Microbial biodegradation was reported as the major mechanism which removes estrogens from engineered and natural polluted environments (Yu et al., 2013; Huang et al., 2014; Liyanage et al., 2024). Despite many studies on estrogens biodegradation, there are still many knowledge gaps regarding the catabolic pathways. Moreover, the involved enzymes, genes, and their regulation mechanisms are not fully characterized yet. Degradation of estrogens by aerobic bacteria has been reported, and many estrogens–degrading bacteria have been isolated from various environments (Yu et al., 2013; Zhang et al., 2015; Olivera and Luengo, 2019; Chiang et al., 2020; Pratush et al., 2020a).

Estrogens, like other steroids, are carbon-rich and highly reduced compounds that are abundant and ubiquitous in the environment. Therefore, they represent a precious carbon and energy source for some microorganisms (Chiang et al., 2020). In fact, some microorganisms (including bacteria, yeasts, fungi, and microalgae) can perform biotransformation reactions on estrogens (Pratush et al., 2020a). However, only dedicated bacteria have the ability to mineralize estrogens (complete degradation to CO₂) (Bergstrand et al., 2016; Holert et al., 2018). Recent studies identified steroid-degrading bacteria and elucidated the biodegradation mechanisms using both culture-dependent and culture-independent approaches.

Bacteria that can degrade estrogens under aerobic conditions were isolated mainly from activated sludge, soil, compost, sandy aquifers, and very little from the marine environment. Although the isolates belong to different phyla, most estrogen-degrading bacteria reported to date belong mainly to the Actinomycetota (Actinobacteria) and Pseudomonadota (Proteobacteria) (Supplementary Tables S1, S2) (Bergstrand et al., 2016; Chiang et al., 2020; Pratush et al., 2020a; Shah et al., 2024).

Key enzymes involved in aerobic estrogen biodegradation and biotransformation are classified in different categories including dehydrogenases, hydroxylases, ring-cleavage dioxygenases, isomerases, monooxygenases, hydratases, and demethylases, as well as cytochrome P450 (Chiang et al., 2020; Pratush et al., 2020a).

Various estrogen biotransformation reactions were reported, including hydroxylation, dehydrogenation, and ring cleavage reactions (Figure 2). Most of the studies have focused on the aerobic biodegradation E1 and E2 as model substrates. On the contrary, E3 and EE2 have not been less thoroughly investigated. Moreover, the biodegradation of estrogens under anaerobic conditions was reported only recently (Wang et al., 2020).

Despite the high structural similarity among estrogens, a diversity of biotransformation reactions and degradation pathways have been reported in the literature. Some of those reactions/pathways were proposed based on the detection and identification of metabolic intermediates. Another important feature of estrogen biodegradation is the fact that estrogens are interconvertible, i.e., one estrogen can be transformed to another during biodegradation or biotransformation. To date, three major catabolic pathways for steroid biodegradation have been reported. Under aerobic conditions, sterols and androgens are degraded via the 9,10-seco pathway, while estrogens are commonly degraded via the 4, 5-seco pathway.

On the contrary, under anaerobic conditions sterols and androgens are catabolized via the 2,3-seco pathway, while the anaerobic degradation of estrogens has not been studied in detail (Chiang et al., 2020; Jacoby et al., 2025). Regardless of the oxygen availability, the three pathways merge at a common key intermediate, namely HIP (3aα-H-4α(3′-propanoate)-7aβ-methylhexahydro-1,5-indanedione), which is subsequently degraded via a common central pathway to produce metabolites that are further degraded through central metabolic networks (Figure 3). In the following, different estrogen biodegradation and biotransformation mechanisms will be presented. For clarity, E2 and E1 will be discussed together because their degradation pathways are linked. Subsequently, EE2 and E3 degradation pathways will be presented separately.

