; Di Nardo, G.; Gilardi, G. Identification of endocrine disrupting chemicals acting on human aromatase. Biochim. Biophys. Acta Proteins Proteom. 2018, 1866, 88–96.
  31. Zhang, C.; Schilirò, T.; Gea, M.; Bianchi, S.; Spinello, A.; Magistrato, A.; Gilardi, G.; Di Nardo, G. Molecular basis for endocrine disruption by pesticides targeting aromatase and estrogen receptor. Int. J. Environ. Res. Public Health 2020, 17, 5664.
  32. Kloas, W. Amphibians as a model for the study of endocrine disruptors. Int. Rev. Cytol. 2002, 216, 1–57.
  33. Scholz, S.; Renner, P.; Belanger, S.E.; Busquet, F.; Davi, R.; Demeneix, B.A.; Denny, J.S.; Léonard, M.; McMaster, M.E.; Villeneuve, D.L.; et al. Alternatives to in vivo tests to detect endocrine disrupting chemicals (edcs) in fish and amphibians—Screening for estrogen, androgen and thyroid hormone disruption. Crit. Rev. Toxicol. 2013, 43, 45–72.
  34. Zhao, J.; Mak, P.; Tchoudakova, A.; Callard, G.; Chen, S. Different catalytic properties and inhibitor responses of the goldfish brain and ovary aromatase isozymes. Gen. Comp. Endocrinol. 2001, 123, 180–191.
  35. Tong, S.K.; Chiang, E.F.; Hsiao, P.H.; Chung, B. Phylogeny, expression and enzyme activity of zebrafish cyp19 (P450 aromatase) genes. J. Steroid Biochem. Mol. Biol. 2001, 79, 299–303.
  36. Hasemann, C.A.; Kurumbail, R.G.; Boddupalli, S.S.; Peterson, J.A.; Deisenhofer, J. Structure and function of cytochromes P450: A comparative analysis of three crystal structures. Structure 1995, 3, 41–62.
  37. Ghosh, D.; Griswold, J.; Erman, M.; Pangborn, W. Structural basis for androgen specificity and oestrogen synthesis in human aromatase. Nature 2009, 457, 219–223.
  38. Di Nardo, G.; Breitner, M.; Bandino, A.; Ghosh, D.; Jennings, G.K.; Hackett, J.C.; Gilardi, G. Evidence for an elevated aspartate pKa in the active site of human aromatase. J. Biol. Chem. 2015, 290, 1186–1196.
  39. Tripathi, S.; Li, H.; Poulos, T.L. Structural basis for effect or control and redox partner recognition in cytochrome P450. Science 2013, 340, 1227–1230.
  40. Liou, S.-H.; Mahomed, M.; Lee, Y.-T.; Goodin, D.B. Effector roles of putidaredoxin on cytochrome P450cam conformational states. J. Am. Chem. Soc. 2016, 138, 10163–10172.
  41. Hollingsworth, S.A.; Batabyal, D.; Nguyen, B.D.; Poulos, T.L. Conformational selectivity in cytochrome P450 redox partner interactions. Proc. Natl. Acad. Sci. USA 2016, 113, 8723–8728.
  42. Zhang, C.; Catucci, G.; Di Nardo, G.; Gilardi, G. Effector role of cytochrome P450 reductase for androstenedione binding to human aromatase. Int. J. Biol. Macromol. 2020, 164, 510–517.
  43. Yoshimoto, F.K.; Guengerich, F.P. Mechanism of the third oxidative step in the conversion of androgens to estrogens by cytochrome P450 19A1 steroid aromatase. J. Am. Chem. Soc. 2014, 136, 15016–15025.
  44. Khatri, Y.; Luthra, A.; Duggal, R.; Sligar, S.G. Kinetic solvent isotope effect in steady-state turnover by CYP19A1 suggests involvement of Compound 1 for both hydroxylation and aromatization steps. FEBS Lett. 2014, 588, 3117–3122.
  45. Lo, J.; Di Nardo, G.; Griswold, J.; Egbuta, C.; Jiang, W.; Gilardi, G.; Ghosh, D. Structural basis for the functional roles of critical residues in human cytochrome P450 aromatase. Biochemistry 2013, 52, 5821–5829.
  46. Di Nardo, G.; Gilardi, G. Human aromatase: Perspectives in biochemistry and biotechnology: Human Aromatase. Biotechnol. Appl. Biochem. 2013, 60, 92–101.