The first E2-degrading bacterium was isolated from activated sludge and identified as Novosphingobium tardaugens sp. nov. (Fujii et al., 2002), and Coombre et al. (1966) proposed the first pathway for E1 degradation by the soil bacterium Nocardia sp. E110. It was proposed that a dioxygenase catalyzes the cleavage of the A-ring of E1 and this pathway was included as a part of E2 degradation pathways reported in subsequent studies, proposing that E2 degradation can be initiated by one of the following reactions: (a) hydroxylation of the A-ring at C4, (b) hydroxylation of a saturated ring, (c) dehydration of the D-ring at C17, and (d) dehydrogenation of the D-ring at C17 (Yu et al., 2013) (Figure 3).

The first mode of degradation was reported by Kurisu et al. (2010) who detected 4-OH-E2 during degradation by the soil isolate Sphingomonas sp. ED8, which suggests that E2 was hydroxylated at C4. The authors proposed that degradation of this hydroxylated product proceeds via meta cleavage. Kurisu et al. (2010) detected other products of E2 degradation by the ED8 strain, including hydroxy-E2, keto-E2, keto-E1, and 3-(4-hydroxyphenyl)-2-hydroxyprop-2-enoic acid. This latter product suggested that E2 degradation was initiated at a saturated ring. However, the authors did not further investigate how the saturated ring was cleaved. In another study, Nakai et al. (2011) used Nitrosomonas europeae to degrade E2 and detected a metabolite identified as estra-1, 3, 5 (10), 16 tetraen-3-ol (estratetra enol, or E0), which results from dehydration at C17 of the D-ring. Nitrosomonas europeae further degraded E0 to non-estrogenic compounds.

Transformation of E2 to E1 via dehydrogenation of C17 at the D-ring was also reported (Yu et al., 2007). Estrone can be further transformed to 4-OH-E1, followed by further degradation via meta cleavage (Kurisu et al., 2010). Lee and Liu (2002) suggested another degradation pathway for E2 when utilized by mixed sewage bacteria based on the detection of a new metabolite designated XI, which contains a lactone structure at the D-ring. All those previous studies proposed some biotransformation reactions, albeit complete pathway for estrogen degradation was not elucidated in terms of metabolic intermediates, genes, and enzymes, thus rendering the assessment of the fate and biodegradation potential of estrogens in the polluted environments challenging. Furthermore, the structural elucidations were mostly based on mass-spectrometry and no ring-cleave products were reported.

Biodegradation of EE2 has been less frequently reported, and most of the available studies were conducted with microbial cultures that can utilize/transform EE2 in the presence of co-substrates, thus following a cometabolic mode of degradation (O’Grady et al., 2009; Pauwels et al., 2008). This could be attributed to recalcitrance of EE2 compared to the natural estrogens (Zhang et al., 2015; Olivera and Luengo, 2019). Co-substrates may strengthen central metabolism, promote cell growth, and supply reducing power, and can even induce the expression of some enzymes that play a role in EE2 metabolism (Gröning et al., 2007; Tran et al., 2013; Fischer and Majewsky, 2014). Currently, no complete EE2 biodegradation pathway is available and knowledge of the involved genes, enzymes are largely lacking.

Only a few bacteria can degrade EE2 as a sole carbon and energy source such as Sphingobacterium sp. JCR5 (Haiyan et al., 2007) and Rhodococcus equi (Yoshimoto et al., 2004). The number of reported microorganisms that can transform or cometabolize EE2 is very limited including Sphingomonas spp., Pseudomonas spp., Nitrosomonas europaea, the cyanobacterium Microcystis novacekii, the microalgal strain Selenastrum capricornutum, and the fungi Phoma sp. and Pleurotus ostreatus (Flint, 2019; Pratush et al., 2020a). Degradation of EE2 occurs under aerobic and anaerobic conditions, albeit higher degradation rates were reported under aerobic conditions (Aris et al., 2014; Cajthaml et al., 2009). EE2 degradation was slow in marine sediments (Ying and Kookana, 2003) and aquifer material (Ying et al., 2003), while in agricultural soils (Colucci and Topp, 2001) and nitrifying activated sludge (Vader et al., 2000) it was more rapidly degraded. Earlier studies reported biotransformation of EE2 by some bacteria and microalgae (Haiyan et al., 2007; Della Greca et al., 2008) including Sphingobium sp. JCR5, Rhodococcus zopfii, and Nitrosomonas europaea EE2 (Shi et al., 2002; Yoshimoto et al., 2004; Haiyan et al., 2007; Aris et al., 2014). One of the possible degradation pathways starts by transforming EE2 to E1, then E3, but no further degradation was reported (Li et al., 2013a, 2013b), or transformation of EE2 to E1, followed by ring cleavage (Cajthaml et al., 2009). At low concentrations, EE2 could be co-metabolized with E1, E2, or E3 in sludge and wastewater; however, no further metabolites were detected (Cajthaml et al., 2009; Writer et al., 2012; Rhee et al., 2014).

Several studies reported biotransformation reactions for EE2 without proposing complete degradation pathways (Figure 9). Yi and Harper (2007) detected 2-OH-EE2 during their studies on EE2 degradation by a nitrifying culture and showed that the A-ring was cleaved before the other rings, probably due to the higher electron density of the A-ring carbon units. These findings suggested that EE2 can be cometabolically transformed under nitrification conditions.

Some algal species can transform EE2 into EE2-glucoside, 3-β-D-glucopyranosyl-2-hydroxy-EE2, 3-β-D-glucopyranoside-6β-hydroxy-EE2, 17α-ethinyl-1,4, estradien-10,17β-diol-3-one, and 6-α-OH-EE2 (Della Greca et al., 2008). Thus, hydroxylation, oxidation, and glycosylation reactions were the main biotransformation reactions. In another study, Haiyan et al. (2007) isolated Sphingobacterium sp. JCR5 strain from activated sludge and showed that it can grow on EE2 as a sole carbon and energy source. Mass spectrometry analysis revealed that EE2 is first transformed to E1, followed by hydroxylation and oxidation reactions at C9, leading to B-ring cleavage. The A-ring is then hydroxylated at C4.

Co-metabolism of EE2 was studied with several bacteria isolated from compost. The isolated bacteria degraded E1, E2, and E3. However, EE2 was not degraded when used as a sole carbon source, albeit it was degraded in the presence of E2, and in this case E1 was detected as a metabolite. No other metabolites were detected, and EE2 removal started after complete removal of E2, and it increased as the initial E2 concentration increased (Pauwels et al., 2008). R. erythropolis and R. equi removed EE2 from minimal medium in the presence of co-substrates (adipic acid and glucose). Two metabolites were identified as phenol and a derivative of EE2 with a higher molecular mass (331 atomic mass unit) where the ethinyl group was absent (O’Grady et al., 2009).

In a study by Ma et al. (2018), lab-scale percolation columns were used to investigate EE2 biodegradation during groundwater recharge with reclaimed water. Based on metabolite analysis, different pathways for EE2 degradation were proposed depending on groundwater recharge mode. The dominant pathway in wetting and drying alternative recharge mode started by oxidation of C17 at the D-ring, which is then hydroxylated and cleaved. Thus, EE2 is transformed to E1 → E3 → 7-hydroxy-1-hydroxymethyl-2-methyl-1,2,3,4,4a,9.10,10a-octahydrophenanathrene-2-carboxylic acid → 2-hydroxy-2,4-diene-1, 6-dioic acid. On the contrary, in a continual recharge mode, EE2 metabolism started by hydroxylation of C4 on the A-ring and then the A- or B-ring is cleaved. This transition in the degradation mechanism was proposed to be related to dissolved oxygen and the structure of the involved microbial community.