  47. Baravalle, R.; Di Nardo, G.; Bandino, A.; Barone, I.; Catalano, S.; Andò, S.; Gilardi, G. Impact of R264C and R264H polymorphisms in human aromatase function. J. Steroid. Biochem. Mol. Biol. 2017, 167, 23–32.
  48. Catalano, S.; Barone, I.; Giordano, C.; Rizza, P.; Qi, H.; Gu, G.; Malivindi, R.; Bonofiglio, D.; Andò, S. Rapid estradiol/ERα signaling enhances aromatase enzymatic activity in breast cancer cells. Mol. Endocrinol. 2009, 23, 1634–1645.
  49. Hayashi, T.; Harada, N. Post-translational dual regulation of cytochrome P450 aromatase at the catalytic and protein levels by phosphorylation/dephosphorylation. FEBS J. 2014, 281, 4830–4840.
  50. Miller, T.W.; Shin, I.; Kagawa, N.; Evans, D.B.; Waterman, M.R.; Arteaga, C.L. Aromatase is phosphorylated in situ at serine-118. J. Steroid. Biochem. Mol. Biol. 2008, 112, 95–101.
  51. Parween, S.; Rojas Velazquez, M.N.; Udhane, S.S.; Kagawa, N.; Pandey, A.V. Variability in loss of multiple enzyme activities due to the human genetic variation P284T located in the flexible hinge region of NADPH cytochrome P450 oxidoreductase. Front. Pharmacol. 2019, 10, 1187.
  52. Hemimi, N.; Shaafie, I.; Alshawa, H. The study of the impact of genetic polymorphism of aromatase (CYP19) enzyme and the susceptibility to polycystic ovary syndrome (575.5). FASEB J. 2014, 28, 575-5.
  53. Jin, J.L.; Sun, J.; Ge, H.J.; Cao, Y.X.; Wu, X.K.; Liang, F.J.; Sun, H.X.; Ke, L.; Yi, L.; Wu, Z.W.; et al. Association between CYP19 gene SNP rs2414096 polymorphism and polycystic ovary syndrome in Chinese women. BMC. Med. Genet. 2009, 10, 139.
  54. Mehdizadeh, A.; Kalantar, S.M.; Sheikhha, M.H.; Aali, B.S.; Ghanei, A. Association of SNP rs.2414096 CYP19 gene with polycystic ovarian syndrome in Iranian women. Int. J. Reprod. Biomed. 2017, 15, 491–496.
  55. Lee, K.M.; Abel, J.; Ko, Y.; Harth, V.; Park, W.Y.; Seo, J.S.; Yoo, K.Y.; Choi, J.Y.; Shin, A.; Ahn, S.H.; et al. Genetic polymorphisms of cytochrome P450 19 and 1B1, alcohol use, and breast cancer risk in Korean women. Br. J. Cancer 2003, 88, 675–678.
  56. Barone, I.; Giordano, C.; Malivindi, R.; Lanzino, M.; Rizza, P.; Casaburi, I.; Bonofiglio, D.; Catalano, S.; Andò, S. Estrogens and PTP1B function in a novel pathway to regulate aromatase enzymatic activity in breast cancer cells. Endocrinology 2012, 153, 5157–5166.
  57. Cornil, C.A.; Ball, G.F.; Balthazart, J. Functional significance of the rapid regulation of brain estrogen action: Where do the estrogens come from? Brain Res. 2006, 1126, 2–26.
  58. Balthazart, J.; Choleris, E.; Remage-Healey, L. Steroids and the brain: 50 years of research, conceptual shifts and the ascent of non-classical and membrane-initiated actions. Horm. Behav. 2018, 99, 1–8.
  59. Forlano, P.M.; Deitcher, D.L.; Myers, D.A.; Bass, A.H. Anatomical distribution and cellular basis for high levels of aromatase activity in the brain of teleost fish: Aromatase enzyme and mRNA expression identify glia as source. J. Neurosci. 2001, 21, 8943–8955.
  60. Diotel, N.; Le Page, Y.; Mouriec, K.; Tong, S.-K.; Pellegrini, E.; Vaillant, C.; Anglade, I.; Brion, F.; Pakdel, F.; Chung, B.-C.; et al. Aromatase in the brain of teleost fish: Expression, regulation and putative functions. Front. Neuroendocrinol. 2010, 31, 172–192.
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