Recent studies on EE2 degradation in synthetic wastewater by microalgae proposed hydroxylation reactions at C2 or C4, followed by ring opening (Wang et al., 2019b) (Figure 10). However, enzymes and genes of the proposed biochemical reactions were not reported. Peng et al. (2023) studied EE2 degradation by Pseudomonas citronellolis SJTE-3 and showed that it can grow on and degrade EE2 only in the presence of co-substrates like ethanol and glucose. The authors also identified two new degradation metabolites. Metabolism of EE2 was inducible, and 8 genes were significantly upregulated in the presence of EE2, including genes encoding a short chain dehydrogenase, a membrane transporter, and a cytochrome P450 hydroxylase, which were crucial for EE2 metabolism.

Guo et al. (2021) studied the effect of rhamnolipid biosurfactants produced by Pseudomonas aeruginosa MIG-N146 on the biodegradation of EE2 in shake flasks containing sediment/water systems. The authors reported enhanced biodegradation of EE2 as the rhamnolipid concentration increased, which was attributed to better mass transfer and stimulation of the indigenous microbial consortium. Three metabolic intermediates were detected including a dehydrogenation product of EE2 at C15-C16 of the D-ring, and a carboxylation product of EE2. This study did not report any data on the involved enzymes or microbial community.

Recently, bacteria affiliated with the genera Aeromonas, Rhizobium, and Paraburkholderia isolated from an acid mine drainage degraded EE2 in a minimal medium and produced several degradation metabolites which were mainly ring cleavage products (Palma and Costa, 2024). The authors mentioned that EE2 was the sole carbon source in the cultures. However, EE2 was added from a methanol stock solution, and it was not clear whether it was evaporated before inoculation. Furthermore, while the aim of the study was to isolate EE2-degrading bacterial consortia, all the enrichment and isolation procedures were performed in a rich medium (LB medium) in the presence of paracetamol.

Compared to E1, E2 and EE2, a limited number of studies addressed E3 biodegradation. In most of those studies, E3 was mentioned in the context of substrate spectrum experiments together with other steroid estrogens. Moreover, those studies reported only E3 biotransformation reactions, albeit a complete E3 mineralization pathway is lacking. For instance, Ke et al. (2007) isolated three bacteria from marine sand in microcosms using E1, E2, E3, and EE2 as sole carbon sources. The isolates were identified as members of the genera Sphingomonas, Agromyces, and Acinetobacter. These bacteria, individually or as a consortium, had different substrate spectra in terms of estrogen degradation. The strain identified as Acinetobacter sp. degraded E3 in aerobic cultures and transiently produced 16α-OH-E1, which was reported as a metabolite of E2 degradation by sewage bacteria (Lee and Liu, 2002) and of E1 degradation by intestinal bacteria (Järvenpää et al., 1980). Moreover, Staphylococcus aureus can reduce 16α-OH-E1 to E3 under anaerobic conditions (Järvenpää et al., 1980). Nitrifying activated sludge and ammonia-oxidizing bacterium Nitrosomonas europaea degraded E1, E2, E3 and EE2 in minimal medium (Shi et al., 2004). However, no further information was reported on the degradation mechanisms. Ueda et al. (2012) isolated and characterized a laccase from the mushroom Pleurotus eryngii and showed that it can remove E3 from aqueous solutions. A Comamonas testosteroni strain isolated from animal waste grew on E3 as a sole carbon and energy source (Liu et al., 2021). Nothing was reported in this study on the specific E3 degradation products, enzymes, or genes. However, genomic analysis revealed the presence of 22 steroid degradation genes. The authors also identified a testosterone degradation gene cluster and a gene encoding 17β-hydroxysteroid dehydrogenase, which catalyzes the transformation of E2 to E1. In the same context, Rhodococcus sp. PI4, a marine isolate, utilized E2, E3, and testosterone as sole carbon sources and induced the expression of a 17β-hydroxysteroid dehydrogenase which transforms E3 to 16α-OH-E1 (Ye et al., 2019).

A Gordonia sp. strain R9 isolated from chicken leachates utilized E3 and other steroid estrogens as sole carbon sources (Liu et al., 2020). Although no information on E3 degradation was reported, the authors detected E3 as a metabolite of E2. Estriol was also reported as a product of E2 degradation by Microbacterium resistens MZT7 (Hao et al., 2022) and Rhodococcus sp. RCBS9 (Hao et al., 2024), and it was not clear how E3 is further metabolized.

A study by Hsiao et al. (2021) showed that a Rhodococcus sp. strain B50 can grow on E3 as a sole carbon source. However, no further information on the mechanism of E3 degradation was reported. Wang P. et al. (2019) isolated a Pseudomonas putida strain SJTE-1 from activated sludge and showed that it can degrade E1, E2, E3 and testosterone as sole carbon sources. Nothing was reported in this study on the E3 degradation mechanisms.

Evidence supporting the conclusion that E3 is degraded by some bacteria via the 4,5-seco pathway was reported in recent studies. We showed that a bacterial consortium obtained from raw domestic sewage transformed E3 to E1 potentially via transient formation of 16α-OH-E1. Further degradation of E1 proceeded via the 4,5-seco pathway and the community structure of the consortium shifted strongly toward Croceicoccus estronivorus which dominated the community after 10 days of incubation (Hashem et al., 2025). Liu et al. (2024) reported the degradation of E3 by Novosphingobium sp. ES2-1 and proposed two alternative pathways for E3 degradation based on metabolite analysis. The first proposed pathway proceeds via E3 transformation to E1, which is further degraded via the 4,5-seco pathway. However, the authors did not clarify how E3 was transformed to E1. They proposed oxidation and dehydration reactions, albeit no biochemical evidence was presented. The other pathway proposed by Liu et al. (2024) also proceeds by transforming E3 to E1, then to 4-OH-E1, which undergoes B-ring cleavage, resembling the 9,10-seco pathway. However, nothing was reported on the genes and enzymes involved in this later pathway. Moreover, the mechanism of the B-ring cleavage was not discussed. The various detected metabolites were not quantified, which is essential to reflect on the relative significance or dominance of the two proposed pathways. Degradation of E3 via the 9,10-seco pathway was also proposed by Thathola et al. (2024) using a psychrotolerant Pseudomonas proteolytica strain based on the detection of E1, 16-OH-E1, in addition to two proposed ring cleavage downstream products: 2-hydroxy2-4-dienevaleric acid, and 2-hydroxy-2,4-diene-1,6-dioic acid. These products were identified only using mass spectrometry and no further biochemical evidence was presented.

In contrast to the aerobic estrogen degradation, biodegradation of estrogens under anaerobic conditions has been very rarely investigated. The low aqueous solubility (1.5 mg/L at room temperature) of estrogens and stability of the aromatic A-ring render estrogens challenging substrates for microbial utilization (Shareef et al., 2006; Wang et al., 2020). Therefore, under aerobic conditions, bacteria depend on oxygenases and oxygen to activate and cleave the aromatic A-ring through the 4,5-seco pathway (Chen et al., 2017; Chiang et al., 2020). Lack of oxygen in anaerobic environments, such as river and marine sediments, prohibits estrogen degradation and makes the process slower compared to aerobic degradation. Hence, anaerobic environments are considered as major reservoirs for estrogens (Hanselman et al., 2003; Czajka and Londry, 2006). Probably, slow degradation rates and difficulties in isolating anaerobic estrogen-degrading bacteria explain why anaerobic estrogen degradation has been less investigated compared to aerobic degradation. Only two bacterial species are known to degrade estrogens under anaerobic conditions, namely, Denitratisoma oestradiolicum and Steroidobacter denitrificans (Fahrbach et al., 2006, 2008).

To elucidate the biochemical mechanisms of anaerobic estrogen degradation, Wang et al. (2020) isolated a denitrifying bacterium, Denitratisoma sp. strain DHT3, from a municipal WWTP and showed that it can efficiently degrade estrogens under denitrifying conditions. The authors sequenced the genome of this strain and performed comparative transcriptomics to identify potential genes involved in anaerobic estrogen degradation using E2- and testosterone-grown cultures. Genes involved in the anaerobic 2,3-seco pathway were expressed at similar levels in both cultures. On the contrary, genes of transport, salvage, and reductive activation of cobalamin were upregulated in the E2 culture. In addition, a gene cluster encoding a putative methyltransferase (emtABCD) was also upregulated in the E2 culture. Gene disruption experiments confirmed that emtA plays a role in anaerobic estrogen degradation by the DHT3 strain.

Interestingly, the emtABCD gene cluster is present in the genome of estrogen-degrading anaerobes, but lacking in steroid-degrading bacteria that cannot utilize estrogens (Wang et al., 2020). Moreover, metabolite analysis of E1-grown cultures of the DHT3 strain revealed several androgenic metabolites including two ring-cleavage products which are characteristic of the 2,3-seco pathway, i.e., 17β-hydroxy-1-oxo-2,3-seco-androstan-3-oic acid (2,3-SAOA) and HIP. These findings suggested that anaerobic estrogen degradation in the DHT3 strain is initiated by conversion of the C₁₈ estrogens to C₁₉ androgens which are further degraded via the 2,3-seco pathway. The key step is the conversion of estrogens to androgens via a methylation reaction catalyzed by a cobalamin-dependent methyltransferase (emtABCD) (Figure 11).

These findings highlight microbe-host metabolic relationships mediated through retroconversion of estrogens into androgens based on the role of sex steroids in bidirectional metabolic interactions between bacteria and their eukaryotic hosts (Karavolos et al., 2013; Markle et al., 2013; vom Steeg and Klein, 2017). Conversion of estrogens into androgens represents the reverse process of estrogen (C₁₈) biosynthesis from androgens (C₁₉) via removal of the C₁₉ angular methyl group, leading to formation of the aromatic A-ring (Miyairi and Fishman, 1985), and it is a thermodynamically challenging reaction which has not been reported in any organism (Wang et al., 2020).

The reverse reaction, namely, the conversion of C₁₉ androgens into C₁₈ estrogens, was recently discovered in strict anaerobic bacteria (Wang et al., 2025). This aromatase-independent estrogenesis is catalyzed by a cobalamin-dependent methyltransferase (AbeABC), which is 57% identical to the EmtAB methyltransferase identified in the denitrifying Denitratisoma spp. Homologs of AbeAB were identified in uncultivated bacteria from marine sediments, suggesting widespread occurrence of anaerobic estrogenesis in anoxic ecosystems. Based on those findings, some ecological implications can be envisaged. For instance, the reversible conversion of estrogens into androgens in anaerobic environments (animal guts and sediments) could potentially affect steroid hormone cycling and fate in these habitats. Furthermore, the unique presence of the abeAB genes in microbial genomes render them a functional biomarker that enables monitoring of ecosystem health and tracking steroid pollution (Wang et al., 2025). A recent study showed that EE2 was transformed to E1 and E2 in an electrochemical anaerobic membrane bioreactor (Lei et al., 2025), suggesting that anaerobic degradation of EE2 may proceed via E2 or E1 conversion to androgen. The discovery of this unprecedented steroid microbial bioconversion paves the way for potential diverse applications in environmental monitoring and bioremediation.

The recent studies and findings on anaerobic estrogen biodegradation reflect an increasing interest in this topic whose understanding has been lagging compared to the aerobic degradation processes. Three overlapping drivers can be envisaged to have motivated researchers to investigate this topic further. The first is purely scientific curiosity to understand how bacteria deal with challenging substrates like steroid estrogens in the absence of oxygen or under oxygen fluctuation conditions. Recalling the unprecedented biochemical aspects revealed by extensive studies on the anaerobic degradation of aromatic compounds (Ismail and Gescher, 2012; Boll and Estelmann, 2020; Boll et al., 2020a, 2020b), enables us to reconcile the importance of this driver. The second driver could be the rising global concerns on steroid estrogens as emerging pollutants and the need to better understand their fate and biodegradation mechanisms particularly in anaerobic environments where they tend to accumulate (Hanselman et al., 2003; Czajka and Londry, 2006). While earlier and recent studies addressed the fate and kinetics of estrogen degradation in anaerobic ecosystems (Muller et al., 2010b; Zheng et al., 2012; Luo et al., 2024), unraveling the biochemical mechanisms is pivotal for the development of efficient bioremediation technologies for polluted anaerobic environments due to the recalcitrance of estrogens in the (Pratush et al., 2020a). The third driver is related to the overwhelming recent interest in understanding the functional relevance of the gut microbiome in health and pathogenesis, which is instrumental toward the development of novel therapies and probiotics (Li et al., 2022; Diviccaro et al., 2025). In this context, it is known that sex steroids mediate bidirectional interactions between bacteria and their eukaryotic hosts (Kverka and Stepan, 2024; Nieto et al., 2025). Moreover, bacteria can alter a host’s sex steroid profile through interconversions (biotransformations), utilization as a carbon and electron source, and affecting endogenous estrogen metabolism in postmenopausal women (Horinouchi et al., 2012; Markle et al., 2013; Ridlon et al., 2013; Fuhrman et al., 2014; Zhang et al., 2025). While the three proposed drivers guide research in different directions with different objectives, they share better understanding of the subject matter as a key enabler and will continue to fuel research on estrogen microbial metabolism and associated applications in health care and environmental biotechnology.

A C₁₃ metabolite (HIP), containing the C/D rings of the steroid substrate, was commonly identified in all bacterial pathways for steroid catabolism. Hence, it is degraded through a common central catabolic pathway which targets the C/D rings (Table 3; Figure 12). After activation by a specific acyl-CoA synthetase (FadD3), the remaining carbons of the steroid B-ring are removed through β-oxidation. This is followed by hydrolytic cleavage the D-ring by an enoyl-CoA hydratase (EchA20), and another hydrolase (IpdAB) opens the C-ring. Interestingly, gene mining studies revealed the presence of conserved gene clusters for HIP degradation in the majority of experimentally verified steroid-degrading bacteria (Chiang et al., 2020).

In addition to biological degradation and transformation, estrogens are exposed to several abiotic factors in the environment, which can lead to some degradation and transformation reactions, including chemical and photodegradation/transformation. These abiotic reactions may affect the fate of estrogens in the environment and can also directly interfere with the susceptibility of estrogens to biodegradation and biotransformation (Griffith et al., 2023). This depends on the nature of the products of the abiotic reactions, which could be more prone to biodegradation or more recalcitrant than the estrogen substrate. For instance, chlorine is commonly applied as a disinfectant during water treatment due to its strong oxidation capability, continuous disinfection, and low cost (Deborde and von Gunten, 2008). However, chlorine often produces undesirable intermediate byproducts and potentially carcinogenic halogenated byproducts (Richardson and Kimura, 2016).

Estrogen degradation during chlorination was reported mostly in lab-scale studies, showing the formation of intermediate products and disinfection byproducts (Deborde et al., 2004; Lee and von Gunten, 2009). At an initial concentration of 50 mg/L, E2 was completely degraded after 10 min during chlorination at pH 7.5 (Hu et al., 2003). In batch tests, E3 degradation during chlorination followed a second-order kinetic (Zhao et al., 2014). Moreover, E1 degradation with free chlorine produced chlorinated estrone by-products (Nakamura et al., 2007). The kinetics and products of E2 chlorination were also studied in a pilot-scale water distribution system (He et al., 2016; Li et al., 2017). Further studies by Li et al. (2017) on the kinetics and mechanism of E2 chlorination in a pilot-scale water distribution system showed that chlorination degraded E2 in three stages: (1) halogenation of the aromatic ring, (2) cleavage of the benzene moiety and chlorine substitution, (3) production of trihalomethanes and halogenated acetic acids from the phenolic intermediates.

During E3 chlorination in a pilot-scale water distribution system, E2, E1, 16α-OH-E1, and chlorinated E3, in addition to four types of disinfection by-products including trihalomethanes, haloacetonitriles, haloketones, and chloral hydrate were identified as intermediate products (Dong et al., 2019). Based on the identified products, three possible pathways for E3 chlorination were proposed (Figure 13). During water chlorination, EE2 was transformed into 4-chloro-EE2 and 2,4-dichloro-EE2. In the presence of Br⁻, the bromo-analogs of the chloro-EE2 derivatives were additionally produced (Lee and von Gunten, 2009). In drinking water, chlorination of E2 produced several transformation products including 2,4-dichloro-E2, monochloro-E1, 2,4-dichloro-E1, and by-product such as 4-[2-(2,6-dichloro-3-hydroxyphenyl)ethyl]-7α-methyloctahydroinden-5-one (Hu et al., 2003).

Chlorinated E1, such as 2-chloro-E1, 4-chloro-E1, 2,4-dichloro-E1, were detected in a sewage treatment plant effluent in Japan. Moreover, in a buffer of pH 7 and 9 combining E1 with sodium hypochlorite, the same chlorinated products were produced in addition to 1,4-estradiene-3,17-dione derivatives resulting from the reaction of 2,4-dichloro-E1 and sodium hypochlorite (Nakamura et al., 2007). Ten oxidation products of E2 were identified during treatment with chloramine in a water distribution system. The identified products were formed by addition or loss of a functional group at different positions. The proposed reactions include electrophilic substitution of the ortho-position of the phenol moiety, elimination reaction between C9 and C11, and a hydrogenation reaction on the benzene ring (He et al., 2016). In general, biodegradation studies on halogenated estrogens are very scarce. Recently, the findings of Griffith et al. (2023) on biodegradation kinetics of free and halogenated estrogens in river water-sediment microcosms suggested that halogenated estrogens degrade more slowly than the corresponding free estrogens. Halogenated E1 derivatives were detected as transformation products of halogenated E2. While detection of halogenated E1 derivatives suggests subsequent metabolism via the most common 4,5-seco pathway, further studies are needed to unravel how dehalogenation of the A-ring occurs before hydroxylation and ring cleavage. Moreover, biodegradation kinetics and pathways in wastewater microcosms containing a mixture of free and halogenated estrogens is worth investigating.

Beside chlorination, advanced oxidation processes are being investigated as a wastewater treatment to eliminate endocrine-disrupting chemicals including estrogens. These processes include ozonation, photocatalysis, Fenton and Fenton-like processes, and they depend on the production of highly reactive hydroxyl radicals to oxidize organic pollutants (Wang and Wang, 2016; Dewil et al., 2017). There is also an interest in the development of chemical-free electrochemical oxidation processes (Dewil et al., 2017). As these processes depend on very strong oxidizing agents, namely hydroxyl radicals, they can lead to several hydroxylated and ring-cleavage transformation products. Even the ethinyl group of EE2 can be removed (Reis et al., 2023).

In the environment, estrogens are prone to direct and indirect photo-transformation reactions, which can play a role in estrogen removal (Chen et al., 2013; Adeel et al., 2017). The most common photo-transformation products of estrogen are hydroxylated derivatives bearing hydroxy groups at various positions (Mazellier et al., 2008; Li Puma et al., 2010; Caupos et al., 2011; Chen et al., 2013; Zhao et al., 2019). In addition, some ring cleavage products were also reported during estrogen photodegradation (Perondi et al., 2020). Susceptibility of estrogens to photodegradation can be exploited to develop concerted photocatalysis-biocatalysis technologies that can enhance estrogens removal, as shown for conjugated E2. Photocatalysis can contribute to deconjugation transformations which make free estrogens available for microbial degradation (Ding et al., 2024).